ABSTRACT

TIRADO-ACEVEDO, OSCAR. Production of Bioethanol from Synthesis Gas Using

Clostridium ljungdahlii as a Microbial Catalyst. (Under the direction of Dr. Amy M.

Grunden).

As world energy consumption increases, the main sources of energy, which are fossil fuels,

are declining. Biofuels are a promising source of sustainable energy. Nevertheless,

feedstocks for biofuels need to be inexpensive and separate from the human food network.

Lignocellulosic biomass has been identified as one such feedstock. An innovative way to

convert lignocellulosic biomass to biofuels is through a process called gasification-

fermentation. There are known microorganisms that can convert synthesis gas components to

biofuels. One such organism is Clostridium ljungdahlii which can utilize CO and H2 and

produce ethanol. The purpose of the research described herein was to identify and describe

fermentation conditions that support higher ethanol production from synthesis gas by C.

ljungdahlii. Earlier reports had shown that certain acetogens could divert electrons and

carbon flux away from acetate and towards a number of reductant sinks including ethanol. In

the present study, we describe a laboratory-isolated C. ljungdahlii strain (strain OTA1). This

strain produces approximately 2-fold more ethanol than the wild-type (WT) strain.

Furthermore, we demonstrate that addition of oxygen to the cultures improves ethanol to

acetate ratios, as well as total ethanol yields in both C. ljungdahlii WT and OTA1 cultures

grown in media with and without reducing agents. In addition, we demonstrate that pre-

adapting C. ljungdahlii cultures in a medium containing fructose also improves ethanol

formation by this bacterium.

Production of Bioethanol from Synthesis Gas Using Clostridium ljungdahlii

as a Microbial Catalyst

by

Oscar Tirado-Acevedo

A dissertation submitted to the Graduate Faculty of

North Carolina State University

in partial fulfillment of the

requirements for the degree of

Doctor of Philosophy

Microbiology

Raleigh, North Carolina

2010

APPROVED BY:

_______________________________ ______________________________

Dr. Amy M. Grunden Dr. Mari S. Chinn

Committee Chair

________________________________ ________________________________

Dr. Jonathan W. Olson Dr. Eric S. Miller

ii

DEDICATION

To God.

To my wife Johanna, for being there for me, loving me unconditionally and standing next to

me through these years as a PhD student even if she did not understand what my research

was about.

To my parents, for giving me a foundation to build my future on. For giving me everything

when they did not have much.

To my kids, all I do, I will do it for you guys.

iii

BIOGRAPHY

Oscar Tirado-Acevedo was born in Rincón, Puerto Rico to Oscar Tirado-Hernández and

Blanca I. Acevedo-Lorenzo. He is the oldest of three children and had a happy childhood

with a very supportive and united family. Growing up in a coastal town he was interested in

marine sciences since the time he was very little. He completed his Bachelors’ and Masters’

degrees from the University of Puerto Rico. Somewhere along the way, he got married to

Johanna M. Crespo-Torres. Because of a lack of employment opportunities in Puerto Rico

and a desire to experience new opportunities abroad, he and his wife moved to the United

States. After a year working at a microbial pathogenesis laboratory in Nashville TN, they

moved to Raleigh NC, to attend NCSU in the Graduate Program in the Microbiology

Department. He received his PhD degree in Microbiology, has a beautiful daughter, is

expecting a son and is moving on to keep searching for even better things in the future.

iv

ACKNOWLEDGEMENTS

I would like to thank my thesis committee members, Dr. Mari S. Chinn, Dr. Jonathan Olson,

Dr. Eric Miller and especially my thesis advisor Dr. Amy M. Grunden for being a source of

ideas, support and knowledge.

I would also like to thank my laboratory colleagues, Dr. Casey M. Theriot, Dr. Alice

M. Lee, Dr. Jimmy L. Gosse, Dr. Anthony A. Devine, and Rushyannah Killens for ideas,

support and laughs, as well as Drs. Julie Du and Mikyoung Ji. Dr. Flickinger for ideas,

technical discussions and letting me use the GC. I also appreciate the training Dr. Gosse

provided me for the GC operation. I do not know if I would have made it without that

instrument.

I offer my thanks to the NCSU Microbiology Department faculty who would open

their office doors for me any time I had a question, especially Dr. Hassan and Dr. Bruno-

Barcena. Also, I would like to thank Cindy Whitehead for keeping me up to date on forms to

fill-out, datelines, registration, etc. Thank you, Cindy.

v

TABLE OF CONTENTS

LIST OF TABLES ..................................................................................................................vii

LIST OF FIGURES................................................................................................................viii

PRODUCTION OF BIOFUELS FROM SYNTHESIS GAS USING MICROBIAL

CATALYSTS............................................................................................................................1

Abstract....................................................................................................................2

Introduction..............................................................................................................3

Syngas Chemical Catalytic Conversion.................................................................16

Syngas Biotransformation......................................................................................19

Metabolic Engineering of Syngas Biotransformation Processes...........................33

Acetogens and Oxidative Stress Response............................................................35

Conclusions............................................................................................................37

References Cited....................................................................................................46

METABOLIC RESPONSE OF CLOSTRIDIUM LJU GDAHLII STRAINS TO OXYGEN

EXPOSURE.............................................................................................................................64

Abstract..................................................................................................................65

Introduction............................................................................................................66

Materials and Methods...........................................................................................68

Results....................................................................................................................74

Discussion..............................................................................................................82

References Cited..................................................................................................101

INFLUENCE OF CARBON SOURCE PRE-ADAPTATION ON CLOSTRIDIUM

LJU GDAHLII GROWTH AND PRODUCT FORMATION.............................................107

Abstract................................................................................................................108

Introduction..........................................................................................................109

vi

Methods................................................................................................................110

Results and Discussion........................................................................................112

Conclusion...........................................................................................................115

References Cited..................................................................................................121

STUDY CONCLUSIONS...............................................................................................123

APPENDIX......................................................................................................................126

vii

LIST OF TABLES

Table 1-1 Composition of synthesis gas derived from various lignocellulosic biomass

sources.................................................................................................................43

Table 1-2 Summary of microbial catalysts capable of producing biofuels from synthesis

gas........................................................................................................................44

Table 1-3 Thermochemical reactions that occur during gasification of biomass.................45

Table 2-1 Effect of O2 on product profiles in C. ljungdahlii strains at maximum

production............................................................................................................96

Table 2-2 Oxidative stress enzyme activities in C. ljungdahlii WT and OTA1..................97

Table 2-3 Pentose Phosphate Pathway enzyme activities in C. ljungdahlii WT and

OTA1...................................................................................................................98

Table 2-4 Ethanol forming enzyme activities in C. ljungdahlii WT and OTA1..................99

Table 2-5 Intracellular pyridine nucleotide pools of C. ljungdahlii WT and OTA1..........100

viii

LIST OF FIGURES

Figure 1-1. Biofuel production from biomass: (A) hydrolysis-fermentation (B1)

gasification-chemical synthesis and (B2) gasification biosynthesis................39

Figure 1-2. Fuel products obtained from synthesis gas transformation..............................40

Figure 1-3. Diagrams of gasification reactors: A) updraft gasifier; B) downdraft gasifier;

C) fluid-bed gasifier.........................................................................................41

Figure 1-4. Actetyl-CoA or Wood-Ljungdahl pathway used in autotrophic growth by

anaerobes..........................................................................................................42

Figure 2-1. C. ljungdahlii cell growth. A) WT reduced, B) OTA1 reduced, C) WT

unreduced, D) OTA1 unreduced.....................................................................87

Figure 2-2. C. ljungdahlii ethanol production. A) WT reduced, B) OTA1 reduced, C) WT

unreduced, D) OTA1 unreduced.....................................................................88

Figure 2-3. C. ljungdahlii acetate production. A) WT reduced, B) OTA1 reduced, C) WT

unreduced, D) OTA1 unreduced.....................................................................89

Figure 2-4. C. ljungdahlii WT syngas utilization when grown in reduced medium; A) 0%

O2, B) 8% O2, C) 12% O2................................................................................90

Figure 2-5. C. ljungdahlii OTA1 syngas utilization when grown in reduced medium; A)

0% O2, B) 8% O2, C) 12% O2.........................................................................91

Figure 2-6. C. ljungdahlii WT syngas utilization when grown in unreduced medium; A)

0% O2, B) 6% O2, C) 10% O2 ........................................................................92

Figure 2-7. C. ljungdahlii OTA1 syngas utilization when grown in unreduced medium;

A) 0% O2, B) 6% O2, C) 8% O2 .....................................................................93

Figure 2-8. C. ljungdahlii fructose utilization. A) WT reduced, B) OTA1 reduced, C) WT

unreduced, D) OTA1 unreduced.....................................................................94

Figure 2-9. Model for the C. ljungdahlii oxidative stress detoxification system. SOD

(superoxide dismutase), Cat (catalase), Rr (rubrerythrin), Trx (thioredoxin)..95

ix

Figure 3-1. Synthesis gas fermentation reactor..................................................................117

Figure 3-2. C. ljungdahlii culture density in the syngas reactor (A) and syngas-fructose

reactor (B) inoculated with cells pre-grown on fructose (circles), syngas-

fructose (squares), or syngas (triangles)........................................................118

Figure 3-3. C. ljungdahlii production of ethanol (A and B) and acetate (C and D) in the

syngas reactor (A and C) and in the syngas-fructose reactor (B and D)

inoculated with cells pre-grown on fructose (circles), syngas-fructose

(squares), or syngas (triangles).......................................................................119

Figure 3-4. C. ljungdahlii fructose utilization (A) in the syngas-fructose reactor inoculated

with cells pre-grown on fructose (circles), syngas-fructose (squares), or

syngas (triangles) and syngas utilization (B) in the syngas reactor inoculated

with cells pre-grown on syngas.....................................................................120

Figure A-1. C. ljungdahlii-WT metabolism in reduced medium. A) Growth, B) fructose

utilization, C) ethanol production, D) acetate production..............................127

Figure A-2. C. ljungdahlii-WT growing in reduced medium headspace, analysis at

different O2 concentrations. A) 0 % O2 B) 4 % O2 C) 6 % O2 D) 8 % O2

E) 12 % O2 ....................................................................................................128

Figure A-3. C. ljungdahlii-WT metabolism in unreduced medium. A) Growth, B) fructose

utilization, C) ethanol production, D) acetate production.............................131

Figure A-4. C. ljungdahlii-WT growing in unreduced medium headspace analysis at

different O2 concentrations. A) 0 % O2 B) 4 % O2 C) 6 % O2 D) 8 % O2

E) 10 % O2.....................................................................................................132

Figure A-5. C. ljungdahlii-OTA1 metabolism in reduced medium. A) Growth, B) fructose

utilization, C) ethanol production, D) acetate production.............................135

Figure A-6. C. ljungdahlii-OTA1 growing in reduced medium headspace analysis at

different O2 concentrations. A) 0 % O2 B) 4 % O2 C) 6 % O2 D) 8 % O2

E) 12 % O2.....................................................................................................136

Figure A-7 C. ljungdahlii-OTA1 metabolism in unreduced medium. A) Growth, B)

1

CHAPTER 1

Literature Review

Production of Biofuels from Synthesis Gas Using Microbial Catalysts*

Oscar Tirado-Acevedo1, Mari S. Chinn2, Amy M. Grunden1

1Department of Microbiology, North Carolina State University, 4548 Thomas Hall, Campus Box 7615, Raleigh, NC 27695-7615; 2Department of Biological and Agricultural Engineering North Carolina State University D S, Weaver Labs 277, Box 7625 Raleigh, NC 27695-7626

*Published in: Advances in Applied Microbiology, 2010 (70) 57-92.

2

ABSTRACT

World energy consumption is expected to increase 44% in the next twenty years.

Today, the main sources of energy are oil, coal and natural gas, all fossil fuels. These fuels

are unsustainable and contribute to environmental pollution. Biofuels are a promising source

of sustainable energy. Feedstocks for biofuels used today such as grain starch are expensive

and compete with food markets. Lignocellulosic biomass is abundant and readily available

from a variety of sources, for example energy crops and agricultural/industrial waste.

Conversion of these materials to biofuels by microorganisms through direct hydrolysis and

fermentation can be challenging. Alternatively, biomass can be converted to synthesis gas

through gasification and transformed to fuels using chemical catalysts. Chemical conversion

of synthesis gas components can be expensive and highly susceptible to catalyst poisoning,

limiting biofuel yields. However, there are microorganisms that can convert the CO, H2 and

CO2 in synthesis gas to fuels such as ethanol, butanol and hydrogen. Biomass gasification-

biosynthesis processing systems have shown promise as some companies have already been

exploiting capable organisms for commercial purposes. The discovery of novel organisms

capable of higher product yield, as well as metabolic engineering of existing microbial

catalysts, make this technology a viable option for reducing our dependency on fossil fuels.

3

1.1 Introduction

There are three main sources of non-renewable fuels: oil, natural gas, and coal. The

oil market is the largest commodity market in the world (Driesprong et al., 2008). However,

in recent years there have been concerns about uncertainty in gasoline prices, oil production

peaking, environmental damage caused by oil spills and emissions from oil combustion as

well as political instability in many major oil producing nations (Basher and Sadorsky, 2006;

Bentley et al., 2007; Sheehan and Himmel, 1999; Wirl, 2008). Coal is an abundant source of

fuel. Nevertheless, coal burning has been linked to environmental pollution and human

health problems including arsenic poisoning (Liu et al., 2002; Ng et al., 2003; Popovic et al.,

2001). Natural gas is the third most used fossil fuel after oil and coal. In addition it is more

efficient and less carbon intensive than any other fossil fuel

(http://www.eia.doe.gov/oiaf/ieo/world.html, 2009; Lochner and Bothe, 2009). Still, natural

gas reserves in the US and Europe are declining rapidly, the gas is difficult to transport, and

the liquefied natural gas market is only 9% of the total natural gas market (Dresselhaus and

Thomas, 2001; Lochner and Bothe, 2009; Yepes Rodríguez, 2008) For these reasons,

countries around the world are exploring different ways to minimize their dependency on

fossil fuels (Barnwal and Sharma, 2005; Vicente et al., 2005; Zhao and Melaina, 2006).

Biomass has been identified among the renewable energy sources to have the highest

potential to minimize some of these problems (Maniatis and Millich, 1998). Biofuels, fuels

(liquid or gas) that can be produced from biomass (organic material produced by plants,

animals or microbes), can help meet energy demands. Biofuels have the advantage that they

4

can be produced from renewable agronomic raw materials using existing farm machinery and

grain distribution systems independent of global location (Herrera, 2006). Biofuels have the

potential to be sustainable, abundant and environmentally friendly energy sources.

With an annual production of approximately 17.3 billion US gallons, ethanol is

currently the biofuel produced in the greatest quantity world-wide, with the majority of

production in Brazil and the United States (Demirbas and Balat, 2006;

http://www.ethanolrfa.org/resource/facts/trade/, 2009). The first automobiles designed by

Henry Ford were fueled by ethanol (Al-Hasan, 2003). Ethanol is produced commercially in

the US and Brazil from corn and sugar cane, respectively (Herrera, 2006). Some other

countries produce it from wheat, and palm oil (Herrera, 2006). This biofuel has been used

(10% blend) as a replacement for methyl-tert-butyl ether (MTBE) as a fuel oxygenate in

gasoline for the last thirty years (Mackay et al., 2006; Sheehan and Himmel, 1999). In

Brazil, all cars run on a 20-26% ethanol blend up to 100% ethanol in flex-fuel vehicles

(Goldemberg et al., 2008). In today’s US market, ethanol can be used in flexible fuel

vehicles as a blend of 85% ethanol and 15% gasoline (E85). Nevertheless, with less than

2000 E85 refueling stations in the whole nation (most states with less than 100 stations total),

presently there is insufficient infrastructure to make this feasible

(http://www.afdc.energy.gov/afdc/fuels/stations_counts.html, 2009; Sheehan and Himmel,

1999).

Butanol is considered a promising biofuel. Biobutanol is part of the widely known

acetone-butanol-ethanol (ABE) fermentation and can be produced from corn, whey permeate

and molasses (Ezeji et al., 2007; Qureshi and Ezeji, 2008). Its production from fermentation

5

was first described by Pasteur in 1869, and it has been produced commercially since the

beginning of the 20th century as part of the ABE fermentation. This process played a crucial

part in both World War I and II (Jones and Woods, 1986). Production of butanol by

fermentation ended in the 1950’s due to lower production cost from petrochemical sources.

Today, annual production of butanol is estimated to be around 350 million gallons

(Shapovalov and Ashkinazi, 2008). Butanol is used mainly as an industrial intermediate to

make chemicals; primarily butyl acrylate, but also butyl acetate, ethylene glycol, and butyl

xanthate among others. Butanol has the possibility of being used in car engines up to 100%

without need for any modifications (Ramey, 2007). The two major drawbacks of butanol

fermentation are the production of side-products (acetone and ethanol) and product inhibition

at low concentrations (Maddox, 1980). Ongoing research is focused on identifying solutions

to these obstacles (Ezeji et al., 2007; Qureshi and Ezeji, 2008).

Hydrogen is viewed as an ideal fuel for future transportation because it can be

converted to energy without production of CO2; the only byproduct of its combustion is

water (Antoni et al., 2007; Schlapbach and Zuttel, 2001). It can be used in fuel cells as well

as in combustion engines (Schlapbach and Zuttel, 2001). To date, hydrogen is produced by

chemical processes such as methane steam reforming (MSR), where steam reacts with

methane over a nickel catalyst to yield H2 and CO, and electrolysis of water. These

techniques are energy intensive and could be detrimental to the environment (Nath and Das,

2004). Other disadvantages are that it is difficult to transport and store, and would require

the development of a new distribution infrastructure (Antoni et al., 2007). Nevertheless, great

improvements have been achieved in these areas (Chalk and Miller, 2006). A more

6

environmentally friendly way of obtaining hydrogen is by microbial production (biohydrogen

generation). However, biohydrogen production is currently still at a laboratory scale, but

recent work has shown that it can be produced from a number of biomass feedstocks

including sugars, and new microbes with improved hydrogen production capabilities are

being isolated and characterized in laboratories around the world (Akhtar and Jones, 2008;

Davila-Vazquez et al., 2008; Maeda et al., 2008b).

The United States government’s Biomass Program adopted a plan to make biofuels

production cost competitive by 2012 and to reduce gasoline consumption in the US by 30%

by 2030 (http://www1.eere.energy.gov/biomass/biofuels_initiative.html, 2007). The Energy

Independence and Security Act of 2007 requests a production of 36 billion gallons of

renewable fuels by 2022 (Sastri and Lee, 2008), three times higher than current production

(Gura, 2009). To help meet this goal, the US government and federal agencies are planning

to spend more than $2 billion to support research and development of advanced biofuel

technologies (Akhtar and Jones, 2008; http://www1.eere.energy.gov/biomass/pdfs/nbap.pdf,

2008). To date, most first generation biofuels are produced from starch. Nevertheless,

research and industry efforts are moving toward production of biofuels from lignocellulosic

biomass.

Starch is the most abundant storage carbohydrate in plants. Starch contained in grains

or plants is a mixture of amylose (10-30%), and amylopectin (70-90%) (Peters, 2006), both

containing alpha-1,4-linked glucose polymers. These polymers differ in that amylose is a

linear glucose chain and amylopectin contains alpha-1,6-side chains. Grain starch

conversion to biofuels via fermentation is well established and is also the most mature

7

technology today (http://www.nrel.gov/biomass/pdfs/39436.pdf, 2006). In the case of corn,

ethanol is produced by either one of two processes, dry grind or wet mill. Most of the

ethanol is produced by dry grind (Bothast and Schlicher, 2005). In this process, grain is

mixed with water, a thermostable alpha-amylase, ammonia, and lime, and the mixture is

heated in a reactor at temperatures at approximately 88°C. This is where the liquefaction

(starch gelatinization and hydrolysis) step occurs. The mixture is then transferred to the

saccharification tank where sulfuric acid is added to lower the pH of the slurry.

Subsequently, glucoamylase is added and the temperature held at 61°C. This process

releases glucose, which in turn can be converted to biofuels such as ethanol and butanol

through fermentation. The entire process is described in detail in (Kwiatkowski et al., 2006).

The feedstock for this process is the most significant cost input (Bothast and Schlicher,

2005); therefore, biofuel production cost is proportionally affected by the feedstock market

price (Kwiatkowski et al., 2006). For example, corn price average ranges from $1.94 to

$3.24 per bushel (McAloon et al., 2002), but have more than doubled in 2008 (Pimentel et

al., 2009). In turn, corn ethanol has doubled as well (O'brien and Wolverton, 2009).

Feedstocks most used in commercial production of biofuels (mostly ethanol) are

sugar cane, corn, and wheat. This process still relies on economic subsidies as high as $8.7

billion per year (Datar et al., 2004; Kowplow, 2006), and there is evidence that ethanol

production from grains gives a negative net energy balance (Patzek et al., 2005; Pimentel et

al., 2007; Pimentel and Patzek, 2005), which results in a non-ideal process. The use of crop

products for biofuels production has also raised some concerns. For example, the use of corn

for ethanol production has increased the prices of US beef, chicken, pork, eggs, breads,

8

cereals, and milk by 10% to 20% (Pimentel et al., 2009) and has been linked with recent food

shortages and riots around the world (Solomon and Johnson, 2009).

Scientists have been investigating the use of more sustainable, accessible and

economic feedstocks, namely lignocellulosic biomass. Sources for these feedstocks can be of

agricultural, forestry, industrial or municipal residue origin as well as dedicated energy crops

such as switchgrass, miscanthus and poplar among others (Clifton-Brown et al., 2004;

Demirbas and Balat, 2006; Kim et al., 2009; Schmer et al., 2008). Lignocellulosic

(cellulosic) biomass is composed of cellulose (14-70%), hemicellulose (9-22%), and lignin

(8-30%). Cellulose, the most abundant biopolymer on Earth (O'Sullivan, 1997), is a fibrous,

hard, impermeable homopolysaccharide composed of glucose units. Hemicellulose, the

second most common biopolymer in nature (Saha, 2003), is a mixture of pentoses (xylan and

arabinan) and hexoses (mannose, glucose and galactose). Hemicellulose is essential for cell

wall integrity (Saha, 2003). Lignin, a biopolymer that is considered to be the most

recalcitrant of known biopolymers to degradation, (Steffen et al., 2007) forms a matrix with

hemicelluloses around the cellulose forming a rigid polymeric network (Kirk and Farrell,

1987). Lignin is made up of the precursor alcohols coumaryl, coniferyl and sinapyl.

The application of lignocellulosic biomass for biofuels production though hydrolysis-

fermentation is very attractive because of its abundance and higher sugar yield compared to

corn starch (Hamelinck et al., 2005). Nevertheless, this technology is not currently

commercially available since this process is still not cost effective (Himmel et al., 2007;

Mosier et al., 2005). A diagram illustrating the basic hydrolysis-fermentation procedure is

shown in Fig. 1-1A. In this process, feedstocks require a chemical pretreatment in which the

9

carbohydrates-lignin network is broken. This pretreatment hydrolyzes hemicelluloses and

also makes cellulose more accessible for enzymatic hydrolysis (Hamelinck et al., 2005).

Briefly, biomass is generally treated at elevated temperatures with chemicals such as dilute

sulfuric acid, sulfur dioxide, ammonia, and lime (Yang and Wyman, 2008). Cellulose is then

converted to glucose monomers by further hydrolyzing with acids or cellulases. The resulting

liquor needs to be treated to remove unwanted components such as acids and degradation

products of C5 and C6 sugars, such as furfural and hydroxymethyl furfural. The liquor is

separated into solid and liquid fractions and the liquid fraction pH is neutralized. Then the

resulting liquid portion is collected and fermented with either yeast or bacteria. Furthermore,

the product is purified by distillation, and dehydration. This topic has been extensively

reviewed previously (Hamelinck et al., 2005; McMillan, 1992; Mosier et al., 2005; Sun and

Cheng, 2002; Yang and Wyman, 2008).

Although extensive work has been done to develop the hydrolysis-fermentation

process for conversion of biomass to biofuels, there are still significant challenges that need

to be addressed before this process can be commercially viable. These challenges include

slow kinetics of breaking down cellulose to glucose, low yields of individual sugars from

hemicelluloses, and removal of lignin (Himmel et al., 2007). To provide adequate rates of

production and total sugar yields, high enzyme concentrations are required for cellulose

hydrolysis. Furthermore, enzyme recovery and recycling, which is necessary to mitigate the

cost of the hydrolysis-fermentation process, is a complicating factor since the enzymes used

for cellulose degradation have a tendency to bind to the residual lignocellulose, and can,

therefore, be lost during the solid-liquid separation (Eriksson et al., 2002). In addition, the

10

lignin released during biomass hydrolysis can be biocidal and its presence often causes

bioreactor failure (Maness and Weaver, 2002), and since this lignin cannot be broken down

into fermentable sugars, 8-30% of the original biomass is not utilized for product formation.

Also, there is the formation of waste streams such as acid pretreatment materials and toxic

compounds found in acidic hydrolysates of biomass (Datar et al., 2004). Moreover, the

biochemical composition and structure of the biomass (cellulose, hemicellulose, and lignin)

dictates the process performance since this influences the final ethanol yield (Hamelinck et

al., 2005). Unlike the sugar-intermediate biosynthesis technology, in synthesis gas-

intermediate biosynthesis, feedstock biochemical composition does not materially affect the

outcome of the process.

Synthesis gas (syngas) is a product of the gasification of biomass. Gasification is a

well established technology where carbonaceous material (usually coal, wood and charcoal)

is cracked at extreme temperatures (700-1000°C). If pure oxygen is used as the oxidant in

the gasifier, then the resulting synthesis gas is rich in CO and H2. If air is used, then the

resulting gas (producer gas) is a mixture of CO, CO2, H2, CH4, N2, some light hydrocarbons

such as C2H2 and C2H4 as well as heavy hydrocarbons known as tars (Do et al., 2007). The

partial oxidation reactions with oxygen that take place within a gasifier are exothermic.

Steam can also be used as an oxidant in indirect gasification. The result of these

thermochemical reactions is an endothermic and often heat transfer limited, but

thermodynamically efficient process (Sipma et al., 2006). The ratio of the components of

synthesis gas varies depending on the biomass source and the gasification conditions

employed (see Table 1-1).

11

Gasification of biomass to provide vehicles with fuel has been in use since the early

1930’s (McKendry, 2002). Due to a petroleum products shortage during World War II, this

technology flourished in some European countries providing fuel for both civilians and

militaries (Dasappa et al., 2004). More recently, in the 1980’s and 1990’s, synthesis gas was

successfully used in the US and Europe for heat and electricity (Faaij, 2006;

http://fossil.energy.gov/programs/powersystems/gasification/gasificationpioneer.html, 2008

). In 2007, the US Department of Defense released plans to use synthesis gas in its

military bases and vehicles

(http://newsblaze.com/story/2006020806383900003.mwir/topstory.html, 2006). Synthesis

gas can also be transformed into a number of different products such as methanol, ethanol,

hydrogen, dimethylether and others via chemical catalysts (Fig. 1-2). This idea is not new

and has been developed in the past few decades (Wilhelm et al., 2001). Nevertheless, these

are expensive processes subjected to high pressures and temperatures (Quinn et al., 2004;

Takeguchi et al., 2000). Furthermore, syngas coming out of the gasifier has many

contaminants leading to catalyst poisoning (Leibold et al., 2008). Some synthesis gas

transformations to biofuels such as ethanol, butanol and hydrogen can be performed using

chemical as well as biological catalysts (Fig. 1-1B.1 and 1-1B.2). Biological processes, while

slower than chemical reactions, have a number of advantages such as higher yields (even

similar to direct fermentation of biomass (Spath and Dayton, 2003)), specificity, and

generally greater potential for decreased catalyst poisoning (Younesi et al., 2008). Some key

biological transformations related to end product formation are irreversible, these also occur

at ambient temperatures and pressures, therefore requiring minimum energy and lower cost

12

(Klasson et al., 1992a; Klasson et al., 1992b). Pioneering work by Dr. Clausen’s group

focused on production of biofuels from synthesis gas using microbial catalysts, and they

proposed methane production from synthesis gas using a mixed culture of methanogens and

hydogenotrophic bacteria (Barik et al., 1988b; Klasson et al., 1990). They also suggested the

possibility of biological production of ethanol from synthesis gas when there was no known

organism that could perform that reaction (Barik et al., 1988a). The number of investigations

on methane production from syngas has declined during the last decade. However, a number

of new organisms capable of producing biofuels from synthesis gas components have been

isolated (Table 1-2). In addition, microbial ethanol production from synthesis gas has

flourished to the point that Coskata, Inc. has a developed a commercial demonstration unit

this year (http://www.cleantech.com/news/4995/coskata-leaks-word-demo-plant-and-r, 2009)

and has plans to have a full commercial plant operating by 2011. INEOS Bio has a pilot

syngas to ethanol production plant in operation (http://www.ineosbio.com/57-

Welcome_to_INEOS_Bio.htm, 2009) and Syngas Biofuels Energy, Inc. is making efforts for

commercialization of biological conversion of syngas to butanol

(http://www.syngasbiofuelsenergy.com/, 2009). Plans and research to make microbial

biohydrogen production from biomass cost competitive by 2020 are also in progress.

Hyvolution, a multi-country effort funded by the European Union is working on the

development of a bioprocess for cost-effective production of hydrogen from biomass

(http://www.biohydrogen.nl/hyvolution, 2009). Also, Sapporo Breweries, Ltd. in

collaboration with the Brazilian government announced plans to operate a plant for

production of hydrogen from industry waste

13

(http://www.tradingmarkets.com/.site/news/Stock%20News/2239395/, 2009). None of these

biohydrogen projects are planning to utilize synthesis gas from biomass as their feedstock;

however, recent published research described briefly in this work (see Section IV. C) appear

very promising in regards to the development of industrial biohydrogen production from

synthesis gas.

1.2 Biomass gasification

Carbonaceous material can be used as feedstocks for gasification to produce syngas

streams. Researchers have experimented (Table 1-1) with feedstocks as diverse as wood,

sawdust, grass straw, nut and cacao shells, olive husks, even meat and bone meal (a by-

product of rendering industries) (Soni et al., 2009). This is important since synthesis gas can

then be produced from local biomass resources providing energy independence to both

developed and developing countries. The gasification of solid waste and biomass is

generally more complex than coal gasification due to the diversity of the carbon based

materials. Biomass gasification also needs to occur at lower temperatures since biomass is

more reactive than coal (Huber et al., 2006).

Synthesis gas produced from biomass contains contaminants such as alkali

compounds, H2S, HCl, NH3, HCN, potassium, sodium, and tars that are detrimental to the

gasifier equipment as well as catalysts for Fischer-Tropsch (F-T) reactions (Tijmensen et al.,

2002). Gasification occurs through a combination of complex reactions that include drying,

pyrolysis, combustion and finally reduction. Pyrolysis occurs in the absence of oxygen and

releases volatiles and gases from the biomass and produces char (solid carbonaceous

14

material). Combustion occurs when the oxidant reacts with carbon and water from the

biomass, producing CO2 and H2O. In the reduction reaction, oxidant as well as CO2 and H2O

produced in the combustion step react with the carbon in char producing CO as well as H2.

Two other important reactions occurring in the gasifier are the water-gas shift reaction in

which water reacts with CO to form H2 and CO2 and methanation, where CO reacts with H2

to form CH4 and H2O. If the combustion to provide heat to the process is generated in a

separate reactor, the process is called indirect gasification. Table 1-3 shows some of the

reactions that take place in gasifiers.

1.2.1. Gasifiers

Although there are many different gasifier designs in use, the three main types are the

updraft, the downdraft and fluidized-bed gasifiers. In the updraft gasifier (Fig. 1-3A), the

feedstock enters from the top of the chamber where it is dried by the syngas leaving the

chamber. Simultaneously, the gas leaving the chamber is cooled by the feedstock. The dried

feedstock travels down the vessel where pyrolysis to char occurs. The char that is generated

continues moving down the gasifier vessel where it is reduced and also interacts with the

oxidant for combustion. Ashes fall through a grate to the bottom of the gasifier. Some of the

advantages of updraft gasifiers are their simple construction, low cost, ability to handle high

moisture and inorganic content, and their high energy efficiency due to the low temperature

of the gas leaving the chamber. Some of the disadvantages are that the resulting syngas can

contain high levels of tars and hydrocarbons. Therefore if this gas will be used for F-T

15

conversion, extensive cleanup is required. Furthermore, updraft gasifiers have a feed size

limit, and slagging (molten residual ash that can be recycled for many applications) potential.

Downdraft gasifiers are very similar to updraft gasifiers (Fig. 1-3B), except that the

feedstock and oxidizer in downdraft gasifiers both enter from the top of the gasifier. The gas

passes though the hot zone combusting the tars and leaving the reactor from the bottom.

Some of the advantages of this design are that it has a fairly simple design and is low cost,

and it produces a relatively cleaner gas with very low tar formation. Some of the

disadvantages are that the system requires low moisture and ash feedstock, can only use

feedstocks within a limited particle size range (between 1-30 cm), and it has low efficiency

because the product gas leaves the gasifier at higher temperatures, which requires an

additional cooling system as compared to an updraft gasifier.

The third type of gasifier is the fluidized bed gasifier (Fig. 1-3C) This is a more

recent configuration, but it is also the most popular type (Bricka, 2007). In this design, the

feedstock size is reduced to a powder, and is mixed with the fluidizing material, which is

usually silica sand, ceramic or alumina. The oxidizer and mixed feedstock enter the reactor

from the bottom, where they form a bed of hot fluid, where most of the conversion to

synthesis gas occurs. One of the biggest advantages of this type of gasifier is the uniform

temperature distribution in the fluid bed. Also, carbon conversion could be up to 100%, and

it is a good design for large scale applications. Some of the problems with this type of

gasifier are that it produces a gas with high suspended particulates that need to be pre-heated

prior to introduction to the fluidized bed and there can be a loss of bed fluidization due to

feedstocks’ ash contents.

16

Gasification technology is significant to chemical or microbial catalytic syngas

conversion process development as quality of the gaseous feedstock influences application,

product formation and system efficiency. Optimization of the gasification unit operation in

combination with downstream catalytic conversion will improve the application of this

thermochemcial approach to bioenergy production.

1.3 Syngas chemical catalytic conversion

Fischer-Tropsch (F-T) is a surface catalyzed polymerization process that uses CHx

monomers formed by hydrogenation of adsorbed CO in order to obtain hydrocarbons with a

broad chain length and functionality (Iglesia, 1997). This process utilizes metal catalysts such

as Co, Ru, Rh and Fe catalysts. F-T plants have been operated around the world since 1938

(Dry, 2002). Syngas can be chemically converted to ethanol and butanol by F-T chemistry

and to hydrogen by the water-gas-shift reaction (WGS) (see Fig. 1-2). A successful chemical

transformation of syngas requires a purified syngas and a fixed CO/H2 ratio for maximum

conversion. Therefore, removal of H2S and organic sulfur compounds as well as excess CO2

is necessary. This cleanup process typically accounts for 60-70% of capital costs for plant

operation (Dry, 2002).

1.3.1. Ethanol Synthesis

In this process synthesis gas is heated to 210-350°C and compressed up to 1-7 MPa.

The gas is converted to alcohol across a fixed bed catalyst as described by Eq. (1). The

17

reaction rate is high, and completion takes seconds to minutes, with up to 60% CO

conversion to ethanol (Wei et al., 2009). Next the gas is cooled, this allows the alcohols to

condense and separate from the unconverted syngas. The liquid alcohols are then further

refined by alcohol separation and purification steps. Some of the catalysts that have been

historically used contain rhodium (Rh), cobalt, molybdenum and others including multi-

component catalysts (Takeuchi et al., 1985). Rh seems to be the best catalytic metal for

synthesis gas to ethanol conversion for its ability to perform all of the four specific functions

that a catalyst should perform. These include: dissociation of the adsorbed CO to carbon and

oxygen, hydrogenation of the adsorbed carbon to methyl species, insertion of non-dissociated

CO into the methyl species to form an adsorbed acyl species, and hydrogenation of the acyl

species to form the ethanol product (Haider et al., 2009). Some of the problems with these

chemical catalysts are their non-selectivity (i.e. a mix of alcohols is produced) and low

conversion yields (Subramani and Gangwal, 2008). Also the high cost of Rh may affect

commercialization of this technique. The reader is referred to references (He and Zhang,

2008; Spivey and Egbebi, 2007) for a full review of syngas to ethanol chemical catalysts.

2CO + 4H2 catalyst CH3CH2OH + H2O (1)

1.3.2 Hydrogen Synthesis

Synthesis gas produced from biomass can also be enriched thermochemically for

hydrogen production. Although hydrogen production yield from biomass is relatively low

(12-14% based on biomass weight), this technology has been viewed as very promising

(Asadullah et al., 2002; Demirbas, 2005). Subjection of synthesis gas to the water-gas shift

18

(WGS) reaction is the most widely used process (Nath and Das, 2003). This reaction

produces H2 and CO. In this process, syngas coming from the gasifier is transferred to a

WGS reactor where the H2 concentration is increased. In this reactor, steam and CO react to

produce H2 and CO2 (Eq. 2). Usually, the operating temperatures for the WGS reaction

range from 330 to 530°C (Tonkovich et al., 1999).

Hydrogen from syngas can also be produced by steam methane reforming (SMR).

Similar to natural gas SMR, in synthesis gas SMR, CH4 and other light hydrocarbons in the

gas are reacted with steam over a catalyst, usually nickel (Eq. 3). This reaction is carried out

at temperatures ranging from 700 to 1100 °C and pressures up to 2.5 MPa (Demirbas, 2007).

Out of all chemical production to hydrogen, production from synthesis gas SMR gives the

highest conversion efficiency and concentration of hydrogen (Ciambelli et al., 2009).

However, again, tars, dust, sulfur alkali compounds and other impurities can cause catalyst

block or poisoning in this process (Ciambelli et al., 2009). Still the cost of hydrogen

production from biomass gasification is three times higher than the current price of hydrogen

from steam methane or natural gas reforming (Balat et al., 2009; Demirbas, 2005).

CO+ H2O catalyst H2 + CO2 (2)

CH4 + H2O catalyst 3H2 + CO (3)

1.3.3 Butanol Synthesis

Over the last several decades, chemical catalytic systems for synthesis of higher

weight alcohols such as butanol, through CO hydrogenation have been developed (the overall

19

reaction is described in Eq. 4). Catalysis for higher alcohol synthesis can be achieved in

two ways. One is to produce methanol and branched alcohols, using modified methanol

synthesis with Cu/ZnO based catalysts (Christensen et al., 2009). These reactions generally

take place at temperatures around 400°C and pressures of 20.0 MPa (Smith and Anderson,

1983). The other process to produce higher weight alcohols is to form straight-chain

alcohols, using MoS2 and Co/Cu-based catalysts (Christensen et al., 2009; Surisetty et al.,

2009). These catalysts usually operate at temperatures of 275–325°C, and pressures of 7.5–

10MPa (Herman, 2000). The biggest disadvantage of these methods is, like with the ethanol

process, low selectivity. This in turn adds cost to the process due to the need for alcohol

separation. A maximum butanol selectivity of 4% of total alcohol was achieved when a

Cu/ZnO catalyst promoted with 0.5% K2CO3 was used (Smith and Anderson, 1983). Sun and

colleagues (Xu et al., 2004) showed that use of a Fe–CuMnZrO2 catalyst resulted in butanol

formation selectivity as CO conversion increased, but the butanol yield was only 7.5%

alcohol by weight.

4CO + 8H2 catalyst C4H9OH + 3H2O (4)

1.4 Syngas Biotransformation

1.4.1 Acetyl-CoA Pathway and Carbon Monoxide Dehydrogenase

Scientists have suggested that before an O2 atmosphere appeared, the first autotrophs

on Earth, organisms able to use CO or CO2 as their sole source of carbon, utilized the acetyl-

20

CoA pathway (Pereto et al., 1999; Ragsdale and Wood, 1991b; Russell and Martin, 2004).

Apparently, this pathway is still exclusively used by anaerobes (Henstra et al., 2007). The

acetyl-CoA or Wood-Ljungdahl pathway (see Fig. 1-4) was first described in the acetogen

Moorella thermoacetica (formerly known as Clostridium thermoaceticum), a heterotroph

capable of producing 3 moles of acetate from one mole of glucose. The pathway, which is

not cyclic like the Calvin cycle or the reverse TCA cycle, is formed by two branches. These

branches have been called the Eastern and Western branches and were the research focus of

Lars Ljungdahl and Harland Wood, respectively (Ragsdale, 1997). The Eastern branch

produces the methyl group of acetyl-CoA and the Western produces the carbonyl group. The

methyl group is formed from the reduction of CO2 to formate which is converted to

formyltetrahydrofolate and is reduced to methyltetrahydrofolate. These steps involve

formate dehydrogenase, and a series of tetrahydrofolate dependent enzymes (Drake, 1994;

Ragsdale and Wood, 1991b). Even though the pathway was described in a heterotroph, it has

been a model for autotrophic growth from CO2 or CO. Briefly, the organism can synthesize

acetyl-CoA by reducing CO2 with electrons produced by a hydrogenase from H2 (Eq. 5). If

CO is used, it acts as both a source of carbon and electrons (Eq. 6) (Drake, 1994). These

reactions are exergonic and therefore allow ATP formation and energy conservation through

both substrate-level phosphorylation and the electron transport chain (Ragsdale and Wood,

1991b).

Carbon monoxide dehydrogenase (CODH) is the central enzyme in this pathway

(Wood et al., 1986), and it has also been characterized in a diverse group of organisms. In

acetogens, this enzyme is responsible for reducing CO2 to CO yielding the carbonyl group of

21

acetyl-CoA and it also catalyzes the final step in synthesizing acetyl-CoA from CH3, CO and

S-CoA. Acetogens convert acetyl-CoA to acetate gaining an ATP. Some of these organisms

can also reduce acetyl-CoA to acetaldehyde and ethanol using electron donors such as

NAD(H) and NADP(H). This results in a net ATP consumption (Klasson et al., 1992b). In

chemolithoautotrophs, this enzyme also enables the utilization of CO as the sole carbon and

electron source by catalyzing the oxidation of CO to CO2 (Eq. 7). In these organisms, energy

is conserved through an electron transport chain. In hydrogenogenic carboxydotrophs, the

CODH reaction is the same as Eq. 7. However, the electrons are transferred to a membrane

associated hydrogenase that combines the generation of hydrogen with the translocation of

protons. This generates the proton gradient needed for ATP formation by ATP synthase. In

aceticlastic archaea, the CODH works in a reverse manner (Eq. 8) in which acetyl-CoA is

formed from acetate. Acetyl-CoA is cleaved and the methyl group is reduced to CH4 and CO

is oxidized to CO2. Energy in this system is also generated through an electron transport

chain. In methanogens able to grow on CO2/H2 or CO, CODH drives the formation of

acetyl-CoA from methyltetrahydrosarcinapterin and CO as well as the oxidation of CO to

CO2.

In Methanosarcina barkeri a hydrogenase couples generation of hydrogen with the

translocation of protons much like the process that occurs in hydrogenogenic

carboxydotrophs. In sulfate reducing bacteria, CODH functions very similarly to the reverse

acetyl-CoA pathway in methanogens. More recently, there has been evidence of a CODH

enzyme in hyperthermophilic archaea able to grow on CO. These CODHs are very similar to

counterparts in methanogens, such as Methanosarcina acetivorans C2A and Methanosarcina

22

mazei Gö1(Lee et al., 2008a). There has even been found a hyperthermophilic bacterium,

Carboxydothermus hydrogenoformans, containing five different forms of this enzyme (Wu et

al., 2005). For a complete review on CODH the reader is referred to (Ferry, 2003).

As is shown in sections IV. A, IV. B and IV. C, microbial production of ethanol,

hydrogen and/or butanol from synthesis gas components depends on CODH, the acetyl-coA

pathway or both. Therefore, we can expect the acetyl-CoA pathway and Carbon Monoxide

dehydrogenase to be critical not only for microorganisms surviving in a CO or CO2

atmosphere, but to the whole biofuels production from synthesis gas process.

2CO2 + 4H2 CH3COOH + 2H2O (5)

4CO + 2H2O CH3COOH + 2CO2 (6)

CO + H2O CO2 + 2H+ + 2e- (7)

CH3COOH CH4 + CO2 (8)

1.4.2 Ethanol from Syngas

Ethanol, butanol, hydrogen, biodiesel and methanol are all currently being evaluated

as next generation biofuels. Of these, ethanol is by far the biofuel produced in the greatest

quantities world-wide (Demirbas and Balat, 2006). Brazil is the number one ethanol

producer in the world with 41% of the total production, followed very closely by the United

States (Herrera, 2006). Most of this ethanol is produced by microbial fermentation of sugars

from either sugar-cane or corn starch. To make ethanol a commercial fuel contender, the

feedstock has to be switched to lignocellulosic biomass. Hydrolysis-fermentation technology

23

has proven to be expensive and labor intensive. Up to 40% of the carbon present in the

biomass is lost in the form of lignin and most microorganisms used in this process are unable

to utilize 5-carbon sugars in the hydrolysates. Plus, ethanol produced this way has not been

able to compete with fossil fuel derivatives like gasoline and diesel. Biomass gasification

can yield up to 100% carbon conversion to gas components and fermentation of syngas to

ethanol has been shown to be commercially feasible

(http://www.cleantech.com/news/4995/coskata-leaks-word-demo-plant-and-r, 2009).

The first microorganism shown to catalyze conversion of synthesis gas components to

ethanol (Eq. 9 and 10) was the acetogen Clostridium ljungdahlii (Barik et al., 1988a) named

after Dr. Lars Ljungdahl to honor his work on clostridia and acetogens. Even though ethanol

production from synthesis gas was detected, the main product was acetate. Initially, a molar

ratio of ethanol to acetate of 1:9 and an ethanol concentration of less than 1 g/L was obtained

in batch cultures (Vega et al., 1989). It was quickly learned that yeast extract had an

influence on the product ratio and that the ethanol production was non-growth related.

Therefore, yeast extract concentration was greatly reduced or eliminated completely and

replaced by cellobiose. This increased both ethanol and cell concentrations. Adding

reducing agents to the media seemed to alter electron flow to NADH formation and in turn,

increasing ethanol production. These first experiments where done in batch cultures. By

applying the culture performance information acquired through experimentation and

operating two continuously stirred tank reactors (CSTR) in series (the first to promote growth

and the second one for increased ethanol production), they were able to improve ethanol

production by 30 fold (Klasson et al., 1991b). A cell recycle apparatus was added to the

24

CSTR and pH was held at 4.5, agitation was set at 450 rpm, gas flow rate was 30 ml/min and

liquid flow rate ranged from 3.5 to 12 ml/h. These modifications increased the cell

concentration from 800 mg/L to 4000 mg/L, and increased ethanol production to 50 g/L with

an ethanol to acetate molar ratio of 21:1 and CO and H2 conversions of 90% and 70%

respectively (Klasson et al., 1993; Phillips et al., 1993a). Investigations also showed that C.

ljungdahlii is quite tolerant of sulfur gases. It is able to grow and uptake CO and H2 in the

presence of up to 2.7% H2S or 5% carbonyl sulfide (Klasson et al., 1993; Smith et al., 1991).

This is relevant since syngas contains a considerable amount of these gases.

Work by Ghasem Najafpour and colleagues has investigated the effect of gas

pressures in ethanol and acetate production by C. ljungdahlii in batch cultures (Najafpour and

Younesi, 2006; Younesi et al., 2005). CO consumption and CO2 production was the highest

when syngas was applied at 1.6 and 1.8 atm, showing that CO uptake is not inhibited by

synthesis gas at high pressures. H2 and CO2 consumption occurred after 72 h of incubation

time in cultures with 1.6 and 1.8 atm of syngas, by which time CO had already been

exhausted. This shows that CO was the preferred substrate for cell growth. This was

expected since free energy is higher when CO is used as a substrate as compared to CO2

(Barik et al., 1988a). Growth and acetate formation were not affected by the higher syngas

pressures. On the contrary, ethanol production was enhanced at syngas pressures of 1.6 and

1.8 atm by four fold. For their system, ethanol yield was only 0.6 g/L and the ethanol to

acetate molar ratio was 0.54.

More recently, different synthesis gas compositions as well as different agitation

speeds and gas flow rates were compared for their effect on ethanol production by C.

25

ljungdahlii (Younesi et al., 2006). Using synthesis gas with a composition of (volume %)

55% CO, 20% H2, 10% CO2 and 15% Ar, a bioreactor agitation speed of 300 rpm and a gas

flow rate of 10 ml/min, CO utilization was only 14% (by volume). When agitation was

increased to 400 rpm, CO utilization only increased to 18%. When pure CO or a CO rich gas

mix (70% CO, 15% H2 and 15% Ar) was used, cell concentration increased by 28%, but CO

utilization remained low. Throughout the experiment a maximum of 6g/L ethanol and 7g/L

acetate was produced.

Since ethanol production by C. ljungdahlii is assumed to be non-growth related, the

effects of nitrogen limitation and low pH on ethanol and acetate production under a nitrogen

atmosphere were investigated (Cotter et al., 2009a). Results showed that C. ljungdahlii

requires vitamins and trace elements to maintain high cell viability in media lacking a

nitrogen source. Also, ethanol and acetate production were significantly lower in resting cells

as compared to growing cultures. Cultures were stable when initial pH of the non-growth

media was lowered to 4.5. Nevertheless, cell viability dropped from 100% in the control pH

(6.8) to 11.1%. Ethanol production was also significantly lower in cultures at low pH as

compared to the control cultures.

The effects of pH and gas flow on growing batch cultures have also been studied,

under a synthesis gas atmosphere (Cotter et al., 2009b). The gas flow rates tested were 5, 7.5

and 10 ml/min and culture pHs were 6.8 and 5.5. It was reported that initial lower pH

conditions resulted in lower cell densities and end product formation. The greatest ethanol

formation occurred during cell growth, which is not in agreement with the theory that ethanol

production is non-growth associated. Higher ethanol yields were obtained at gas flows of 7.5

26

ml/min with an ethanol to acetate ratio of 1:8. CO2 concentration coming out of the reactor

increased in all treatments tested. CO and H2 were consumed with CO2 consumption

occurring at the later stages of the culture growth.

A few years after C. ljungdahlii was described, Clostridium autoethanogenum,

another acetogen able to produce ethanol from CO was isolated from rabbit feces (Abrini et

al., 1994). Synthesis gas fermentation studies with C. autoethanogenum are limited.

However, ethanol production from syngas components has been shown to be only a fraction

of the molar yield of C. ljungdahlii, and the ethanol to acetate molar ratio was around 1:1or

less (Abrini et al., 1994; Cotter et al., 2009a; Cotter et al., 2009b). In their non-growing

experiments Chinn and colleagues, (Cotter et al., 2009a) observed that C. autoethanogenum

culture densities decreased substantially, but were 100% viable. The C. autoethanogenum

cells also consumed xylose, meaning that cells were metabolically active even under nitrogen

limitation conditions. Nevertheless, ethanol and acetate production were lower than when

cells were grown in the presence of a nitrogen source. In batch cultures with constant

synthesis gas flow, higher ethanol yields were achieved at a gas flow rate of 10 ml/min with

an ethanol to acetate molar ratio of 1:16. CO and H2 were consumed and CO2 was produced

in similar quantities across all flow rates (Cotter et al., 2009b).

The only published research showing growth and ethanol production from actual

biomass producer gas has been done with the acetogen Clostridium carboxidivorans P7

(Datar et al., 2004; Liou et al., 2005). This organism (originally named bacterium P7) was

isolated from an agricultural settling lagoon and was extensively studied because of its ability

to produce six times more ethanol than acetate (Rajagopalan et al., 2002). In these studies,

27

the researchers operated the reactor by switching from bottled gas to producer gas. In the first

stages, the culture cells were grown in batch liquid with continuous bottled synthesis gas

flow at 180 ml/min and pH was controlled at 5.9. When cell growth plateaued, continuous

liquid flow at 1.5 ml/min was initiated. Acid production was shown to be growth associated

and ethanol production was minimal at this stage. Both H2 and CO were consumed during

cell growth. When the reactor was operated with producer gas, cell growth and H2 uptake

stopped, CO consumption decreased, and cells began to wash out. Nevertheless, ethanol

production increased and acid production decreased, indicating that ethanol production was

non-growth associated. When the reactor was returned to bottled syngas and liquid batch

mode, cells began to grow again indicating that the cells had not lost viability. CO uptake

started again, but not H2 consumption. The group reported that this H2 uptake inhibition

could be caused by nitric oxide (NO) and acetylene present in the producer gas used (Ahmed

et al., 2006; Ahmed and Lewis, 2007). NO is a known hydrogenase inhibitor, and in this

organism, it affects H2 uptake at concentrations above 40 ppm (Ahmed and Lewis, 2007).

Therefore, the producer gas used may need to be processed to remove NO to concentrations

below 40 ppm. Nevertheless, their results seem promising for the development of

commercial processes.

Yet another bacterium has been isolated for its ethanol production from synthesis gas.

In this case a thermophile, Moorella sp. HUC22-1 (Sakai et al., 2004). Even though

lowering pH and cell recycle have improved ethanol production from 1mM to 15 mM, the

ethanol to acetate molar ratio obtained was 1:45 (Sakai et al., 2005). An acetaldehyde

dehydrogenase (Aldh) and three alcohol dehydrogenases (AdhA, AdhB, and AdhC) have

28

been described from this organism (Inokuma et al., 2007; Sakai et al., 2004). Aldh was

shown to catalyze the thioester cleavage of acetyl-CoA, as well as the thioester condensation

from CoASH and acetaldehyde. It also was shown to have activity towards both NADP(H)

and NAD(H), but activity towards NAD(H) was determined to be eight fold higher. AdhA

was observed to catalyze the NADP(H)-dependent reduction of acetaldehyde as well as the

oxidation of ethanol. This enzyme can also use NAD(H) as a cofactor but with decreased

activity. AdhB was active only when NADP(H) was used as a cofactor. AdhC showed no

activity with any of the cofactors used. Both AdhA and AdhB were active towards reduction

of a variety of aldehydes. Surprisingly, the highest activities were toward n-butylaldehyde

and isobutylaldehyde, even though this organism has not been shown to produce butanol.

Finally the study reported higher aldh gene expression when cells were grown on H2/CO2,

but lower adhABC expression in cells grown on H2/CO2 than cells grown on fructose.

Another possible use for the discussed autotrophic microorganisms is the conversion

of CO2 emitted from industrial operations such as fossil-fueled power plants. This CO2 can

be converted to ethanol with a suitable electron donor. Shu and Wiesner (Shu and Wiesner,

2008) ran a simulation using either C. ljungdahlii or Moorella sp. HUC22-1 for conversion

of CO2 from a power plant to ethanol. They determined that H2 was the most favorable

electron donor when compared to acetate or methane since the reaction occurs at near

ambient temperatures. Hydrogen can be produced by microorganisms (Section IV. C), or

with current technologies like steam methane reforming. They propose to use an aerated

membrane reactor (AMR) as the bioconversion vessel. As described by Shu and Weiner

(2008), inside the AMR there is a membrane bundle that liquid media and gases pass

29

through. Bacteria are attached to the membranes as well as suspended in the medium. This

reactor does not require agitation like the others, thus reducing operational costs. In their

simulation, the ethanol yield was significantly lower than yields obtained by the Clausen

group (Klasson et al., 1993). Nevertheless, when C. ljungdahlii conversion was modeled,

ethanol production costs by this method appeared to be less than the current ethanol market

price. These findings appear very promising by themselves; imagine if even higher ethanol

yields were reached.

2CO2 + 6H2 bacteria CH3CH2OH +3H2O (9)

6CO + 3H2O bacteria CH3CH2OH + 4CO2 (10)

1.4.3 Hydrogen from Syngas

Hydrogen is a versatile fuel as it can be used for power generation as well as a

transportation fuel (in association with fuel cells). It is considered to be a clean fuel since

water is the only byproduct when it is burned. It has been proposed that hydrogen will likely

be the most common alternative fuel used for automotive consumption in the next 25 years

(Demirbas, 2007). Natural gas and coal SMR are the least expensive known technologies for

H2 production. This process results in a gas mixture of mainly CO and H2. Subsequently, CO

is chemically converted to CO2 by the water-gas shift (WGS) reaction producing additional

H2. Syngas may also be converted to hydrogen via the WGS reaction. At a production cost

of up to $50/GJH2 (www.iea.org/Textbase/techno/essentials.htm, 2007), these processes do

not make hydrogen a viable replacement for fossil fuels at present (Ismail et al., 2008).

30

Hydrogen can also be produced by biological systems. Biological hydrogen

production is environmentally friendly and requires less energy input compared to chemical

processes (Ismail et al., 2008). Rhodospirillum rubrum, Rhodopseudomonas palustris P4,

Citrobacter sp. and Rubrivivax gelatinosus CBS are photoautotrophic and

chemoheterotrophic microorganisms capable of performing the gas-water shift (GWS)

reaction (Ismail et al., 2008; Jung et al., 1999; Klasson et al., 1992b; Maness and Weaver,

2002; Markov and Weaver, 2008; Najafpour et al., 2003; Najafpour et al., 2004). Two

enzymes that mainly contribute to the GWS reaction in these organisms are CODH and

hydrogenase. The former catalyses the oxidation of CO, and hydrogenase mediates the

reduction of protons to H2 (Maness et al., 2005; Najafpour et al., 2004). Biological GWS is

thermodynamically favorable at room temperature, (CO + H2O �H2 + CO2 ∆G= - 20

kJ/mol) and atmospheric pressure, compared to the chemical catalysis where a two-stage

process is required as well as high temperature (>200°C) (Benemann, 1999). Therefore,

minimum energy requirements and low process cost are expected (Ismail et al., 2008).

R. rubrum grows quickly and reaches high cell concentrations that uptake CO more

rapidly than other similar organisms capable of performing the GWS reaction (Klasson et al.,

1992b; Najafpour et al., 2003; Najafpour et al., 2004). It also tolerates small amounts of O2

and sulfur often present in syngas (Klasson et al., 1992b). As a result, this strain is the

favorite organism for studies investigating biohydrogen production from syngas. R. rubrum

requires a light source for growth; however, H2 production is independent of light intensity

(Najafpour et al., 2004). An organic carbon source is needed for this organism to efficiently

consume CO. The highest CO consumption (90-97 %) has been determined to occur when

31

R. rubrum is provided with acetate as the organic carbon substrate (1-2 g/l), resulting in a

98% hydrogen production yield (Najafpour and Younesi, 2007; Najafpour et al., 2004).

Investigations involving continuously stirred tank reactors (CSTR) of R. rubrum and with

continuous CO flow resulted in hydrogen yields and CO conversions of 87% and 95%

theoretical values, respectively (Younesi et al., 2008). This type of bioreactor was stable

for continuous operation for 27 days (Ismail et al., 2008). Most hydrogen production from

syngas research has been done using artificial syngas. A mixture of pure gases is normally

present in synthesis gas at a fixed concentration (e.g. 56.0% N2, 17.2% CO, 16.3% CO2, and

8.8% H2). In a study by DiSpirito and colleagues, R. rubrum growing on artificial syngas and

on “real” producer syngas was compared. It was determined that producer syngas had no

negative effect on growth rates, biomass production, hydrogen production or carbon

monoxide consumption (Do et al., 2007).

R. gelatinosus CBS is another promising strain for its use in syngas-to-hydrogen

conversion. In the presence of CO, this organism carries out the WGS reaction in both light

and dark conditions (Maness and Weaver, 2002). These cells are capable of converting

100% of CO in the gas phase to H2 in the dark (Markov and Weaver, 2008). Its tolerance for

oxygen (Maness and Weaver, 2002) and its capacity to use CO as the sole carbon and energy

source (Maness et al., 2005; Markov and Weaver, 2008) make it an attractive biocatalyst.

32

1.4.4 Butanol from Syngas

Butanol, like ethanol, can be produced from fermentable sugars, synthesis gas and

glycerol. Butanol has a number of notable qualities that make it a suitable alternative fuel. Its

energy content is 30% more than ethanol (Qureshi and Ezeji, 2008). It can be mixed with

gasoline in any proportion or be used as the sole fuel component (100% butanol) in

unmodified car engines (Ramey, 2007). It carries less water; and therefore, it can be

transported through existing gasoline pipelines (Dürre, 2007). Reports of biological butanol

formation date back to Louis Pasteur. He reported an alcohol product from a clostridial

culture (Dürre, 2007). The acetone-butanol-ethanol (ABE) fermentation was essential during

World War I. Acetone was needed to prepare munitions, and it was in great shortage at the

time. Production of acetone by fermentation meant a constant supply of acetone to Britain

and its allies (Dürre, 2007). C. acetobutylicum has been the model organism for research in

ABE fermentation from sugars, but other species have also been extensively investigated.

Some of the most studied are Clostridium beijerinckii, Clostridium

saccharoperbutylacetonicum, and Clostridium saccharobutylicum. Butyribacterium

methylotrophicum is an anaerobe capable of using 1-carbon compounds such as CO2 (in the

presence of H2), CH4, and formate as carbon sources in addition to fermentable substrates

like glucose, sucrose and glycerol (Zeikus et al., 1980). It also possesses the advantageous

ability to produce butanol from synthesis gas (Grethlein et al., 1990; Lynd et al., 1982;

Zeikus et al., 1980). It is one of the most versatile CO utilizing bacteria (Grethlein et al.,

1991). Other fermentation products are ethanol, acetate and butyrate. The first attempts at

33

investigating this strain for butanol production from CO yielded concentrations of 1.4 g/L

(Worden et al., 1991). After some changes in fermentation set up, such as operation at pH

5.5 and continuous cell cycle, butanol production from CO was improved by more than

200% (Grethlein et al., 1991). Nevertheless, with classic ABE strains producing butanol at

more than 400 g/l (Lee et al., 2008b), B. methylotrophicum is not yet a contender for

commercial biobutanol production. Another interesting, but less studied strain for butanol

production, is C. carboxidivorans P7. Being able to produce up to 4 times more ethanol than

butanol from CO or producer gas, this strain has mostly been studied for its ethanol

production capabilities (Section IV. B) (Datar et al., 2004; Liou et al., 2005; Rajagopalan et

al., 2002).

1.5 Metabolic Engineering of Syngas Biotransformation Processes

Escherichia coli strain KO11 is the classic example of metabolic engineering for

improved biofuels production in a microorganism. In this organism, Ingram and colleagues

successfully integrated the pyruvate decarboxylase and alcohol dehydrogenase II genes from

Zymomonas mobilis (Ohta et al., 1991). This strain has been the foundation of a significant

number of studies (Underwood et al., 2002; Yomano et al., 2008; Yomano et al., 1998).

After genetic tools were developed and its genome sequenced, Z. mobilis has become an

important part of recent bioethanol production research. Since this organism can only utilize

simple C6 sugars, research has been focused on providing it pathways for xylose and

arabinose utilization (Rogers et al., 2007).

34

Metabolic engineering in E. coli has also been applied for enhanced hydrogen

production. Over-expression of the native formate hydrogen lyase (FHL) in E. coli resulted

in about three times more hydrogen production than the wild type strain (Yoshida et al.,

2005). Through a series of deletions and mutations, Wood and colleagues successfully un-

regulated FHL, inhibited hydrogen uptake and redirected glucose metabolism towards

formate resulting in around a five-fold hydrogen production increase (Maeda et al., 2007).

They also randomly bioengineered the HycE hydrogenase (produces hydrogen from formate)

to obtain a 23-fold increase in hydrogen production from E. coli (Maeda et al., 2008a). It

was recently shown that similar to E. coli, Citrobacter amalonaticus Y19 possess a FHL

complex (Kim et al., 2008). This finding potentially provides many opportunities for

engineering this bacterium. Some work has also been done in an effort to redirect metabolism

towards H2 production in R. palustris (Rey et al., 2007).

To date, metabolic engineering studies focused on butanol producing bacteria have

targeted enhancing butanol production, increasing tolerance to solvents and selecting for

butanol over other products. Findings from these studies have provided insight into how to

meet some of the challenges of using metabolic engineering strategies to improve

biotransformation of synthesis gas. For example, regular plasmid vectors are degraded in

ABE Clostridia due to DNA restriction patterns in these bacteria. Also genetic tools for these

strains are not as developed as with E. coli or Bacillus subtilis. Some of these hurdles have

been overcome (Heap et al., 2007; Mermelstein et al., 1992) and to date, some of the aims

have been met by inactivating acetate and butyrate kinases, the solvent formation repressor

35

gene solR, and over-expression of heat shock proteins among other approaches (Green et al.,

1996; Harris et al., 2001; Tomas et al., 2003).

Just recently, attention has been given to modifying or introducing butanol (or its

isomers) production pathways in E. coli. Yukawa and colleagues (Inui et al., 2008)

successfully cloned and expressed butanol production genes from C. acetobutylicum ATCC

824. Similar approaches have been taken with yeast (Steen et al., 2008). Also, higher

isobutanol production has been observed in E. coli cells over-expressing Alcohol

Dehydrogenase A from Lactococcus lactis (Atsumi et al., In press).Throughout the literature,

many microorganisms have been genetically manipulated for enhance biofuel production.

However, synthesis gas has not been the source of energy or growth for organisms in these

experiments. This is probably due to the lack of genetic information and tools for syngas

utilizing organisms. The work of Nishio and colleagues (Inokuma et al., 2007) is a good start

on identifying what enzymes are more active under syngas fermentation and which ones are

not, helping to identify good candidate genes for genetic modifications. Metabolic

engineering of these microorganisms may further facilitate biomass conversion to biofuels as

well as lowering the cost of these processes.

1.6. Acetogens and oxidative stress response

The acetyl-CoA pathway used by acetogens relies on enzymes that are highly

susceptible to O2 (Ljungdhal, 1986; Ragsdale and Wood, 1991b). Some of these enzymes

are: formate dehydrogenase, methylene-H4folate reductase, disulfide reductase, and carbon

monoxide dehydrogenase (Ljungdhal, 1986). Therefore, acetogens have been classified as

36

anaerobes, even obligate anaerobes (Drake et al., 2008). Nevertheless, acetogens have been

isolated from environments that experience transient O2 exposure such as leaf litter, forest

soil, termite guts, and plant roots (Brioukhanov and Netrusov, 2007; Kusel et al., 2001).

Thus, it is not surprising that an array of different Reactive Oxygen Species (ROS) protecting

enzymes have also been detected in acetogens and characterized.

Moorella thermoacetica (Clostridium thermoaceticum), the model “obligately”

anaerobic homoacetogen, studied for the elucidation of the acetyl-CoA pathway contains a

five-gene cluster encoding for rubrerythrin (Rbr), rubredoxin oxidoreductase (Rbo), two

rubredoxins (Rub) and a type A flavoprotein suggested to protect this bacterium from ROS

(Das et al., 2001). In a study by Drake and colleagues, M. thermoacetica, Sporomusa

silvacetica and Clostridium magnum were able to grow and reduce O2 concentrations up to

1.9 % (v/v) in the gas phase (Karnholz et al., 2002). These three organisms exhibited NADH

oxidase as well as peroxidase activities. However, superoxide dismutase and catalase

activities were not detected. Sporomusa termitida and Sporomusa sp. strain TmAO3, both

isolated from termite guts and Acetobacterium woodii, isolated from estuarine sediments

show H2-dependent O2 reduction. An NADH oxidase was recently isolated and

characterized from Clostridium aminovalericum. This was the first enzyme of this type

characterized from the genus Clostridium (Kawasaki et al., 2005). In this the same study,

genes encoding for ROS protecting enzymes such as glutathione peroxidase, thioredoxin,

Rbr, peroxidase, various SODs and thioredoxin reductase were found in the genome of both

Clostridium aminovalericum and C. acetobutylicum.

37

It has been shown that acetogens can divert electrons from acetate formation routes

to alternative terminal electron-accepting processes when exposed to O2 (Drake et al., 2008).

For example M. thermoacetica can use nitrate as e- acceptor. Additionally, Clostridium

glycolicum RD-1 is capable of diverting e- for the formation of H2 and lactate. Furthermore,

this organism shows a 2-fold increase in ethanol formation concomitant with approximately

2-fold reduction in acetate production. These results are encouraging as researchers might be

able to increase ethanol production from synthesis gas at the expense of acetate formation by

exposing these acetogens to certain amounts of O2. This would be beneficial especially for

improving biofuel production from organisms where genetic systems are not available. On

the other hand, it could also be used in combination with metabolic engineering of organisms

where genetic systems are already developed.

1.7. Conclusions

Current US ethanol production from corn starch is not sustainable. It directly impacts

food and feed markets, depends on subsidies, and has questionable energy balances. Biofuels

from cellulosic biomass have promise to be a renewable alternative to fossil fuels. Industry

can take advantage of feedstocks such as energy crops, agricultural residues as well as

industrial and municipal waste. Developing technologies around lignocellulosic biomass

feedtsocks can make biofuels cost competitive without negatively altering global food and

environmental affairs.

38

Fermentation of synthesis gas obtained from biomass has proven to be a viable

approach to produce biofuels. When compared to lignocellulose hydrolysis-fermentation,

gasification-fermentation has higher product yields and lower energy input (Piccolo and

Bezzo, 2009; Wei et al., 2009). Microbial catalysts generally can manage synthesis gas

contaminants making them more robust than some chemical catalysts. Nevertheless, quality

of the synthesis gas coming out of the gasifier has to be closely monitored since

contaminants such as NO, acetylene and O2 can inhibit activities from these microbes.

Seemingly most of these problems have been overcome since a commercial demonstration

plant utilizing this process is already in operation, with possibilities of several other

companies opening new plants in the near future. Furthermore, microbial ethanol production

from CO2 emitted from power plants seems promising and would support value-added,

environmentally-conscious fuel production.

Even though a number of microorganisms have been isolated with abilities to convert

synthesis gas to biofuels, the list is still small and extensive physiological and metabolic

research is limited for most of these isolates. Consequently, metabolic tools and techniques

are non-existent for many of these microbes. The fact that some acetogens produce higher

ethanol concentration as a response to O2 exposure opens up new possibilities for

improvement of biofuels production. Furthermore, the combination of new and better

isolates with advancement in metabolic engineering would improve gasification-fermentation

process costs, product yields, and overall performance.

39

Figure 1-1. Biofuel production from biomass: (A) hydrolysis-fermentation (B1) gasification-chemical synthesis and (B2) gasification-biosynthesis.

40

Figure 1-2. Fuel products obtained from synthesis gas transformation, modified from Spath and Dayton (2003).

41

Figure 1-3. Diagrams of gasification reactors: A) updraft gasifier; B) downdraft gasifier; C) fluid-bed gasifier.

42

Figure 1-4. Actetyl-CoA or Wood-Ljungdahl pathway used in autotrophic growth by anaerobes.

43

Table 1-1. Composition of synthesis gas derived from various lignocellulosic biomass sources.

Composition vol %

Source CO CO2 H2 N2 CH4 Other

Switchgrass1 14.7 16.5 4.4 56.8 4.2 3.4

Pine wood chips2 16.1 13.6 16.6 37.6 2.7 13.4

Willow3 9.4 17.1 7.2 60.5 3.3 2.5

Cacao shells3 8 16 9 61.5 2.3 3.2

Dairy biomass4 8.7 15.7 18.6 56 0.6 0.4

Kentucky bluegrass straw5 12.9 17.4 2.6 64.2 2.1 0.8

Demolition wood/paper residue3 9.2 16.1 6.1 63.2 2.8 2.6 1- (Datar et al., 2004) 2- (Corella et al., 1998), 3- (van der Drift et al., 2001), 4- (Gordillo and Annamalai, In press, doi:10.1016/j.fuel.2009.07.018 ), 5- (Boateng et al., 2007).

44

Table 1-2. Summary of microbial catalysts capable of producing biofuels from synthesis gas

Microorganism Source of isolation Growth temperature Biofuel Reference

Acetobacterium woodii Estuary sediment Mesophilic Ethanol (Balch et al. 1977; Buschhorn et al. 1989)

Butyribacterium methylotrophicum Sewage digester Mesophilic Ethanol, butanol (Zeikus et al. 1980)

Caldanaerobacter subterraneus subsp. pacificus Marine hot vent Thermophilic H2 (Sokolova et al. 2001)

Carboxydocella themoautotrophica Hot spring Thermophilic H2 (Sokolova et al. 2002)

Carboxydothermus hydrogenoformans Hot swamp Thermophilic H2 (Gerhardt et al. 1991)

Citrobacter sp. Y9 Wastewater sludge digester Mesophilic H2 (Oh et al. 2003)

Clostridium autoethanogenum Rabbit feces Mesophilic Ethanol (Abrini et al. 1994)

Clostridium glycolicum RD-1 Sea grass roots Mesophilic Ethanol (Kusel et al. 2001)

Clostridium ljungdahlii Chicken waste Mesophilic Ethanol (Barik et al. 1988; Tanner et al. 1993)

Clostridium carboxidivorans Lagoon sediment Mesophilic Ethanol, butanol (Liou et al. 2005)

Moorella sp. HUC22-1 Mud Thermophilic Ethanol (Sakai et al. 2004)

Rhodopseudomonas palustris P4 Wastewater sludge digester Mesophilic H2 (Jung et al. 1999)

Rhodospirillum rubrum Dead mouse residue Mesophilic H2 (Zurrer and Bachofen 1979; Gest 1995)

Rubrivivax gelatinosus Lake sediment Mesophilic H2 (Uffen 1976)

Thermincola carboxydiphila Hot spring Thermophilic H2 (Sokolova et al. 2005)

Thermococcus strain AM4 Hydrothermal vent Thermophilic H2 (Sokolova et al. 2004b)

Thermolithobacter carboxidivorans Hot spring Thermophilic H2 (Sokolova et al. 2007)

Thermosinus carboxidivorans Hot spring Thermophilic H2 (Sokolova et al. 2004a)

45

Table 1-3. Thermochemcial reactions that occur during gasification of biomass

Reaction Chemistry Enthalpy change

Partial Oxidation C + 1/2O2 CO -268 MJ/kg mole

Complete Oxidation C + O2 CO2 -406 MJ/kg mole

Water Gas Reaction C + H2O CO + H2 +118 MJ/kg mole

Water Gas Shift Reaction CO + H2O CO2 + H2 -42 MJ/kg mole

Methane Formation CO + 3H2 CH4 + H2O -88 MJ/kg mole

46

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CHAPTER 2

Metabolic Response of Clostridium ljungdahlii Strains to Oxygen Exposure

Oscar Tirado-Acevedo1, Jimmy L. Gosse2, Mari S. Chinn2, and Amy M. Grunden1

1Department of Microbiology, North Carolina State University, 4548 Thomas Hall, Campus Box 7615, Raleigh, NC 27695-7615; 2Department of Biological and Agricultural Engineering North Carolina State University D S, Weaver Labs 277, Box 7625 Raleigh, NC 27695-7626

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ABSTRACT

Research in the biofuels area has been moving away from feedstocks that compete

with food supplies. Lignocellulosic biomass has been identified among the renewable energy

sources to have the highest potential to minimize dependency on fossil fuels. Any

carbonaceous material can be converted to syngas (a mixture of CO, CO2, N2, and H2) by

means of gasification. Clostridium ljungdahlii ferments syngas to ethanol resulting in the

conversion of the CO and H2 present in the syngas to ethanol and acetic acid. ATP synthesis

resulting from acetate production skews the final liquid products to higher acetate production

compared to ethanol. There have been previous studies reporting that certain acetogens can

produce higher amounts of ethanol compared to acetate when oxygen is added to the

cultures. For this project, a range of oxygen concentrations was added to the culture

headspace of both C. ljungdahlii wild type and the laboratory derived OTA1 strains. In the

present study, we demonstrate that these acetogens can grow in the presence of oxygen and

that oxygen is consumed in the course of the syngas fermentations. Furthermore, we

demonstrate that addition of oxygen to the cultures improves ethanol to acetate ratios, as well

as total ethanol yields in cultures grown in media with and without reducing agents. In

addition, cell extracts generated from the syngas fermentation cultures exposed to the varying

amounts of oxygen were assayed for oxidative stress enzyme activities, and NADH oxidase

activity was observed; however, catalase activity was not detected. NAD(P)H concentrations

decrease after cultures reach mid-log growth phase suggesting these adenine nucleotides

were utilized in response to oxygen exposure and for acetyl-CoA reduction to ethanol.

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2.1. Introduction

Acetogenic bacteria (acetogens) form acetate as a final product of fermentation.

This group of organisms is very significant. They are ubiquitous in the environment and

reside at an important step of the anaerobic food chain. Acetogens are indirectly involved in

green-house gas formation. Acetate is considered to be the primary precursor for methane

(CH4) formation (Chauhan and Ogram, 2006). Methane is the second most important

greenhouse gas after CO2 (Frankenberg et al., 2005; Thauer et al., 2008). Furthermore,

approximately 70% of the CH4 produced on earth is produced by methanogens from acetate

(Chauhan and Ogram, 2006). Acetogens can metabolize a wide range of organic substrates

such as sugars, formate, methanol and amino acids (Morton et al., 1993). They are an

important part of the carbon cycle on Earth because of their ability to utilize CO and CO2

through the acetyl-CoA pathway as energy and carbon sources. Acetogens have also been

shown to participate in the anaerobic metabolism of methyl-tert-butyl-ether (MTBE)

(Mackay et al., 2007) as well as the degradation of halogenated compounds (Zinder, 2010).

These compounds are particularly problematic due to their persistence in the environment,

bioaccumulation in various organisms, and cytotoxic effects on wildlife and humans (Luo et

al., 2008; Teuten et al., 2005).

Acetogens have shown potential economical uses as well. Recently, it has been

demonstrated that some acetogens are able to degrade cellulose, converting it directly to

acetate (Karita et al., 2003; Wolin et al., 2003). Also, acetogens have the ability to convert

sugars and synthesis gas (a mixture of mainly CO2, CO and H2) to acetic acid (a valuable

industrial chemical), and/or ethanol, butanol and hydrogen (important biofuels) among other

67

products (Allen et al., 2009; Cotter et al., 2009a; Cotter et al., 2009b; Heiskanen et al., 2007;

Kundiyana et al., 2010; Munasinghe and Khanal; Tirado-Acevedo et al., 2010). Significant

advances have been made by companies for commercialization of syngas fermentation

technologies (Gnansounou and Dauriat, 2010; Munasinghe and Khanal).

Clostridium ljungdahlii (ATCC 49587T) was the first acetogen isolated for its ability

to produce ethanol and acetate from CO (Barik et al., 1988a; Tanner et al., 1993a). Ethanol

to acetate production ratios reported have been as low as 1:9 and as high as 21:1 (Vega et al.,

1989). Besides CO, this bacterium also utilizes a number of other carbon sources including

pyruvate, fructose, and xylose among others. Although a number of studies have been

performed to evaluate the ability of C. ljungdahlii to ferment syngas components and to

improve ethanol production by this organism, few studies have been done from a functional

genomics perspective. However, recently the C. ljungdahlii genome sequence was

published, and a genetic system developed for heterologous butanol production in C.

ljungdahlii was presented (Köpke et al., 2010). It has been found that in addition to the

acetyl-CoA and ethanol fermentation pathways, the C. ljungdahlii genome has genes

encoding reactive oxygen species detoxification enzymes, Pentose Phosphate Pathway (PPP)

enzymes and aldehyde oxidoreductase (AOR).

Acetogens have traditionally been classified as strict anaerobes (Fuchs, 1986),

nevertheless, they have been isolated from different aerobic or microaerobic environments

(Drake et al., 2008). It has been demonstrated these bacteria are equipped with an assortment

of oxidative stress enzymes and that certain acetogens can even reduce oxygen by different

means (Boga and Brune, 2003; Das et al., 2005; Karnholz et al., 2002; Küsel et al., 2001). In

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addition, it has been suggested that exposing acetogens to microaerobic conditions trigger a

shift in electron flow towards ethanol, lactate, H2, and/or NH4+ production instead of acetate

formation (Drake and Daniel, 2004; Drake et al., 2008; Küsel et al., 2001). Clostridium

ljungdahlii was isolated from a microaerobic environment (Tanner et al., 1993a). In

addition, preliminary data suggests that this bacterium can tolerate relatively high

concentrations of O2 in the gas phase. Therefore, the objectives of this study were to

determine the tolerance, metabolic, and physiological response of C. ljungdahlii toward O2.

2.2 Materials and Methods

2.2.1 Organisms and inoculum preparation

Clostridium ljungdahlii (ATCC 55383), an acetogen isolated from chicken waste, and

Clostridium ljungdahlii-OTA1, a laboratory derived strain, were grown on a modified

Reinforced Clostridial Medium (mRCMfs). This medium contained (per liter, pH 6.8):

proteose peptone (10 g), beef extract (10 g), yeast extract (3 g), fructose (5 g), 5 mg/mL

resazurin solution (0.1%, w/v final), 50 ml PETC salt solution [(per L): NH4Cl (20 g), KCl

(2 g), MgSO4.7H2O (4 g), NaCl (16 g), KH2PO4 (2 g), and CaCl2 (0.4 g)], 10 ml PETC trace

element solution [(per L): nitrilotriacetic acid (2 g), MnCl2.4H2O (1.3 g), FeSO4.7H2O

(0.4 g), CoCl2.6H2O (0.2 g), ZnSO4.7H2O (0.2 g), CuCl2.2H2O (0.02 g), NiCl2.6H2O

(0.02 g), Na2MoO4.2H2O (0.02 g), Na2SeO3 (0.02 g), and Na2WO4.2H2O (0.025 g)] and 10

ml modified Wolfe’s vitamin solution [(per L): biotin (2 mg), folic acid (2 mg), pyridoxine

(10 mg), thiamine HCl (5 mg), riboflavin (5 mg), nicotinic acid (5 mg), calcium D (+)

pantothenate (5 mg), vitamin B12 (5 mg), p-aminobenzoic acid (5 mg), and lipoic acid

69

(thioctic acid) (5 mg)]. The medium was prepared and dispensed in 160 ml serum bottles (50

ml medium/bottle). Bottles were sealed with butyl rubber stoppers and aluminum seals,

connected to a vacuum manifold using a needle and a 0.22 µm filter and made anaerobic by

cycling three times (30 s cycles) between vacuum headspace evacuation and sparging with

artificial synthesis gas (a pure bottled mixture containing 20% CO, 20% CO2, 10% H2 with

N2 balance). The bottles were then autoclaved for 30 min (121 °C, 15 psig). Prior to

inoculation, when needed, cysteine–HCl (1 ml 2.5% w/v, 0.5 g final) and sodium sulfide (1

ml 2.5% w/v, 0.5 g final), were added to the bottles. The culture bottles were then inoculated

with 5 ml freezer stock cells and incubated with shaking at 100 rpm at 37°C. After 24 h, 2.5

ml of culture were transferred to fresh medium and incubated as above. After 12 h, 2.5 ml of

actively growing cells were transferred to fresh medium and incubated as above. After

inoculation, any excess pressure was released from the bottles by inserting a needle

connected to a 0.22 µm filter and a hose with its end immersed in a water trap. After 12 h of

incubation, O2 was injected by syringe at the indicated concentration (headspace volume).

Liquid and gas samples were taken at 0, 12, 24, 48, and 72 h. Results are reported as an

average of six replicates.

2.2.2 Gas and liquid product analysis

Headspace gas samples were collected using a sample lock, 5 ml gas-tight syringe

(SGE, Ringwood, Australia). The samples were analyzed by gas chromatography using an

Agilent 7890A gas chromatograph containing a 13823 molsieve 5 Å zeolite molecular sieve

packed stainless steel column (6 ft x 1/8 in. i.d. Supelco) and a thermal conductivity detector.

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Argon was used as the carrier gas at a flow rate of 30 ml/min. The injector and detector

temperatures were 150 °C and 250 °C respectively, and the oven temperature was 160 °C.

Liquid samples (500 µl) were acidified with 125 µl 25% m-phosphoric acid. The

acidified samples were centrifuged for 10 min at 14,000 x g at room temperature. The sample

supernatants were analyzed for ethanol and acetate concentrations by gas chromatography

using an Agilent 7890A gas chromatograph containing a 0.25 µm J & W DB-FFAP column

(30 m x 0.32 mm i.d.) and a flame ionization detector. Argon was used as the carrier gas at a

flow rate of 30 ml/min. The injector and detector temperatures were 250 °C and the oven

temperature 160 °C.

2.2.3 Intracellular enzyme assays

Cell extracts were prepared as follows, 694 ml of sterile, modified un-reduced RCMfs

in 2 L stoppered bottles were inoculated with 5% (total volume) mid-log C. ljungdahlii

culture. These cultures were grown as above for 12 h to an O.D.600 of 0.3 – 0.4.

Experimental bottles were treated with 6 % O2 (headspace volume) and returned to the

incubator. After 2 h (early response) or 24 h incubation (later response), cultures were

transferred to an anaerobic chamber and dispensed into centrifuge bottles. The bottles were

sealed and centrifuged (25 min at 11,000 x g, 4 °C). The cell pellets were washed with 40 ml

anaerobic potassium phosphate buffer (50 mM, pH7.0) and centrifuged as described above.

Cell pellets were resuspended in 3 ml anaerobic potassium phosphate buffer (50 mM, pH7.0)

per gram of wet cell pellet. Five hundred microliters of the cell suspension were transferred

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to 2 ml screw cap tubes. The cell suspensions were either processed immediately for assays

or stored at -80 °C.

Five hundred microliters of 0.1 mm disruption glass beads (RPI, Illinois, USA) were

added to the tubes. Samples were vortexed horizontally (10 min, maximum speed) using a

MoBio vortex adapter 13000-V1 for Vortex-Genie 2® (MoBio, California, USA), and

centrifuged (10 min, 21,000 x g, 4 °C). Supernatants were collected and kept on ice for

assays. All assays were carried out in 3 ml total volume at 25 °C in anaerobic cuvettes

except for the superoxide dismutase assay that was performed aerobically in 96-well plates.

All enzyme assays are reported as an average of six replicates, whereas the nucleotide pool

measurements represent three replicates.

NADH or NADPH oxidase was assayed as described by Stanton and Jensen, 1993.

Reactions were initiated by adding cell extract (10-100 µg protein). Catalase was assayed by

measuring the disappearance of H2O2 at 340 nm (Hassan and Fridovich, 1978). The assay

contained, 2 ml 50 mM potassium phosphate buffer pH 7.0 and 1 ml 59 mM H2O2 solution

(150 µL 30% H2O2 in 25 mL 50 mM, pH 7.0 potassium phosphate buffer). This mix should

give an Abs240 of 0.858 ± 0.02. The catalase reactions were initiated by adding cell extract

(10-100 µg protein). Peroxidase was assayed as described by Poole and Ellis, 1996, with the

following modification, 200 µM NADH or NADPH (final concentration) was used. SOD

activity was determined using a commercially available WST SOD assay kit (Dojindo

Laboratories, Kumamoto, Japan) (Peskin and Winterbourn, 2000). This kit uses the water-

soluble tetrazolium salt WST-1 which produces a formazan dye upon reduction with a

superoxide anion. This reduction rate is linearly related to xanthine oxidase activity, which is

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inhibited by SOD. Therefore, the 50% inhibition activity of SOD can be determined by the

colorimetric method. Sample volumes were adjusted so that a 10 µg protein amount was

loaded per well. Enzyme activity, defined as percent inhibition of WST-1 reduction, was

calculated according to the manufacturer’s protocol using the formula SOD activity

(inhibition rate %) = {[(Ablank 1 - Ablank 3) - (Asample - Ablank 2)]/ (Ablank 1 - Ablank 3)} x 100,

where blank 1 lacks sample solution, blank 2 lacks xanthine oxidase solution and blank 3

lacks both sample and xanthine oxidase solutions.

Alcohol dehydrogenase assays contained 1.5 ml 100 mM Tris-HCl pH 8.5, 0.5 ml 2M

ethanol, and 1 ml of 0.025 M NAD+ or NADP+. Reactions were initiated by adding cell

extract (10-100 µg protein) and the appearance of NADH or NADPH was measured at 340

nm. Acetaldehyde dehydrogenase assays were performed as described in Clark and Cronan,

1980, in which the appearance of NADH or NADPH was measured at 340 nm.

Glucose-6-P dehydrogenase and 6-P-gluconate dehydrogenase were assayed as

described by Sugimoto and Shiio, 1987a and Sugimoto and Shiio, 1987b by measuring the

increase in absorbance at 340 nm of NADPH with the following modification, the reaction

mixture contained 50 mM Tris-HCl buffer, pH 7.5. One unit of the enzyme activity was

defined as the amount catalyzing the formation of 1 nmol of NADPH per min.

2.2.4 Quantitative analysis of intracellular pyridine nucleotide pools

Pyridine nucleotides were extracted as in (Wimpenny and Firth, 1972). In brief,

NAD+ and NADP+ were extracted with HCl, while NADH and NADPH were extracted with

KOH from cells after 2 h (early response) or 24 h (later response) of 6 % headspace v/v

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oxygen exposure. Cells were grown on un-reduced mRCMfs as above. The concentrations

of pyridine nucleotides in extracts were measured by spectrophotometric enzyme assays as in

(Klingenberg, 1985).

2.2.5 Additional analytical methods

Growth was measured as optical density at 600 nm. Protein quantification in cell

extracts was performed using Bio-Rad’s Protein Assay Dye Reagent (Bio-Rad, Hercules,

CA, USA), according to manufacturer’s instructions. The amount of fructose present in the

culture media was determined by high-performance liquid chromatography as follows.

Liquid samples (600 µl) were centrifuged (10 min, 14,000 x g, room temperature).

Supernatant was filtered though a 0.22 µm syringe filter and analyzed using a Shimadzu

LC20 liquid chromatograph containing a HPX-87H column maintained at 65 °C. Sulfuric

acid (5 mM) was used as the eluent at a rate of 0.6 ml/min.

To determine cell density 11 ml of culture were sampled from growing cells. One ml

was used for O.D.600 reading while 10 ml of culture were filtered through a disposable pre-

weighed and pre-dried 22 µm syringe filter. Flow through was discarded. Filter and cells

were washed by passing 10 ml 50 mM phosphate buffer pH 7.0. Filter with cells was then

incubated at 80°C. These were weighed every 24 h until constant weight. The optical

densities were plotted against their corresponding dry cell weights yielding a linear

relationship between measured O.D.600 nm and culture densities (mg/L). The relationship

between optical density and dry cell weight was found to be 312 and 353 mg dry cells/L per

O.D.600 unit for C. ljungdahlii-WT and C. ljungdahlii-OTA1 respectively.

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2.3. Results

2.3.1 Effects of oxygen on C. ljungdahlii growth

In all of our experiments, cultures reached stationary phase by 48 h. C. ljungdahlii

strains were unexpectedly resistant to various concentrations of oxygen in the gas phase (Fig.

2-1 and Appendix Figs. A-1, A-3, A-5 and A-7). C. ljungdahlii-WT grew in reduced

medium containing up to 8% O2 with no apparent differences in cell density when compared

to control cultures (Fig. 2-1A). C. ljungdahlii-OTA1 also grew in reduced medium

containing up to 8% O2 in the gas phase. Nevertheless, growth was significantly reduced

(p<0.05) in these cells after 12 h of O2 exposure when compared to control cultures (Fig. 2-

1B). Cell growth was inhibited when either WT or OTA1 cultures were exposed to 12% O2

in reduced medium.

Reducing agents are regularly used to remove O2 traces from the media as well as to

lower the redox potential in the medium (Chalmers and Taylor-Robinson, 1979). Because of

that the oxygen exposure growth studies performed for cells grown with reducing agents in

their media cannot necessarily be relied upon to provide an accurate assessment of the

Clostridium cells resistance to oxygen exposure. Therefore, we removed both cysteine and

ammonium sulfide from the medium (un-reduced medium) and proceeded to expose C.

ljungdahlii to different O2 concentrations. Absence of reducing agents in the medium did not

have an effect in growth for either of the C. ljungdahlii species, since, as in control

experiments, cell numbers were similar in both reduced and unreduced media (Fig 2-1C and

D). C. ljungdahlii species were also capable of growing in unreduced media containing

relatively high amounts of O2 in the gas phase (Fig. 2-1 C, D, and Appendix Figs. A-3 and A-

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7). As in reduced media, C. ljungdahlii-WT showed no differences in growth when the

culture was exposed to O2 compared to control cultures (Fig. 2-1C). As expected, cultures

grown in un-reduced media showed less resistance to O2 than cells grown in reduced media.

Unlike C. ljungdahlii-WT, C. ljungdahlii-OTA1 growth was significantly reduced (p<0.05)

when cells were exposed to 6% O2 when compared to control cultures (Fig. 2-1 D). C.

ljungdahlii-WT and –OTA1 cultures failed to continue growth when exposed to 10% O2 and

8% O2 respectively, indicating that the OTA1 strain was more sensitive to O2 exposure than

the WT strain.

2.3.2 Effects of oxygen on ethanol production

Both in control as well as O2 exposed conditions, C. ljungdahlii –OTA1 produced

significantly higher (p < 0.05) concentrations of ethanol than C. ljungdahlii-WT (Fig. 2-2).

Ethanol production in C. ljungdahlii-WT reached its peak at 683 mg/L after 48 h of growth

in reduced medium (Fig. 2-2A). When these cells were exposed to 8% O2 in the gas phase,

the amount of ethanol produced was significantly higher (p < 0.05). Ethanol production

continued beyond 48 h with a production of 1.4 g/L at 72 h. This shows approximately

100% increase in ethanol production when compared to control cells. C. ljungdahlii-OTA1

strain growing in reduced control medium also reached maximum ethanol production at 48 h

with 1.2 g/L produced (Fig. 2-2B). It is important to note that in control experiments, OTA1

produced more than 75 % more ethanol than the WT strain (p < 0.05). As with WT, when C.

ljungdahlii-OTA1 cells were exposed to 8% O2, the amount of ethanol produced was

significantly higher (p < 0.05). Ethanol accumulation continued past 48 h with a production

76

of 2.0 g/L at 72 h; a 66 % increase compared to the 0% oxygen control experiments and 46%

higher when compared to C. ljungdahlii-WT exposed to 8% O2 (p < 0.05). Neither strain had

significant ethanol production when the headspace was supplemented with 12% O2 because

of poor growth (Fig 2-2A and B). Generally, ethanol production by both C. ljungdahlii-WT

and OTA1 strains ceased after 72 h (data not shown). For this reason experiments were

ended after 72 h of growth.

When grown in un-reduced medium, C. ljungdahlii strains produced less ethanol

when compared to cells grown in reduced media (Fig. 2-2C, D). These results were expected

as reducing agents have previously been shown to enhance ethanol production from syngas

(Atiyeh et al., 2009; Klasson et al., 1992c). Ethanol production in C. ljungdahlii-WT strain

reached 247 mg/L at 24 h and showed minimal increases in ethanol accumulation at later

time points (Fig. 2-2C). When these cells were exposed to 8% O2 in the gas phase, ethanol

production was maintained at increasing levels until 48 h. These cultures also had

significantly higher (p < 0.05) ethanol production (741 mg/L) than the 0% O2 control

experiment. A 200 % increase in ethanol production in C. ljungdahlii-WT control versus O2

treated cultures in un-reduced medium was observed. The C. ljungdahlii-WT strain was

unable to produce ethanol when the headspace was supplemented with 10% O2. In control

experiments, C. ljungdahlii-OTA1 strain continued ethanol production until reaching a

maximum (509 mg/L) after 48 h (Fig. 2-2D). When the OTA1 cultures were exposed to 6%

O2, ethanol production was increased by approximately 120% (1109 mg/L). When we

compare both strains exposed to 6% O2, C. ljungdahlii –OTA1 produced 50 % more ethanol

77

than the wild type strain. When exposed to 8% O2 ethanol production was negligible for the

OTA1 strain.

2.3.3 Effects of oxygen on acetate production

C. ljungdahlii, as do most acetogens, gains an ATP molecule when acetate is made

from acetyl-CoA (Russell and Martin, 2004). Therefore, because of this energetic benefit, it

produces considerably more acetate than ethanol. We see this in our experiments as C.

ljungdahlii strains produce as much as 22 times more acetate than ethanol (Fig. 2-2, 2-3).

When growing in reduced medium, C. ljungdahlii-WT strain, accumulated acetate until 48 h

of growth in the absence of O2 in the gas phase (Fig. 2-3A). When these cells were exposed

to 8% O2 the rate of acetate production decreased; however, the acetate concentration reached

similar amounts as in control experiments (Fig. 2-3A). C. ljungdahlii-OTA1 strain growing

in reduced medium also stopped acetate production by 48 h in control experiments (Fig. 2-

3B). As in the wild type strain, acetate formation rate decreased when C. ljungdahlii-OTA1

cells were exposed to 8% O2. Nevertheless, the final acetate concentration was comparable

to control experiments (Fig. 2-3B). Under both control and O2 exposed conditions, C.

ljungdahlii-WT produced significantly (p < 0.05) more acetate than C. ljungdahlii-OTA1,

33% and 45% higher amounts, respectively. Both strains cease acetate production when 12%

O2 was added in the gas phase because of poor culture growth.

In unreduced medium with no O2 in the headspace, C. ljungdahlii-WT produced the

highest observed acetate concentration of 7.3 g/L. Acetate production continued until 72 h

(Fig. 2-3C). Again, cells exposed to O2 showed a decrease in acetate production rate;

78

however, acetate concentration reached similar levels compared to control cells. C.

ljungdahlii -OTA1 growing in unreduced medium continued to produce acetate until 48 h.

When these cells were exposed to 6% O2, acetate continued to accumulate until 72 h of

growth. However, acetate in O2 exposed cells was 35% lower (p < 0.05) than in control cells

(Fig. 2-3D). In unreduced medium C. ljungdahlii-WT also produced significantly (p < 0.05)

more acetate than C. ljungdahlii-OTA1, 33% and 53% higher amounts in control and O2

exposure experiments, respectively. We observed that under all our conditions, regardless of

the medium, C .ljungdahlii-OTA1 produced significantly less acetate (p < 0.05) than the wild

type strain (Fig. 2-3). Acetate production in both wild type and OTA1 strains did not occur

when cells were exposed to 10% and 8% O2 respectively. Table 2-1 summarizes the effects

of O2 exposure on liquid products accumulation for the C. ljungdahlii-WT and –OTA1

strains.

2.3.4 Effects of oxygen on syngas utilization

Both C. ljungdahlii-WT and –OTA1 strains growing in reduced medium showed

continuous H2 and CO consumption until cultures reached stationary phase (Fig. 2-4A, 2-

5A). Consumption rate for these gases was 0.004 mmol/h and 0.015 mmol/h, respectively.

CO2 showed a net production throughout the length of the experiment at a rate of 0.025

mmol/h and 0.045 mmol/h for the WT and OTA1 strain, respectively. When cultures were

exposed to 8 % O2, O2 was completely reduced after 36 h of exposure (Fig. 2-4B, 2-5B).

Consumption rates for H2 and CO were not affected by O2 exposure. Nevertheless, the CO2

production rate increased (0.069 mmol/h and .061 mmol/h for C. ljungdahlii-WT and –

79

OTA1, respectively) when cultures were exposed to 8% O2 (Fig. 2-4B, 2-5B). Neither

consumption of CO and H2 nor production of CO2 were observed for cultures supplemented

with 12% O2, (Fig. 2-4C, 2-5C).

C. ljungdahlii-WT and OTA1 growing in unreduced medium also showed

continuous H2 and CO consumption until cultures reached stationary phase (Fig. 2-6A, 2-

7A). Consumption rates for H2 and CO were very similar for both strains (0.0072 and 0.0075

mmol/h for H2 and 0.018 and 0.019 mmol/h for CO for WT and OTA1, respectively. Again

CO2 was produced for the duration of the experiment and was accumulated at a rate of 0.018

mmol/h and 0.025 mmol/h in C. ljungdahlii-WT and OTA1cultures, respectively. When

cultures were exposed to 6% O2, O2 was completely reduced after 36 h of exposure (Fig. 2-

6B, 2-7B). Again, consumption rates for H2 and CO were not affected by O2 exposure; also

the CO2 production rate increased (0.031 mmol/h and .043 mmol/h for C. ljungdahlii-WT

and –OTA1 respectively) when cultures were exposed to 6% O2 (Fig. 2-6B, 2-7B). Neither

consumption nor production of gases was observed for cultures supplemented with 10 % O2,

(Fig. 2-6C, 2-7C).

2.3.5 Effects of oxygen on fructose utilization

The medium used for growth was supplemented with 5 g/L fructose as this carbon

source improves C. ljungdahlii growth, ethanol production and syngas utilization (see

chapter 3). Fructose utilization by C. ljungdahlii strains in control cultures was completed

by 48 h of growth (Fig. 2-8). A reduction of the fructose utilization rate was observed when

the cells were exposed to 8% and 6% O2 in reduced and unreduced medium, respectively.

80

Nevertheless, fructose was always completely utilized by 72 h. When C. ljungdahlii strains

growing in reduced media were exposed to 12% O2, fructose utilization did not appear to

occur. The same results were obtained when C. ljungdahlii-WT and –OTA1 strains growing

in unreduced media were exposed to 10% and 8% O2, respectively.

2.3.6 Enzyme activities associated with oxygen tolerance and ethanol production and

determination of cellular pyridine nucleotide pools

Crude cell extracts from both control and O2 exposed C. ljungdahlii strains had

superoxide dismutase, peroxidase, and oxidase activities. Catalase activity, which detoxifies

hydrogen peroxide, was not detected in any of the cell extracts (Table 2-2). C. ljungdahlii

WT strain had higher SOD activities than the OTA1 strain (24 and 9.5 U/mg for WT and

OTA1 respectively). SOD activity was not detected at early logarithmic phase in control

samples (14 h). However, activity was apparent at mid-logarithmic phase (36 h). SOD was

induced by O2 exposure as activity was detected in exposed cells at early logarithmic phase.

C. ljungdahlii strain crude extracts generally showed higher oxidase and peroxidase

activities (1.5 – 2.4 fold) when NADPH was used as electron donor (Table 2-2).

Nevertheless, this activity was not observed after 36 h of culture growth. Both oxidase and

peroxidase activities were detected early in the growth phase. Activities decreased after mid-

logarithmic growth phase and did not show induction after cells were exposed to O2. In

general, oxidase activity was higher in crude extracts from C. ljungdahlii strain OTA1 and

peroxidase higher in crude extracts from the WT strain.

81

The pentose phosphate pathway plays a crucial role for cells combating oxidative

stress. Specially one of its key enzymes, glucose-6-phosphate-dehydrogenase (G6PDH),

provides reducing equivalents in the form of NADPH for oxygen species reduction

(Lundberg et al., 1999; Pomposiello et al., 2001). Therefore, both NADPH producing

enzymes in the pathway, G6PDH and 6-phospho-gluconate dehydrogenase (6PGDH) were

assayed. Both enzymes were detected in crude extracts from both C. ljungdahlii species

exposed to 6 % O2 (Table 2-3). Activity was detected in extracts after 2 h (14 h of growth) of

O2 exposure but not after 24 h (36 h of growth) of O2 exposure. Importantly, activity was not

detected in cell extracts of control samples that had not been exposed to oxygen. The C.

ljungdahlii-WT strain showed higher G6PDH and 6PGDH activities than the OTA1 strain.

To try to understand why ethanol production was higher in O2 exposed cells,

ethanol production enzymes and pyridine nucleotide pools were measured. NAD+ dependent

aldehyde dehydrogenase (ALDH) activity was found in crude extracts from all samples

(Table 2-4). On the other hand, no activity was detected when NADP+ was provided as the e-

acceptor. ALDH activity was higher in crude extracts from control cells during early

logarithmic phase. However, activity was higher in crude extracts from O2 exposed cells

during mid-logarithmic phase. NAD+ dependent alcohol dehydrogenase (ADH) activity was

also observed in crude extracts from all samples assayed (Table 2-4). When NADP+ was

used as e- acceptor, activity was only detected in crude extracts from C. ljungdahlii strains

during early logarithmic phase. In addition, it was barely detected in samples from strain

OTA1 during mid-logarithmic growth phase. ADH activity was higher in C. ljungdahlii-

OTA1 crude extracts when compared to the WT strain. Once more, activity was higher in

82

extracts made from cells in early logarithmic phase. However, in O2 exposed cells, ADH

activity remained at steady levels after 24 h of O2 exposure, while it dropped more than half

in control cells.

The total pool size of NADH and NAD+ was approximately 2.5 fold higher in C.

ljungdahlii-WT strain control samples during early logarithmic growth phase compared to

the OTA1 strain (Table 2-5). Also, the total pool size of NADPH and NADP+ was higher in

the WT strain control samples at early logarithmic growth phase when compared to the

OTA1 strain (approximately 3 fold higher). In general, the total pool size of NAD(P)H and

NAD(P)+ for both strains decreased as cell numbers increased. In both strains, the

NADPH/NADP+ ratio increases upon exposure to O2 whereas NADH/NAD+ generally

decrease. This is more apparent in the OTA1 strain compared to WT (Table 2-5).

2.4. Discussion

We have shown that C. ljungdahlii strains are capable of tolerating and

enzymatically reducing O2 at concentrations up to 8% in the gas phase (Fig. 2-1). We also

demonstrate that exposure to O2 increases ethanol production as well as decreases acetate

formation (Fig. 2-2, 2-3 and Table 2-1). These results are not surprising as acetate producing

bacteria have been shown to tolerate oxygen in previous studies (Hardy and Hamilton, 1981).

Also, Drake and colleagues have described an acetogen, Clostridium glycolicum RD-1,

capable of tolerating up to 4% O2 in the gas phase in agitated cultures and showing increased

ethanol production at the expense of acetate in glucose-supplemented medium (Kusel et al.,

83

2001). However, we have reported that C. ljungdahlii can tolerate and reduce O2

concentrations of up to 8% in the headspace in agitated unreduced medium (see appendix)

and produces up to 10 fold more ethanol than was reported for C. glycolicum RD-1.

Furthermore, this study demonstrates that C. ljungdahlii is producing ethanol and acetate

while still consuming synthesis gas components (H2 and CO) (Fig. 2-4 and 2-5), presumably

by using the acetyl-CoA pathway, which is known to have enzymes that are readily

inactivated by O2 (Ragsdale and Wood, 1991b). Our results, therefore, suggest that C.

ljungdahlii species have effective enzymes to protect its metabolism from limited oxidative

stress.

Numerous anaerobes posses an array of oxidative stress protection enzymes. These

range from the classic enzymes such as SOD, catalase and NADH oxidase, to more recently

described, unusual enzymes such as SOR, thioredoxin reductase, and rubrerythrin among

others (Brioukhanov et al., 2002; Das et al., 2001; Jenney et al., 1999). In fact, the C.

ljungdahlii genome contains genes coding for at least six oxidative stress protection enzymes

(Köpke et al., 2010). These include SOD, catalase, NADPH flavin oxidoreductase, NADH

oxidase, rubredoxin and rubrerythrin. Cell extracts of C. ljungdahlii showed activities for

SOD, oxidase and peroxidase (using either NADH or NADPH as an e- donor), but not

catalase (Table 2-2). The C. ljungdahlii genome does not contain genes coding for NADPH

oxidase nor peroxidase; nevertheless, NADPH flavin oxidoreductase can show activity

towards O2 (reducing it to H2O2) (Bruchhaus et al., 1998). Also rubrerythrin has peroxidase

activity using NADPH as an e- donor (Coulter et al., 1999). These enzymes (with the

84

exception of SOD), appear to be constitutively expressed as we could detect relatively high

activities in control samples of both C. ljungdahlii strains.

Classic oxidative stress regulator systems such as SoxRS or PerR are not evident in

the genome of this organism. Nevertheless, we saw an increase in NADPH/NADP+ ratio in

C. ljungdahlii-WT strain as a response to O2 exposure (Table 2-5). In addition, oxidase and

peroxidase activities on both WT and OTA1 strains are higher when NADPH is used as e-

donor. NADPH plays a role in biosynthetic anabolism reactions and is required for many

detoxification reactions in bacteria; furthermore, it is produced mainly by the pentose

phosphate pathway (PPP) in response to oxidative stress (Briolat and Reysset, 2002; Kabir

and Shimizu, 2006). The C. ljungdahlii genome has genes that encode for the main enzymes

involved in the PPP. We show that both glucose-6-phosphate dehydrogenase and 6-

phosphogluconate dehydrogenase (responsible for producing NADPH) activities in crude

extracts are up-regulated after exposing the cells to O2 (Table 2-3). These results show

evidence for some level of regulation of oxidative stress response. On the other hand, we do

not observe an increase in NADPH/NADP+ ratio in strain OTA1. This suggests that we may

have missed this increase (happened between O2 exposure and 2 h sampling time) or that

NADPH consumption increased due to O2 exposure. There is evidence that demonstrates

NADPH turnover rate (consumption) can increase as a response to oxidative stress (Emerson

et al., 2003). Higher SOD activity in crude extracts from C. ljungdahlii-WT compared to the

OTA1 strain might explain the higher tolerance to O2 shown by the former versus the latter.

A proposed model for C. ljungdahlii oxidative stress detoxification system is shown in figure

2.9. It has been shown that addition of reducing agents to the culture medium alters e-

85

flow to increase the formation of NAD(P)H which in turn directs carbon and acid to alcohol

production (Dürre et al., 1995; Klasson et al., 1991a). We confirm this theory in our

experiments as ethanol production in C. ljungdahlii-WT and –OTA1 species is

approximately 2.5 fold higher when cultures are grown in reduced medium (Table 2-1).

Exposure of C. ljungdahlii to O2 appears to have similar effects on ethanol formation as with

addition of reducing agents. O2 exposed cultures produced ethanol concentrations

comparable to cultures in reduced medium and more than 2-fold higher than the control

unreduced cultures (Table 2-1). In addition, ethanol forming enzymes remain at relatively

higher levels after 24 h of O2 exposure compared to control cultures (Table 2-4).

Furthermore, it appears that the oxygen and reducing agents effect is additive as cells

growing in reduced media and exposed to O2 produced the most ethanol.

NAD(H) and NADP(H) levels are strongly regulated in cells by the NAD kinase

(encoded in C. ljungdahlii genome) and these levels will fluctuate depending on the growth

condition of the cell (Grose et al., 2006). For example, NADH is toxic to the cell during

oxidative stress as it contributes electrons for iron reduction, thereby facilitating the hydroxyl

radical generating Fenton reaction (Imlay and Linn, 1988; Woodmansee and Imlay, 2002).

This triggers NAD+ destruction/consumption followed by de novo synthesis. However,

NADP(H) has not been shown to enhance the Fenton reaction (Grose et al., 2006; Hashida et

al., 2010). Upon restoration of anaerobic conditions, the cell could have excess NADPH and

reducing power that were accumulated in response to the oxidative stress conditions, (Kuřec

et al., 2009; Wimpenny and Firth, 1972) which then must be balanced. We see this as

NAD(P)H concentrations decline at 24 h after O2 exposure (Table 2-5). Under conditions in

86

which C. ljungdahlii has high levels of NAD(P)H, it might reduce acetate to aldehyde and

then ethanol by the action of aldehyde oxidoreductase (AOR) enzymes encoded in the

genome. This could also assist the cell in energy conservation by forming ATP by acetate

production then acetaldehyde and ethanol (Köpke et al., 2010). This might explain an

increase of ethanol formation in cultures exposed to oxidative stress.

We have isolated a mutant strain (OTA1) that produces approximately 2-fold more

ethanol than the WT strain, most likely due a higher ethanol dehydrogenase activity (Table 2-

4). We have also shown that is possible to increase ethanol production along with decreasing

acetate formation by exposing C. ljungdahlii to O2. The fact that C. ljungdahlii can tolerate

high O2 concentrations and simultaneously produce higher ethanol concentration, is

significant for its use in the biofuels industry. It is likely that in a syngas fermentation

facility these cells will encounter O2 contamination in the gas stream, either directly from the

syngas or O2 intrusion along the syngas collection/stream process (Datar et al., 2004; Zainal

et al., 2002). Not only would these cells survive this O2 intrusion, the process would benefit

from it, lowering the cost by eliminating the use of reducing agents and stringent anaerobic

environment.

87

0 12 24 36 48 60 720

500

1000

15000% O2

8% O2

12% O2

Culture density (mg/L)

Time (h)

0 20 40 60 800

500

1000

15000% O2

8% O2

12% O2

Time (h)

Culture Density (mg/L)

0 20 40 60 800

500

1000

15000% O2

6% O2

10% O2

Time (h)

Culture Density (mg/L)

0 20 40 60 800

500

1000

15000% O2

6% O2

8% O2

Time (h)

Culture Density (mg/L)

A

DC

B

Figure 2-1. C. ljungdahlii cell growth. A) WT reduced, B) OTA1 reduced, C) WT unreduced, D) OTA1 unreduced.

88

0 12 24 36 48 60 720

500

1000

1500

2000

25000% O2

8% O2

12% O2

Time (h)

Ethanol (mg/L)

0 12 24 36 48 60 720

500

1000

1500

2000

25000% O2

8% O2

12% O2

Time (h)

Ethanol (mg/L)

0 12 24 36 48 60 720

500

1000

1500

2000

25000% O2

6% O2

10% O2

Time (h)

Ethanol (mg/L)

0 12 24 36 48 60 720

500

1000

1500

2000

25000% O2

6% O2

8% O2

Time (h)

Ethanol (mg/L)

C D

BA

Figure 2-2. C. ljungdahlii ethanol production. A) WT reduced, B) OTA1 reduced, C) WT unreduced, D) OTA1 unreduced.

89

0 12 24 36 48 60 720

2000

4000

6000

8000

100000% O2

8% O2

12% O2

Time (h)

Acetate (mg/L)

0 12 24 36 48 60 720

2000

4000

6000

8000

100000% O2

8% O2

12% O2

Time (h)

Acetate (mg/L)

0 12 24 36 48 60 720

2000

4000

6000

8000

100000% O2

6% O2

10% O2

Time (h)

Acetate (mg/L)

0 12 24 36 48 60 720

2000

4000

6000

8000

100000% O2

6% O2

8% O2

Time (h)

Acetate (mg/L)

C D

BA

Figure 2-3. C. ljungdahlii acetate production. A) WT reduced, B) OTA1 reduced, C) WT unreduced, D) OTA1 unreduced.

90

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

A B

C

Figure 2-4. C. ljungdahlii WT syngas utilization when grown in reduced medium; A) 0% O2, B) 8% O2, C) 12% O2

91

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 20 40 60 800

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

A B

C

Figure 2-5. C. ljungdahlii OTA1 syngas utilization when grown in reduced medium; A) 0% O2, B) 8% O2, C) 12% O2

92

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

A

C

B

Figure 2-6. C. ljungdahlii WT syngas utilization when grown in unreduced medium; A) 0% O2, B) 6% O2, C) 10% O2

93

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 20 40 60 800

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

A

C

B

Figure 2-7. C. ljungdahlii OTA1 syngas utilization when grown in unreduced medium; A) 0% O2, B) 6% O2, C) 8% O2

94

0 12 24 36 48 60 720

1

2

3

4

5

6

70% O2

8% O2

12% O2

Time (h)

Fructose (g/L)

0 12 24 36 48 60 720

1

2

3

4

5

6

70% O2

8% O2

12% O2

Time (h)

Fructose (g/L)

0 12 24 36 48 60 720

1

2

3

4

5

6

70% O2

6% O2

10% O2

Time (h)

Fructose (g/L)

0 12 24 36 48 60 720

1

2

3

4

5

6

70% O2

6% O2

8% O2

Time (h)

Fructose (g/L)

D

B

C

A

Figure 2-8. C. ljungdahlii fructose utilization. A) WT reduced, B) OTA1 reduced, C) WT unreduced, D) OTA1 unreduced.

95

Figure 2-9. Model for the C. ljungdahlii oxidative stress detoxification system. SOD (superoxide dismutase), Cat (catalase), Rr (rubrerythrin), Trx (thioredoxin).

96

Table 2-1. Effect of O2 on product profiles in C. ljungdahlii strains at maximum production

Strain Medium Condition (% O2) EtOH (g/L) Ac (g/L) EtOH/Aca

ratio

Total liq. Prod. (g/L)

C. ljung.-WT

Reduced 0% 0.68 4.9 0.14 5.7

8% 1.4 4.5 0.30 5.9

Unreduced 0% 0.29 7.2 0.05 7.6

6% 0.81 6.7 0.17 7.5

C. ljung.-OTA1

Reduced 0% 1.3 3.3 0.40 4.6

8% 2.0 2.5 0.80 4.5

Unreduced 0% 0.54 4.9 0.11 5.4

6% 1.1 3.2 0.35 4.3 aEtOH is ethanol, Ac is acetate

97

Table 2-2. Oxidative stress enzyme activities in C. ljungdahlii WT and OTA1

aN.D. is not detected

98

Table 2-3. Pentose Phosphate Pathway enzyme activities in C. ljungdahlii WT and OTA1

Strain Condition Time (h)

Enzyme Activity (mU/mg of protein)

Glucose-6-P DH

6-P-gluconate DH

C. ljungdahlii-WT Control 14 N.D.a N.D.

36 N.D. N.D. O2 exposed 14 0.70 ± 0.14 0.76 ± 0.19

36 N.D. N.D. C. ljungdahlii-OTA1 Control 14 N.D. N.D.

36 N.D. N.D. O2 exposed 14 0.30 ± 0.25 0.50 ± 0.21

36 N.D. N.D. aN.D. is not detected.

99

Table 2-4. Ethanol forming enzyme activities in C. ljungdahlii WT and OTA1

Strain Condition Time (h)

Enzyme Activities (mU/mg of protein)

ALDH ALDH ADH ADH

(NAD+) (NADP+) (NAD+) (NADP+) C. ljungdahlii-WT Control 14 102 ± 29 N.D.a 89 ± 32 67 ± 5

36 40 ± 0.3 N.D. 18 ± 3 N.D. O2 exposed 14 25 ± 11 N.D. 82 ± 16 73 ± 3

36 58 ± 2 N.D. 29 ± 5 N.D. C. ljungdahlii-OTA1 Control 14 98 ± 46 N.D. 131 ± 40 1 ± 1

36 23 ± 2 N.D. 58 ± 1 N.D. O2 exposed 14 27 ± 1 N.D. 87 ± 28 1 ± 1.5

36 64 ± 8 N.D. 86 ± 5 N.D. aN.D. is not detected.

100

Table 2-5. Intracellular pyridine nucleotide pools of C. ljungdahlii WT and OTA1

101

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CHAPTER 3

Influence of carbon source pre-adaptation on Clostridium ljungdahlii growth and product formation*

Oscar Tirado-Acevedo1†, Jacqueline L. Cotter2†, Mari S. Chinn2 and Amy M. Grunden1

1-Department of Microbiology, North Carolina State University, 4548 Thomas Hall, Campus Box 7615, Raleigh, NC 27606, USA 2-Department of Biological and

Agricultural Engineering, North Carolina State University, 277 Weaver Labs, Campus Box 7625, Raleigh, NC 27695-7625, USA. † O.T.A. and J.L.C. contributed equally to this

work.

*Submitted for publication to: Bioresource Technology

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ABSTRACT

Clostridium ljungdahlii was grown on different carbon sources including syngas only,

syngas-fructose and fructose only to identify ideal pre-adaptation conditions for ethanol and

acetate production from subsequent cultures grown in reactors containing syngas only or

fructose-syngas substrates. In syngas only reactors, cultures pre-adapted to fructose had

faster growth rates and higher ethanol and acetate formation than cells pre-grown on syngas

or syngas-fructose. In syngas-fructose reactors, cultures did not show significant growth or

acetate production differences under pre-adaptation treatments. Nevertheless, in these

reactors, syngas and syngas-fructose pre-adapted cultures showed higher ethanol production

than fructose pre-adapted cultures. Among the pre-adaptation carbon sources tested, fructose

showed better results in syngas only reactors. However, the presence of syngas in the pre-

adaptation cultures proved to be the better method overall for ethanol production.

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3.1. Introduction

Conversion of synthesis gas to liquid fuels by biological catalysts has been suggested as a

promising technology to achieve oil independence (Zeikus, 1980). In this method, cellulosic

biomass and other carbon sources (coal, industry and/or municipal waste, etc.) are converted

to synthesis gas (syngas) by a well known process called gasification (Bridgwater, 1995).

This syngas is then fed to a microorganism which can utilize it as a carbon and energy

source. The by-product of this fermentation could be ethanol, butanol or other commodity

chemicals. There have been a number of microorganisms isolated for use in this process. In

addition, some companies are en route to commercializing their methods (Tirado-Acevedo et

al., 2010).

Extensive research has been focused on optimizing bioreactor designs and media

development (Hurst and Lewis, 2010; Phillips et al., 1993b; Worden et al., 1991). Recent

bioreactor processes have made use of micro-bubble gas dispersers, cell recycling, and

addition or removal of reducing agents and yeast extract, respectively. However, only a few

studies have examined the effects that seed cultures pre-adapted to different carbon sources

and carbon source in the production culture media have on product formation and cell yield

(Klasson et al., 1992c).

Clostridium ljungdahlii, which has been studied in regard to synthesis gas fermentation

(Phillips et al., 1993b), was isolated from chicken yard waste and was the first

microorganism described to be able to ferment syngas components to ethanol via the acetyl-

CoA pathway (Tanner et al., 1993b). In addition to utilizing CO and H2, C. ljungdahlii can

use sugars such as fructose as a carbon source. The purpose of this study was to compare

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cell growth and product formation from C. ljungdahlii grown on different carbon sources

(syngas and sugar combinations) while using cells pre-adapted to sugar, syngas and mixed

sugar-syngas substrates in a continuous gas feed, batch liquid system.

3.2. Methods

3.2.1. Organism and medium preparation

C. ljungdahlii (ATCC 55353) was obtained from the American Type Culture Collection.

The bacterium was grown at 37 ºC on a modified Reinforced Clostridial Basal medium with

additional salts, vitamins, and trace elements adopted from the ATCC 1754 PETC medium

(RCM.NA.SVE) in a nitrogen atmosphere unless otherwise indicated (Cotter et al., 2009a).

Cultures provided syngas (7.5 ml/min ) were grown in a fermentation reactor (Fig.3.1)

(Cotter et al., 2009b). The bottled syngas used was an artificial mix of 50% N2, 20% CO,

20% CO2, and 10% H2. When added to medium in combination with syngas, fructose was

used at 2.5 g/L. All culture transfers were done at 5 % inoculum (v/v).

2.2. Inoculum preparation using different carbon sources

C. ljungdahlii was grown on RCM.NA.SVE (which from here on will be referred to

simply as medium). Cultures were initiated from a freezer stock (-80°C) and incubated at

37°C for 72 hours before 3 serial transfers every 24 hours to achieve steady growth in

medium containing fructose (5 g/L). Active cells (472 mg dry cells/ml) were transferred to

different carbon sources [medium-syngas, medium-syngas-fructose (2.5 g/L) or medium-

fructose (5g/L)] to start the seed cultures for the fermentation studies. Fermentation reactors

containing medium-syngas and medium-syngas-fructose at a working volume of 250 ml were

inoculated with the three different seed cultures (472 mg dry cells/ml; 5% v/v). Each

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medium seed culture combination was grown in triplicate. Cell growth was monitored and

liquid and gas samples were taken for analysis every 6-8 hours.

3.2.3. Substrate and end product analysis

Headspace gases were analyzed by gas chromatography on a Carbosieve S-II, 100/120

mesh stainless steel column (10’ x 1/8”: 3.05 m x 0.3175 cm) using a thermal conductivity

detector (Shimadzu GC-17A), Ethanol and acetate concentrations were determined in

acidified samples by gas chromatography (Shimadzu GC-17A, Kyoto, Japan). The

chromatography column was packed with Supelco SP 1000 (1% H3 PO4, 100/120 mesh) and

analytes were quantified using a flame ionization detector (Cotter et al., 2009a; Cotter et al.,

2009b).

The amount of fructose present in the experimental cultures was determined by high-

performance liquid chromatography (Shimadzu LC20) as follows: liquid samples (600 µL)

were centrifuged (10 min, 18,400 x g, room temperature). Supernatant was filtered though a

0.22 µm syringe filter into a crimp vial and analyzed on a HPX-87H column (65°C) using a

refractive index detector. Sulfuric acid (5 mM) was used as the eluent at a rate of 0.6

ml/min.

3.2.4 Statistical analysis

Cell growth and liquid product analysis from the different inoculum sources were

evaluated using the General Linear Model (GLM) in SAS® Version 9.1 (SAS Inc., Cary, NC,

USA). Assessment of statistical significance for pre-adaptation on different carbon sources

and different production growth substrates was set at P < 0.05.

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3.3. Results and discussion

3.3.1. Growth of C. ljungdahlii and product formation in syngas reactors

C. ljungdahlii cultures were pre-adapted on three different carbon source combinations

(syngas, syngas-fructose or fructose). Subsequently, these cells were transferred to

fermentation reactors with either syngas or syngas-fructose medium. In syngas only reactors,

cells pre-adapted on fructose produced the highest cell densities reaching maximum growth

after 24 hours (Fig. 3-2A). This observation is interesting since one would expect a lag-time

given that these cells would have to switch from fructose fermentation to syngas

fermentation. Syngas and syngas-fructose pre-adapted cells reached maximum densities

after 40 and 56 hours, respectively. Cells pre-adapted on syngas-fructose showed the longest

lag-time, contributing to the increased time to reach peak growth, where fructose only

cultures had a growth rate of 10.4 mg/ml/h, while syngas and syngas-fructose cultures both

had growth rates of 5 mg/ml/h. Interestingly, after 56 hours cell density was statistically the

same independent of pre-adaptation medium, possibly due to product inhibition or depletion

of an essential nutrient.

Ethanol production in syngas reactors was significantly higher (P < 0.05) when cells

where pre-adapted on fructose, reaching a maximum of 2.2 mM after 64 hours of growth

(Fig. 3-3A). Ethanol production in these cells seems to be non-growth associated since these

cells reached stationary phase after 24 hours. Cells pre-adapted on syngas also reached

maximum ethanol yield after 64 hours, while cells pre-adapted on syngas-fructose reached a

maximum after 48 hours. Although acetate is considered to be a growth associated product,

it was continuously accumulated during the growth phase as well as after the stationary phase

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was reached (Fig. 3-3C). Cells pre-adapted on fructose produced significantly higher acetate

concentrations (P < 0.05) with a maximum of 34.3 mM. Again, in syngas reactors, cells pre-

adapted to syngas-fructose show a lag time in acetate production when compared to fructose

or syngas pre-adapted cells but reached comparable concentrations by the end of the

experiment.

3.3.2. Growth and product formation in syngas-fructose reactors

No significant growth differences were observed among the three inoculum sources in the

syngas-fructose reactors (Fig. 3-2B). These cultures did not show a lag-time and reached a

maximum cell density after 24 hours. Cell growth rates were approximately 43 mg/L/h for

syngas and fructose pre-adapted cultures and 37 mg/L/h for syngas-fructose pre-adapted

cultures. Independent of inoculum source, cells growing in syngas-fructose reactors had

significantly higher cell yields (P < 0.05) as compared to cells growing in syngas reactors

(787 mg/L vs. 332 mg/L).

Ethanol concentration in syngas-fructose reactors was statistically lower compared to

cells that were pre-adapted on fructose (Fig 3-3B). Pre-adaptation had no effect on acetate

production as all cultures produced statistically the same amount of acetate. Independent of

pre-adaptation, both ethanol and acetate production seem to be growth associated as

production slows down when cells reach stationary phase. It had been previously suggested

that ethanol production is associated with non-growth conditions (Klasson et al., 1992c). We

observe this in syngas only medium but not when fructose is present (Fig. 3-3A and B).

Independent of the inoculum source, cells grown in syngas-fructose reactors produced

significantly more ethanol and acetate than cells grown in syngas reactors (Fig. 3-3), with as

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much as 8- and 1.5-fold more ethanol and acetate, respectively. This trend has also been

observed with addition of 5 g/L of fructose to syngas-grown cultures, where increases up to

25% and 35 % for ethanol and acetate concentration respectively were seen compared to

syngas alone (unpublished data). This may indicate that an addition of a small concentration

of sugar in the medium could improve C. ljungdahlii ethanol yields. A number of low-cost

sugar sources have been identified that could act as a carbon supplement for this process

without interfering with food supply demands (Jiang et al., 2009; Lawford and Rousseau,

1997). The use of low concentration sugar supplementation may be worth examining as a

method to improve commodity chemical production from C. ljungdahlii.

3.3.3. Gas and fructose analysis

Headspace gas composition was monitored in both syngas and syngas-fructose

fermentations. Figure 3-4A illustrates a representation of headspace gases exhausting from

the reactor. CO and H2 consumption at 8 to 24 hours coincides with a CO2 increase. This

indicates that C. ljungdahlii is most likely using the acetyl-CoA pathway for the metabolism

of CO and H2, thereby providing the cell with carbon for biomass and energy production as

well as reducing equivalents (Ragsdale and Wood, 1991a). Nevertheless, by the end of the

culture growth, the head-space gases return to near starting concentrations. This observation

may be related to several conditions, including cells that are no longer consuming gases for

growth and metabolism or that the gas flow rate exceeds the cells’ rate of consumption.

More complete utilization of the syngas supplied could be achieved by changing the reactor

design to include gas recirculation (Nie et al., 2008).

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Fructose utilization was analyzed throughout the experiment for the syngas-fructose

reactors. Fructose was consumed at equal rates independent of inoculum source and was

exhausted by 24 hours (Fig. 3-4B). Fructose utilization and liquid product synthesis appear

to be tightly associated as disappearance of fructose in the medium correlates with reduction

of ethanol and acetate production. Nevertheless, after fructose is completely consumed,

ethanol production rate is still higher on syngas-fructose reactors when compared to syngas

reactors. This indicates that fructose is most likely providing the cell with an excess of

reducing equivalents , which in turn could be utilized to reduce acetate to ethanol (Kopke et

al., 2010).

3.4. Conclusions

In syngas reactors, inoculum source was found to have an effect on growth rate and final

cell concentration as well as ethanol formation. In these reactors, the highest cell

concentration and ethanol production was found in reactors with cells pre-adapted with

fructose. These reactors produced around 2.5 and 1.5 times more ethanol than reactors

inoculated with syngas and syngas-fructose pre-adapted cells, respectively. Fructose pre-

adaptation likely provides higher levels of energy equivalents (ATP/GTP) and reducing

power (NADH/NADPH) to the cells, which enables the cells to efficiently utilize a carbon

source (syngas) that initially requires an energy and reductant input (Kopke et al., 2010).

In syngas-fructose reactors, inoculum source did not appear to have an impact on cell

density, liquid product yield, nor fructose utilization. This suggests that independent of pre-

adaptation, in syngas-fructose reactors, cells have the capacity to readily utilize fructose as

carbon and electron source. Overall, syngas-fructose reactors had approximately 2.5 times

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higher cell densities and 8 and 1.5 times higher ethanol and acetate, respectively than syngas

reactors. This indicates that a small amount of fructose may be needed to increase ethanol

yields in our system as excess reducing equivalents translate to better ethanol yields.

However, fructose is an expensive carbon source and is also part of the human food supply,

but there are cheap alternative carbon sources that could be suitable for C. ljungdahlii culture

(Gullón et al., 2008; Jiang et al., 2009).

Syngas utilization in the reactors could not be conclusively established possibly due to

the gas flow rate being greater than an accurately measureable cell gas utilization rate. If

quantification of gas consumption in the continuous gas feed system is desired, changes in

reactor design and gaseous substrate flow patterns would be needed. Therefore, it was

concluded that for syngas-fructose reactors, pre-adaptation is not necessary as the overall

fermentation was not affected. Nevertheless, in syngas only reactors a fructose pre-

adaptation will be beneficial as it increases cell numbers and liquid products.

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Figure 3-1. Synthesis gas fermentation reactor.

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Figure 3-2. C. ljungdahlii culture density in the syngas reactor (A) and syngas-fructose reactor (B) inoculated with cells pre-grown on fructose (circles), syngas-fructose (squares), or syngas (triangles).

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Figure 3-3. C. ljungdahlii production of ethanol (A and B) and acetate (C and D) in the syngas reactor (A and C) and in the syngas-fructose reactor (B and D) inoculated with cells pre-grown on fructose (circles), syngas-fructose (squares), or syngas (triangles).

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Figure 3-4. C. ljungdahlii fructose utilization (A) in the syngas-fructose reactor inoculated with cells pre-grown on fructose (circles), syngas-fructose (squares), or syngas (triangles) and syngas utilization (B) in the syngas reactor inoculated with cells pre-grown on syngas.

121

REFERE-CES

Bridgwater, A. V., 1995. The technical and economic feasibility of biomass gasification for power generation. Fuel. 74, 631-653. Cotter, J. L., Chinn, M. S., and Grunden, A. M., 2009a. Ethanol and acetate production by Clostridium ljungdahlii and Clostridium autoethanogenum using resting cells. Bioprocess. Biosyst. Eng. 32, 369-380. Cotter, J. L., Chinn, M. S., and Grunden, A. M., 2009b. Influence of process parameters on growth of Clostridium ljungdahlii and Clostridium autoethanogenum on synthesis gas. Enzyme. Microb. Technol. 44, 281-288. Gullón, B., Yáñez, R., Alonso, J. L., and Parajó, J. C., 2008. l-Lactic acid production from apple pomace by sequential hydrolysis and fermentation. Bioresour. Technol. 99, 308-319. Hurst, K. M., and Lewis, R. S., 2010. Carbon monoxide partial pressure effects on the metabolic process of syngas fermentation. Biochem. Eng. J. 48, 159-165. Jiang, L., Wang, J., Liang, S., Wang, X., Cen, P., and Xu, Z., 2009. Butyric acid fermentation in a fibrous bed bioreactor with immobilized Clostridium tyrobutyricum from cane molasses. Bioresour. Technol. 100, 3403-3409. Klasson, K. T., Ackerson, M. D., Clausen, E. C., and Gaddy, J. L., 1992. Bioconversion of synthesis gas into liquid or gaseous fuels. Enzyme. Microb. Technol. 14, 602-608. Kopke, M., Held, C., Hujer, S., Liesegang, H., Wiezer, A., Wollherr, A., Ehrenreich, A., Liebl, W., Gottschalk, G., and Durre, P., 2010. Clostridium ljungdahlii represents a microbial production platform based on syngas. P.N.A.S. 107, 13087-13092. Lawford, H., and Rousseau, J., 1997. Corn steep liquor as a cost-effective nutrition adjunct in high-performance Zymomonas ethanol fermentations. Appl. Biochem. Biotechnol. 63-65, 287-304. Nie, Y., Liu, H., Du, G., and Chen, J., 2008. Acetate yield increased by gas circulation and fed-batch fermentation in a novel syntrophic acetogenesis and homoacetogenesis coupling system. Bioresour. Technol. 99, 2989-2995. Phillips, J., Klasson, K., Clausen, E., and Gaddy, J., 1993. Biological production of ethanol from coal synthesis gas. Appl. Biochem. Biotechnol. 39-40, 559-571.

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Ragsdale, S. W., and Wood, H. G., 1991. Enzymology of the Acetyl-CoA Pathway of CO2 Fixation. Crit. Rev. Biochem. Mol. Biol. 26, 261-300. Tanner, R. S., Miller, L. M., and Yang, D., 1993. Clostridium ljungdahlii sp. nov., an Acetogenic Species in Clostridial rRNA Homology Group I. Int. J. Syst. Bacteriol. 43, 232-236. Tirado-Acevedo, O., Chinn, M. S., and Grunden, A. M., Production of Biofuels from Synthesis Gas Using Microbial Catalysts. In: I. L. Allen, S. Sima, M. G. Geoffrey, Eds.), Advances in Applied Microbiology, Vol. Volume 70. Academic Press, 2010, pp. 57-92. Worden, R. M., Grethlein, A. J., Jain, M. K., and Datta, R., 1991. Production of butanol and ethanol from synthesis gas via fermentation. Fuel. 70, 615-619. Zeikus, J. G., 1980. Chemical and Fuel Production by Anaerobic Bacteria. Annu. Rev. Microbiol. 34, 423-464.

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CHAPTER 4

STUDY CO-CLUSIO-S

Fossil fuels are harmful to the environment and human health, and the fossil reserves

are dwindling at a dramatic rate. In addition, today’s biofuels sources such as corn and sugar

cane are expensive, depend on government subsidies and can increase food prices as less of

these crops are made available for human consumption as a result of their use for fuel

production. Biofuels from cellulosic biomass have been identified as one suitable

alternative to fossil fuels and first generation biofuel corn ethanol (hydrolysis-fermentation).

Synthesis gas has been used as fuel for decades due to being affordable and can be obtained

from any type of biomass regardless of its composition. Being a gas, its use as a

transportation fuel is not ideal. Converting it to liquid would condense its energy and make it

easier to store and to transport.

Fermentation of synthesis gas obtained from biomass has proven to be a viable

approach to produce biofuels. This technology has higher product yields and lower energy

input than lignocellulose hydrolysis-fermentation. Researchers have isolated several

microorganisms capable of converting synthesis gas to more suitable, higher energy fuels.

These biocatalysts can manage most synthesis gas contaminants making them more robust

than some chemical catalysts. Nevertheless, contaminants such as NO, acetylene and O2 can

inhibit activities from these microbes. The industry and the government have been very

interested in this process and a commercial demonstration plant is already in operation

(Coskata Inc. Madison, Pennsylvania).

124

The syngas-fermenting bacterium C. ljungdahlii has been the subject of a number of

investigations to improve its ethanol yields. Nevertheless, extensive physiological and

metabolic research is limited for this organism. Several acetogens have been shown to

produce higher ethanol concentration as a response to O2 exposure. This opens up new

possibilities for improvement of biofuels production.

During our research to increase ethanol production by C. ljungdahlii, one of our aims

was to demonstrate that C. ljungdahlii could increase ethanol formation from synthesis gas as

a response to O2 exposure. We have also isolated a mutant strain (OTA1). This strain

produces approximately 2-fold more ethanol than the WT strain. Both strains increase

ethanol production along with decreasing acetate formation when exposed to O2. This is

significant for C. ljungdahlii’s use in the biofuels industry as O2 contamination is likely to

occur in the synthesis gas.

We also demonstrate that inoculum source for syngas reactors has an effect on growth

rate and final cell concentration as well as ethanol formation. Fructose pre-adaptation

improves cell concentration and ethanol production, most likely providing higher levels of

energy and reducing power to the cells. Fructose is an expensive carbon source; however, an

alternative inexpensive sugar may be utilized to enhance ethanol production instead.

Given the lack of genetic tools available for C. ljungdahlii, we have had to rely on other

methods to increase ethanol production from synthesis gas. We were successful in doing this

by exposing the cells to O2 and pre-adapting the cells to fructose. The fact that this organism

increases ethanol formation at the expense of acetate in the presence of O2 is beneficial in an

125

industrial production setting since it will lower the cost by eliminating the use of reducing

agents and the requirements for stringent anaerobic conditions.

126

APPE-DIX

This appendix section contains additional figures from C. ljungdaglii oxygen

exposure study presented in chapter 2. These include growth, ethanol and acetate formation,

headspace analysis and fructose utilization.

127

0 12 24 36 48 60 720

500

1000

1500

2000

0% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Culture Density (mg/L)

0 12 24 36 48 60 720

1

2

3

4

5

6

70% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Fructose (g/L)

0 12 24 36 48 60 720

500

1000

1500

2000

25000% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Ethanol (mg/L)

0 12 24 36 48 60 720

2000

4000

6000

8000 0% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Acetate (mg/L)

A

DC

B

Figure A-1. C. ljungdahlii-WT metabolism in reduced medium. A) Growth, B) fructose utilization, C) ethanol production, D) acetate production.

128

Figure A-2. C. ljungdahlii-WT growing in reduced medium, headspace analysis at different O2 concentrations. A) 0 % O2 B) 4 % O2 C) 6 % O2 D) 8 % O2 E) 12 % O2

129

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

A B

C D

130

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

E

131

0 12 24 36 48 60 720

500

1000

15000% O2

4% O2

6% O2

8% O2

10% O2

Time (h)

Culture Density (mg/L)

0 12 24 36 48 60 720

1

2

3

4

5

6

70% O2

4% O2

6% O2

8% O2

10% O2

Time (h)

Fructose (g/L)

0 12 24 36 48 60 720

500

1000

1500

2000

25000% O2

4% O2

6% O2

8% O2

10% O2

Time (h)

Ethanol (mg/L)

0 12 24 36 48 60 720

2000

4000

6000

8000 0% O2

4% O2

6% O2

8% O2

10% O2

Time (h)

Acetate (mg/L)

A

DC

B

Figure A-3. C. ljungdahlii-WT metabolism in unreduced medium. A) Growth, B) fructose utilization, C) ethanol production, D) acetate production

132

Figure A-4. C. ljungdahlii-WT growing in unreduced medium, headspace analysis at different O2 concentrations. A) 0 % O2 B) 4 % O2 C) 6 % O2 D) 8 % O2 E) 10 % O2

133

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

C D

A B

134

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

E

135

0 12 24 36 48 60 720

2000

4000

6000

8000

0% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Acetate (mg/L)

0 12 24 36 48 60 720

500

1000

1500

20000% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Culture Density (mg/L)

0 12 24 36 48 60 720

500

1000

1500

2000

25000% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Ethanol (mg/L)

0 12 24 36 48 60 720

2

4

6

80% O2

4% O2

6% O2

8% O2

12% O2

Time (h)

Fructose (g/L)

C

A

D

B

Figure A-5. C. ljungdahlii-OTA1 metabolism in reduced medium. A) Growth, B) fructose utilization, C) ethanol production, D) acetate production

136

Figure A-6. C. ljungdahlii-OTA1 growing in reduced medium, headspace analysis at different O2 concentrations. A) 0 % O2 B) 4 % O2 C) 6 % O2 D) 8 % O2 E) 12 % O2

137

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)Gas concentration (mmoles)

A

D

B

C

138

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles) E

139

0 12 24 36 48 60 720

500

1000

15000% O2

2% O2

4% O2

6% O2

8% O2

Time (h)

Culture Density (mg/L)

0 12 24 36 48 60 720

1

2

3

4

5

6

70% O2

2% O2

4% O2

6% O2

8% O2

Time (h)

Fructose (g/L)

0 12 24 36 48 60 720

500

1000

1500

2000

2500

0% O2

2% O2

4% O2

6% O2

8% O2

Time (h)

Ethanol (mg/L)

0 12 24 36 48 60 720

2000

4000

6000

8000 0% O2

2% O2

4% O2

6% O2

8% O2

Time (h)

Acetate (mg/L)

A

DC

B

Figure A-7. C. ljungdahlii-OTA1 metabolism in unreduced medium. A) Growth, B) fructose utilization, C) ethanol production, D) acetate production

140

Figure A-8. C. ljungdahlii-OTA1 growing in unreduced medium, headspace analysis at different O2 concentrations. A) 0 % O2 B) 2 % O2 C) 4 % O2 D) 6 % O2 E) 8 % O2

141

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)Gas concentration (mmoles)

A

DB

B

142

0 12 24 36 48 60 720

1

2

3

4H2

O2

CO

CO2

Time (h)

Gas concentration (mmoles)

E