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IMPROVING THE PERFORMANCE OF Escherichia coli KO 11 DURING THE
FERMENTATION OF XYLOSE TO ETHANOL
STUART A. UNDERWOOD
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
Stuart A. Underwood
This work is dedicated to my wife, Beverly, and my family. The years of their endless
love and support made this work possible.
I would like to thank my mentor, Dr. Lonnie O. Ingram, for his insight and
guidance in my academic development. His inspiration and general enthusiasm for
science will have lasting effects on the development of my career.
My deepest appreciation is extended to the other members of my graduate
committee: Dr. K. T. Shanmugam, Dr. Julie A. Maupin-Furlow, Dr. Gregory W. Luli and
Dr. Jon D. Stewart. Without their guidance and sagacity, this work would not have been
I am grateful to Lorraine Yamano, Dr. Shengde Zhou and Dr. Fernando
Martinez-Morales for their help in learning the intricacies of molecular biology. Many
thanks to Sean York and Alfredo Martinez for their help in learning our fermentation
processes. I would also like to thank Dr. Marian L. Buszko for his guidance with NMR
My warmest thanks to the other members of Dr. Ingram's laboratory and the
biomass research group for the many insightful discussions about scientific matters.
The text and figures in Chapter 2 and Chapter 3, in part or in full, are reprints of
the material as it appears in Applied and Environmental Microbiology (vol. 68, pp.
1071-1081 and pp. 6263-6272, respectively).
TABLE OF CONTENTS
ACKNOWLEDGMENTS ................. .......................... iv
LIST OF TABLES ............... .............................. viii
LIST OF FIGURES .......... .. ...................... .. .......... ix
ABSTRACT ............ ............................................. xi
1 INTRODUCTION ............ ......................... ...... 1
Lignocellulose as a Carbohydrate Source ............................ 4
Adaptation to High Sugar Environments ............................. .. 5
Xylose versus Glucose Metabolism ................ ................. 8
Pyruvate Dissimilation ............... .......................... 10
Engineering E. coli for Ethanol Production ............................. 15
Deleterious Effects of Metabolic Engineering ........................... 16
Project Goals .......... .. ...................... ............ 18
2 FLUX THROUGH CITRATE SYNTHASE LIMITS THE GROWTH OF
ETHANOLOGENIC Escherichia coli KO 11 DURING XYLOSE
FERMENTATION ................................... ............ 26
Introduction ........... ........................................26
Materials and Methods ..........................................27
Microorganisms and Media ................ ................... 27
Fermentation Conditions .................................... 28
Aerobic Growth Studies ........................................29
Analytical M ethods ........................................... 29
G enetic M ethods .............................. ..... ........ 30
NAD(P)H/NAD(P)+ ratio .......................................31
Enzyme Assays .............. .................... ......... 31
Results ................... ...................... ............. 32
Macro-nutrient Limitation. ......... ...... ................... 32
Energy Limitation ............. ...... .......................... 33
Metabolic Imbalance Relieved by Addition of Pyruvate or Acetaldehyde. 33
Pyruvate as a Source of Carbon Skeletons for Biosynthesis ............. 35
Whole-cell Fluorescence ....................................... 37
Citrate Synthase, a Link Between NADH and 2-Ketoglutarate. .......... 39
Discussion ........... .........................................40
3 GENETIC CHANGES TO OPTIMIZE CARBON PARTITIONING IN
ETHANOLOGENIC Escherichia coli KO11 ........................... 53
M materials and M ethods ............................................ 54
Microorganisms and Media ................ ................... 54
Ferm entation ............ ........................ ........ 54
Analytical M ethods ........................................... 54
Genetic Methods .............. ................................ 55
Construction of pLOI2065 Containing a Removable Tetracycline Resistance
Cassette .............. .................... .............56
Nucleotide Sequence Accession Number ......................... 56
Construction of SU102 Containing an Insertion Mutation in ackA ........ 56
Construction of SU104 Containing a Deletion in adhE ................. 57
Results and Discussion ............................................ 58
Acetate Addition Stimulates Growth and Ethanol Production by Reducing Net
Acetate Production During Sugar Metabolism .................... 58
Stimulation of Growth and Ethanol Production by Added Pyruvate Can Be
Primarily Attributed to Increased Acetate Production. .............. 59
Stimulation of Growth and Ethanol Production by Acetaldehyde Can Be
Attributed to Increased Acetyl-CoA. .......................... 62
Stimulation of Growth and Ethanol Production by Inactivation of Non-
biosynthetic Pathways Which Consume Acetyl-CoA ............... 64
Conclusions ........... ........................................66
4 A DEFICIT IN PROTECTIVE OSMOLYTES IS RESPONSIBLE FOR THE
DECREASED GROWTH AND ETHANOL PRODUCTION DURING XYLOSE
FERMENTATION .............. .............................77
Introduction ............... .............................. 77
M materials and M ethods ............................................ 79
Microorganisms and Media. ................. ................ 79
Fermentation. ........... ................... ............ 79
13C NMR. ........... .................... ............ 80
Analytical Methods .................. ............ ........... 81
Results and Discussion .......... ............. ....... 81
Citrate Synthase Flux Limits the Biosynthesis of Glutamate, a Primary
Intracellular Osmolyte .................. ...................... 81
Genetic Changes to Optimize Carbon Partitioning Increased the Glutamate
Pool ............................................... 84
Glutamate Accumulation Functions in Osmoprotection ................ 85
Replacement of Glutamate by Other Osmoprotectants ................. 86
Betaine from Difco Yeast Extract Restores Growth in Luria Broth
Conclusions ............... ...............................88
5 GENERAL DISCUSSION AND CONCLUSIONS ...................... 96
Increased Acetyl-CoA Availability Stimulated Growth ................... 96
Some TCA Intermediates Increase Growth. ............................ 98
Citrate Synthase-A Unifying Hypothesis. ........................... 99
Future Prospects for Metabolic Engineering ........................... 104
REFERENCES ............. .......................................... 106
BIOGRAPHICAL SKETCH ............................................ 119
LIST OF TABLES
Table pa e
2-1. Effects of additives on the composition of fermentation products ............. 43
2-2. Effects of additives on growth and ethanol production by KO 11 .............. 44
3-1. Strains and plasmids used in Chapter 3 .................................. 68
3-2. Effects of mutations and additives on cell yield and ethanol productivity ........ 69
4-1. Intracellular accumulation of protective osmolytes by KO 11 ................. 90
LIST OF FIGURES
1-1. Glucose transport by the phosphotransferase system. ....................... 20
1-2. Glycolysis. ....................................................... .21
1-3. Xylose transport in E. coli ................ ......................... 22
1-4. Xylose metabolism .......... ................... ............23
1-5. Reactions of the pyruvate dehydrogenase complex ....................... 24
1-6. Fermentation pathways ofE. coli ................ ................... 25
2-1. Comparison of maximal cell densities ............... ................ 45
2-2. Comparison of growth and ethanol production from glucose and xylose ........ 46
2-3. Effects of added pyruvate and acetaldehyde .......................... 47
2-4. Initial effects of added TCA pathway intermediates ..................... .. 48
2-5. Effect ofmetabolites on whole-cell fluorescence ....................... 49
2-6. B. subtilis citZ increases the growth and ethanol production .................. 50
2-7. Relationship between cell yield and fermentation performance ................ 51
2-8. Fermentation and TCA pathway ....................................... 52
3-1. Allosteric control of central metabolism ................................. 70
3-2. Plasmids used to construct mutations ................................... 71
3-3. Effect of media additions and mutations ................................. 72
3-4. Effect of media additions and mutations on organic acid production ........... 73
3-5. Metabolism of added acetaldehyde and pyruvate during fermentation .......... 75
3-6. Partitioning of carbon among competing pathways ......................... 76
4-1. Carbon flow during fermentation of xylose by E. coli KOl l.................. 91
4-2. Major intracellular osmolytes accumulated by ethanologenic E. coli during
fermentation .................................... .. ...........92
4-3. Effect of osmoprotectants (1.0 mM) on maximum cell concentration ........... 93
4-4. Effects of Betaine and DMSP on growth and ethanol production .............. 94
4-5. The chemical structure ofbetaine and DMSP ............................. 95
Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
IMPROVING THE PERFORMANCE OF Escherichia coli KO 11 DURING THE
FERMENTATION OF XYLOSE TO ETHANOL
Stuart A. Underwood
Chair: Lonnie O. Ingram
Co-Chair: Keelnatham T. Shanmugam
Major Department: Microbiology and Cell Science
The large-scale conversion of lignocellulose to fuel ethanol would greatly reduce
the U.S. dependence on imported oil. To facilitate this need, Escherichia coli has been
genetically engineered for the homofermentative production of ethanol from all
constituent sugars of lignocellulose. However, high levels of complex nutrients are
required for rapid fermentation of xylose, the second most abundant sugar in
lignocellulose. With low levels of complex nutrients, the rate of xylose fermentation was
limited by the growth of the biocatalyst. In a mineral salts medium containing 1% corn
steep liquor as a nutrient source (90 g liter1 xylose), growth was limited by an imbalance
in the partitioning of carbon between ethanol production and biosynthetic pathways.
Citrate synthase was shown to catalyze the specific growth-limiting reaction. The
allosteric controls of citrate synthase regulate carbon flow through the oxidizing arm of
the TCA pathway, ultimately producing 2-ketoglutarate and glutamate. Functionally
expressing citrate synthase II (citZ) from Bacillus subtilis stimulated growth due to its
different allosteric and kinetic properties. Acetyl-CoA server as an antagonist to the
NADH-mediated allosteric inhibition of the E. coli citrate synthase. Supplementing the
medium with pyruvate, acetate, acetaldehyde, 2-ketoglutarate or glutamate increased
growth and ethanol production by activating, relieving or bypassing the allosteric
regulation of the E. coli citrate synthase. Conservation of acetyl-CoA by mutating acetate
kinase (AackA) also increased growth and ethanol production, presumably by increasing
the availability of acetyl-CoA (activating citrate synthase). In addition to biosynthetic
needs, large intracellular pools of glutamate (>20 mM) function as a protective osmolyte.
During growth in the high osmotic environment of the corn steep liquor medium
containing 0.6 M xylose, intracellular glutamate was low (< 10 mM) and cells grew
poorly, consistent with a glutamate deficiency. The addition of glutamate to the medium
and all approaches that stimulated citrate synthase increased the high intracellular pool of
glutamate during growth in this medium. Supplementing with other protective osmolytes,
such as betaine and dimethylsulfoniopropionate, restored growth without affecting the
intracellular pool of glutamate and appear to act directly as alternative osmolytes. These
results indicate that the poor growth and ethanol production in 1% cor steep liquor
medium (0.6 M xylose), the apparent requirement for high levels of nutrients without a
specific auxotrophic requirement and the beneficial effects of increased intracellular
glutamate all result from the requirement for high levels of protective osmolytes. Under
these conditions, the growth of the biocatalyst (E. coli) and ethanol production are limited
by insufficient levels of intracellular osmoprotectants rather than the synthesis of
glutamate, per se.
The production of fuel ethanol from renewable feedstocks could potentially
decrease the U.S. dependance on imported oil as well as decrease the release of fossilized
carbon into the atmosphere as carbon dioxide (CO2), a greenhouse gas. Blends of 95%
ethanol with gasoline are effective motor fuels, as demonstrated by Brazil's use of such
blends for more than 20 years prior to securing inexpensive sources of fossil fuels. In the
year 2002, approximately 140 billion gallons of gasoline were consumed in the United
States, most of which was derived from foreign oil. Approximately 2.9 billion gallons of
ethanol are produced annually in the U.S., slightly more than 2% of the gasoline
consumed. While the volume of ethanol produced increases each year, demands for
ethanol and energy also increase. For example, the phasing out of the gasoline oxygenate
methyl tertiary-butyl ether (MTBE) over the next several years will further increase the
demand for fuel ethanol, an alternate oxygenate. A substantial increase in ethanol
production must be achieved to replace MTBE with 10% ethanol.
Today, most of the ethanol derived from fermentation uses cornstarch as the
feedstock with yeast as the biocatalyst. Competing demands for cornstarch and variable
crop yields cause price volatility. Feedstock is the major contributor to the cost of current
ethanol processes. The cost of ethanol production must remain low in order for it to be an
economically competitive automobile fuel. The necessity for a less expensive, lower
demand feedstock is obvious. Agricultural wastes (corn stover, sugarcane bagasse, wheat
straw, etc.) are relatively inexpensive sources of carbohydrates that can be converted to
ethanol (Amtzen and Dale 1999; Ingram and Doran 1995; Ingram et al. 1999; Zaldivar et
al. 2002). More than 200 billion gallons of ethanol could be produced using these
lignocellulosic materials, sufficient to replace all of the gasoline burned by automobiles in
the United States (Arntzen and Dale 1999). As these agricultural wastes have little or no
competing uses, they offer long-term solutions to the necessity for inexpensive
However, there is no known organism in nature capable of fermenting all of the
various hexose and pentose components ofbiomass to ethanol. This difficulty is further
compounded by the complex, polymeric and somewhat variable structure of the
lignocellulosic biopolymers (Clarke 1997). Harsh treatments are required to breakdown
these sugar polymers into suitable substrates for fermentation. During these processes,
furfural, hydroxymethylfurfural, acetate, and many other cytotoxic byproducts are
released into the resulting solutions. An organism must tolerate the environmental
conditions created by these treatments to be an effective biocatalyst. With advances in
molecular biology, genetically engineering a desirable microorganism to produce ethanol
should be possible.
There are essentially two approaches to engineering an organism for the
production of ethanol from lignocellulosic residues. Either an ethanol producing
microorganism could be engineered to use all of the various sugars or a microorganism
already capable of fermenting all of these sugars could be engineered to produce
exclusively ethanol. The former approach has been pursued by many groups through the
engineering ofSaccharomyces cerevisiae or Zymomonas mobilis (deficient in pentose
metabolism) to utilize these carbohydrates by expressing heterologous transport and
metabolic pathways (Aristidou and Penttila 2000; Chotani et al. 2000; Gong et al. 1999).
While high productivities have been reported for both organisms in optimal conditions,
yeasts capable of fermenting both xylose and arabinose have not been reported in the
literature. Z. mobilis, a very fastidious organism, is not environmentally hardy, and the
harsh conditions resulting from the pretreatment of the lignocellulose severely hinders its
One of the most studied and characterized organisms, Escherichia coli is an
excellent candidate for genetic engineering. The complete genetic sequence has been
published (Blattner et al. 1997), and much is known about its physiology (Neidhardt et al.
1990). The utility of this organism in industrial processes is second only to yeast. Typical
of enteric organsism, E. coli is capable of fermenting both the pentoses and hexoses
present in lignocellulose. However, E. coli is a mixed acid fermenter, producing lactate,
acetate, ethanol, format and succinate as its major fermentation products. Previous work
in our laboratory engineered the metabolism ofE. coli to produce exclusively ethanol
(Ohta et al. 1991).
The sugars of hemicellulose hydrolysates, containing mostly xylose, were
fermented by the engineered E. coli strain, with yields approaching 100% (0.51 g ethanol
/ g sugar) (Asghari et al. 1996; Lawford and Rouseau 1996; Martinez et al. 1999; York
and Ingram 1996a; York and Ingram 1996b). However complex, expensive nutrients
(Luria broth) are required to obtain these high yields. High levels of inexpensive nutrients
are required to replace these rich nutrients (Asghari et al. 1996; Lawford and Rouseau
1996; Martinez et al. 1999; York and Ingram 1996a; York and Ingram 1996b), but this
creates waste management problems and increases cost. Fermentations which use low
levels of complex nutrients or no nutritional supplements would be most desirable for
Corn steep liquor (CSL), a by-product from the wet milling of corn, is an
inexpensive nutrient source with demonstrated utility in industrial processes.
Fermentations of hemicellulose hydrolysate with CSL as the nutrient source exhibited
dose-dependent change in ethanol productivities (Martinez et al. 1999). To equal the
ethanol productivity achieved with Difco nutrients (5 g liter' yeast extract and 10 g liter'
tryptone), 50 g liter-' CSL (wet weight; 50% solids) were required. The goal of this
present study is to understand the basis of the need for complex nutrients and develop
physiological and genetic solutions to circumvent this requirement.
Lignocellulose as a Carbohydrate Source
Most of the dry weight biomass is lignocellulose, composed of cellulose,
hemicellulose, pectin and lignin (Clarke 1997). Cellulose, the most abundant polymer on
the planet, is a homopolymer of cellobiose (P-1,4-glucose) and represents 20-50% of the
dry weight of plant matter. Lignin is a polymer of aromatic alcohols, comprising 10-20%
of the dry weight of plant biomass. Representing only 1-10% of the dry weight, pectin is a
methylated homopolymer of galacturonic acid. Hemicellulose is a complex, branched
polymer ofhexoses (glucose, galactose, mannose, rhamnose, and fucose) and pentoses
xylosee and arabinose). This polymer represents 20-40% of the plant dry weight and is the
most easily solubilized component of lignocellulose.
The sugars ofhemicellulose are released as monomers through a variety of
hydrolysis procedures, but dilute acid hydrolysis is currently the preferred method
(Ingram et al. 1999). This procedure uses moderate heat and low pH to release the sugars
of hemicellulose into solution as monomers (Grohmann et al. 1985). The exact ratio of
sugars in these hydrolysates can vary considerably depending on the feedstock, but xylose
is the most prevalent sugar in hydrolysates of hard woods and grasses (sugarcane, wheat
straw, etc.). Generating a concentrated sugar solution during hydrolysis is a formidable
challenge, but a goal of 100 g liter-' total sugar monomers in hemicellulose hydrolysates
should be achievable. Most of the studies presented here have used 90 g liter-' xylose as
the fermentation substrate.
Adaptation to High Sugar Environments
Many industrial fermentation processes operate as either batch fermentations
(with all required nutrients and substrates supplied initially) or fed-batch fermentations
(multiple additions of nutrients; requires concentrated feed solutions). As the hydrolysis
ofhemicellulose produces sugar streams up to 100 g liter-' (Ingram et al. 1999), their
fermentation to ethanol favors a batch fermentation process to avoid the additional cost of
concentrating these sugar streams and potentially concentrating growth inhibitory
compounds. However, this relatively high sugar concentration requires E. coli to adapt to
this higher osmolarity.
The rapid accumulation of potassium is the first response ofE. coli and related
organisms to an increase in the osmotic strength of the medium. Within a minute after an
increase in osmotic pressure, glutamate (a negatively-charged amino acid) synthesis is
increased to provide charge balance for the accumulated potassium (McLaggan et al.
1994). The short time between the accumulation of potassium and the biosynthesis of
glutamate, suggests that the onset of glutamate biosynthesis is a result of allosteric
regulation (<5 min) rather than genetic induction (10-20 min). Additionally, the
accumulation of glutamate in response to osmotic stress was found to be dependent on
the presence of K in the medium (McLaggan et al. 1994).
Escherichia coli has two biosynthetic pathways for glutamate. Under a nitrogen
limitation (0.1 mM ammonium), glutamate synthase-glutamine synthetase has been
shown to be the predominant glutamate biosynthetic pathway (Pahel et al. 1978). During
growth in excess nitrogen, glutamate dehydrogenase (GDH), a pathway that does not
consume ATP, is the primary glutamate biosynthetic pathway (Helling 1994).
Additionally, GDH is activated by K (Measures 1975). This allosteric regulation of GDH
has been proposed to be responsible for osmotically activated glutamate biosynthesis
The intracellular concentration of K can be as high as 800 mM in E. coli during
growth in media of high osmolarity (Cayley et al. 1991; Cayley et al. 1992). Cells
deficient in glutamate accumulation have demonstrated growth defects during osmotic
challenge (Csonka 1988; McLaggan et al. 1991; Yan et al 1996) due to an inability to
maintain sufficient K (Yan et al. 1996). The large increases in intracellular potassium
and glutamate are transient, and their levels begin to decrease to 20-50 mM as trehalose
or other protective osmolytes accumulate in the cytoplasm (Dinnbier et al. 1988; Giaever
et al. 1988). However, glutamate pools remain elevated during growth the higher osmotic
conditions (Yan et al. 1996).
For the long-term adaptation to media of high osmolarity, E. coli synthesizes
trehalose (Boos et al. 1990; Dinnbier et al. 1988; Giaever et al. 1988) or accumulates
other charge-neutral (zwitterionic) compatible solutes (betaine, proline ectoine,
dimethylsulfonioproprionate, etc.) (Csonka and Hanson 1991). E. coli has a limited
capacity for biosynthesis of these compounds. Although E. coli is incapable of de novo
betaine biosynthesis, choline can be oxidized to betaine. However, this process is
restricted to aerobic growth (Landfald and Strom 1986). Some organisms synthesize
proline for long-term osmoadaptation (Kawahara et al. 1989). However, the y-glutamyl
kinase step in proline biosynthesis is subject to strong feedback inhibition in E. coli,
preventing the biosynthesis of this protective osmolyte (Csonka 1988; Smith 1985; Smith
et al. 1984). Thus, many of the protective osmolytes accumulated by E. coli must be taken
from their environment.
E. coli and related organisms have two primary transport systems for protective
osmolytes during osmotic stress, ProP and ProU (Randall et al. 1995). The ProP system
uses the proton gradient maintained by the cell to drive the uptake ofosmoprotectants.
This low affinity system (Km for proline is 0.3 mM) also transports many other
osmoprotectants (Lucht and Bremer 1994). The ProU transport system consists of a
periplasmic binding protein with a high affinity for betaine (Km 1.3 [IM), a
membrane-spanning component and a membrane bound enzyme which hydrolyzes ATP
for the active transport of betaine (Lucht and Bremer 1994).
A hierarchy for osmoprotectants has been empirically established for E. coli,
primarily for salt-mediated osmotic stress (Randall et al. 1995). Although there have been
conflicting reports concerning the validity of this hierarchy for sugar-mediated osmotic
stress (Glaasker et al. 1998), betaine is generally regarded as the most effective protective
osmolyte for E. coli. In at least one report, the ability of betaine to restore growth during
osmotic challenge with different carbon sources was dependent on the particular sugar
(Dulaney et al. 1968). Thus, the sugar-mediated osmotic stress anticipated for
fermentations of hemicellulose hydrolysates (100 g liter-' sugar) may require the
accumulation of different osmolytes.
Xylose versus Glucose Metabolism
The reactions involved in the transport and metabolism of glucose are well
understood and outlined in Figures 1-1 and 1-2. Transport of glucose into the E. coli
cytoplasm is mediated by a phospho-transferase system (PTS). The energy and phosphate
required for translocation and phosphorylation of PTS sugars comes from
phosphoenolpyruvate (PEP). An additional ATP is required for the phosphorylation of
fructose-6-phosphate to fructose-1,6-bisphosphate. Thus, to metabolize glucose, an initial
investment of 2 ATP equivalents (1 ATP and 1 PEP) is required.
Fructose-1,6-bisphosphate is cleaved into dihydroxyacetone-phosphate and
glyceraldehyde-3-phosphate. These two molecules are interconverted via triose-phosphate
isomerase. For the production of pyruvate, the terminal product of glycolysis,
glyceraldehyde-3-phosphate is oxidized and phosphorylated to form
1,3-bisphosphoglycerate by glyceraldehyde-3-phosphate dehydrogenase. During this step,
nicotinamide adenine dinucleotide (NAD ) is reduced to NADH. The high group-transfer
potential of the phosphate bond on carbon 1 is used in the production of ATP from ADP
in the proceeding reaction catalyzed by phosphoglycerate kinase. The reactions leading to
the formation of phosphoenolpyruvate do not result in any further energy yield or
In converting glyceraldehyde-3-phosphate to PEP, 1 ATP and 1 NADH are
produced. The conversion of PEP to pyruvate is either carried out via the
phosphotransferase system or through an ATP yielding reaction catalyzed by a pyruvate
kinase. The net reaction ofglycolysis can be written as follows:
glucose + 2NAD+ + 2ADP + 2 Pi 2 pyruvate + 2NADH + 2H' + 2 ATP
The production of pyruvate from 1/2 molecule glucose yields a net of 1 ATP and 1
reducing equivalent (NADH). Reducing equivalents are often considered as pools of both
their reduced and oxidized forms, and their ratio is indicative of the metabolic state of the
cell (respiration or fermentation) (de Graef et al. 1999; Snoep et al. 1990). Though the
ratio of the reduced to oxidized form (NADH/NAD+ ratio) varies widely with different
growth conditions, the absolute concentration of the two forms remains relatively
constant (de Graef et al. 1999).
In contrast to glucose, xylose is transported into the cell by either a proton
symport pathway (xylE) or an ATP dependant transporter (xylFGH) (Song and Park 1997;
Tao et al. 2001; Fig. 1-3). During fermentation, the cellular proton gradient is maintained
presumably by energy-consuming reactions (F1/F0 ATPase, for example). Thus, a proton
symport pathway is fueled indirectly by the hydrolysis of ATP. The transport of each
xylose is energized by the hydrolysis of 1 ATP. Once inside the cell, xylose is converted
into xylulose by xylose isomerase. Xylulose is then phosphorylated by xylulokinase,
utilizing the hydrolysis of a second ATP. Regardless of the pathway, xylose uptake and
activation phosphorylationn) require energy derived from the hydrolysis of 2 ATP
molecules. In contrast, glucose transport uses a single ATP equivalent (PEP) for both
transport and activation.
Intracellular xylulose-5-phosphate is metabolized by the pentose-phosphate
pathway (Fig 1-4). Through a series of reactions catalyzed by transketolase and
transaldolase, xylose is converted into intermediates of glycolysis (fructose-6-phosphate
and glyceraldehyde-3-phosphate). For every 6 xyloses consumed (30 carbon atoms), 4
fructose-6-phosphates and 2 glyceraldehyde-3-phosphates are produced. These molecules
are further metabolized by glycolysis to ultimately yield 10 molecules of pyruvate. Thus,
all 30 carbon atoms which began in xylose are converted into pyruvate.
The energy (ATP) and reducing equivalents (NADH) produced from the reactions
common to xylose and glucose metabolism are the same regardless of the original
substrate. However, since xylose is a pentose and requires separate energy for transport
and activation, growth on xylose results in a relatively low ATP yield. The transport and
activation of 6 xylose molecules (30 carbons) requires 12 ATPs, assuming 1 ATP is
required for transport regardless of the pathway. In the conversion of this
xylulose-5-phosphate to 10 molecules of glyceraldehyde-3-phosphate, an additional 4
ATPs are consumed. The conversion of these 10 molecules of
glyceraldehyde-3-phosphate to pyruvate yields 20 ATPs. The net gain of energy in the
conversion of 6 molecules of xylose to 10 pyruvate is 4 ATPs. An equal amount of
glucose on the basis of moles of carbon (5 molecules; 30 carbon atoms) produces a net of
10 ATPs during conversion to 10 molecules of pyruvate. The net energy gain for glucose
catabolism is 1 ATP per pyruvate, 2.5-fold more ATP than from xylose catabolism.
While there is a considerable difference in ATP yield between xylose and glucose, only
one NADH is produced per pyruvate from either glucose and xylose.
A facultative anaerobe, E. coli accomplishes redox balance either by respiration
(Gennis and Stewart 1996) or fermentation (Bock and Sawers 1996). During respiration,
reducing equivalents are oxidized when their electrons are donated to the primary
oxido-reductases of the electron transport system. These electrons are passed between the
various proteins of the electron transport chain and ultimately used to reduce a terminal
electron acceptor (oxygen during aerobic respiration, for example). The energy from these
reducing equivalents is preserved in the form of a proton gradient established by the
concomitant translocation of H' from the cytosol to the periplasm. This proton gradient is
used to produce ATP via the F1/F0 ATPase.
The pyruvate dehydrogenase complex (PDH) catalyzes the oxidative metabolism
of pyruvate to acetyl-Coenzyme A (acetyl-CoA) and CO2, with the formation of 1
reducing equivalent (NADH). This complex consists of three activities: pyruvate
decarboxylase (aceE), acetyltransferase (aceF), and lipoate dehydrogenase (Ipd) (Fig.
1-5). The decarboxylation of pyruvate to an enzyme-bound acetyl moiety by the pyruvate
decarboxylase of this complex requires a thiamine pyrophosphate (TPP) cofactor, a
carrier of the "active" acetaldehyde. The acyl moiety is then transferred to an acyl-carrier
through the reduction of a disulfide bond. The resulting thioester has a high group transfer
potential and is transferred to Coenzyme A, an acetyltransferase reaction. The disulfide
which accepts the acetaldehyde from the TPP must be regenerated through an
oxidation/reduction reaction. NAD+ is reduced to NADH as the sulfhydryl group is
oxidized, forming the required disulfide bond. This reaction is subject to strong feedback
inhibition by NADH (Hansen and Henning 1966).
During aerobic growth, the tricarboxylic acid (TCA) cycle is responsible for the
total oxidation of acetyl-CoA to CO2 (Cronan, Jr. and LaPort 1996). The first step in this
cycle, citrate synthase, is also the rate controlling step (Lee et al. 1994; Walsh and
Koshland, Jr. 1985). This enzyme catalyzes the condensation of acetyl-CoA and
oxaloacetate to form citrate (Weitzman 1981). Citrate synthase is primarily regulated by
allosteric controls, activated by acetyl-CoA and inhibited either by NADH and 2-
ketoglutarate (Gram-negative) or ATP (Gram-positive, archea and eukaryotes) (Weitzman
1981). This provides a link between the energetic needs of the cell and the generation of
reducing equivalents (and ultimately ATP) through the TCA cycle.
The TCA cycle is also a source of carbon skeletons for biosynthesis. More than
half of the amino acids made by the cell are derived from intermediates of the TCA cycle
(Neidhardt et al. 1990). Oxaloacetate must be regenerated for the continued cyclic action
as intermediates are drawn into biosynthesis. This anapleurotic reaction is catalyzed by
phosphoenolpyruvate carboxylase in E. coli. The biosynthetic needs and metabolic state
of the cell dictate the activity of this reaction through allosteric control. Acetyl-CoA,
fructose-1,6-bisphosphate and GTP are activators of this enzyme (Izui et al. 1981), while
malate and aspartate (products of oxaloacetate utilizing reactions) are inhibitors (Izui et
al. 1981). Acetyl-CoA and oxaloacetate are co-substrates for citrate synthase. Thus, the
allosteric activation of phosphoenolpyruvate carboxylase and citrate synthase by
acetyl-CoA links the availability of the two co-substrates for citrate synthase.
During fermentation, no external terminal electron acceptors are available for
respiration, resulting in the accumulation of NADH to higher levels than during
respiration (de Graef et al. 1999). For redox balance to be maintained, intracellular
metabolites serve as electron acceptors. As NAD+ regeneration becomes difficult, NADH
generation is not favored. The formation of acetyl-CoA from the pyruvate dehydrogenase
reaction is inhibited by the high NADH/NAD+ ratio (Hansen and Henning 1966),
necessitating an alternate, non-oxidative route to acetyl-CoA generation. The
non-oxidative cleavage of pyruvate to acetyl-CoA and format is catalyzed by pymvate
formate-lyase (PFL; Knappe and Sawers 1990).
PFL activity is relative to the metabolic state of the cell, similar to PDH and
citrate synthase. However, PFL activity is regulated by post-translational modification
enzymes which are allosterically controlled. The protein is translated in an inactive form.
An oxygen-labile free radical is placed on a glycine residue by a PFL-activase enzyme
(pflA) forming the active PFL enzyme (Conradt et al. 1984). To protect the enzyme from
irreversible inactivation by oxygen, the multi functional alcohol dehydrogenase (adhE)
also has a PFL-deactivase activity to remove the oxygen-labile free radical (Kessler et al.
1991). The PFL-deactivase activity of adhE is inhibited by NADH (Kessler et al. 1992),
linking the activation state of PFL to the metabolic state of the cell as described by the
NADH/NAD+ ratio. When oxygen supply is limited, NADH accumulates and inhibits
PDH and the PFL deactivase activity. This causes a shift in flux to acetyl-CoA from
oxidative pyruvate cleavage (PDH) to the non-oxidative cleavage (PFL).
In contrast to respiration, acetyl-CoA is an electron acceptor during fermentation.
The two-step reduction of acetyl-CoA to ethanol is catalyzed by alcohol dehydrogenase
(adhE; Fig. 1-6), regenerating 2 NAD Glycolysis produces only one NADH per
pyruvate. Thus, the native alcohol production pathway results in an NADH deficit. This is
overcome by converting one of the acetyl-CoA to acetate, producing an additional ATP
by substrate-level phosphorylation. In E. coli grown under anaerobic fermentation
conditions with glucose as the carbon and energy source, equal amounts of acetate and
ethanol are produced.
Lactic acid is often produced by E. coli during fermentation in addition to acetate
and ethanol, primarily as active growth slows and stationary growth. Pyruvate is reduced
in a single-step reaction catalyzed by lactate dehydrogenase (LDH, IdhA gene product;
Bunch et al. 1997), resulting in the re-oxidation of 1 NADH per lactate produced. The
pathway for lactate production in E. coli is controlled by allosteric regulation, activated
by pyruvate (Tarmy and Kaplan 1968). In conditions of surplus supply of pyruvate, the
lactate pathway is activated. There is an associated energetic loss as a result of lactate
production compared to the co-production of acetate and ethanol, as no ATP is made in
the reduction ofpyruvate to lactate.
In contrast to respiration, the TCA cycle is interrupted at 2-ketoglutarate
dehydrogenase due to transcriptional regulation during fermentation (Iuchi and Lin 1988).
The resulting pathway has two sides, the reductive (leading to succinate production) and
the oxidative (stopping at 2-ketoglutarate). For succinate production during fermentation,
the anapleurotic pathway for oxaloacetate production (PPC) is the first step. As described
previously, there are multiple allosteric effectors of this enzyme which control its
physiological activity. Oxaloacetate is converted to malic acid through the reverse activity
of the malate dehydrogenase, regenerating 1 NAD+ (Bock and Sawers 1996). Fumarase
catalyzes the conversion ofmalate to fumarate. Fumarate is reduced to succinate by a
fermentation specific fumarate reductase (frdBACD gene products) with the oxidation of
a reduced menaquinone (Cronan, Jr. and LaPort 1996). The oxidative side of the TCA
pathway provides carbon skeletons for biosynthesis (Neidhardt et al. 1990).
Engineering E. coli for Ethanol Production
The enteric bacterium E. coli can use all of the sugar constituents of
lignocellulose, while the natural ethanol producing Saccharomyces cerevisiae and
Zymomonas mobilis are limited to growth on hexoses,. Wild-type E. coli produces
ethanol from the two step reduction of acetyl-CoA oxidizing two NADH to NAD As a
result, acetate (no further reduction required) is made in approximately equal amounts to
ethanol. However, Z. mobilis and S. cerevisiae produce ethanol from pyruvate through a
pathway which only re-oxidizes one NADH. The irreversible, non-oxidative cleavage of
pyruvate into acetaldehyde and carbon dioxide is catalyzed by pyruvate decarboxylase
(PDC). Acetaldehyde is reduced to ethanol, oxidizing 1 NADH. Thus, for each pyruvate
that is converted to ethanol via this pathway, one NADH is re-oxidized. With this
stoichiometry, all of the pyruvate generated by glycolysis can be converted to ethanol
without the necessity of other oxidized products to maintain redox balance.
Previous studies demonstrated that the Z. mobilis genes involved in ethanol
production are expressed well in E. coli (Ingram and Conway 1988). These genes (pdc
and adhB) were used to construct a synthetic operon which was integrated into the
chromosome for increased genetic stability of the recombinant strain (Ohta et al. 1991). A
deletion was introduced in fumarate reductase to decrease succinate production,
problematic in xylose fermentation. The resultant strain, designated K 11, fermented
both pentoses and hexoses to ethanol with yields approaching 100% of total sugars
present (0.51 g ethanol/g sugar = 100% theoretical yield) during fermentation in
laboratory media containing excess complex nutrients. In addition to the alterations to the
fermentation profile, there were some notable effects on growth physiology. In broth
cultures, comparatively high cell yields were achieved. On solid media, colonies
exhibited a raised morphology, similar to yeast.
Deleterious Effects of Metabolic Engineering
The engineering of metabolic pathways for the production of industrial chemicals
as an alternative to chemical synthesis has been performed for a variety of chemicals
(Chotani et al. 2000). Metabolic engineering for renewable chemicals such as ethanol
(Ingram et al. 1999), acetate (Causey et al, 2003), lactate (Bianchi et al. 2001; Chang et
al. 1999a; Dien et al 2001; Kyla-Nikkila et al. 2000; Zhou et al. 2002; Zhou et al. 2003),
propanediol (Nakamura et al. 2000; Tong et al. 1991), adipic acid (Niu et al. 2002) and
succinate (Donnelly et al 1998a; Donnelly et al. 1998b; Vemuri et al. 2002) have focused
primarily on product yields. The metabolic engineering of these new products has often
resulted in unexpected changes which increased the need for complex nutrients and
decreased potential utility (Bunch et al. 1997; Chang et al. 1999a; Chao and Liao 1994;
Chao et al. 1993; Martinez et al. 1999).
Undesirable changes such as reduced growth, decreased glycolytic flux and low
volumetric productivity are generally attributed to a lack ofATP (Gokarn et al. 2000; Xie
et al. 2001), creation of futile cycles (Chao and Liao 1994; Chao et al. 1993; Patnaik et al.
1992), changes in intracellular metabolite pools or a metabolic imbalance (Aristidou et al.
1992; Bunch et al. 1997; Chang et al. 1999b; Contiero et al. 2000; Liao et al. 1996; Yang
et al. 1999a; Yang et al. 1999b; Zhou et al. 2002; Zhou et al. 2003). Often, these
detrimental effects are masked by abundant complex nutrients in laboratory media and are
only apparent in mineral salts or low-nutrient media (Bunch et al. 1997; Chao and Liao
1994; Chao et al. 1993; Martinez et al. 1999).
Salmonella typhimurium was engineered for succinate production by increasing
the expression ofpyc encoding pyruvate carboxylase (Xie et al. 2001). Although
succinate production increased, growth rate declined by 18% and glycolytic flux
decreased by 40%. Similar results were reported for an analogous construction in E. coli
(Gokarn et al. 2000). Donnelly and coworkers (1998a and 1998b) isolated E. coli mutants
which produced 5-times more succinate than the parent strain, and again growth rate was
impaired. Growth rate and cell yield were also decreased by engineering E. coli for the
production of 3-deoxy-D-arabinoheptulosonate 7-phosphate (DAHP) by over-expression
ofpps (phosphoenolpyruvate synthase) (Patnaik and Liao 1994; Patnaik et al. 1992). This
inhibition of growth was more pronounced in minimal medium (Chao and Liao 1994).
Acetate production during the aerobic growth ofE. coli on sugars has been
correlated with a decline in metabolic activity and reduced expression of heterologous
genes (Aristidou et al. 1995; Bauer et al. 1990; Chang et al. 1999b; Luli and Strohl 1990).
Many approaches have been employed to decrease acetate production and increase
recombinant products (Aristidou et al. 1995; Barbosa and Ingram 1994; Bauer et al. 1990;
Chang et al. 1999b; Contiero et al. 2000; Yang et al. 1999a). Mutations in the primary
acetate pathway (pta, phosphotransacetylase; ackA, acetate kinase) increased the yield of
recombinant proteins, but usually reduced cell growth. The detrimental effect on growth
was attributed to the accumulation of metabolic intermediates such as acetyl-CoA or
acetyl-phosphate. An alternative approach, channeling pyruvate away from acetate by
expressing the Bacillus subtilis alsA gene encoding acetolactate synthase, also reduced
acetate production by 80% and increased product yields, but again reduced cell growth
(Yang et al. 1999a). Other attempts to decrease acetate production by increased
expression of IdhA (lactate dehydrogenase) were ineffective in rich medium (Yang et al.
1999b). In mineral salts medium, over-expression of IdhA was accompanied by a severe
growth limitation (Bunch et al. 1997).
Lactate dehydrogenase (IdhA) has been expressed to divert carbon away from
acetate accumulation. Despite the relatively high Km for pyruvate, lactate production
increased by 50% (Yang et al. 1999b). Interestingly, the amount of acetate produced in
these fermentations was not altered. However, these studies were conducted in a rich
laboratory medium. In a mineral salts medium (M9), expression of LDH resulted in
severe growth defects (Bunch et al. 1997). These growth defects were attributed to a
decrease in pyruvate availability necessary for growth.
Though strain KO 11 is prototrophic, high levels of complex nutritional
supplements are required for the rapid fermentation of sugars to ethanol (Martinez et al.
1999). For example, during the fermentation of 90 g liter-' xylose to ethanol, the addition
of CSL as a nutritional supplement (0-50 g liter') had a dose dependent effect on final
cell concentration. With the increase in biocatalyst concentration, there was a
proportional increase in fermentation rate and decrease in required fermentation time.
Although the addition of 50 g liter' CSL is not cost-prohibitive, the handling of this much
material on the scale of an industrial fermentation could be problematic and generate
excessive cost for waste disposal.
These studies will examine the basis for the high nutrient requirement for KO 11
for the rapid conversion of sugar to ethanol. As a starting point, growth and ethanol
production will be evaluated in a medium containing 1% CSL, 90 g liter' xylose, and
mineral salts. Physiological and genetic approaches will be used to characterize the
growth limitation. Solutions for solving this limitation will be presented. Further work
will demonstrate the basis for a specific biosynthetic pathway under the conditions tested.
Knowledge gained in these studies will have applications in the further development of
the commercial production of ethanol from plant biomass by metabolically engineered E.
coli and will significantly contribute to the field of metabolic engineering by emphasizing
the importance of metabolic intermediates down-stream of the product forming node.
n-i vmil v.cZISCZ
Periplasm P-HPr nzyme PEP
cBrr pH ptS i
G Enzyme HPr nzyme I- Pyruvate
Figure 1-1. Glucose transport by the phosphotransferase system. The phosphate from PEP
passes through a cascade of enzymes and ultimately to intracellular glucose. The
hydrolysis of 1 ATP equivalent (PEP) is used to energize transport and activate
Figure 1-2. Glycolysis. The enzymes and genes that catalyze the conversion of glucose to
pyruvate are as follows: (1) phosphoglucose isomerase,pgi; (2)
phosphofructokinase,pflkA; (3) fructose-6-phosphate aldolase,fba; (4) triose
phosphate isomerase, tpi; (5) glyceraldehyde-3-phosphate dehydrogenase, gapA;
(6) phosphoglycerate kinase,pgk; (7) phosphoglycerate mutase, gpmA orpgml;
(8) enolase, eno; (9) phosphotransferase system; (10) pyruvate kinase, pykA or
H+ H+ H
\ xyl Outside
Xylose Xyl ATP H+ ADP
nH+ Xylulose Xylulose-SP
Xylose ATP ADP +P,
Figure 1-3. Xylose transport in E. coli. In contrast to glucose, the transport and activation
of xylose is not coupled, each requiring energy.
Figure 1-4. Xylose metabolism. Xylose is metabolized to intermediates of glycolysis (in
bold) by the pentose-phosphate pathway. For the sake of carbon balance, 6 xylose
are converted to 4 fructose-6-phosphate and 2 glyceraldehyde-3-phosphate. Note
that neither ATP nor reducing equivalents are produced or consumed in this
C -C -CH3
Enzyme I-TPP -c -CH,
Enzyme 2- R- I
oEny Enzyme 1-TPP
Enzm 2 -C-CH3
Enzyme 2 R.
Transa cety- tn Coe nzym e A-SH
Coenzyme A-S~ C -CH-
Enzyme 3- FAD
NADH + H+
Figure 1-5. Reactions of the pyruvate dehydrogenase complex. Enzyme I (aceE) catalyzes
the oxidative decarboxylation of pyruvate through a pyruvate dehydrogenase
activity. The thiamine pyrophosphate bound activated acetaldehyde is passed to
the lipoate transacetylase (Enzyme 2; aceF), and ultimately to Coenzyme A. To
regenerate the reduced lipoate, the FAD bound to Enzyme 3 (dihydrolipoate
dehydrogenase; Ipd) is reduced. NAD+ is reduced to NADH by this FADH,,
allowing for another cycle.
Enzyme 3 FADH2
Coenzyme A Acetyl-CoA
j 2 Lactate
Ace P [Acetalhyde]
Figure 1-6. Fermentation pathways of E. coli. (1) pyruvate kinase, pykA orpykF or PTS
sugar transport; (2) pyruvate formate-lyase, pflB; (3) phosphotransacetylase, pta;
(4) acetate kinase, ackA; (5) PFL-deactivase / alcohol dehydrogenase /
acetaldehyde dehydrogenase, adhE; (6) PEP carboxylase, ppc; (7) malate
dehydrogenase, mdh; (8) fumarase,fumB; (9) fumarate reductase,frdABCD; (10)
lactate dehydrogenase, IdhA.
FLUX THROUGH CITRATE SYNTHASE LIMITS THE GROWTH OF
ETHANOLOGENIC Escherichia coli KO 11 DURING XYLOSE FERMENTATION
Our laboratory has previously engineered E. coli strain B for the production of
ethanol from pentose-rich, hemicellulose syrups by expressing high levels of Zymomonas
mobilis pdc (pyruvate decarboxylase) and adhB (alcohol dehydrogenase) (Ingram et al.
1999; Ohta et al. 1991). This strain was chosen for metabolic engineering because of its
hardiness, wide substrate range, and ability to grow well in mineral salts medium without
organic nutrients (Alterthum et al. 1989; Luli and Strohl 1990). During xylose
fermentation, ATP yield in E. coli is low (-0.67 ATP per xylose) due to separate energy
requirements for uptake and phosphorylation (Tao et al. 2001). Unlike most genetically
engineered strains of E. coli, KO 11 grew to higher densities than the parent in both
mineral salts and complex media (Martinez et al. 1999). Initial studies with Luria broth
demonstrated rapid and efficient conversion of sugars to ethanol by KO 11, with yields
approaching 95% of the theoretical maximum. However, volumetric productivity and
ethanol yields were considerably lower in mineral salts medium without complex
nutrients (Lawford and Rouseau 1996; Martinez et al. 1999; Moniruzzaman and Ingram
1998; York and Ingram 1996a; York and Ingram 1996b).
Supplementing mineral salts medium with complex nutrients significantly
increased ethanol production. The least expensive complex nutrient, corn steep liquor,
supported growth rates and ethanol productivities near those for Luria broth but only
when provided at high concentrations (5% w/v). Although not prohibitively expensive,
the addition of high levels of complex nutrients adds to the cost of ethanol production and
increases the requirements for waste treatment.
The lower rate of ethanol production volumetricc productivity) in minimal media
(compared to Luria broth) resulted from low cell densities and reduced expression of
recombinantpdc and adhB (lower metabolic activity). Inorganic components did not
appear to be limiting and no specific auxotrophic requirements could be identified
(Martinez et al. 1999). Reduced expression of heterologous genes was attributed to
"biosynthetic burden", the competitive reduction in synthesis of heterologous products
due to de-repression of native genes for biosynthetic enzymes (Martinez et al. 1999). In
this study, I have used a mineral salts medium containing 1% CSL and investigated the
basis of the requirement for higher levels of nutrients during xylose fermentation. Four
hypotheses were examined as the basis for the decreased growth in the CSL medium: 1)
availability of macro-nutrients; 2) loss of a biosynthetic pathway due to metabolic
engineering; 3) insufficient ATP during xylose fermentation; and 4) an imbalance in
Materials and Methods
Microorganisms and Media
E. coli B (ATCC 11303) and an ethanologenic derivative, strain KO11 (Ohta et al.
1991), were used in all fermentation experiments. KO11 contains a deletion in thefrd
region (anaerobic fumarate reductase) which eliminates succinate production. Genes
encoding the Zymomonas mobilis ethanol pathway (pdc, adhB) and chloramphenicol
acetyltransferase (cat) were integrated into thepfl gene (chromosome) by a single
cross-over event resulting in a functional, full length pfl gene downstream. Both E. coli B
and K 11 are prototrophic. Stock cultures were stored in glycerol at -750C. Working
cultures were transferred daily on solid medium containing mineral salts and 1% CSL.
Xylose (2%) and chloramphenicol (alternating between 40 and 600 mg liter-') were
included in solid media for K011.
A citrate synthase mutant, E. coli W620 (glnV44, gltA6, galK30, pyrD36,
spdL129, thi-1), was obtained from the E. coli Genetic Stock Center (CGSC # 4278) and
used to test expression of the B. subtilis citZ gene (citrate synthase). This strain contains a
gltA6 mutation (citrate synthase) that prevents growth on M9 medium containing thymine
and glucose (Herbert and Guest 1968).
Corn steep liquor medium (CSL+X) contained (per liter in distilled water): 10 g of
corn steep liquor (-50% solids), 1 g of KH2PO4, 0.5 g of K2HPO4, 3.1 g of (NH4)2SO4, 0.4
g of MgCL2,6H20, and 20 mg of FeCl3*6H20. A one-liter stock solution of CSL was
prepared by dilution of 200 g with distilled water, adjustment to pH 7.2 with 50% NaOH
and steam sterilization. Before use, the sterile stock solution of CSL was aseptically
clarified by centrifugation (10,000 x g, 5 minutes). Mineral solutions were prepared as
described previously (Martinez et al. 1999). Broth cultures and fermentations contained
9% (w/v) xylose medium, unless indicated otherwise. In some experiments, Luria broth
containing xylose was included for comparison.
Seed cultures (100 ml in 250 ml-flask) were grown 14-16 hours at 350C with
agitation (120 rpm). Cells were harvested by centrifugation (5,000 x g, 5 min) and used as
an inoculum to provide an initial concentration of 33 [ig ml-' dry weight (0.1 OD550nm).
Fermentation vessels contained a total volume of 350 ml ( 350C, 100 rpm). Cultures were
maintained at pH 6.5 by the automatic addition of 2N KOH (Moniruzzaman and Ingram
1998). For strain B, 6 N KOH was used to maintain pH after the initial 24 h.
Supplements were added with distilled water as necessary (10 ml total volume).
Organic acids and amino acids were neutralized with NaOH, sterilized by filtration and
added at a final concentration of 2 mg ml-'. Acetaldehyde was added at a final
concentration of 0.25 mg ml-' or 0.5 mg ml-'. Cell mass, ethanol, organic acids and sugars
were monitored at 24 h intervals.
Aerobic Growth Studies
Cells were grown with aeration in 250 ml, baffled flask (350C, 220 rpm)
containing 50 ml of CSL+X medium. A range of sugar concentrations was tested (0.5% -
5%) to determine maximal cell density under conditions of sugar excess. Media were
inoculated directly using cells grown on solid media (18-24 h). Ethanol and cell mass
were measured after 16 h. For comparison, Luria broth containing 5% (w/v) xylose was
Cell mass was estimated as OD550nm using a Baush & Lomb Spectronic 70 (1
OD550 = 0.33 mg ml-' dry cell weight). Ethanol was measured by gas chromatography
using a Varian model 3400 CX as described previously (Moniruzzaman and Ingram
1998). Organic acids and sugars were measured by HPLC using a HP 1090 Series II
chromatograph equipped with a BioRad Aminex HPX-87H ion exclusion column (450C,
4 mM H2SO4, 0.5 ml minf', 10 pl injection) and dual detectors (refractive index monitor
and UV detector at 210 nm).
Fermentation products were also analyzed by NMR to confirm the identities of
HPLC peaks. Broth samples were centrifuged to remove cells. Supernatants (0.9 ml) were
mixed with deuterium oxide (0.1 ml) and sodium 3-(trimethylsilyl)propionate (10 mM
internal standard) in 5 mm sample tubes. Proton spectra were obtained using a modified
Nicolet NT300 spectrometer in the Fourier transform mode (Buszko et al. 1998) as
follows: frequency, 300.065 MHz; excitation pulse width, 5 ps; pulse repetition delay, 3
s; spectral width, 3.6 KHz. A minimum of 100 acquisitions were obtained for each
The citZ gene encoding B. subtilis citrate synthase II has been previously
described (Jin and Sonenshein 1994). This gene was amplified by PCR (forward primer,
5'-TGTGCTCTTCCATGTTTTTACAACACTGTTAAAG-3'; reverse primer,
5'-TTGCTCTTCGTTAGGCTCTTTCTTCAATCG-3') using genomic DNA from B.
subtilis strain YB886 as the template (Barbosa and Ingram 1994). Primers were added to
the Taq PCR Master mix (Qiagen) as recommended by the manufacturer. Conditions of
thermal cycling were as follows: 1) two initial cycles with denaturation at 940C (60 s),
annealing at 500C (60 s) and elongation at 680C (90 s); 2) twenty-eight cycles with
denaturation at 940C (10 s), annealing at 700C (60 s), and elongation at 680C (90 s); and
3) a final elongation step at 720C (10 min). The PCR product (1.5 kbp) was cloned into
pCR2.1-TOPO (Invitrogen) using ampicillin (50 [ig ml1) for selection. Colonies were
screened for size and ability to complement the gltA mutation ofE. coli W620 on
glucose-minimal medium (Herbert and Guest 1968). The citZ gene was also confirmed by
DNA sequencing using a LI-COR model 4000L sequencer (Middendorf et al. 1992).
Whole cell fluorescence was used as a relative measure of reduced nucleotides in
situ (Tartakosvsky et al. 1996; Trivedi and Ju 1994). Since only the reduced form of
NAD(P)H fluoresces at 460 nm, an immediate decrease in the fluorescence of fermenting
cells is interpreted as a decline in the level of NAD(P)H and the NAD(P)H/NAD(P)+
ratio. Cells were grown for 12 h in CSL+X medium, harvested by centrifugation (5,000 x
g, 5 min) and washed 3 times in mineral salts. The pellet was then suspended in mineral
salts solution at a concentration of 1.0 OD550. Emission at 460 nm (excitation at 340 nm)
was recorded at 5 s intervals using an Aminco-Bowman Series 2 Luminescence
Spectrometer. Cells were energized by the addition of 1% xylose resulting in an
immediate increase in fluorescence, primarily due to the increase in NADH/NAD+ ratio.
Test compounds were added at a final concentration of 2 mg ml-' (organic acids, amino
acids) or 0.25 mg ml-' acetaldehydee) using distilled water as a control. Results for each
test compound were expressed relative to the xylose-dependent increase in fluorescence.
Control experiments confirmed that quenching of cellular fluorescence did not occur
when additives were mixed with energy-deficient cells (without xylose).
Citrate synthase was assayed using a modification of the method described
previously (Evans et al. 1993, Faloona and Srere 1969). Cultures were grown in one-liter
flasks (250 ml Luria broth) for 16 h at 350C (150 rpm). Cells were harvested by
centrifugation, washed 3 times in buffer containing 50 mM Tris-Cl (pH 8.0) and 20%
glycerol and suspended in 2 volumes of same buffer. Cell-free preparations were made by
two passages through a French Pressure cell (20,000 psi) followed by treatment with
-100 ag ml' deoxyribonuclease I. Cell debris was removed by centrifugation (15,000 x g,
Ih, 40C). The supernatant was dialyzed against 20 mM Tris-Cl and 20% glycerol. Each
assay (1 ml) contained 20 mM Tris-Cl (pH 8.0), 10 mM KC1, 1 mM
5',5'-dithio-bis-(2-nitrobenzoic acid), 10 mM oxaloacetate and 0.5 mM acetyl-CoA.
Reactions were initiated by the addition of cell lysate and monitored for 300 s at 412 nm.
Specific activity was reported as pmol of reduced coenzyme A produced per min per mg
Previous studies have shown that up to 5% CSL is needed to support anaerobic
growth and ethanol production at a rate near that of Luria broth (Martinez et al. 1999).
Based on a comparison of E. coli elemental composition (Taylor 1946), the
macro-nutrient salts in CSL+X medium should provide sufficient nitrogen and
phosphorus to support the growth of at least 5 mg ml-' dry weight. During anaerobic
growth in pH-stats with CSL+X (Fig. 2-1), the maximal cell density for KO11 was only
about 1 mg ml-', 33% lower than the parent E. coli B (1.5 mg ml-') in CSL+X medium
and only 25% of that reached by KO 11 (4 mg ml-') in Luria broth plus sugar (Martinez et
al. 1999, York and Ingram 1996b, York and Ingram 1996b). During aerobic growth with
the same nutrients, however, KO 11 grew to a maximum density of 2.7 mg ml-'. This is
almost two-fold higher than E. coli B under the same conditions and 2.7-fold higher than
KO 11 during anaerobic growth. Together, these results indicate that the anaerobic growth
of KO 11 in 1% CSL with 9% xylose is not limited by the availability of macro-nutrients
(ie. N, P, etc.) or by the inactivation of a biosynthetic pathway due to genetic
manipulation. However, metabolic engineering of the ethanol pathway does appear to
contribute to the reduced growth of KO 11 in this medium under anaerobic conditions.
The separate energy requirement for uptake (ATP-dependent transporter or proton
symport) and phosphorylation (xylulokinase) results in a low net yield of ATP from
xylose fermentation (0.67 ATP per xylose), 33% of the yield from glucose (2 ATP per
glucose) (Tao et al. 2001). An additional ATP can also be produced from pyruvate by the
acetate pathway. Since KOl1 produced less acetate than strain B (Table 2-1), the growth
ofKO 11 could be limited by the availability of ATP. To test this hypothesis, I compared
the growth of KO11 in 1% CSL containing 9% xylose with growth in 1% CSL containing
9% glucose (Fig. 2-2A and Fig. 2-2B). Glucose was fermented to ethanol at a higher rate
than xylose. However, the cell yield of KO 11 was identical for both sugars, despite the
3-fold difference in net ATP production. Although cell densities were low, cells remained
metabolically active for at least 96 h and produced most of the ethanol after growth had
Metabolic Imbalance Relieved by Addition of Pyruvate or Acetaldehyde.
Pyruvate serves a dual role during fermentation, as a source of carbon skeletons
for biosynthesis and as a source of electron acceptors acetaldehydee) to allow continued
ATP production by glycolysis. During sugar fermentation to ethanol, one NADH is
produced per pyruvate. Each NADH must be oxidized by reducing an electron acceptor
such as acetaldehyde or by biosynthetic reactions (Mat-Jan et al. 1989). In a growing wild
type E. coli, partitioning of pyruvate between biosynthesis and redox needs is presumed
to be balanced for optimal growth. Metabolic engineering of the ethanol pathway
contributed to the reduced growth of KO 11 only under fermentative conditions,
consistent with a metabolic imbalance resulting from uncontrolled utilization of pyruvate
for ethanol production. This possibility was confirmed by the addition of pyruvate to
CSL+X medium. Pyruvate addition resulted in a dose-dependent increase in cell growth
and ethanol production that was particularly evident after 24 h (Fig. 2-3A and Fig. 2-3B).
With 2 mg ml-' of added pymvate, growth and ethanol production were twice that of the
control without pyruvate addition (Table 2-2). Supplementing with pyruvate did not cause
a buildup of TCA intermediates or acidic fermentation products (Table 2-1). Note that
format was produced in all fermentations, confirming the that the pfl gene encoding
pyruvate formate-lyase remains functional in KO 11.
The addition ofpyruvate to media has been shown to increase the intracellular
pyruvate pool in E. coli (Yang et al. 2001), increasing the ratio of potential electron
acceptors for the oxidation of NADH (from glycolysis). When added at a level of 2 mg
ml-', pyruvate was metabolized concurrently with cell growth during the first 24 h after
inoculation (Table 2-1). The pyruvate-dependent increase in cell mass (-1 mg ml-') was
roughly equivalent to half of the added pyruvate (Table 2-2, Fig. 2-3A). Remaining
pyruvate is presumed to be metabolized to acetaldehyde by recombinant Z. mobilis
pyruvate decarboxylase. Since acetaldehyde has been previously shown to stimulate
growth and ethanol production by yeasts (Walker-Caprioglio et al. 1985) and Z. mobilis
(Stanley et al. 1997), it seemed possible that the stimulation of cell growth by pyruvate
could be mediated in part by an increase in acetaldehyde from pyruvate (Table 2-2, Fig.
2-3C and Fig. 2-3D). Concentrations of acetaldehyde above 0.50 mg ml' were toxic.
With lower concentrations of acetaldehyde (0.25 and 0.50 mg ml-'), cell growth and
ethanol production were increased. Like pyruvate, added acetaldehyde was fully
metabolized during the initial 24 h after inoculation (Table 2-1). A near optimal level of
acetaldehyde was provided by 2 additions of 0.25 mg ml' each to CSL+X medium
(initially and after 12 h). This was almost as effective as pyruvate (2 mg ml-') in
stimulating ethanol production and also caused a 65% increase in cell mass. The basis for
the increase in cell growth is not readily explained by the limited routes for acetaldehyde
metabolism in E. coli as compared to those for pyruvate, a key central metabolite. These
results provide evidence that the beneficial effect of added pyruvate results primarily
from an increase in electron acceptors.
Pyruvate as a Source of Carbon Skeletons for Biosynthesis.
The pyruvate-stimulated increase in cell growth reflects a two-fold increase in the
flow of carbon into biosynthesis. Pyruvate and upstream metabolites in glycolysis are
used for the biosynthesis of approximately half of cellular constituents. Pools for these
upstream intermediates may increase when pyruvate is added, increasing availability for
biosynthesis. Pyruvate (and phosphoenolpyruvate) is also converted to a series of
biosynthetic intermediates by the TCA pathway and linking reactions. The TCA pathway
provides half of the carbon skeletons for cell protein. None of the TCA intermediates can
be produced readily from acetaldehyde by biosynthetic reactions. Note that the TCA
pathway is not cyclic during fermentation. This pathway is interrupted between
2-ketoglutarate and succinate by ArcAB-mediated repression of genes (sucAB) encoding
2-ketoglutarate dehydrogenase (Iuchi and Lin 1988). One side of the TCA pathway
produces 2-ketoglutarate, the precursor for the glutamic acid family of amino acids,
polyamines, among others. Precursors such as oxaloacetate on the other side of the TCA
pathway are derived from phosphoenolpyruvate. Oxaloacetate is used for synthesis of the
aspartic acid family of amino acids, etc. The addition of pyruvate could potentially
increase the flow of carbon into both sides.
TCA pathway intermediates were tested as additives to CSL+X medium.
Utilization of these additives was investigated using HPLC and NMR (Table 2-2, Fig.
2-4). All except two, succinate (100% remaining) and isocitrate (78% remaining), were
metabolized efficiently during the initial 24 h of fermentation (Table 2-1). Additions of
malate and fumarate resulted in a similar small increase in fumarate, but did not stimulate
growth or ethanol production. Despite the potential interconversion of these
intermediates, fumarate did not accumulate when oxaloacetate was added. Addition of
aspartate, the transamination product of oxaloacetate, was similarly ineffective. Indeed,
addition of oxaloacetate, malate, fumarate and aspartic acid reduced growth and ethanol
production. In contrast, 2-ketoglutarate was almost as effective as pyruvate in stimulating
growth and ethanol production by KO 11. A similar stimulation was also observed for
glutamate, the transamination product of 2-ketoglutarate.
TCA intermediates that are immediate precursors of 2-ketoglutarate were not
beneficial. Isocitrate was not readily metabolized. Citrate was metabolized but had no
effect on growth and ethanol production. Growth with added citrate was accompanied by
an accumulation of fumarate and a high acetate/formate ratio similar to that with pyruvate
(Table 2-1). Note that this ratio is near unity for other fermentations with TCA
intermediates, providing a clue to the ineffectiveness of citrate. The addition of citrate
may induce citrate lyase (Furlong 1987; Lutgens and Gottschalk 1980; Schneider et al.
2000), an enzyme that cleaves citrate into an equimolar mixture of oxaloacetate and
acetate. Oxaloacetate is readily metabolized to furmarate. Both fumarate and acetate were
higher in fermentations with added citrate than with 2-ketoglutarate and other TCA
intermediates, consistent with the induction of citrate lyase. Induction of this enzyme is
presumed to block the beneficial effects of this TCA intermediate for biosynthesis.
When considered together, studies with added TCA intermediates provide
evidence that the beneficial effect ofpyruvate for growth and ethanol production by
KO 11 in CSL+X medium results in large part from an increase in the flow of carbon
skeletons into 2-ketoglutarate and subsequent products of biosynthesis. However,
investigations with added pyruvate and acetaldehyde provided evidence that an increase
in electron acceptors was arguably of primary importance for the beneficial effect of
pyruvate. For both to be possible, both must be mediated by a common mechanism.
The ratio of NAD(P)H/NAD(P)+ has been shown to alter cellular patterns of
metabolic flux (de Graef et al. 1999). NAD(P)H is an allosteric inhibitor of many
enzymes including pyruvate dehydrogenase (Graham et al. 1989), phosphotransacetylase
(Suzuki 1969), malate dehydrogenase (Sanwal 1969) and citrate synthase (Faloona and
Srere 1969; Weitzman 1966). In KO11, the addition of acetaldehyde or pyruvate
(metabolized to acetaldehyde by recombinant pyruvate decarboxylase) would be expected
to decrease the level of NAD(P)H and the NAD(P)H/NAD(P)+ ratio by increasing the
pool of acetaldehyde available for reduction to ethanol. This has been investigated in
non-growing cells by examining the effects of these additives on whole-cell fluorescence.
(Fig. 2-5A and Fig. 2-5B).
Fluorescence changes in responses to additives were immediate and stable as
shown for acetaldehyde (Fig. 2-5A). Relative fluorescence increased when fermentation
was initiated by the addition of xylose, and decreased immediately upon the addition of
acetaldehyde (alcohol dehydrogenase) and pyruvate (pyruvate decarboxylase plus alcohol
dehydrogenase), consistent with expected changes in the oxidation of NADH. The
fluorescence of energized cells also decreased immediately upon the addition of
2-ketoglutarate and oxaloacetate (Fig. 2-5B). The apparent decline in NAD(P)H in
response to these two TCA pathway intermediates may be due to reductive amination
oxaloacetatee and 2-ketoglutarate). Addition of the respective amino acid products,
glutamic acid and aspartic acid, did not cause a similar change. With added oxaloacetate,
malate dehydrogenase provides an additional opportunity for NADH oxidation.
Additions of malate, fumarate, succinate, citrate and isocitrate did not
significantly alter whole cell fluorescence. Together, these data demonstrate that three
compounds which increased the growth and fermentation of KO 11 in CSL+X medium
acetaldehydee, pyruvate and 2-ketoglutarate) also decreased the NAD(P)H/NAD(P)+ ratio
in cells. Compounds which did not decrease this ratio were not beneficial. Oxaloacetate
was an exception. Although this compound decreased the NAD(P)H/NAD(P)+, growth
and fermentation were retarded. The negative effects of added oxaloacetate may be
attributed to the induction of pyruvate carboxykinase. Together with
phosphoenolpyruvate carboxylase, this enzyme creates a futile cycle for ATP (Chao et al.
1994; Chotani et al. 2000 ). ATP yields are low for xylose and ATP wasted by this futile
cycle may offset any potential benefits from increased oxidation of NADH.
Citrate Synthase, a Link Between NADH and 2-Ketoglutarate.
In E. coli (gltA) as in most Gram-negative bacteria, citrate synthase is
allosterically inhibited by NADH and activated by acetyl-CoA (Weitzman 1981). The
activity of this enzyme serves to regulate the flow of carbon into the 2-ketoglurate side of
the TCA pathway, linking the cellular abundance of NADH and acetyl-CoA to the
production of 2-ketoglutarate for biosynthesis (Faloona and Srere 1969; Lee et al. 1994;
Walsh and Koshland, Jr. 1985). This enzyme integrates both beneficial effects of added
pyruvate, increased electron acceptors acetaldehydee) and increased carbon skeletons in
the 2-ketoglutarate arm of the TCA pathway (2-ketoglutarate). The allosteric control of
this enzyme by NADH could restrict the flow of carbon into the biosynthesis of
2-ketoglutarate and other products. This hypothesis can be readily tested by expressing an
NADH-insensitive recombinant citrate synthase gene in K 11.
The primary citrate synthase in Gram-positive bacteria is allosterically regulated
by ATP and relatively insensitive to NADH (Jin and Sonenshein 1996). Since an
over-abundance of ATP is not anticipated during xylose fermentation (Tao et al. 2001),
expression ofB. subtilis citZ in KO11 would be expected to increase carbon flow into the
oxidizing arm of the TCA pathway. Primers were used to clone the citZ gene (including
ribosomal binding site) into pCR2.1-TOPO to produce pLOI2514. Plasmid pLOI2514
was found to complement a gltA mutation in E. coli W620 on plates containing M9
minimal media supplemented with glucose and thymine. Citrate synthase activity (0.08 U
mg protein-') was also confirmed in strain W620(pLOI2514) and absent in strain W620
Expression of citZ in KO 11 (pLOI2514) increased growth and ethanol production
by approximately 75% (Fig. 2-6) in comparison to the control with vector alone,
KO 1 (pCR2.1-TOPO). The low level of NADH-insensitive citrate synthase produced
from pLOI2514 was almost as effective as pyruvate, acetaldehyde and 2-ketoglutarate
additions in stimulating growth. Thus the allosteric regulation of the native citrate
synthase by high NADH appears to limit the flow of carbon skeletons into biosynthesis in
The rate of ethanol production and ethanol yield are important factors in
determining the cost of large-scale fermentation processes. For KO 11, both of these are
directly related to the extent of growth of the biocatalyst (Fig. 2-7). In CSL+X medium,
more than half of the ethanol was produced after cells entered stationary phase (Fig. 2-1).
By increasing cell densities, fermentation times can be reduced without sacrificing
ethanol yield. However, previous studies with KO 11 have shown that high levels of
complex nutrients were needed for cell growth and rapid ethanol production (Martinez et
al. 1999; York and Ingram 1996b; York and Ingram 1996b). This apparent requirement
for high levels of complex nutrients now appears to reflect a regulatory error in the
partitioning of pyruvate skeletons between competing requirements for the oxidation of
NADH and biosynthesis. Our study demonstrates that the growth of KO 11 was not
limited by nutrients, a lack of biosynthetic enzymes, or insufficient ATP from xylose
metabolism (0.67 ATP per xylose). During the fermentation of 9% xylose, growth was
limited by a lack of carbon skeletons for the biosynthesis of products derived from
2-ketoglutarate. Growth and ethanol production were increased by the addition of
pyruvate or 2-ketoglutarate, but not by the addition of oxaloacetate, malate or fumarate.
The apparent starvation for carbon skeletons to produce 2-ketoglutarate was also
alleviated by the addition of acetaldehyde, consistent with an involvement of NADH or
NADH/NAD+ ratios. The ratio ofNADH/NAD+ is typically higher during fermentation
than during oxidative metabolism (de Graef et al. 1999). High levels of NADH serve as
an allosteric inhibitor of citrate synthase, the first committed step for the production of
2-ketoglutarate and a likely bottleneck for the biosynthesis of many amino acids (Walsh
and Koshland, Jr. 1985). Addition of acetaldehyde decreased the NADH/NAD+ ratio by
increasing the pool of electron acceptors, potentially increasing the function of the native
citrate synthase in vivo. This hypothesis was confirmed, in part, using the B. subtilis citZ
gene encoding an NADH-insensitive citrate synthase (Jin and Sonenshein 1994; Jin and
Sonenshein 1996). Expression of citZ in KO 11 stimulated growth and ethanol production
by almost two-fold, substantially reducing the need to supply high levels of complex
The pattern of carbon flow in KO 11 is summarized in Figure 2-8. Expression of
high levels of Z. mobilis pdc and adhB redirect pyruvate away from native fermentation
pathways (pyruvate formate-lyase, lactate dehydrogenase) and into ethanol, even in the
presence of competing native enzymes (Ohta et al. 1991). Since the Km of pyruvate
decarboxylase for pyruvate is approximately one-tenth that of the competing enzyme,
pyruvate formate-lyase, production of acetyl-CoA would also be limited. Integration of
the ethanol-production genes into the chromosomalpfl gene may further contribute to this
problem by reducing the level of pyruvate formate-lyase activity. Although pyruvate
dehydrogenase has a Km for pyruvate that is equal to that of pyruvate decarboxylase,
pymvate dehydrogenase is expressed at low levels during fermentation and is
allosterically inhibited by the high levels of NADH present during fermentation (de Graef
et al. 1999; Graham et al. 1989). In CSL+X medium, a portion of cellular pyruvate was
converted to acetyl-CoA by KO 11 during the first 24 h as evidenced by the accumulation
of acetate as a fermentation product. These acetate levels are presumed to be in excess of
The addition of either pyruvate or acetaldehyde dramatically stimulated the
growth of KO 11 in CSL+X medium. At least three sites of allosteric regulation may
contribute to the increase in growth. Both pyruvate dehydrogenase (de Graef et al. 1999;
Graham et al. 1989) and citrate synthase (Faloona and Srere 1969; Weitzman 1981) are
allosterically inhibited by NADH. Oxidation of NADH from glycolysis during the
reduction of acetaldehyde from added pyruvate or added acetaldehyde would tend to
decrease the NADH/NAD+ ratio. This should reduce the allosteric inhibition ofpyruvate
dehydrogenase and citrate synthase by NADH. Additional acetyl-CoA from pyruvate
dehydrogenase would supplement that produced by pyruvate formate-lyase and increase
the pool of acetyl-CoA a substrate for citrate synthase and an allosteric antagonist of
NADH inhibition of the native citrate synthase (Weitzman 1981). Together, these
regulatory circuits would feed-forward to promote the flow of additional carbon skeletons
into 2-ketoglutarate and subsequent products of biosynthesis.
Table 2-1. Effects of additives on the composition of fermentation products (24 h) in 1% CSL+X medium (9% xylose).
% of additive Fermentation Products (mM)
Medium or strain Additive
remaining Fumarate Succinate Lactatea Formate Acetate Ethanol
CSL+X medium <0.01 <0.3 9.6 <1.5 <1.0 <1.0
E. coli B, parent None <0.01 53.4 27.7 95.8 91.4 71
Sodium pyruvate (2 mg ml'1)
Acetaldehyde (2 x 0.25 mg ml'1)
Citric acid (2 mg ml ')a
Isocitric acid (2 mg ml'1)
Sodium 2-ketoglutarate (2 mg ml'1)
Oxaloacetic acid (2 mg ml'1)
Sodium malate (2 mg ml'1)
Sodium fumarate (2 mg ml'1)
Sodium succinate (2 mg ml'1)
E. coli KO 11
Table 2-2. Effects of additives on growth and ethanol production by KO 11 in 1% CSL+X medium (9% xylose).
SConcentration Cell Yieldc o ton Maximum Ethanol Yieb
Additive No. of Ethanol Yieldb (%)
mg ml1- mM Average % of Control g liter' h-1 % of Control mg ml1- % of Control
None (control) 26 -- 0.94 0.15 100 0.38 0.08 100 31.12 4.64 100 61
Sodium pyruvate 3 0.5 4.6 1.32 0.32 140 0.55 0.17 144 37.60 5.91 121 74
Sodium pyruvate 3 1.0 9.1 1.49 0.30 158 0.66 0.17 173 40.09 5.53 129 79
Sodium pyruvate 15 2.0 18.2 1.99 0.20 212 0.81 0.14 213 44.22 2.69 142 87
Sodium pyruvate 3 4.4 36.4 2.08 0.08 221 0.83 0.01 218 43.23 1.50 139 85
Acetaldehyde (half initially + 4 0.5 (total) 11.4 1.55 0.07 165 0.60 0.35 158 44.05 1.89 142 86
half after 12 h)
citric acid 2 2.0 10.4 0.88, 0.75 86 0.35, 0.33 89 30.00, 27.59 93 56
isocitric acid 2 2.0 6.6 1.11, 0.88 105 0.51, 0.36 113 36.45, 29.59 106 65
sodium 2-ketoglutarate 5 2.0 11.9 1.89 0.18 201 0.84 0.08 221 41.54 1.24 133 81
oxaloacetic acid 2 2.0 15.1 0.75, 0.75 80 0.28, 0.28 74 23.58, 23.64 76 46
sodium malate 2 2.0 11.2 0.76, 0.84 85 0.27, 0.27 71 23.89, 24.97 79 48
sodium fumarate 2 2.0 12.5 0.80, 0.96 91 0.37, 0.32 92 27.15, 29.84 92 56
sodium succinate 3 2.0 7.4 1.00 0.12 106 0.41 0.05 108 35.00 1.31 112 69
potassium glutamate 3 2.0 10.8 1.66 0.17 177 0.66 0.13 174 43.14 0.13 139 85
aspartic acid 2 2.0 15.0 0.85, 0.82 88 0.33, 0.32 84 28.40, 27.40 90 55
a Volumetric Productivity was calculated as the average hourly rate of ethanol production between 24 h and 48 h after inoculation.
When less than 3 replicates are presented, the values of each replicate are shown.
b Yield is expressed as a percentage of the theoretical yield (100% = 0.51 g ethanol per g xylose).
5 25 I Anaerobic
: ^ H5: 5Aerob c I
Figure 2-1. Comparison of maximal cell densities achieved during aerobic and anaerobic
growth in 1% CSL mineral salts medium containing either xylose or glucose. Thin
lines representing the standard error of the mean are shown for averages with
three or more replicates.
0.5- --9% Xylo se
S-A- 9% Glucose
0 24 48 72 96
-m- 9% Xyl ose
-- 9% Glucose
0 24 48 72 96
Figure 2-2. Comparison of growth and ethanol production from glucose and xylose by E.
coli KO11 during the fermentation of 9% sugar in 1% CSL mineral salts medium.
A. Growth. B. Ethanol. Thin lines representing the standard error of the mean are
shown for averages with three or more replicates.
0 24 48
S -u-No addition
-&-0.25 g Acetaldehyde
---0.50 g Acetaldehyde
--o-2x 0.25 g Acetaldelh
0 24 48 72
S0.25 g Acetaldehyde
--0.50 g Acetaldehyde
S--2x 0.25 g Acetaldelyc
0 24 48 72
Figure 2-3. Effects of added pyruvate and acetaldehyde on growth and ethanol production
by E. coli KO11 in CSL+X medium. A. Cell growth with added pyruvate. B.
Ethanol production with added pyruvate. C. Cell growth with added acetaldehyde.
D. Ethanol production with added acetaldehyde. Thin lines represent the standard
error of the mean.
Figure 2-4. Initial effects of added TCA pathway intermediates on growth and ethanol
production by E. coli KO11 (24 h). A. Growth. B. Ethanol. Thin lines represent
the standard error of the mean.
A 1% Xylose 0.25 g/L Acetaldehyde
0 60 120 180 240 300
O 100 B
Figure 2-5. Effect of metabolites on whole-cell fluorescence. A. Effects of acetaldehyde
on the xylose-dependent increase in fluorescence (time course). B. Effects of
metabolites on the xylose-dependent increase in fluorescence. Values in B are
expressed as a percentage of the xylose-dependent increase in the fluorescence of
whole cells observed in the presence of both xylose and the indicated additive.
Note that a decrease in the xylose-dependent fluorescence is interpreted as a
decrease in NAD(P)H and the NAD(P)H/NAD(P)+ ratio.
-- K011 (pCR2.1-TOPO)
0.0 -T-KOll (pLOI2514)
II' I I
0 24 48 72 96
-eKO 11 (pCR2.1-TOPO)
0A --KO 11 (pLOI2514)
0 24 48 72 96
Figure 2-6. B. subtilis citZ increases the growth and ethanol production of KO 1l in
CSL+X medium. A. Growth. B. Ethanol. Thin lines represent the standard error of
m Ethanol Yield
A Volumetric Productivity
0.0 0.5 1.0 1.5 2.0
Cell Mass (g/L)
Figure 2-7. Relationship between cell yield and fermentation performance. In this plot,
results were combined from fermentations with CSL+X medium alone and with
supplements. A computer-generated polynomial was used to approximate cell
yields. Results from a linear regression analysis are shown for volumetric
productivity. Dotted lines represent the the 95% confidence intervals.
glyceraldehyde-3-P -------------------------- XYLOSE 52
;NADH t; acetyl-CoAl t
NAD 17 (deleted)
C02 -'r1 NADPH
Figure 2-8. Fermentation and TCA pathway. Unless noted otherwise, enzymes listed are
native to E. coli. Key to enzymes: 1. pyruvate kinase (pykA, pykF); 2. pyruvate
formate-lyase (pflB); 3. pyruvate dehydrogenase (aceEF,lpd); 4.
phosphotransacetylase (pta); 5. acetate kinase (ackA); 6. alcohol/aldehyde
dehydrogenase (adhE); 7. Z. mobilis pyruvate decarboxylase (pdc); 8. Z. mobilis
alcohol dehydrogenase II (adhB); 9. lactate dehydrogenase (IdhA); 10.
phosphoenolpyruvate carboxylase (ppc); 11. citrate synthase (gltA); 12. aconitase
(acn); 13. isocitrate dehydrogenase (icd); 14. glutamate dehydrogenase (gdhA);
15. malate dehydrogenase (mdh); 16. fumarase (fumB); 17. fumarate reductase
(frdABC); 18. aspartate transaminase (aspA); 19. aspartase (aspC). Arrows
beneath citrate synthase indicate inhibition of activity by NADH and antagonism
of NADH inhibition by acetyl-CoA.
GENETIC CHANGES TO OPTIMIZE CARBON PARTITIONING IN
ETHANOLOGENIC Escherichia coli KO 11
Citrate synthase, a key enzyme in the partitioning of carbon into biosynthesis
(Walsh and Koshland, Jr. 1985), was shown to be growth limiting for KO11 (Chapter 2).
Native citrate synthase is allosterically inhibited by high levels of NADH typical of
fermentation (Weitzman 1981). Growth and ethanol production were substantially
improved in KO 11 by expression of an NADH-insensitive citrate synthase (citZ) from
Bacillus subtilis. A similar stimulation of growth and ethanol production was observed
during low aeration (oxidation of NADH) and with the addition ofpyruvate,
2-ketoglutarate, and acetaldehyde.
An alternative approach to enhance citrate synthase activity in KO 11 is to increase
available substrate pools oxaloacetatee and acetyl-CoA). In vitro, acetyl-CoA has been
shown to serve as an allosteric activator ofphosphoenolpyruvate carboxylase (Izui et al.
1981) for the production of oxaloacetic acid and to relieve the allosteric inhibition of
citrate synthase by NADH (Weitzman 1981). In this chapter, I demonstrate that
physiological and genetic approaches which increase the availability of acetyl-CoA for
biosynthesis stimulate cell growth and ethanol production. These results were used to
engineer a second generation biocatalyst, strain SU102, in which a small additional
portion of substrate carbon was redirected from fermentation products to cellular
Materials and Methods
Microorganisms and Media
Strains and plasmids used in this study are listed in Table 3-1. KOl and its
derivatives (SU102 and SU104) are prototrophic. Working cultures of ethanologenic
strains were transferred daily on solid medium (1.5% agar) containing mineral salts, 2%
xylose, and 1% CSL (Chapter 2). Stock cultures were stored frozen at -750C. Luria-agar
plates were used for the maintenance of other strains. Ampicillin (50 [ig/ml), kanamycin
(50 lig/ml) and tetracycline (5 or 10 [ig/ml) were added as appropriate.
Seed cultures and fermentations (350C and 150 rpm) were grown in mineral salts
medium containing 1% CSL and 9% xylose (CSL+X medium; Chapter 2). Fermentations
were maintained at pH 6.5 by automatic addition of 2N KOH (Moniruzzaman and Ingram
1998). Supplements were filter sterilized as concentrates and added directly to
fermentation broth. Samples were removed during fermentation for the measurement of
cell mass, ethanol, organic acids and sugars.
Cell mass was estimated from the optical density at 550 nm using a Bausch &
Lomb Spectronic 70 spectrophotometer (1 OD50 = 0.33 mg ml-' dry cell weight). Ethanol
and acetaldehyde were measured by gas chromatography (Varian 3400CX)
(Moniruzzaman and Ingram 1998). Organic acids and sugars were analyzed by HPLC
(Hewlett Packard 1090 series II chromatograph equipped with refractive index and UV210
detectors) with a BioRad Aminex HPX-87H ion exclusion column. Maximum volumetric
productivity in mmol liter' h-' was estimated as the first derivative of ethanol production
using PSI-Plot software (Poly Software International, Salt Lake City, Utah). Specific
productivity was estimated by dividing volumetric productivity by cell mass; units are
mmol (gram cell dry weight)-' hour'.
Standard methods were used for plasmid construction, DNA amplification (PCR),
transformation, electroporation and P1 phage transduction (Miller 1992; Sambrook and
Russell 2001). Primers (ORFmers) for the amplification of the E. coli ackA and adhE
coding regions were purchased from the Sigma Genosys (The Woodlands, TX). These
primers included SapI sites at both ends of the amplified product. Chromosomal DNA
from E. coli W3110 (ATCC 27325) served as the template for amplification. This strain
was also used as an intermediate during the construction of adhE deletion in KO 11.
Chromosomal insertion of deleted genes (adhE and ackA) was facilitated by
inserting a tet gene flanked by FRT sites for removal of the antibiotic marker by the
chlorotetracycline-inducible FLP recombinase (pFT-A) in the final construct
(Martinez-Morales et al. 1999; Posfai et al. 1997). Integration of linearized DNA was
facilitated by using pKD46 (temperature conditional) containing an arabinose-inducible
red recombinase (Datsenko and Wanner 2000). Putative deletion mutants were selected
for tetracycline resistance (5 mg liter-') and screened for appropriate antibiotic resistance
markers. At each step, mutants were verified by analyses of PCR and fermentation
Construction of pLOI2065 Containing a Removable Tetracycline Resistance Cassette
To facilitate antibiotic removal after chromosomal integration, a reusable cassette
was constructed from the tet gene of pKNOCK-Tc (Alexeyev 1999) and the FRT sites in
pSG76-A and pSG76-K (Posfai et al. 1997). Both FRT sites were oriented in the same
direction to allow efficient in vivo excision of the tet gene by theflp-encoded
recombinase (Martinez-Morales et al. 1999). This cassette was inserted into a modified
pUC18 to produce pLOI2065 (Fig. 3-2). Plasmid pLOI2065 contains two EcoRl sites and
two Smal sites oriented to allow the isolation of the FRT-tet-FRT cassette as a Smal to
EcoRl fragment for directional insertion, as a blunt fragment (Smal) and as a sticky-ended
Nucleotide Sequence Accession Number
The sequence for plasmid pLOI2065 has been deposited in GenBank under
acquisition number AF521666.
Construction of SU102 Containing an Insertion Mutation in ackA
Strain SU102 was made by introducing the ackA mutation directly into KO11.
The PCR-amplified coding region of ackA was cloned into pCR2.1-TOPO. After
digestion with EcoRl, the 1.2 kbp fragment containing the ackA coding region was ligated
into the unique EcoRl site of pLOI2302. A recombinant plasmid was selected in which
the direction of transcription of lac and ackA genes were opposite. The ackA gene was
disrupted by digestion with EcoRV (1 site) and the insertion of a 1.7 kbp Smal fragment
from pLOI2065 containing a tet gene flanked by two FRT sites for FLP recombinase. A
2.8 kbp AscI fragment containing ackA'-FRT-tet-FRT- 'ackA was isolated from this
plasmid and ligated into the AscI site ofpLOI2224 containing a conditional R6K replicon.
The resulting plasmid, pLOI2375 (Fig. 3-2), was used as a template for PCR
amplification of the 2.8 kbp AscI fragment with ackA primers. After purification by
phenol extraction, amplified DNA was used for electroporation ofE. coli KO 1 (pKD46)
expressing phage lambda red recombinase (Datsenko and Wanner 2000). Recombinants
were selected for tetracycline resistance. Plasmid pKD46 was eliminated by growth at
400C. The integrated tet gene was deleted using pFT-A expressing theflp recombinase
(Martinez-Morales et al. 1999; Posfai et al. 1997). After removal of this plasmid by
growth at 400C, the resulting strain containing a mutation in ackA (insertion of 98 bases
including stop codons in all three reading frames) was designated SU102.
Construction of SU104 Containing a Deletion in adhE
A mutation in adhE was initially constructed in W3110 prior to P1 transduction
into KO 11. The PCR-amplified coding region of adhE (2.7 kbp) was cloned into
pCR2.1-TOPO. A recombinant plasmid was selected in which the transcription of lac and
adhE were oriented in the same direction. The central region of the adhE gene (1.1 kbp)
was deleted by digestion with HinCII (2 sites) and replaced with a 1.6 kbp Smal fragment
from pLOI2065 containing the FRT-tet-FRT cassette (1.7 kbp) to produce pLOI2803 (Fig.
3-2). After digestion of pLOI2803 with both PvuI and Scal, this plasmid served as a
template to amplify the 3.2 kbp region containing adhE'-FRT-tet-FRT- 'adhE using adhE
primers. This amplified DNA was used for electroporation. Recombinants were selected
for tetracycline resistance. Plasmid pKD46 was eliminated by growth at 420C.
P1 transduction was used to transfer the adhE mutation in W3110 to KO 11. To
circumvent differences in restriction systems, the adhE::tet mutation was transduced into
a restriction-negative (modification-positive) derivative ofE. coli B (strain WA837) prior
to transduction into K 11. The tetracycline resistance gene was deleted from the K 11
derivative using pFT-A expressing theflp recombinase (Martinez-Morales et al. 1999;
Posfai et al. 1997). After removal of this plasmid by growth at 400C, the resulting strain
containing an internal deletion in adhE was designated SU104.
Results and Discussion
Acetate Addition Stimulates Growth and Ethanol Production by Reducing Net Acetate
Production During Sugar Metabolism.
During the aerobic metabolism of sugars by E. coli, acetate production has been
associated with a decrease in growth rate. Considerable effort has been made to minimize
acetate production as a means of increasing cell density and the production of
recombinant proteins (Aristidou et al. 1995; Bauer et al. 1990; Chang et al. 1999;
Contiero et al. 2000; Yang et al. 1999a; Yang et al. 1999b). The addition of as little as 2 g
liter-' sodium acetate (24 mM) has been shown to decrease growth rate during oxidative
sugar metabolism (Luli and Strohl 1990). During xylose fermentation by KO 11, however,
the addition of acetate stimulated growth and ethanol production (Fig. 3-3A and B; Table
3-2). A portion of the added acetate was initially consumed, in contrast to control
fermentations where acetate was continuously produced (Fig. 3-4A). Rates of acetate
production declined during subsequent incubation in both control and
acetate-supplemented fermentations. Although almost twice as much sugar was
metabolized by acetate-supplemented fermentations than by control fermentations (no
additions), net acetate production in the acetate-supplemented culture (7.0 mmol liter'1)
was less than half that of the control (18.6 mmoles liter-') after 72 h.
Previous studies have shown that the reversible phosphotransacetylase-acetate
kinase pathway can serve as a route for entry of added acetate into the intracellular pool
of acetyl-CoA (Brown et al. 1977; Higgins and Johnson 1970). Additional acetate uptake
activity may be provided by the inducible acetyl-CoA synthetase, although this gene is
typically repressed under fermentative conditions (Kumari et al. 1995). Thus, the
stimulation of growth and ethanol production by added acetate is presumed to result from
the increased availability of acetyl-CoA. Under anaerobic conditions, the primary role of
the TCA pathway is to supply carbon skeletons for biosynthesis. Increasing the
availability of acetyl-CoA would promote biosynthesis by relieving the NADH-mediated
allosteric inhibition of citrate synthase (Weitzman 1981) and by serving as an allosteric
activator of phosphoenolpyruvate carboxylase (Izui et al. 1981).
Stimulation of Growth and Ethanol Production by Added Pyruvate Can Be Primarily
Attributed to Increased Acetate Production.
The stimulation of growth and ethanol production by pyruvate reported previously
(Chapter 2) appeared quite similar to the effects of added acetate (Fig. 3-3A and B).
Analysis of products during fermentation provided further evidence of a related
mechanism of action for acetate and pyruvate (Fig. 3-4 A-E). With the exception of
format (Fig. 3-4B), profiles of organic acids were similar for acetate and
pyruvate-supplemented cultures. Both were distinctive from the control lacking
supplements. Control fermentations produced lower levels of lactate than
pyruvate-supplemented and acetate-supplemented fermentations during the initial 72 h
(Fig. 3-4C). Addition of pyruvate stimulated the production of acetate to levels equivalent
to that of acetate-supplemented fermentations (Fig. 3-4A). In both pyruvate and
acetate-supplemented fermentations, acetate concentrations were approximately 2-fold
higher than in the control after 36 h. Acetate concentrations in all fermentations remained
relatively constant during further incubation.
Most of the supplemental pyruvate (22 mM) was metabolized during the initial 3
h of incubation (Fig. 3-5) although the benefits for growth and ethanol production
persisted throughout fermentation. During the initial 3 h, the largest change was an
increase in acetate (Fig. 3-6A). Smaller pyruvate-dependent increases were observed for
ethanol, format, lactate and acetaldehyde. Biosynthetic needs were estimated to be small
(increase of approximately 0.06 mg dry cell weight liter') and did not represent a
significant sink for the added pyruvate (2 g liter'). The partitioning of pyruvate between
these different fermentation products (and biosynthesis) in KO 11 is generally regarded as
the result of 5 competing reactions: pyruvate decarboxylase (PDC), pyruvate
formate-lyase (PFL), pyruvate dehydrogenase (PDH), lactate dehydrogenase (LDH) and
phosphoenolpyruvate carboxylase (PPC). On a triose basis, relative activities can be
estimated from the distribution of fermentation products (Fig. 3-1; de Graef et al. 1999).
The large pyruvate-dependent increase in acetate after 3 h (Fig. 3-6A) reflects an increase
in acetyl-CoA production (PFL and PDH activities). In the absence of format hydrogen
lyase induction (Bock and Sawers 1996), format production provides an independent
measure of PFL activity and exhibited a modest increase compared to acetate. These
results indicate that PDH activity (estimated as acetate minus format) serves as the
primary source of additional acetyl-CoA during the metabolism of added pyruvate (Fig.
3-6B). Production of ethanol also increased immediately after the addition of pyruvate
due to an increase in the production of acetaldehyde by PDC. Increased pyruvate
oxidation by PDH is presumed to provide the additional NADH required to reduce
acetaldehyde produced from added pyruvate. The increase in LDH activity (estimated as
lactate production) can be attributed to substrate activation (Tarmy and Kaplan 1968).
Elevated extracellular levels of acetate in pyruvate-supplemented fermentations may
serve to increase intracellular acetyl-CoA pools, extending the period of growth and
thereby increasing the volumetric rate of ethanol production.
The channeling of pyruvate to acetyl-CoA and acetate by the addition of pyruvate
can be readily explained based on known allosteric controls (Fig. 3-1). Pyruvate is both a
substrate for acetyl-CoA production and a strong allosteric activator of
phosphotransacetylase (Suzuki 1969). Addition of pyruvate has also been shown to
increase acetaldehyde and decrease the level of NADH (Chapter 2), an allosteric inhibitor
of phosphotransacetylase (Suzuki 1969) and PDH (de Graef et al. 1999; Hansen and
Henning 1966). These actions would also tend to increase the partitioning of carbon into
Higher levels of succinate and fumarate (3-fold to over 10-fold, respectively) were
produced by acetate- and pyruvate-supplemented fermentations (Fig. 3-4C and D). PPC
(Izui et al. 1981) and citrate synthase (Weitzman 1981) are both activated by acetyl-CoA
and link the supply of this important intermediate to fermentation and biosynthesis. Under
anaerobic conditions, the reductive portion of the TCA pathway is used to produce
succinate. Due to the deletion of fumarate reductase (frd) in KO 11, little succinate was
produced and a small amount of fumarate accumulated. The increases in succinate and
fumarate levels in acetate and pyruvate-supplemented fermentations may result from an
excess of citrate. Excess citrate can be cleaved into acetate and oxaloacetate by an
inducible citrate lyase (Lutgens and Gottschalk 1980). Additional succinate can be
produced from isocitrate by isocitrate lyase (Weitzman 1981).
Pyruvate and free CoA are co-substrates for format production by PFL (Fig.
3-1). Formate levels increased during the initial 12 h of incubation in all fermentations
and declined thereafter (Fig. 3-4B). The decline in format can be attributed to the
formate-inducible format hydrogen lyase (Bock and Sawers 1996). Supplementing with
acetate and pyruvate had opposite effects on format production (Fig. 3-4B), higher
concentrations in pyruvate-supplemented fermentations and lower levels in
acetate-supplemented fermentations in comparison to those of the control. Both
differences are in general agreement with the central role of acetyl-CoA in metabolism
(Chang et al. 1999b; Contiero et al. 2000; Kirkpatrick et al. 2001). In
acetate-supplemented fermentations, format production by PFL may be limited by a lack
of free CoA. Conversely, higher format levels produced by pyruvate-supplemented
fermentations may result from an increase in free CoA due to the allosteric activation of
phosphotransacetylase by pyruvate (Suzuki 1969).
Stimulation of Growth and Ethanol Production by Acetaldehyde Can Be Attributed to
Growth and ethanol production were also stimulated by acetaldehyde (Chapter 2;
Fig 3-3A and B). At concentrations above 5.6 mM, acetaldehyde strongly inhibited
growth. It was empirically determined that stimulation equivalent to that of pyruvate
could be achieved by the addition of 11.2 mM acetaldehyde, 5.6 mM initially and 5.6 mM
after 12 h of fermentation (Chapter 2). Previous studies also demonstrated that the
addition of acetaldehyde caused a rapid decrease in the intracellular concentration of
NADH (Chapter 2).
The initial portion of added of acetaldehyde was metabolized within 3 h (Fig.
3-5A). During this time, ethanol increased by an amount equal to 70% of the added
acetaldehyde (Fig. 3-6A). Increased pyruvate flux through PDH appears to provide the
additional NADH required for acetaldehyde reduction (Fig. 3-6B). The second
acetaldehyde addition was metabolized within 1 h (Fig. 3-5A) although benefits persisted
throughout fermentation (Fig. 3-3A and B). Following the second addition, production of
acetate and ethanol was increased while format production was reduced. The persisting
benefit of acetaldehyde additions for growth and ethanol production are presumed to
result from an increase in the intracellular acetyl-CoA pool as a consequence of higher
extracellular levels of acetate. High levels of NADH and global regulation by ArcA and
FNR (de Graef et al. 1999) may also limit PDH function in the absence of supplements.
Increased production of acetyl-CoA by PDH (and perhaps increased synthesis of PDH)
would be expected in response to NADH oxidation.
The production of format by PFL may be limited by competition with PDH for
free CoA. Patterns of organic acid production in acetaldehyde-supplemented cultures
provide further support for a mechanism of action similar to that for pyruvate and acetate
(Fig. 3A-E). Acetate levels were higher in all three supplemented cultures than in the
unsupplemented control. Each supplemented fermentation also produced higher levels of
succinate, lactate and fumarate than the control.
Stimulation of Growth and Ethanol Production by Inactivation of Non-biosynthetic
Pathways Which Consume Acetyl-CoA.
Acetyl-CoA serves as the single most important intermediate for cellular
biosynthesis, providing over half of the cellular carbon during sugar metabolism
(Neidhardt et al. 1990). Previous studies have shown that cell growth is limited by the
availability of carbon skeletons during the fermentation of xylose (Chapter 2), a limitation
which was relieved (Fig. 3-1A and B) by supplements which increase the extracellular
levels of acetate (acetate, pyruvte, acetaldehyde). During fermentation (Fig. 3-1), two
pathways drain acetyl-CoA from the intracellular pool but provide limited benefit to
biosynthesis. Acetyl-CoA can be reduced to acetaldehyde and ethanol by alcohol
dehydrogenase E (adhE) as an alternative route for NADH oxidation in KO 11 (Fig. 3-1).
Acetyl-CoA can also be converted to acetate by phosphotransacetylase (pta) and acetate
kinase (ackA), increasing the production of ATP. Mutations in these pathways were
investigated as a means of sparing acetyl-CoA for biosynthetic needs.
Inactivation ofackA rather thanpta was chosen to minimize potential problems
associated with global regulation. Acetyl-P is proposed to serve as an important global
regulator in E. coli (Bouche et al. 1998; Kirkpatrick et al. 2001; McCleary et al. 1993),
affecting gene expression and fundamental processes such as the turnover of RpoS.
During oxidative metabolism, inactivation of the acetate pathway (pta, ackA) is
detrimental to growth (Chang et al. 1999b; Contiero et al. 2000; Kirkpatrick et al. 2001).
Although not fully understood, this detrimental effect has been attributed to depletion of
free CoA due to low rates of acetyl-CoA turnover (Chang et al. 1999b). In contrast to that
found in previous studies concerning oxidative metabolism, inactivation of ackA (SU102)
stimulated growth and ethanol production during the fermentation of xylose (Fig. 3-3C
and D). An adhE mutation in strain KO11 (SU104) was of no benefit during xylose
fermentation. Together, these results suggest that ADH contributes little to metabolism in
KO11. The beneficial effect of inactivating ackA is presumed to result from an increase in
the availability of acetyl-CoA for biosynthesis, the genetic equivalent of adding acetate,
pyruvate, or acetaldehyde.
Strains SU104 (adhE mutant) and SU102 (ackA mutant) were also tested in
fermentations with supplements that had been shown to increase the growth and ethanol
production in KO 11 (Table 3-2). Addition of acetate, pyruvate and acetaldehyde to
SU104 increased growth and ethanol production indicating that the native alcohol
dehydrogenase (adhE) was not essential for this response. Growth and ethanol production
by SU102 (ackA) without supplements were equivalent to that of KO11 with
supplements. The addition of pyruvate, acetate, 2-ketoglutarate, or acetaldehyde to SU102
provided little further improvement in growth or ethanol production.
HPLC analysis of organic acids revealed similarities in the patterns of fumarate
(Fig. 3-3J) and succinate (Fig. 3-31) production between SU102 (ackA mutant) and KO11
supplemented with acetate, pyruvate or acetaldehyde (Fig. 3-4E). The ackA mutation in
SU102 also increased lactate production (Fig. 3-41) and delayed the production of format
(Fig. 3-4G) and acetate (Fig. 3-4F). The delay in format production in SU102 could
result from increased acetyl-CoA, reducing the pool of free CoA (co-substrate for PFL)
analogous to acetate-supplemented KO 11 (Fig. 3-4B). Both acetate addition and
mutations in the acetate pathway have been shown to cause a similar repression of 37
genes (Kirkpatrick et al. 2001), attributed to an increase in the acetyl-CoA pool.
Inactivation of acetate kinase (SU102) caused an initial delay in acetate
production but did not block later synthesis. The pathway responsible for acetate
production during the latter stages of fermentation remains unknown but may be the
result of spontaneous dephosphorylation of acetyl-P as previously proposed (Brown et al.
1977) or from induction of cryptic enzyme(s). Despite the potential benefit of increased
ATP production by acetate kinase, the increased drain of acetyl-CoA to acetate through
this pathway appears to be more detrimental for growth and ethanol production by KO 11
than the reduction in ATP. With the exception of acetate (Fig. 3-4F), the production of
fermentation products by the adhE mutant (strain SU104) was essentially the same as for
the parent strain, K 11 (Fig. 3-4A). Acetate production by SU104 continued throughout
fermentation and reached higher final concentrations than K 11.
Increasing the availability of acetyl-CoA stimulated growth and ethanol
production from xylose by prolonging the growth phase of ethanologenic E. coli. The
resulting increase in biocatalyst rather than an increase in cellular activity was responsible
for the increased rate of ethanol production (Table 3-2). Similar benefits were obtained by
minimizing the loss of acetyl-CoA as acetate (ackA mutation) and by increasing
intracellular levels of acetate (supplementing with acetate, pyruvate, or acetaldehyde).
Inactivation of the native E. coli alcohol/aldehyde dehydrogenase (adhE) had little effect
indicating that this pathway has limited function in ethanologenic KO 11.
ATP production during xylose fermentation does not appear to limit growth or
cell yield in KO11. Including the energy required for xylose uptake and activation, less
than 1 ATP (net) is produced from the metabolism of each xylose converted to ethanol
(Tao et al. 2001; Chapter 2). During the initial 12 hours of growth, up to 31% of the ATP
(net) produced by KO11 is provided by the acetate pathway (calculated by assuming 1
ATP per acetate from acetate kinase and 0.4 ATP per pyruvate from glycolysis).
Disruption of this pathway (ackA) in SU102 increased cell yield by 2-fold (Table 3-2).
Thus, the partitioning of carbon skeletons rather than the production of ATP appears to
limit the growth of ethanologenic E. coli during xylose fermentation.
The mechanism for the stimulation of growth in ethanologenic E. coli KO 11 is
consistent with established patterns of allosteric regulation although further controls of
gene expression (FNR, ArcA) may also contribute to the observed effects. More than half
the amino acids produced in the cell are derived from the TCA pathway. Flux through
this pathway is controlled by PPC and citrate synthase (Lee et al. 1994; Walsh and
Koshland, Jr. 1985), activities which can be stimulated by acetyl-CoA (Weitzman 1981).
The individual addition of pyruvate, acetate, and acetaldehyde increased the extracellular
levels of acetate which can in turn serve to elevate intracellular pools of acetyl-CoA by
reversible reactions. Additional benefits of supplements include a reduction in the level of
NADH (added pyruvate and acetaldehyde), an allosteric inhibitor of citrate synthase
which is antagonized by high levels of acetyl-CoA. Based on these results with mutants
and with supplements, we conclude that the regulation of acetyl-CoA production and
consumption can be used to make small changes in the partitioning of carbon between
biosynthesis and fermentation during the ethanol production by E. coli KO11.
Table 3-1. Strains and plasmids used in Chapter 3.
Strain or Plasmid Relevant Characteristics
frd cat pfl' pfl::(Z. mobilis pdc' adhB')
rB m,+ gal met
Reference or source
Ohta etal. 1991
y P exo repAlOl pSC101 repliconts (red recombinase)
blaflp pSC101 repliconts (FLP recombinase)
bla kan ColEl
tet R6K (pir dependent replicon)
kan FRT R6K (pir dependent replicon)
bla FRT R6K (pir dependent replicon)
bla FRT-tet-FRT ColEl
kan R6K (pir dependent replicon)
bla ColEl (EcoRI flanked by AscI sites)
ackA::FRT-tet-FRTkan R6K (pir dependent replicon)
adhE::FRT-tet-FRTkan ColE 1
Datsenko and Wanner 2000
Posfai et al. 1997
Posfai et al. 1997
Posfai et al. 1997
Zhou and Ingram 1999
Table 3-2. Effects of mutations and additives on cell yield and ethanol productivity.
Cell Mass Ethanol
Concentration Maximum Time l a Maximum Max VP b Sp. Prod.c Theoretical
Strain and additive (mM) N (g liter') (h) (h-1) (mM) (mmol liter' h-1) (mmol g-1 h-') Yield d (%)
a Specific growth rate at 2 h.
b VP, maximum volumetric productivity, mM ethanol produced per liter per hour.
' Specific productivity at 12 h, mmol ethanol produced per gram cell dry weight per hour.
d Theoretical yield from 91g liter' xylose (1.667 mmol ethanol/mmol xylose).
e Half added initially, half added after 12 h.
fND, not determined. Estimates of specific and volumetric productivity were not calculated due to the limited number of sampling times.
glyceraede-3-P ------------------ XYLOSE
ID -- NAD'
NAD 1 NADH NADH NAD'
@AcCoA Npywrate acetaBehyde ETHANOL
10, 3(DNADH RDA te 6,AD'
10AcCoA 2 : 6 NADH
CO, acetyl-CoA 4H I acetyl-P 5 acetale
W oaloacetate 0 NADH- cta
NADH Ad aoA n- citraa
SATP ADP+P 12
17 17 + NH4' I R\ NAAfl
RH1 ghltamaite 2-loetogtalate isocitbae
NADP' NADPH CO,
Figure 3-1. Allosteric control of central metabolism. Unless noted otherwise, enzymes
listed are native to E. coli. Enzymes: 1. pyruvate kinase (pykA, pykF); 2. pyruvate
formate-lyase (pflB); 3. pyruvate dehydrogenase (aceEF,lpd); 4.
phosphotransacetylase (pta); 5. acetate kinase (ackA); 6. alcohol/aldehyde
dehydrogenase (adhE); 7. Z. mobilis pyruvate decarboxylase (pdc); 8. Z. mobilis
alcohol dehydrogenase II (adhB); 9. lactate dehydrogenase (IdhA); 10.
phosphoenolpyruvate carboxylase (ppc); 11. citrate synthase (gltA); 12. aconitase
(acn); 13. isocitrate dehydrogenase (icd); 14. glutamate dehydrogenase (gdhA);
15. glutamine synthetase (glnA); 16. malate dehydrogenase (mdh); 17. fumarase
(fumB); 18. fumarate reductase (frdABCD); 19. aspartate transaminase (aspA); 20.
aspartase (aspC). o indicates allosteric activation, o indicates allosteric inhibition.
m '.kA EcoRVmISm
Figure 3-2. Plasmids used to construct mutations in KO 11. FRT sites allow in vivo
excision of the tet gene after integration using FLP recombinase (flp). A. Plasmid
pLOI2065 containing a tet gene flanked by FRT sites. B. Plasmid pLOI2375
containing an interrupted ackA gene.
l "JO lJ -__
I jSynk for A ad B -abols fa C rad D
U 0.5 ) K0No Adi 0.5a K11 NoAdd
-0-KOll I+ e SJ104
S-o- K11+ Acet~ ---- SUl ad
OD0 -Ko- K11O+ cAdehydte OJO-- -o-- -SU12 + Pynwrate
0 12 24 36 48 60 72 84 96 0 12 24 36 48 60 72 84 96
Time (h) Time (h)
1000 10 D
800 :R 800 -
200- 200 -
0 12 24 36 48 6 72 84 6 0 12 24 36 48 60 72 84 96
Tim e (i) Time (1I
Figure 3-3. Effect of media additions and mutations on growth (A, C) and ethanol
production (B, D). Symbols for A and B: 0, KO11 no additive; A, K1 1+
pyruvate; 0, KO11 + acetate; and O KO11+ acetaldehyde. Symbols for C and
D: 0, KO11 no additive; 0, SU102 (ackA) no additive; A, SU104 (adhE) no
additive; and 0, SU102 + pyruvate.
-- _--~_OII_+_Ae____-l 0 UU 2 Pv___
m-KOll + ANo ta1 0 ; 31I2PM v
-ci-- KOll + AcetaHeyde -SUlU 1 Py iivate
0 12 24 36 48 60 72 84 96 0 12 24 36 48 60 72 84
I0 12 24 36 48 60 72 84
0 12 24 48 72 84 0 1224 48 7284
Figure 3-4. Effect of media additions and mutations on organic acid production: acetate
(A/F), format (B/G), lactate (C/H), fumarate (D/I) and succinate (E/J). Symbols
for A-E: U, KO 1l no additive; A, KOl + pyruvate; 0, KO 1l + acetate; and O,
KOl 1+ acetaldehyde. Symbols for F-J: U, KOl 1 no additive; O, SU102 no
additive; A, SU14 no additive; and SU102 + pyruvate.
Figure 3-4. Effect of media additions and mutations on organic acid production: acetate
(A/F), format (B/G), lactate (C/H), fumarate (D/I) and succinate (E/J). Symbols
for A-E: 0, KO11 no additive; A, KO11+ pyruvate; C, KO11 + acetate; and 7,
KO11+ acetaldehyde. Symbols for F-J: 0, KO11 no additive; 7, SU102 no
additive; A, SU104 no additive; and C, SU102 + pyruvate.
0 12 24 36 48 60 72 84 96
1.4 I I I I
0 12 24 36 48 60 72 84 96
0 12 24 36 48 60 72 84 96
0 12 24 36 48 60 72 84 96
Figure 3-4. continued.
I I I
0 3 6 9 12 15 18 21 24
I I I I
6 12 18 24
Figure 3-5. Metabolism of added acetaldehyde and pyruvate during fermentation.
Pyruvate addition is indicated by the large arrow. Acetaldehyde additions (5.6 mM
each) are indicated by the large arrow (initial addition) and the small arrow
(second addition at 12 h). A. Utilization of added pyruvate and acetaldehyde.
Symbols: A, pyruvate utilization by KO 1; 0, pyruvate utilization by SU102; O,
acetaldehyde utilization by KO 11; E, acetaldehyde in KO 11 broth with no
additions; and @, pyruvate in KO 11 broth with no additions. B. Effect of second
acetaldehyde addition on production of fermentation products by KO 11. Symbols:
U, cell mass; A, ethanol; @, format; 0, lactate; A, acetate; *, succinate; and 0,
a.) ( D a.) ( (D 6
e 4M 4 e 4M W1 0 -d
2 I lPyruva e B
FEIJT l Acetaldehyde
PPC/PPS LDH PFL PDH PDC
Figure 3-6. Partitioning of carbon among competing pathways during the initial 3 h of
fermentation. A. Fermentation products after 3 h. B. Relative activity of primary
enzymes that partition 3-carbon intermediates carbon (pyruvate and
phosphoenolpyruvate) through competing pathways. Relative activities were
estimated using fermentation products, expressed as [imol of product per ml
during the initial 3 h of incubation. Endogenous production of acetate in
acetate-supplemented fermentations was assumed to be equal to that for the
control fermentation without additives. Pyruvate decarboxylase activity was
assumed to be equal to ethanol production, except for the
acetaldehyde-supplemented fermentations where it could not be calculated.
Pyruvate dehydrogenase was calculated as the difference between acetate and
format production. Lactate dehydrogenase, pyruvate formatelyase, and
phosphoenolpyruvate carboxylase activities were assumed to equal to the
production of lactate, format, and succinate, respectively. Abbreviations: PPC,
phosphoenolpyruvate carboxylase; PPS, phosphoenolpyruvate synthetase; LDH,
lactate dehydrogenase; PFL, pyruvate formatelyase; PDH, pyruvate
dehydrogenase; PDC, pyruvate decarboxylase.
A DEFICIT IN PROTECTIVE OSMOLYTES IS RESPONSIBLE FOR THE
DECREASED GROWTH AND ETHANOL PRODUCTION DURING XYLOSE
Maintaining inexpensive sources of fuels and commodity chemicals for the U. S.
is a matter of national security. Increasing the production of fuel ethanol offers a potential
solution to this problem. The conversion of lignocellulose to fuel ethanol and other
chemicals typically derived from petroleum would decrease the U. S. dependance on
imported oil (Artzen and Dale 1999). Enteric bacteria are noted for their broad range of
growth substrates, including all the sugars present in the polymers of lignocellulose.
Escherichia coli, a microbial platform for the commercial production of amino acids and
recombinant proteins (Chotani et al. 2000; Akesson et al. 2001), was previously
engineered for the production ethanol by integrating a synthetic operon containing the
ethanol pathway from Z. mobilis (pdc and adhB) into the chromosome (Ohta et al. 1991).
The resultant strain, designated KO 11, fermented all the sugar constituents of
lignocellulose to ethanol with yields approaching 100% (Ohta et al. 1991; Martinez et al.
1999; Ingram et al. 1999).
During batch fermentations with strain KO 11, the volumetric rate of ethanol
production was directly related to the growth of the biocatalyst (Martinez et al. 1999;
Chapter 2; Chapter 3). Cell yield for both the ethanologenic strain and its parent (E. coli
B) was dependent upon the availability of nutrients in a variety of media, despite the
absence of any specific auxotrophic requirements. During the batch fermentation of
xylose (90 g liter1) to ethanol by strain KO 11, cell growth appeared to be limited by the
availability of carbon skeletons derived from the citrate arm of the anaerobic TCA
pathway (Chapter 2; Chapter 3) (Fig. 4-1).
During fermentative metabolism, the TCA pathway is interrupted by the
repression of 2-ketoglutarate dehydrogenase (Iuchi and Lin 1988). The ultimate product
of this pathway, 2-ketoglutarate, is a substrate for glutamate biosynthesis. During growth
in minimal media, glutamate is the most abundant free amino acid in the cytoplasm of E.
coli (Cayley et al. 1991). In addition to its roles in metabolism and protein synthesis,
glutamate biosynthesis is part of the primary response to osmotic stress (Csonka 1989;
Csonka and Hanson 1991). The high osmolarity of the medium used for ethanol
production (0.6 M xylose) would be expected to increase the requirement for glutamate as
a protective osmolyte.
Potassium ions are rapidly accumulated by E. coli in response to osmotic stress.
This is rapidly followed by the accumulation of glutamate (McLaggan et al. 1994), a
negatively charged amino acid and protective osmolyte, to balance the positive charge of
the accumulated potassium. In the closely related organism Salmonella typhimurium,
cells that are restricted in their ability to synthesize glutamate have been demonstrated to
grow poorly during osmotic stress (Csonka 1988; Yan et al. 1996). Mutations preventing
glutamate production were associated with the inability to balance the charge of
intracellular potassium. The resulting decrease in steady-state potassium levels has been
proposed to limit cell growth (Yan et al. 1996).
Alternate protective osmolytes such as glycine betaine (hereafter referred to as
betaine), proline, taurine, dimethylsulfoniopropionate and many others can be
accumulated from the environment. A hierarchy for these osmoprotectants was
empirically determined for salt stress (Randall et al. 1995). Although there have been
conflicting reports concerning this hierarchy for sugar-mediated osmotic stress (Glaasker
et al. 1998), betaine is generally regarded as the most effective protective osmolyte for E.
coli. The effectiveness of betaine for restoration of growth has been shown to cary with
the sugar used for osmotic stress (Dulaney et al. 1968)..
In this study, NMR was used to examine changes in the intracellular pools of
osmolytes in response to genetic changes and nutrient supplements that stimulated cell
growth and ethanol production. Low cell yield and low ethanol production in the absence
of supplements appears to result from a deficit in intracellular glutamate or alternative
Materials and Methods
Microorganisms and Media.
The ethanologenic E. coli strains KO 11 and SU102 (KO 11 AackA; Chapter 3) are
prototrophic. Working cultures were transferred daily on solid medium (1.5% agar)
containing mineral salts, 2% xylose, and 1% corn steep liquor (CSL+X medium; Chapter
2) alternating between 40 mg liter' and 600 mg liter' chloramphenicol. Stock cultures
were stored frozen at -700C in 40% glycerol.
Seed cultures (350C and 120 rpm) and fermentations (350C and 100 rpm) were
grown in either mineral salts medium containing 1% corn steep liquor and 9% xylose
(Chapter 3) or Luria broth (10 g liter -1 tryptone, 5 g liter-' yeast extract, 5 g liter-' NaC )
with 9% xylose. For fermentations, sufficient cell mass to achieve and initial
concentration of 33 mg liter' were harvested by centrifugation (5000 x g; 5 min) and
suspended in appropriate fresh medium. Fermentations were maintained at pH 6.5 by
automatic addition of 2N KOH (Moniruzzaman and Ingram 1998). Stock solutions (100
mM) ofbetaine (Sigma, St. Louis, MO) and dimethylsulfoniopropionate (TCI America,
Portland, OR) were dissolved in deionized water and filter sterilized directly into the
fermentation vessel. Glutamate and acetate were added as described previously (Chapter
2; Chapter 3).
Intracellular osmolytes were analyzed by NMR (Park et al. 1997). After a 24 h
incubation in the fermentation chamber, 700 mL culture was harvested by centrifugation,
washed twice in mineral salt solution containing NaCl (0.6 M) and resuspended in 3
volumes of ethanol (95%). Suspensions were rocked gently 16-24 hr at 40C. Cellular
debris was removed by centrifugation (4"C, 10,000 x g, 30 min). Extracts were dried
under vacuum, disloved in deionized water, dried under vacuum, resuspended in 33%
D20 and filtered (0.2[im) Acetone (10[iL) was used as an internal reference. Data was
obtained using a modified Nicolet NT300 spectrometer operating in the Fourier transform
mode as follows: 75.46 MHz; excitation pulse width, 25 us; pulse repetition delay, 40s;
spectral width 18 kHz and broadband (bi-level) decoupling of protons. For cell extracts,
at least 1000 scans were obtained.
Cell mass was determined from the optical density at 550 nm using a Bausch &
Lomb Spectronic 70 spectrophotometer (1 OD550= 0.33 g liter' dry cell weight), and
ethanol was measured by gas chromatography (Varian 3400CX) (Moniruzzaman. and
Ingram 1998). For the quantitation of the compounds detected by NMR, a standard curve
was generated. The average of all the chemical shift values are reported normalized to
Results and Discussion
Citrate Synthase Flux Limits the Biosynthesis of Glutamate, a Primary Intracellular
E. coli typically maintains large cytoplasmic pools of potassium, glutamate and
trehalose during growth in media containing high concentrations of sugars or salt (Cayley
et al. 1991; Lewis et al. 1990). Previous studies with the ethanologenic E. coli strain
KO 11 demonstrated that the flux through citrate synthase, the first step in glutamate
biosynthesis, limited growth and ethanol production during the fermentation of 90 g liter'
xylose (0.6 M) to ethanol (Chapter 2; Chapter 3). This limitation was proposed to be due
to the drain ofpyruvate to ethanol by the recombinant pathway (pyruvate
decarboxylase-alcohol dehydrogenase) which has a higher affinity for pyruvate than
pyruvate formate-lyase or pyruvate dehydrogenase, competing pathways for aceyl-CoA
biosynthesis (Chapter 2; Chapter 3). Supplementing the CSL+X medium with acetate
increased the availability of acetyl-CoA (Chapter 3), an activator of citrate synthase
(Weitzman 1966). During fermentations with strain K 11, supplementing the CSL+X
medium with glutamate increased the final cell yield and ethanol productivity (Chapter 2)
by bypassing the growth-limiting citrate synthase.
The intracellular osmolyte pools were compared between conditions of lower
growth (CSL+X medium without additives) and higher growth (supplemented with
glutamate or acetate) to investigate whether the inability of strain KO 11 to accumulate
glutamate resulted from restricted citrate synthase flux (Fig. 4-2; Table 4-1).
Fermentations without additives accumulated only proline, while those supplemented
with acetate (activating citrate synthase) or glutamate (bypassing citrate synthase) resulted
in approximately 2-fold higher growth and ethanol productivity. Cells from these
fermentations accumulated approximately the same level of intracellular glutamate,
supporting the hypothesis that a deficit in the accumulation of this protective osmolyte
The intracellular proline, a known osmoprotectant (Csonka 1989;Csonka and
Hanson 1991), was likely accumulated from the CSL provided as a source of complex
nutrients. Though some organisms synthesize proline in response to osmotic challenge
(Kawahara et al. 1989), E. coli can only accumulate this protective osmolyte by active
transport (Smith et al. 1984). However, mutants have been isolated that are less sensitive
to the feedback-inhibition of the proline biosynthetic pathway (Smith 1985). Strains
expressing these genes accumulated high levels of intracellular proline (derived from
glutamate) and were more resistant to high osmotic environments (Csonka 1981; Csonka
et al. 1988). Presumably, the intracellular proline in strain KO11 may have resulted from
a similar spontaneous mutation. Accordingly, such a mutation would also reduce the
Glutamate was accumulated in fermentations with increased growth yield, and the
intracellular concentration of proline decreased (Table 4-1). This further supports the
hypothesis that the intracellular proline was taken up from the medium and was not a
result of biosynthesis. If the drain of intracellular glutamate for proline production had
starved the cells for glutamate, supplementing the medium with proline should have a
sparing effect on the consumption of glutamate. However, proline addition only increased
the intracellular proline pool without affecting either growth or ethanol production (Table
Glutamate is a product of proline degradation (McFall and Newman 1996), but
the degradation of proline has been shown to be inhibited in media of high osmotic
strength (Csonka 1988). Supplementing the medium with an excess ofproline (17 mM)
should provide excess proline for glutamate production. However, the absence of
increased growth and intracellular glutamate in these fermentations confirms the
previously observed inhibition of proline degradation during osmotic stress (Csonka
The accumulation of proline was previously shown not to affect glutamate pools
(Cayley et al. 1992). During experiments in a glucose-mineral salts medium buffered with
MOPS and high NaC1, the primary osmolytes accumulated by E. coli were K glutamate,
MOPS and trehalose (Cayley et al. 1991; Cayley et al. 1992; Lewis et al. 1990). Cultures
in this medium supplemented with proline accumulated this protective osmolyte and
reduced the biosynthesis of trehalose (Cayley et al. 1992). However, the intracellular
glutamate concentration was not significantly altered by the accumulation of proline.
Thus, the intracellular accumulation of proline (from the CSL) by strain KO 11 should not
have affected the glutamate requirement.
Strain KO11 failed to synthesize detectable levels of the osmoprotectant trehalose
(<10 mM) under these conditions. While the presence of proline would have decreased
the synthesis of trehalose (Cayley et al. 1992), significant trehalose should have been
detected. This may be a result of growth on xylose, a pentose. Perhaps the relatively low
ATP yield from xylose catabolism (0.4 ATP/pyruvate) restricts the gluconeogenic
production of glucose and uridine diphosphate-glucose, substrates for trehalose
Genetic Changes to Optimize Carbon Partitioning Increased the Glutamate Pool.
The functional expression of citZ by strain KO 11 (pLOI2514) was previously
shown to increase growth and ethanol production. To confirm that the expression of this
enzyme aides in glutamate accumulation, the intracellular osmolyte pool during
fermentations in the CSL+X medium were analyzed (Fig 4-2; Table 4-1). Similar to the
fermentations supplemented with glutamate or acetate, cells from these fermentations had
an increased glutamate pool. The intracellular accumulation of proline was similar to that
of other experiments with increased growth yields. Thus, the expression of citZ provided
more citrate, ultimately increasing glutamate biosynthesis and the glutamate pool.
A mutation in the primary acetate production pathway (Apta) was previously
shown to increase glutamate biosynthesis (Chang et al. 1999b). This likely resulted from
the accumulation of acetyl-CoA, an activator of citrate synthase (Weitzman 1966).
Acetyl-CoA is also an activator ofphosphoenolpyruvate carboxylase (ppc) (Izui et al.
1981), the controlling step in the biosynthesis of oxaloacetate and co-substrate for citrate
synthase. Thus, the biosynthesis of citrate is regulated, in part, by the availability of
acetyl-CoA. In an analogous study presented here, blocking acetate production (Aack)
increased the intracellular glutamate pool level (Fig. 4-2, Table 4-1).
Glutamate Accumulation Functions in Osmoprotection.
Three different osmoprotectants were tested for their ability to restore growth and
ethanol production, replacing the additional glutamate requirement.(Fig. 4-3). The
addition of 1.0 mM betaine or dimethylsulfoniopropionate (DMSP) increased the cell
yield and ethanol production similar to experiments where glutamate production had been
increased. Taurine, a weak osmoprotectant for E. coli (McLaggan and Epstein 1991),
failed to increase growth or ethanol production. Neither betaine nor presumably DMSP
should provide a source of glutamate. Thus, supplying osmoprotectants to the medium
replaced the need for the accumulation of intracellular glutamate.
To determine the optimal concentration required to restore growth, betaine and
DMSP were added from 0.1-2.0 mM and 0.1-1.0 mM, respectively (Fig. 4-4). Growth
and ethanol were increased in a dose-dependant manner in each instance. The maximum
stimulation of growth and ethanol production by betaine was at the highest level of
betaine tested, 2.0 mM. However, only 0.25 mM DMSP was required for maximum
benefit. Surprisingly, during the fermentation of xylose to ethanol, DMSP is 10-fold more
effective in restoring growth and ethanol production than betaine. Though this is contrary
to previous reports that betaine is the most effective protective osmolyte (Randall et al.
1995), these studies were done using high levels NaCl for osmotic challenge. The ability
of betaine to restore growth has been reported to vary during osmotic challenge with
different sugars (Dulaney et al. 1968). Thus, a specific protective osmolyte may be more
effective during challenge by different osmolytes (sugars and salts).
Replacement of Glutamate by Other Osmoprotectants.
The intracellular osmolyte pools of cells in fermentations supplemented with
betaine and DMSP were examined by NMR. Cells from fermentations supplemented with
betaine were found to contain only detectable levels of betaine (Fig. 4-2), consistent with
previous studies (Cayley et al. 1992). This was likely a result of the properties of the
osmotically activated transport pathways. While the Km of ProP and ProU for proline are
0.3 mM and 2 tiM, respectively, ProU has a 1.3 [iM Km for betaine (Lucht and Bremer
1994). The high level of betaine in the medium (2.0mM) coupled with the low Km of the
primary betaine transport pathway (ProU) explains the exclusive accumulation of this
protective osmolyte. Additionally, the ProP system has a periplasmic, high-affinity
betaine binding protein (KD 1 tiM) which aids in the accumulation of this preferred
Cells from the fermentations supplemented with 0.25 mM DMSP accumulated
both proline and DMSP. There are three possible explanations for the contemporaneous
accumulation of proline and DMSP under these conditions. It is possible that there is an
independent transporter for DMSP. However, DMSP is structurally similar to betaine
(Fig. 4-5), and it is likely transported by the same mechanism. Possibly, the accumulation
of both proline and DMSP results from the Km for proline and DMSP being more similar
to each other. Alternatively, the low level of DMSP in the medium (0.25 mM) compared
to the concentration of betaine used (2.0 mM) may have allowed for the proline (equal
concentrations in both experiments) to more effectively compete for transport by ProP
Neither the betaine nor DMSP supplemented fermentations contained detectable
levels of glutamate (>10 mM; Fig 4-2), confirming that the high glutamate pool was not
needed for biosynthesis, per se. Thus, the requirement for additional glutamate is
presumed to be associated with adaptation to the higher sugar environment. This is
consistent with previous reports using strains ofE. coli or S. typhimurium which were
deficient in glutamate biosynthesis (McLaggan et al. 1994; Csonka et al. 1994; Yan et al.
1996). While the growth of these strains was poor in high osmotic environments, normal
growth was observed in more optimal osmotic environments. Thus, the observed
deficiency in glutamate biosynthesis in strain KO11 was attributed to the inability to
accumulate large quantities of intracellular glutamate for osmoadaptation but not
necessarily for macromolecular biosynthesis.
Betaine from Difco Yeast Extract Restores Growth in Luria Broth Fermentations.
Dulaney and coworkers (1968) demonstrated that Difco yeast extract, a
component of Luria broth (10 g liter-' tryptone, 5 g liter-' yeast extract and 5 g liter-'),
contains betaine by extracting and fractionating Difco yeast extract. Xylose fermentations
with these nutrients yielded high biocatalyst concentrations and high ethanol productivity
(Table 4-1). Cells harvested from these fermentations at 12 h (when the sugar
concentration would be similar to that of the CSL fermentations at 24h) accumulated
proline and betaine were accumulated by strain KO11 (Fig 4-2). The ratio of proline to
betaine was much higher in Luria broth than in CSL+X medium supplemented with 2.0
mM betaine, thus allowing proline to more effectively compete with betaine for transport
by the osmotically active tranporters, ProP and ProU. The lower growth yield in
betaine-supplemented fermentations indicated that although betaine aided in restoring
growth, it was not as effective as the rich Difco nutrients. Some other nutrient in the Luria
broth may be necessary for even higher growth yields (>2 g liter'). The high availability
of carbon skeletons, essential vitamins and minerals in the Luria broth would also have a
sparing effect on all biosynthetic pathways. The higher growth yield and ethanol
productivity observed in fermentations with these nutrients may have resulted from this
general sparing effect.
Growth and ethanol production in the CSL+X medium was restricted by a deficit
in the accumulation of protective osmolytes. Glutamate, typically accumulated in
response to osmotic stress, failed to accumulate due to restricted citrate biosynthesis.
Supplementing the medium with potassium glutamate (2 g liter') bypassed this limitation
and restored the intracellular glutamate pool. Alternately, the addition of sodium acetate
(2 g liter'), an activator of citrate synthase and precursor of glutamate biosynthesis,
restored the intracellular glutamate pool and increased growth and ethanol production.
Together, these results suggested that a deficit in the production of glutamate restricted
Betaine and DMSP increased the growth and ethanol production in a
dose-dependent manner when added to the medium. NMR analysis of the intracellular
osmolytes during these fermentations demonstrated the accumulation of these protective
osmolytes. While the addition of DMSP to the CSL+X medium resulted in the
accumulation of both proline and DMSP, cells from betaine supplemented cultures