Improving Fermentation of Pretreated Biomass by Reducing Inhibitor Concentrations and Increasing Microbial Biocatalyst Tolerance

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Improving Fermentation of Pretreated Biomass by Reducing Inhibitor Concentrations and Increasing Microbial Biocatalyst Tolerance
Geddes, Ryan D
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University of Florida
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Doctorate ( Ph.D.)
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University of Florida
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Microbiology and Cell Science
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Biomass ( jstor )
Enzymes ( jstor )
Ethanol ( jstor )
Ethanol production ( jstor )
Fermentation ( jstor )
Plasmids ( jstor )
Polyamines ( jstor )
Pretreatment ( jstor )
Sugars ( jstor )
Toxicity ( jstor )
Microbiology and Cell Science -- Dissertations, Academic -- UF
biomass -- detoxify -- ethanol -- furfural -- hydrolysate -- inhibitors -- lignocellulose -- polyamines
bibliography ( marcgt )
theses ( marcgt )
government publication (state, provincial, terriorial, dependent) ( marcgt )
born-digital ( sobekcm )
Electronic Thesis or Dissertation
Microbiology and Cell Science thesis, Ph.D.


Lignocellulosic biomass provides an abundant source of carbohydrates that can be utilized for the production of fuels and chemicals. However, plant biomass is resistant to degradation, resulting in the need for pretreatment processes that allow access to the fermentable sugars. During pretreatment, compounds inhibitory to fermentation are generated. Among these, furfural has been shown to be a key inhibitory compound, acting synergistically with other inhibitors. Comparison of mRNA profiles of two furfural-resistant mutants (EMFR9 and EMFR35) to their parent, strain LY180, provided evidence for the importance of polyamine transporters in conferring tolerance to furfural. Each mutant contained a single polyamine transporter gene (potE or puuP) that was up-regulated by over 100-fold compared to the parent LY180, and a mutation that silenced the expression of yqhD. Based on the phenotype of these two genetic changes in the mutants, furfural tolerance was reconstructed in strain LY180. The genes encoding 8 polyamine transporters in E. coli were cloned and tested for their effect on furfural toxicity. Half of these increased furfural tolerance (PotE, PuuP, PlaP and PotABCD). Supplementing AM1 medium with individual polyamines (agmantine, putrescine and cadaverine) also increased furfural tolerance to a limited extent. In pH-controlled fermentations, plasmids containing polyamine transporter genes were shown to promote the metabolism of furfural and substantially reduce the time required to complete fermentation. A second process-based approach for improving hydrolysate fermentatbility was employed. Tube culture assays were designed that focus on comparing biological, chemical and physical treatments of hydrolysate. Six strategies were investigated that reduced toxicity in hemicellulose hydrolysates of sugarcane bagasse. These strategies included the use of vacuum evaporation, laccase enzymes, high pH treatment (ammonium hydroxide), bisulfite and increasing cell mass either by increased inoculum or by providing a low level of aeration (0.01 vvm). High pH was the most beneficial single treatment, increasing the minimum inhibitory concentration from 15% (control) to 70% hydrolysate. Tube culture assays of toxicity for ethanol production proved to be excellent predictors of performance in pH-controlled fermenters. A combination of treatments completely eliminated all inhibitory activity in hydrolysate, rendering it as fermentable as laboratory mineral salts medium without hydrolysate. ( en )
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Thesis (Ph.D.)--University of Florida, 2014.
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by Ryan D Geddes.

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© 2014 Ryan Geddes


To my wife Claudia for believing in me and to my son Alessandro for being my constant source of inspiration


4 ACKNOWLEDG E MENTS I would like to express my deep gratitude to Dr. Lonnie Ingram for giving me the opportunity to join his lab and to conduct my research with the benefit of his mentorship and guidance . I would also lik e to thank my committee members Dr. Claudio Gonzalez, Dr. Julie Maupin Furlow , Dr. K.T. Shanmugam and Dr. Spyros Svoronos for their i nvaluable advice and support throughout my time in graduate school. Special thanks to Dr. Svoronos for helping and encouraging me to pursue my d octoral d egree. Also special thank s to Dr. Shanmugam who always found time to listen to another one of my ideas and provide some much needed perspective. I thank Dr. Xuan Wang, Dr. Huaba o Zheng, Lorraine Y o mano and Sean York who were instrumental in providing me with the skills and tools n ecessary to further my research. I am eternally grateful to all my family for their love and support and for the many sacrifices they have made on my behalf. Finally, I thank my loving wife Claudia and my charismatic son Al essandro for loaning me the time to continue my education.


5 TABLE OF CONTENTS page ACKNOWLEDGEMENTS ................................ ................................ ............................... 4 LIST OF TABLES ................................ ................................ ................................ ............ 7 LIST OF FIGURES ................................ ................................ ................................ .......... 8 ABSTRACT ................................ ................................ ................................ ..................... 9 CHAPTER 1 INTRODUCTION ................................ ................................ ................................ .... 11 Biofuels ................................ ................................ ................................ ................... 11 Lignocellulosic Biom ass ................................ ................................ .......................... 12 Pretreatment of Lignocellulosic Biomass ................................ ................................ 14 Biological Pretreatment ................................ ................................ .................... 16 Phys ical Pretreatment ................................ ................................ ...................... 17 Chemical Pretreatment ................................ ................................ ..................... 18 Hydrolysate Toxicity and Mechanism of Inhibition ................................ .................. 19 Common Inhibitory Compounds Present in Dilute Acid Hydrolysates .............. 19 Inhibitory Mechanisms of Hydrolysate Compounds ................................ .......... 20 Organic Acids ................................ ................................ ................................ ... 21 Aldehydes ................................ ................................ ................................ ......... 22 Phenolic Compounds ................................ ................................ ....................... 25 Enzymatic Hydrolysis of Lignocellulosic Biomass ................................ ................... 27 Fermentation of Lignocellulosic Sugars ................................ ................................ .. 28 Objectives ................................ ................................ ................................ ............... 29 2 POLYAMINE TRANSPORTERS AND POLYAMINES INCREASE FURFURAL TOLERANCE DURING XYLOSE FERMENTATION WITH ETHANOLOGENIC Escherichia coli STRAIN LY180 ................................ ................................ ............. 34 Improving Hydrolysate Tolerance of Microbial Bioctalysts ................................ ...... 34 Materials and Methods ................................ ................................ ............................ 36 Results ................................ ................................ ................................ .................... 40 Discussion ................................ ................................ ................................ .............. 47 3 COMBINING TREATM ENTS TO IMPROVE THE FERMENTATION OF SUGARCANE BAGASSE HYDROLYSATES BY ETHANOLOGENIC Escherichia coli LY180 ................................ ................................ ............................ 61 Chemical and Process Methods to Reduce Hydrolysate Toxicity ........................... 61 Materials and Methods ................................ ................................ ............................ 63 Results and Discussion ................................ ................................ ........................... 67


6 4 CONCLUSIONS ................................ ................................ ................................ ..... 83 LIST OF REFERENCES ................................ ................................ ............................... 87 BIOGRAPHIC AL SKETCH ................................ ................................ .......................... 105


7 LIST OF TABLES Table page 1 1 Compostion of lignocellulosic biomass types. ................................ .................... 31 1 2 Carbohydrate and chemical composition of lignocellulosic biomass types. ........ 31 1 3 Major inhibitory compounds present in hardwood, softwood, and agricultural residue dilute acid hydrolysates of lignocellulosic biomass types. ...................... 32 2 1 Bacterial strains, plasmids, and primers. ................................ ............................ 49 2 2 Primers for RT PCR and sequence confirmation. ................................ ............... 51 2 3 Microarry expression ratios comparing the relative exp ression of polyamine transporter genes in furfural resistant mutants (EMFR9 and EMFR35) to the parent LY180. ................................ ................................ ................................ ..... 52 2 4 mRNA levels of potE and puuP in EMFR9 and EMFR35 relative to LY180. ...... 52 3 1 Sugar and inhibitor concentrations after hydrolysate treatments. ....................... 76 3 2 Estimated costs for hydrolysate treatments. ................................ ....................... 77


8 LIST OF FIGURES Figure page 1 1 Production of inhibitors by dehydration of 6 carbon sugars and 5 carbon sugars to hydroxymethylfurfural and furfural, respectively, during dilute acid pretreat ment of lignocellulosi c biomass ................................ .............................. 33 2 1 Comparison of furfural tolerance in the parent LY180 and mutants isolated from serial transfers in AM1 containin g furfural. ................................ ................. 53 2 2 Effect of polyamine transporter plasmids on furfural tolerance in LY180 ............ 54 2 3 Effect of puuP and potE deletions on furfural tolerance ................................ ...... 55 2 4 Alignment of Illumina sequencing reads from EMFR35 on the puuP region of LY180 (parent) ................................ ................................ ................................ .... 56 2 5 Reconstructing furfural tolerance in LY180 based on EMFR9 and EMFR35 ...... 57 2 6 Effect of polyamine supplements (1, 5 or 10 mM) on furfural (10 mM) tolerance in LY180 ................................ ................................ .............................. 58 2 7 Effect of polyamine transporter plasmids on the fermentation of strain LY180 in AM1 medium containing furfural (10 mM) and100 g/L xylose ......................... 59 2 8 Effect of polyamine transporter expression on the growth of E.coli LY180 in the absence of furfural (48 h incubation). ................................ ........................... 60 2 9 Effect of polyamine supplements on the growth of LY180 in the absence of furfural (48 h incubation). ................................ ................................ .................... 60 3 1 Metrics used to evaluate toxicity in treated hydrolysates ................................ .... 78 3 2 Effect of chemical and process treatments on ethanol production in hemicellulose hydrolsates using LY180 (MICEt 50 and MICEt 100 values) ............ 79 3 3 Effect of vacuum evaporation on furfural removal and fermentation of hemicellulose hydrolysate (50% dilution). ................................ ........................... 80 3 4 Fermentation of treated hydrolysate with LY180 ................................ ................ 81 3 5 Fermentation of 90% hydrolysates by LY180 ................................ ..................... 82


9 Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy IMPROVING FERMENTA TION OF PRETREATED BIOMASS BY REDUCING INHIBITOR CONCENTRATIONS AND INCREASING MICROBIAL BIOCATALYST TOLERANCE By Ryan Geddes December 2014 Chair: Lonnie Ingram Major: Microbiology and Cell Science Lignocellulosic biomass provides an abundant source of carbohydrates that can be utilized for the production of fuels and chemicals. However, lignocellulosic biomass is resistant to degradation, resulting in the need for pretreatment processes that allow a ccess to the fermentable sugars. During pretreatment, compounds inhibitory to fermentation are generated. Among these, furfural has been shown to be a key inhibitory compound, acting synergistically with other inhibitors. Comparison of m RNA profiles of two furfural resistant mutants (EMFR9 and EMFR35) to their parent , strain LY180 , provided evidence for the importance of polyamine transporters in conferring tolerance to furfural. Each mutant contained a single polyamine transporter gene ( potE or puuP ) that was up regulated by over 100 fold compared to the parent LY180, and a mutation that blocked the expression of yqhD . Based on the phenotype of these two genetic changes in the mutants , furfural tolerance was reconstructed in strain LY180. The genes en coding 8 polyamine transporters in E. coli were cloned and tested for their effect on furfural toxicity . Half of these increased furfural tolerance (PotE, PuuP, PlaP and PotABCD). Supplementing


10 AM1 medium with individual polyamines (agmantine, putrescine and cadaverine) also increased furfural tolerance to a limited extent. In pH controlled fermentations, plasmids containing polyamine transporter genes were shown to promote the metabolism of furf ural and substantially reduce the time required to complete fermentation. A second process based approach for improving hydrolysate fermentability was employed. Tube culture assays were designed that focus on comparing biological, chemical and physical tr eatments of hydrolysate. Six strategies were investigated that reduce d toxicity in hemicellulose hydrolysates of sugarcane bagasse. These strategies included the use of vacuum evaporation, laccase enzymes, high pH treatment (ammonium hydroxide), bisulfite and i n creasing cell mass either by increased inoculum or by providing a low level of aeration (0.01 vvm) . High pH was the most beneficial single treatment, increasing the minimum inhibitory concentration from 15% (control) to 70% hydrolysate. Tube culture assays of toxicity for ethanol production proved to be excellent predictors of performance in pH controlled fermenters. A combination of treatments completely eliminated all inhibitory activity in hydrolysate, rendering the hydrolysate as fermentable as laboratory mineral salts medium.


11 CHAPTER 1 INTRODUCTION Biofuels Fossil fuels are the primary source of energy worldwide but are limiting . S everal leading energy experts including the US Energy Information Administration have predicted that global oil production will peak prior to the year 2050 with a steep decline thereafter ( 1 ). At the same time, global energy demand is expected to continue to increase by at least 20% compared to 2009 driven mainly by grow ing economies like China and India ( 2 ). The US is the largest consumer of petroleum in the world, using an average of 19 million barrels of petroleum per day of which 65% is imported. Petroleum accounts for 40% of the total US energy use . Up to 71% of the imported petroleum is used in the transportation sector to fuel cars and trucks ( 3 ). Liquid fuel demand for the transportation sector is anticipated to rise dramatically due to an estimated 2.3 billion additional cars running worldwide by the year 2050 (1 .9 million from emerging markets ; 3 ). A large proportion of imported oil is obtained from 4 5 countries (Russia, Saudi Arabia, Libya and Iran) which could set up difficult situations for the importing countries due to fluctuations in the price of crude o il ( 4 ) arising from inflation and political atmosphere contributing significantly to global warming. The transportation sector contributes 25% of the global energy related ca rbon dioxide emissions ( 5 ). Biofuels are the most promising option for reducing oil demand and carbon dioxide emissions caused by the transportation sector. Cellulosic biofuels could reduce green house gas


12 emissions ( GHG ) by as much as 90% compared to pet roleum based fuels ( 6 , 7 ). An advanced biorefinery could be energy positive providing energy to the grid and requiring no fossil fuel input , because the plant material used to make the energy is capture d in the product and the remaining residue. A carbon n eutral process is made possible by t he establishment of a cycle where biomass feedstock is grown using the carbon dioxide emitted during the biofuels production process ( 7 ) . In an attempt to reduce US trade deficits, decrease dependence on oil imports, in crease energy security, lower greenhouse gas emissions and create domestic jobs, the Energy Policy Act of 2005 and 2007 was passed by Congress. The Energy Policy Act of 2007 mandates the production of 36 billion gallons of ethanol per year by 2022 with 44% derived from cellulosic sources ( 8 ). Lignocellulosic Biomass The use of non food sources for biofuels production is favored due to rising world demand for food. Lignocellulosic biomass represents a non food source and is the most abundant renewable resou rce on Earth. Another advantage of lignocellulosic biomass is its presence all over the world and the potential for growing local sources without estimates that there is 1. 3 billion tons of biomass available annually which could be converted to 130 billion gallons of liquid fuel ( 9 ). The types of lignocellulosic biomass can be grouped into three major categories: 1. softwoods (conifers), 2. hardwoods (angiosperm trees) and 3. grasses (Poaceae family). Lignocellulose is plant cell wall material that is composed of three major polymers: hemicellulose, cellulose and lignin (10) . Unlike starch from corn kernels these polymers represent the structural components of plants which are designed to be


13 resistant to chemical and biological degradation (11) . The exact distribution and composition of the cellulose, hemicellulose and lignin depends on various factors including the type of plant, its age, the part of the plant and growth c ondition (10) . Plant cell material can be broken down into individual cell walls (primary, secondary and tertiary) each composed of distinct ratios of the 3 major polymers (10) . The primary plant cell wall is present on the outside of the plant cell and is primarily composed of lignin. The secondary cell wall contains a majority of the polysaccharide and lignin polymers (10) . Cellulose is a crystalline homopolymer made up of D glucose units connected by 1,4 glycosidic bonds. The glucose units contain hyd roxyl groups that hydrogen bond with neighboring cellulose chains creating a crystalline tightly packed structure (12) . The individual chains can vary in length from 500 to 15 , 000 residues ( 12 ). These chains can create tightly packed bundles called element ary microfibrils made up of 20 40 chains (12) . These elementary microfibrils can in turn hydrogen bond with each other to produce macrofibrils containing amorphous and crystalline regions coated by hemicellulose, pectin and lignin (12) . The resulting matri x creates a cell wall with high tensile and mechanical strength (12) . The cellulose content of sofwoods, hardwoods and agricultural residues are 41 50%, 31 53% and 24 50% respectively (Table 1 1 ; 10 ) . The main differences between the various types of ligno cellulose are in the individual sugars that make up the heteropolymer hemicellulose and the side chains of the aromatic polymer, lignin (Table 1 2 ; 10 ) . The lignocellulose structure is similar to reinforced concrete. The cellulose macrofibrils are like th e steel bars coated with a flexibile neoprene like lignin and hemicellulose matrix ( 13 ). Hemicellulose is a more


14 complex heteropolymer of different carbohydrates and having various side chain compounds attached. The major components of this heteropolymer a re xylan, mannan, glucan and galactan (10) . Sofwoods typically contain the highest amount of lignin and are also considered the most difficult to break down into individual carbohydrates (14) . Agricultural residues contain the least amount of lignin and a re in turn less resistant to chemical and biological hydrolysis (14) . Lignin is the second most abundant polymer on Earth. It forms covalent linkages with hemicellulose providing mechanical strength to plants ( 10 ). L ignin is made up of aromatic units covalently linked through ether and carbon carbon bonds forming a complex non defined structure ( 10 ). Lignin is made up of 3 major building blocks that are incorporated in the form of the phenylpropanoids, p hydroxyphenyl, guaiacyl and syringyl. The phenyl propanoids can be incorporated to varying degrees and can have different side chain constituents. Softwoods, hardwoods and grasses evolved separately and are found to contain different amounts and types of lignin and hemicellulose structures ( 10 ). Pretreat ment of Lignocellulosic Biomass The cost effective breakdown of lignocellulosic biomass into sugars that can be fermented continues to be a substantial hurdle (15) . The process must be cost competitive with the very well known and optimized production of g asoline as fuel for vehicles. Major cost drivers are raw materials, enzymes, sugar yields and ethanol yields (15) . Enzyme companies such as Novozymes and Genencor have achieved major cellulase cost reductions for second generation processes over the last 10 years (16) . The estimated enzyme use costs have been claimed to be reduced up to 20 fold to


15 $0.50/gallon of ethanol or lower (16). However, the cost of enzymes still needs to be reduced since constraints exist in other parts of the process such as pretr eatment conditions. The generation of inhibitors during pretreatment limits fermentation efficiency , and the optimum pretreatment condition for enzyme hydrolysis is rarely the same as that for maximum fermentation yields (17) . Enzyme catalyzed conversion o f lignocellulose to monomeric sugars is a slow process unless the cellulosic biomass has undergone some form of pretreatment to disrupt the naturally recalcitrant plant cell wall structures (18) . Pretreatment is a crucial and highly influential step in the biomass to sugars and ethanol process because it has a direct impact on many downstream processes including enzyme hydrolysis sugar yields, fermentation efficiency, energy demand and wastewater treatment (19) . The type of pretreatment employed will determ ine the specific manner in which the crystallinity, surface area, porosity and chemical composition of the biomass substrate are altered (20) . The pretreatment of lignocellulosic materials seeks to effectively alter the biomass structural elements to incr ease cellulase digestibility while minimizing the generation of inhibitors, the loss of sugars and the use of chemicals (18) . Meeting all these requirements under the same pretreatment condition has proven to be a formidable challenge (17) . A vast quantity of research has focused on discerning the most effective types of pretreatment for various biomass types (21) . Pretreatments can be categorized into biological, physical, physico chemical and chemical. Each pretreatment system has its own advantages and d isadvantages and the most efficacious pretreatment will depend on the substrate and specific downstream process requirements.


16 Biological Pretreatment Naturally occurring microorganisms such as white , soft and brown root fungi are capable of degrading lignocellulosic biomass under mild environmental conditions with no chemical additions (22) . However, the process is much slower than the combination of pretreatment and enzymatic hydrolysis (22) . These microorganisms produce and release enzymes into their external milieu to deconstruct biomass into sugars they can use as a food source (22) . Fungi and bacteria capable of degrading lignocellulose are specific to the species of wood to be degraded ( 23 ). Softwoods contain lumens that are completely surrounded by cell walls requiring microorganisms capable of penetrating through the cell wall to reach inside the lumen (23) . On the other hand, the vessels of hardwoods have open ends where microorganisms can enter the wood cells freely and degrade the cell walls ( 23) . The chemical structure of hardwood and softwood cell walls also differ s as previously mentioned and require the microorganisms to synthesize unique cellulolytic cocktails (23 24) . The brown rot fungi act mostly by hydrolyzing the cellulose and hemicel lulose but can also modify lignin in its non oxidized form ( 24 ). In contrast to white and soft rot fungi, brown rot fungi extensively hydrolyze the wood cell walls irrespective of whether or not the substrates are used for metabolism ( 25 ). Researchers have shown the presence of small low molecular weight hydroxyl radicals produced by fungi such as phenolates, oxalate and aromatic compounds in the early phases of brown rotting ( 26 33 ) . These compounds are thought to act as ferric ion chelators, reducing agen ts and redox cycling compounds used by cellulolytic enzymes for hydrolysis of lignocellulose ( 34 ). Lignin is the main structural barrier to enzymatic and biological attack of lignocellulosic biomass. The key enzymes produced by wood degrading microorganism s are the lignin degrading


17 enzymes lignin peroxidase, manganese peroxidase and laccase ( 35 36 ). These enzymes require various mediators and cofactors to catalyze their respective reactions. Laccase uses oxygen as an electron acceptor while peroxidases use hydrogen peroxide during lignin degradation and modification ( 3 5 ). Physical Pretreatment Physical pretreatments reduce the particle size, porosity and cellulose crystallinity of lignocellulosic biomass without the addition of chemicals (3 7 ) . The most com mon methods are comminuition (chipping, milling and grinding), pyrolysis and irradiation (38) . During the milling process, the size of lignocellulosic materials decrease s from 10 30 mm to 0.2 2 mm ( 39 ). Milling increases surface area and decreases cellulos e crystallinity although it requires high energy input. In general, as particle size decreases during milling, energy consumption increases exponentially ( 40 ). Different milling technologies exist including hammer, ball and knife milling (41) . The choice o f milling technology is driven by the particular physical and chemical properties of the material including moisture content. Some researchers have suggested that milling processes can disrupt the chemical bonds between lignin and the structural carbohydra tes of plants increasing the accessibility of the cellulolytic enzymes ( 42 ). In many applications, milling as a stand alone pretreatment is cost prohibitive due to high energy demand and enzyme costs. Pyrolysis has gained popularity in the last decade wi th the advent of start ups ( e.g . Evergent Technologies, KioR) applying the technology to a variety of feedstocks. The biomass is heated to temperatures exceeding 300 ° C causing plant structural polymers to breakdown into volatile gases, pyrolysis oil and a lignin rich biochar ( 43 44 ).


18 Irradiation is a less commonly studied physical pretreatment method but has the added advantages of high heating efficiencies and reduced operating costs ( 45 ). Irradiation of rays can disrupt glycosidic bonds withi n cellulose chains but the presence of lignin may be required ( 46 47 ). Microwave pretreatment is another type of irradiation but it uses water to release acetyl groups naturally present in the biomass. The released acetate acts as a catalyst in the hydroly sis of hemicellulose ( 47 ). Irradiation methods have not been widely adopted due to high capital costs ( 47 ). Chemical Pretreatment Methods of chemical pretreatment include acid hydrolysis, alkaline hydrolysis, oxidative delignification, ozonolysis, wet oxi dation and solvent extraction. Acid pretreatment of lignocellulosic material has been the most widely studied with dilute sulfuric acid being the most common ( 48 5 1 ). Acid pretreatment uses either concentrated or dilute acid to disrupt the tightly packed r ecalcitrant structure of lignocellulose. Acid hydrolysis has been used to treat diverse feedstocks ranging from switchgrass ( 52 53 ) and corn stover ( 54 ) to spruce ( 55 ) and poplar ( 56 57 ). Other types of acids have also been investigated, such as nitric ( 58 59 ), hydrochloric ( 60 62 ) and phosphoric ( 6 3 67 ). Acid hydrolysis removes hemicellulose, disrupts the lignin carbohydrate complex by redistributing lignin and significantly improves cellulose enzymatic digestibility ( 68 ). In general, dilute acid pretreatment (0.2 4% w/w) can be performed at high temperatures (160 ° C 220 ° C) and short times ( e.g. 5 minutes) or at low temperatures (120 ° C 160 ° C) and long times ( e.g. 1 hour) ( 69 78 ). Although acids are ideal catalyst for the fractionation of lignoce llulosic materials, their use in processes has some significant disadvantages: the corrosive nature of acids necessitates


19 expensive reactor construction and resulting pretreatment hydrolysates contain inhibitory sugar degradation products. The optimum pret reatment condition var ies with each biomass type and a balance must be struck between minimum sugar degradation and maximum enzymatic hydrolysis yields. Hydrolysate Toxicity and Mechanism of Inhibition Common Inhibitory Compounds Present in Dilute Acid Hyd rolysates Though many inhibitory compounds are produced during dilute acid hydrolysis of lignocellulose, some have been proven to be present at higher levels and/or are particularly inhibitory to the fermenting organism (7 9 ) . Inhibtory compounds found in h emicellulose hydrolysate are from 3 major classes: furans, organic acids, and phenolic compounds (7 9 ) . As shown in Figure 1 1 , furans are produced by the dehydration of pentose and hexose sugars resulting in furfural and hydroxymethyl furfural production, respectively (7 9 8 2 ) . Organic acids ( such as acetic, formic and levulinic acid ) can be produced by the degradation of furans and by the cleavage of acid side chains present in the hemicellulose structure. Various phenolic compounds are produced from the b reakdown of lignin polymer. Hydrolysis of t he different classes of biomass (hardwood, softwood, and agricultural residue) may produce slightly different concentrations of inhibitory compounds (7 9 , 83 ) . T able 1 3 below lists the major inhibitory compounds present in hardwood, softwood, and agricultural residue dilute acid hydrolysates ( 79 86 ) . Softwood hemicellulose contains more glucose and mannose units than hardwood or agricultural residue hemicellulose whi ch contain more xylose units ( 10, 7 9 ). Therefore, dilute acid pretreatment of hardwood and agricultural residue may produce more furfural than that of softwood but would depend on the severity of the


20 pretreatment. The hemicellulose of hardwoods and agricul tural residues are also typically more acetylated than that of softwoods ( 10, 7 9 ). Thus, d ilute acid hydrolysates of hardwoods and agricultural residues may contain more acetic acid than those of softwood. There are two major types of lignin: guaiacyl and guaiacyl syringyl ( 10, 7 9 ). The distinction is in the side chain found in the phenylpropanoid skeleton. The guaiacyl syringyl lignins contain a methoxy group in the 3 and 5 carbon position while the guaiacyl lignins only contain a methoxy group at the 3 c arbon position ( 10, 7 9 ). Softwood lignin is mostly made up of the guaiacyl units and hardwood lignin is primarily made up of the guaiacyl syringyl type. Palmqvist and Hahn Hagerdal (2000) reported that syringaldehyde and syringic acid were found in hardwoo d hydrolysates ( 7 9 ). T he y also reported the presence of vanillin and vanillic acid formed by the degradation of the guaiacyl propane units of willow, spruce, poplar, pine, and red oak lignins ( 7 9 ). They also found that most of the lignin derived phenolic c ompounds were made up of 4 hydroxybenzoic acid constituents in hardwood hydrolysates ( 7 9 ). Inhibitory Mechanisms of Hydrolysate Compounds The fine balance , between the harsh conditions required during pretreatment due to the recalcitrance of lignocellulos e and the protection of solubilized sugars from further exposure to those conditions , is very difficult to maintain. Consequently, hydrolysates are rarely devoid of compounds that inhibit subsequent fermentation processes. Substantial research has been dev oted to the study of key inhibitors present in hydrolysates, method of toxicity and detoxification strategies. Hydrolysates vary in their degree of inhibition and biocatalysts respond differently to the inhibitors ( 7 9 86 ).


21 Biocatalysts experience an aggregate inhibition effect imposed by the specific combinations of inhibitors present in hydrolysates. Organic A cids Acetate is one of the most abundant inhibitors in hydrolysates, present at concentrations ranging from 2 15 g/L ( 8 7 ). However, several oth er organic acids are produced in the pretreatment process including some derived from structural components of the lignocellulose or from sugar degradation products. Zaldivar and Ingram (1999) investigated the effects of organic acids (acetic, ferulic, gal lic, 4 hydroxybenzoic, syringic, vanillic, furoic, formic, levulinic, caproic) on the growth of ethanologenic E. coli strain LY01 (8 8 ) . They found a high correlation between the hydrophobicity of the organic acids and their inhibitory effect (pH 7.0). The order of inhibition was: caproic > ferulic > feroic > 4 hydroxybenzoic > formic > acetic > levulinic. In general, higher initial fermentation pH ( e.g. pH 7.0 v. pH 6.0) and higher temperatures ( e.g. 40 ° C v. 30 ° C) were observed to result in improved growth during organic acid stress (88) . Palmqvist et al. (1996) and Roe et al. (1998) suggested that organic acids penetrate the cell membrane in an undissociated form and dissociate after entering the cytoplasm (8 9 90 ) . Th e dissociated acid can then cause a coll apse of the proton gradient in the cell , resulting in a decrease of ATP ( primary energy source of the cell ) . Increasing initial cell density of ethanologenic E. coli strain LY01 by up to 40 fold did not overcome acid inhibition (88) . Binary combinations of organic acids displayed an additive toxicity that more closely mimic hydrolysate conditions ( 8 8 ). Interestingly, the combination of the two most abundant inhibitors in dilute acid hydrolysates furfural


22 and acetate resulted in a synergistic inhibition of growth (80% reduction). Membrane leakage was not found to be a major inhibitory mechanism for organic acids (88) . Aldehydes Furfural has been identified as one of the key inhibitors affecting fermentation of dilute acid hydroly sates ( 89 91 ). The presence of furfural and furan derivatives prolong s the lag phase during the initial 24 h ours of fermentation in both bacteria and yeast ( 91 9 6 ). In addition, the extent of the lag was a function of the initial furfural concentration in S. cerevisiae hydrolysate fermentations ( 94 95 ). S. cerevisiae ( 91, 94 95 ) and E. coli ( 92 93 ) may be metabolizing and detoxifying furfural to a less toxic compound via an unknown mechanism. Miller et al . (2009) isolated furfural resistant strains of E. co li that were deficient in furfural reduction by the NADPH dependent oxidoreductases (9 7 ) YqhD and DkgA. This finding was counterintuitive as these enzymes can catalyze the reduction of furfural to the less toxic compound furfuryl alcohol. The authors perfo rmed 53 serial transfers in pH controlled vessels under constant furfural stress. The resulting strain was able to produce the same level of ethanol as its parent strain (no furfural), in the presence of up to 1.0 g/L furfural. A 72 h lag was observed in t he parent strain under similar conditions (1.0 g/L furfural) . Comparison of mRNA profiles revealed 12 oxidoreductases that were up or down regulated at least two fold following the addition of 0.5 g/L furfural (9 7 ) . Of particular interest were four oxidor eductase genes that were downregulated in the presence of furfural. Three of these, yqhD, dkgA, and yqfA, decreased furfural tolerance when overexpressed from plasmids. Deletion of yqhD as well as expression of the NADH/NADPH transhydrogenase, PntAB, increased furfural tolerance (9 7 ) . Similar


23 global transcript analysis in S. cerevisiae identified a 5 hydroxymethylfurfural reductase, ADH6, involved in tolerance to HMF ( 96 ). The effect of varying carbon ( e.g. glucose and xylose) and nutrient source ( e.g. yeast extract) have also been investigated ( 95 ). The use of glucose and yeast extract improved the growth of E. coli in the presence of furfural. The proposed mechanism of glucose and yeast extract benefit was the generation of more NADPH available for bi osynthesis and reduction in biocatalyst biosynthetic needs , respectively (95) . Subsequent studies by Miller et al . (2009) using the parent strain LY180, revealed that sulfur assimilation was one of the targets of furfural inhibition (9 8 ) . The NADPH pools required for amino acid and other biosynthetic reactions were being depleted during the reduction of furfural by oxidroreductases with a low K m for NADPH fold and approx fold when E. coli was exposed to 0.5 g/L furfural (9 8 ) . The genes were divided up by functional groups. Amino acid and nucleotide biosynthesis genes were particularly altered (>20% of total genes in the functional grou p) in response to furfural. Most of the genes were downregulated. Growth of E. coli was improved by supplementation with 0.1 mM of 5 out of 20 amino acids tested. The order of benefit was: cysteine > methionine > serine and arginine > histidine. In additio n, expression of genes involved in sulfur assimilation w as generally increased. The authors hypothesized that E. coli cells were deficient in sulfur containing amino acids during furfural stress due to the inhibition of sulfate metabolism. Adding sulfur co ntaining amino acids, amino acid precursors as well as sulfur compounds increased furfural tolerance (9 8 ) .


24 Wang et al. (2011) identified a beneficial oxidoreductase, FucO, capable of reducing furfural to the less toxic furfuryl alcohol (9 9 ) . The unique feature of this enzyme was that it did not compete with biosynthetic NADPH pools like the previously characterized YqhD ( 9 7 ) but rather it used the anaerobically abundant NADH as the reductant. FucO was known as 1,2 propanediol reductase , however, in vitro studies and crystal structure analysis revealed that it can also reduce furfural using NADH as a cofactor. Purified FucO also exhibited 5 hydroxymethyl furfural reductase acitivity. Furfural tolerance was increased by 50% in the presence of 15 mM furfural when fucO was overexpressed from a plasmid in ethanologenic E. coli LY180 ( 9 9 ). Global transcript analysis was used to search for additional unknown oxidoreductases that can use NADH as a cofactor ( 100 ). Four oxidoreductases, aldA, ydhABC, yeiTA, and ucpA , were upregulated at least 3 fold and had potential NADH binding domains. The expression of ucpA improved growth by 50% in the presence of 15 mM furfural. However, none of the oxidoreductases exhibited furan reductase acitivity in vitro with either NADH o r NADPH ( 100 ) . The exact function of UcpA and its mechanism for conferring furfural tolerance is unknown. The search for a genetic basis to furfural tolerance has also uncovered genes related to DNA biosynthesis. Zheng et al . (2012) isolated the thymidylate synthase gene, thyA , from genomic libraries of three bacterial species: Bacillus subtilis , E. c oli and Zymomonas mobilis (10 1 ) . The thyA containing plasmids were enriched when transformants were grown on medium containing furfural. The authors demonstrated that the growth of E. coli in the presence of furfural is improved by overexpression of thyA a nd supplementation with thymine, thymidine or 5,10 methylenetetrahydrofolate.


25 Increased tolerance was proposed to be due to increased pyrimidine deoxyribonucelotides involved in DNA repair ( 101 ). The exact target(s) of furfural inhibition via pyrimidine de oxribonucleotide depletion remains unknown. However, Zheng et al . (2012) proposed several possible mechanisms: competition for NADPH pool required for deoxyribonucleotide de novo synthesis, direct DNA damage and inhibition of enzymes involved in the folate cycle (10 1 ) . Shahabudin et al . (1991) have reported that furfural can cause DNA damage (10 2 ) . The authors isolated a nuclease from pea seeds capable of specifically targeting A T rich regions of DNA. The nuclease was used to hydrolyze most of the A T ric h sites on native calf thymus DNA. The result was a marked decrease in S 1 nuclease hydrolysis of calf thymus DNA after furfural and hydroxymethylfurfural treatment (molar ratio: 1 mole DNA base pair/16 moles of furan). The authors proposed that S 1 nuclease remov ed the substrate furans tact on ( i.e. A T rich regions of DNA). S ubsequent studies demonstrated that plasmids (pBR322 and pBluescript SK) treated with furfural carried mutat ions ( 10 3 10 4 ). The transformation efficiency of the plasm i ds also d ecreased after furfural treatment . Phenolic Compounds Phenolic compounds are released by partial solubilization of lignin during dilute acid pretreatment ( 105 ) . The resulting phenolic compounds have varying degrees of toxicity to the fermenting biocatalyst. Toxicity is related to the substituents on the phenol ring structure. Phenolic aldehydes have been found to be particularly toxic to biocatalysts ( 88, 93, 10 5 10 8 ). The types of phenolic compounds present in pretreatment liquors are dependent on the biomass used due to differences in lignin structures of softwood, hardwood and grasses ( 10 9 ). The hemicellulose and cellulose


26 bonds with lignin as well as the ratio of syringyl, guiacyl and hydroxyl content will vary with biomass source ( 10 9 ). S. cerevisiae has the native ability to convert phenolic aldehydes to the less toxic corresponding alcohols ( 1 10 ). The order of toxicity of phenolic compound forms is: aldehyde > acid > alcohol ( 88, 93, 105 107 ). Zha et al (2014) identified lignocellulosic biomass inhibitors present in 24 different types of hydrolysates before and after fermentation ( S. cerevisiae CEN.PK 113 7D) using GC MS. S. cerevisiae CEN.PK 113 7D was grown in YPD (yeast extract peptone dextrose) , challenged with various phenolic compounds and growth was compared to the reference fermentation in minimal salts media containing 20 g/L glucose (105) . Lignocellulose inhibitors were added at 0.2, 0.5 and 1.0 g/L concen t rations , and cell growth was measured in a BioScreen C Analyzer (OD 430 580 nm , pH 5.0, 30 ° C). Vanillin (1.0 g/L) was reported to extend the lag phase in fermentation much like furfural ( 105 ). In contrast, the acid form, vanillic acid, did not impact the growth of S. cerevisae . Benzaldehyde and benzoic acid acted mainly by reducing the growth rate ( 105 ). Additional acid phenols such as p coumaric acid and 3 phenyl lactic acid had no effect on growth ( 105 ). Nine phenolic compounds were identified in wet alkaline oxidation pretreated wheat straw and tested for possible inhibitory effects on ethanol production by S. cerevisiae ATCC 96581 ( 111 ). Vanillin, 4 hydroxyacetophenone and acetovanillone at 10 mM conce n t rations resulted in 53 67% reduction in productivity. The phenolic acids, 4 hydroxyvanillic and 4 hydroxysyringic acid as well as syringaldehyde and acetosyringone were less inhibitory resulting in less than a 16% decrease in productivity ( 111 ).


27 The mech anism of phenolic compound inhibition has not been fully elucidated ; h owever, some researchers have proposed that phenolic compounds affect proper protein to lipid ratio incorporation in to the cell membrane (Keweloh et al 1990). Laccases and peroxidases ca n oxidize soluble phenolic compounds in hydrolysate through radical mediated polymerization ( 112 ). Enzymatic Hydrolysis of Lignocellulosic Biomass Cellulases are enzymes that catalyze a series of synergistic reactions used to 1,4 glycosidic bonds found in cellulose. The depolymerization of lignocellulose requires several enzyme types including endoglucanases (1,4 D glucan glucanhydrolases), exoglucanases (1,4 D D glucoside glucohydrolases) (10) . Cellulases contain three general structural domains: cellulose binding domain, interlinker domain and catalytic domain ( 113 114 ) . Enzymatic hydrolysis of cellulose is composed of three basic steps: adsorption of the enzyme onto the cellulose fiber, catalytic depolymerization of cellulose and desorption of the enzyme ( 114 ) . Depolymerization begins by the adsorption of endoglucanases onto the amorphous regions of cellulose ( 115 ) . The cellulose fibril bundles contain amorphous regions that are more susc eptible to chemical and biological attack than the more abundant crystalline regions. The degree of crystallinity of cellulose has been used as a means to predict the resulting enzyme hydrolysis efficiency ( 116 117 ) . Studies have reported a relationship be tween increased surface area and porosity resulting in increased sugar yields after enzyme hydrolysis ( 118 120 ) . The presumed mechanism is the ability for enzymes to penetrate more sites and the presence of more contact area for enzymatic attack.


28 Endogluc anases differ in the presence or absence of a cellulose binding domain (CBD) ( 121 ) . CBD acts as a head that binds to the hydrophobic cellulose surface tightly and directs the processive movement of the cellulase along the cellulose fibril ( 122 123 ) . The CBD can act as a kind of wedge peeling individual elementary fibrils apart and thus allowing the processive movement of cellulase along the chain ( 124 ) . Researchers believe that the endoglucanases lacking CBDs are the first line of attack ( 125 ) . These end oglucanases create nicks within the amorphous parts of the cellulose chain and open up more ordered inner layers for CBD containing endoglucanases to hydrolyze. The resulting hydrated cellulose chain ends are the substrates for the catalytic domain of cell obiohydrolases/exoglucanases (CBH). Some CBHs attack from the reducing end containing free hemiacetal groups at C1 and others from the non reducing end containing the free hydroxyl at C4 ( 124 ) . The concerted action of endo and exo glucanases create s solub le glucose, cellobiose and cellodextrins that can be glucosidases ( 117 ) . Fermentation of Lignocellulosic Sugars The cost effective conversion of lignocellulosic biomass into fuels and che micals depends on the ability of biocatalysts to co utilize the major sugars present in hydrolysates following hemicellulose solubilization after dilute acid pretreatment and cellulase hydrolysis . S. cerevisiae strains have been genetically modified to fer ment xylose in lignocellulose hydrolysates ( 126 128 ). As previously mentioned, hydrolysates contain fermentation inhibitors that must be removed or converted to less toxic compounds to achieve high productivity in fermentation. Microorganisms have also bee n isolated that are more resistant to furfural ( 97 98, 129 131 ) , hydroxymethylfurfural ( 131 1 33 ) , acetate ( 134 ) and phenolic compounds ( 135 136 ) . The co utilization of two of


29 the most abundant sugars in hydrolysates, glucose and xylose, is particularly challenging due to a combination of catabolite repression and the energy burden associated with detoxification of inhibitors and anaerobic fermentation of hydrolysates. However, biocatalysts have been developed that utilize both glucose and xylose d uring f ermentation of hydrolysate ( 126 128, 137 ). Objectives The overall objective of this study is to improv e the production of fuels and chemicals from lignocellulosic biomass by increasing the fermentability of hydrolysates produced during the process. Two ge neral approaches were investigated to overcome the inhibitory effects on fermentation caused by compounds generated during pretreatment. The first approach improves the biocatalyst robustness by engineering the organism to be either more tolerant to inhib itors or to directly aid in the detoxification of inhibitor y compounds . An example of the former is the overexpression of thyA in E. coli which has been demonstated to improve furfural tolerance and is though t to be involved in the repair of furfural induc ed damage to DNA ( 10 1 ). In the latter case a number of oxidoreductases have been shown to reduce aldehydes such as furfural to their less toxic alcohol derivatives ( 93, 97, 99, 100, 106, 130 ) . This study i nvestigates a novel approach to furfural tolerance by examining genes (polyamine transporters) discovered to confer tolerance in the presence of furfural and examines their role in the de velopment of a furfural tolerant biocatalyst. The second approach involves mit igating inhibitor concentration and toxicity in hemicellulose hydrolysates prior to fermentation. For this approach a number of promising detoxification strategies were investigated by utilizing a n assay designed to


30 allow direct comparisons of the differen t treatments, alone or in combination . Th is study also strenghtens the validity of the assay results by investigat ing fermentation profiles of the best treatments identified. Evaluati on of the treatment combinations allow d esign and process configuration i mprovements .


31 T able 1 1. Compostion of lignocellulosic biomass types . Biomass Type Cellulose (% DW) Hemicellulose (% DW) Lignin (% DW) Hardwood 41 50 11 33 19 30 Softwood 31 53 19 36 17 24 Agricultural Residue 24 50 12 38 6 29 References 10, 8 3 Table 1 2. Carbohydrate and chemical composition of lignocellulosic biomass types. Biomass Type Hemicellulose Structure Hardwood Glucuronoxylan Xylan backbone 10% of the xylose units have substituents such as glucuronic acid or methyl glucuronic acid 1:2 ratio of glucose to mannose 7 acetyl groups per 10 xylose units (ester linkages) Softwood Galactoglucomannans Principal component is glucomannan Arabinofuranose linked to the backbone 1:3 ratio of glucose:mannose 1:10 galactose unit/glucose unit Alpha 1,6 galactose links to the glucose mannose chain 1 acetyl group per 3 4 mannose units Arabinoglucuronoxylans Agricultural Residue Glucuronoxylan Xylan backbone 10% of the xylose units have substituents such as glucuronic acid or methyl glucuronic acid 1:2 ratio of glucose to mannose 7 acetyl groups per 10 xylose units (ester linkages) Reference 10


32 T able 1 3 . Major inhibitory compounds present in hardwood, softwood, and agricultural residue dilute acid hydrolysates of lignocellulosic biomass types. Type of Toxic Compound Toxic Compound Hardwood Softwood Agricultural Residue Furans Furfural 5 Hydroxymethyl Furfural P P P P P P Organic Acids Acetic Acid Formic Acid Levulinic Acid P P P P P P P P P Phenolic Compounds 4 Hydroxybenzaldehyde p Coumaric Acid Ferulic Acid Syringaldehyde Vanillin 4 Hydroxybenz oi c Acid Syringic Acid Vanillic Acid P P P P P P P P P P P P P P P P ( P )= presence of the compound in hydrolysate. References 79 83. ( )=no reported presence of the compound in hydrolysate.


33 Figure 1 1. Production of inhibitors by dehydration of 6 carbon sugars and 5 carbon sugars to hydroxymethylfurfural and furfural , respectively , during dilute acid pretreatment of lignocellulosic biomass. Lignin solubilizes into phenolic monomers that can also inhibit biocatalysts.


34 CHAPTER 2 POLYAMINE TRANSPORTERS AND POLYAMINES INCREASE FURFURAL TOLERANCE DURING XYLOSE FERMENTATION WITH ETHANOL OGENIC Escherichia coli STRAIN LY180 Improving Hydrolysate Tolerance of Microbial Bioctalysts Lignocellulosic biomass can be used as a renewable carbohydrate feedstock for the production of fuels and chemicals ( 82 ) . Unlike starch which is used by plants for temporary energy storage, lignocellulose is a structural component designed to resist microbial enzymes and chemical deconstruction. A pretreatment step such as dilute acid hydrolysis is required to increase the accessibility of cellula se enzymes for more efficient carbohydrate hydrolysis ( 138 ). Dilute acid pretreatment is effective at open ing the lignocellulose structure by hydrolyzing hemicellulose into pentose monomers (91,98,139 141) . Dilute acid pretreatments also produce unwanted s ide products such as furfural and hydroxymethylfurfural (HMF) that inhibit fermentation and increase process cost ( 91,98,139 141 ). Increasing concentrations of furans have been correlated with toxicity of hydrolysates. Addition of furfural to detoxified hy drolysates has been shown to restore toxicity ( 80, 92 ). Genes have been described that have products allowing increase d resistance of bacterial and yeast biocatalysts to furfural and hydroxymethylfurfural ( 91 , 97, 99,101, 132, 141 148 ). However to date, none have completely solved the toxicity problem. In yeast, overexpression of the transcription factor Yap1 has been shown to increase tolerance to both furfural and HMF ( 147 ). Disruption of pentose phosphate pathway (PPP) genes ( ZWF1, GND1, RPE1 and TKL1 ) led to increased sensitivity to furfural and HMF in Saccharomyces cerevisiae ( 142 ). Disruption of PPP genes is proposed to


35 decrease the levels of NADPH, the cofactor for many enzymes involved in cellular defense mechanisms and biosynthesis ( 142 ). A mutate d form of the alcohol dehydrogenase ADH1 from S. cerevisiae was shown to confer tolerance by reducing HMF to 2,5 bis hydroxymethylfuran using NADH instead of NADPH ( 148 ). Microarray analysis of mRNA transcripts in a furfural resistant mutant of Escherich ia coli (strain EMFR9) revealed that the NADPH dependent YqhD is involved in furan tolerance ( 97 ). Native expression of chromosomal yqhD was found to inhibit the growth of E. coli in media containing furfural ( 97 98 ). Blocking the functional expression of yqhD increased resistance to furfural. The detrimental effect of YqhD (low K m for NADPH) has been attributed to competition for NADPH, limiting biosynthesis. In contrast, increased expression of an NADH dependent oxido reductase gene ( fucO ) was beneficial f or furfural tolerance ( 99 ). Both enzymes reduce furfural (aldehyde) to the less toxic furfuryl alcohol. Recently , a new genomic tool, multi SCale Analysis of Library Enrichments (SCALE), was developed and using this analysis, f our genes in E. coli that inc rease furfural tolerance were identified ( 145 ): thyA (DNA biosynthesis), lpcA (lipopolysaccharide biosynthesis) , groES and groEL (GroES EL chaperonin complex). Increased expression of the groES , groEL , grpE and c lpB genes ( 149 150 ) also i ncrease d ethanol tolerance. The precise mechanism of furfural toxicity is unknown. Although methods have been developed to mitigate or remove inhibitors ( 151 ), all increase process complexity. Furfural has been shown to cause strand breaks in DNA ( 152 ), damage membranes ( 153 ) and react with other cellular components ( 142 , 154 ).


36 Polyamines can bind both nucleic acids and phospholipid membranes, reducing exposure of these vital cellular constituents to furfural. In E. coli , intracellular levels of polyamines are regulated by biosynthesis, degradation, and expression of transporter genes ( 155 ). T his study report s that plasmids containing polyamine transporter genes and polyamine supplements can be used to increase furfural tolerance in ethanologenic E. coli LY180. Material s and Methods Strains, media, and growth conditions. Table 2 1 contains a list of strains, plasmids, and primers used in this study. Table 2 2 lists primers used for RT PCR reactions and for sequencing genes cloned into plasmids. Ethanologenic E. coli stra in LY180 ( E. coli W derivative; [ 97 ]) and its derivatives were used to investigate furfural tolerance for growth and ethanol production. A furfural resistant mutant of LY180 was previously isolated after 53 serial transfers of cultures in the presence of furfural and designated EMFR9 ( 97 ). A second strain, EMFR35, was subsequently isolated from the same enrichment culture after 107 transfers. Both mutants contained the same IS10 insertion in yqhC , silencing expression of yqhD ( 130 ). Strains and pTrc99A bas ed plasmids were constructed using Luria Bertani medium. After construction, cultures were grown in AM1 mineral salts medium ( 158 ). All media were supplemented with xylose (20 g /L for solid medium, 50 g /L for broth cultures, and 100 g /L for pH controlled f ermentations). Ampicillin (50 mg /L ) and isopropyl D thiogalactopyranoside (0.1 mM IPTG) were added as needed. Stocks of these were prepared in 70% ethanol (v/v) . Chromosomal deletion of puuP and potE were constructed using the method of Datsenko and Wann er ( 156 ). Constructions were confirmed by DNA sequencing and PCR analysis.


37 Furfural tolerance. Tolerance was examined by measuring growth and ethanol production after 48 h (37°C) using tube cultures (13 mm by 100 mm) containing 4 ml of AM1 medium. Ampicil lin (12.5 mg /L ), furfural, isopropyl D thiogalactopyranoside (IPTG) and other supplements were added as indicated. Inocula were grown overnight on AM1 xylose plates (solid medium). Cells from f resh colonies were resuspended in growth medium and adjusted to an OD 550nm of 1.0. Tube cultures were inoculated to an initial OD 550nm of 0.1 (43 mg dcw /L ) and incubated in a reciprocating water bath (50 oscillations min 1 ). Construction of polyamine transporter plasmids. Genes encoding PotE and PuuP were amplified (including ribosomal binding site and terminator region) from strain E. coli W (ATCC 9637) chromosomal DNA by using polymerase chain reaction (PCR). These fragments were cloned into the NcoI and BamHI sites of pT rc99A to produce pLOI5249 and pLOI5412 , respectively. Genes encoding six o ther polyamine transporters were also cloned in a similar manner using primers with flanking restriction sites (Table 2 1). After ligation, plasmids were transformed into E. coli TOP 10F . Plasmids were purified using a QIAspin Spin Miniprep kit (Qiagen Valencia, CA). Clones were verified by digestion with restriction enzymes, gel analysis of PCR products and sequencing. Data and analyses for tube experiments. Cell mass was measured as OD 550nm using a Bausch & Lomb Spectronic 70 spectrophotometer. An OD 550nm value of one is equivalent to 430 mg dcw / L . Ethanol was measured using an Agilent 6890N gas chromatograph (Palo Alto, CA) equipped with flame ionization detectors and a 15 m HP P lot Q Megabore column. Data presented are averages for three or more


38 experiments with standard deviations. Note that the presence of ampicillin and plasmids caused a small reduction in growth and ethanol production. Fermentations. The effect of furfural ad dition on pH controlled fermentations was investigated as described previously ( 97 ) using AM1 medium supplemented with 10 0 g /L xylose . Ethanol was measured by gas liquid chromatography. Cell mass was estimated from measurements of optical density at 550 nm . Furfural was measured using a Beckman Coulter DU 800 spectrophotometer ( 159 ). Results represent an average of three or more experiments with standard deviations. Genome Sequencing. Genomic DNA samples from LY180, EMFR9 and EMFR35 were purified according to the bacterial genomic DNA isolation protocol from the DOE Joint Genome Institute ( Next generation sequencing was performed using Illumina paired end short read technology provided by the Tufts University Core facility (Boston, MA). Se quence data for LY180 was assembled using Geneious software (Auckland, New Zealand). Sequencing of EMFR9 and EMFR35 has not been completed. Genome annotation of LY180 was provided by the Prokaryotic Genomes Automatic Annotation Pipeline (PGAAP) of the Nati onal Center for Biotechnology Information (NCBI). The fully assembled genome of LY180 has been deposited in NCBI GenBank ( under the accession number CP006584. Microarray analysis. Expression of mRNA was analyzed as previously described ( 97 , 1 30). For each data set, 4 cultures were grown separately in 500 ml fermenters (350 ml working volume) and sampled at an OD 550nm of 1.5 (650 mg dcw /L ) prior to furfural addition. Furfural was added at 5 mM for EMFR9 and LY180 con trol


39 and 15 mM for EMFR35 and LY180. Cultures were sampled again after 15 min of incubation with furfural. Each sample was processed separately for RNA extraction. RNA from each set of 4 samples was pooled and submitted to Roche NimbleGen for microarray a nalyses. Data from NimbleGen were imported into ArrayStar (DNA Star) for analysis. Each experiment was performed twice and averaged. Expression results for each gene are reported as the arithmetic ratio of mutant transcripts divided by the parent LY180. M icroarray data accession number. Microarray data for gene expression in LY180, EMFR9, and EMFR35 have been deposited in NCBI Gene Expression Omnibus ( with GEO series accession numbers GSE17786 and GSE46442. RT PCR Methods. RNA was isolated (Qiagen RNeasy Protect Bacteria Mini Kit) from cultures grown to OD 550nm of 1.5 (650 mg dcw /L ) as described in the previous section. Residual genomic DNA was removed by DNase treatment using the Ambion Turbo DNA free Kit (Life Tec hnologies, Grand Island, NY). Real time RT PCR was performed in 25 µL reactions, each containing 125 ng of RNA (Qiagen QuantiTect SYBR Green RT PCR Kit). Primer sequences for RT PCR analysis of polA , potE and puuP are listed in Table 2 2 . The number of cyc les to reach the threshold (C T value) was measured for each primer pair in triplicate. Using expression of polA gene as a reference for comparisons among strains, fold differences were calculated by dividing expression in the mutant by expression in the pa rent. Results represent an average of 3 reactions with standard deviations.


40 Results Isolation of furfural resistant mutants. Two furfural resistant mutants of ethanologenic LY180 were isolated after 53 and 107 serial transfers in minimal medium conta ining furfural, EMFR9 ( 97 ) and EMFR35, respectively. Furfural tolerance of these strains was compared by measuring growth and ethanol production in tube cultures (Fig ure 2 1A and 2 1B). For each strain, curves for growth and ethanol production exhibited si milar trends as a function of furfural concentration. The parent LY180 was the most sensitive to furfural. Furfural tolerance was highest for EMFR35, followed by EMFR9. For these strains, the minimal inhibitory concentration (MIC; Figure 2 1C) of furfural w as 20.0 mM (EMFR35), 16 mM (EMFR9) and 12.5 mM (LY180). Up regulation of polyamine transporters in furfural resistant mutants . Microarray studies of oxidoreductase encoding mRNA in EMFR9 previously identified silencing of yqhD (NADP dependent furfural reductase) as an important mutation for furfural tolerance ( 97 , 1 30). This mutation was present in both EMFR9 and EMFR35, consistent with a common ancestor. Further investigation of the EMFR9 microarray data revealed a 100 fold u p regulation of potE expression (putrescine uptake) relative to the parent LY180 (Table 2 3 ) , which was the largest increase among all the genes in this strain. Although E. coli contains 8 polyamine transporters encoded by 18 genes, only the expression of potE exhibited a large expression increase in EMFR9 relative to LY180 (Table 2 3 ). Up regulation of potE in EMFR9 was confirmed using RT PCR (27 fold up regulation) (Table 2 4 ). Addition of furfural did not affect potE expression (Table 2 3 ; microarray co mparison) in EMFR9 as compared to LY180 (parent). EMFR35 was isolated after 107 serial transfers in AM1 medium containing furfural, a continuation of the same enrichment culture used to isolate EMFR9 (53


41 transfers). Unlike EMFR9, expression of potE was no t up regulated in EMFR35 relative to the parent LY180 (microarray comparison). However, a different polyamine transporter ( puuP ) was expressed at 700 fold higher levels in EMFR35 than in the parent LY180 (Table 2 3 ). This unusually large up regulation of puuP in EMFR35 was confirmed by RT PCR using polA expression as a reference (Table 2 4 ). RT PCR measurement confirmed that puuP was highly expressed in EMFR35, over 1500 fold higher than in the parent LY180. Smaller changes were also noted in EMFR35 for t wo other polyamine transporters (Table 2 3 ; microarray comparisons), a 10 fold increase in plaP expression and a potential furfural induced increase in expression of the potABCD operon. Expression ratios for genes encoding other polyamine transporters in EMFR9 and EMFR35 were low with an average expression ratio of 1.1 relative to LY180. In each furfural resistant mutant (EMFR9 and EMFR35), a single polyamine transport gene exhib ited the largest increase in expression among all chromosomal genes, relative to the parent LY180. Plasmids containing polyamine transporters increase furfural tolerance in LY180 . The unusually high expression of a single polyamine transporter in each fur fural resistant mutant suggests that these transporters may contribute to furfural resistance. To test this hypothesis, all 8 polyamine transporters in E. coli were cloned into pTrc99A and transformed into LY180. LY180 containing empty vector served as a c ontrol. The largest difference in growth and ethanol production w as observed with 10 mM furfural for potE and puuP (Figure 2 2A, 2B, and 2 2C). At this furfural concentration, cell mass


42 and ethanol were 4 fold higher with pLOI5249 (PotE) and pLOI5412 (PuuP ) than with LY180 containing empty vector. Figure 2 2C shows a comparison of furfural tolerance (10 mM) for LY180 containing individual polyamine transporter plasmids. Four transporter plasmids more than doubled the growth of LY180 (empty vector control) in the presences of 10 mM furfural. Three of the beneficial transporters (PotE, PuuP and PlaP) are single gene proton symports for putrescine uptake ( 160 ). The fourth, potABCD , encodes an ATP dependent ABC transporter for spermidine and putrescine ( 160 ). T he four beneficial transporters increased growth relative to the empty vector control in the following order: PuuP > PotE > PlaP > PotABCD. Half of the transporter plasmids (PotFGHI, YdcSTUV, CadB, MdtJI) did not affect furfural tolerance. The two transpo rters that were highly expressed in furfural resistant mutants based on RT PCR and microarrays ( potE in EMFR9 and puuP in EMFR35) were also the most beneficial for furfural tolerance in LY180 when expressed from plasmids pLOI5249 and pLOI5412, respectively . With all polyamine transporters, l eaky expression of polyamine transporter genes from pTrc99A plasmids was sufficient to confer an incr ease in furfural tolerance (Figure 2 2C). Addition of IPTG decreased furfural tolerance with all transporter construct s ( Figure 2 2C), but caused a small increase in furfural tolerance in the empty vector control. Addition of IPTG decreased the growth of all strains carrying cloned transporter genes even in the absence of furfural ( Figure 2 8 ). Induction of membrane prote ins could lead to membrane destabilization . Similar detrimental effects with inducer have been observed for several E. coli genes including pntAB ( 98 ) and trehalose biosynthetic genes ( 161 ).


43 Deletion of chromosomal potE and puuP in LY180 increased furfural sensitivity . The importance of native puuP and potE genes was examined by constructing deletions in LY180, designated RG100 ( puuP ), RG101 ( potE ) and RG102 ( puuP, potE ). Furfural tolerance in these strains was compared by measuring cell yield a fter 48 h with 10 mM furfural ( Figure 2 3A). Deletion of puuP decreased cell yield by 70% (0.15 g dcw /L for RG100 as compared to 0.50 dcw /L for LY180). Deletion of potE alone also decreased furfural tolerance in comparison to LY180 but to a lesser extent. Deletions of both puuP and potE from LY180 decreased growth in 10 mM furfural by 80%, indicating that native expression levels of these genes contribute to furfural tol erance in the parent. Deleting potE in EMFR9 reduced furfural tolerance. The large increase in potE expression in EMFR9 (relative to LY180) was also presumed to contribute to furfural tolerance. Deletion of the potE coding region in EMFR9 to produce strain RG105 resulted in a substantial loss of furfural tolerance (Fig ure 2 3B and 2 3C). With 10 mM furfural, cell yield and ethanol production of RG105 were reduced by over 70% compared to EMFR9 . T he MIC for RG105 was reduced from 15 mM furfural to 12.5 mM fur fural. Addition of plasmid pLOI5249 encoding PotE restored furfural resistance in strain RG105 to near that of EMFR9 containing empty vector, confirming the importance of this gene. Gene duplication contributes to increased expression of puuP in EMFR35. At tempts to delete the puuP gene in EMFR35 were unsuccessful using the same genetic tools employed successfully with LY180 and EMFR9. Although the kanamycin gene was readily integrated, this integration did not remove the chromosomal puuP


44 gene. A persistent full length coding region for puuP remained in the chromosome. These observations suggest that multiple copies of puuP are present in the chromosome. M ultiple chromosomal copies of genes have previously been identified by examining chro mosomal coverage using DNA sequence reads ( 162 ). Illumina sequence reads for EMFR35 were mapped against LY1 80 using Geneious software (Figure 2 4). No mutation w as found in the primary sequence of puuP or in the nearby puuR region (repressor of puu genes ). However, this comparison of sequence reads revealed that an 8.8 kbp block of chromosomal DNA containing the puuP gene is duplicated at least 4.5 fold in comparison to flanking sequences. The repressor puuR was outside of the amplified region and appears t o be present as a single copy. Amplification of puuP without amplification of the putative repressor ( puuR ) may be responsible for the increase in expression observed in EMFR35 (Table 2 3 ). Beneficial tandem repeats are often selected under stress conditio ns and can collapse when environmental conditions improve. Tandem repeats are frequently associated with increased resistance to antibiotics (36) but could also contribute to furfural tolerance. No gene amplification w as observed in the potE region, plaP region, or potABCD region in furfural resistant mutants (EMFR9 and EMFR35). No specific repressor is known for these genes. The mechanism by which expression of potE is increased in EMFR9 and expression of plaP (10 fold) and potABC (2 to 3 fold) are increased in EMFR35 remain s unknown but may involve unidentified regulatory gene ( s ) . Reengineering furfural tolerance based on genetic traits in EMFR9 and EMFR35. Two genetic traits were identified in each of the furfural resistant mut ants


45 (EMFR9 and EMFR35), silencing of yqhD expression by a regulatory mutation ( 130 ) and up regulation of a polyamine transporter gene ( potE and puuP , respectively). Strains with analogous traits were constructed by transforming plasmids expressing polyami ne transporter genes into an LY180 derivative containing a yqhD deletion (XW092). Furfural tolerance in the resulting strains was compared to th e mutants EMFR9 and EMFR35 (Figure 2 5). LY180 containing empty vector (pTrc99A) was the most sensitive to furfu ral followed by X W092 (pTrc99A) (Figure 2 5A and 2 5B). Addition of the potE plasmid (pLOI5249) to XW092 increased furfural tolerance to near that found in EMFR9 containing pTrc99A indicating that both genetic traits are needed. Results with the puuP plasm id (pLOI5412) were simil ar (Figure 2 5C and 2 5D). Strain XW092 (pLOI5412) was also more resistant to furfural than LY180 (pTrc99A) and EMFR9 (pTrc99A). However, EMFR35 (pTrc99A) was more furfural tolerant than XW092 (pLOI5412) indicating that additional m utations are involved. Constructs with a single genetic trait for furfural tolerance were more resistant to furfural than the parent containing empty vector, and more sensitive to furfural inhibition than constructs carrying two traits. Figure 2 1C summari zes MIC values for different strains and constructs. Supplementing with agmantine, putrescine and cadaverine increased furfural tolerance. E. coli contains 3 polyamines (in order of abundance): putrescine, > spermidine, > cadaverine ( 164 ). Each of these was tested as a supplement for LY180 in AM1 medium containing 10 mM furfural (Fig ure 2 6A). Agmatine, a precursor of putrescine, was also included. All polyamines except spermidine caused a small increase in furfural tolerance. The addition of spermidine reduced furfural tolerance,


46 though only a small decrease in growth was observed when spermidine was added in the absence of furfural (Figure 2 9 ). Th e addition of 5 mM putrescine increased growth and ethanol production in AM1 medium containing furfural (Fig ure 2 6B and 2 6C). The combination of suboptimal levels of putrescine (1.0 mM) with transporter plasmids ( potE or puuP ) increased the growth of LY1 80 in the presence of 10 mM furfural (Figure 2 6D). In both cases, combinations were more effective than putrescine or transporter plasmid alone. The beneficial activity of plasmid borne polyamine transporters for furfural tolerance appears to be partially replaced by supplementing with polyamines. Batch fermentations with furfural. In the absence of furfural (AM1 medium containing 100 g /L xylose), growth and ethanol production began immediately with all LY180 constructs (Figure 2 7A and 2 7B). Xylose ferm entation was complete within 48 h. Addition of 10 mM furfural completely inhibited the growth and fermentation of LY180 (pTrc99A) for over 96 h. Furfural was partially metabolized to furfuryl alcohol (Fig ure 2 7C; 97 ) during the initial 24 h of incubation. Thereafter, the rate of furfural metabolism declined progressively. Less than half of the furfural was metabolized by LY180 (pTrc99A) after 96 h. In contrast, LY180 (pLOI5249; potE ) and LY180 (pLOI5412; puuP ) completed the fermentation of xylose (100 g /L ) in AM1 medium supplemented with 10 mM furfural within 96 h (Figure 2 7A, 2 7B, and 2 7C). Furfural addition (10 mM) caused an initial lag of 48 h during which furfural was fully metabolized. After this lag, rates of growth and ethanol production with LY180 (pLOI5249) and LY180 (pLOI5412) were similar to the LY180 (pTrc99A) control lacking furfural. Plasmid based expression of polylamine


47 transporters ( puuP and potE ) increased the metabolism of furfural, decreased the initial lag, and decreased the time required to complete xylose fermentation. Discussion Inhibitors such as furfural are formed during dilute acid pretreatment of lignocellulosic biomass ( 91, 98, 139 141 ). These pose a significant challenge for the fermentation of hemicellulose sugars int o fuels and chemicals ( 80, 151 ). Furfural resistant mutants of E. coli ( 98 99 ) and yeasts ( 165 ) have been described that overexpress oxidoreductases which reduce furfural to the less toxic furfuryl alcohol ( 93, 106 ). Furfural has been shown to cause single strand breaks and DNA mutagenesis, primarily in AT rich regions ( 152 ). Allen et al. ( 153 ) have reported reactive oxygen species (ROS), which are also known to damage DNA, proteins and phospholipid membranes, accumulate in yeast cells during growth in the presence of furfural. In E. coli , polyamines protect DNA from damage by oxidative stress agents ( 166 167 ). Overexpression of thyA, thymidylate synthase, has also been demonstrated to increase furfural resistance in E. coli ( 101 ) and may facilitate repair o f furfural damaged DNA. Many genes that are beneficial for furfural tolerance in E. coli ( fucO, pntAB, ucpA ; 144 ) directly or indirectly promote the NADH dependent reduction of furfural to the less toxic and less reactive compound, furfuryl alcohol ( 93, 106 ). Given the reactive nature of furfural, cellular damage may be ongoing until this compound has been fully metabolized. In our studies, furfural concentrations above 3 mM appear sufficient to prevent growth and metabolism in the presence or absence of furfural tolerance genes ( 97 101, 144 ). Plasmids with polyamine transporter genes appear to fu nction in a similar manner (Figure 2 7C). With 10 mM furfural and 100 g /L xylose, furfural metabolism by LY180 (empty vector control) slows progressively after the first 24 h and


48 does not reach completion even after 96 h. In contrast, LY180 containing either PotE or PuuP plasmids continues to metabolize furfural at a relatively constant rate until the process has been completed. Polyamines are essential for cell division ( 160 , 168 ). Concentrations are maintained within a specific range ( 169 ) and elevated during exponential growth ( 170 ). Polyamines bind anionic structures such as plasma membranes, ribosomes, DNA, etc. ( 167 , 171 ). This binding protects DNA from damage by some reactive compounds (41), regulates gene expression and modulates translational fidelity ( 169 , 172 ). Excess polyamines are toxic, displacing magnesium from r ibosomes and inhibiting protein synthesis ( 173 174 ). Our studies suggest that polyamine transporters ( such as PotABCD, PlaP, PotE and PuuP ) and polyamine supplements in the medium protect cellular processes from furfural damage and allow cells to complete the reduction of furfural to the less toxic furfuryl alcohol. Increased expression of polyamine transporters and polyamine supplements represent a new approach to increase furfural tolerance in E. coli and may prove useful with other organisms.


49 T able 2 1 . Bacterial strains, plasmids, and primers . Strain, plasmid, or primer Relevant characteristic(s) Reference or source E. coli strains E. coli W LY180 RG100 RG101 RG102 RG105 EMFR9 EMFR35 XW092 TOP10F Plasmids pLOI5249 pLOI5408 pLOI5409 pLOI5410 pLOI5411 pLOI5412 pLOI5414 pLOI5415 pKD4 pKD46 pCP20 pTrc99A Primers potABCD cloning potFGHI cloning ydcSTUV cloning plaP cloning potE cloning puuP cloning mdtJI cloning cadB cloning wild type frdBC ::( frg Zm celY Ec ) ldhA ::( frg Zm casAB Ko ) adhE ::( frg Zm estZ Pp FRT) ackA ::FRT rrlE ::( pdc adhA adhB FRT) mgsA ::FRT LY180 puuP LY180 potE LY180 puuP potE EMFR9 potE LY180 furfural resistant mutant ( yqhD silenced) LY180 furfural resistant mutant ( yqhD silenced) LY180 yqhD F [ lacI q Tn 10 (Tet r )] mcrA ( mrr hsdRMS mcrBC lacZ M15 lacX74 recA1 araD139 ( ara leu ) 7697 galU galK rpsL endA1 nupG potE gene in NcoI BamHI digested pTrc99A potABCD gene in AvaI XbaI digested pTrc99A potFGHI gene in EcoRI XbaI digested pTrc99A ydcSTUV gene in EcoRI XbaI digested pTrc99A plaP gene in EcoRI XbaI digested pTrc99A puuP gene in EcoRI XbaI digested pTrc99A mdtJI gene in EcoRI XbaI digested pTrc99A cadB gene in EcoRI XbaI digested pTrc99A bla FRT kan FRT (Red recombinase), temperature conditional replicon FLP + c I857 + p R Rep ts , bla , catF pTrc bla oriR rrnB lacI q For CCCCCGGGCAAGGTGGTTAACCACAAACC Rev GCTCTAGACGAATTGAAAATTAGCGTGTAA For CGGAATTCGTTAACGAACTTTCAGAAGGAA Rev GCTCTAGAATTTGTGTCAGCAGATATAGCCA For CGGAATTCGAACAATTAATTACGACAGGAGTAAG Rev GCTCTAGACAGCGGTTTGCCACAATTAC For CGGAATTCGCGACGGTTATCACCGTAAA Rev GCTCTAGATGCGATTATTTTTCGCGAGA For CATGCCATGGAACCTGTTGCCAGGTTTTGCA Rev CGGGATCCAGCTTCCTCGGTGAAGAACA For CGGAATTCCAAACCTTATTACGCAGGGGAG Rev GCTCTAGACATGTTGGGCTTCTTCGCTG For CGGAATTCACTTTGGTTTCGCTGAATTAAG Rev GCTCTAGAAGGCGGGATATCCTGAAGAT For CGGAATTCTGACCCGGACTCCAAATTCAA Rev GCTCTAGAACAACGGCAGGTTCTCGTTCA ATCC 9637 97 This study This study This study This study 97 E. Miller 144 Invitrogen (Carlsbad, CA) This study This study This study This study This study This study This study This study 156 156 157 Laboratory collection This study This study This study This study This study This study This study This study






52 Table 2 3 . Microarry expression ratios comparing the relative expression of polyamine transporter genes in furfural resistant mutants (EMFR9 and EMFR35) to the parent LY180 . Gene Expression Ratio (Mutant / LY180) a Transporter Gene EMFR9 (0 mM Furfural) EMFR9 (5 mM Furfural) EMFR35 (0 mM Furfural) EMFR35 (15 mM Furfural) ABC Transporter Spermidine/ Putrescine potA potB potC potD 0.8 1.1 0.9 0.8 1.0 1.3 1.1 1.3 0.8 0.7 0.5 1.2 2.2 3.0 3.1 1.0 ABC Transporter Putrescine potF potG potH potI 1.0 1.0 0.8 0.8 0.7 0.7 0.8 0.9 0.9 1.0 1.2 1.3 1.4 1.2 1.1 1.2 ABC Transporter Spermidine/ Putrescine ydcS ydcT ydcU ydcV 1.0 1.0 0.9 0.9 1.0 1.3 1.2 1.1 1.3 1.5 0.9 1.0 1.1 1.6 1.4 1.2 Putrescine Symporter plaP 2.4 1.9 1.7 10 Putrescine Symporter potE 120 100 1.0 0.8 Putrescine Symporter puuP 0.9 1.1 750 590 Spermidine Antiporter mdtJ mdtI 1.3 1.7 1.1 1.5 1.0 1.0 0.3 1.9 Cadaverine Symporter cadB 0.6 1.4 0.8 0.7 a Comparisons were made using total RNA from cells grown in the presence and absence of furfural. Table 2 4. mRNA levels of potE and puuP in EMFR9 and EMFR35 relative to LY180. Relative mRNA Level (Mutant / LY180)* Transporter Gene LY180 (0 mM Furfural) EMFR9 (0 mM Furfural) EMFR35 (0 mM Furfural) Putrescine Symporter potE 1.0 ± 0.0 27.2 ± 1.5 1.0 ± 0.1 Putrescine Symporter puuP 1. 0 ± 0.0 4.2 ± 0.3 1500 ± 197.0 * polA was used as a reference gene to normalize expression values for all strains.


53 Figure 2 1. Comparison of furfural tolerance in the parent LY180 and mutants isolated from serial transfers in AM1 containing furfural. EMFR9 was isolated after 53 serial transfers in AM1 medium containing xylose (100 g / L) and furfural. EMFR35 was isolated from a later stage of the same culture but after 107 serial transfers. A ) Growth in different concentrations of fu rfural; B ) Ethanol production in different concentrations of furfural; C ) Summary comparison of furfural tolerance (MIC) of different strains . Genetic traits conferring furfural tolerance included plasmid based expression of polyamine transporters potE and puuP (pLOI5249 and pLOI5412 , respectively) and deletion of oxidoreductase yqhD from the parent LY180, producing strain XW092. Constructs with a single genetic trait were more resistant to furfural than the parent containing empty vector, and more sensitive to furfural than constructs carrying two traits.


54 F igure 2 2. Effect of polyamine transporter plasmids on furfural tolerance in LY180. Plasmids containing potE (pLOI5249) or puuP (pLOI5412) increased the MIC for growth and ethanol production. LY180 containing empty vector (pTrc99a) ser ved as a control. A ) Growth in different concentrations of furfural; B ) Ethanol production in different concentrations of furfural; and C ) Growth of LY180 containing plasmids with each of the 8 polyamine transporters (10 mM furfural). Four of the 8 polyami ne transporters were beneficial for growth in the presence of 10 mM furfural (without addition of IPTG). Addition of 0.1 mM IPTG decreased furfural tolerance of all constructs containing polyamine transporters on plasmids. Note that IPTG caused a small inc rease in growth of LY180 containing empty vector (pTrc99A).


55 F igure 2 3. Effect of puuP and potE deletions on furfural tolerance. A ) Cell yield . Comparison of LY180 and derivatives with a deletion in puuP (strain RG100), potE (strain RG101), and both genes (strain RG102); B ) Cell yield ; and C ) Ethanol production. Comparison of EMFR9 and a mutant containing a potE deletion (RG105). Transporter deletions in the LY180 chromosome reduced furfural tolerance below that of the unmodified parent. Deletion of p otE in EMFR9 reduced furfural tolerance to that of the parent, LY180 (Fig ure 2 1, 2 3B. and 2 3C). Addition of pLOI5249 containing potE substantially restored furfural tolerance in strain RG105 carrying a potE deletion (Fig ure 2 3B and 2 3C).


56 F igure 2 4. Alignment of Illumina sequencing reads from EMFR35 on the puuP region of LY180 (parent). Sequence base pair numbers are shown at the bottom. Sequence coverage of the 8.8 kbp region containing puuP is increased 4.5 fold compared to puuR and flanking regions of the genome. Hatch marked numbers indicate average sequence reads for regions within arrows. Genes listed in the bottom have been modified to allow labeling and are not to scale.


57 F igure 2 5 . Reconstructing furfural tolerance in LY180 based on EMFR9 and EMFR35. A ) Growth of XW092 and EMFR9; B ) Ethanol production with XW092 and EMFR9; C ) Growth of XW092 and EMFR35; D ) Ethanol production with XW092 and EMFR35. Strain XW092 is a derivative of L Y180 with a deletion in yqhD, similar to the silencing of yqhD expression in EMFR9 and EMFR35. This strain was used as a host for plasmids expressing polyamine transporters. Plasmids pLOI5249 (PotE) and pLOI5412 (PuuP) were added and the resulting constructs compared to EMFR9 and EMFR35, respectively. Furfural resist ance of XW092(pLOI5249) was equivalent to that of EMFR9 containing empty vector indicating that these two mutations are sufficient to reconstitute furfural tolerance. In contrast,XW092(pLOI5412) was less resistant to furfural than EMFR35 containing empty v ector. Additional unknown mutations appear to be needed to fully reconstruct the level of furfural resistance present in EMFR35 .


58 F igure 2 6. Effect of polyamine supplements (1, 5 or 10 mM) on furfural (10 mM) tolerance in LY180. Cell yield after 48 h (A, B, and D) and ethanol production (C) were used as measures of furfural tolerance. A ) Polyamine supplements (1.0, 5.0 and 10.0 mM) added to AM1 medium containing furfural; B ) and C ) Effect of putrescine (5 mM) on furfural tolerance; and D ) C ombined effects of putrescine (1.0 mM) and a polyamine transporter plasmid (PotE, pLOI5249; PuuP, pLOI5412).


59 F igure 2 7 . Effect of polyamine transporter plasmids on the fermentation of strain LY180 in AM1 medium containing furfural (10 mM) and100 g /L xylose. A ) Growth; B ) Ethanol production C ) Furfural metabolism during fermentation.


60 F igure 2 8 . Effect of polyamine transporter expression on the growth of E.coli LY180 in the absence of furfural (48 h incubation). F igure 2 9. Effect of polyamine supplements on the growth of LY180 in the absence of furfural (48 h incubation).


61 CHAPTER 3 COMBINING TREATMENTS TO IMPROVE THE FERMENTATION OF SUGARCANE BAGASSE HYDROLYSATES BY ETHANOLOGENIC Escherichia coli LY180 Chemical and Process M ethods to Reduce Hydrolysate Toxicity Lignocellulosic biomass is a non food source of carbohydrates and is an attractive feedstock for biorefineries wanting to produce renewable fuels, solvents, plastics, and chemicals. Sources of lignocellulose include ag ricultural residues, municipal green waste, energy crops, wood processing waste, and others (175). Unlike starch , which is used as an energy storage polymer by green plants, lignocellulose is a structural component that has evolved to resist chemical and b iological deconstruction (176). Some form of pretreatment is essential for the effective enzymatic digestion of the cellulosic component (18, 76, 116, 177). Dilute acid hydrolysis at elevated temperatures is quite effective at increasing the available surf ace area for enzymatic hydrolysis of cellulose, while also hydrolyzing most hemicellulose s into monomeric sugars (18, 138, 178). Major bottlenecks in the conversion of lignocellulose to bioproducts include the generation of fermentation inhibitors during dilute acid pretreatment and the need for high cellulase enzyme loadings (79, 85, 139 140, 179). Inhibitory compounds formed in clude furans, organic acids, aldehydes, alcohols and phenolic compounds (79 82). Though increased pretreatment severity improves enzymatic digestibility, this method also increases the generation of inhibibitory side products that inhibit microbial activity and require detoxification (85, 139). Many methods have been reported for the mitigation of t oxins and improvement of biomass sugar fermentation and can be categorized broadly as biological, chemical and physical treatments (139, 151) . Saccharomyces cerevisiae NRRL Y 1536 and 424A


62 (LNH ST) fermentation of steam pretreated mixed har dwood was improved by the removal of phenolic compounds (180). Kim et al. (2011, 2013; [180 181]) determined that the removal of phenolic compounds through activated charcoal or ethyl acetate enhanced fermentation and relieved enzyme inhibition. Alkaline p H treatement of hemicellulose hydrolysate with calcium hydroxide (over liming), sodium hydroxide, potassium hydroxide or ammonium hydroxide has been shown to reduce toxicity of hydrolysates using yeasts and ethanologenic Escherichia coli (80, 92, 182) . In most cases, however, high pH treatment also resulted in significant sugar destruction (80, 92, 182) . Increasing the pH to pH 9.0 with ammonium hydroxide was demonstrated to decrease hydrolysate toxicity with minimal sugar loss (183 , 184) . In addition , seve ral reduced sulfur compounds have been investigated for their ability to improve growth and fermentation of hemicellulose hydrolysates (81, 185) . Sodium meta bisulfite (converts to two bisulfite molecules in aqueous solution) has also been shown to decreas e hydrolysate toxicity (81,185) . Some ethanologenic microorganisms have the native ability to partially detoxify hydrolysates (135 136, 186 187) . However, even strains engineered with inhibitor detoxifying genes require high levels of inoculum to be effective (79, 88, 187 189) . Non ethanologenic microorganisms also produce enzymes that metabolize some of the toxic chemicals in hydrolysate (135 136, 190 192) . White rot fungi (able to grow on decaying plant material) produce laccase enzymes that couple the oxidation of various phenolic compounds to the reduction of oxygen (186, 193 194) . Oxidation of phenolic compounds is thought to create free radicals that lead to polymerization,


63 thereby removing these toxic compounds from solution (186, 195) . Studies have shown improved fermentability of hydrolysates treated with laccase enzymes (186, 193 194) . Processes can be configured to include physical steps that reduce hydrolysate toxicity (196). Volatile inhibitory compounds can be removed from hydrolysate by evaporation under vacuum (85, 151, 196) . The resulting hydrolysate is more concentrated in sugar and non volatile inhibitors but has reduced levels of furfural, hydroxymethyl furfural, acetic acid and vanillin (85, 151, 196 197) . The addition of small amo unts of air to the culture or headspace has been shown to promote the fermentation of hemicellulose hydrolysates. Low levels of areatioin results in increased cell mass and ethanol productivity but reduce yields (198 201) . In this investigation, we have examined six methods that improve fermentation of dilute acid hydrolysates of hemicellulose individually and in combination. A combination of treatments was developed that fully eliminated toxicity for ethanologenic E. coli , allowing the fermentation of hemicellulose sugars at a rate equivalent to that of pure xylose in the same mineral salts medium. Materials and Methods Preparation of s ugarcane bagasse hydrolysate . Sugarcane bagasse was generously provided by Florida Cryst als Corporation (Okeelanta, F L ). Hemicellulose hydrolysates of sugarcane bagasse were prepared at the University of Florida Biofuels Pilot Plant. Bagasse was soaked for 4 h in a 14 fold excess of 0.5% phosphoric acid (w/w including moisture in bagasse), de watered to 50% moisture using a model CP 4 screw press (Vincent Corporation, Tampa, FL) and loaded into a steam pretreatment reactor designed by Dr. Guido Zacchi (202). Dilute acid impregnated bagasse was treated for 5 min at 190 ° C , as previously described (185) . Pretreated bagasse


64 contained 70% moisture (30% dry weight including fiber and solubles). Hemicellulose hydrolysate (syrup) was separated from pre treated bagasse by using a CP 4 screw press (Vincent Corporation, Tampa, FL). A glass fiber filter (Wh atman GF/D, 15 mm Clarified hydrolysates were diluted and used as sugar and nutrient sources during experiments that evaluated toxicity. Multiple batches of hydrolysate we re used during the course of this study. The average composition for 5 batches was: (g/L): xylose (48 ± 7), glucose (2.7 ± 0.7), arabinose (4.5 ± 0.5), galactose (2.9 ± 0.5), acetate (3.7 ± 1.6), furfural (1.6 ± 0.5), hydroxymethyl furfural (0.0) and total monomer sugars (58.4 ± 8.5). Strains, media, and growth conditions . Ethanologenic E. coli strain LY180 ( E. coli W derivative; [97]) was used in this study of hemicellose hydrolysate toxicity. LY180 contains a single set of chromosomally integrated genes from Zymomonas mobilis ( pdc , adhA , adhB ) for ethanol production. Key genes in competing pathways ( ldhA , adhE , ackA , frdBC , mgsA ) have been deleted. A cellulase gene ( celY ) from Erwinia chrysanthemi and cellobiose transport and utilization genes from Klebsiella oxytoca ( casABC ) have also been added to this strain. Cultures were grown in AM1 medium (158) containing 20 g/L xylose (plates) or 50 g/L xylose (broth). Optical density (OD 550nm ) measurements in hemicellulose hydrolysates are obscured by col or and color changes during fermentation, especially at higher concentrations of hydrolysate. With LY180 and other ethanologenic E. coli strains and low inocula, cell growth is essential for ethanol production. Both parameters follow similar trends (97 99, 101, 203). In this study, ethanol production was used as the primary measure of hydrolysate performance.


65 Toxicity of untreated and treated hydrolysates . Relative toxicity of hydrolysate was evaluated by measuring ethanol produced after 48 h (37°C) in tub e cultures (13 mm by 100 mm) containing 4 ml of AM1 medium (158) (50 g/L xylose) supplemented with various concentrations of hydrolysate (0% hydrolysate to 90% hydrolysate) containing the components of AM1 medium (ammonia neutralization, 1.5 mM MgSO 4 , 2 mM KCl, 1 mM betaine and AM1 trace metals; approximately 50 g/L total sugars). Inocula were grown overnight on AM1 xylose plates (solid medium). Cells from f resh colonies were re suspended in AM1 medium and adjusted to an OD 550nm of 1.0. Tube cultures wer e inoculated to an initial OD 550nm of 0.1 (43 mg dcw/L) and incubated in a reciprocating water bath (50 oscillations / min, 37°C). Since inocula and media components represented up to 10% of the volume, t he highest concentration of hydrolysate that could be tested was 90% . Hydrolysate adjusted to pH 6.3 with ammonium hydroxide served as a control for maximum toxicity . Relative toxicity was evaluated by determining the hydrolysate concentrations (treated and untreated control) that inhibited ethanol production by 50% (ICEt 50 ) and by 100% (MICEt 100 ). All tube cultures and tests were prepared as biological triplicates and all experiments were repeated at least twice. Hydrolysate treatments . Six approaches were evaluated for their effects on hydroly s ate toxicity a lone and in combination. For combinations, treatments were performed in the following order: 1) vacuum treatment; 2) laccase; 3) high pH treatment (HpH); 4) bisulfite addition; 5) increased inoculum; and 6) aeration. For high pH exposure ( HpH ) treatment , hydrolysate was adjusted to pH 9.0 by addition of ammonium hydroxide (5 N) and stored at room temperature overnight before inoculation


66 (16 h). Ammonia treated hydrolysate fell to approximately pH 7.5 during incubation (184). For bisulfite treatment, a fr eshly prepared solution was added to culture tubes immediately before inoculation (1 mM final concentration of bisulfite or 0.5 mM sodium metabisulfite). Hydrolysate adjusted to pH 6.3 immediately before inoculation served as the untreated control for all experiments. A Buchi Rotavapor R110 evaporator (Flawil Switzerland) equipped with a Cole Palmer aspirator pump Model 7049 00 (Chicago, Illinois) was used (55°C) to remove volatiles (vacuum treatment) prior to pH adjustment of hydrolysate. Unless otherwise indicated, hydrolysate was evaporated to 50% by weight and restored to original weight by adding deionized water before testing for toxicity. Laccase (Novozymes NS 22127) was generously provided by Novozymes North America, Inc. (Franklinton, North Carolin a). For laccase treatment, hydrolysate was adjusted to pH 5.0 with ammonium hydroxide (5 N). Laccase (5 U/ml) was added to hydrolysate (50 ml in 250 ml shake flasks) and incubated in a water bath (50°C, 150 rpm) for 3 h. A laccase unit, U, is defined as t he amount of laccase that catalyzes the 7.5, 30 °C ). When needed, aeration was provided in tube cultures using an incubator Fe rmentation. Fermentations were conducted in pH controlled vessels (500 ml) with a 300 ml working volume (37°C, 150 rpm; [184]). Cultures were maintained at pH 6.5 by the automatic addition of 2M potassium hydroxide. A modified AM1 medium (AM1 90% hydrolysa te containing approximately 50 g/L total sugar) was prepared using hemicellulose hydrolysate as the sole source of fermentable sugar. Trace metals,


67 MgSO 4 (1.5 mM final), KCl (2 mM final) and betaine (1 mM final) were added from 1000X stock solutions (158). Hydrolysates were treated in various ways as indicated to reduce toxicity. Seed cultures were grown overnight (16 h) in AM1 medium supplemented with 50 g/L xylose. Fermentations were initiated by inoculating to an initial OD 550nm of 0.1 (43 mg dcw/L) and monitored for up to 120 h. For cultures requiring low level aeration ( micro aeration ) , subsurface sparging (0.01 vvm, 3 m l /min) was provided by using a peristaltic pump. Chemical analysis. Sugars, furans, and organic acids were analyzed using two Agilent T echnologies 1200 series HPLC systems, one equipped with a BioRad (Hercules, CA) Aminex HPX 87P ion exclusion column and the other equipped with a BioRad Aminex HPX 87H column (184). Ethanol was measured using an Agilent Technologies 6890 N Network gas chro matography system with a wide bore HP PLOT Q column (J&W Scientific, Folsom, CA; [184]). Total f urans (furfural and hydroxymethyl furfural) were measured by ultraviolet absorption (159) and by HPLC. Moisture content was determined using a Kern model MLB 5 0 3 moisture analyzer (Balingen, Germany). Statistical analysis. Prism software (Graphpad, San Diego, CA) was used to perform two way ANOVA (analysis of variance) comparisons of data. Differences were judged significant when P values for the null hypothesi s were 0.05. Results and Discussion Ethanol production (MICEt 100 and ICEt 50 ) as a measure of hydrolysate toxicity . A standardized procedure was developed to compare hydrolysate treatments for toxicity using culture tubes containing dilutions of hydrolysate in AM1 mineral salts medium (50 g/L xylose) and a low inoculum. Hydrolysate adjusted to pH 6.3 with ammonium hydro xide immediately before inoculation served as the control, exhibiting


68 maximum toxicity. In previous studies (88, 93, 97 98, 106), OD 550nm , MIC and IC 50 were used as metrics. However, these measurements of optical density in hydrolysate medium are complicat ed by dark color, color changes during incubation, and precipitation of lignin in some cases. Since the shapes of curves for cell mass during growth and cumulative ethanol production are similar in AM1 medium without hydrolysate (Figure 3 1A), analogous me trics for this study were b ased on ethanol production: 1) hydrolysate concentration at which ethanol production was completely inhibited (MICEt 100 ); and 2) 50% inhibition of ethanol production (ICEt 50 ). Figure 3 1B illustrates the graphical estimation of MICEt 100 and ICEt 50 using control hydrolysate (adjusted to pH 6.3 and inoculated) and hydrolysate after HpH as examples. The higher MICEt 100 and ICEt 50 values after HpH exposure indicate a reduction in hydrolysate toxicity. Single and combination treat ments of hydrolysate . Many treatments have been described previously that reduce hydrolysate toxicity for yeasts (186 189, 198 199, 201) and alcohol producing bacteria (92, 196, 200). Six of these treatments were compared for their ability to reduce the t oxicity of dilute acid hydrolysates of sugarcane bagasse (Figure 3 2A). When tested individually, HpH treatment was the most beneficial for reduction of hydrolysate toxicity. With HpH treatment, MICEt 100 and ICEt 50 concentrations of hydrolysate were increa sed by at least 5 fold over the control, allowing fermentation (ethanol production) in 70% hydrolysate. Previous studies have shown that high pH treatments are also effective using sodium hydroxide, potassium hydroxide, and calcium hydroxide (92, 182). Unl ike previous reports, however, HpH treatment using ammonia caused very little sugar destruction (Table 3 1).


69 Addition of 1 mM bisulfite had the second largest benefit when tested alone, increasing MICEt 100 from 15% hydrolysate (control) to 25% hydrolysate and ICEt 50 from 4.8% hydrolysate (control) to 16.3% hydrolysate (3 fold). When tested alone, addition of air, increasing inoculum size by 4 fold, and removal of volatiles with vacuum each caused only a small decrease in toxicity. Laccase treatment alone p rovided little benefit for fermentation as a single treatment. All ICEt 50 values for treated hydrolysates were significantly higher than the untreated hydrolysate control. All MICEt 100 values were significantly higher than the control except for laccase a lone and air alone. Combination treatments were investigated using HpH, bisulfite, laccase and vacuum evaporation (Figure 3 2B). All were beneficial in comparison to the control (Figure 3 2A). Vacuum treatment with bisulfite was the most effective binary c ombination, increasing the MICEt 100 by over 2 fold and the ICEt 50 by over 4 fold as compared to control (Figure 3 2A). Neither bisulfite with laccase nor vacuum with laccase increased the MICEt 100 or the ICEt 50 more than bisulfite alone (Figure 3 2A). The beneficial effect of laccase was observed only for combinations that included prior vacuum treatment. The combination of laccase, vacuum and bisulfite was significantly better than any binary combination (p<0.05). Vacuum treatment appears to remove volatil e inhibitors that mask the benefit of laccase. None of the combinations tested without HpH exposure (Figure 3 2B) were as effective as the HpH alone (Figure 3 2A). Binary combinations that included HpH treatment (Figure 3 2B) were all significantly less toxic than HpH, as indicated by high ICEt 50 values and MICEt 100 values at or above 90% hydrolysate. The combination of


70 vacuum and HpH treatment was more effective than the ternary combination of laccase, HpH, and bisulfite. In most cases, each added treatm ent caused an incremental decrease in toxicity (increase in ICEt 50 concentration), consistent with the presence of multiple inhibitors and mechanisms of action. Laccase was ineffective except when combined with prior vacuum treatment or with HpH treatment. Hydrolysate exposed to H pH combined with three other treatments (vacuum, laccase, and bisulfite) had the highest metrics and lowest toxicity, an ICEt 50 of over 80% hydrolysate and a MICEt 100 of greater than 90% hydrolysate. Reducing the extent of vacuum evaporation . Half the weight of hydrolysate was evaporated in the initial vacuum evaporation experiments. With this treatment, all furfural and 25% of the acetate were removed (Figure 3 3A; Table 3 1). However, less evaporation may be desirable for some ap plications. Further tests were conducted to determine the extent of furfural removal at different stages of evaporation (Figure 3 3A). The initial furfural concentration (20 mM) was reduced by 90% after only 10% evaporation of hydrolysate. Furfural was com pletely removed after 20% or more evaporation (Figure 3 3A). The effect of different vacuum treatments on toxicity was also examined. After each vacuum treatment, deionized water was added to replace lost weight and the hydrolysate was exposed to HpH (Fig ure 3 3A). Toxicity was measured as ethanol production in treated hydrolysate diluted with an equal volume of AM1 medium (50 g/L xylose) with and without bisulfite. This hydrolysate dilution (50%) partially inhibited ethanol production. Ethanol production was increased by more than 3 fold after 10% evaporation (results with and without bisulfite), concurrent with the removal of 90% of


71 the furfural. A further increase in ethanol production over 15% was observed with the complete removal of furfural by 20% e vaporation. This small increase in ethanol production may also reflect removal of less volatile compounds such as phenolics and acetic acid (Figure 3 3A). In all cases, ethanol production was lower without bisulfite, consistent with independent modes of ac tion. The concurrent reduction of toxicity and removal of furfural by vacuum evaporation suggests that furfural is an important inhibitor in hydrolysate. However, unquantified compounds of equal or greater importance may also be removed by evaporation. E xa mining the effect of adding 20 mM furfural to vacuum treated hydrolysate (50% evaporation) tested this possibility (Figure 3 3B). Restoring the initial level of furfural to vacuum treated hydrolysate precisely restored the toxicity for ethanol production, establishing furfural as the dominant inhibitor in HpH treated hydrolysate (with and without bisulfite treatment). These results provide further evidence that furfural plays a central role as an inhibitor of growth and fermentation in dilute acid hydrolysa tes. Effect of selected treatments on sugar and inhibitor concentrations . Sugar and inhibitor concentrations were measured in hydrolysates after various combinations of treatments (Table 3 1). To facilitate comparisons, all hydrolysates were diluted to 90 % with AM1 medium analogous to the medi um used in tube fermentations. Vacuum treatment (50% evaporation) of hydrolysate reduced furfural to undetectable amounts and also reduced acetate concentrations by 25% compared to the control (pH 6.3). When vacuum ev aporation was combined with laccase and bisulfite treatment, no further reductions in acetate were observed. All treatment combinations conducted


72 without HpH caused very little sugar loss (<1%). HpH treatment reduced furfural concentrations by 36% while ac etate concentrations remained unchanged. A s mall amount of sugar w as lost during HpH treatment, approximately 0.6 g/L. However, this sugar loss with ammonia based HpH (pH 9) was much lower than previously reported for high pH treatments using lime ( 10% 2 0% of total sugar; [ 92, 183 ] ). No further loss of sugar was observed when HpH was combined with vacuum evaporation, laccase or bisulfite treatments. Increasing inoculum size and aeration improved ethanol production . All hydrolysates exposed to HpH (alone or in combinations) were less toxic than untreated hydrolysate (Figure 3 2A and 3B). Two of the least toxic combinations were used to examine the effects of inoculum size and aeration: 1) vacuum evaporation + HpH + bisulfite ; and 2) vacuum evaporation + la ccase + HpH + bisulfite (Figure 3 4A and 4C). Increasing inoculum size by up to 4 fold that of the standard inoculum resulted in a small incremental increase in ethanol production. A similar increase in ethanol production with hydrolysate and high inocul um has been reported for yeasts (187 189). Increased inoculum size may increase the rate of inhibitor metabolism. However, a threshold of toxicity must be overcome before this increase in inocula can be effective (79, 88). Addition of air (Figure 3 4B and 4 D) also provided an incremental benefit, increasing ethanol production in 90% hydrolysate by over 2 fold for hydrolysates treated with all four methods (vacuum evaporation + laccase + HpH + bisulfite ). Air promotes an increase in cell mass, energy producti on, and allows redox balance, factors that may minimize damage from toxins in hydrolysate (198 201). Aeration of the most highly -


73 treated hydrolysate produced more ethanol (17 g/L) than tube cultures of LY180 with laboratory AM1 medium containing 50 g/L xyl ose without hydrolysate (8.5 g/L ethanol; Figure 3 1A). Fermentation of hydrolysate (90%) in pH controlled vessels . The effect of the various hydrolysate treatments w as further investigated using pH controlled fermenters. LY180 grown in AM1 minimal medium with 50 g/L xylose was able to completely utilize xylose within 36 h (Figure 3 5A), achieving a corresponding maximum ethanol titer of 23 g/L (Figure 3 5B). In contrast , sugar utilization and ethanol production was completely inhibited in 90 % hydrolysate even after HpH exposure, with or without bisulfite. However, vacuum evaporation combined with HpH treatment allowed fermentation in 90% hydrolysate to proceed, producin g 20 g/L ethanol after 120 h with nearly complete sugar utilization. Further improvements in productivity were observed when bisulfite and laccase were also included (vacuum+HpH+bisulfite and vacuum+laccase+HpH+bisulfite), producing 21 g/L ethanol in 120 h and 96 h, respectively. Lower final ethanol titers with hydrolysate are due to lower sugar content (90% hydrolysate). Micro aeration (0.01 vvm) dramatically improved fermentation of treated hydrolysate ( vacuum+lacc a se+ HpH+bisulfite). With this combined a pproach, fermentation was completed within 36 h, equivalent to the fermentation of laboratory xylose in AM1 medium without hydrolysate. All sugars present in the sugarcane bagasse hemicellulose hydrolysate were completely utilized (Figure 3 5C). This combi nation of treatments with micro aeration fully eliminated the toxicity of hemicellulose hydrolysate for ethanologenic E. coli LY180.


74 Operating costs associated with treatment options. H emicellulose hydrolysate treatment strategies were examined for their c ost effectiveness (Table 3 2) . The six treatments evaluated were selected because they were believed to be relatively inexpensive options for the detoxification of hydrolysate. Bisulfite addition was calculated to be the least expensive treatment ( less tha n 1 cent per gallon of ethanol produced ) . High pH treatment using a mmonia to produc e the required ammonium hydroxide was another relatively inexpensive treatment option (5 cents per gallon of ethanol). A mmonia addition also serves as a nitrogen source for the ethanologenic strain LY180. Laccase was the most expensive option analyzed resulting in a cost of almost $15 per gallon of ethanol . At this cost exogenous addition of laccase would be impractical . However, our experiments clearly show ed th at laccas e was an effective treatment for the detoxification of hydrolysate. Expression of a functional laccase enzyme directly from the microbial biocatalyst c ould eliminate any cost associated with laccase addtion. Concentrating hydrolysate by v acuum evaporatio n (20% 50%) was very effective at furfural removal (Figure 3 3A). Using a triple effect evaporator to remove 20% hydrolysate (by weight) was calculated to cost $0.38 per gallon of ethanol. However t he cost of 20% vacuum evaporation could be lowered or es sentially eliminated through the recovery of wasted heat energy in a biorefinery. The h eat energy supplied to the pretreatment hydroly z er alone is over 3 times that required during vacuum evaporation. Process integration aimed at recovering heat energy thr oughout a biorefinery c ould be employed to offset the cost of running the evaporation unit. Also, there are potential savings associated with concentrating the hydrolysate sugars which


75 would lead to higher ethanol titers and reduced energy costs during dis tillation and recovery of the product. The other strategies used for hydrolysate detoxification were l ow level aeration and increased inoculum. Micro aeration costs would be primariy a ssociated with capital expense s for piping . High agitation associated with aerobic fermentations ; w ould not apply in the case of anaerobic fermentatioins supplied with a low level of aeration. L ow level aeration (0.01vvm) c ould be supplied to fermenter s by the same compressor system that actuates pneumatic valves. A process that combin ed treatment strategies was able to fully eliminate hydrolysate toxicity (Figure 3 5). Provided process improvements can be accomplished through laccase expression from the biocatalyst and by heat energy and process integration in a biorefienry , a c ombin ation of all six treatments could have associated costs of as little as $0.06 per gallon of ethanol (current selling price of $2.00 per gallon).


76 Table 3 1. Sugar and inhibitor concentrations after hydrolysate treatments . Sugars (g/L) Inhibitors (g/L) Treatment Glucose Xylose Galactose Arabinose Total Sugars Furfural Acetate Control 3.2 43.8 3.1 4.1 54.2 2.0 5.1 V 3.2 43.8 3.0 4.1 54.1 ND 3.7 V, B 3.2 43.6 3.0 4.1 53.9 ND 3.7 V, L, B 3.2 43.6 3.0 4.0 53.9 ND 3.7 HpH 3.2 43.4 3.0 4.1 53.6 1.3 5.1 V, HpH 3.2 43.4 3.0 4.0 53.5 ND 3.7 V, HpH, B 3.2 43.4 3.0 4.0 53.5 ND 3.7 V, L, HpH, B 3.2 43.3 3.0 4.0 53.5 ND 3.7 Hydrolysat e a 2.7 ± 0.7 48.0 ± 7.0 2.9 ± 0.5 4.5 ± 0.5 58.4 ± 8.5 1.6 ± 0.5 3.7 ± 1.6 V, vacuum evaporation ND, not detectable B, bisulfite L, laccase HPH, high pH treatment a a verage from 5 batches of hydrolysate


77 Table 3 2 . Estimated costs for hydrolysate treatments. Treatment Chemical or Process Energy or Materials Cost ( Source ) Estimated Cost ($/Gallon EtOH) V a vacuum evapor ation $ 0.0 821/kWh e 0.38 L b l accase e nzyme $ 30.00 /kg f 14.85 HpH c a mmonia $ 0.60 /kg g 0. 05 B d s odium b isulfite $ 0.66 /kg h 0.008 a vacuum evaporation to remove 20% hydrolysate ( by weight) including all furfural b laccase (5 U/ml) c high pH treatment (pH increase from 6.3 to 9.0 with ammonia ) d bisulfite (1 mM) e source US Energy Information Administration f source Alibaba g source ICIS h source ICIS


78 F igure 3 1. Metrics used to evaluate toxicity in treated hydrolysates. A ) G rowth and ethanol production in AM1 medium (50 g/L xylose). B ) Examples of hydrolysate plots illustrating the estimation of MICEt 100 (hydrolysate concentration that completely blocks ethanol production) and ICEt 50 (hydrolysates concentration that causes a 50% inhibition of ethanol production). These metrics are based on ethanol production rather than OD 550nm due to color in the hydrol ysate, changes in color during incubation, and precipitation. Ethanol production was measured after 48 h. Two curves are shown as examples: untreated hydrolysate adjusted to pH 6.3 e ( ).


79 F igure 3 2. Effect of chemical and process treatments on ethanol production in hemicellulose hydrolsates using LY180 (MICEt 50 and MICEt 100 values). A ) Single hydrolysate treatments; B ) Combination of hydrolysate treatments. The open triangle above bars indicates that the MICEt 100 is greater than 90%, the highest concentration which can be tested.


80 F igure 3 3 . Effect of vacuum evaporation on furfural removal and fermentation of hemicellulose hydrolysat e (50% dilution): A ) Correlation between furfural removal and ethanol production by LY180 (48 h, 50% hydrolysate; HpH treatment with or without bisulfite) in tube cultures; and B ) LY180 fermentation (48 h, 50% hydrolysate, HpH exposure, with and without bisulfite) with and without vacuum treatment. Furfural concentrations were restored to original levels in 50% hydrolysate (10 mM) to investigate its effect on restoring toxicity .


81 F igure 3 4 . Fermentation of treated hydrolysate with LY180. Combinations of increasing inoculum size, and aeration were tested using two combination treatments of hemicellulose hydrolysates (90%). Figure A and B (vacuum, HpH, and bisulfite treatments); figure C and D (vacuum, laccase, HpH, and bisulfite treatments). No air was supplied to A and C . Micro aeration (0.01 vvm) was provided to B and D.


82 F igure 3 5 . Fermentation of 90% hydrolysates by LY180. A ) Sugar utilization, B ) Ethanol production, C ) Utilization of individual sugars after V, L, HpH, B, Air treatment. Abbreviations for hydrolysate treatments: V, vacuum evaporation; L, laccase; HpH, high pH treatment; B, bisulfite; and Air, micro aeration (0.01 vvm).


83 CHAPTE R 4 CONCLUSIONS Fossil fuels are a finite resource, distributed uneaqually across the world. Countries depend heavily on fossil fuels for their energy needs as well as for production of many of the commodities in everyday use. Agricultural and forest resid ues are much more widely available to countries and regioins. Improved utilization of renewable feedstocks such as lignocellulosic biomass can allow more sustainable production of valuable fuels and chemicals. However challenges remain in depolymerizing li gnocellulosic biomass into fermentable sugars . Among the main challenges are the production of inhibitory compounds during biomass pretreatment that retard fermentation, the need for high loading of costly cellulase enzymes and contamination which affects fermentation performance. Reducing inhibitor concentration or toxicity has a direct benefit to the biocatalyst during the fermentation process. However, indirect benefits are also realized. A more robust biocatalyst that is able to outperform other organisms in inhibitor rich environments can decrease the chance of contamination. Indeed studies have indicated that contaminati on is a significant problem , especially in cases where lowered cellulose enzyme loading lead to lengthy hydrolysis times for the release of cellulose sugars ( 205, 206 ) . Another indirect benefit is that by reducing inhibitor concentration or toxicity a more severe pretreatment can be employed that increases accessibility to carbohydrates while maintaining fermentability of the resulting hydrolysate . This result s in lowered enzyme doses necessary for hydrolys is of cellulose sugars. Overexpression of polyamine transporters w as shown to improve performance of an ethanologenic E. coli biocatalyst in the presence of a major inhibitory compound


84 (furfural) found in hydrolysate. Of the 8 transporter s in E . coli known to transport polyamines, improveme n ts could be obs evered when 4 of these were overexpressed . The largest improvement w as achieved by expressing potE and puuP . Supplementing the fermentation medium with polyamine compounds also allowed small improvements in growth and ethanol production. Studies have previ ously described deletion of an oxidoreductase gene yq h D, associated with the conversion of furfural to the less toxic furfural alcoho l , as causing increased furfural toleran ce in the evolved furfural tolerant strain EMFR9 ( 97 98 , 130 ). The data presented in this study showed that the furfural tolerance phenotype of the evolved strain (EMFR9) could be fully reconstructed by just two genetic traits (deletion of yqhD and overexpression of the polyamine transporter potE ). Expression of polya mine transporters and the addition of polyamine compounds provide a novel approach to furfural tolerance by protecting DNA, cell membranes and other cellular components from the damaging effects of furfural. As a complimentary strategy to increased biocata lyst tolerance for improved hydrolysate fermentability a number of chemical and physical detoxification treatments were also investigated. H igh pH treatment , carried out by adjusting the pH of hemicellulose hydrolysate up to 9.0 using ammonium hydroxide an d allowing it to settle to pH 7.5 after 16 hours was most effective treatment for improving fermentability. O ther types of high pH treatments have been described previously such as overlimi n g with calcium hydroxide ( 9 2 ) . However, these procedures generate large quantities of byproducts and have been shown to cause s ugar loss of up to 10% ( 92 ) . A 35 % reduction in furfural concentration was observed using ammonium hydroxide high pH treatment with only a 1% corresponding loss of sugar. The importance of furfu ral


85 concentration in hydrolysates was underscored by an experiment in which it was remov ed during vacuum evaporation an d an equivalent amount of furfural was added back to the sample. By doing so t oxicity was fully restored, indicating that though other inhibitors are removed by vacuum treatment , furfural was sufficient to cause the inhibition observed. E ach treatment was effective either alone or in combination at providing a benefit to fermentation . C ombinations of treatments resulted in incremental inc reases in fermentability of hydrolysate . Th ese results reflect multiple inhibitor types with m ultiple mod es of toxicity being present in hydrolysate . A process combining h igh pH with other simple treatmnets (vacuum evaporation, laccase, bisulf i te) was eff ective at allowing fermentation of 90% hydrolysate ( diluted only by nutrients a dded for fermentation ) . With t he addition of air (sparged at 0. 0 1 vvm ) to this combination , ethanol was produc ed at the same rate as a fermentation in AM1 mineral salts medium without any added hydrolysate , i ndicat ing toxicity had been fu lly eliminated . Improved biocatalyst tolerance and hydrolysate detoxification strategies wer e both employed successfully to i ncrease the fermentability of hemicellulose hydrolysate . These stra tegies ultimately serve to reduce the complexity of processing lignocellulosic biomass by allowing sugars released during pretreatment to be combined with those released during enzymatic hydrolysis and co fermented to valu e added produc t s. Hemicellulose h ydrolysate is a complex mixture of sugars with a diverse set of inhibitory compounds. The various treatments investigated in ths study allow hydrolysates with different inhibitor profiles to be generated. F uture studies targeting


86 specific inhibitor types f or the discovery of genetic traits able to confer further biocatalyst tolerance are possible after select treatments .


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105 BIOGRAPHICAL SKETCH Ryan Geddes was born in Lionel Town, Clarendon, Jamaica in 1980. He attended St. Thomas Moore Preparatory School and later Glenmuir Hi gh School in May Pen, Clarendon where he graduated in summer 1998. He moved to the United States in spring 1999 to pursue a Bachelor of Science in Chemical Engineering degree at the er 2003. While studying at UF, he met his wife Claudia Geddes and together they have a son Alessandro. After graduating, Ryan joined Myriant, an industrial biotechnology company that produces renewable platform chemicals, working as a Technology Transfer E ngineer. In August 2010 he returned to the University of Florida to join Dr. Lonnie doctoral degree in December 2014. He currently resides in Raleigh, North Carolina with his wife and son and continues his work in the field of industrial biotechnology as a scientist with Novozymes a world leader in industrial enzyme production.