Process development and metabolic engineering to enhance 2,3- butanediol production by Paenibacillus polymyxa DSM 365
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Christopher Chukwudi Okonkwo
Graduate Program in Animal Sciences
The Ohio State University
2017
Dissertation Committee:
Thaddeus C. Ezeji, Advisor
Ramesh Selvaraj
Katrina Cornish
Ana Alonso
Copyrighted by
Christopher Chukwudi Okonkwo
2017
Abstract
2,3-Butanediol (2,3-BD) is a platform chemical with vast industrial applications; particularly for its use in the production of 1,3-butadiene (1,3-BD), the monomer from which synthetic rubber is manufactured. Currently, 2,3-BD production is by chemical synthesis using petroleum-derived feedstocks such as propylene, acetylene, butene and butane. Microbial 2,3-BD fermentation is aimed at producing 2,3-BD renewably, and potentially reduce dependency on finite petroleum-derived feedstocks. However, fermentative production of 2,3-BD is hampered by (1) cost of food-based substrates; (2) low 2,3-BD titer, yield and productivity during 2,3-BD fermentation stemming from formation of competing products such as exopolysaccharides (EPS), ethanol, lactic, formic and acetic acids; and (3) high cost of 2,3-BD purification, due partly to additional purification steps necessary to remove 2,3-BD co-products especially EPS prior to 2,3-
BD recovery. The objectives of this study were conceived to examine use of process design, alternative substrates and metabolic engineering, to enhance 2,3-BD production.
Chapter 3 (objective 1) focused on identification of key fermentation parameters that influence 2,3-BD fermentation by Paenibacillus polymyxa and optimization of them for maximum 2,3-BD production. The study examined the impact of yeast extract, tryptone, ammonium acetate, ammonium sulfate, and crude glycerol concentration, and inoculum size and fermentation temperature on 2,3-BD production by P. polymyxa. The
ii results showed that only three parameters (tryptone, temperature and inoculum size) had significant effects on 2,3-BD production by P. polymyxa. The three factors were optimized and 2,3-BD production by P. polymyxa increased from ~27 g/L to 51.1 g/L in batch bioreactor cultures and from 47 g/L to 68.5 g/L in fed-batch cultures. The improvement in 2,3-BD production by P. polymyxa was accompanied by 11% and 19% reduction in ethanol and EPS formation, respectively, when compared to the un- optimized fermentation medium and conditions. Due to the inability of P. polymyxa to produce more than 6% 2,3-BD in fed-batch cultures, and the attendant increase in the accumulation of acetoin (the precursor from which 2,3-BD is biosynthesized) in the bioreactor, chapter 4 (objective 2) focused on understanding 2,3-BD-mediated feedback inhibition during 2,3-BD fermentation. This study evaluated the response of P. polymyxa to high 2,3-BD concentrations during growth and 2,3-BD fermentation. Cultures of P. polymyxa were challenged with levo-2,3-BD (20, 40 and 60 g/L) at 0 h fermentation in a glucose medium. The inhibition of P. polymyxa growth by levo-2,3-BD was concentration dependent, triggering total growth inhibition when the concentration of
2,3-BD attained 60 g/L. Furthermore, when P. polymyxa was challenged with incremental 2,3-BD concentrations (20, 40 and 60 g/L at 12, 24 and 36 h, respectively) to mimic 2,3-BD accumulation during fermentation, 2,3-BD was reconverted to acetoin when its concentration reached 60 g/L, possibly to alleviate 2,3-BD toxicity.
Chapter 5 (objective 3) evaluated the feasibility of using readily available non- food lignocellulosic biomass (LB) as substrate for 2,3-BD fermentation. Pretreatment of
LB to release fermentable sugars is accompanied by the generation of lignocellulose-
iii derived microbial inhibitory compounds (LDMICs) such as furfural, hydroxymethyl- furfural (HMF), and phenolic compounds which inhibit growth and pose a significant roadblock to LB use as substrates. The study investigated the ability of P. polymyxa to use LB-based agricultural residue, wheat straw hydrolysate (WSH), for the production of
2,3-BD. Prior to testing the fermentability of WSH to 2,3-BD, the ability of P. polymyxa to co-metabolize the representative mixed sugars (glucose, xylose and arabinose) of WSH was evaluated. The results show that P. polymyxa simultaneously co-metabolized the mixed sugars (glucose, xylose and arabinose) component of LB to 2,3-BD without exhibiting signs of carbon catabolite repression characteristics. Batch fermentations conducted using 60%, 80%, and 100% WSH, and a glucose-based control, showed that the growth of P. polymyxa increased 17%, 27% and 32% in 60%, 80% and 100% WSH, respectively, relative to the glucose control medium. 2,3-BD production in 60%, 80% and
100% WSH was 32, 31 and 23 g/L, respectively, which was comparable to the 32 g/L obtained in the glucose-based control. The enhanced growth in WSH suggests that P. polymyxa might have sequestered additional carbon from LDMICs. Hence, the ability of
P. polymyxa to use LDMICs as sole carbon sources was investigated. The growth of P. polymyxa in HMF increased 2.4-fold relative to the control with no carbon source which suggested that P. polymyxa might have utilized HMF in WSH for cell biomass accumulation. In addition, P. polymyxa showed robust tolerance to furfural and phenolic compounds (coumaric acid, vanillic acid and vanillin) during fermentation.
Chapter 6 (objective 4) explored a metabolic engineering strategy to deactivate the EPS production pathway of P. polymyxa and drastically reduce or eliminate EPS
iv production during 2,3-BD fermentation. The study identified a levansucrase gene which encodes levansucrase, the enzyme responsible for EPS biosynthesis in P. polymyxa. The results showed that the levansucrase gene was successfully disrupted, and the resulting P. polymyxa levansucrase null mutant showed 34% and 54% increases in growth in sucrose and glucose media, respectively. Additionally, the P. polymyxa levansucrase null mutant grown in sucrose and glucose media produced 6.4- and 2.4-fold lower EPS, respectively, than that produced by the P. polymyxa wildtype. The observed decrease in EPS formation by the levansucrase null mutant may be a direct cause of the 4-27% increase in 2,3-BD yield, and 4-128% increase in 2,3-BD productivity observed during 2,3-BD fermentation.
Interestingly, the levansucrase null mutant remained genetically stable over fifty generations with no observable decrease in growth, 2,3-BD and EPS formation.
Collectively, our results show that P. polymyxa levansucrase null mutant has potential for improving the economics of large-scale microbial 2,3-BD production.
v
Acknowledgments
I am deeply indebted to my advisor Dr. Thaddeus Ezeji for his support, guidance and encouragement throughout the course of my graduate program. I would like to thank
Dr. Ezeji specially for providing me the privilege and opportunity to conduct research under his mentorship. Dr. Ezeji offered me impeccable support through difficult times, research wise and offered me a research assistantship for over three years, which allowed me grow as a scientist. The training I received under his tutelage and mentorship has transformed me.
I am eternally grateful to Dr. Victor Ujor for guiding me through good laboratory practices, scientific writing and for his invaluable advice and suggestions towards overcoming experimental challenges during the course of my graduate program. I also wish to thank him for meticulously reading the first draft of my PhD dissertation. I wish to thank a friend and colleague, Dr. Chidozie Agu for introducing me to the laboratory during the first few weeks of my graduate program and for being a wonderful team mate.
I wish to thank members of my committee; Dr. Ramesh Selvaraj, Dr. Katrina
Cornish and Dr. Ana Alonso for agreeing to serve on my committee and for their invaluable advice and constructive comments that led to the completion of this research. I am honored to have you all in my committee. I appreciate all past and present graduate students, faculty members and staff in the Department of Animals Sciences, The Ohio vi
State University. I specially thank the Department of Animal Sciences for the invaluable contribution to the research funding that made my research work possible. I thank
OARDC Graduate Research Enhancement Competitive Grants Program (OARDC
SEEDS Grant) for research funding.
I greatly appreciate Drs. Gabriel and Jane Okafor for sharing information leading to this research opportunity and for their encouragement and good will. I wish to express my special gratitude to my friends, Mary Mbah, Mrs. Dana Ujor, Drs. Segun and Foluke
Awe, Gloria Okpala and Sarah Emereonye for their moral support and encouragement. I would like to express my gratitude to Prof. Obioma Njoku, Prof. Lawrence Ezeanyika,
Prof. Edwin Alumanah and Prof. Ferdinand Chilaka for their moral support.
My heartfelt gratitude goes to my parents, Mr. and Mrs. Christopher C. Okonkwo for their love and support. I also wish to thank my siblings for their prayers and moral support. Above all, my profound gratitude goes to almighty God for giving me the grace and endurance to complete this research.
vii
Vita
2008…………………………………………B.Sc. Biochemistry, University of Nigeria
2012………………………………………... M.Sc. Biochemistry, University of Nigeria
2013 to 2017 ...... Graduate Research Associate, Department
of Animal Science, The Ohio State
University
Publications
Okonkwo CC, Ujor V, Mishra, P, and Ezeji TC (2017) Process development for enhanced 2,3-butanediol production by Paenibacillus polymyxa DSM 365. Fermentation
(Accepted).
Okonkwo CC, Ujor CV, and Ezeji TC (2017) Investigation of relationships between 2,3- butanediol toxicity and production during growth of Paenibacillus polymyxa. New
Biotechnology, 34:23-31
viii
Okonkwo CC, Azam, MM, Ezeji TC, and Qureshi N (2016) Enhancing ethanol production from cellulosic sugars using Scheffersomyces (Pichia) stipitis. Bioprocess and
Biosystems Engineering, 39(7): 1023-1032
Ujor V, Okonkwo C, and Ezeji TC (2016) Unorthodox methods for enhancing solvent production in solventogenic Clostridium species. Applied Microbiology and
Biotechnology, 100: 1089-1099
Fields of Study
Major Field: Animal Sciences
Area of Specialization: Applied microbiology–bioproducts, biofuels, bioassay
development, and metabolic engineering
ix
Table of Contents
Abstract ...... ii
Acknowledgments...... vi
Vita ...... viii
Publications ...... viii
Fields of Study ...... ix
List of Tables…………………………………………………………………………..xviii
List of Figures…………………………………………………………………………...xxi
Chapter 1: Introduction ...... 1
References ...... 5
Chapter 2: Literature review ...... 8
2.1 Introduction ...... 8
2.2 Brief history of microbial 2,3-BD production ...... 10
2.3 2,3-Butanediol production methods ...... 11
2.4 Major 2,3-BD-producing microorganisms...... 12
2.5 Process development for enhanced 2,3-BD production ...... 14
2.6 Types of EPS and structures ...... 18 x
2.7 Ecological relevance of EPS biosynthesis ...... 20
2.8 Relationship between 2,3-BD production and biosynthesis of EPS ...... 21
2.9 Molecular genetics of EPS production ...... 23
2.10 EPS mitigation strategies: manipulations of nutrients and fermentation conditions 24
2.10.1 Oxygen availability and agitation ...... 25
2.10.2 Osmotic stress ...... 26
2.10.3 Culture pH ...... 27
2.10.4 Fermentation temperature ...... 28
2.10.5 Nitrogen sources ...... 28
2.11 EPS mitigation strategies: potential for strain development ...... 30
2.12 Challenges of microbial 2,3-BD production from lignocellulosic biomass 40
2.13 Microbial 2,3-BD recovery ...... 45
2.14 Simultaneous 2,3-butanediol fermentation and recovery ...... 48
2.15 Conclusions and perspectives...... 49
References ...... 50
Chapter 3: Process development for enhanced 2,3-butanediol production by
Paenibacillus polymyxa DSM 365 ...... 84
3.1 Abstract ...... 84
3.2 Introduction ...... 85
xi
3.3 Materials and methods ...... 88
3.3.1 Experimental methods ...... 88
3.3.1.1 Microorganism and culture preparation ...... 88
3.3.1.2 Batch and fed-batch fermentations ...... 90
3.4 Analytical methods ...... 91
3.5 Experimental design and data analysis ...... 92
3.5.1 Plackett-Burman design ...... 92
3.5.2 Path of steepest ascent ...... 94
3.5.3 Box-Behnken design and response surface methodology ...... 94
3.6 Results and discussion ...... 95
3.6.1 Plackett-Burman design ...... 95
3.6.2 Path of steepest ascent design ...... 98
3.6.3 Box-Behnken design and response surface methodology ...... 99
3.6.4 Experimental validation of the optimized medium and conditions in batch and fed- batch fermentations ...... 102
3.7 Conclusions ...... 105
References ...... 106
Chapter 4: Investigation of relationship between 2,3-butandiol toxicity and production during growth of Paenibacillus polymyxa ...... 120
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4.1 Abstract ...... 120
4.2 Introduction ...... 121
4.3 Materials and methods ...... 123
4.3.1 Microorganisms and culture conditions ...... 123
4.3.2 Fed-batch fermentations ...... 124
4.3.3 Batch fermentation, and levo- and meso-2,3-BD toxicity bioassay ...... 125
4.3.4 Analytical methods ...... 126
4.4 Statistical analysis and calculations ...... 128
4.5 Results ...... 128
4.5.1 Production of 2,3-BD by P. polymyxa in fed-batch cultures ...... 128
4.5.2 Tolerance of P. polymyxa to levo-2,3-BD during 2,3-BD fermentation ...... 129
4.5.3 Levo-2,3-BD supplementation alters the ratio of levo- to meso-2,3-BD produced by P. polymyxa ...... 132
4.6 Discussion ...... 134
4.7 Conclusions ...... 139
References ...... 140
Chapter 5: 2,3-Butanediol production from lignocellulosic biomass: Impact of microbial inhibitors on Paenibacillus polymyxa growth and 2,3-butanediol production ...... 152
xiii
5.1 Abstract ...... 152
5.2 Introduction ...... 153
5.3 Materials and methods ...... 155
5.3.1 Microorganism and culture conditions ...... 155
5.3.2 Pretreatment of Wheat straw ...... 156
5.3.3 Enzymatic hydrolysis of WS slurry……………… ...... 157
5.3.4 Pure individual and mixed sugar fermentations ...... 158
5.3.5 Fermentation of wheat straw hydrolysate ...... 158
5.3.6 Growth of P. polymyxa on pure LDMICs as sole carbon sources ...... 159
5.4 Analytical methods ...... 160
5.5 Statistical methods...... 161
5.6 Results ...... 161
5.6.1 Pretreatment and hydrolysis of wheat straw biomass ...... 161
5.6.2 2,3-BD production in pure individual and mixed sugars ...... 162
5.6.3 P. polymyxa growth and 2,3-BD production in wheat straw hydrolysate ...... 164
5.6.4 Using LDMICs as sole carbon sources ...... 165
5.7 Discussions ...... 166
5.8 Conclusions ...... 172
References ...... 173
xiv
Chapter 6: Molecular inactivation of exopolysaccharide biosynthesis in
Paenibacillus polymyxa DSM 365 for enhanced 2,3-butanediol accumulation ...... 189
6.1 Abstract ...... 189
6.2 Introduction ...... 190
6.3 Materials and Methods ...... 193
6.3.1 Microorganisms and culture conditions ...... 193
6.3.2 Genomic DNA extraction ...... 193
6.3.3 PCR amplification to generate levansucrase inactivation constructs ...... 195
6.3.4 Construction of recombinant plasmid ...... 197
6.3.5 Restriction digestion ...... 197
6.3.6 DNA ligation ...... 197
6.3.7 Preparation of E. coli JM 109 competent cells ...... 198
6.3.8 Transformation of competent E. coli JM 109 cells ...... 199
6.3.9 Preparation and electroporation of competent P. polymyxa protoplasts ...... 200
6.3.10 Characterization of P. polymyxa levansucrase null mutant ...... 202
6.3.11 Analytical methods ...... 202
6.3.12 Levansucrase assay ...... 204
6.3.13 Growth rate and generation time of levansucrase null mutant ...... 205
6.3.14 Stability of P. polymyxa levansucrase null mutant ...... 206
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6.3.15 Statistical analysis and calculations ...... 207
6.4 Results ...... 208
6.4.1 Inactivation of levansucrase gene in P. polymyxa DSM 365...... 208
6.4.2 Effect of levansucrase disruption on EPS formation ...... 209
6.4.3 Effect of calcium chloride supplementation on growth, sugar utilization, 2,3-BD yield and productivity ...... 210
6.4.4 Stability of levansucrase null mutant ...... 213
6.5 Discussion ...... 215
6.5.1 Effect of levansucrase inactivation on the growth of P. polymyxa levansucrase null mutant ...... 216
6.5.2 Effect of levansucrase inactivation on EPS biosynthesis by P. polymyxa levansucrase null mutant ...... 218
6.5.3 Effect of CaCl2 supplementation on sugar utilization, 2,3-BD yield and productivity by P. polymyxa levansucrase null mutant ...... 221
6.5.4 Stability of P. polymyxa levansucrase null mutant ...... 224
6.6 Conclusions ...... 225
References ...... 225
Chapter 7: Conclusions and recommendations ...... 255
7.1 Conclusions ...... 255
7.2 Recommendations ...... 260 xvi
References ...... 263
Bibliography ...... 264
Apendix A: One factor-at-a-time experiments ...... 300
Apendix B: CaCO3 supplementation in wildtype and levansucrase null mutant cultures ...... 209
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List of Tables
Table 2.1. EPS production by major 2,3-BD producers...... 75
Table 2.2. Comparative techniques for 2,3-BD recovery and purification ...... 77
Table 3.1. Statistical analysis of Plackett-Burman design results showing effect of medium components and fermentation conditions on 2,3-BD production by P. polymyxa
...... 111
Table 3.2. The path of steepest ascent experimental design and 2,3-BD production by P. polymyxa ...... 112
Table 3.3. Box-Behnken design and response results for 2,3-BD production ...... 113
Table 3.4. ANOVA for 2,3-BD production by P. polymyxa according to the response surface quadratic model (lack of fit is not significant) ...... 114
Table 3.5. The product profiles of P. polymyxa grown in batch and fed-batch fermentations under optimized conditions ...... 115
Table 3.6. Comparison of 2,3-BD concentrations obtained in this study to those of other studies using P. polymyxa ...... 116
Table 4.1. Yields, productivities, and sugar utilization in cultures of P. polymyxa DSM
365 challenged with levo-2,3-BD ...... 144
Table 5.1. Saccharification profile during enzymatic hydrolysis of acid-pretreated wheat straw ...... 178 xviii
Table 5.2. Lignocellulose-derived microbial inhibitory compounds (LDMICs) detected in wheat straw hydrolysate ...... 179
Table 5.3. Comparison of growth, 2,3-BD concentration, yield, productivity, and maximum ethanol and acetoin concentrations in individual and mixed sugar cultures . 180
Table 5.4. Comparison of growth, 2,3-BD concentration, yield and productivity, and ethanol and acetoin concentrations in WSH and glucose control ...... 181
Table 5.5. Sugar utilization profiles of P. polymyxa in WSH ...... 182
Table 6.1. Comparison of protein sequence between P. polymyxa DSM 365 levansucrase and other P. polymyxa strains with complete genome sequence using NCBI Blastp algorithm for alignment ...... 233
Table 6.2. Comparison of nucleotide sequence between P. polymyxa DSM 365 levansucrase and other P. polymyxa strains with complete genome sequence using NCBI
Blastn algorithm for alignment ...... 234
Table 6.3. List of primers and PCR strategies used to generate levansucrase inactivation construct ...... 235
Table 6.4. List of microorganisms, vectors and enzymes used in this study and their respective characteristics and sources ...... 237
Table 6.5. Substrate consumed, growth, maximum products, 2,3-BD yield and productivity during sucrose fermentation by P. polymyxa DSM 365 wildtype and levansucrase null mutant ...... 238
xix
Table 6.6. Substrate consumed, growth, maximum products, 2,3-BD yield and productivity during glucose fermentation by P. polymyxa DSM 365 wildtype and levansucrase null mutant ...... 239
Table 6.7. Comparison of fermentations with and without erythromycin supplementation during stability test of P. polymyxa levansucrase null mutant ...... 240
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List of Figures
Figure 2.1. Comparison of 2,3-BD produced by wildtype and serrawettin mutant strains of Serratia marcescens ...... 79
Figure 2.2. Schematic representation of proposed EPS biosynthesis in P. polymyxa from different substrate sources ...... 80
Figure 2.3. Characterization of EPS according to net charges ...... 81
Figure 2.4. Chemical structures of exopolysaccharides ...... 82
Figure 2.5. Fermentation of LB-derived sugars to 2,3-BD ...... 83
Figure 3.1. Contour and response surface plots ...... 117
Figure 3.2. Contour and response surface plots ...... 118
Figure 3.3. The fermentation profiles of P. polymyxa using optimized culture medium and conditions ...... 119
Figure 4.1. Fed-batch 2,3-BD fermentation by P. polymyxa ...... 146
Figure 4.2. Tolerance of P. polymyxa to levo-2,3-BD challenge at 0 h ...... 147
Figure 4.3. Tolerance of P. polymyxa to levo-2,3-BD pulse-fed at 12, 24, and 36 h .... 148
Figure 4.4. The profiles of levo- and meso-2,3-BD in cultures of P. polymyxa following levo-2,3-BD challenge (0 h-addition and pulse-feeding) ...... 149
Figure 4.5. Relative inhibitory effects of levo- and meso-2,3-BD on the growth of P. polymyxa ...... 150 xxi
Figure 4.6. Schematic 2,3-BD feedback inhibition representation during 2,3-BD fermentation in P. polymyxa DSM 365...... 151
Figure 5.1. HPLC chromatogram of WSH sugar analyses before and after enzymatic hydrolysis ...... 183
Figure 5.2. HPLC chromatogram of WSH sugar with several LDMIC peaks after wheat straw pretreatment and hydrolysis ...... 184
Figure 5.3. Production of 2,3-BD in single and mixed sugar media ...... 185
Figure 5.4. Sugar utilization profile of P. polymyxa in single and mixed sugar fermentations...... 186
Figure 5.5. Fermentation profile of P. polymyxa in WSH ...... 187
Figure 5.6. Use of LDMICs as sole carbon sources by P. polymyxa ...... 188
Figure 6.1. Levansucrase inactivation construct generation ...... 241
Figure 6.2. Gel image showing colony PCR of P. polymyxa levansucrase null mutant 242
Figure 6.3. The schematic representation of step-by-step procedure employed to evaluate the stability of the P. polymyxa levansucrase null mutant ...... 243
Figure 6.4. Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in sucrose cultures without CaCl2 supplementation...... 244
Figure 6.5. Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in sucrose cultures with 0.2 g/L CaCl2 supplementation ...... 245
Figure 6.6. Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in sucrose cultures with 0.4 g/L CaCl2 supplementation ...... 246
xxii
Figure 6.7. Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in glucose cultures without CaCl2 supplementation ...... 247
Figure 6.8. Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in glucose cultures with 0.2 g/L CaCl2 supplementation ...... 248
Figure 6.9. Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in glucose cultures with 0.4 g/L CaCl2 supplementation ...... 249
Figure 6.10. Stability test: product profile in sucrose-based medium and 0.4 g/L CaCl2 with 35µg/ml erythromycin supplementation ...... 250
Figure 6.11. Stability test: product profile in sucrose-based medium and 0.4 g/L CaCl2 without erythromycin supplementation ...... 251
Figure 6.12. Agarose (1%) gel images od stability test colony-PCR of P. polymyxa levansucrase null mutant ...... 252
Figure 6.13. Replica plates showing genetic stability test of P. polymyxa levansucrase null mutant ...... 253
Figure 6.14. Determination of generation (doubling time) of P. polymyxa levansucrase null mutant ...... 254
Figure A.1. One-factor at a time experiments showing effect of crude glycerol on 2,3-BD production by P. polymyxa ...... 301
Figure A.2. One-factor at a time experiments showing effect of temperature on 2,3-BD production by P. polymyxa ...... 302
Figure A.3. One-factor at a time experiments showing effect of inoculum size on 2,3-BD production by P. polymyxa ...... 303
xxiii
Figure A.4. One-factor at a time experiments showing effect of ammonium sulfate on
2,3-BD production by P. polymyxa...... 304
Figure A.5. One-factor at a time experiments showing effect of ammonium acetate on
2,3-BD production by P. polymyxa...... 305
Figure A.6. One-factor at a time experiments showing effect of tryptone on 2,3-BD production by P. polymyxa ...... 306
Figure A.7. One-factor at a time experiments showing effect of yeast extract on 2,3-BD production by P. polymyxa ...... 307
Figure A.8. Competing products generated during 2,3-BD fermentation in un-optimized medium and conditions ...... 308
Figure B.1. Fermentation profile of levansucrase null mutant in glucose medium supplemented with 4 g/L CaCO3...... 310
Figure B.2. Fermentation profile of levansucrase null mutant in glucose medium supplemented with 4 g/L CaCO3...... 311
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Chapter 1: Introduction
The compound, 2,3-Butanediol (2,3-BD), is a feedstock chemical with numerous industrial applications including its potential use in the synthesis of 1,3-butadiene (1,3-
BD), the monomer of synthetic butadiene-based rubbers. Further, 2,3-BD may be used as an additive in aviation fuel. Other 2,3-BD uses include synthesis of methyl ethyl ketone
(MEK), a high energy fuel additive, and as solvent in the manufacture of printing inks, resins and lacquers (Celinska and Grajek, 2009). Additionally, 2,3-BD is used as antifreeze due to its low freezing point of -60 °C (Soltys et al., 2001). Owing to its high reactivity, 2,3-BD can be converted to diacetyl, a flavor enhancing compound used in the food processing industry and in the manufacture of pharmaceutical and personal care products (Bartowsky and Henschke, 2004; Celinska and Grajek, 2009). The price of 2,3-
BD is estimated at ~USD 7.89/lb and the market is worth more than USD 975 million annually (Transparent Market Research, 2012). The global market for the downstream products of 2,3-BD and MEK, is estimated at USD 2.56 billion and is expected to reach
USD 3.64 billion in 2020, while the 1,3-BD market is estimated to reach USD 33.01 billion by 2020 (Grand View Research, 2015; 2016; Global Market Insights, 2016).
At present, 2,3-BD is produced at commercial scale through cracking of petroleum-derived hydrocarbons. However, instabilities in the price of crude oil together with its finite nature have re-awakened interest in alternative methods for producing 1 petrochemical-derived products, including 2,3-BD. 2,3-BD can be produced from sugars and lignocellulosic biomass (LB) by microbial fermentation. Microbial production of 2,3-
BD using sugars and LB hydrolysates as substrates is undermined by co-generation of interfering products, which divert carbon away from 2,3-BD biosynthesis, and complicate
2,3-BD recovery (down-stream processing) from spent broth. The co-products typically generated during 2,3-BD fermentation include exopolysaccharides (EPS), ethanol, acetoin, lactic, formic and acetic acids. Consequently, carbon diversion from 2,3-BD to these co-products results in decreased 2,3-BD titer, yield and productivity. Moreover, accumulation of competing products significantly impacts the cost of 2,3-BD recovery due to additional down-stream processing steps. Further, ethanol, lactic and formic acids are inhibitory to 2,3-BD-producing microorganisms, thus, their accumulation in the fermentation broth retards the growth of fermenting microorganisms and 2,3-BD biosynthesis. Additionally, the majority of known 2,3-BD producers are pathogens, hence their use in large-scale industrial fermentation is inherently unsafe (Ji et al., 2011;
Celinska and Grajek, 2009). Collectively, aforementioned challenges impede the commercialization of 2,3-BD production by fermentation.
Biological production of 2,3-BD in the US favors the use of non-pathogenic microorganisms. This criterion makes P. polymyxa, a non-pathogenic 2,3-BD producer, an attractive candidate for industrial-scale 2,3-BD production. However, 2,3-BD production by P. polymyxa is also plagued by low yield and productivity, largely due to diversion of carbon to competing products such as EPS, lactic acid, ethanol and formic acid. On a positive note, P. polymyxa can utilize a wide range of sugars such as glucose,
2 xylose and arabinose (the major sugars derived from lignocellulosic biomass) for the production of 2,3-BD. Notably, lignocellulosic biomass is a source of relatively cheap renewable substrates that have shown promise in reducing the overall production cost of biofuels and chemicals. More importantly, P. polymyxa has the ability to produce 98% of the optically active levorotary isomer of 2,3-BD (Nakashimada et al., 2000; Yu et al.,
2011). Economically, the levorotary isomer is more desirable than the mesorotary and the dextrorotary isomers due to its special applications in asymmetric synthesis of valuable chiral specialty chemicals, of which 1,3-BD is the most prominent (Celinska and Grajek,
2009; Häßler et al., 2012; Li et al., 2013). These attributes make P. polymyxa a preferred microorganism for industrial-scale 2,3-BD fermentation.
P. polymyxa was known as Bacillus polymyxa until 1993 when Ash et al. (1993) established the genus Paenibacillus to accommodate the former ‘group 3’ species of the genus Bacillus that comprises over 30 species of facultative anaerobic, endospore- forming, neutrophilic, peri-flagellated, heterotrophic, low G+C Gram positive bacilli
(Ash et al., 1991). According to the 16S rRNA gene sequence analysis, P. polymyxa is closely related to the group 3 bacilli, which represents a phylogenetically distinct group exhibiting high intragroup sequence relatedness, and is slightly related to B. subtilis (Ash et al., 1991; Lal and Tabacchioni, 2009).
Despite the great potential of P. polymyxa for industrial 2,3-BD production, only one study has been conducted on metabolic engineering of P. polymyxa for efficient 2,3-
BD production. This is due to limited availability of genetic information on P. polymyxa.
In a study by Zhang et al. (2016), the authors successfully introduced an exogenous
3
NAD+-dependent formate dehydrogenase (FDH) gene from Candida boidinii into P. polymyxa ZJ-9 to regenerate NADH (NAD+→NADH) for 2,3-BD formation. FDH introduction increased intracellular NADH/NAD+ ratio in recombinant P. polymyxa, which manifested in increased 2,3-BD production; increasing from 33.4 g/L to 36.8 g/L and 47.5 g/L to 51.3 g/L in batch and fed-batch cultures, respectively.
The goal of this work is to develop a hyper-2,3-BD-generating bioprocess using a combination of statistical design process and metabolic engineering approaches. This goal will be encompassed in two major objectives of the study, which are 1) to understand 2,3-BD fermentation and improve the process for enhanced 2,3-BD production and 2) to metabolically engineer P. polymyxa DSM 365 to minimize or eliminate EPS production during 2,3-BD fermentation, and maximize 2,3-BD production.
Exclusion of EPS from the process would improve the economics of 2,3-BD fermentation as majority of carbons used for the production of EPS would most likely be channeled to the 2,3-BD biosynthesis pathway and improve 2,3-BD yield. To accomplish these objectives, the work plan employed a multi-pronged approach that involved: 1) use of statistical models to optimize medium constituents (nutrients) and conditions during 2,3-
BD fermentation; 2) determination of P. polymyxa tolerance to 2,3-BD during 2,3-BD fermentation; 3) evaluate the feasibility of using readily available non-food lignocellulosic biomass (LB) as substrate for 2,3-BD fermentation; and 4) development of genetic tools to metabolically engineer P. polymyxa. While there have been studies conducted on nutrient manipulations to reduce competing co-products formation during
2,3-BD fermentation, there has never been any report of successful deletion or
4 inactivation of genes in P. polymyxa to reduce co-product formation during 2,3-BD fermentation. Therefore, this research work combines process development with the use of metabolic engineering to reduce the formation of undesirable EPS during 2,3-BD fermentation and enhance 2,3-BD titer, yield and productivity.
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Häßler, T, Schieder, D, Pfaller, R, Faulstich, M, Sieber, V (2012) Enhanced fed-batch
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(R, R)-2,3-butanediol production by Paenibacillus polymyxa ATCC12321. Appl
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7
Chapter 2: Literature review
2.1 Introduction
2,3-Butanediol (2,3-BD) has broad industrial applications; the most prominent is its potential use for the synthesis of 1,3- butadiene (1,3-BD), the monomer of synthetic butadiene rubber, currently produced by cracking of petroleum. In addition to its potential use as a precursor of this synthetic rubber monomer, 2,3-BD can be used as aviation fuel additive due to its high octane rating and antifreeze property (freezing point is -60 °C)
(Garg and Jain, 1995). The energy content of 2,3-BD is 27,189 J/g, which is comparable to established biofuels such as ethanol (29,055 J/g), methanol (22,081 J/g) and butanol
(32,010 J/g) (Flickinger, 1980; Szwaja and Naber, 2010), an attribute that positions 2,3-
BD as a viable fuel in light of rising global energy demand. Further, a number of 2,3-BD derivatives have huge industrial applications in the feedstock chemical sector. For instance, diacetyl, the partial dehydration product of 2,3-BD, is used as a flavoring agent in food industries and methyl ethyl ketone (MEK), the condensation product of 2,3-BD, is used for the manufacture of some types of adhesives, printing inks and lubricating oils
(Villet, 1981).
Due to the global need for synthetic rubber, the demand for 1,3-BD is about 9 million metric tons per year and this volume is expected to increase by 3% yearly (White,
2007). To meet this demand and ensure stability in the availability of 1,3-BD feedstock, 8 there is a critical need to establish an industrial-scale 2,3-BD fermentation process to reduce over-dependency on petroleum-derived 1,3-BD. To transition from the current laboratory scale fermentative production of 2,3-BD to commercial scale, it is imperative to increase the productivity and yield of 2,3-BD to levels that make economic sense.
Towards this goal, research efforts currently focus on exploiting metabolic perturbations in 2,3-BD producing microorganisms during fermentation along with use of metabolic engineering strategies to increase utilization of cheap substrates such as lignocellulose- derived sugars (LDS) and non-food based substrates for efficient 2,3-BD production.
Fermentative 2,3-BD production is accompanied by competing products that reduce 2,3-BD yield and productivity. Microbial producers of 2,3-BD intrinsically co- produce acetic, lactic, and formic acids, ethanol and EPS alongside 2,3-BD (Häßler et al., 2012; Guo et al., 2010; Cho et al., 2012; Zhang et al, 2010b). Consequently, carbon substrates are diverted away from the 2,3-BD pathway, which adversely affects 2,3-BD yield and productivity. Of all the competing products generated during 2,3-BD formation,
EPS is the most predominant, accounting for about 20% of the total product output
(Häßler et al., 2012). In addition to diminishing 2,3-BD yield and productivity, EPS increases viscosity of the fermentation broth and slows down oxygen transfer during fermentation, clogs bioreactor lines, and decreases mixing efficiency in the bioreactor
(Seviour et al., 2011; Humphrey, 1998). Additionally, EPS reduces the efficiency of 2,3-
BD recovery from the spent broth after fermentation. To rid the fermentation broth of
EPS during downstream processing additional purification steps/processes are required, which increase the overall cost of 2,3-BD production (Häßler et al., 2012).
9
Although the production of EPS during 2,3-BD fermentation and the associated problems have received little attention, the majority of 2,3-BD-producing microorganisms are known to synthesize significant amounts of EPS during growth and
2,3-BD production. As a result, there are limited data on the amounts of EPS generated during 2,3-BD fermentation and the biochemical interplay between EPS formation and
2,3-BD production. To increase 2,3-BD yield and productivity to a scalable titer, and improve the economics of the process, it might be necessary to manipulate 2,3-BD- producing microorganisms, fermentation media and culture conditions to abolish or significantly reduce EPS production as well as accumulation of other competing products such as ethanol, lactic and acetic acids. Here, we highlight the challenges that EPS biosynthesis poses to 2,3-BD fermentation and purification while proffering possible strategies to mitigate EPS accumulation during 2,3-BD fermentation, with a view to improving 2,3-BD yield and productivity.
2.2 Brief history of microbial 2,3-BD production
The first microbe reported to produce 2,3-BD was Klebsiella pneumoniae
(formerly known as Aerobacter aerogenes or Klebsiella aerogenes) and the study was carried out by Harden and Walpole in 1906 (Magee and Kosaric, 1987; Ji et al., 2011a).
In 1926, Paenibacillus polymyxa (formerly known as Bacillus polymyxa) was observed to accumulate 2,3-BD in liquid cultures (Garg and Jain, 1995; Ji et al., 2011a), which influenced Fulmer et al. to propose the first industrial microbial production of 2,3-BD in
1933 (Celinska and Grajek, 2009; Fulmer et al. 1933). During World War II (WWII), shortages in rubber supply and 1,3-BD triggered intense research efforts on 2,3-BD
10 fermentation and its conversion to 1,3-BD to meet rising demands; and the process of microbial biosynthesis of 2,3-BD and its conversion to 1,3-BD were satisfactorily improved. However, after WWII, the abundance of cheap fossil fuel derived 1,3-BD led to neglect and demise of microbial 2,3-BD production (Celinska and Grajek, 2009; Ji et al., 2011a). However, the surge in the cost of petroleum and petroleum-derived products in early 1970s reignited interest in microbial 2,3-BD production. Presently, a handful of
2,3-BD biosynthesis microorganisms has been identified while development of non- native 2,3-BD microorganisms is also being vigorously pursued by genetic engineering
(Celinska and Grajek, 2009).
2.3 2,3-Butanediol production methods
Prior to the advent of microbial 2,3-BD biosynthesis, the only available route for
2,3-butanediol production was through synthesis using fossil-derived feedstocks such as propylene, acetylene, butane and butene. Commercially, 2,3-BD is synthesized by the hydrolysis of 2,3-butylene oxide (2,3-epoxybutane). Synthesis of 2,3-butylene oxide is achieved when n-butane and 2-butene (obtained from hydrocarbon cracking) are mixed in a ratio of 1:3 and passed over oxygen and inert gases at 675 °F under high pressure. After complete oxidation of n-butane and 2-butene, a flowing stream of water is passed over the reaction mixture resulting in the production of 2,3-butylene oxide as the major product, with mixtures of hydroxyl methyl ethyl ketone (MEK) released as minor reaction products (Mitchell and Robertson, 1954). The use of cis-and trans-2-butene gives rise to cis- and trans-2,3-butylene oxide, respectively, and the isomers of 2-butene determine the stereoisomer of 2,3-BD to be synthesized. The 2,3-butylene oxide is
11 separated and further hydrolyzed to 2,3-BD. Hydrolysis of trans-2,3-epoxybutane results in the production of meso-2,3-BD, whereas a racemic mixture of levo- and meso-2,3-BD is achieved by the hydrolysis of cis- epoxybutane (O’Neil, 2001).
Today, other methods of 2,3-BD production including chemical synthesis and fermentative CO2 hydrogenation. Recently, a new method of 2,3-BD synthesis from glucose without the use of microorganisms has been reported (Ge et al., 2011). In this process, glucose is catalytically hydrogenated to sorbitol which is further cracked at high temperature to 2,3-BD. This method is rather inefficient, as mixtures of other products such as ethylene glycol and propylene glycol are also generated from the process. The caveat to this method is the high cost of 2,3-BD purification, which entails exclusion of the co-generated polyols. This is particularly important owing to the high boiling points of 2,3-BD (~180 °C), ethylene glycol (197.3 °C) and propylene glycol (188.2 °C), which are very similar and, hence, pose formidable difficulty to the separation process.
However, this method was reported to yield more than 97% propylene glycol, ethylene glycol and 2,3-BD, with 2,3-BD constituting 5% of the mixture (Xu, 2005). Changchun
Dacheng Group, a Chinese company, is working towards adopting this technology for
2,3-BD production from corn-based glucose, with the target of 60,000 tons of 2,3-BD per year (Ge et al., 2011).
2.4 Major 2, 3-BD-producing microorganisms
Numerous microorganisms especially bacteria and yeasts can convert sugars to
2,3-BD, even though the amounts generally produced by microorganisms are below levels capable of supporting industrial-scale 2,3-BD production. Some bacteria, however,
12 have demonstrated the capacity to achieve up to 10% 2,3-BD yield during fermentation.
For instance, fed-batch fermentation using Klebsiella pneumoniae resulted in the production of 150 g/L meso-2,3-BD in a glucose-based medium (Ma et al., 2009), and cultures of Enterobacter aerogenes and Serratia marcescens synthesized 110 and 152 g/L 2,3-BD from glucose and sucrose-based media, respectively (Zeng et al., 1991;
Zhang et al., 2010b). Notably, these are class II microorganisms that pose considerable dangers to humans due to their pathogenicity (Celinska and Grajek, 2009). Other class II
2,3-BD producers include Klebsiella oxytoca, Raoultella ornithinolytica and
Enterobacter cloacae. The pathogenicity of these microorganisms makes them unattractive for industrial-scale applications. However, on a more positive note, some non-pathogenic Class I microorganisms are capable of producing considerable amounts of 2,3-BD. Major class I 2,3-BD-producers include Paenibacillus polymyxa (formerly
Bacillus polymyxa), Bacillus licheniformis, and B. amyloliquefaciens. Interestingly, a maximum titer of 111 g/L 2,3-BD has been reported for P. polymyxa DSM 365 in a sucrose-based medium (Häßler et al., 2012). Further, 144 and 132.9 g/L 2,3-BD have been achieved using B. licheniformis and B. amyloliquefaciens, respectively, from glucose-based cultures (Jurchesscu et al., 2013; Yang et al, 2013). These class I 2,3-BD producers are capable of achieving 2,3-BD titers comparable to their class II pathogenic counterparts and, as such, are currently receiving considerable attention for industrial- scale microbial 2,3-BD production. P. polymyxa has a significant advantage over other major 2,3-BD producers due to its ability to synthesize up to 98% of the levo-2,3-BD isomer. This isomer is more desirable than the meso form because it is easier to dehydrate
13 to 1,3-BD (Nakashimada et al., 2000; de Mas et al, 1988). Conversely, the class II microorganisms predominantly produce the meso isomer of 2,3-BD, whereas some produce equal amounts of meso- and levo-isomers (Ma et al., 2009; Syu, 2001).
2.5 Process development for enhanced 2,3-BD production
A range of factors (including nutrients and fermentation conditions) have been identified as having major influence on microbial 2,3-BD production. Yu and Saddler
(1982) pioneered studies on the effect of acetic acid supplementation on 2,3-BD production. Fermentations conducted with the addition of 0.5% acetic acid to the fermentation medium significantly improved the growth of K. pneumoniae and increased
2,3-BD production from ~4.2 g/L to 14 g/L (~3-fold increase). In another study, a 40% increase in 2,3-BD production was achieved when 150 mM acetate was added to batch cultures of P. polymyxa (Nakashimada et al., 2000). Further increase in 2,3-BD production was obtained following simultaneous glucose and acetate addition to fed- batch cultures of P. polymyxa (Nakashimada et al., 2000).
The underlying mechanism of the acetate-mediated increase in 2,3-BD biosynthesis was traced to the dynamics of the 2,3-BD biosynthesis pathway. Acetate is one of the by-products of 2,3-BD fermentation. The presence of acetate in its ionized form in fermentation culture induces the expression of acetolactate synthase (ALS), a key enzyme of the 2,3-BD biosynthesis pathway (Stormer. 1977; Ji et al., 2011b). ALS catalyzes pyruvate conversion to acetolactate which further is converted to acetoin via the activity of acetolactate decarboxylase. Butanediol dehydrogenase further converts acetoin
14 to 2,3-BD. Thus, it is likely that acetate addition to the fermentation broth favors acetolactate formation, thereby promoting 2,3-BD production. In light of reports that acetate supplementation increases 2,3-BD biosynthesis, Zhang et al. (2010a) pursued optimization of levels of acetate and other nutrient factors such as yeast extract, sucrose, sodium citrate, ammonium hydrogen phosphate, magnesium sulfate and manganese sulfate in the fermentation medium for improved 2,3-BD production. The study showed that supplementation of 4 g/L acetate and 33.3 g/L yeast extract increased 2,3-BD production by S. marcescens from 9.4 g/L to 44.7 g/L in batch cultures. In a similar study, Li et al. (2013a) tested different concentrations of acetate, yeast extract, corn steep liquor powder (CSLP), potassium phosphate, triammonium citrate and magnesium sulfate, and showed that 6.5 g/L acetate, 5.8 g/L yeast extract, 14.7 g/L CSLP increased
2,3-BD production by B. licheniformis from 18.9 g/L to 42.3 g/L.
Fermentation conditions have also been manipulated to increase 2,3-BD production. Conditions such as agitation speed, temperature, and culture pH greatly impact microbial 2,3-BD production (Ji et al., 2011a; Celinska and Grajek, 2009). Indeed, agitation speed regulates oxygen supply to bacterial cells during fermentation impacting growth and product formation. Owing to the micro-aerobic nature of 2,3-BD fermentation, oxygen supply to bacteria cultures is manipulated to create an initial oxygen-rich environment for enhanced biomass accumulation, and then switched to an oxygen-limiting environment once significant cell biomass is developed. The switch to oxygen-limiting environment favors 2,3-BD biosynthesis (Celinska and Grajek, 2009).
Recently, some studies employed a two-stage agitation speed control strategy to create
15 different levels of aeration in the bioreactor to achieve higher 2,3-BD production. For instance, Ji et al. (2009) started 2,3-BD fermentation using K. oxytoca at 300 rpm for the first 15 h of fermentation and switched to 200 rpm for the rest of the fermentation period.
The two-stage aeration strategy led to 16% and 20% increases in cell biomass during 2,3-
BD production compared to fermentation conducted at 100 rpm. In addition, acetoin concentration decreased by 67% compared to when the fermentation was conducted at
100 rpm. Li et al. (2013a) previously performed a similar two-stage agitation speed control study using B. licheniformis. Switching agitation speed from 400 rpm after 10 h of fed-batch fermentation to a lower agitation speed of 200 rpm for the rest of fermentation increased 2,3-BD production from 45.7 g/L (at constant 400 rpm) to 115.7 g/L (two-stage agitation speed control). The improved 2,3-BD production corresponds to a sharp decrease in acetoin accumulation, which decreased from 45.1 g/L (conducted at constant 400 rpm) to 2.2 g/L (two-stage agitation speed strategy).
Temperature and culture pH are the other fermentation conditions that have drawn research attention towards enhancing 2,3-BD production. Generally, the optimum temperature range for microbial 2,3-BD production is 30-39 °C, and the optimal culture pH falls in the range of 5.5-6.3 (Celinska and Grajek, 2009). According to Biebl et al.
(1998), lowering the temperature of K. pneumoniae culture from 35 to 30 °C substantially decreased ethanol formation while improving 2,3-BD production. Further, starting 2,3-
BD fermentation at a pH of about 7 and a temperature of about 30 °C appear to be the best conditions for maximum 2,3-BD production by K. pneumoniae (Biebl et al., 1998).
Conversely, Barret et al. (1983) tested temperatures in the range of 30 - 37 °C on 2,3-BD
16 production by K. pneumoniae and E. aerogenes, and reported that the highest 2,3-BD yield achieved by K. pneumoniae was at 33 °C, which was 5% and 203% more than the yields obtained at 30 °C and 37 °C, respectively. For E. aerogenes, the highest 2,3-BD yield was achieved at 37 °C and pH 7. In a similar study conducted by Perego et al.
(2003), 39 °C was optimal for 2,3-BD production by E. aerogenes. For P. polymyxa, the optimal temperatures for 2,3-BD production are in the range of 30 - 39 °C at pH of 6.0
(Dai et al., 2014; Marwoto et al., 2002; Nakashimada et al., 1998). The optimal temperatures for strains of B. licheniformis vary from 37 - 50 °C. When 2,3-BD fermentation was conducted in the temperature range of 37 - 60 °C using B. licheniformis
10-1-A, it was observed that 50 °C was the optimal temperature for 2,3-BD production
(Li et al., 2013a). 2,3-BD production by B. licheniformis 10-1-A at this temperature increased ~ 200% when compared to the fermentation conducted at 37 °C (Li et al.,
2013a). Li et al. (2013a) further tested the pH optimum of B. licheniformis 10-1-A for
2,3-BD production. At pH 7, 2,3-BD production increased to 49.6 g/L compared to 4.5 g/L obtained at pH 8.
To exploit the pH preference of acetoin reductase/butanediol dehydrogenase of B. subtilis to regulate acetoin and 2,3-BD production, Zhang et al. (2014) applied a two- stage pH based strategy. Notably, while pH of 6.5 favors acetoin reduction to 2,3-BD, pH of 8.5 favors the oxidation of 2,3-BD to acetoin. Priya et al. (2016) pursued combined optimization of a two-stage pH and agitation strategy in a bioreactor to improve 2,3-BD production by E. cloacae. Fermentations started at initial pH of 7.5 and held for 10 h before switching to pH 6.5 for the rest of the fermentation led to increased 2,3-BD
17 production from 65.2 g/L (under constant pH of 7.5) to 75.02 g/L (in a two-stage pH strategy), with a concomitant decrease in ethanol production from 8.23 g/L to 3.24 g/L.
To date, however, combined optimization of medium components (nutrients) and culture conditions has not been investigated. There is a possibility of interaction between nutrients and culture conditions, which may engender increased 2,3-BD production. This hypothesis forms the basis of the process development aspect of this work.
2.6 Types of EPS and structures
Exopolysaccharides (EPS) produced by 2,3-BD-producing microorganisms are extracellular polymeric substances that exist in various forms - ionic EPS with a net positive (acidic) or negative (basic) charge on the surface of the polymers and neutral
EPS with no net charge (Fig. 2.3). The ionic extracellular polymeric substances synthesized by bacteria include alginates and xanthan which are predominantly made by
Pseudomonas and Azotobacter spp, and Xanthomonas campestris, respectively (Hay et al., 2013; Schmid and Sieber, 2015; Vuyst and Degeest, 1999), whereas neutral EPS such as dextran, levan, and other EPS types, are mainly synthesized by P. polymyxa and B. licheniformis (Häßler et al., 2012; Liu et al., 2010a; Liu et al., 2010b). The ionic EPS possesses carboxylic acid residues derived mainly from pyruvate or uronic acids which enable EPS to form complexes with metallic ions. The negative charges are also conferred by the sugar nucleotide monomers of some EPS. It is thought that EPS- producing bacteria utilize the net negative charge on the surface of EPS as a mechanism for sequestering heavy metals in their natural habitats (Ordax et al., 2010). EPS interact
18 and bind to divalent cations very strongly, thus, playing a pronounced role in metal flocculation (More et al., 2014). The affinity of EPS for different cations is dependent on the type of EPS and cations present. Indeed, the EPS produced by Paenibacillus spp. was shown to sequester heavy metals such as Pb, Cu, Zn, Ni, Co and Cd (Morillo et al.,
2006).
EPS confers competitive advantage to EPS producing microorganisms. For instance, EPS exhibits some surface-active properties, which enable bacterial species that secrete them to emulsify hydrocarbons in water. EPS binds strongly to the surface of hydrocarbon droplets enabling emulsification (Sutherland, 1990). The bacterium is then able to use the emulsified hydrocarbons as an energy source. The surfactant activity of bacterial EPS has been employed in oil spill clean-up programs (bio-remediation; More et al., 2014; Gutierrez et al., 2013).
Structurally, EPS can exist as homo-polysaccharides with the repeating unit being a specific monosaccharide or a derivative of a monosaccharide, while some exist as hetero-polysaccharides with the polymer having a number of different monosaccharide units or the derivatives of monosaccharides or a combination of both as the repeating units (Fig 2.4). Dextran, pullulan and curdlan are examples of homopolymers. Curdlan, which is synthesized by P. polymyxa (Rafigh et al., 2014), is composed of a single glycosyl-linkage (1,3- linkage), whereas pullulan and dextran contain two (1,6- and 1,4- linkages) and three or more (1,6-; 1,2-; 1,3- or 1;4- linkages) different types of glycosyl- linkages, respectively (Shingel, 2004; Monsan et al., 2001). Examples of hetero-polymers include levan and xanthan (Fig 2.4).
19
2.7 Ecological relevance of EPS biosynthesis
A considerable number of major 2,3-BD producing microorganisms have been isolated from different sources where they were reported to use EPS as a means of adaptation to unfavorable environmental conditions (Kostakioti et al., 2013). In
Klebsiella and Serratia sp, capsular polysaccharides and extracellular lipopeptides are used for host invasion, colonization and for protection against desiccation (Zhang et al.,
2010b; Bryan et al., 1986), whereas Paenibacillus species use EPS as a form of support and attachment to plant hosts in natural environments (Bezzate et al., 2001). Some bacteria use EPS to either establish communities through biofilm formation or grow on already synthesized structural polysaccharide matrices (Fuente-Nunez et al., 2014), enabling propagation in their native environments. These biofilms and structural polysaccharide matrices confer on the bacteria some form of resilience to adverse environmental conditions, thereby allowing them to thrive at extreme temperatures, and sometimes, in extreme pH conditions. Microbial EPS is particularly important industrially with wide-ranging applications in various industries such as in the food processing, cosmetics and pharmaceutical industries (Moon et al., 2006; Zanchetta et al.,
2003; Mansel, 1994), as well as for the removal of toxic metal ions from wastewater and crude oil-contaminated lands (Chu and Kim, 2006; Shi et al., 2006). However, EPS are unwanted in microbial 2,3-BD production due to their negative impact on 2,3-BD fermentation and separation.
Some studies have suggested that bacteria cannot re-use EPS as an energy or carbon source (Daigger and Leslie-Grady, 1982) due to claims that most bacteria which
20 synthesize EPS lack the requisite enzymes for converting the EPS to utilizable sugars.
Contrary to this notion, it has been shown that some bacteria can utilize EPS as carbon and energy sources (Salazar et al., 2008). For instance, P. polymyxa, can express levanase, an enzyme that hydrolyzes levan (EPS) to fructose residues; this possibly may explain the observed decrease in the concentration of synthesized levan during 2,3-BD fermentation under carbon limitation (Raza et al., 2011). In addition, some strains of
Bifidobacteria can utilize the heteropolysaccharides (containing glucose, galactose and mannose) that they produce in the intestines of man and animals by converting them to utilizable volatile fatty acids (Salazar et al., 2008).
2.8 Relationship between 2,3-BD production and biosynthesis of EPS
Most EPS synthesized by major 2,3-BD-producing bacteria can be classified as biofilms, extracellular or capsular polysaccharides. The EPS synthesized during 2,3-BD fermentation typically vary in amount, depending on the sugar substrate used and the micronutrients available in the medium, in addition to the fermentation conditions employed (Lee et al., 1997; Häßler et al., 2012). For instance, sucrose is the predominant substrate (precursor) for EPS production by P. polymyxa and B. licheniformis (Häßler et al., 2012; Liu et al., 2010b; Lee et al., 1997), with a production of up to 54 g/L EPS by P. polymyxa during 2,3-BD fermentation (Häßler et al., 2012). Sucrose is known to be the preferred substrate for EPS production by P. polymyxa, however, EPS is produced by cultures of P. polymyxa grown on other non-sucrose substrates (Lee et al., 1997). In a fermentation study using 20 g/L different carbohydrates, P. polymyxa synthesized 18 g/L
21
EPS from sucrose and 9, 4.8, 4.8, 4.6 and 6 g/L EPS from glucose, fructose, galactose, lactose and soluble starch, respectively (Table 2.1).
Production of EPS during 2,3-BD fermentation is not limited to P. polymyxa as studies have shown that Klebsiella and Serratia species synthesize capsular polysaccharides and extracellular lipo-peptides during 2,3-BD fermentation. The production of large amounts of capsular polysaccharides during 2,3-BD fermentation by
K. pneumoniae increased fermentation broth viscosity and impacted cell removal from the broth during 2,3-BD downstream processing (Cho et al., 2012). Capsular polysaccharides and other EPS increase the viscosity of fermentation broth and negatively impact filtration during downstream processing (Guo et al., 2010). When fermentation is conducted in a fed-batch mode, EPS production by 2,3-BD microorganisms increases with increases in 2,3-BD production. For instance, 54.8 g/L
EPS was generated during fed-batch fermentation with K. pneumoniae sp. pneumoniae
(Ramirez-Castillo and Uribelarrea, 2004), which further underpins the diversion of carbon to EPS by Klebsiella species and away from 2,3-BD biosynthesis.
Serrawettin W1, an extracellular lipopeptide, synthesized by Serratia spp during
2,3-BD fermentation diminishes the 2,3-BD production capacity of S. marcescens.
Serrawettin W1 is a non-ionic biosurfactant, that cause foaming of the fermentation broth during 2,3-BD production. Foaming causes loss of broth, cells and products during fermentation and can also increase the risk of contamination (Stanbury et al., 1995).
22
2.9 Molecular genetics of EPS production
The chemical composition of bacterial EPS is dependent on the genetic repertoire of the producing microorganisms and on their ecological niches (Sutherland, 2001).
Multiple genes are thought to be involved in bacterial EPS biosynthesis, however, only a few of them have been identified in P. polymyxa, K. pneumoniae, K. oxytoca and Serratia marcescens. In Bacillus spp. and other Gram positive bacteria, the EPS genes are borne on a large epsA-O operon known as the 15-operon (Nagorska et al., 2010). The genes of the epsDEFGHJ operon encode enzymes that catalyze EPS polymerization in the bacterial cell, whereas the genes of the epsABCIK operon encode the proteins necessary for the EPS export (Looijesteijn et al., 1999). A section of the epsA-O operon found in
Bacillus spp has been annotated in P. polymyxa (Hou et al., 2016; Jeon et al., 2010). The eps operon presumed to be exopolysaccharide biosynthesis cluster in P. polymyxa SC2 consist of twelve genes (Hou et al., 2016). Levansucrase, the enzyme that catalyzes the extracellular synthesis of levan, is encoded by the sacB gene. The sacB gene encodes a protein with a 499 amino acid sequence in P. polymyxa strain CF43 (Bezzate et al.,
2001). A different gene, sacC, encodes levanase, the enzyme that degrades levan (Shida et al., 2002).
The synthesis of capsular polysaccharide in Klebsiella spp is controlled by the chromosomal cps region of this bacterium and is comprised of nineteen open reading frames. The cps operon in Klebsiella is employed in the synthesis of GDP-mannose pyrophosphorylase, phosphomanomunase and other EPS biosynthesis enzymes (Arakawa et al., 1995). The capsular polysaccharide of Klebsiella spp is a colanic acid-containing
23 hexasaccharide repeating units comprised of D-glucose, D-galactose, L-fucose and D- glucuronic acid in the ratio of 1:2:2:1 (Ophir and Gutnick, 1994; Markovitz, 1977). In
Serratia spp, pswP has been identified as the central gene for serrawettin biosynthesis
(Sunaga et al, 2004; Li et al., 2005). The pswP encodes a phosphopantetheinyl transferase
(PPTase) that transfers a phosphopantetheinyl group from coenzyme A to the peptidyl carrier protein during serrawettin biosynthesis (Li et al, 2005). This occurs during post- translational modification of serrawettin. Serrawetin is a cyclodepsipeptide (lipopeptide) composed of two molecules of serratamic acid (D-3-hydroxydecanoyl-L-serine). Due to the absence of serratamic acid spots on the thin-layer chromatograms of serrawettin W1 mutant strains of Serratia marcescens, it was suggested that nonribosomal peptide synthetases (NRPS family) may be involved in serrawettin biosynthesis (Pradel et al,
2007). The gene encoding NRPS was predicted to be swrA of S. marcescens because S. marcescens swrA mutants were deficient for serrawettin W2 production (Pradel et al,
2007; Lindum et al, 1998; Matsuyama et al, 1992).
2.10 EPS mitigation strategies: manipulations of nutrients and fermentation conditions
Metabolic stressors may contribute to EPS biosynthesis during 2,3-BD fermentation. Established stressors that affect EPS production during fermentation include culture pH, temperature, oxygen availability, and agitation speed (Raza et al.,
2011; Liu et al., 2009; Bandaiphet and Prasertsan, 2006). Other factors such as substrate types and nutrient availability have been reported to influence EPS biosynthesis (Raza et
24 al., 2011; Jefferson, 2004). Meticulous manipulation of stressors could reduce EPS biosynthesis and possibly maintain or improve 2,3-BD production.
2.10.1 Oxygen availability and agitation
Oxygen availability influences both EPS and 2,3-BD production (Lee et al., 2007;
Celinska and Grajek, 2009; Bandaiphet and Prasertsan, 2006). Several reports have attested that fermentation conducted under micro-aerobic (limited oxygen) conditions led to improved 2,3-BD titer, yield and productivity (Celinska and Grajek, 2009; Voloch et al., 1985). 2,3-BD-producing bacteria are largely facultative organisms, hence, they thrive in both aerobic and limiting oxygen environments. Agitation of fermentation medium ensures proper mixing of nutrients and dissolution of oxygen in the broth
(Survase et al., 2007). The speed of agitation during fermentation determines the amount of oxygen available to the fermenting organisms. Low agitation speed results in limited oxygen supply, whereas a high agitation speed ensures high oxygen supply. Since 2,3-BD fermentation is a micro-aerobic process, conducting fermentation under limited oxygen supply may enhance 2,3-BD production and decrease EPS formation. While limited oxygen supply favors 2,3-BD production, it may result in reduced cell biomass buildup.
Further, the rate of NADH oxidation (NADH → NAD+) via respiration is reduced during fermentation under limited oxygen supply, which may negatively affect glycolysis given the critical role of NAD+ in the oxidative step of the glycolytic pathway (Inui et al.,
2004). However, 2,3-BD fermentation under limited oxygen supply also lead to increase ethanol production (Voloch et al., 1985). High oxygen supply favors EPS production
(Lee et al., 1997). Considering that aeration is commonly associated with agitation speed
25
(Celinska and Grajek, 2009), the use of two-way agitation speed control could also reduce EPS and CPS production. Starting fermentation at a high agitation speed and switching to a low speed after sufficient cell biomass develops may reduce EPS formation during 2,3-BD fermentation.
2.10.2 Osmotic stress
Osmotic stress is one of the factors that trigger bacterial production of EPS
(Bandaiphet and Prasertsan, 2006). Many EPS-producing bacteria synthesize EPS for protection against desiccation and osmotic stress stemming from high concentrations of dissolved solutes (Robertson and Firestone, 1992). Under osmotic stress, bacteria respond in very specific ways that alter expression of certain genes and cellular processes (Zamfir and Grosu-Tudor, 2014) including the expression of EPS biosynthesis genes (for EPS- producing bacteria). Consequently, carrying out 2,3-BD fermentation in medium with low salt concentration may reduce EPS formation. Some metal ions act as cofactors for enzymes that produce EPS (Suntherland, 1977; Domenico et al., 1989). Metal ions that enhance EPS production include Ca2+ and Fe3+ (Raza et al., 2011; Suntherland, 2001;
Meng and Futterer, 2003; Domenico et al., 1989). For instance, Ca2+ plays a critical role in maintaining the structural integrity and stability of levansucrase (Meng and Futterer,
2003). However, metals such as Mg2+, Mn2+ or K+ stimulate growth but not EPS production (Grobben et al., 2000). Supplementation of fermentation medium with metal chelating agents such as EDTA (ethylenediaminetetraacetic acid) and EGTA [ethylene- bis(oxyethylenenitrile)tetraacetic acid] can bind metals and make them unavailable for
EPS biosynthesis. Indeed, supplementation of fermentation medium with sodium
26 salicylate and EGTA as chelating agents resulted in drastic reduction of capsular polysaccharide production by K. pneumoniae strains without affecting cell growth
(Domencio et al., 1989). Consequently, it is imperative to reduce or eliminate the use of
Ca2+ and Fe3+ in 2,3-BD fermentation, replace them them with less EPS-inducing ions
(Co2+, Mg2+, Mn2+ or K+) to minimize EPS biosynthesis.
2.10.3 Culture pH
2,3-BD production is pH-dependent and the pH optima for 2,3-BD biosynthesis ranges from 6.0-6.8 (Nakashimada et al., 2000; Perego et al., 2000) although few studies did report a pH optimum of 5.5 (Celinska and Grajek, 2009). The pH optimum for 2,3-
BD production appears to be species-specific, with Klebsiella and Enterobacter species preferring pH below 6, whereas pH values above 6 are more favorable to Paenibacillus species (Biebl et al., 1998; Voloch et al., 1985; Celinska and Grajek, 2009). Changes in pH during fermentation affect the activities of 2,3-BD and EPS biosynthesis enzymes.
Furthermore, EPS production by Paenibacillus spp. is favored by pH ranges between 6.5 and 8 (Liu et al., 2009; Raza et al., 2011), whereas E. cloacae prefers pH 7 (Bandaiphet and Prasertsan, 2006), a pH of 7.5 promotes biosynthesis of capsular polysaccharide by
K. pneumoniae (Domenico et al., 1989). Since pH changes affect various 2,3-BD- producing microorganisms differently, modulating fermentation culture pH should be performed in a way that would minimize EPS production while enhancing 2,3-BD production. For instance, conducting 2,3-BD fermentation below pH of 6.5 for P. polymyxa would minimize EPS formation.
27
2.10.4 Fermentation temperature
Fermentation temperature is a critical factor for EPS biosynthesis (Fett, 1993;
Sutherland, 2001). The optimal temperature for EPS production varies with bacterial species/strain. Generally, the optimal temperature for EPS biosynthesis ranges from 24 and 30 °C (Raza et al., 2011; Liu et al, 2009). For instance, the optimal temperature for levan biosynthesis by P. polymyxa was reported to be at 30 °C (Han and Clarke, 1990), whereas for some Bacillus spp., the temperature ranges from 30 and 50 °C (Nakapong et al., 2013; Belghitha et al., 2012). The optimal temperature for capsular polysaccharide production by Klebsiella spp. is 30 °C (Mengistu et al., 1994), although higher temperatures (up to 40 °C) have been reported to support the growth of K. pneumoniae with diminished CPS production (Mengistu et al., 1994). Since 30 ºC is the optimal temperature for EPS production by P. polymyxa, conducting fermentations at 35 °C may reduce EPS formation. A previous report indicates that temperature of 42 °C can support growth of P. polymyxa (Marwoto et al., 2002). Thus, production of 2,3-BD by P. polymyxa in temperatures above 35 °C could reduce EPS production.
2.10.5 Nitrogen sources
EPS biosynthesis is influenced largely by nitrogen availability in the fermentation medium. The available nitrogen is expressed as a ratio to the total carbon in the fermentation medium (C/N ratio). High C/N ratio enhances the synthesis of capsular polysaccharides by Klebsiella spp and E. aerogenes (Ramirez-Castillo and Uribelarrea,
2004; Farres et al., 1997; Sutherland, 1983; Dudman and Wilkinson, 1956; Duguid and
Wilkinson, 1953). Fermentation medium with high C/N ratio appears to stimulate EPS
28 producing microbes to convert the excess carbon to EPS (Duguid and Wilkinson, 1953).
However, a different scenario appears to apply in the case of P. polymyxa where conflicting conclusions have been drawn in this regard. Häßler et al. (2012) demonstrated that at 5 g/L yeast extract EPS production by P. polymyxa is enhanced whereas when yeast extract in the fermentation medium was increased to 60 g/L a drastic reduction in
EPS production was observed. Conversely, in a study where the nitrogen content of the fermentation medium was monitored, production of EPS by P. polymyxa ceased when the nitrogen in the medium was exhausted (Lee et al., 1997). In fact, the authors observed rapid degradation of EPS when nitrogen in the medium exhausted (Lee et al., 1997). In another study, Liu et al. (2009) tested the effects of 10 g/L each of yeast extract, beef extract, meat peptone and several inorganic nitrogen sources on EPS biosynthesis by P. polymyxa. The culture containing yeast extract produced the maximum EPS. The authors suggested that yeast extract may contain EPS stimulatory compounds which directly influence EPS production by P. polymyxa. The conflicting conclusions on effects of nitrogen on EPS production by different groups may stem from varying culture conditions employed by the researchers. To minimize capsular polysaccharide biosynthesis by Klebsiella spp., E. aerogenes and other pathogenic 2,3-BD producers, it may be rational to increase the nitrogen content of 2,3-BD fermentation medium.
However, with P. polymyxa, increasing the nitrogen content of the medium may not necessarily result in reduced EPS production.
29
2.11 EPS mitigation strategies: potential for strain development
Modulation and optimization of culture conditions and fermentation medium components offer considerable promise for reducing EPS biosynthesis in 2,3-BD- producing bacteria. Unfortunately, some factors that reduce EPS biosynthesis also reduce growth and 2,3-BD production. Development of non-EPS-producing, hyper-2,3-BD producing mutants could completely eliminate the problems posed by EPS to 2,3-BD fermentation. Towards achieving this goal, key EPS biosynthesis genes need to be identified for targeted genetic manipulation. Cutting-edge gene knockout/knockdown, mRNA inactivation or suppression as well as gene editing techniques, which have undergone significant advancements recently, hold substantial promise towards generating EPS null mutants with hyper 2,3-BD production capability.
In addition to formation of capsular polysaccharides (CPS) and lipopolysaccharides (LPS) during 2,3-BD fermentation, K. pneumoniae synthesizes fimbriae, a virulence factor used for adhesion to surfaces (Huynh et al., 2015). Fimbriae expression facilitates the ability of K. pneumoniae to form biofilms which results from the accumulation of CPS and LPS (Schroll et al., 2010; Balestrino et al., 2008; Schembri et al., 2005). The deletion of fim, cps and lps genes in K. pneumoniae would reduce expression and biosynthesis of fimbriae, CPS and LPS with the following benefits (1) increased efficiency in substrate conversion to 2,3-BD, with enhanced 2,3-BD yield as substrates which would have been used for the syntheses of CPS, LPS and fimbriae are blocked and only available for 2,3-BD synthesis; (2) reduced cost of downstream processing and improved cost of 2,3-BD fermentation; and (3) the removal of the
30 virulence factor in K. pneumoniae and oxytoca would increase its prospects of being adopted for industrial scale 2,3-BD fermentation. In the chromosome of K. pneumoniae, several open reading frames (ORFs) designated as cps genes are arranged in clusters although, the number of ORFs in the cps gene clusters is strain specific. Expression of cps genes in K. pneumoniae leads to synthesis of various enzymes responsible for CPS formation. Since there are several ORF in the cps gene clusters, attempts to disrupt all
ORFs in the chromosomal cps gene cluster may result in impaired K. pneumoniae growth, which would ultimately impact 2,3-BD production. A specific ORF in the cps gene clusters has been targeted in K. pneumoniae with a view to reducing CPS formation during 2,3-BD and 1,3-propanediol (1,3-PD) fermentation. Guo et al. (2010) disrupted the ORF3 of cps gene clusters in K. pneumoniae CGMCC 1.6366 and this led to approximately 17% and 7% improvement in 2,3-BD and 1,3-PD production, respectively.
However, cell growth was unaffected, whereas broth viscosity reduced dramatically with potential to improve downstream processing. In the study, the resulting strain (cps null mutant) exhibited greater ease of cell separation from the broth and CPS, when compared to the wild type. Further, rmpA and its isoform, rmpA2 genes, were identified in a plasmid of K. pneumoniae. The genes were shown to regulate CPS production in K. pneumoniae (Cheng et al., 2010; Hsu et al., 2010). The deletion of rmpA/rmpA2 in K. pneumonia NTUH-K2044 revealed that expression of cps gene clusters and capsule production were only enhanced by rmpA gene (Cheng et al., 2010; Hsu et al., 2010).
Therefore, deletion of rmpA gene in the plasmid of K. pneumoniae would be a better
31 option to abolish CPS production during 2,3-BD fermentation than attempting deletion of the entire cps gene clusters.
Studies on the elimination of virulence factors in K. pneumonia and oxytoca yielded promising results for applying Klebsiella species in industrial 2,3-BD fermentation. The wabG and fimA genes, responsible for LPS and fimbriae formation, respectively, in K. pneumoniae, and the wabG gene of oxytoca were inactivated via homologous recombination. The K. pneumoniae and oxytoca wabG null mutants showed impaired capsule formation, and K. pneumoniae fimA null mutant showed no fimbriae formation (Huynh et al, 2015; Jung et al., 2013). Following wabG and fimA inactivation in K. pneumoniae, the growth rate and 2,3-BD production capabilities of the mutants were unaffected (Huynh et al, 2015), however, Jung et al. (2013) observed a 28 % reduction in 2,3-BD production by the K. pneumoniae wabG mutant even though the cell growth was unaffected. Production of acetoin and lactate in the K. pneumoniae wabG mutant was negligible when compared to the wildtype. Although yield was not determined, it is plausible that the wabG mutant produced higher 2,3-BD yield relative to the wildtype due to lesser production of competing products.
Similarly, metabolic engineering approach has been extended to S. marcescens
H30 where the swrW gene, coding for serrawettin W1 synthase, was inactivated.
Foaming during 2,3-BD fermentation reduced significantly and the 2,3-BD production capacity of S. marcescens H30 srwW null mutant in a fed-batch fermentation using an optimized medium reached 152 g/L (Zhang et al., 2010a; 2010b). The observed improvement in 2,3-BD production in the srwW mutant could be partly attributed to
32 improved channeling of carbon to 2,3-BD over serrawettin biosynthesis (Fig 2.1). It is also probable that inactivation of srwW gene in S. marcescens H30 may have improved the tolerance of the srwW mutant to 2,3-BD, thereby facilitating the accumulation of up to 152 g/L 2,3-BD in the bioreactor.
The other native 2,3-BD producers that synthesize large amounts of EPS during
2,3-BD fermentation include B. licheniformis, B. amyloliquefaciens, P. polymyxa and E. aerogenes. Nonetheless, there are no reports to date on metabolic engineering of these microorganisms to reduce EPS production during 2,3-BD fermentation. The major EPS produced by B. amyloliquefaciens, licheniformis and P. polymyxa species is levan.
Levansucrase is the key enzyme involved in levan biosynthesis in these microorganisms.
Therefore, targeting levansucrase for deletion in these microorganisms might improve
2,3-BD yield by rechanneling excess carbon that is currently metabolically invested in
EPS biosynthesis to 2,3-BD production. In addition, this would ultimately reduce the cost of downstream processing of 2,3-BD. Levan is the only characterized EPS synthesized by
B. licheniformis, amyloliquefaciens and P. polymyxa species, although it may not be the only EPS produced by these species. EPS production by P. polymyxa has been observed in fermentations involving numerous sugar types (Lee et al., 1997; Raza et al., 2011).
Bioinformatics-assisted analysis of the genome of P. polymyxa reveals the presence of genes coding for three putative exopolysaccharide export proteins and several sugar- nucleotide interconversion proteins, which indicate that P. polymyxa may produce more than one EPS type (Fig 2.2). Indeed, there are some reports on synthesis of EPS by P. polymyxa that are composed of mannose, galactose, and glucose in the ratios of
33
1.23:1.14:1 with monosaccharides and glucuronic acid ratio of 7.5:1 (Raza et al., 2011), and other EPS composed of mainly mannose, fructose and glucose (Liu et al., 2009). It is imperative to identify and target the key genes responsible for EPS biosynthesis in native
2,3-BD producers. E. aerogenes (formerly Aerobacter aerogenes) synthesizes four serologically identical capsular polysaccharides made of repeating units of four tetrasaccharide consisting of glucose, mannose, glucose and glucuronic acid (Gahan et al., 1967). Again, there is a dearth of information on the genetics of exopolysaccharide production in E. aerogenes. Analysis of the genome of E. aerogenes would likely provide insight as to what genes to target to reduce the virulence of E. aerogenes and in turn, enhance 2,3-BD production capacity.
Another important target to alter to enhance 2,3-BD production is to increase intracellular NADH/NAD+ flux in 2,3-BD producing microorganisms. Enhancing the intracellular levels of NADH improves biosynthesis of 2,3-BD which is subject to redox potential. Acetoin conversion to 2,3-BD requires NADH and the direction of the reaction catalyzed by 2,3-butanediol dehydrogenase is dependent on the intracellular
NADH/NAD+ ratio (Johansen et al., 1975; Magee and Kosaric, 1987; Blomqvist et al.,
1993). High intracellular NADH/NAD+ ratio favors 2,3-BD production (acetoin→2,3-
BD) and a low ratio favors the reverse reaction (acetoin ← 2,3-BD). Increasing intracellular NADH/NAD+ ratio in 2,3-BD producing microorganisms using exogenous chemicals such as ascorbic acid and sodium borohydride (NaBH4) has been shown to enhance 2,3-BD biosynthesis (Dai et al., 2014). Indeed, while exogenous addition of
NADH to E. aerogenes cultures altered the overall intracellular NADH/NAD+ ratio and
34 increased 2,3-BD, acetate, and ethanol production with concomitant reduction in hydrogen production, exogenous addition of NAD+ triggered the opposite effects (Zhang et al., 2009a). However use of NADH or compounds that promote NADH regeneration to increase 2,3-BD production will add to the overall cost of 2,3-BD production by fermentation. Therefore, introduction of genes whose protein products boost
NADH/NAD+ levels in 2,3-BD-producing microorganisms might be a better approach.
For example, heterologous introduction of the Vitreoscilla hemoglobin (VHb) gene into
E. aerogenes prolonged cell viability and enhanced acetoin and 2,3-BD production by up to 83%, although the mechanism of VHb-induced increased cell viability and product accumulation remains unknown (Geckil et al, 2004). However, while the intracellular
NAD+/NADH levels in the E. aerogenes vgb strain carrying the VHb gene was not determined, Geckil et al. (2004) suggested that the observed increases may have resulted from indirect changes in NAD+/NADH and ADP/ATP ratios engendered by the ability of
VHb to make interior cellular compartment more oxidized, thereby altering the carbon flux within central metabolic pathways (Geckil et al., 2004; Celinska and Grajek, 2009).
High intracellular NADH/NAD+ ratio in bacteria may enhance production of fermentation products but may also adversely affect microbial growth. An approach to increasing the intracellular NADH levels in 2,3-BD producing organisms without altering the NADH/NAD+ ratio is to co-express NADH and NAD+-regenerating enzymes. To maintain high NADH levels without affecting the NADH/NAD+ ratio, Yang et al. (2013) co-overexpressed the glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and 2,3- butanediol dehydrogenase (BDH) genes of B. amyloliquefaciens. GAPDH converts
35 glyceraldehyde-3-phosphate to 1,3-bisphosphoglycerate with concomitant reduction of
NAD+ to NADH, whereas BDH utilizes the NADH generated from GAPDH to oxidize acetoin while releasing 2,3-BD and NAD+. The B. amyloliquefaciens mutant with the gapdh and bdh overexpressed genes generated a record 2,3-BD concentration of 132.9 g/L in 45 h with a productivity of 2.95 g/L/h (Yang et al., 2013). In another study, Yang et al. (2015) disrupted the NADH oxidase (nox) gene of B. subtilis 168 with heterologous insertion of Candida boidinii’s formate dehydrogenase gene (fdh). NOX oxidizes
+ available NADH to NAD and H2, whereas FDH catalyzes CO2 production from formate while reducing NAD+ to NADH. Disruption of B. subtilis nox not only makes NADH available but the expression of fdh gene ensures increased intracellular NADH pool when the cultures were supplemented with sodium formate. Sodium formate addition ensures continuous supply of NADH from NAD+. This approach has previously been applied to increase levels of NADH and NAD+ without altering the intracellular NADH/NAD+ ratio by heterologous expression of formate dehydrogenase (fdh) gene of C. boidinii in K. oxytoca for efficient conversion of glycerol to 1,3-PD (Zhang et al., 2009b).
Considering that 2,3-BD is biosynthesized in a mixed acid pathway, deletion of genes that encode competing products such as ethanol, formic and lactic acid in major
2,3-BD-producing microorganisms may improve 2,3-BD titer and yield. Collectively, reduced accumulation of EPS, ethanol, formic and lactic acids would ultimately obviate the need for some additional steps during 2,3-BD recovery.
Further, genes from native 2,3-BD producing microorganisms may be transferred to non-native 2,3-BD producers. The non-native 2,3-BD producing microorganisms could
36 be selected for heterologous gene transfers based on the criteria that such microorganisms are devoid of EPS production capabilities, while producing minimal amounts of 2,3-BD competing products. The non-native 2,3-BD producers should as well be able to utilize lignocellulosic-derived sugars and tolerate, to a significant extent, microbial inhibitory compounds that are generated during lignocellulosic biomass pretreatment and hydrolysis. It would be an added advantage to select non-native 2,3-BD producing microorganisms that can utilize diverse but cheap substrates for industrial 2,3-BD fermentation. Use of alternative, cheap and easily accessible substrates for industrial 2,3-
BD production would improve the economics of 2,3-BD production.
Recently, genes responsible for the competing pathways of 2,3-BD production were deleted in E coli JM109 while the 2,3-BD genes from pyruvate were assembled in a high-copy plasmid, pEnBD, with a constitutive promoter. The mutant strain generated low amounts of acetate, ethanol, and succinate under a low oxygen aeration condition.
However, while the sugar utilization and 2,3-BD production by the generated mutant were high under aerobic condition, its growth under microaerophilic condition was slow but had more than 50% increase in 2,3-BD yield, relative to the wildtype (Li et al.,
2010a).
Similarly, introduction of two NADH-regenerating enzymes (glucose and formate dehydrogenases) and 2,3-butanediol dehydrogenase in E. coli increased the intracellular levels of NADH with the generation of 31.7 g/L 2,3-BD, which translates to a yield of
89.9% from diacetyl. Thus, increasing the intracellular levels of NADH is a driver for
2,3-BD production. Interestingly, production of organic acids (acetic, lactic, and gluconic
37 acids) was abolished completely when the levels of NADH in the cell were high (Wang et al., 2013).
Saccharomyces cerevisiae is an important non-native 2,3-BD producer, which has potential for industrial applications. However, ethanol, one of the competing products of
2,3-BD fermentation, is the major product of S. cerevisiae and would limit 2,3-BD yield in yeast. Pyruvate decarboxylase (pdc) is the key enzyme for ethanol production in S. cerevisiae and catalyzes pyruvate conversion to acetaldehyde which is subsequently converted to ethanol by alcohol dehydrogenase. When PDC was targeted for deletion in
S. cerevisiae for efficient 2,3-BD production, the inactivation of S. cerevisiae’s pdc completely abolished ethanol production (Lian et al., 2014). When exogenous acetolactate synthase, acetolactate decarboxylase and endogenous butanediol dehydrogenase genes (the 2,3-BD gene cassettes) were introduced in the S. cerevisiae pdc-deficient mutant, approximately 100 g/L levo-2,3-BD was produced from glucose and galactose substrates (Lian et al., 2014). Nonetheless, the pdc-deficient S. cerevisiae exhibited growth defect stemming from lack of acetyl-coA, the precursor for fatty acids and some amino acids, in the cytosol of S. cerevisiae (Pronk et al., 1996; Steensmati et al., 1996; Flikweert et al., 1999). Accumulation of cytosolic NADH stemming from redox imbalance due to blockage of ethanol production pathway where acetaldehyde acts as ‘electron sink’ to dissipate NADH and regenerate NAD+ for glycolysis continuation
(Van Maris et al., 2004; Kim et al., 2013). The available acetyl-coA in the pdc-deficient
S. cerevisiae is synthesized only in the mitochondria since pyruvate dehydrogenase is solely found in the mitochondria and cannot permeate the mitochondria membranes (Van
38
Roermund et al., 1995; Kim et al., 2016). To overcome poor growth in pdc-deficient S. cerevisiae while enhancing 2,3-BD production, Kim et al. (2016) overexpressed pdc genes of Crabtree-negative yeasts (Candida tropicalis and Kluyveromyces marxianus) and NADH oxidase from Lactococcus lactis (noxE) in the mitochondria of pdc-deficient
S. cerevisiae. Notably, the Crabtree effect occurs when respiration is inhibited and completely overtaken by the fermentative process during the exponential growth phase of yeast grown aerobically on highly glucose or fructose concentrated media (De Deken,
1966). Since the PDC of C. tropicalis exhibited lower activity under high concentrations of glucose and fructose, and aeration, compared to Crabtree positive yeasts (van Urk et al., 1990; Postma et al., 1989; De Deken, 1966), Kim et al. (2016) expressed the pdc gene of C. tropicalis in pdc-deficient S. cerevisiae followed by gene expression optimization under different promoters, and obtained approximately 121.8 g/L 2,3-BD in a fed-batch fermentation. Although, glycerol accumulated in the cultures due to excess NADH, the expression of noxE of L. lactis in the pdc-deficient S. cerevisiae resulted in decreased glycerol production with accumulation of acetoin (Kim et al., 2016). The noxE gene, encoding the NADH oxidase NoxE, catalyzes NADH conversion to NAD+ and hydrogen gas, and its expression in pdc-deficient S. cerevisiae decreased the intracellular level of
NADH required for acetoin reduction to 2,3-BD. However, the expression of noxE was not optimized under different promoters. Therefore, optimization of NoxE expression in pdc-deficient S. cerevisiae may result in a balance between NADH oxidation to NAD+ and hydrogen and its availability for acetoin reduction to 2,3-BD. However, manipulations of PDC expression together with the expression of NADH oxidase in pdc-
39 deficient S. cerevisiae resulted in improved cell growth and production of 154.3 g/L 2,3-
BD, amounting up to 81 % theoretical 2,3-BD yield (Kim et al., 2016).
Heterologous gene transfers do not come without challenges. In some occasions, the final heterologous gene products maybe somewhat different to the hosts native gene products, and might be caused by the expression strategies employed (Siest et al., 1993).
In some instances, foreign genes that are cloned into a non-native microorganism may have dissimilar specific signature sequences recognized by the restriction endonucleases of the heterologous host or the foreign gene could introduce extra restriction sites not recognized by the host (Thomas and Nielsen, 2005). These could result in the destruction of the foreign genes. However, the problem can be overcome through methylation of the foreign DNA (Moser et al., 1993).
2.12 Challenges of microbial 2,3-BD production from lignocellulosic biomass
The use of conventional sugars derived from food crops such as corn and sugarcane to produce biofuels and chemicals has sparked the food versus fuel debate.
Biofuels production from food crops have been identified as a major player in the 2006-
2008 commodity price boom including the hike in food prices (Zilberman et al., 2012;
Richard et al., 2012). For instance, increased biofuel production from food crops in the
United States contributed to 20-25 % increase in the price of corn and 7-8 % increase in the price of soybeans between 2001 and 2007 (Zilberman et al., 2012). One major factor that influences the food/fuel prices is the demand for biofuels. Indeed, energy prices, economic growth, and increased demand for biofuels increase demand for food crops used in biofuel production, with concomitant increase in food prices (Zilberman et al.,
40
2012; Richard et al., 2012). For instance, an increase in energy price will have direct impact on the cost of transportation, which may serve as incentive for increasing biofuel production with overall impact on food prices (Mitchell, 2008).
To reduce the impact of producing biofuels and chemicals from food crops on food prices, it is important to use non-food crops and agricultural residues for biofuel and chemical production. One important example of agricultural residues is readily available cheap lignocellulosic biomass (LB) which is also an attractive feedstock for 2,3-BD production (Lee et al., 2015). LB offers great promise to replace food crops as substrates for the production of biofuels and chemicals, and reduce cost. Notably, LB is composed mainly of cellulose and hemicellulose embedded in a lignin matrix (Lee et al., 2004). The lignin matrix helps protect the cellulose and hemicellulose components of LB from microbial degradation. Thus, the lignin matrix must be deconstructed to help hydrolytic enzymes gain access to cellulose and hemicellulose components of LB and facilitate their hydrolysis to fermentable sugars. However, pretreatment methods used in the deconstruction of the lignin matrix release lignocellulose-derived microbial inhibitory compounds (LDMICs) that inhibit the growth of fermentation microorganisms (Delgenes et al., 1996; Palmqvist and Hahn-Hagerdal, 2000). Examples of LDMICs generated during LB pretreatment include furans (furfural and 5-hydroxylmethylfural), acetic acid, phenolic compounds including vanillin, syringylaldehyde, cinnamaldehyde, and ferulic, coumaric and vanillic acids (Fig 2.5; Ujor et al., 2015; Palmqvist and Hahn-Hagerdal,
2000). LDMICs from LB hydrolysates can be removed prior to fermentation. However, current methods of LDMICs removal prior to fermentation are cost ineffective and often
41 result in the loss of fermentable sugars (Frazer and McCaskey, 1989; Martinez et al.,
2000).
The effect of LDMICs on ethanol and butanol producing microorganisms have been studied extensively, however, studies on the effects of various LDMICs on 2,3-BD producing microorganisms and 2,3-BD production are sparse. Some studies have reported the use of LB hydrolysates for 2,3-BD production but unclear on whether or not
LDMICs were removed prior to 2,3-BD fermentation. However, some other studies have investigated the effects of 2,3-BD production from LB using different 2,3-BD producing microorganisms. A study conducted by Wu et al. (2013) evaluated the effects of acetic acid (0, 5 and 10 g/L), furfural (0, 0.5 and 1 g/L), HMF (0, 1 and 2 g/L), and combinations thereof on the growth of K. oxytoca and 2,3-BD production using a central composite design (CCD). Acetic acid, furfural and HMF were found to have negative effects on the growth of K. oxytoca, with furfural being the most toxic. From the model generated from the CCD, furfural affected 2.3-BD yield whereas HMF had no effect on
2,3-BD yield. Acetic acid concentration of ≤ 7.5 g/L enhanced 2,3-BD yield and the authors concluded that removal of furfural prior to LB hydrolysate fermentation would improve cell biomass and 2,3-BD yield by K. oxytoca.
In another study, Huang et al. (2013) investigated the production of 2,3-BD by
Klebsiella sp. using detoxified or conditioned rice straw hydrolysate (RSH). The RSH was detoxified with Ca(OH)2 (overliming-treatment hydrolysate) or conditioned with
NaOH. The Ca(OH)2 treated RSH contained no furfural or HMF, and 85% of sugars was utilized by Klebsiella sp. resulting in 2,3-BD yield of 3.2 g/g. The NaOH-treated RSH
42 contained 0.51 and 0.49 g/L furfural and HMF, respectively, and sugar utilization in the
NaOH treated RSH was lower when compared to the Ca(OH)2-treated RSH. Although sugar utilization by Klebsiella sp. grown in NaOH-treated RSH was low, 2,3-BD yield was ~28% more than that obtained in the Ca(OH)2 treated RSH. The authors suggested that the presence of salts generated from the NaOH and inhibitors may have enhanced sugar conversion to 2,3-BD resulting in increased 2,3-BD yield when compared to the
Ca(OH)2- treated RSH.
Investigation on the effects of LDMICs on the growth and 2,3-BD production by
E. aerogenes revealed that furfural (1-4 g/L) and HMF (1-3 g/L) decrease the growth of
E. aerogenes and 2,3-BD production (Lee et al., 2015). Growth and 2,3-BD production were completely inhibited at 5 g/L each of furfural and HMF. Lee et al. (2015) further tested the effects of coumaric acid and syringaldehyde (phenolic compounds) on the growth of E. aerogenes and 2,3-BD production. Coumaric acid drastically inhibited growth of E. aerogenes and 2,3-BD production at 0.5 g/L coumaric acid, and experienced complete growth inhibition and 2,3-BD production at 1.5 g/L coumaric acid. Further, >
1.5 g/L syringaldehyde completely inhibited the growth of E. aerogenes. The assay of acetolactate synthase, acetolactate decarboxylase and butanediol dehydrogenase (the 2,3-
BD biosynthesis enzymes) in the presence of furfural, HMF and phenolic compounds show significant reduction in enzyme activities of E. aerogenes with increasing concentrations of inhibitors, suggesting the negative impacts LDMICs have on 2,3-BD biosynthesis. For P. polymyxa, furfural and HMF concentrations below 1 g/L improved the production of 2,3-BD by P. polymyxa S-07 but when concentration of furfural or
43
HMF exceeds 1 g/L, 2,3-BD production by P. polymyxa S-07 was significantly inhibited
(Wang and Chen, 2014). Phenolic compounds at concentrations above 2.6 g/L severely affected the growth of P. polymyxa S-07 and 2,3-BD production (Wang and Chen, 2014).
Carbon catabolite repression (CCR) is another challenge facing the use of LB hydrolysates as substrate for the production of biofuels and chemicals as CCR phenomenon may cause incomplete or prolonged sugar utilization during fermentation.
Slow or incomplete sugar utilization leads to decreased 2,3-BD yield and productivity
(Bothast et al., 1999; Kim et al., 2010). Some 2,3-BD producing microorganisms experience CCR in which glucose represses the expression of enzymes and transporter proteins for other sugars in the LB hydrolysate. For instance, E. cloacae and K. oxytoca showed CCR effect during fermentation of corn fiber hydrolysate and mixed glucose and xylose to 2,3-BD, respectively, where xylose was utilized only when glucose was completely depleted (Saha and Bothast, 1999; Ji et al., 2009). To eliminate CCR effect in
K. oxytoca, Ji et al. (2011b) overexpressed crp (in) gene that encodes mutant cAMP receptor protein, CRP (in), that does not require cAMP to function in K. oxytoca. In the presence of glucose and xylose, PTS (EIIAGlc) protein of K. oxytoca is dephosphorylated and in the dephosphorylated state binds to non-PTS permeases and inhibit the uptake of non-PTS sugars (Gosset, 2005; Ji et al., 2011b). However, when glucose is absent, the
IIA component of PTS (EIIAGlc) is in its phosphorylated state bound to adenylate cyclase, leading to the biosynthesis of cAMP. Further, cAMP binds to CRP and triggers the induction of non-PTS sugar uptake (Gosset, 2005; Ji et al., 2011b). Therefore, Ji et al.
(2011b) overexpressed a mutant CRP that does not require cAMP to induce uptake of
44 xylose in K. oxytoca. The K. oxytoca mutant harboring the crp(in) gene showed simultaneous utilization of glucose and xylose resulting in approximately 29% increase in
2,3-BD productivity when compared to the wildtype. To make 2,3-BD production from
LB more economical, it is important to identify or develop microorganisms with intrinsic/extrinsic capability of LDMICs tolerance during conversion of LB hydrolysates to 2,3-BD. Further, it is crucial to develop CCR- negative 2,3-BD microorganisms for efficient LB hydrolysate conversion to 2,3-BD.
2.13 Microbial 2,3-BD recovery
Several techniques for recovering 2,3-BD from fermentation broth have been reported. Simple distillation has been used extensively for industrial-scale recovery of bioproducts from fermentation broths. However, bioproducts that are commonly purified by distillation must possess acceptable physical attributes e.g., hydrophobicity/ hydrophilicity, volatility and low boiling points. Unfortunately, recovery of 2,3-BD by conventional distillation methods is hindered by two important physical properties of 2,3-
BD– its high boiling point (177 to 184°C) and water affinity. Consequently, 2,3-BD distillation must be conducted for a prolonged duration and at high temperatures to effectively remove bound water molecules (Maddox, 1996). This process is untenable due to high energy cost as well as resultant discolored distillate. The dark distillate likely stems from heat-induced conversion of fermentation medium components and residual by-products of colored fermentation intermediates. To generate colorless 2,3-BD with acceptable market standard, the distillate must be subjected to downstream purification
45 processes. These additional steps renders distillation an ineffectual purification method for commercial-scale 2,3-BD production.
Some alternative 2,3-BD recovery strategies reported in the literature include: pervaporation, solvent extraction, reverse osmosis, aqueous two-phase system (ATPS), reactive 2,3-BD extraction, and steam stripping (Qureshi et al., 1994; Li et al., 2010b; Li et al., 2013b; Xiu and Zeng, 2008). Pervaporation, or vacuum membrane distillation, facilitates selective vaporization of 2,3-BD from impure mixtures through a membrane.
The process is performed under vacuum, which creates a concentration gradient between the opposite phases of the membrane and allows selective passage of 2,3-BD (Neel,
1991). Qureshi et al. (1994) used a nonporous polytetrafluoroethylene (PTFE) membrane under vacuum to concentrate up to 430 g/L 2,3-BD from fermentation broth in a fed- batch culture. However, the technique does not recover all of the 2,3-BD from fermentation broth.
Solvent or liquid-liquid extraction is based on a principle that certain solutes can solubilize and partition in specific ratios between immiscible aqueous and organic solvents (Anthemidis and Ioannou, 2009). 2,3-BD partitions in solvents such as ethyl acetate, diethyl ether and dodecanol. These solvents have been used for extraction of 2,3-
BD from fermentation broth (Eiteman and Gainer, 1989; Maddox, 1996; Syu, 2001).
However, solvent extraction technique does not recover all 2,3-BD from fermentation broth (68 to 75% recovery; Anvari and Khayati, 2009; Tsao, 1978). Similar to solvent extraction, aqueous two-phase system (ATPS) or salting-out techniques rely on hydrophilic solvents (e.g., ethanol, methanol, or isopropanol) and inorganic salts to
46 recover 2,3-BD from fermentation broth. This technique is based on the partitioning of
2,3-BD between two-phases created by the hydrophilic solvent and inorganic salt solution
(Jiang et al., 2009). Inorganic salts such as potassium phosphate, potassium carbonate, or ammonium sulfate salts have been used in ATPS to enrich up to 87% 2,3-BD (Li et al.,
2010b; Jiang et al., 2009; Sun et al., 2009). Unfortunately, glucose and other fermentation products may also partition with 2,3-BD, which necessitates a secondary purification method. Also, there is no information on large-scale ATPS use.
An integrated scheme that uses both solvent extraction and pervaporation to recover 2,3-BD from fermentation broth was proposed (Shao and Kumar, 2009). The solvent, 1-butanol, was used to extract a mixture of 2,3-BD and water from fermentation broth. Then, 1-butanol and water were selectively excluded with the use of a composite membrane consisting of polydimethylsiloxane (PDMS) and chitosan (Shao and Kumar,
2009). Steam stripping and reverse osmosis for 2,3-BD recovery have also been tested
(Table 2.2). Use of aldehydes in reactive 2,3-BD extraction have been demonstrated (Li et al., 2013c). This technique uses ion-exchange resin HZ732 (as catalyst) and aldehydes
(as both reactants and extractants) to recover 2,3-BD from fermentation broth. Examples of aldehydes employed in the method include n-butylaldehyde, propionaldehyde and acetaldehyde (Li et al., 2013c). The technique is based on the principle that the two hydroxyl groups on 2,3-BD can enable formation of dioxolanes with aldehydes. The dioxolane formed in the process extracts into excess aldehydes or organic solvents e.g., cyclohexane. Next, the dioxolanes produced are hydrolyzed to aldehyde, 2,3-BD and water (in the absence of the catalyst) after which 2,3-BD is recovered by vacuum
47 distillation. However, the method has some drawbacks (see Table 2.2). Owing to the complexity of microbial 2,3-BD recovery from fermentation broth, methods that can convert microbial 2,3-BD in the fermentation broth directly to an easily extractable intermediate compound will reduce cost of 2,3-BD production. Reactive extraction of
2,3-BD is possible only when 2,3-BD is reversibly converted into a volatile compound that can be readily extracted.
Another potential strategy to recover 2,3-BD from fermentation broth is to convert 2,3-BD directly to methyl ethyl ketone (MEK) while it is still in the fermentation broth using acid that has sulfonic groups covalently bound to a matrix as catalyst
(Emerson et al; 1982; Tran and Chambers, 1987; Maddox, 1996). The MEK produced can be readily recovered by distillation due to its low boiling point (~80 °C). The drawback to this method is that MEK cannot be reconverted to 2,3-BD. However, the technique is handy when MEK and 1,3-BD are the desired end products of microbial 2,3-
BD production. A major application of 2,3-BD is in the production of MEK, a feedstock chemical used in surface coatings, chemical intermediates, printing inks, and adhesives.
2.14 Simultaneous 2,3-butanediol fermentation and recovery
Microorganisms that produce 2,3-BD can re-utilize the compound as a carbon source, which leads to less extractable 2,3-BD (Okonkwo et al., 2017; Ghosh and
Swaminathan, 2003). Techniques that facilitate simultaneous 2,3-BD fermentation and recovery (SFR) are required to prevent re-utilization of 2,3-BD after production but these methods have challenges. For example, in situ recovery techniques that require
48 temperatures above the fermentation temperature would kill the fermenting microorganism and add to the heating cost of 2,3-BD production during SFR. While solvent/liquid-liquid extraction and pervaporation may likely be adapted and incorporated in the SFR process, use of solvents with high partition coefficients and low toxicity to the fermenting microorganisms are critical for effective use of liquid-liquid extraction process. Oleyl alcohol is one such solvent with low toxicity on K. pneumoniae. Its use in
SFR increased 2,3-BD production from 17.9 g/L in conventional fermentation to 23.01 g/L in extractive 2,3-BD fermentation (Anvari and Khayati, 2009). However, the toxic effect of oleyl alcohol has not been tested in other 2,3-BD major producers.
2.15 Conclusions and perspectives
The unpredictable petroleum economy coupled with the finite nature of crude oil reserves, and environmental issues associated with fossil fuel use, warrant the development of alternative processes with low carbon footprint for the production of normally petrochemical-derived products such as 2,3-BD. Development of hyper 2,3-BD producing microbial strains that produce less competing products may support increases in 2,3-BD titer and yield sufficient for industrial-scale production. Also, increasing the ability of 2,3-BD producing microbes to utilize a wide range of substrates (e.g. lignocellulosic biomass) will be an added advantage and may help bring commercial microbial 2,3-BD production to fruition. Given that the cost of 2,3-BD production is largely influenced by costs associated with 2,3-BD recovery from the fermentation broth, it is expected that the use of cheap and readily available substrates and 2,3-BD producers
49 that do not make EPS, LPS or CPS, will drastically reduce the cost of microbial 2,3-BD production. To achieve this goal and develop next generation bio-catalysts for efficient
2,3-BD fermentation processes, a holistic approach that combines metabolic perturbation in 2,3-BD producing microorganisms, deletion of metabolic pathways for the production of competing products, introduction of efficient 2,3-BD biosynthesis machinery, optimization of fermentation medium and conditions, may be necessary.
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Table 2.1. EPS production by major 2,3-BD producers Species/strains Substrate 2,3-BD 2,3-BD EPS (g/L) EPS type Fermentation Reference (g/L) productivi mode ty (g/L/h) Klebsiella Glucose 50.9 0.51 Yes Capsular Fed-batch Cho et al., 2012 pneumoniae exopolysaccharide K. oxytoca Glucose 34.1 0.34 No - Fed-batch Cho et al., 2012 K. pneumoniae Glucose 31.3 0.66 Yes Lipopolysaccharid Batch Jung et al., 2013 KCTC 2242 e Serratia Glucose 44.0 1.83 Yes Serrawettin Batch Zhang et al., 2010b marcescens K. pneumoniae Glucose 9.6* - Yes; 34.9* - Fed-batch Ramirez-Castillo and sp. pneumoniae Uribelarrea, 2004 K63 Paenibacillus Sucrose 52.0 1.08 Yes. 54 - Fed-batch Häßler et al., 2012 polymyxa DSM
75 365
Bacillus Sucrose - - Yes; 47.45 Levan Batch Liu et al., 2010a. licheniformis 8- 37-0-1 B. licheniformis Kimchi - - Yes; 8.02 Exopolysaccharide Batch Song et al., 2011 KS-20 s B. polymyxa Sucrose - - Yes; 54.6 Exopolysaccharide Fed-batch Lee et al., 1997 KCTC 8648P s B. polymyxa Glucose - - Yes; 9.0 Exopolysaccharide Batch Lee et al., 1997 KCTC 8648P s B. polymyxa Fructose - - Yes; 4.8 Exopolysaccharide Batch Lee et al., 1997 KCTC 8648P s
Continued below table
75
Table 2.1 Continued above table
Species/strains Substrate 2,3-BD 2,3-BD EPS (g/L) EPS type Fermentation Reference (g/L) productivi mode ty (g/L/h) B. polymyxa Galactose - - Yes; 4.8 Exopolysaccharide Batch Lee et al., 1997 KCTC 8648P s B. polymyxa Lactose - - Yes; 4.6 Exopolysaccharide Batch Lee et al., 1997 KCTC 8648P s B. polymyxa Soluble - - Yes; 6.0 Exopolysaccharide Batch Lee et al., 1997 KCTC 8648P starch s P. polymyxa Xylose - - Yes; 3.34 Exopolysaccharide Batch Liu et al., 2009 EJS-3 s P. polymyxa Maltose - - Yes; 3.61 Exopolysaccharide Batch Liu et al., 2009 EJS-3 s
76
76
Table 2.2. Comparative techniques for 2,3-BD recovery and purification. Separation Principle of operation/application Caveats References techniques/met hods
Distillation/ev Distillation and evaporation are used for Requires deproteinization and desalting prior to Maddox, aporation removal of large volume of water from distillation which adds to 2,3-BD recovery cost; 1996; fermentation broth prior to 2,3-BD recovery. removal of large quantity of water is an energy intensive process; recovered 2,3-BD is impure, colored and viscous and needs additional purification steps.
Pervaporation Non-porous PTFE membrane has been shown Application of pervaporation to fermentation Qureshi et to concentrate 2,3-BD allowing passage of broth results in membrane fouling by proteins al., 1994 water and butanol when subjected under and salts. The technique does not recover all 2,3- reduced pressure. BD.
77
Liquid-liquid Some organic solvents allows 2,3-BD to No solvents that can extract 2,3-BD have been Tsao, 1978; extraction/ distribute in them and enable extraction of 2,3- found; sugars and some fermentation solvents Eiteman Solvent BD. Nonetheless, the degree of 2,3-BD can distribute in solvents used for 2,3-BD and Gainer, extraction distribution in the solvents vary which affects separation; use of non-volatile solvents for 2,3- 1989, 2,3-BD recovery BD recovery makes purification of 2,3-BD from the solvents difficult.
Reactive Aldehydes reacts reversibly with 2,3-BD to Requires fermentation broth pretreatment to Li et al., extraction form dioxolanes in the presence of HZ732 ion- remove proteins and cells; 2,3-BD is collected 2012; exchange resin as catalyst. Dioxolanes are into the aqueous phase after dioxolanes 2013b; readily hydrolyzed to aldehydes and 2,3-BD hydrolysis and requires vacuum-distillation for 2013c further purifications thus, increasing cost of purification. Corrosion caused by acidity of the system delimits industrial application Continued below table
77
Table 2.2 Continued above table
Separation Principle of operation/application Caveats References techniques/met hods
Aqueous two- The technique uses the principle that short 2,3-BD does not distribute completely in the Jiang, et phase system chain alcohols can form stable and adjustable alcohol phase and thus is not entirely recovered. al., 2009; (ATPS) two-phase system with inorganic salts There is lack of information on the scale-up of Li et al., therefore allowing 2,3-BD to distribute the process. 2010b; between the two-phase system enabling 2,3- BD recovery
Steam Steam stripping is used to remove volatile 2,3-BD is recovered with large amount of water Blom et al., stripping organic compounds from water. The technique and impurities which require further purification 1945; has been applied to recover 2,3-BD under steps. Wheat et pressure when passed through a stripping al., 1948 78 column in a counter-current manner.
Reverse Water from broth is forced through a semi- Concentration polarization is a major problem Stanbury et osmosis permeable membrane in a direction that is which results from fermentation products that are al., 1995. opposite the osmotic forces via an applied retained in the membrane pressure to concentrated 2,3-BD that is further Sridhar, extracted using solvents. 1989
78
Figure 2.1. Comparison of 2,3-BD produced by wildtype and serrawettin mutant strains of Serratia marcescens and the corresponding foam heights obtained during 2,3-BD fermentation. SM and Serrawetin M represent serrawetin mutant strain whereas WT represents the wildtype strain (Zhang et al., 2010a; Zhang et al., 2010b).
79
Figure 2.2. Schematic representation of proposed EPS biosynthesis in P. polymyxa from different substrate sources (modified from Looijesteijn et al., 1999). 1, levansucrase; 2, PEP/PTS; 3, phosphoglucose isomerase; 4, 6-phosphofructokinase; 5, α- phosphoglucomutase; 6, deoxy-TDP-glucose pyrophosphorylase; 7, dTDP-rhamnose biosynthetic enzyme system; 8, UDP-glucose pyrophosphorylase; 9, UDP-galactose-4- epimerase; 10, fructose PEP/PTS; 11, 1-phosphofructokinase; 12, glycolytic enzymes; 13 and 14, eps polymerase and exporting proteins; 15, FBPase; 16, pentose transporter permease; 17, xylose isomerase; 18, xylulose kinase; 19, pentose phosphate enzymes.
80
Figure 2.3: Characterization of EPS according to charges. Modified from Lembre et al.
(2012), Vuyst and Degeest (1999), Rehm (2010).
81
Figure 2.4: Chemical structures of Exopolysaccharides. (A) and (B) represent hetero- polymers and (C) and (D) represents homo-polymers. Modified from Ferreira et al.
(2015), Vereyken et al. (2003), Kadajji and Betageri (2011), Sakurai et al. (2005).
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Figure 2.5. Fermentation of LB-derived sugars to 2,3-BD. A. Biotechnological conversion of sugars from LB to 2,3-BD and its use as feedstock for producing synthetic rubber and plastics. B. Sugars obtained from lignocellulosic biomass can be fermented to
2,3-BD, however, LDMICs generated during LB pretreatment inhibit microbial growth and product (2,3-BD) formation. Modified from Clifford, 2017; Bracchita, 2012; UK
Essays, 2013.
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Chapter 3: Process development for enhanced 2,3-butanediol production by Paenibacillus polymyxa DSM 365
3.1 Abstract
While chiral 2,3-butanediol (2,3-BD) is currently receiving remarkable attention because of its numerous industrial applications in synthetic rubber, bioplastics, cosmetics, and flavor industries, 2,3-BD-mediated feedback inhibition of Paenibacillus polymyxa
DSM 365 limits accumulation of higher concentrations of 2,3-BD in the bioreactor during fermentation. Box-Behnken design, Plackett-Burman design (PBD) and response surface methodology were employed to evaluate the impacts of seven factors including tryptone, yeast extract, ammonium acetate, ammonium sulfate, and glycerol concentrations, and temperature and inoculum size on 2,3-BD production by P. polymyxa
DSM 365. Results showed that three factors; tryptone, temperature and inoculum size significantly influence 2,3-BD production (p< 0.05) by P. polymyxa. The optimal levels of tryptone, inoculum size and temperature as determined by Box-Behnken design and response surface methodology were 3.5 g/L, 9.5% and 35 °C, respectively. The optimized process was validated in batch and fed-batch fermentations in a 5-L Bioflo 3000
Bioreactor, and 51.1 and 68.5 g/L 2,3-BD, were obtained, respectively. Interestingly, production of exopolysaccharides (EPS), an undesirable co-product, was reduced by 19% when compared to fermentation conducted with un-optimized medium and conditions.
These results underscore an interplay between medium components and fermentation
84 conditions, leading to increased 2,3-BD production and decreased EPS production by P. polymyxa. Collectively, our findings demonstrate both increased 2,3-BD titer, a fundamental prerequisite to potential commercialization of fermentative 2,3-BD production using renewable feedstocks, and reduced flux of carbons towards undesirable
EPS production.
3.2.1 Introduction
The compound, 2,3-Butanediol (2,3-BD) is an industrial platform chemical with vast industrial applications, particularly for its potential use in the synthesis of 1,3- butadiene (1,3-BD), a monomer of synthetic rubber. Other applications of 2,3-BD include synthesis of methyl ethyl ketone (MEK), a fuel additive with higher heat of combustion than ethanol, and as solvents for lacquer and resins (Celinska and Grajek, 2009). Further,
2,3-BD is used as an antifreeze due to its low freezing temperature (-60 °C; Soltys et al.,
2001), an ink additive, a chemical feedstock for the production of acetoin and diacetyl, vital flavor enhancers in the food industry (Bartowsky and Henschke, 2004), and an additive in aviation fuel. Due to the finite nature of petroleum and the need to reduce society’s dependence on petroleum-derived feedstocks for industrial processes, it has become imperative to develop sustainable feedstocks such as 2,3-BD from renewable resources. At present, 2,3-BD is produced from hydrocarbons by cracking butane and 2- butene followed by hydrolysis to 2,3-BD (Mitchell and Robertson, 1954; O’Neil, 2001).
Recently, the often encountered instabilities in crude oil price have re-ignited interest in fermentative 2,3-BD production from cheap renewable feedstocks. Multifarious research
85 efforts are currently underway to increase the yield, titer, and productivity of microbe- derived 2,3-BD. These include metabolic engineering of producer organisms to produce and tolerate higher 2,3-BD concentrations, and optimization of fermentation media components and conditions for maximal 2,3-BD accumulation in the broth. In this study, we sought to optimize 2,3-BD production by Paenibacillus polymyxa DSM 365
(hereafter referred to as P. polymyxa) by assessing the impacts of both medium components and fermentation conditions on 2,3-BD accumulation.
P. polymyxa was chosen for this study due to its non-pathogenicity and capacity to produce 98% levo 2,3-BD; the industrially preferred 2,3-BD isomer due to its properties, which make it amenable to important chemical reactions that generate key industrially applicable products, such as dehydration to 1,3-BD (for synthetic rubber production), dehydrogenation to acetoin or diacetyl (flavor enhancers and essential components in fragrances), ketalization to methyl tert-butyl ether (fuel additive) and esterification to 2,3-BD diester (used as a precursor in the synthesis and compounding of cosmetics, drugs, and thermoplastic polymers (Celinska and Grajek, 2009; Garg and Jain,
1995).
To assess the 2,3-BD production capacity of P. polymyxa, we first conducted batch fermentations in 100 ml Pyrex bottles, which resulted in a maximum 2,3-BD concentration of 24 g/L (Okonkwo et al, 2017). Batch fermentation in the bioreactor (6-
L) produced 27 g/L 2,3-BD, whereas fed-batch fermentation (in the bioreactor) resulted in 47 g/L, despite excess glucose supply (Okonkwo et al., 2017). Therefore, we rationalized that in addition to other factors, 2,3-BD-mediated feedback inhibition might
86 pose a significant roadblock to the accumulation of 2,3-BD during fermentation, and this assumption was confirmed by assaying 2,3-BD toxicity against P. polymyxa in fermentation cultures (Okonkwo et al., 2017). We observed that 2,3-BD exerts a concentration-dependent toxicity on P. polymyxa with ~50 g/L 2,3-BD as the toxic threshold above which cell growth stalls considerably and the accumulated 2,3-BD is converted backwards to acetoin, the precursor of 2,3-BD, probably to alleviate 2,3-BD toxicity (Okonkwo et al., 2017). In addition, a significant portion of sugar substrates are diverted to exopolysaccharide (EPS) production during 2,3-BD fermentation, thereby lowering 2,3-BD yield and complicating its recovery from the fermentation broth
(Okonkwo et al., 2017; Häßler et al., 2012). Therefore, if P. polymyxa 2,3-BD fermentation is to reach industrial-scale, it is critical to determine the optimal conditions and medium components necessary for commercially-viable 2,3-BD accumulation and tolerance during fermentation. Further, cheaply available substrates such as glycerol, which is currently accumulated in excess as a by-product of biodiesel production
(Johnson and Taconi, 2007; Gerpen, 2005) holds significant promise towards improving the economics of 2,3-BD fermentation, either as a sole carbon source or as a supplement of glucose or other sugars. In fact, glycerol has been shown to support 2,3-BD production by Klebsiella pneumoniae as a sole carbon source (Petrov and Petrova, 2010; Biebl et al.,
1998). Thus, we investigated the optimal conditions and medium components for high
2,3-BD production by P. polymyxa using a glycerol-supplemented medium. In addition to lowering overall substrate cost, glycerol catabolism furnishes the cell with additional
NADH (Lin, 1976; Neijssel et al., 1975), which supplies extra reducing power for 2,3-BD
87 dehydrogenase, the final enzyme of the 2,3-BD pathway, which consumes NADH during conversion of acetoin to 2,3-BD (Voloch et al., 1983).
Previous optimization studies focused largely on enhancing 2,3-BD production.
These studies either targeted medium components only, or fermentation conditions without a holistic evaluation of both parameters (medium components and fermentation conditions; Häßler et al., 2012; Gao et al., 2010). Medium components and fermentation conditions such as temperature, inoculum size, pH and aeration rate most reasonably interact during fermentation to engender 2,3-BD production. Therefore, in this study, select medium components and fermentation conditions were assessed collectively for their capacity to enhance 2,3-BD production by employing various optimization strategies. Plackett-Burman experimental design, path of steepest ascent method, Box-
Behnken experimental design and response surface methodology strategies were employed to optimize 2,3-BD production by P. polymyxa. The medium components tested in this study include yeast extract, tryptone, ammonium acetate, ammonium sulfate and crude glycerol; whereas the fermentation conditions that were extensively investigated include temperature and inoculum size. These factors were shown to influence 2,3-BD production by P. polymyxa from our one-factor-at-a-time experiments.
3.3 Materials and Methods
3.3.1. Experimental methods
3.3.1.1. Microorganism and culture preparation
P. polymyxa was obtained from the German Collection of Microorganisms and
Cell Culture, Braunschweig, Germany (DSMZ- Deutsche Sammlung von
88
Mikroorganismen und Zellkulturen). Lyophilized cells were reactivated by inoculating into Luria bertani (LB) broth, grown overnight (12 h), and then stored as glycerol (50% sterile glycerol) stock at –80 °C until further use. Glucose, yeast extract and tryptone were prepared and sterilized separately at 121°C for 15 min followed by cooling to 50 °C prior to mixing with filter-sterilized components (buffer and trace element solution), and this mixture forms the final pre-culture medium. For inoculum preparation, 1 ml of P. polymyxa glycerol stock was inoculated into 30 ml of pre-culture medium. The pre- culture medium contained (g/L); 20.0 glucose, 5.0 yeast extract (YE; Sigma-Aldrich, St louis, MO), and 5.0 tryptone (Sigma-Aldrich, St. Louis, MO), 0.2 MgSO4, and 3.0
(NH4)2SO4. The pre-culture was supplemented with 0.9 ml of phosphate buffer (pH 6.5) and 0.09 ml trace element solution. The phosphate buffer (pH 6.5) contained (g/L); 3.5
KH2PO4, 2.75 K2HPO4, while the trace element solution was prepared by dissolving 0.4 g
FeSO4 in 3 ml 25% HCl, followed by the addition of 500 ml double-distilled H2O and
(g); 0.8 H3BO3, 0.04 CuSO4.5H2O, 0.04 NaMoO4.2H2O, 5.0 MnCl2.4H2O, 0.1
ZnSO4.7H2O, 0.08 Co (NO3)2.6H2O, 1.0 CaCl2. 2H2O, and 0.01 biotin. The trace element solution was made up to 1 L with double-distilled H2O. The pre-culture was incubated aerobically at 37 °C and 200 rpm for 10-12 h in an incubator shaker (New Brunswick
Scientific, Edison, NJ). When optical density (OD600nm) of the pre-culture reached 1.0 -
1.2, 30 ml (10 ml each) of actively growing cells were distributed in three 250 ml flasks containing 90 ml sterile pre-culture medium each and incubated for another 2-3 h until
OD600nm reached 1.0 - 1.2, and then the pre-culture was transferred to production medium.
89
Phosphate buffer and trace element solution were prepared separately and filter-sterilized using 0.22 µm PES filter (Corning Incorporated, Corning, NY).
3.3.1.2. Batch and fed-batch fermentations
Batch and fed-batch fermentations were conducted in a 5-L Bioflo 3000
Bioreactor (New Brunswick Scientific, Edison, NJ) with a 2-L starting volume. The bioreactor was equipped with sensors for measuring pH and temperature and stirrers for medium agitation. The medium was continuously stirred by means of 2 Rushton impellers (3-plate). Fermentations were conducted aerobically by sparging sterile air into the medium at a flow rate of 150 ml/min through a 0.2 µm PTFE Acro®50 sterile filter
(Pall Corporation, Ann Arbor, MI) using a Masterflex L/S® Pump (Cole-Parmer
Instrument Company, Vernon Hills, IL) through the top of the bioreactor into the fermentation medium. In addition to concentrations of the medium components studied, the production medium contained (g/L): 120 glucose, 3.5 KH2PO4, 2.75 K2HPO4, 0.2
MgSO4, 0.05 CoCl2, 10.0 3-(N-morpholino) propanesulfonic acid (MOPS), and 6 ml of trace element solution (described above: microorganism and culture preparation). All medium components were prepared separately and then mixed under aseptic conditions.
Fermentation was started at an initial pH of 6.5±0.1 and pH was externally controlled with 12.5% NH4OH or 6.5 N H3PO4 when pH dropped below 6.0±0.1 or increased above
6.5±0.1. The fermentation medium was stirred at 300 rpm and the culture was fed when broth glucose concentration fell below 15 g/L for fed-batch fermentations. Each round of
90 sugar feeding was accompanied by addition of half-strength of the other medium components (buffer and trace element solutions).
3.4 Analytical methods
® Cell growth was determined by measuring optical density (OD600) in a DU
Spectrophotometer (Beckman Coulter Inc., Brea, CA). Changes in pH were measured using an Acumen® Basic pH meter (Fischer Scientific, Pittsburgh, PA). The concentrations of 2,3-BD, acetoin, ethanol, and acetic acid were determined using a
7890A Agilent gas chromatograph (Agilent Technologies Inc., Wilmington, DE) equipped with a flame ionization detector (FID) and a J x W 19091 N-213 capillary column [30 m (length) x 320 µm (internal diameter) x 0.5 µm (HP-Innowax film thickness)]. The carrier gas was nitrogen, and the inlet and detector temperatures were maintained at 250 and 300 °C, respectively. The oven temperature was programmed to span from 60 to 200 °C with 20 °C min-1 increments, and a 5-min hold at 200 °C.
Samples (1 µL) were injected with a split ratio of 10:1.
Glucose concentration was determined by HPLC using a Waters 2796
Bioseparations Module equipped with an Evaporative Light Scattering Detector (ELSD;
Waters, Milford, MA) and a 9 µm Aminex HPX-87P column; 300 mm (length) x 7.8 mm
(internal diameter) connected in series to a 4.6 mm (internal diameter) x 3 cm (length)
Aminex deashing guard column (Bio-Rad, Hercules, CA). The column temperature was maintained at 65 °C. The mobile phase was HPLC-grade water maintained at a flow rate
91 of 0.6 mL/min. The amounts of EPS produced were measured using a previously described method (Okonkwo et al., 2017).
3.5.1 Experimental design and data analysis
Other authors have previously reported that yeast extract, ammonium acetate, ammonium sulfate, glycerol, and tryptone, as well as temperature and inoculum size influence microbial 2,3-BD production (Gao et al., 2010; Anvari and Motlagh, 2011;
Zhang et al., 2010; Perego et al., 2003; Marwoto et al., 2002). Some of these studies concluded that using high concentrations of expensive yeast extract (up 60 g/L) was crucial for optimal 2,3-BD production. Therefore, we first conducted one-factor-at-time experiments to evaluate the degrees of effect exerted by these factors on the 2,3-BD production capacity of P. polymyxa. The one-factor-at-a-time experiments underlined the effects of yeast extract, ammonium acetate, ammonium sulfate, glycerol, and tryptone, as well as temperature and inoculum size on 2,3-BD production by P. polymyxa; hence, we employed Plackett-Burman design, path of steepest ascent, Box-Behnken and response surface methodology for further optimization studies.
3.5.2 Plackett-Burman design
Placket-Burman design allowed for the evaluation of important factors that influence 2,3-BD production based on the assumption that the selected factors do not interact. In this design, each factor was defined at two levels; a high (+1) and a low (−1),
92 which represent two different concentrations or condition set points. The actual experimental values were defined according to the equation;
Xi = xi – x0/ ∆xi (I = 1, 2, 3,…, k)
(1)
From the equation above, Xi is the defined value of an independent factor such as inoculum size, temperature, glycerol, tryptone, yeast extract, ammonium acetate and ammonium sulfate, xi is the real value of an independent factor, and x0 represents the real value of an independent factor at a center point value. Further, ∆xi is the difference between the real value at the center point (x0) and the real values at the lower or upper point of an independent factor. The data obtained from the design were fitted to a first- order model for 2,3-BD production as shown in the equation below;
Y = β0 + ∑βi Xi
(2)
Y is the concentration of 2,3- BD obtained from each experimental run, while β0 is the intercept, and Xi represents the ith factor (X1 – X7; see Table 3.1) and βi is the regression coefficient of each factor (X1 – X7; see Table 3.1) (Khuri and Mukhopadhyay,
2010). The resulting data were then analyzed and fitted to a linear regression using
Design Expert software package (version 10.0, Stat-Ease, Inc. Minneapolis, USA).
Analysis of variance (ANOVA) at 95% confidence interval (P< 0.05) was used to determine significant factors. The significant factors were chosen for path of steepest ascent experiment.
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3.5.3 Path of steepest ascent
The path of steepest ascent method enables the determination of three optimal levels at which to further optimize each of the significant factors obtained from the
Plackett-Burman design. This is carried out by moving the center point value of each factor sequentially along the path of steepest ascent until no further increase in 2,3-BD is obtained; i.e., the center point value of each selected factor is either increased or decreased until the maximum 2,3-BD achieved begins to decline (Box et al., 1978; Chang et al., 2002).
3.5.4 Box-Behnken design and response surface methodology
The optimal nutrient concentrations and fermentation conditions for maximum
2,3-BD production were determined by employing Box-Behnken design and response surface methodology. The three significant factors - tryptone, temperature and inoculum size - selected from the Plackett-Burman design were varied at three levels. These factors were shown to exert the most significant effects on 2,3-BD production. Box-Behnken design, when integrated with response surface methodology quantifies the relationship between the independent input factors and the obtained response surfaces (Kwak, 2005).
In this study, the association between the responses and the three important factors were determined according to the second order polynomial function:
2 Y = β0 + ∑βi Xi + ∑βii X i + ∑βij Xi Xj (i,j = 1,2…,k)
(3)
In the equation above, Y is the calculated 2,3-BD response function and β0 is the estimated regression coefficient of the fitted response at the center point of the design,
94 while Xi represents the corresponding actual value factors for inoculum size, temperature and tryptone. The regression coefficient for the linear terms is represented by βi, whereas
βij is the interaction effect and βii is the quadratic effect. Design Expert software package
(Version 10.0, Stat-Ease Inc., Minneapolis, MN) was used to calculate and analyze the second-order polynomial coefficients. Analysis of variance (ANOVA) was used to test the significance of independent factors (tryptone, inoculum size, temperature) and their interactions at an alpha (α) level of 0.05.
3.6 Results and discussion
3.6.1 Plackett-Burman design
To obtain optimized medium and fermentation conditions for improved 2,3-BD production by P. polymyxa, medium components and fermentation conditions were evaluated. Nutrient components including tryptone, glycerol, ammonium sulfate, and other fermentation parameters (temperature and inoculum size) were studied to determine the extent to which these factors impact 2,3-BD production by P. polymyxa. All media contained crude glycerol. Crude glycerol served a dual purpose in the medium - a source of carbon and an additional source of NADH, as glycerol catabolism generates two additional molecules of NADH, relative to glucose on a molar basis (Lin, 1976; Neijssel et al., 1975). The reducing equivalent furnished by NADH is critical for 2,3-BD biosynthesis, as NADH is required for the reduction of acetoin to 2,3-BD (Xiao and Xu,
2007; Li et al., 2013).
Tryptone was used as an organic nitrogen source in addition to yeast extract, whereas ammonium acetate and ammonium sulfate served as inorganic sources of
95 nitrogen. For fermentation conditions, temperature and inoculum size were selected for investigation. Like most microbial processes, 2,3-BD biosynthesis is enzyme-controlled, therefore, fermentation temperature impacts substrate consumption and 2,3-BD production, because enzyme activity is temperature-dependent (Perego et al., 2003;
Marwoto et al., 2002). Inoculum size has been reported to increase substrate utilization with improved 2,3-BD production and yield (Perego et al., 2003; Nilegaonkar et al.,
1992). To our knowledge, this is the first report of combined optimization of fermentation nutrients and conditions for enhanced 2,3-BD production by P. polymyxa.
To determine the factors with significant influence on 2,3-BD production, Plackett-
Burman design was employed to test each of the factors at two levels. Our preliminary experiments involving each factor (one-factor-at-a-time experiment) showed that the concentrations of yeast extract, tryptone, ammonium acetate, ammonium sulfate, and glycerol that had significant influence on 2,3-BD production were approximately 5.0, 5.0,
4.0, 3.0, and 7.0 g/L, respectively (Fig A.1-A.8). Furthermore, the approximate temperature and inoculum size that exerted marked effects on 2,3-BD accumulation from the one-factor at a time experiments were 36 °C and 8 %, respectively (Fig A.1-A.8).
The results obtained from our one-factor at a time experiments informed the selection of two levels for each factor, which were then tested using the Plackett-Burman design. The two levels for each of the factors were determined by employing equation 1 above; designated as low and high levels, respectively, as shown in Table 3.1. The
ANOVA generated from the experimental runs using Plackett-Burman design is shown in
Table 3.1. The linear regression coefficient of the model, R2 was 0.9993 and the adjusted
96 determination coefficient, Adj R2, was 0.9951, which are both significantly close to unity, indicating the robustness of the model for further studies. R2 measures variations in 2,3-
BD response that are explained by the tested factors for a linear regression model, whereas Adj R2 is the measure of goodness-of-fit for the model. As shown in Table 3.1, the p-values for tryptone, temperature and inoculum size were 0.0377, 0.0348 and
0.0260, respectively, indicating that tryptone, temperature and inoculum size were the most important factors that influenced 2,3-BD production by P. polymyxa among the factors studied, at 95% confidence interval.
Among the medium components tested, tryptone exerted the most significant effect on 2,3-BD production by P. polymyxa. However, it was observed that further increase in the concentration of tryptone in the fermentation medium increased biomass formation without increasing 2,3-BD production as shown by the t-value of tryptone
(Table 3.1). The pattern observed with tryptone was not unusual considering that tryptone supplies amino acids for protein (including enzymes) biosynthesis, as well as serving as a source of nitrogen for the biosynthesis of nucleic acids. In light of this, we speculated that a concentration threshold may exist for tryptone, within which both biomass and 2,3-BD accumulation by P. polymyxa occur optimally. On the other hand, increasing the temperature of the fermentation medium reduced 2,3-BD production by P. polymxa as revealed by the t-value of temperature in Table 3.1. Temperature regulation is essential for cells to function optimally, and when temperature falls below, or exceeds, the optimum range for an organism, cellular metabolism is impeded, which in this case, adversely affected 2,3-BD production (Garg and Jain, 1995). Increasing inoculum size in
97 the fermentation medium was found to positively influence 2,3-BD production by P. polymyxa as shown by the t-value of inoculum (Table 3.1). Inoculum size determines the population of viable cells in the fermentation medium at time zero, and the greater the number of cells in the medium at time zero, the shorter the lag phase of growth, which ultimately translates to a faster conversion of substrates to 2,3-BD.
3.6.2 Path of steepest ascent design
The path of steepest ascent method was used to determine the optimum levels for each of the three significant factors obtained by Plackett-Burman design. The optimum level for each of the factors is critical for Box-Behnken design and response surface methodology. For the path of steepest ascent study, the center point (value) between the low and high levels in the Plackett-Burman design was employed. The center point value for each factor in the Plackett-Burman design was moved along a path that ensures increase in 2,3-BD production. The direction for which the center point of each factor is moved was informed by the t-values shown in Table 3.1. The effects of tryptone and temperature were negative (-6.99 and -7.88, respectively), whereas that of inoculum size was positive (+10.29). These imply that to increase 2,3-BD production by P. polymyxa, the center point of each factor with positive effect needs to be increased sequentially while that with a negative effect is to be decreased until no further increase in 2,3-BD production is observed. The center points for tryptone, temperature and inoculum size were 5 g/L, 36 °C and 8%, respectively. Consequently, tryptone and temperature were decreased sequentially from 5 g/L and 36 °C to 2.5 g/L and 32 °C, respectively, while inoculum size was sequentially increased from 8% to 10.5% until no further increase in
98
2,3-BD production was observed (Table 3.2). From Table 3.2, the best three experiments in terms of 2,3-BD concentration were experiments 2, 3 and 4. The levels of each factor corresponding to experiments 2, 3 and 4 in the path of steepest ascent were selected for further optimization using Box-Behnken design and response surface methodology.
Based on these, the low, center and high values, respectively selected for temperature were 33, 34 and 35°C , and those of inoculum size were 8.5, 9.0 and 9.5%, whereas the selected concentrations for tryptone were 3.5, 4.0 and 4.5 g/L.
3.6.3 Box-Behnken design and response surface methodology
Based on the results obtained from Plackett-Burman design and path of steepest ascent method, Box-Behnken design was used to conduct 15 experimental runs to further optimize the levels of temperature, tryptone and inoculum size as shown in Table 3.3.
Data obtained from the design matrix were analyzed using multiple regressions and a second order polynomial equation model was obtained as shown below:
2 2 Y = 51.97 + 0.64X1 - 0.16X2 + 1.17X7 + 2.36X1X2 – 0.79X1X7 +0.64X2X7 - 2.31X1 - 0.71X2
2 -0.059X7
(4)
Where Y was the predicted response, X1, X2 and X7 were the defined values of inoculum size, temperature and tryptone, respectively.
Analysis of variance (ANOVA) was used to test the statistical significance of the model as shown in Table 3.4. The model had a p-value of 0.0017 which is far less than
0.05 (indicative of significance). The regression coefficient, R2 of the model was 0.9750 and the adjusted determination coefficient, Adj R2, of the model was 0.9300, implying
99 that 93% of variation in the response can be explained by the model. The lack of fit p- value was 0.5904, indicates that lack of fit was not significant, and confirms that the model was adequate for predicting 2,3-BD production. The lack of fit test is used to compare residual errors to the pure errors and gives an F value for the model
(Muthukumar et al., 2003). The F-value of the model was 21.65, which is low, thereby confirming that the model is significantly robust.
To further evaluate the optimal levels of the individual factors, the significance of each factor and their interaction terms were tested using F-test, and the corresponding P- values for each of the model terms are shown in Table 3.4. Model terms with a P-value less than 0.05 are statistically significant. The model terms for inoculum size, tryptone, inoculum size and temperature (inoculum size x temperature), inoculum size and tryptone
(inoculum size x tryptone), and the quadratic term, inoculum size and inoculum size
(inoculum size) 2 were all found to be significant. As shown in Table 3.4, tryptone exerted the largest effect on 2,3-BD production by P. polymyxa amongst the individual terms studied, whereas inoculum size and temperature (inoculum size x temperature) exhibited the greatest effect when the individual and interaction terms were compared.
The interaction effects of factors were also evaluated by response surface methodology. The response surface is a three-dimensional plot that graphically represents the regression equation and shows relationships between the response and the independent factors (Bas and Boyaci, 2007; Liu et al., 2010). Concave or convex response surfaces show that the maximum or minimum response is located within the experimental region, whereas a saddled surface shows a relative maximum and a relative
100 minimum response, respectively (Gao et al., 2010; Bezerra et al., 2008). Further, the contour plots are two-dimensional representations of the response surface, which enhance visual interpretation of the response surface (Liu et al., 2010; Bezerra et al., 2008). Plots showing elliptical contours indicate significant interactions between the independent factors and the center of the smallest ellipse refers to a point of maximum or minimum response (Bas and Boyaci, 2007). Also, plots with circular contours show that the interactions between the independent factors are negligible (Liu et al., 2010).
In the present study, the interaction between temperature and inoculum size when tryptone is maintained at the center value (4.0 g/L) shows a concave surface (Fig. 3.1a), suggesting the presence of an apparent optimum condition. The corresponding elliptical contour plot shows that the interaction between temperature and inoculum size has significant effect on 2,3-BD production by P. polymyxa (Fig. 3.1b). Additionally, the interaction between inoculum size and tryptone when temperature is kept at the center point value also shows a concave surface (Fig. 3.1c) with an elliptical contour (Fig.
3.1d), thereby indicating a significant interaction. Conversely, the interaction between temperature and tryptone shows a contour that is not elliptical and, therefore, is not significant (Fig. 3.2b). The optimum levels for the factors where maximum 2,3-BD production by P. polymyxa is predicted were obtained from the elliptical contour plot of
Fig. 3.1b, where strong interactions were observed. The maximum levels of inoculum size and temperature were indicated at the point of intersection between the major and minor axes confined by the smallest ellipse in Fig. 3.1b (Gao et al., 2010; Tanyildizi et al., 2005; Myers et al., 2009). The optimum conditions for maximum predicted 2,3-BD
101 production were calculated when the coordinates of the important points (from Fig. 3.1b) were inserted into Eq. (3), and the partial derivatives set to zero. The maximum predicted
2,3-BD was 51.5 g/L, which corresponds to a temperature of 34.98 °C and an inoculum size of 9.45%. The concentration of tryptone at this maximum predicted 2,3-BD was 3.5 g/L. The contour plots showing interactions between inoculum size and tryptone and between temperature and tryptone were not fully elliptical. The lack of a perfect elliptical contour is an indication that little or minimal interaction exists between the factors under evaluation. Thus, the optimized fermentation medium and conditions obtained in this study were 9.5% inoculum size, 3.5 g/L tryptone, and a temperature of 35 °C with the addition of yeast extract, 5 g/L; ammonium acetate, 4 g/L; NH4SO4, 3 g/L, crude glycerol, 7 g/L; KH2PO4, 3.5 g/L; K2HPO4, 2.75 g/L; CoCl2, 0.05 g/L, MgSO4, 0.2 g/L;
MOPS, 10 g/L; and trace element solution.
3.6.4 Experimental validation of the optimized medium and conditions in batch and fed-batch fermentations
The optimized fermentation medium and conditions obtained from the analyses above were then used to conduct batch and fed-batch fermentations in a 5-L bioreactor to validate the fermentation medium and conditions. In each case, fermentation was conducted with a starting volume of 2 L. As shown above, apart from inoculum size, temperature and tryptone, all the other factors that were tested by the Plackett-Burman experimental design were kept at their center point values. Batch and fed-batch fermentations were conducted in triplicate. The concentration of 2,3-BD obtained from the mean of three biological replicates using the optimized medium and conditions was
102
51.1 g/L, which was 99% of the predicted maximum 2,3-BD concentration of 51.5 g/L by response surface methodology. The yield and productivity of 2,3-BD obtained in the batch fermentation were 0.42 g/g and 1.70 g/L/h, respectively (Table 3.5). The batch fermentation profile in Fig. 3.3a shows complete glucose utilization, which is an indication of efficient glucose conversion to 2,3-BD with minimal formation of competing products. Thus, in addition to increased 2,3-BD production, the concentrations of ethanol, acetic acid, EPS and acetoin were considerably diminished (Tables 3.5 and
Fig A.1-A.8), when compared to non-optimized fermentations (Okonkwo et al., 2017).
Due to reduced diversion of carbon to the EPS and ethanol biosynthesis pathways, and most likely, increased carbon flux from acetoin and acetic acid to 2,3-BD production during batch fermentation by P. polymyxa, 2,3-BD yield increased from 0.32 g/g glucose under un-optimized fermentation conditions (Okonkwo et al., 2017) to 0.42 g/g glucose under optimized condition, accounting for a 31% increase in 2,3-BD yield.
To further determine the maximum 2,3-BD that P. polymyxa can accumulate using the optimized fermentation medium and conditions, a fed-batch fermentation was conducted. During the fed-batch process, glucose was intermittently replenished in the culture, accompanied by addition of half-strength of the other nutrient components until no further increase in 2,3-BD production or glucose consumption was observed. The maximum 2,3-BD obtained from the mean of three independent fed-batch fermentations was 68.5±4.3 g/L with yield and productivity of 0.34 g/g and 0.70 g/L/h, respectively
(Table 3.5; Fig 3.3b). Similarly, the concentrations of competing products namely acetoin, ethanol and acetic acid were considerably reduced, indicating that the optimized
103 process enabled efficient conversion of substrate carbon to the desired product, 2,3-BD.
Nonetheless, acetoin was observed to increase towards the end of the fermentation, at which point, a corresponding decline in 2,3-BD was observed; a mechanism that P. polymyxa is thought to adopt for reducing the toxicity of 2,3-BD at an elevated concentration (Okonkwo et al., 2017). Notably, the yield and productivity of 2,3-BD reduced as the fermentation mode was switched from batch to fed-batch (Table 3.5). This is not unusual considering that greater amounts of glucose are consumed in the fed-batch cultures, some of which is funneled to cell maintenance and growth. In fact, cell dilution effect resulting from intermittent feeding of glucose and other nutrients into the broth engenders a lag phase, albeit transiently. This triggered brief cell growth in which a portion of the glucose may have been used for growth thus, diverts glucose away from
2,3-BD biosynthesis, momentarily. On the other hand, the concentration of acetoin in the fed-batch fermentation increased 3-fold whereas ethanol and acetic acid increased 1.2- fold, when compared to the batch fermentation (Table 3.5). A portion of the additional glucose fed into the fed-batch cultures was further converted to ethanol and acetic acids, which increased relative to batch cultures, not fed additional glucose. Interestingly, optimized culture medium and conditions resulted in a 19% reduction in EPS production in both the batch and fed-batch fermentations (Table 3.5; Fig A.1-A.8).
A comparison of 2,3-BD concentration obtained in this study to those from other studies is summarized in Table 3.6. Different strains of P. polymyxa have been shown to possess the enzymatic repertoire needed to metabolize several carbon sources to 2,3-BD, with yields ranging from 0.33 to 0.51 g/g glucose (Table 3.6). The 2,3-BD yield of 0.42
104 and productivity of 1.70 obtained in the batch fermentation in this study compares favorably to those reported by other authors. More importantly, it is worthy of note that the yield and productivity obtained in this study were achieved using lower amount of organic nitrogen in the forms of tryptone and yeast extract when compared to the other studies, a critical economic consideration for large-scale operations.
3.7 Conclusions
Based on the results from Box-Behnken design and response surface methodology, an optimized medium (7 g/L crude glycerol included) and culture conditions for enhanced 2,3-BD production by P. polymyxa were developed. The optimized conditions were validated in batch and fed-batch fermentations, leading to production of 51.1 and 68.5 g/L, respectively, of 2,3-BD. These represent 89% and 46% increases in 2,3-BD production in batch and fed-batch cultures, respectively, with attendant diminished generation of competing co-products, especially EPS, relative to the un-optimized fermentations. The results presented here underline the interplay between medium components, culture conditions and product-mediated toxicity (feedback inhibition), as the earlier determined toxic threshold of 2,3-BD (50 g/L) on P. polymyxa in a non-optimized medium (Okonkwo et al., 2017) was significantly exceeded in this work (68.5 g/L). The glycerol incorporated in the fermentation medium used in this study may contribute to 2,3-BD biosynthesis via improved NADH regeneration, especially in the optimized medium, relative to the un-optimized control medium. Collectively, we demonstrate that lower amounts of the expensive organic nitrogen sources, tryptone and
105 yeast extract, can be used for optimal 2,3-BD production. This represents significant reduction in operating cost in efforts to commercialize biological production of 2,3-BD.
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Table 3.1: Statistical analysis of Plackett-Burman design results showing effect of medium components and fermentation conditions on 2,3-BD production by P. polymyxa.
Factor Low High % t value p value level (-1) level (+1) Contribution * X1: Inoculum size (%) 6 10 39.56 10.29 0.0260 * X2: Temperature (°C) 35 37 23.23 -7.88 0.0348 X3: CH3COONH4 (g/L) 3 5 4.87 3.61 0.0753 X4: (NH4)2SO4 (g/L) 2 4 9.30 4.99 0.0595 X5: Glycerol (g/L) 5 10 0.29 0.89 0.2112 X6: Yeast extract (g/L) 5 7 4.51 3.47 0.1174 * X7: Tryptone (g/L) 5 7 18.23 -6.99 0.0377 R2= 0.9993, Adj R2= 0.9951 *Statistical significance
111
Table 3.2: The path of steepest ascent experimental design and 2,3-BD production by P. polymyxa Run Factors 2,3-BD (g/L) Inoculum size (%) Temperature (°C) Tryptone (g/L) 1 8.0 36 5.0 46.6 2 8.5 35 4.5 47.9 3 9.0 34 4.0 52.0 4 9.5 33 3.5 48.3 5 10 32 3.0 45.5
112
Table 3.3: Box-Behnken design and response results for 2,3-BD production Run Coded values Actual values 2,3-BD X1 X2 X7 X1 (%) X2 (°C) X7 (g/L) (g/L) 1 -1 0 +1 8.5 34 4.5 50.7 2 +1 0 +1 9.5 34 4.5 51.0 3 -1 +1 0 8.5 35 4.0 46.2 4 0 0 0 9.0 34 4.0 52.6 5 0 -1 -1 9.0 33 3.5 51.0 6 -1 -1 0 8.5 33 4.0 51.0 7 0 0 0 9.0 34 4.0 52.0 8 +1 -1 0 9.5 33 4.0 47.0 9 0 +1 +1 9.0 35 4.5 52.7 10 +1 0 -1 9.5 34 3.5 50.1 11 +1 +1 0 9.5 35 4.0 51.6 12 0 +1 -1 9.0 35 3.5 49.2 13 0 0 0 9.0 34 4.0 51.4 14 0 -1 +1 9.0 33 4.5 51.9 15 -1 0 -1 8.5 34 3.5 46.7 X1, inoculum size; X2, temperature; X7, tryptone.
113
Table 3.4: ANOVA for 2,3-BD production by P. polymyxa according to the response surface quadratic model (lack of fit is not significant).
Factors Sum of Degree of Mean of squares F value p value squares freedom Model 61.55 9 6.84 21.65 0.0017* * X1 3.32 1 3.32 10.50 0.0230 X2 0.21 1 0.21 0.66 0.4539 * X7 10.90 1 10.90 34.52 0.0020 * X1 X2 22.23 1 22.23 70.38 0.0004 * X1 X7 2.46 1 2.46 7.80 0.0383 X2 X7 1.64 1 1.64 5.19 0.0718 2 * X1 19.64 1 19.64 62.18 0.0005 2 X2 1.87 1 1.87 5.91 0.0592 2 X7 0.013 1 0.013 0.040 0.8487 Residual 1.58 5 0.32 Lack of fit 0.87 3 0.29 0.82 0.5904 Pure error 0.71 2 0.35 Cor. Total 63.13 14 R2= 0.9750, Adj R2= 0.9300; Cor. = Corrected; *Statistical significance.
114
Table 3.5: The product profiles of P. polymyxa grown in batch and fed-batch fermentations under optimized conditions.
Product Batch fermentation Fed-batch fermentation profile Max. conc. Yield Productivit Max. conc. Yield Productivi (g/L) (g/g) y (g/L/h) (g/L) (g/g) ty (g/L/h) 2,3-BD 51.1±0.6 0.42±0.01 1.70±0.02 68.5±4.3 0.34±0.03 0.70±0.04 Ethanol 6.6±0.2 0.06±0.00 0.22±0.01 8.2±0.5 0.06±0.00 0.17±0.01 Acetoin 4.0±0.1 0.03±0.00 0.08±0.00 12.0±2.4 0.05±0.01 0.10±0.02 Acetic acid 1.5±0.1 0.01±0.00 0.03±0.00 1.8±0.7 0.01±0.00 0.03±0.01 EPS 5.0±0.2 0.04±0.00 0.41±0.01 4.7±0.0 0.02±0.00 0.39±0.00 Glucose 120.5±1.6 N/A N/A 200.0±8.5 N/A N/A consumed N/A: Not applicable. Error bars show standard deviations of means (n=3).
115
Table 3.6: Comparison of 2,3-BD concentrations obtained in this study to those of other studies using P. polymyxa.
Carbon 2,3-BD 2,3-BD 2,3-BD Growth Organic P. Fermentation References source (g/L) yield (g/g) productivi OD600nm nitrogen polymyxa mode ty (g/L/h) used (g/L) strain Glucose 51.1±06 0.42±0.01 1.70±0.02 11.3±0.1 YE, 5; DSM 365 Batch This work tryptone, 3.5 Glucose 68.5±4.3 0.34±0.03 0.70±0.04 11.9±0.1 YE, 5; DSM 365 Fed-batch This work tryptone, 3.5 Raw inulin 37.6±0.3 0.51 0.89 26.8±0.4* YE, 3; ZJ-9 Batch Gao et al., 2010 extract from peptone, 2 Jerusalem artichoke 116 tubers Sucrose 111.0 ND 2.06 23† YE, 60 DSM 365 Fed-batch Häßler et al., 2012 Glucose 71.7 ND 1.33 13 YE, 10 CJX518 Fed-batch Dai et al., 2014 Glucose 16.5 0.33 2.01 9.5 YE, 15 ICGEB200 Batch Adlakha and 8 Yazdani, 2015 Inulin 51.3 ND ND NS YE, 6; ZJ-9 (XG- Fed-batch Zhang et al., Peptone, 3 1) 2016 Inulin 36.8 ND ND 11† YE, 6; ZJ-9 (XG- Batch Zhang et al., Peptone, 3 1) 2016 * † Unit in g/L, Determined at OD660nm, NS-Not shown, ND- not detected. The data shown are maximum product concentrations
and cell growth achieved during fermentation.
116
Figure 3.1. Contour and response surface plots. (a) The response surface plot; (b) the resultant contour plot showing the effects of temperature and inoculum size on 2,3-BD production by P. polymyxa with tryptone fixed at 4 g/L; (c) response surface plot and (d) the resultant contour plot depicting the effects of tryptone and inoculum size on 2,3-BD production by P. polymyxa with temperature fixed at 34 °C.
117
Figure 3.2. Contour and response surface plots. (a) The response surface plot and (b) the resultant contour plot (b) showing the effects of tryptone and temperature on 2,3-BD production by P. polymyxa with inoculum size fixed at 9%.
118
119
Figure 3.3. The fermentation profiles of P. polymyxa using optimized culture medium and conditions. (a) Batch and fed-batch
(b) fermentations. Error bars show standard deviations of means (n=3).
119
Chapter 4: Investigation of relationship between 2,3-butanediol toxicity and production during growth of Paenibacillus polymyxa
4.1 Abstract
Understanding the capacity of Paenibacillus polymyxa DSM 365 to tolerate increasing concentrations of 2,3-butanediol (2,3-BD) is critical to engineering a 2,3-BD- overproducing strain. Hence, we investigated the response of P. polymyxa to high 2,3-BD concentrations. In fed-batch cultures (6-L bioreactor) 2,3-BD was accumulated to a maximum concentration of 47 g/L despite the presence of residual 13 g/L glucose in the medium. Concomitantly, accumulation of acetoin, the precursor of 2,3-BD, increased after maximum 2,3-BD concentration was reached, suggesting that 2,3-BD was reconverted to acetoin after the concentration tolerance threshold for 2,3-BD was exceeded. Cultures of P. polymyxa were then challenged with levo-2,3-BD (20, 40 and 60 g/L) at 0 h in a glucose medium, and a concentration dependent growth inhibition response to levo-2,3-BD was observed. The growth of P. polymyxa was completely inhibited by 60 g/L levo-2,3-BD. Furthermore, P. polymyxa was challenged with incremental 2,3-BD concentrations (20, 40 and 60 g/L at 12, 24 and 36 h, respectively) to mimic 2,3-BD accumulation during fermentation. Interestingly, 2,3-BD was reconverted to acetoin when its concentration reached 60 g/L, possibly to alleviate 2,3-BD toxicity.
Collectively, our findings indicate that 2,3-BD-mediated toxicity is a major metabolic
120 impediment to 2,3-BD overproduction, making it an important metabolic engineering target towards rational design of a 2,3-BD-overproducing strain.
4.2 Introduction
The instability of petroleum price coupled with the finite nature of crude oil has reignited interest in bio-products as renewable replacements for their petroleum-derived counterparts. Among such products, biologically-derived 2,3-butanediol (2,3-BD) is currently receiving tremendous attention because of its multifaceted industrial applications (Ji et al., 2011). For instance, 2,3-BD is a crucial feedstock chemical in the production of 1,3-butadiene (1,3-BD), a monomer of synthetic rubber, currently produced by cracking of petroleum (Celinska and Grajek, 2009). In addition, 2,3-BD is used as an anti-freeze due to its extremely low freezing point of - 60 °C (Soltys et al., 2001).
Further, methyl ethyl ketone an important fuel additive is a derivative of 2,3-BD
(Emerson et al., 1982), which also serves as a feedstock chemical for diacetyl production, an important flavor enhancer in the food industry (Bartowsky and Henschke, 2004).
Different bacterial species including multiple Klebsiella species, Paenibacillus polymyxa,
Bacillus licheniformis, and Bacillus amyloliquefaciens, have been shown to produce 2,3-
BD from sugars (Ji et al., 2011). P. polymyxa was chosen for this study because of its non-pathogenicity. Most 2,3-BD producers, particularly Klebsiella species, are pathogenic, which makes them unattractive for industrial application. In addition, P. polymyxa produces up to 98% levorotary 2,3-BD (levo-2,3-BD), the 2,3-BD isomer better suited to industrial applications due to its chiral nature, which makes it more
121 amenable to a wide range of desirable chemical reactions (Sadhu et al., 1984; De Mas et al., 1988; Nakashimada et al., 2000; Gao et al., 2010; Yu et al., 2011). For example, levo-
2,3-BD is easily dehydrated to form 1,3-butadiene (1,3-BD).
Apparently, yields and titers of 2,3-BD during bacterial fermentation remain low, thereby impeding efforts at commercializing fermentative 2,3-BD production. Co- production of multiple interfering products such as ethanol, formate, lactate, acetate, acetoin, and exopolysaccharides (EPS) account in part for low 2,3-BD yield during fermentation because carbons are diverted away from the 2,3-BD biosynthesis pathway
(De Mas et al., 1988; Häßler et al., 2012; Li et al., 2013; Papoutsakis and Meyer, 1985).
Levan is an EPS produced by P. polymyxa during 2, 3-BD fermentation. Levan is comprised of fructans linked predominantly by β (2-6) and β (2-1) glycosidic bonds; and its production during fermentation turns the medium into a sticky, colloidal solution that makes product recovery difficult (Chaudhary et al., 1996; Miasnikov, 1997).
Additionally, the co-products, formate, lactate, and ethanol drastically affect cell growth by disrupting intracellular pH and denaturing enzymes and membranes when produced in significant amounts during fermentation (Kashket, 1987; Piard and Desmazeaud, 1991;
Warnecke and Gill, 2005; Kwon et al., 2011). These co-products also interfere with downstream processing of 2,3-BD, thereby impacting the cost of 2,3-BD purification and ultimately, overall production cost. Consequently, efforts at improving fermentative 2,3-
BD production have focused considerably on media manipulations/optimization, and strain improvement targeted at reducing the production of competing products
(Nakashimada et al., 2000; Dai et al., 2014). Although some progress has been made with
122 different groups reporting 2,3-BD titers between 15 and 80 g/L in batch cultures and 19.5 to 111 g/L in fed-batch cultures (Nakashimada et al., 2000; Häßler et al., 2012; Li et al.,
2013), commercialization of 2,3-BD fermentation remains to be actualized. Increasing
2,3-BD concentration during fermentation is a critical prerequisite for commercialization.
To this end, we rationalized that in addition to the effects of interfering co-products, other factors may contribute to the inability of 2,3-BD producing microorganisms to accumulate 2,3-BD titers above a certain concentration threshold.
We rationalized that a likely reason for this may be 2,3-BD-mediated toxicity to the fermenting microorganisms. Hence, in this study, we investigated the tolerance of P. polymyxa DSM 365 to increasing 2,3-BD concentrations added at 0 h and pulse-fed at 12,
24 and 36 h of fermentation. Our results suggest that 2,3-BD-mediated toxicity likely poses a roadblock to the accumulation of higher 2,3-BD concentrations during fermentation. Therefore, we infer that understanding the mechanism of action of this roadblock might aid efforts targeted at engineering 2,3-BD-overproducing strains.
4.3 Materials and methods
4.3.1 Microorganism and culture conditions
Paenibacillus polymyxa DSM 365 used in this study was procured from the
German Collection of Microorganisms and Cell Culture, Braunschweig, Germany
(DSMZ- Deutsche Sammlung von Mikroorganismen und Zellkulturen). The lyophilized stock was reactivated by inoculating into Luria Bertani (LB) broth, grown overnight (12 h), and then stored as glycerol stock (50 % sterile glycerol) at – 80 °C. Inocula for
123 fermentation were prepared by inoculating 1 ml of P. polymyxa stock into 30 mL of pre- culture medium containing (g/L); 20.0 glucose, 5.0 yeast extract (YE; Sigma-Aldrich, St louis, MO), and 5.0 tryptone (Sigma-Aldrich, St. Louis, MO), and 3.0 (NH4)2SO4. The pre-culture was supplemented with 0.09 ml of phosphate buffer (pH 6.5) and 0.09 ml trace element solution. The phosphate buffer (pH 6.5) contained (g/L); 3.5 KH2PO4, 2.75
K2HPO4, 0.2 MgSO4, while the trace element solution was prepared by dissolving 0.4 g/L
FeSO4 into 3 mL 25 % HCl, followed by addition of 500 ml double-distilled H2O and then addition of (g/L); 0.8 H3BO3, 0.04 CuSO4.5 H2O, 0.04 NaMoO4.2 H2O, 5.0 MnCl2.4
H2O, 0.1 ZnSO4.7 H2O, 0.08 Co(NO3)2.6 H2O, 1.0 CaCl2. 2 H2O and 0.01 biotin. When the optical density (OD600nm) of the pre-culture reached 1.0-1.2, 10 ml of actively growing cells were transferred into 90 mL of sterile pre-culture medium and incubated aerobically for another 2-3 h until OD600nm reached 1.0-1.2, after which it was transferred to the production media. Glucose, yeast extract and tryptone were prepared and sterilized separately at 121°C for 30 min. Phosphate buffer and trace element solution were prepared separately and filter-sterilized using 0.22 µm PES filter (Corning Incorporated,
Corning, NY). All media components were constituted after sterilization.
4.3.2 Fed-batch fermentations
Fed-batch fermentations were conducted in a 6-L Bioflo 3000 Bioreactor (New
Brunswick Scientific, Edison, NJ). The fermenter was equipped with sensors for measuring pH, agitation speed, and temperature. Mixing was achieved by means of 2
Rushton impellers (3-plate). Sterile air was sparged into the medium through a 0.2 µm
124
PTFE Acro®50 sterile filter (Pall Corporation, Ann Arbor, MI) using a Masterflex L/S®
Pump (Cole-Parmer Instrument Company, Vernon Hills, IL) connected to the top of the bioreactor at a flow rate of 150 mL/min. The production media contained (g/L): 100 glucose, 5.0 YE, 5.0 tryptone, 3.0 (NH4)2SO4, 3.5 KH2PO4, 2.75 K2HPO4, 0.2 MgSO4,
1.5 NH4 acetate, 0.05 CoCl2, 10.0 3-(N-morpholino) propanesulfonic acid (MOPS), and 6 mL of the trace element solution. All medium components were prepared separately and filter-sterilized using a 500 mL polystyrene non-pyrogenic sterile filter bottle (0.2 µm;
Corning Incorporated, NY), with the exception of glucose, YE and tryptone that were separately autoclaved at 121 °C for 15 min. After cooling, the medium components were constituted under aseptic condition. The fed-batch fermentations were initiated with a starting working volume of 2 L (inoculated with 10 % v/v seed culture) at an initial pH of 6.5±0.1 that was externally controlled with 12.5 % NH4OH or 6.5 N H3PO4 when pH dropped below 6.0±0.1 or increased above 6.5±0.1. The fermentation medium was stirred at 300 rpm and the bioreactor was fed when broth glucose concentration dropped below
20 g/L. Each feeding was accompanied by addition of half strength of all the original medium components.
4.3.3 Batch fermentation, and levo- and meso-2,3-BD toxicity bioassay
The tolerance of P. polymyxa to levo-2,3-BD was conducted in batch cultures in sterile 125 ml Pyrex bottles containing 30 ml of the production medium. Since levo-2,3-
BD is by far the predominant isomer produced by P. polymyxa, 2,3-BD toxicity bioassay was first conducted with this isomer. The production medium contained 80 g/L glucose,
125 while the other medium constituents were same as in the medium described above for the fed-batch process. Two separate sets of experiments were conducted to test the tolerance of P. polymyxa to levo-2, 3-BD. First, levo-2,3-BD was added to the medium at 0 h. In this set of experiments, levo-2,3-BD was added to triplicate cultures to final concentrations of 0, 20, 40 and 60 g/L levo-2,3-BD, respectively. The medium with 0 g/L levo-2,3-BD served as control. Second, levo-2,3-BD was pulse-fed into the fermentation medium in a step wise manner aimed at mimicking the pattern of 2,3-BD accumulation by P. polymyxa during fed-batch fermentation. Levo-2,3-BD pulse-feeding was commenced at 12 h, when growth had reached sufficient cell density to withstand possible levo-2,3-BD-mediated toxicity. Levo-2,3-BD was pulse-fed at 12, 24 and 36 h to make up 2,3-BD concentrations in the cultures to 20, 40 and 60 g/L, respectively. Twenty seven milliliters of the production medium was inoculated with 3 ml of pre-culture (10% inoculum; OD600nm of 1.0-1.2). All experiments were started at pH 6.5 and incubated in an Innova™ 4000 rotary shaker (New Brunswick Scientific, Edison, NJ) agitated at 200 rpm, and temperature was maintained at 37 °C. To test whether both levo- and meso-2,3-
BD are equally toxic to P. polymyxa, cultures of P. polymyxa were supplemented in triplicate with 20, 40 and 60 g/L of either levo- or meso-2,3-BD at 0 h and cell growth was measured by monitoring cell density as described below (analytical methods).
4.3.4 Analytical methods
® Cell growth was determined by measuring optical density (OD600) in a DU
Spectrophotometer (Beckman Coulter Inc., Brea, CA). Changes in pH were measured
126 using an Acumen® Basic pH meter (Fischer Scientific, Pittsburgh, PA). The concentrations of 2,3-BD (levo- and meso-), acetoin, ethanol, and acetic acid were determined using a 7890A Agilent gas chromatograph (Agilent Technologies Inc.,
Wilmington, DE, USA) equipped with a flame ionization detector (FID) and a J x W
19091 N-213 capillary column [30 m (length) x 320 µm (internal diameter) x 0.5 µm
(HP-Innowax film thickness)]. The carrier gas was nitrogen, and the inlet and detector temperatures were maintained at 250 and 300 °C, respectively. The oven temperature was programmed to span from 60 to 200 °C with 20 °C min-1 increments, and a 5-min hold at
200 °C. Samples (1 µL) were injected with a split ratio of 10:1.
Glucose concentrations were determined by HPLC using a Waters 2796
Bioseparations Module equipped with an Evaporative Light Scattering Detector (ELSD;
Waters, Milford, MA) and a 9 µm Aminex HPX-87P column; 300 mm (length) x 7.8 mm
(internal diameter) connected in series to a 4.6 mm (internal diameter) x 3 cm (length)
Aminex deashing guard column (Bio-Rad, Hercules, CA). The column temperature was maintained at 65 °C. The mobile phase was HPLC-grade water maintained at a flow rate of 0.6 mL/min. The EPS produced during fermentation was quantified as previously described (Zhang et al., 2002). Culture broth was centrifuged at 8,000 x g for 10 min to pellet the cells while EPS remained in the supernatant. EPS was precipitated with 95% cold ethanol (4 °C), 3 x the volume of the supernatant. The supernatant-ethanol mixture was kept overnight at 4 °C, followed by centrifugation at 8,000 x g for 10 min. The EPS pellet was dried in the oven at 60 °C and weighed afterwards on a Mettler AE 166 weighing balance (Mettler, Toledo, OH).
127
4.4 Statistical analysis and calculations
Analysis of variance (ANOVA) using Tukey’s method for pairwise comparisons between treatments was conducted using Minitab 16 (Minitab Inc., State College, PA). Maximum product concentrations, yields and productivities were analyzed at 95 % confidence interval.
4.5 Results
4.5.1 Production of 2,3-BD by P. polymyxa in fed-batch cultures
To determine the 2,3-BD production capacity of P. polymyxa, fed-batch fermentations (three separate experiments) were conducted. Fresh glucose medium was fed into the bioreactor when concentrations fell below 20 g/L (Fig. 4.1B). The sum of levorotary and mesorotary 2,3-BD is presented as total 2,3-BD. Approximately 47 g/L total 2,3-BD was produced by P. polymyxa in fed-batch fermentations (Fig. 4.1A). The concentration of 2,3-BD increased with time until 72 h when concentration began to decrease, with concomitant increase in acetoin concentration (Fig. 4.1A). Prior to 72 h, acetoin concentrations remained less than 6 g/L and increased to 10.7 g/L afterwards
(Fig. 4.1A). Similar to 2,3-BD, ethanol concentration increased steadily until 72 h when the concentration plateaued. Residual glucose concentration in the range of 10 – 16 g/L remained in the fermentation broth at the end of fermentation (Fig. 4.1B). Extra glucose medium was fed into the bioreactor at 24, 42, and 60 h, thereby increasing glucose concentrations to 53, 45, and 35 g/L, respectively (Fig. 4.1B). The rate of 2,3-BD production reduced markedly after 60 h before a drop in concentration was observed at
128
72 h (Fig. 4.1A). A total of 199 g/L of glucose was consumed by P. polymyxa during fed- batch fermentation. EPS concentration increased with time and plateaued after 72 h fermentation (Fig. 4.1A). However, it appears that a fraction of the produced EPS was utilized during the course of the fermentation given the oscillatory trend of the EPS concentration in the bioreactor (Fig. 4.1A).
4.5.2 Tolerance of P. polymyxa to levo-2,3-BD during 2,3-BD fermentation
To evaluate the tolerance of P. polymyxa to levo-2, 3-BD (the major 2,3-BD isomer produced by this microorganism) during fermentation, different concentrations of levo-2,3-BD (0, 20, 40, and 60 g/L) were either added to the fermentation medium at 0 h or pulse-fed into the culture at 12, 24 and 36 h. The concentrations of choice and time points of levo-2,3-BD addition were informed by the patterns of 2,3-BD accumulation observed in the fed-batch experiment (Fig. 4.1A). Levo-2,3-BD affected the growth of P. polymyxa in a concentration dependent manner. While 20 g/L 2,3-BD added at 0 h had no effect on the final cell density in cultures of P. polymyxa relative to the control (0 g/L
2,3-BD), a 46% reduction in exponential growth rate (at 12 h) was observed with an extended lag phase in cultures supplemented with 20 g/L 2,3-BD (Fig. 4.2A). With 40 and 60 g/L levo-2,3-BD, cell growth reduced 32% and 100% (at 36 h), respectively, relative to the control (Fig. 4.2A). The rate of 2,3-BD production was approximately 1.2- and 1.9-fold faster in the control than in cultures supplemented with 20 and 40 g/L levo-
2,3-BD, respectively, where extended concentration-dependent lags in 2,3-BD production were observed (Fig. 4.2B). Maximum total 2,3-BD concentrations (this refers to 2,3-BD
129 accumulated by the growing cells, excluding supplemented 2,3-BD) accumulated in the cultures treated with 0 and 20 g/L levo-2,3-BD were approximately 23 and 19 g/L. In constrast, the cultures supplemented with 40 g/L levo-2,3-BD produced only 58% of total
2,3-BD of that of the control with no levo-2,3-BD supplementation (Fig. 4.2B). The cultures challenged with 60 g/L levo-2,3-BD at 0 h did not produce 2,3-BD. With the exception of cultures challenged with 60 g/L levo-2,3-BD, which exhibited no growth, hence no metabolic activity, acetoin concentrations showed an irregular pattern, increasing and decreasing at different time points (Fig. 4.2C). Interestingly, initial rise and fall in acetoin concentrations (12 h) occurred only in cultures challenged with 20 g/L levo-2,3-BD (6 g/L), whereas acetoin increased continuously in 40 g/L levo-2,3-BD- challenged cultures (Fig. 4.2C). Acetoin concentrations increased sharply towards the end of the fermentation, beginning at 48 h for the controls and the cultures challenged with 20 g/L levo-2,3-BD, and at 12 h for the 40 g/L levo-2,3-BD-challenged cultures
(Fig. 4.2C). Although residual glucose concentrations in the 0 g/L- and 20 g/L levo-2,3-
BD-challenged cultures were the same, the initial rate of glucose utilization was 3.9-fold faster in the control fermentations (12 h of fermentation; (Fig. 4.2D)). Residual glucose concentration in the cultures supplemented with 40 g/L levo-2,3-BD at 0 h was 9.5- and
7.6-fold higher than those in cultures supplemented with 0 g/L and 20 g/L, respectively.
Despite high glucose consumption in the 20 g/L levo-2,3-BD supplemented cultures, 2,3-
BD yield was considerably low. This is likely as a result of diversion of carbon to the
EPS and ethanol biosynthetic pathways (Table 4.1).
130
In an attempt to mimic the pattern of 2,3-BD production by P. polymyxa, we pulse-fed levo-2,3-BD into the fermentation broth at different time points. Pulse-feeding began at 12 h (late exponential phase) when the cultures had grown significantly (Fig.
4.3A). Pulse-feeding of levo-2,3-BD exerted less effect on cell growth and 2,3-BD production by P. polymyxa than in cultures challenged with levo-2,3-BD at 0 h.
Following the initial pulse-feeding (12 h), cell density of P. polymyxa cultures decreased
(Fig. 4.3A). Whereas 2,3-BD concentration decreased in P. polymyxa cultures challenged with levo-2,3-BD (pulse-feeding from 12 – 36 h and 0 h supplementation), this effect was more pronounced in cultures challenged with levo-2,3-BD at 0 h. For instance, cultures of
P. polymyxa pulse-fed between 12 and 36 h to bring the total concentration of levo-
2,3BD to 20 g/L (12 h), 40 g/L (24 h), and 60 g/L (36 h) exhibited a 19% decrease in total 2,3-BD concentration (relative to the controls, Fig. 4.3B), while cultures challenged with levo-2,3-BD (20 and 40 g/L) at 0 h showed decrease (16 and 42%, respectively) in total 2,3-BD concentration (Fig. 4.2B). Further, cultures challenged with 60 g/L levo-2,3-
BD at 0 h were completely inhibited while the same concentration at 36 h under pulse- feeding only decreased cell growth, glucose utilization, and 2,3-BD production by 16, 5 and 19%, respectively, relative to the controls. Concomitantly, acetoin concentration increased approximately 2-fold in the challenged cultures relative to the controls after maximum total 2,3-BD concentration was attained in the challenged culture through 2,3-
BD supplementation and production (Fig. 4.3C). The glucose profiles in both the challenged and unchallenged cultures were similar pre-challenge. However, upon initial levo-2,3-BD pulse-feeding, the rate of glucose utilization decreased 1.1-fold in the pulse-
131 fed cultures when compared to the controls (Fig. 4.3C). Similarly, residual glucose concentration in the challenged cultures was 1.6-fold higher than those in the control fermentations.
4.5.3 Levo-2,3-BD supplementation alters the ratio of levo- to meso-2,3-BD produced by P. polymyxa.
Addition of levo-2,3-BD to P. polymyxa cultures regardless of whether the addition was at 0 h or by pulsed-feeding, altered the ratio of levo- to meso-2,3-BD produced during fermentation, and this effect appears to be concentration-dependent.
While <20 g/L levo-2,3-BD added at 0 h did not alter production of the levo-2,3-BD isomer, resulting in a ratio of levo-2,3-BD to meso-2,3-BD of 23:0 (levo-2,3-BD, 23.0 g/L; meso-2,3-BD, 0 g/L), the ratios of produced 2,3-BD in the 20 g/L levo-2,3-BD- supplementated culture at 0 h decreased from approximately 25:1 (levo-2,3-BD, 14.5 g/L; meso-2,3-BD, 0.6 g/L) to 11: 1 (levo-2,3-BD, 9.7 g/L; meso-2,3-BD, 0.9 g/L) from 60 to
72 h of fermentation. Further, the ratio of levo- to meso-2,3-BD decreased from approximately 12:1 (levo-2,3-BD, 11.7 g/L; meso-2,3-BD, 0.9 g/L) to 4:1 (levo-2,3-BD,
8.3 g/L; meso-2,3-BD, 2.3 g/L) when the concentration of supplemented levo-2,3-BD at
0 h increased to 40 g/L (Fig. 4.4A). For the pulse-feeding experiments, the control cultures (0 g/L levo-2,3-BD supplementation) produced 24.3 g/L levo-2.3-BD and 0 g/L meso-2,3-BD, resulting in a levo-2,3-BD to meso-2,3-BD ratio of 24:0 (Fig. 4.4B).
However, when cultures were pulse-fed with levo-2,3-BD leading to a final 2,3-BD concentration of 60 g/L, the concentrations of 2,3-BD isomers produced at the end of the
132 fermentation were 15.6 g/L levo-2,3-BD and 2.6 g/L meso-2,3-BD, resulting in levo-2,3-
BD to meso-2,3-BD ratio of approximately 6:1 (Fig. 4.4B).
Increased production of meso-2,3-BD by P. polymyxa following levo-2,3-BD supplementation led us to ask whether the change in the levo-2,3-BD to meso-2,3-BD ratio produced resulted from lower toxicity of the meso-2,3-BD isomer than levo-2,3-BD.
Lower toxicity of the meso isomer would explain increased accumulation of the meso isomer following addition of levo-2,3-BD to fermentation cultures, as a means of alleviating stresses stemming from levo-2,3-mediated feedback inhibition. Thus, we evaluated the effects of different concentrations (20, 40 and 60 g/L) of the two isomers
(meso- and levo-2,3-BD) on the growth of P. polymyxa using cultures with no 2,3-BD supplementation (0 g/L 2,3-BD) as controls. When the fermentation medium was supplemented with 20 g/L meso- and levo-2,3-BD, the meso-2,3-BD treated cultures exhibited the same maximum optical density as the control cultures, while the maximum growth attained by the levo-2,3-BD-treated cultures showed a slight reduction in optical density (2.3%) relative to the control and the meso-2,3-BD treated cultures (Fig. 4.5A).
At 40 g/L, the maximum growth achieved by the levo-2,3-BD treated cultures was 24% lesser than that reached by the meso-2,3-BD treated cultures (Fig. 4.5A). More strikingly, while 60 g/L levo-2,3-BD completely inhibited the growth of P. polymyxa, cultures supplemented with 60 g/L meso-2,3-BD grew after an extended lag phase (Fig.
4.5C).
133
4.6 Discussion
Final product concentrations achieved during fermentation significantly influence the economics of large-scale biotechnological operations. The base chemical, 2,3-BD is a versatile petroleum-derived raw material, and as global fossil fuel reserve drops, biological routes for 2,3-BD production are attracting increasing attention as renewable alternatives to fossil-based feedstock. Increasing 2,3-BD concentration in the fermentation broth will exert a significant impact on the commercialization of biological
2,3-BD production. In this study, we investigated the effect of 2,3-BD-mediated feedback inhibition on product concentrations obtained in cultures of P. polymyxa. As expected, a higher concentration of total 2,3-BD was achieved in fed-batch fermentations in the bioreactor than in batch fermentations in shake flasks. Additional glucose supply, mixing, and pH control in the reactor are some of the factors responsible for increased product accumulation in the bioreactor (fed-batch) than in the flasks (batch). Clearly, increased
2,3-BD concentration during fermentation by P. polymyxa significantly limits product accumulation to below the toxic threshold of 50 g/L. This is evidenced by (i) termination of fermentation when total 2,3-BD concentration reached ~47 g/L in fed-batch cultures, despite glucose repletion in the broth (Fig. 4.1A), (ii) increased accumulation of acetoin in fed-batch cultures when total 2,3-BD concentration reached ~47 g/L (Fig. 4.1A) and in the early stages of batch cultures supplemented with 20 and 40 g/L levo-2,3-BD at 0 h
(Fig 4.2C), and in the later stages of batch cultures pulse-fed with levo-2,3-BD (Fig
4.3B, 4.3C); (iii) a dose-dependent decrease in glucose utilization with levo-2,3-BD
134 supplementation (0, 20, 40, 60 g/L; Figs. 4.2D, 4.3D), and (iv) an altered ratio of levo- to meso-2,3-BD produced by P. polymyxa in levo-2,3-BD-challenged cultures (Fig. 4.4).
To better understand the oscillatory trend of the EPS concentration in the bioreactor during fermentation (Fig. 4.1A), we searched the genome of P. polymyxa for the gene that codes levanase, the enzyme that catalyzes the hydrolysis of levan to fructose residues (Alvaro-Benito et al., 2010). Our bioinformatic analysis showed that P. polymyxa possesses a single copy of the levanase gene (1593 bp) that encodes a polypeptide with 530 amino acid residues. A fraction of EPS produced by P. polymyxa via the activity of levansucrase may have been hydrolyzed to fructose residues by levanase, and thus, may account for the decrease in the EPS concentration in the bioreactor as the fermentation progressed.
In the fed-batch fermentations, 2,3-BD production terminated at 72 h with concomitant increase in acetoin production and accumulation. Notably, P. polymyxa cells were still viable (60 – 100 h) given the upward trend of acetoin accumulation and glucose consumption after 72 h fermentation. Indeed, after glucose (25 g/L) supplementation at
60 h, only minimal increase in 2,3-BD concentration was observed (8%), while acetoin concentration increased 3-fold during the same period (Fig. 4.1). Biosynthesis of 2,3-BD proceeds via acetoin reduction by 2,3-BD dehydrogenase, a dual-functional enzyme that also converts 2,3-BD back to acetoin (Johansen et al., 1975; Magee and Kosaric, 1987;
Blomqvist et al., 1993; Lu et al., 2014). While the P. polymyxa 2,3-BD dehydrogenase can catalyze both forward (acetoin to 2,3-BD direction) and reverse (2,3-BD to acetoin direction) reactions, the ratio of 2,3-BD to acetoin produced during fermentation seems to
135 suggest that the forward reaction is favored. However, it appears that the reverse reaction becomes more favorable when the immediate environment is saturated with 2,3-BD (Fig.
4.6). This may explain why 2,3-BD challenged P. polymyxa cultures produced predominantly acetoin after the maximum tolerable limit for 2,3-BD concentration was attained (Fig. 4.3C).
Conceivably, conversion of 2,3-BD to acetoin is a stress-mitigating mechanism evolved by P. polymyxa to alleviate 2,3-BD toxicity since acetoin is less inhibitory to P. polymyxa than 2,3-BD (bioassay data not shown). Additionally, accumulation of acetoin in the cultures supplemented with levo-2,3-BD at 0 h coincided with a lag in cell growth
(Fig. 4.2A), especially in the cultures challenged with 40 g/L levo-2,3-BD, indicating probable reduction of 2,3-BD toxicity (by conversion to acetoin) to facilitate cell growth.
This interpretation is in line with the acetoin production profile of P. polymyxa cultures pulse-fed with levo-2,3-BD (Fig. 4.3C) or control cultures after relatively high concentrations of 2,3-BD had accumulated in the cultures (Fig. 4.2C). Whereas acetoin accumulation occurred early during fermentation in cultures supplemented with 2,3-BD at 0 h, considerable accumulation of acetoin was not observed until the latter stages of fermentation in cultures pulse-fed with levo-2,3-BD at 12, 24 and 36 h (Fig. 4.2C and
4.3C). With incremental increases in levo-2,3-BD in the pulse-fed fermentations, 2,3-
BD-mediated toxicity markedly increased when total 2,3-BD concentration in the fermentation broth exceeded the toxic threshold (48 g/L), thereby necessitating backward conversion to acetoin possibly to alleviate 2,3-BD toxicity (Fig. 4.3C). In all cases where we observed reduction in 2,3-BD concentration after the toxic threshold had been
136 exceeded, it was accompanied by increase in acetoin concentration. Clearly, backward conversion of 2,3-BD contributes to this trend (increase in acetoin concentration).
However, it is noteworthy that acetoin accumulation after 2,3-BD concentrations exceeded toxic levels may stem in part from acetoin biosynthesis from pyruvate. This is logical given that acetoin is less toxic than 2,3-BD, and so acetoin biosynthesis is less likely to stall due to 2,3-BD toxicity. Reductions in 2,3-BD concentration do not completely account for increases in acetoin concentration, which would most plausibly stem from biosynthesis via pyruvate.
Further, the patterns of glucose consumption in the levo-2,3-BD-challenged cultures support the observation that the greater the concentration of 2,3-BD in the fermentation broth the lower the growth of P. polymyxa and 2,3-BD production (Fig. 4.2 and 4.3). The presence of 2,3-BD (> 20 g/L) in the broth in the early stages of fermentation led to a decreased rate of glucose utilization and consequently reduction in total glucose consumed at the end of fermentation (Fig. 4.2D). Interestingly, while the test and control fermentations (levo-2,3-BD-challenged and unchallenged cultures) exhibited similar glucose utilization profile prior to pulse-feeding (0 – 12 h, Fig. 4.3D), the rate of glucose utilization decreased in the treatment cultures following the first round of levo-2,3-BD pulse-feeding (20 g/L at 12 h), resulting in 3.4 % decrease in total glucose consumed at the end of fermentation relative to the control fermentations.
Interestingly, levo- and meso-2,3-BD isomers exerted different degrees of growth inhibition on P. polymyxa. Our results show that meso-2,3-BD isomer is less inhibitory to
P. polymyxa than the levo-2,3-BD isomer (Fig. 4.5). Although P. polymyxa produces the
137 levo-2,3-BD isomer predominantly during fermentation, the concentration of the meso-
2,3-BD isomer in the fermentation medium increased towards the end of the fermentation when concentrations of levo-2,3-BD isomer had reached toxic levels (Fig. 4.2C and
4.3C). A similar pattern (delayed production of meso-2,3-BD) has been reported previously for P. polymyxa ZJ-9 (Gao et al., 2010). This may be additional stress- mitigating mechanism to reduce levo-2,3-BD mediated toxicity in line with the results obtained in the current study.
The mechanism by which P. polymyxa switches from levo- to meso-2,3-BD biosynthesis upon accumulation of toxic concentrations of levo-2,3-BD is not clear, and this requires further study. However, a likely mechanism for this switch lies in the dual- pronged nature of the 2,3-BD pathway (Fig 4.6C). For instance, butanediol dehydrogenase, the enzyme that catalyzes acetoin reduction to 2,3-BD has been shown to generate the two isomers of 2,3-BD (Zhang et al.,2016) by utilizing different acetoin stereoisomers as substrates (Ui et al., 1986). When acetolactate decarboxylase acts on α- acetolactate, R-acetoin is generated, which is then reduced to levo-2,3-BD by butanediol dehydrogenase. Conversely, S-acetoin generated from diacetyl is reduced to meso-2,3-
BD, also by butanediol dehydrogenase (Fig 4.6C). Since levo-2,3-BD concentrations reduced during fermentation with concomitant increases in acetoin and meso-2,3-BD concentrations (Figs. 4.2 and 4.3), it is likely that at high concentrations of levo-2,3-BD, butanediol dehydrogenase oxidizes levo-2,3-BD back to R-acetoin. Therefore, accumulation of R-acetoin by backward conversion (of levo-2,3-BD) and biosynthesis (of acetoin) via pyruvate may limit the activity of acetolactate decarboxylase, which would
138 lead to spontaneous (non-catalytic) conversion of α-acetolactate to diacetyl (Fig 4.6C).
Spontaneous conversion of α-acetolactate to diacetyl is well documented in the literature
(Haukeli and Lie, 1978). Ultimately, diacetyl is converted to S-acetoin and subsequently to meso-2,3-BD by butanediol dehydrogenase (Fig 4.6C).
4.7 Conclusion
Our results underscore the role of 2,3-BD-mediated feedback inhibition in limiting the accumulation of higher concentrations of 2,3-BD in fermentation cultures of
P. polymyxa. If biological production of 2,3-BD is to reach large-scale commercialization, it is therefore, imperative to tackle this bottleneck through process development and rational design of a 2,3-BD over-producing strain (metabolic engineering). In light of the results presented herein, overcoming 2,3-BD toxicity and abolishing catalysis of 2,3-BD dehydrogenase in the reverse direction (2,3-BD conversion to acetoin) are key metabolic engineering targets worth pursuing. Further, since the levo-2,3-BD is the industrially desirable isomer of 2,3-BD, production of small amounts of meso-2,3-BD may represent considerable losses in large-scale operations.
Therefore, eliminating the meso-2,3-BD biosynthesis route may further improve the economics of 2,3-BD fermentation.
139
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Table 4.1: Yields, productivities, and sugar utilization in cultures of P. polymyxa DSM 365 challenged with levo-2, 3-BD (0,
20, 40, 60 g/L).
2,3-BD Glu. Acetoin Glu. (g/L) (g/L) Treatment Conc. Yield Prod. Used Conc. Yield Prod. Used (g/L) (g/g) (g/L.h) (g/L) (g/g) (g/L.h) Control (0 22.8±0.6a 0.32±0.01a 0.63±0.02a 71.1±3.3a 8.6±0.1a 0.11±0.00a 0.12±0.00a 80.1±2.5a g/L levo- 2,3BD) 20 g/L levo- 19.3±1.4b 0.27±0.02b 0.53±0.04 70.3±0.9a 15.8±0.2b 0.20±0.01b 0.22±0.00b 80.5±1.2a 2,3BD b
40 g/L levo- 13.3±1.0c 0.24±0.02b 0.22±0.02c 54.6±0.7b 16.2±0.5b 0.28±0.01c 0.22±0.01b 57.4±0.4b
144 2,3BD d c c c d c c 60 g/L levo- 0.0± 0.0 0.00±0.00 0.00±0.00 0.0±0.0 0.0±0.0 0.00±0.00 0.00 ±0.00 0.0±0.0 2,3BD d
P-values 0.0 0.00 0.00 0.00 0.00 0.00 0.00 0.00
Continued below table
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Table 4.1 Continued above table.
EPS Glu. Ethanol Glu. (g/L) (g/L) Treatment Conc. Yield Prod. Used Conc. Yield Prod Used (g/L) (g/g) (g/L.h) (g/L) (g/g) (g/L.h) Control (0 g/L 8.2±0.6a 0.11±0.01a 0.17± 0.01a 71.1± 3.3 a 4.5± 0.3a 0.07± 0.00a 0.19±0.01a 61.1±1.8a levo-2,3BD) 20 g/L levo- 8.8±1.4a 0.13±0.02a 0.24± 70.3± 0.9a 4.6± 0.2a 0.07± 0.00a 0.13±0.01b 70.3±0.9b 2,3BD 0.04b 40 g/L levo- 7.9±0.1a 0.49±0.04b 0.33± 54.6± 0.7b 1.7± 0.0b 0.04±0.00b 0.05±0.00c 41.9±1.0c 2,3BD 0.01c 60 g/L levo- 0.0±0.0b 0.00±0.00c 0.00± 0.00d 0.0± 0.0c 0.0± 0.0c 0.00±0.00c 0.00± 0.00d 0.0±0.0 d 145 2,3BD
P-values 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Tukey’s pairwise comparisons between treatments for each parameter (maximum products, yield and productivity) were conducted.
Treatments with different superscripts within a column are significant at p < 0.05. Glu and EPS represent glucose and exopolysaccharides,
respectively.
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Figure 4.1: Fed-batch 2,3-BD fermentation by P. polymyxa. Concentrations of fermentation products (total 2,3-BD, acetoin, ethanol, exopolysaccharide: EPS; A) and glucose utilization profile (B) during fed-batch fermentation by P. polymyxa. Error bars show standard deviations of means of three biological replicates.
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Figure 4.2: Tolerance of P. polymyxa to levo-2,3-BD challenge at 0 hr. A: Optical density; B: Total 2,3-BD concentrations with different treatments; C: Acetoin profile; D:
Glucose profile. Cultures were challenged with 0, 20, 40, and 60 g/L levo-2,3-BD at 0 h.
Error bars show standard deviations of means of three biological replicates.
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Figure 4.3: Tolerance of P. polymyxa to levo-2,3-BD pulse-fed at 12, 24, and 36 hrs. A:
Optical density; B: Total 2,3-BD concentrations with different treatments; C: Acetoin profile; D: Glucose profile. Cultures were challenged with 20, 40, and 60 g/L levo-2,3-
BD at12, 24, and 36 h, respectively. Error bars show standard deviations of means of three biological replicates.
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Figure 4.4: The profiles of levo- and meso-2,3-BD in cultures of P. polymyxa following levo-2,3-BD challenge (0 hr-addition and pulse-feeding). A: 0 hr addition of levo-2,3-
BD; B: Pulse-feeding of levo-2,3-BD. Broken lines represent meso-2,3-BD with different scales (secondary axis). Since meso-2,3-BD was absent in most cultures, the values (0 g/L) are merged with the x-axis, hence, not visible. Error bars show standard deviations of means of three biological replicates.
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150 Figure 4.5: Relative inhibitory effects of levo- and meso-2,3-BD on the growth of P. polymyxa. A: The growth profiles of P.
polymyxa challenged with 20 g/L levo- and mso-2,3-BD, relative to the control (0 g/L 2,3-BD); B: The growth profiles of P.
polymyxa challenged with 40 g/L levo- and mso-2,3-BD, relative to the control; C: The growth profiles of P. polymyxa
challenged with 60 g/L levo- and mso-2,3-BD, relative to the control. Error bars show standard deviations of means of three
biological replicates.
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Figure 4.6. Schematic representation of 2,3-BD-mediated feedback inhibition during fermentation by P. polymyxa. (A) The prominent operating pathways during 2,3-BD biosynthesis from sugars as carbon sources; (B) activity of butanediol dehydrogenase catalyzing the reversible acetoin-2,3-BD interconversions. At low 2,3-BD levels in cultures, the rate of acetoin production and conversion to 2,3-BD is relatively constant
(rate of acetoin production equals rate of its conversion to 2,3-BD Bi), whereas above toxic 2,3-BD threshold, 2,3-BD is re-converted to acetoin by the culture (Bii); (C)
Proposed mechanism for the switch from levo- to meso-2,3-BD formation in P. polymyxa
(modified, based on Ui et al., 1986). Figures represent the following enzymes: α- acetolactate synthase, 1; α-acetolactate decarboxylase, 2; butanediol dehydrogenase, 3; diacetyl reductase, 5. The arrows in green and blue represent levo- and meso-2,3-BD formation pathways. The red arrow shows irreversible reduction of diacetyl to R-acetoin by butanediol dehydrogenase.
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Chapter 5: 2,3-Butanediol production from lignocellulosic biomass: Impact of microbial inhibitors on Paenibacillus polymyxa growth and 2,3-butandiol production
5.1 Abstract
Lignocellulosic biomass (LB) is a cheap, readily available and an attractive substrate for economical large-scale fermentative production of 2,3-BD. However, lignocellulose-derived microbial inhibitory compounds (LDMICs) generated during LB pre-treatment pose a significant roadblock to large-scale use of LB in fermentation.
Current methods for removal of LDMICs prior to fermentation are not economical.
Hence, there is need to identify and improve the genetics of microorganisms with intrinsic capability to tolerate LDMICs during fermentation of LB to 2,3-BD. In this study, we investigated the fermentative conversion of LDMIC-replete wheat straw hydrolysate (WSH) to 2,3-BD using Paenibacillus polymyxa, an established metabolic workhorse for 2,3-BD production. First, we demonstrated that P. polymyxa can simultaneously co-utilize pure representative lignocellulosic sugars of WSH with 3% and
18% increase in 2,3-BD yield and productivity, respectively, relative to the fermentations conducted with glucose control. Batch fermentations were then conducted using 60%,
80%, and 100% WSH, and glucose controls. P. polymyxa exhibited considerable tolerance to LDMICs as evidenced by growth which increased 17%, 27% and 32% when compared to the glucose controls. 2,3-BD production was 32, 31 and 23 g/L in 60%, 80%
152 and 100% WSH, respectively. These 2,3-BD concentrations compared favorably to 32 g/L 2,3-BD obtained in the glucose control medium without LDMICs. Further, P. polymyxa showed tolerance to 0.5, 1 and 2 g/L hydroxymethylfurfural, and furfural, and
0.5 g/L each of coumaric acid, vanillic acid and vanillin when used in fermentation media as sole carbon sources. Collectively, these results underscore the potential of P. polymyxa for large-scale fermentation of LB to 2,3-BD.
5.2 Introduction
The use of fossil fuels for energy and chemicals has generated debate on the impacts of fossil fuel consumption on the environment. More importantly, fossil fuel supplies are finite, and with the growing world population expected to reach 9 billion by
2050, global energy demand is expected to increase from 12 billion ton of oil equivalent
(t.o.e) to 18 billion t.o.e by 2035, with the current rate of energy consumption (Chu and
Majumdar, 2012). The use of renewable energy resources and technologies are expected to provide cleaner and environmental friendly energy that will address imminent energy shortages (Dincer, 2000). Currently, as alternatives to fossil fuels, renewable and sustainable techniques for the production of fuels and chemicals are receiving tremendous research attention. In addition, production of biofuels and chemicals from renewable resources is expected to minimize CO2 emission from fossil fuel use (Naik et al. 2010). However, the production of renewable fuels and chemicals is impeded by high substrate cost. Food crops such as corn, barley, rye, sugarcane, soybean, sunflower and canola are the predominantly used substrates for biofuel and chemical production in the
United States, Brazil and Europe (Ajanovic, 2011). Use of food crops for biofuel and
153 chemical production has led to the food versus fuel debate with proponents of food arguing that the use of food crops for biofuel production has led to a hike in food prices
(Ajanovic, 2011). Therefore, the use of nonfood substrates for fermentative production of biofuels and fine chemicals e.g., 2,3-butanediol is imperative. Nonfood feedstock options for renewable fuel and chemical production include energy crops and sustainably harvested wood and forest residues (Tilman et al. 2009).
2,3-Butanediol (2,3-BD) is an important chiral platform chemical with diverse industrial applications including the manufacture of solvents such as methyl ethyl ketone
(MEK; an organic solvent for lacquers and resins), gamma-butyrolactone (GBL; a flavoring and cleaning solvent), and 1,3-butadiene (1,3-BD), a synthetic rubber monomer
(Xiu and Zeng et al. 2008; Celinska and Grajek, 2009; Kopke et al. 2011). Further, the unique properties of 2,3-BD allow its use as (1) anti-freeze due to the low freezing point
(-60 °C) of the levorotatory isomer,(2) ‘octane booster’ for gasoline owing to its high octane rating, (3) liquid fuel because of its heating value of 27,198 Jg-1 (Flickinger, 1980;
Soltys et al. 2001; Celinska and Grajek, 2009).
Common agro wastes with potential for fermentative production of 2,3-BD include corn stover, wheat, rye, and rice straw biomass, which are presently abundant
(Tilman et al. 2009). However, these lignocellulosic agro wastes must be pretreated to release the sugars embedded in the recalcitrant lignin matrix. Pretreatment methods commonly employed to deconstruct the lignin matrix for easy access of hydrolytic enzymes to cellulose and hemicellulose to release fermentable sugars are accompanied by the generation of lignocellulose-derived microbial inhibitory compounds (LDMICs e.g.
154 furfuraldehydes, vanillin, and syringic acid; Delgenes et al., 1996; Palmqvist and Hahn-
Hagerdal, 2000). Among other inhibitory effects, LDMIC disrupt the intracellular redox balance of fermenting microbes and impair their ability to convert lignocellulosic biomass (LB)-derived sugars to desired fermentation products (Ujor et al., 2015;
Palmqvist and Hahn-Hagerdal, 2000). Removal of LDMIC from LB hydrolysates prior to fermentation is desirable; however, pre-detoxification techniques also lead to loss of fermentable sugars and increase in the overall fermentation cost (Frazer and McCaskey,
1989; Martinez et al., 2000; Agu et al., 2016). Simultaneous fermentation and detoxification is a more desirable approach, and so there is need for microorganisms with intrinsic capability to tolerate LDMICs during LB fermentation to 2,3-BD.
In this study, we investigated the ability of Paenibacillus polymyxa, a major non- pathogenic 2,3-BD producer, to tolerate LDMICs while converting wheat straw hydrolysate (WSH) to 2,3-BD. 2,3-BD fermentations were conducted with 60%, 80%,
100% WSH and sugar-only controls. Cell growth, 2,3-BD, acetoin, ethanol and acetic acid production by P. polymyxa were evaluated. Further, the tolerance of P. polymyxa to
LDMICs when used as sole carbon sources was investigated.
5.3 Materials and Methods
5.31 Microorganism and culture conditions
Paenibacillus polymyxa DSM 365 used in this study was procured from the
German Collection of Microorganisms and Cell Culture, Braunschweig, Germany
(DSMZ- Deutsche Sammlung von Mikroorganismen und Zellkulturen). The lyophilized stock was reactivated by inoculating into Luria Bertani (LB) broth, grown overnight (12
155 h), and then stored as glycerol stock (50 % sterile glycerol) at – 80 °C. Inocula for fermentation were prepared by inoculating 1 ml of P. polymyxa stocks into 30 mL of pre- culture medium containing (g/L); 20.0 glucose, 5.0 yeast extract (YE; Sigma-Aldrich, St louis, MO), and 5.0 tryptone (Sigma-Aldrich, St. Louis, MO), and 3.0 (NH4)2SO4. The pre-culture was supplemented with 0.09 ml of phosphate buffer (pH 6.5) and 0.09 ml trace element solution. The phosphate buffer (pH 6.5) contained (g/L); 3.5 KH2PO4, 2.75
K2HPO4, 0.2 MgSO4, while the trace element solution was prepared by dissolving 0.4 g/L
FeSO4 into 3 mL 25 % HCl, followed by addition of 500 ml double-distilled H2O and then addition of (g/L); 0.8 H3BO3, 0.04 CuSO4.5 H2O, 0.04 NaMoO4.2 H2O, 5.0 MnCl2.4
H2O, 0.1 ZnSO4.7 H2O, 0.08 Co(NO3)2.6 H2O, 1.0 CaCl2. 2 H2O and 0.01 biotin. When the optical density (OD600nm) of the pre-culture reached 1.0-1.2, 10 ml of actively growing cells were transferred into 90 mL of sterile pre-culture medium and incubated aerobically for another 2-3 h until OD600nm reached 1.0-1.2, after which it was transferred to the production media. Glucose, yeast extract and tryptone were prepared and sterilized separately at 121°C for 30 min. Phosphate buffer and trace element solution were prepared separately and filter-sterilized using 0.22 µm PES filter (Corning Incorporated,
Corning, NY). All media components were constituted after sterilization.
5.3.2 Pretreatment of Wheat straw
Wheat straw (WS) biomass used in this study was obtained from the agricultural farm of OARDC, Ohio State University, Wooster, Ohio. WS was pulverized to fine particles (1 mm) using a Thomas-Wiley Mill (Thomas Scientific, Swedesboro, NJ, USA).
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The predominant components of WS per biomass dry weight, include cellulose 35-45%, hemicellulose, 30-35% and lignin, 9-18% (Nigam, 2001; Qureshi, et al. 2007). The pulverized WS biomass weighing 221.96 g with a moisture content of 6% was mixed with 1000 ml of 1% (v/v) sulfuric acid in a glass pan (32 x 20 cm) to obtain 15% (w/v) solids loading. The glass pan and its content was covered with aluminum foil and weighed prior to autoclaving at 127°C for 1 h. After autoclaving, water lost due to autoclaving was replaced by adding distilled water to the pan and the WS slurry obtained after acid pretreatment was allowed to cool to room temperature (25 °C), followed by pH adjustment to 5.0 with ammonium hydroxide. The WS slurry was then stored at -20 ºC until use.
5.3.3 Enzymatic hydrolysis of WS slurry
To the acid-pretreated WS slurry, Cellulase (8 ml), Viscozyme (4 ml), Novozyme
188 (2.5 ml), and 0.8 g xylanase (Sigma-Aldrich, St. Louis, MO) were added and the mixture was incubated in a Gyrotory water bath shaker (New Brunswick Scientific,
Edison, NJ) at 50 °C and 80 rpm for 120 h. After hydrolysis, the wheat straw hydrolysate
(WSH) was centrifuged to remove solid debris, and filtered with Whatman filter (11 µm pore size, 110 mm diameter; Whatman International Ltd, Maid Stone, England) to further remove solid residues. Thereafter, the clear WSH was filter-sterilized by passing through a 0.2 µm sterile filter (250 mL volume; Corning Inc., NY). The filter-sterilized WSH was stored in a pre-sterilized screw-capped Pyrex bottle at -20 °C until further use. The WSH was analyzed for sugars and LDMIC composition using High Performance Liquid
Chromatography (HPLC).
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5.3.4 Pure individual and mixed sugar fermentations
Prior to conducting fermentation with WSH, fermentations were conducted with pure individual and mixed sugars. The experiments included glucose-only (control), xylose-only, arabinose-only, xylose-arabinose mixture (1:1) and glucose-xylose- arabinose mixture (8:11:1). The sugar ratio for glucose, xylose and arabinose mixture was informed by the sugar composition of the WSH. The fermentation was conducted in triplicate and each fermentation contained a final sugar concentration of 100 g/L. In addition to concentrations of sugars, the fermentation medium contained 3.5 g/L
KH2PO4, 2.75 g/L K2HPO4, 0.2 g/L MgSO4, 0.05 g/L CoCl2, 10 g/L 3-(N-morpholino) propanesulfonic acid (MOPS), and 6 ml of trace element solution (described above: microorganism and culture preparation). The pH of the fermentation medium was adjusted to 6.6±0.1 using 5 M NaOH. All medium components were prepared separately and then mixed under aseptic condition. Fermentation media were inoculated with actively growing (12 h) P. polymyxa pre-culture to an initial OD600nm of 0.2 -0.3 and fermentation was conducted in triplicate in an Innova™ 4000 incubator shaker (New
Brunswick Scientific, Edison, NJ) at 35 ºC and 200 rpm. Samples (2 mL) were collected at 0 h and every 12 h until fermentation terminated. Samples were analyzed for cell growth, 2,3-BD, acetoin, ethanol, acetic acid and residual sugars.
5.3.5 Fermentation of wheat straw hydrolysate
The WSH contained 38.5 g/L glucose, 55.9 g/L xylose and 4.5 g/L arabinose totaling 98.9 g/L. Batch fermentations were carried out in a 125 mL screw-capped
Pyrex™ bottles with 30 mL fermentation volume. WSH fermentation media were
158 prepared using different volume ratios of WSH to sterile water. The ratios of WSH to sterile water used included 60:40, 80:20 and 100:0 each of which was supplemented with
5 g/L YE, 3.5 g/L tryptone, 3.0 g/L (NH4)2SO4, 3.5 g/L KH2PO4, 2.75g/L K2HPO4,
0.2g/L MgSO4, 1.5 g/L NH4 acetate, 0.05 g/L CoCl2, 10 g/L 3-(N-morpholino) propanesulfonic acid (MOPS), 3 mL of the trace element solution per liter of WSH and glucose (to a final sugar concentration of 100 g/L). The nutrient components added to each medium was prepared as concentrated stock. Glucose was added to each medium to a total final sugar concentration of 100 g/L. The fermentation media were adjusted to a pH of 6.6 using 5 M NaOH prior to inoculation with actively growing (12 h) P. polymyxa pre-culture cell pellets to an initial OD600nm of 0.2 -0.3. Fermentation was conducted in triplicate in an Innova™ 4000 incubator shaker (New Brunswick Scientific, Edison, NJ) at 35 °C and 200 rpm. Two mL samples were collected at 0 h and every 12 h for 84 h.
Fermentation samples were analyzed for cell growth, 2,3-BD, acetoin, ethanol and acetic acid production and residual sugars.
5.3.6 Growth of P. polymyxa on pure LDMICs as sole carbon sources
P. polymyxa pre-culture was harvested by centrifuging at 3,500 rpm and 4 ºC for
5 min, and then inoculated to an initial OD600nm of 1.0-1.5 in mineral medium. The mineral medium contained 0.5 g/L yeast extract; 3.5 g/L KH2PO4, 2.75 g/L K2HPO4, 0.2 g/L MgSO4 and 3 mL trace element solution per 1000 mL mineral medium. Carbon sources used include furfural (0.5, 1 and 2 g/L), 5-hydroxymethylfurfural (HMF; 0.5, 1 and 2 g/L), 0.5 g/L each of p-coumaric acid, vanillic acid and vanillin. The cultures were incubated at 35 °C and 200 rpm for 178 h.
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5.4 Analytical methods
® Cell growth was determined by measuring optical density (OD600) in a DU
Spectrophotometer (Beckman Coulter Inc., Brea, CA). Changes in pH were measured using an Acumen® Basic pH meter (Fischer Scientific, Pittsburgh, PA). The concentrations of 2,3-BD, acetoin, ethanol, and acetic acid were determined using a
7890A Agilent gas chromatograph (Agilent Technologies Inc., Wilmington, DE, USA) equipped with a flame ionization detector (FID) and a J x W 19091 N-213 capillary column [30 m (length) x 320 µm (internal diameter) x 0.5 µm (HP-Innowax film thickness)]. The carrier gas was nitrogen, and the inlet and detector temperatures were maintained at 250 and 300 °C, respectively. The oven temperature was programmed to span from 60 to 200 °C with 20 °C min-1 increments, and a 5-min hold at 200 °C.
Samples (1 µL) were injected with a split ratio of 10:1.
Sugar concentrations were determined by HPLC using a Waters 2796
Bioseparations Module equipped with an Evaporative Light Scattering Detector (ELSD;
Waters, Milford, MA) and a 9 µm Aminex HPX-87P column; 300 mm (length) x 7.8 mm
(internal diameter) connected in series to a 4.6 mm (internal diameter) x 3 cm (length)
Aminex deashing guard column (Bio-Rad, Hercules, CA). The column temperature was maintained at 65 °C. The mobile phase was HPLC-grade water maintained at a flow rate of 0.6 mL/min.
Changes in the concentration of HMF, furfural, p-coumaric acid, vanillic acid and vanillin were determined by HPLC using a Waters2796 Bioseparations Module equipped with Photodiode Array Detector (PDA; Waters, Milford, MA) and a 3.5-µm Xbridge
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C18, 150 mm x 4.6 mm column (Waters, Milford, MA). Samples were eluted at a flow rate of 0.6 mL/min using a gradient mobile phase of acetic acid [0.3% (v/v) in HPLC- grade water] and HPLC-grade methanol (Agu et al., 2016).
5.5 Statistical methods
The maximum growth, 2,3-BD production, 2,3-BD yield, 2,3-BD productivity, ethanol and acetoin production in 60%, 80% and 100% WSH cultures were compared to the glucose cultures using the General Linear Model of Minitab 17 (Minitab Inc., State
College, PA). One way Analysis of variance (ANOVA) using Tukey’s pairwise comparisons between treatments were conducted at 95 % confidence interval and treatments with a p<0.05 were considered significant.
5.6 Results
5.6.1 Pretreatment and hydrolysis of wheat straw biomass
The generation of LDMICs during LB pretreatment hampers the ability of fermenting organisms to utilize the released sugars in LB. In the present study, we evaluated the ability of P. polymyxa to convert LB to 2,3-BD and compared the results to the glucose control without inhibitors. The time course of sugar release during wheat straw pretreatment and hydrolysis is shown in Table 5.1. The total sugar released during wheat straw hydrolysis was 98.9 g/L in the ratio of 8.5:12.5:1 for glucose, xylose, and arabinose, respectively. The use of dilute sulfuric acid (1% v/v) pretreatment and subsequent enzyme hydrolysis released 0.66 g of total sugars per gram of wheat straw biomass. The LDMICs generated during the pretreatment process. Furfural was the most abundant LDMIC generated during WS pretreatment and accounted for 84% of the total
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LDMICs generated. The high hemicellulose content of WS led to the release of more than 61% pentose sugars from which a large amount of furfural was generated (Tables
5.2; Fig 5.2). Ferulic acid produced during the pretreatment process was 7% of the total
LDMICs, the next most abundant LDMIC to furfural (Table 5.2; Fig 5.2). Ferulic acid is the product of lignin deconstruction and belonged to the phenolic class of LDMICs.
5.6.2 2,3-BD production in pure individual and mixed sugars
Prior to conducting 2,3-BD fermentation using WSH, the ability of P. polymyxa to metabolize representative sugars of lignocellulosic biomass was investigated. The sugars tested included glucose (control), xylose, arabinose, a combination of arabinose and xylose in the ratio of 1:1, and a mixture of glucose, xylose and arabinose in the ratio of 9:12:1, respectively. The ratios of glucose, xylose and arabinose were informed by the composition of sugars derived from WS biomass (Tables 5.1; Fig 5.1). The total sugar concentration in each treatment was approximately 100 g/L. P. polymyxa grew rapidly in the glucose control cultures and in treatment containing glucose, xylose and arabinose mixture (Fig 5.3A). P. polymyxa growth in the pentose sugars was slower, however; maximum growth in the pentose sugars was between OD600nm 6.3-7.3 (Fig 5.3A). In the glucose control culture, P. polymyxa produced 32.3 g/L 2,3-BD with yield and productivity of 0.33 g/g and 0.54 g/L/h, respectively (Table 5.3). Similarly, 2,3-BD was produced in cultures containing individual and mixed sugars in the range of 28.2 -32.2 g/L (Fig 5.3B), whereas 2,3-BD yield and productivity obtained in the individual and mixed sugars were between 0.29 - 0.34 g/g and 0.41- 0.72 g/L/h, respectively (Table
5.3). The highest concentration of 2,3-BD (32.3 g/L) was obtained in the glucose control
162 cultures, whereas the least concentration (28.2 g/L) was observed in arabinose cultures
(Table 5.3). The rate of 2,3-BD production was faster in the glucose-xylose-arabinose medium, resulting in 1.3-fold increase in productivity relative to the glucose control cultures (Table 5.3).
The sugar utilization profiles of individual and mixed sugar fermentations are shown in Fig 5.4. In the control cultures, glucose utilization was rapid, and maximum growth was attained in 24 h (Fig 5.4A). Similarly, sugar utilization in fermentation with glucose, xylose and arabinose as a mixture was rapid, and all three sugars were metabolized simultaneously. The presence of glucose in the fermentation medium did not exert a repressive effect on the utilization of other sugars (carbon catabolite repression).
However, the rate of glucose utilization was 1.7-fold more than that of xylose (Fig 5.4A).
Residual xylose (~ 13 g/L) remained at the end of the fermentation whereas glucose and arabinose were completely utilized at 36 h and 72 h of fermentation, respectively (Fig
5.4A). It is noteworthy, however, that the starting concentration of arabinose in the mixed sugar medium was ~5 g/L. Fermentations with pentose sugars only (xylose or arabinose), were slow with a corresponding extended growth lag phase (Figs 5.4C – 5.4E).
Subsequently, fermentation improved resulting in accumulation of maximum cell biomass at 24 h with 2,3-BD production in the range of 28.2 –30.2 g/L (Figs 5.3A, 5.3B).
In the xylose-arabinose mixture (1:1), xylose and arabinose were utilized almost at the same rate during the initial 24 h of fermentation, however, arabinose was consumed 1.1- fold more than xylose although, xylose (~9 g/L) and arabinose (~6 g/L) remained at the end of fermentation (Fig 4B).
163
5.6.3 P. polymyxa growth and 2,3-BD production in wheat straw hydrolysate
To determine the extent of cell growth that P. polymyxa is capable of achieving in
WSH, fermentations were conducted with 60%, 80% and 100% WSH diluted with sterile distilled water. The growth profile of P. polymyxa in WSH is shown in Fig 5.5A. The growth of P. polymyxa was affected by WSH in a concentration dependent manner. P. polymyxa had an extended lag phase of growth in 80% and 100% WSH relative to the glucose control. The initial exponential growth rate (24 h) of P. polymyxa in 80% and
100% WSH decreased 16 % and 46%, respectively, compared to the glucose control (Fig
5.5A). Nonetheless, the growth of P. polymyxa in 80% and 100% WSH improved during the remaining period of fermentation (Fig 5.5A). Overall, the maximum growth of P. polymyxa increased 17%, 27% and 32% in 60%, 80% and 100% WSH, respectively, relative to glucose (Table 5.4).
2,3-BD was produced in all WSH cultures, however; the 2,3-BD production rate was affected by the concentration of WSH (60, 80 or 100%). The initial 2,3-BD production rate (24 h) in 60%, 80% and 100% WSH decreased 47%, 11%, and 100%, respectively, when compared to glucose (Figs 5.5A and 5.5B) due to the extended lag phase in WSH (Fig 5.5B). The maximum 2,3-BD produced on glucose was 32.0 g/L whereas 32.5, 31.2 and 23.4 g/L 2,3-BD were produced in the 60%, 80% and 100%
WSH, respectively. Similarly, accumulation of acetoin, the precursor of 2,3-BD, increased 2%, 66% and 112% with 60%, 80% and 100% WSH, respectively, when compared to the glucose cultures (Table 5.4). Ethanol production in 60% and 80% WSH increased 33% and 12%, respectively, relative to the glucose cultures (Fig 5.5D). Acetate
164 was completely re-assimilated in the glucose-only medium, whereas acetate re- assimilation decreased by less than 22% with WSH (Fig 5.5E).
The starting glucose, xylose and arabinose ratios in 60%, 80% and 100% WSH were 31: 12.5: 1, 20: 12.5:1 and 13: 12.5: 1, respectively (Table 5.5). The glucose concentration in all the WSH cultures varied owing to the use of glucose to bring the total sugar concentration to 100 g/L following the dilution of WSH to 60% and 80 %.
Consistently, all the cultures grown in WSH contained residual glucose, xylose and arabinose at the end of fermentation. Residual glucose, xylose and arabinose in 60%,
80% and 100% WSH were in the ratios of 4:1.5:1, 3:1.8:1 and 4:3:1, respectively.
However, 2,3-BD yield in 60%, 80% and 100% WSH decreased 0, 3 and 18%, respectively, relative to the glucose cultures (Table 5.4). Further, 2,3-BD productivity decreased 15%, 2% and 47% in the 60%, 80% and 100% WSH, respectively, relative to the glucose cultures (Table 5.4).
5.6.4 Using LDMICs as sole carbon sources
Due to the high cell biomass obtained in fermentations containing WSH, we speculated that P. polymyxa may use LDMICs as carbon sources. To test this premise, P. polymyxa was grown in mineral medium containing LDMICs as sole carbon sources.
Reduction in LDMICs concentrations were observed in P. polymyxa cultures, however, majority of reductions observed for LDMICs did not significantly translate to increase in cell growth. As shown in Fig 5.6A, P. polymyxa cultures containing 0.5 – 2 g/L HMF showed a prolonged lag growth phase before growth was observed in cultures containing
1 and 2 g/L HMF. Growth increased 2.4- and 2.8-fold, respectively, when compared to
165 the control without a carbon source. The period of increased cell growth (48 h – 96 h) in the cultures containing 1 and 2 g/L HMF corresponded to the time points when HMF disappeared completely from the cultures (Fig 5.6D). P. polymyxa growth decreased 1.6-,
1.9- and 3-fold in cultures containing 0.5, 1 and 2 g/L furfural, respectively, relative to the control without this carbon source (Fig 5.6B). However, 98%, 63% and 22% furfural were metabolized in cultures containing 0.5, 1 and 2 g/L furfural, respectively (Fig 5.6E).
Similarly, the growth of P. polymyxa decreased 2.4-, 1.1- and 1.9-fold with coumaric acid, vanillic acid and vanillin (0.5 g/L each), respectively, compared to control (Fig
5.6C). Despite the observed poor growth, P. polymyxa metabolized 94% of coumaric acid whereas 59% and 68% of vanillic acid and vanillin were metabolized, respectively (Fig
5.6F).
5.7 Discussions
The high cost of sugar substrates used in microbial fermentation is a major factor that affects the overall cost of fermentation-derived products including 2,3-BD (Yang et al., 2015). LB is a promising alternative substrate for 2,3-BD production to reduce the overall cost of microbial 2,3-BD fermentation. However, utilization of inexpensive and readily available LB for 2,3-BD production is currently unfeasible owing to microbial inhibition of fermenting microorganisms by LDMICs generated during LB pretreatment.
Chemical detoxification of LB hydrolysates prior to fermentation further complicates 2,3-
BD fermentation and adds to the overall cost of 2,3-BD production. In addition, fermentable sugars are lost during chemical removal of LDMIC (Martinez et al., 2000;
Chandel et al., 2011). The objective of this study was to test the ability of P. polymyxa to
166 convert un-detoxified WSH to 2,3-BD. Our approach included (i) to evaluate the ability of P. polymyxa to convert the representative (individual and mixed) sugars to 2,3-BD, (ii) to test the growth of P. polymyxa and 2,3-BD production in 60%, 80% and 100% WSH, and (iii) assess the ability of P. polymyxa to metabolize LDMICs as sole carbon sources.
Fermentations conducted with pure sugars showed that P. polymyxa can simultaneously metabolized glucose, xylose and arabinose to 2,3-BD (Fig 5.3B and 5.4), although glucose was better utilized, relative to xylose and arabinose (Fig 5.4A). This observation suggests that the uptake of xylose and arabinose by P. polymyxa is not affected by the presence of glucose in the fermentation medium; thus, P. polymyxa does not appear to exhibit carbon catabolite repression. In bacteria such as Klebsiella oxytoca and Enterobacter cloacae, the presence of glucose represses the uptake of other sugars such as xylose and arabinose (Ji et al., 2009; Saha and Bothast, 1999). The ability of P. polymyxa to simultaneously co-utilize mixed sugars suggests that P. polymyxa may be carbon catabolite repression (CCR) negative. CCR-negative microorganisms have tremendous potential for the fermentation of LB hydrolysates (Kim et al., 2010). Further,
P. polymyxa does not show preference between xylose and arabinose. Both pentose sugars were consumed almost at the same rate when supplied concomitantly in the medium, similar to fermentations conducted with xylose or arabinose, as single sugar substrates (Fig 5.4D and 5.4E). Moreover, the final total sugars utilized in the mixed sugar fermentations (glucose-xylose-arabinose and xylose-arabinose mixtures) were fairly similar to the total sugars utilized with individual sugars at the end of fermentation
(Fig 5.4). This suggests that the sugar type or combination of sugars in the fermentation
167 medium does not affect overall sugar utilization by P. polymyxa. In parallel, 2,3-BD was produced with all the individual and mixed pure sugars tested, which highlights the potential of P. polymyxa for industrial-scale LB fermentation to 2,3-BD.
Following the results obtained with pure sugars, we tested sugar utilization by P. polymyxa in WSH. As observed with pure sugars, P. polymyxa utilized glucose, xylose and arabinose simultaneously during fermentation of WSH. Interestingly, total sugars consumed increased with 60% and 80% WSH, in which ~9 g/L residual sugars remained post fermentation, relative to ~13 g/L residual sugars with pure mixed sugars (glucose- xylose-arabinose). This represents 31% increase in total sugar utilization with 60% and
80% WSH; further highlighting improved metabolism with WSH (60% and 80%), when compared to pure sugars. However, with 100% WSH, LDMICs appeared to exert a slight increase in toxicity on P. polymyxa, as evidenced by 46% decrease in sugar utilization, relative to the pure sugar controls (Fig. 5.4A; Table 5.5). It is relevant to mention though that cell growth increased 28% with 100% WSH when compared to the pure sugar controls. The overall growth profiles with 60% and 80% WSH were similar and comparable to the 100% WSH, while showing a 14% increase relative to the pure sugar controls (Table 5.3 and 5.4). Clearly, the level of growth observed with WSH, particularly 100% WSH, suggests that additional carbon sources may have been acquired by P. polymyxa from an additional carbon source, most likely the LDMICs. As discussed below, there are indications that P. polymyxa utilizes some LDMICs as carbon sources, in addition to detoxification (biotransformation to less toxic forms).
168
A likely mechanism for the observed reduction in sugar utilization in the WSH relative the pure sugar controls might be ATP depletion. For instance, Banerjee et al.
(1981) showed that furfural (>1 g/L) inhibits the activities of the glycolytic enzymes, triosephosphate dehydrogenase, hexokinase and phosphofructokinase. Biotransformation of furfural (>2.25 g/L) increased the rate of ATP consumption 3-fold (Horváth et al.,
2003), and glycolytic enzymes such as hexokinase and phosphofructokinase require ATP as a cofactor for activity. Hence, likely depletion of intracellular ATP as a result of
LDMIC detoxification (biotransformation) may account for the observed reduction in sugar consumption by P. polymyxa in WSH, relative to the pure sugar controls. Analysis of ATP levels in cells of P. polymyxa grown in WSH and pure sugars would shed more light on any likely role that ATP depletion due to LDMIC detoxification may play in reducing sugar transport and utilization in cultures grown in WSH.
The sulfuric acid pretreated WS biomass generated furfural, HMF, ferulic acid, coumaric acid, vanillic acid, cinnamaldehyde, syringic acid and hydroxybenzaldehyde as shown in Table 5.2. To evaluate the effects of LDMICs in WSH on P. polymyxa, fermentations were conducted with 60%, 80% and 100% WSH. With all the WSH concentrations tested, P. polymyxa demonstrated superior growth relative to the glucose- only control. In fact, overall growth in WSH resulted in enhanced cell biomass accumulation with WSH (Fig 5.3A). However, an extended lag phase was observed with
80% and 100% WSH earlier in the fermentation. This is indicative of LDMIC-mediated toxicity on the growth of P. polymyxa. This pattern is typical of LDMIC-replete cultures as previously reported by Zhang and Ezeji (2012), where the growth of Clostridium
169 acetobutylicum was impeded until furfural, the LDMIC studied by the authors was detoxified to concentrations below the toxic threshold (2 g/L). Considering the variety of
LDMICs generated when LB is pretreated and detoxified, it is likely that one or more of the inhibitory compounds present in WSH was toxic to P. polymyxa, thus warranting detoxification prior to the onset of cell growth (Fig 5.5A). Increase in the overall growth of P. polymyxa (post early lag phase), however, suggests that P. polymyxa may utilize some of the LDMICs in WSH as carbon sources.
2,3-BD production was affected by WSH in a concentration-dependent manner.
The production of 2,3-BD in the WSH decreased with concomitant increase in acetoin accumulation (Table 5.4, Fig 5.5C). Biosynthesis of 2,3-BD proceeds via the activity of
2,3-BD dehydrogenase, which requires NADH for acetoin reduction to 2,3-BD (Johansen et al., 1975; Magee and Kosaric, 1987; Blomqvist et al., 1993; Okonkwo et al., 2017).
Interestingly, increasing concentration of WSH led to increased accumulation of acetoin
(Fig 5.5C). Whereas biosynthesis of 2,3-BD is NADH-dependent, acetoin biosynthesis is not NADH-dependent. Conversely, LDMIC detoxification consumes high amounts of
NADH (Palmqvist and Hahn-Hagerdal, 2000; Ujor et al., 2015). In light of this, it is plausible that reduction in the amounts of NADH available for 2,3-BD dehydrogenase- catalyzed conversion of acetoin to 2,3-BD (due to NADH consumption for LDMIC detoxification) accounts for the reduced biosynthesis of 2,3-BD and increased acetoin accumulation with increasing concentration of WSH. Notably, the control (sugar-only medium), 80% and 60% WSH produced similar 2,3-BD profiles, while production (of
2,3-BD) reduced 27% when the concentration of WSH was increased to 100%.
170
Therefore, it does appear that P. polymyxa exhibits significant tolerance to LDMICs in
WSH below a toxic threshold – 100% WSH. This is not unusual as Clostridium beijerinckii and C. acetobutylicum have been previously reported to detoxify 2 g/L furfural with modest increase in product (butanol) production (Zhang and Ezeji, 2012;
Ezeji et al., 2007). However, when furfural concentration was increased to 3 and 4 g/L, cell growth and product accumulation reduced, considerably (Zhang and Ezeji, 2012;
Ezeji et al., 2007). Whereas 100% WSH did not exert any observable effect on the growth of P. polymyxa, 2,3-BD production reduced significantly. This further underscores the tolerance of this organism to WSH, albeit with specific effect on 2,3-BD biosynthesis; most plausibly LDMIC-mediated NADH consumption.
As mentioned earlier, the growth profiles observed with WSH relative to those observed with pure sugar controls and the respective amounts of sugars consumed in these cultures indicated that P. polymyxa might utilize some LDMICs as carbon source, in addition to a robust tolerance/detoxification mechanism. To test this premise, fermentations were conducted with pure individual LDMICs as sole carbon sources. Fig
5.6 shows the LDMICs utilization profiles of P. polymyxa. The growth of P. polymyxa in all the LDMICs tested produced a concentration-dependent extended lag in growth (Fig
5.6). However, the growth of P. polymyxa in 1 and 2 g/L HMF increased dramatically at
72 h and 84 h, respectively, with a corresponding decrease in HMF concentration (Fig
5.6A and 5.6D). This supports the premise that P. polymyxa might utilize HMF in WSH for cell biomass accumulation. Further studies are necessary to confirm that P. polymyxa utilizes HMF as a carbon source. While 2 g/L HMF resulted in increased cell optical
171 density for P. polymyxa, 2 g/L furfural completely inhibited growth. This suggests that
HMF may be less toxic to P. polymyxa than furfural (Figs 5.6A and 5.6B). Alternatively,
P. polymyxa may possess a dedicated pathway/mechanism for HMF mineralization, without a corresponding mechanism for furfural consumption. A similar preference for
HMF over furfural has been observed in Cupravidus brasiliensis, which has been demonstrated to possess a unique ability to metabolize furans (Agu et al., 2016;
Koopman et al., 2010). Phenolic compounds have been shown to be more toxic than furans - furfural and HMF – even at lower concentrations than furans (Palmqvist and
Hahn-Hagerdal, 2000; Ezeji et al., 2007). Although lower concentrations of phenolic inhibitors (vanillin, vanillic and coumaric) were tested in this study, they were all found to completely inhibit the growth of P. polymyxa (Fig 5.6). It can be concluded then that
P. polymyxa lacks both a mechanisms for detoxifying phenolic compounds and the requisite pathway for mineralization of phenolic inhibitory compounds.
5.8 Conclusions
Development of an integrated 2,3-BD production process that utilizes microbial strains capable of simultaneous co-utilization of lignocellulose-derived sugars while at the same time tolerating or utilizing LDMICs as carbon sources would be a spring board for economic and sustainable large-scale 2,3-BD production. Our results show that P. polymyxa has the metabolic capability to simultaneously co-metabolize representative sugars of lignocellulosic biomass. To the best of our knowledge, this is the first report of mixed sugar fermentation by P. polymyxa. The growth of P. polymyxa was dramatically enhanced in WSH and 2,3-BD production in 60% and 80% WSH was not significantly
172 affected by the presence of LDMICs. In fact, P. polymyxa showed considerable tolerance to LDMICs and may have the metabolic machinery to utilize some of the inhibitors as sole sources of carbon. Overall, P. polymyxa has potential for industrial-scale 2,3-BD production from cheap lignocellulosic biomass.
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Table 5.1: Saccharification profile during enzymatic hydrolysis of acid-pretreated wheat straw.
Time HPLC method (g/L) (h) Glucose Xylose Arabinose Total 0 - 50.8 3.9 54.5 60 38.3 51.9 4.2 94.4 96 38.4 59.8 4.1 101.6 120 38.5 55.9 4.5 98.9
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Table 5.2: Lignocellulose derived microbial inhibitory compounds (LDMICs) detected in wheat straw hydrolysate.
LDMICs Amount (mg/L) Cinnamaldehyde 48.5 Ferulic acid 269.7 Coumaric acid 0.9 Syringaldehyde 44.8 Vanillin 85.7 Syringic acid 39.6 Hydroxybenzaldehyde 49.5 Vanillic acid 32.6 Furfural 3125.2 HMF 27.7
179
Table 5.3. Comparisons of growth, 2,3-BD concentration, yield, and productivity, and maximum ethanol and acetoin concentrations in individual and mixed sugar cultures.
Treatments Max. Max. 2,3- 2,3-BD 2,3-BD Ethanol Acetoin growth BD (g/L) yield (g/g) prod. (g/L) (g/L) (OD600nm) (g/L/h) Glucose 8.2±0.2 32.3±0.5 0.33±0.01 0.54±0.01 4.7±0.1 7.5±0.9 (control) Glu-Xyl- 8.1±0.3 32.2±0.4 0.34±0.01 0.72±0.08 5.2±0.1 8.4±1.6 Ara Xylose 7.3±0.3 29.4±2.8 0.31±0.03 0.41±0.04 5.8±0.1 8.6±1.1 Arabinose 6.3±0.3 28.2±1.7 0.29±0.02 0.46±0.03 5.5±0.1 8.6±0.3 Xyl-Ara 6.7±0.1 30.2±0.9 0.32±0.02 0.42±0.01 5.4±0.0 8.0±0.8 .
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Table 5.4. Comparisons of growth, 2,3-BD concentration, yield, and productivity, and ethanol and acetoin concentrations in WSH and glucose control.
Treatments Max. Max. 2,3- 2,3-BD 2,3-BD Ethanol Acetoin growth BD (g/L) yield (g/g) prod. (g/L) (g/L) (OD600nm) (g/L/h) Glucose 7.9±0.4a 32.0.±2.3a 0.33±0.03a 0.53±0.45a 4.9±0.5a 4.1±0.7a control 60% WSH 9.2±0.21ab 32.5±0.2a 0.33±0.00a 0.45±0.00a 6.5±0.1b 4.2±0.1a 80% WSH 10.0±0.8b 31.2±3.4ab 0.32±0.05a 0.52±0.06a 5.5±0.8ab 6.8±0.5b 100% WSH 10.4±0.1b 23.4±0.6b 0.27±0.01a 0.28±0.01b 2.5±0.3c 8.7±0.1c P-value 0.006 0.034 0.314 0.006 0.002 0.001 Tukey’s pairwise comparisons were conducted between treatments (maximum growth, maximum products, yield and productivity). Treatments with different superscripts within a column are significant at p < 0.05.
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Table 5.5. Sugar utilization profiles of P. polymyxa in WSH
Sugar concentration Treatments Glucose 60% WSH 80% WSH 100% control WSH Initial Glucose 107.1±2.2 76.4±0.6 62.9±2.8 52.7±2.7 sugar Xylose - 30.8±0.3 39.3±1.7 49.6±2.5 conc. (g/L) Arabinose - 2.5±0.0 3.2±0.1 4.0±0.2 Total 107.1±2.2 109.6±0.9 105.4±4.6 106.3±5.4 Residual Glucose 10.9±0.5 6.0±0.1 4.8±0.1 9.3±0.8 sugar Xylose - 2.1±0.0 2.9±0.0 7.4±0.6 conc. (g/L) Arabinose - 1.4±0.0 1.6±0.0 2.4±0.2 Total 10.9±0.5 9.6±0.1 9.2±0.1 19.1±1.6 Sugar Glucose 96.1±2.7 70.3±0.6 58.2±2.7 43.4±3.5 consumed Xylose - 28.7±0.2 36.5±1.7 42.3±3.2 (g/L) Arabinose - 1.0±0.0 1.6±0.1 1.5±0.4 Total 96.1±2.7 100.0±0.8 96.3±4.5 87.2±7.1
182
200.00 Before saccharification 240.00 After saccharification 180.00 220.00 160.00 200.00 140.00
180.00
xylose - 12.961 xylose - 12.862 120.00 160.00
140.00 100.00 glucose - 11.953
LSU 120.00 LSU 80.00 100.00
60.00 80.00
40.00 60.00
arabinose - 15.430 arabinose 40.00 20.00 - 15.551 arabinose 20.00 0.00 0.00
0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00 0.00 2.00 4.00 6.00 8.00 10.00 12.00 14.00 16.00 18.00 20.00 22.00 Minutes Minutes
Figure 5.1: HPLC chromatogram of WSH sugar analyses before and after enzymatic hydrolysis.
183
2.20
2.00
1.80 furfural - 9.561 furfural 1.60
1.40 8.058
1.20 AU 1.00
0.80
0.60 Ferulic Ferulic acid - 22.600
0.40
24.084 26.716
0.20 21.276
23.761
23.584
4.960
23.168
24.328
3.700
3.037
6.389
Vanillin Vanillin - 17.717
4.216
16.533
Syringic acid - 15.888
Hydroxybenzaldehyde - 15.716 Hydroxybenzaldehyde
Vanillic acid - 15.076
HMF HMF - 7.477
3.528
Syringaldehyde - 18.571 Syringaldehyde
3.303
10.436 12.760
4.597
Cinnamaldehyde - 25.160 Cinnamaldehyde
5.582
13.816
12.474 14.050
12.275
11.827
5.910 8.436 11.542
11.393 13.197
10.773
13.548
8.685
11.088 19.575
Coumaric acid Coumaric - 20.139
28.959 0.036 0.00
0.00 5.00 10.00 15.00 20.00 25.00 30.00 Minutes
Figure 5.2: HPLC chromatogram of WSH with several LDMIC peaks after wheat straw pretreatment and hydrolysis.
184
185
Figure 5.3. Production of 2,3-BD in single and mixed sugar media. A: Optical density; B: 2,3-BD; C: Acetoin; D: Ethanol;
and E: Acetic acid concentrations in mixed and individual sugars. Error bars show standard deviations of means of three
biological replicates.
185
186
Figure 5.4. Sugar utilization profile of P. Polymyxa in single and mixed sugar fermentations. A: Glucose, xylose and arabinose
sugar mixture; B: Xylose and arabinose sugar mixture; C: Glucose; D: Xylose and E: Arabionose. Error bars show standard
deviations of means of three biological replicates.
186
187
Figure 5.5. Fermentation profile of P. polymyxa in WSH. A: Optical density; B: 2,3-BD; C: Acetoin; D: Ethanol and E: Acetic
acid concentrations of different dilutions of WSH. Error bars show standard deviations of means of three biological replicates.
187
188
Figure 5.6. Use of LDMICs as sole carbon sources by P. polymyxa. A: P. polymyxa growth in HMF; B: P. polymyxa growth in
furfural; C: P. polymyxa growth in phenolic compounds; D: HMF utilization; E: Furfural utilization and F: Phenolic
compounds utilization. Error bars show standard deviations of means of three biological replicates.
188
Chapter 6: Molecular inactivation of exopolysaccharide biosynthesis in Paenibacillus polymyxa DSM 365 for enhanced 2,3-butanediol accumulation
6.1 Abstract
Formation of exopolysaccharides (EPS) during 2,3-butanediol (2,3-BD) fermentation by Paenibacillus polymyxa decreases 2,3-BD yield, increases medium viscosity and impacts 2,3-BD downstream processing. Therefore, additional purification steps are required to rid the fermentation broth of EPS prior to 2,3-BD purification, which adds to the production cost. To eliminate EPS production during 2,3-BD fermentation, we explored a metabolic engineering strategy to disable the EPS production pathway of
P. polymyxa, thereby increasing 2,3-BD yield and productivity. The levansucrase gene which encodes levansucrase, the enzyme responsible for EPS biosynthesis in P. polymyxa, was successfully disrupted. The resulting P. polymyxa levansucrase null mutant showed 34% and 54% increases in growth with 6.4- and 2.4-fold decreases in
EPS formation in sucrose and glucose cultures, respectively. The observed decreases in
EPS formation by the levansucrase null mutant may account for the 27% and 4% increase in 2,3-BD yield, and 4% and 128% increases in 2,3-BD productivity when grown on sucrose and glucose media, respectively. Genetic stability of the levansucrase null mutant was further evaluated. Interestingly, the levansucrase null mutant remained genetically stable over fifty generations with no observable decrease in growth, 2,3-BD and EPS
189 formation with or without antibiotic supplementations. Collectively, our results show that
P. polymyxa levansucrase null mutant has potential for improving the economics of large- scale microbial 2,3-BD production.
6.2 Introduction
Considering the finite nature of fossil fuels, recurrent instability in oil price and the environmental concerns associated with oil consumption; there is an urgent need to develop sustainable alternatives to fossil fuels and their derivatives. Over the past few decades, significant attention has been devoted to the development of alternative sources of fuels and chemicals. 2,3-Butanediol (2,3-BD) is an industrial platform chemical that is generated via cracking of petroleum-derived hydrocarbons. 2,3-BD has wide industrial applications. For instance, 2,3-BD can be used as a feedstock chemical in the production of 1,3-butadiene (1,3-BD), the monomer from which synthetic rubber is produced
(Celinska and Grajek, 2009; Ji et al., 2011). 2,3-BD can also be used as a feedstock for producing methyl ethyl ketone (MEK), a fuel additive which has a higher heat of combustion than ethanol and as solvent from which resins and lacquers can be produced
(Celinska and Grajek, 2009; Ji et al., 2011). Additionally, 2,3-BD has massive potential as a feedstock for the synthesis of a host of numerous pharmaceuticals, cosmetics, paints, and food preservatives (Syu, 2001; Garg and Jain, 1995)..
Several microorganisms have been shown to possess the metabolic machinery to convert carbohydrates to 2,3-BD. However, 2,3-BD is produced via mixed acid fermentation pathway where other products such as ethanol, acetoin, lactic, formic and acetic acids in addition to exopolysaccharides (EPS) are co-generated. These co-products
190 compete with 2,3-BD for substrates and pyruvate resulting in decreased 2,3-BD production (Guo et al., 2014; Zeng et al., 1991). Several studies have focused on the manipulation of fermentation medium composition and fermentation conditions as means of reducing the accumulation of competing products during 2,3-BD fermentation
(Okonkwo, et al, 2017; Priya et al., 2016; Häßler et al., 2012; Ji et al., 2009; Biebl et al.,
1998). Although, significant progress has been made, accumulation of competing products remains a significant challenge to large-scale production of 2,3-BD. This stems from the fact that considerable levels of co-products are still accumulated in the fermentation broth during 2,3-BD fermentation. Further, genetic manipulation of 2,3-BD producers has been explored previously to inactivate lactate dehydrogenase, alcohol dehydrogenase and pyruvate-formate lyase genes, key genes that encode enzymes involved in the biosynthesis of lactate, ethanol and formic acids, respectively (Jung et al.,
2012; Jung et al., 2014; Guo et al., 2014; Jantama et al., 2015). Nevertheless, majority of these studies were conducted with pathogenic 2,3-BD producers which are not ideal for industrial-scale biotechnological applications, as they pose significant health hazards to humans. Thus, we focused on genetic manipulation of Paenibacillus polymyxa, a non- pathogenic 2,3-BD producer. P. polymyxa was specifically chosen for this study due to its non-pathogenicity and the ability to synthesize levo-2,3-BD, the much desired 2,3-BD isomer owing to its excellent optical attributes that makes it easily dehydrated to 1,3-BD
(Nakashimada et al., 2000; de Mas et al, 1988). The other 2,3-BD isomers are meso- and dextro-2,3-BD, which are the major fermentation products of the predominantly
191 pathogenic 2,3-BD producers such as Klebsiella spp, Enterobacter aerogenes, and
Serratia marcescens (Ji et al., 2011; Celinska and Grajek, 2009).
During 2,3-BD fermentation, P. polymyxa synthesizes the exopolysaccharide, levan; a fructose polymer with numerous fructose units in β-(2, 6)-linkages (Donot et al.,
2012). Typically, P. polymyxa produces more than 50 g/L EPS during fermentation
(Häßler et al., 2012), and this accounts for about 20% of the total consumed carbon.
Consequently, EPS biosynthesis reduces 2,3-BD titer and yield by diverting carbon away from 2,3-BD biosynthesis. In addition, EPS formation during 2,3-BD fermentation constitutes a major nuisance by clogging of reactor lines which affects proper mixing of fermentation broth, and most importantly, complicates 2,3-BD downstream processing.
Additional purification steps would be required to eliminate EPS prior to 2,3-BD extraction at industrial-scale, which ultimately adds to the overall cost of production.
Collectively, reduction in 2,3-BD yield due to EPS formation and the attendant impact on downstream processing adversely affect the economics of 2,3-BD fermentation.
Consequently, it is imperative to abolish EPS biosynthesis in P. polymyxa with a view to re-directing substrate carbon to 2,3-BD biosynthesis for improved titer and yield.
Levan is the only known and characterized EPS synthesized by P. polymyxa.
Levansucrase plays a key role in levan production in P. polymyxa by serving as a conduit for the transfer of fructosyl residues to a growing levan chain (Liang and Wang, 2015). P. polymyxa produces EPS as a means of attachment to plant roots, the natural habitat of this microorganism (Lal and Tabacchioni, 2009; Haggag, 2007; Lebuhn et al., 1997).
Comparative analysis of nucleotide and protein sequences of P. polymyxa DSM 365
192 levansucrase relative to other strains of P. polymyxa with complete genome sequence information were performed to ascertain the number of copies of levansucrase present in
P. polymyxa DSM 365. Comparisons were conducted due to absence of complete genome information on P. polymyxa DSM 365. The results of this study are shown in Tables 6.1 and 6.2. P. polymyxa DSM 365 possesses a single copy of levansucrase gene with an open reading frame of 1497 bp. To eliminate EPS formation during 2,3-BD fermentation, the P. polymyxa levansucrase gene was targeted for inactivation. Using homologous recombination, we report a pioneer work on the inactivation of levansucrase gene of P. polymyxa. The P. polymyxa levansucrase null mutant developed in this study was evaluated for growth, 2,3-BD production, substrate consumption, 2,3-BD yield and productivity. Further, stability of the levansucrase null mutant was evaluated.
6.3 Materials and methods
6.3.1 Microorganisms and culture conditions
Paenibacillus polymyxa DSM 365 used in this study was procured from the
German Collection of Microorganisms and Cell Culture, Braunschweig, Germany
(DSMZ- Deutsche Sammlung von Mikroorganismen und Zellkulturen). The lyophilized stock was reactivated by inoculating into Luria Bertani (LB) broth, grown overnight (12 h), and then stored as glycerol stock (50 % sterile glycerol) at – 80 °C. The microorganisms, vectors and enzymes used in this study are shown in Table 6.4.
6.3.2 Genomic DNA extraction
The genomic DNA of P. polymyxa DSM 365 was extracted using a previously established standard procedure (Sambrook and Russell, 2001). Briefly, P. polymyxa cells
193 were grown overnight (12 h) in a pre-culture medium (Okonkwo et al., 2017) until cell
OD600nm reached 0.7. The cells were harvested and centrifuged at 10,000 x g and 4 °C for
10 min. The cells were suspended in Tris-HCl-EDTA (TE) buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0) and vortexed until cells were completely re-suspended in the TE buffer. Zirconia/Silica beads (0.1 mm, BioSpec Products, Inc., Bartlesville, OK) were added to the cells to a final concentration of 50% (w/v). The cells in the mixture were lysed using a TissueLyzer LT (Qiagen, Hilden, Germany) at 50 oscillations per second for 2 min. The cell lysate was centrifuged at 10,000 x g for 10 min and the supernatant was transferred to a clean Eppendorf tube. Three hundred and fifty microliters of phenol and chloroform (1:1) were added to the supernatant and vortexed vigorously for 30 s followed by centrifugation at 10,000 x g for 10 min. The genomic DNA (gDNA) contained in the supernatant was carefully collected without disturbing the aqueous and chloroform interphase where the cell debris and proteins are deposited. The phenol/chloroform extraction procedure was repeated several times until no visible protein layer was noticeable between the aqueous and chloroform interphase. Finally, 175
µl of chloroform was added to the supernatant containing the gDNA and was vortexed vigorously for 30 s followed by centrifugation at 10,000 x g for 10 min. The supernatant was collected in a new Eppendorf tube and then placed on ice for 10 min. The gDNA was precipitated with 1 mL of ice-cold absolute ethanol added down the side of the Eppendorf tube containing the gDNA and was centrifuged at 10,000 x g for 10 min. The gDNA was washed with 70% (v/v) ethanol and centrifuged at 10,000 x g for 10 min. The gDNA was air-dried at room temperature after decantation of the ethanol. The air-dried gDNA was
194 re-constituted in 20 µl of nuclease-free water. The quality and concentration of the gDNA was determined by NanoDrop using Epoch BioTek (BioTek Instruments, Inc., Winooski,
VT). The gDNA was stored at -20 °C until use.
6.3.3 PCR amplification to generate levansucrase inactivation constructs
PCR primers for levansucrase gene were designed to amplify the entire levansucrase gene with incorporation of XhoI and BamHI restriction sites at the appropriate locations. The PCR primers for the amplification of the erythromycin gene were designed to incorporate ribosomal binding site, spacer and transcription terminator sequences. The design was such that the PCR primers will amplify short sequences (~210 bp) upstream and downstream levansucrase gene designated as LevFragA and LevFragB, respectively. Primers used to generate the constructs and their characteristics are shown in Table 6.3. First, the entire levansucrase gene was amplified from the genomic DNA of
P. polymyxa DSM 365 using LevFragA_fwd and LevFragB_rev primer pair. Then,
LevFragA and LevFragB gene fragments were amplified using Lev-FragA_fwd and
LevFragA_rev, and LevFragB_fwd and LevFragB_rev, respectively, and the gel-purified levansucrase gene amplicon was used as template. The erythromycin gene was amplified using the primer pair, Erm_fwd and Erm_rev1 and pMutin plasmid (BGSC, Columbus,
OH) as template. The erythromycin gene amplicon from the PCR reaction was gel- purified and used as template for PCR reaction using Erm_fwd and Erm_rev2. The use of
Erm_rev2 primer in the second amplification of erythromycin gene ensures the complete addition of the entire transcription terminator sequence downstream of the erythromycin gene sequence. PCR was used to amplify levansucrase, LevFragA, LevFragB and
195 erythromycin (ERM) genes. The PCR protocol and conditions are shown in Table 6.3.
PCR and gene splicing by overlap extension using PCR or gene SOEing (SOEing-PCR) were carried out in a Bio-Rad iCycler™ Thermal Cycler (Bio-Rad, Hercules, CA) using
PrimeStar® GXL DNA polymerase (Clontech-Takara, Mountain View, CA). A 50 µl reaction contained 5X PrimeStar® GXL buffer (10 µl), dNTPs (0.25 mM), Primers (0.5
µM each), DNA template (~5 ng/ µl) and GXL DNA polymerase (1 µl). PCR was run using the following conditions: (1) initial denaturation, 98 °C for 2 min; (2) 98 °C for 20 s (1 cycle); (2) 98 °C for 30 s, annealing temperature of primers for 30 s, 72 °C for 1 min
(35 cycles); (3) final extension, 72 °C for 10 min; (4) hold, 4 °C for 10 min (1 cycle).
Nested PCR was used for one-step SOEing-PCR reaction with the following conditions;
(1) initial denaturation, 98 °C for 2 min; (2) 98 °C for 30 s, annealing temperature of templates overlap region for 30 s; 72 °C for 30 s (5 cycles); (3) 98 °C for 30 s, annealing temperature of primers, 72 °C for 30 s (30 cycles); (4) final extension, 72 °C for 5 min;
(5) hold, 4 ° C for 10 min.
Next, splicing by overlap PCR extension (SOEing-PCR) reactions were used to link LevFragA and ERM genes to generate LevFragA-ERM construct. LevFragA and
ERM genes served as templates in the second PCR cycle of the nested PCR to generate
LevFragA-ERM construct that was further amplified using specific primers
(LevFragA_fwd and Erm_rev2) for LevFragA-ERM in the third and fourth nested PCR cycles. The PCR product of the previous one-step SOEing-PCR reaction, LevFragA-
ERM, and LevFragB fragment, were used as templates to generate LevFragA-ERM-
LevFragB construct in another one-step SOEing-PCR reaction.
196
6.3.4 Construction of recombinant plasmid
The levansucrase inactivation construct (LevFragA-ERM-LevFragB) was ligated into pGEM®7Zf(+), a high copy number plasmid in E. coli JM109 which behaves as a non-replicative vector in P. polymyxa. pGEM®7Zf(+) possesses filamentous phage f1 origin of replication recognized by E.coli but not by P. polymyxa and hence, this vector is used to produce circular single stranded DNA (ssDNA) that enhances homologous recombination (Yanisch-Perron et al., 1985). The presence of phage f1 origin of replication and the ability of pGEM®7Zf(+) to be replicated into stable circular DNA in
E. coli is important for its application in the inactivation of genes via homologous recombination in P. polymyxa and other gram positive bacteria.
6.3.5 Restriction digestion
pGEM®7Zf(+) and LevFragA-ERM-LevFragB were restricted independently with XhoI and BamHI (New England biolabs, Ipswich, MA) in a 50 µl reaction. The reaction mixture consisted of 5 µl CutSmart buffer (New England biolabs, Ipswich, MA),
1 µl XhoI, 0.02 µg/ µl DNA and the reaction volume was made up to 49 µl with nuclease- free water (Amresco®, Solon, OH). The mixture was incubated at 37 °C for 1 h, and 1 µl
BamHI was added and incubated for additional 1 h at 37 °C. The restricted plasmid and
LevFragA-ERM-LevFragB construct were purified by agarose gel electrophoresis using
GenCatch and advanced PCR extraction kit (Epoch Life Science, Sugar Land, TX).
6.3.6 DNA ligation
The purified restriction products, LevFragA-ERM-LevFragB and pGEM®7Zf(+) were ligated in a 20 µl reaction to generate the recombinant pGEM®7Zf(+) carrying the
197 levansucrase inactivation construct, LevFragA-ERM-LevFragB. The ligation reaction mixture consisted of 2 µl T4 DNA ligase buffer (New England biolabs, Ipswich, MA), 1
µl T4 DNA ligase (New England biolabs, Ipswich, MA), plasmid (pGEM®7Zf(+)) and
LevFragA-ERM-LevFragB insert in a ratio of 1:5 with final DNA concentration between
0.02-0.1 pmol. The reaction volume was made up with nuclease-free water (Amresco®,
Solon, OH). The reaction mixture was incubated overnight at 16 °C, heat inactivated at
65 °C for 10 min, then chilled on ice for 20 min prior to transformation of competent E. coli JM 109 with recombinant pGEM®7Zf(+) carrying the levansucrase inactivation construct, LevFragA-ERM-LevFragB (See section 6.3.8).
6.3.7 Preparation of E. coli JM 109 competent cells
E. coli cells were grown overnight in sterile Luria-Bertani (LB) medium at 37 °C and 250 rpm. Twenty milliliters of the culture was inoculated into 180 ml of sterile LB medium and incubated at 37 °C and 250 rpm until cell optical density (OD600nm) reached
0.6. The culture was transferred aseptically into four pre-chilled 50 ml sterile centrifuge tubes and placed on ice for 30 min. The culture was centrifuged at 3,500 rpm and 4 °C for
10 min. The cell pellet was washed with 0.1 M MgCl2 solution and centrifuged at 3,500 rpm and 4 °C for 7 min. The resulting cell pellet was washed with 0.1 M CaCl2 solution and centrifuged at 3,500 rpm and 4 °C for 7 min. The supernatant was discarded and the cells were re-suspended in 700 µl of 0.1 M CaCl2 solution. The competent E. coli JM 109 cells were suspended in 30% glycerol and stored at -80 °C until use.
198
6.3.8 Transformation of competent E. coli JM 109
The ligated pGEM®7Zf(+) and LevFragA-ERM-LevFragB (recombinant pGEM®7Zf(+)) was purified using GenCatch advanced PCR extraction kit (Epoch Life
Science, Sugar Land, TX). The purified product was used to transform competent E. coli
JM 109 cells. The purified recombinant pGEM®7Zf(+) (50 ng) was added to 50 µl of competent E. coli JM109 cells previously placed on ice for 20 min. The transformation mixture was mixed by gently tapping the base of the Eppendorf tube containing E. coli
JM109 cells and the recombinant plasmid. The mixture was placed on ice for 20 min and then heat-shocked at 42 °C for 1 min. The cells were chilled on ice for 5 min followed by addition of 450 ml pre-warmed (37 °C) super optimal broth with catabolite repression
(SOC) medium (2% tryptone, 0.5% yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM
MgCl2, 10 mM MgSO4 and 20 mM glucose). Glucose addition to SOC medium increases transformation efficiency of E. coli via repression of genes involved in catabolism of other carbon sources thus, improving E. coli growth (Gorke and Stulke, 2008). The cells were incubated at 37 °C and 250 rpm for 1 h after which the cells were plated on LB agar supplemented with 50 µg/ml ampicillin, and 5-bromo-4-chloro-3-indolyl-β-D- galactopyranoside (X-gal) and isopropyl-β-D-1-thiogalactopyranoside (IPTG) to a final concentration of 20 mg/ml and 1 mM, respectively. The plates were incubated at 37 °C for 12 h after which white colonies were picked and screened for the presence of correct insert. Colony PCR and gel electrophoresis were used to screen for colonies with correct insert. Colonies with the correct insert were grown in LB medium supplemented with 50
µg/ml ampicillin and the recombinant plasmid was isolated and purified using GenCatch
199 plus plasmid DNA miniprep kit (Epoch Life Science, Sugar Land, TX). Restriction digestion was further used to confirm the orientation of the insert. The recombinant plasmid was isolated in large quantity (390 µg), suspended in 500 µl of nuclease-free water (Amresco®, Solon, OH) and stored at -20 °C prior to use.
6.3.9 Preparation and electroporation of competent P. polymyxa protoplasts
Following initial unsuccessful attempts to transform competent P. polymyxa cells with the recombinant pGEM®7Zf(+) harboring the levansucrase inactivation construct via electroporation, competent P. polymyxa protoplasts were used instead. The cell wall of P. polymyxa was removed by a previously described method (Inukai et al., 1993) with slight modifications. Briefly, P. polymyxa cells were grown in tryptic soy broth (TSB) for
12 h until cell optical density (OD600nm) reached 0.7. The cells were harvested and placed in 50 ml centrifuge tubes pre-chilled on ice for 20 min and then washed twice with 50 mM Tris-Maleate buffer (pH 7.1) containing 2 mM dithiothreitol followed by centrifugation at 3,500 rpm and 4 °C for 7 min. The cell pellets were harvested and re- suspended in Tris-Maleate buffer (pH 7.1) containing 0.6 M sucrose, 5 mM MgCl2 and
300 µg/ml lysozyme (Amresco®, Solon, OH). The cell suspension was incubated in an
ISOTEMP 220 water bath (Fischer Scientific, Pittsburg, PA) for 60 min at 37 °C to make
P. polymyxa protoplasts. P. polymyxa protoplasts were harvested by centrifugation at
3,500 rpm and 4 °C for 7 min. P. polymyxa protoplasts were made competent by washing the protoplasts twice with 10% polyethylene glycol (PEG-8000) and re-suspended in 500
µl of 10% PEG-8000. The competent P. polymyxa protoplasts were transformed with recombinant pGEM®7Zf(+) harboring the levansucrase inactivation construct via
200 electroporation. Twenty microliters (100 µg DNA) of the recombinant plasmid was gently mixed with 100 µl of competent protoplasts in a pre-chilled 0.2 cm electroporation cuvette and was placed on ice for 5 min. Electroporation was performed at 2.5 kV, 25 µF capacitance and infinite resistance as previously described (Zhou and Johnson, 1993) in a
Bio-Rad Gene Pulser Xcell™ electroporator (Bio-Rad, Hercules, CA). Electric pulse was delivered to the protoplasts between 2.5 and 4.1 milliseconds. Following electroporation, the protoplasts were placed on ice for 5 min and then 500 µl of TSB was added and the mixture was incubated at 35 °C for 6 h to allow the protoplasts to recover. The recovered cells were plated on tryptic soy agar (TSA) supplemented with 35 µg/ml erythromycin and incubated at 35 °C for 16- 24 h. Colonies were picked and mixed with 50 µl of TSB.
They were re-plated on a fresh TSA plate supplemented with 35 µg/ml erythromycin and incubated at 35 °C for 12 h. Then, fresh colonies were picked and colony PCR technique was used to screen for the presence of erythromycin gene. The colonies with erythromycin gene were transferred to TSB supplemented with 50 µg/ml erythromycin.
The cells were harvested and genomic DNA was extracted as previously described. PCR was performed to screen for the presence of LevFragA-ERM, ERM and ERM-LevFragB fragments using genomic DNA as template. The presence of all three fragments confirms that the levansucrase gene was successfully inactivated via double-cross homologous recombination. A portion of the P. polymyxa cultures with inactivated levansucrase gene were stored in 50% glycerol at -80 °C.
201
6.3.10 Characterization of P. polymyxa levansucrase null mutant
The P. polymyxa levansucrase null mutant was characterized for cell growth, EPS and 2,3-BD production. Batch fermentations were conducted in sucrose- and glucose- based media. One milliliter of 50% glycerol stock of P. polymyxa levansucrase null mutant was inoculated into 30 ml of pre-culture medium supplemented with 35 µg/ml erythromycin and incubated at 35 °C and 200 rpm for 6 h until cell optical density
(OD600nm) reached 1.0-1.2. The actively growing P. polymyxa levansucrase null mutant
(10%, v/v) was inoculated into the fermentation medium containing 100 g/L sucrose or glucose supplemented with 35 µg/ml erythromycin. The pre-culture and fermentation medium components used in this study have been reported previously (Okonkwo et al.,
2017). The fermentation medium was further supplemented with 0-0.4 g/L CaCl2.
Wildtype P. polymyxa was prepared as described for the levansucrase null mutant without erythromycin supplementation. Batch 2,3-BD fermentations were conducted in loosely-capped 125 ml Pyrex culture bottles with 30 ml fermentation volume. All experiments were carried out in triplicate and 2 ml samples were collected at 0 h and then, every 12 h until the fermentation terminated. Samples were analyzed for cell growth, culture pH, EPS, 2,3-BD, acetoin, acetic acid and ethanol production.
6.3.11 Analytical methods
® Cell growth was determined by measuring optical density (OD600) in a DU
Spectrophotometer (Beckman Coulter Inc., Brea, CA). Changes in pH were measured using an Acumen® Basic pH meter (Fischer Scientific, Pittsburgh, PA). Concentrations of fermentation products, 2,3-BD , acetoin, ethanol, and acetic acid were quantified using a
202
7890A Agilent gas chromatograph (Agilent Technologies Inc., Wilmington, DE, USA) equipped with a flame ionization detector (FID) and a J x W 19091 N-213 capillary column [30 m (length) x 320 µm (internal diameter) x 0.5 µm (HP-Innowax film thickness)] as previously described (Okonkwo et al., 2017).
Sugar concentrations, sucrose, glucose and fructose were quantified by High
Performance Liquid Chromatography (HPLC) using a Waters 2796 Bioseparations
Module equipped with an Evaporative Light Scattering Detector (ELSD; Waters,
Milford, MA) and a 9 µm Aminex HPX-87P column; 300 mm (length) x 7.8 mm
(internal diameter) connected in series to a 4.6 mm (internal diameter) x 3 cm (length)
Aminex deashing guard column (Bio-Rad, Hercules, CA). The column temperature was maintained at 65 °C. The mobile phase was HPLC-grade water (Waters Corporation,
Milford, MA) maintained at a flow rate of 0.6 mL/min as described previously (Okonkwo et al., 2016).
The EPS produced during fermentation was quantified using methods modified from Zhang et al. (2002). Culture broth was centrifuged at 8,000 x g for 10 min to pellet the cells while EPS remained in the supernatant. The EPS in the supernatant was precipitated with 95 % ethanol (4 °C), 10 x the volume of the supernatant. The supernatant-ethanol mixture was kept overnight at 4 °C followed by centrifugation at
8,000 x g for 10 min. The EPS pellet was dried in the oven at 60 °C and reconstituted in distilled water. The EPS containing solution was vortexed vigorously to ensure complete dissolution of the EPS. The EPS was then quantified by Phenol-sulfuric acid method
(Nielsen, 2010; Dublois et al., 1956). Briefly, 25 µl of 80% phenol was added into test
203 tubes A containing 1 mL glucose standards (0-0.1 g/L) and test tubes B containing 1 mL diluted EPS samples. The mixture was vortexed briefly and 2.5 ml concentrated sulfuric acid (Fischer Scientific, Pittsburg, PA) was added to the mixture. The mixture was left to stand for 10 min. The text tubes containing the mixture were incubated at 25 °C for 10 min. After incubation, the mixture was gently vortexed and absorbance was measured at
490 nm against reagent blank prepared as the samples. A standard curve was generated by plotting the values of glucose concentration (X-axis) against absorbance (OD490nm) (Y- axis) and EPS concentration were interpolated from the standard curve.
6.3.12 Levansucrase assay
Levansucrase activity in the levansucrase null mutant and wildtype P. polymyxa was measured using culture supernatant, using a method modified from Euzenat et al.
(1997). P. polymyxa samples (levansucrase null mutant and wildtype) were collected at the exponential growth phase when maximum EPS is produced. The sample was centrifuged for 20 min at 8,600 x g and 4 °C. The supernatant from each sample was divided into two. One portion of the supernatant was used to quantify EPS as described above and the EPS obtained was designated as [EPS]B. The other portion was used to assay levansucrase activity and the total EPS produced after levansucrase activity assay was designated as [EPS]A. The reaction mixture for the levansucrase activity assay consisted of 400 µl of 1 M sucrose in 50 mM phosphate buffer (pH 6.0) and 100 µl of culture supernatant. The mixture was incubated at 35 °C for 1 h. Following levansucrase activity, EPS was precipitated with 90% ethanol (4 °C) and the EPS was subsequently
204 quantified and expressed as [EPS]A. The EPS produced during levansucrase activity was determined from the equation below:
[EPS]L = [EPS]A - [EPS]B
Where [EPS]L represents the concentration of EPS synthesized during levansucrase assay.
The concentrations of proteins in the supernatants were determined by Bradford method (Bradford, 1976). One unit of levansucrase activity was defined as the milligram of protein that catalyzed the formation of one micromole of EPS (levan) per min at 35 °C in 1 M sucrose solution.
6.3.13 Growth rate and generation time of levansucrase null mutant
To determine stability of the levansucrase null mutant, growth curve was first obtained. Cells were grown to exponential phase (10 h) in the pre-culture medium
(Okonkwo et al., 2017), then harvested and washed twice with sterile distilled water by centrifugation at 5,000 x g for 3 min. The cell pellet was constituted into several dilutions and their optical densities (OD600nm) were measured against sterile distilled water as blank. Each cell dilution was centrifuged in a pre-weighed Eppendorf tube at 10,000 x g for 10 min and the supernatant discarded. The Eppendorf tubes with the cell pellets were dried to a constant weight in TempCon™ Oven (American Scientific Products, McGraw
Park, IL) at 50 °C. The Eppendorf tubes with the cells were weighed and the weight of the cells at each OD600nm reading was determined. A standard curve was generated from a plot of cell biomass (mg/L) against absorbance at OD600nm. The standard curve was used to convert optical density measurements at OD600nm to cell biomass. The growth curve
205 was obtained by growing levansucrase null mutant in pre-culture medium and the cell biomass was measured at several time points until the cells reached death phase of growth. Then, the cell biomass was plotted against time. The generation (doubling) time of levansucrase mutant was determined from the exponential phase of the growth curve.
6.3.14 Stability of P. polymyxa levansucrase null mutant
The stability of levansucrase null mutant was determined in the presence and absence of 35 µg/ml erythromycin for 50 generations. For stability test under antibiotic pressure, levansucrase null mutant cells were grown in pre-culture medium supplemented with 35 µg/ml erythromycin until OD600nm reached 1.0-1.2. The actively growing cells
(10%, v/v) were transferred into fermentation medium containing 100 g/L sucrose supplemented with 35 µg/ml erythromycin and this generation was regarded as G0
(generation zero). Several subcultures (every 3 h or 2 generations) were made from G0 until G50 (generation 50) was attained, and in each case, cultures were supplemented with
35 µg/ml erythromycin. It should be noted that the generation time of P. polymyxa levansucrase null mutant is 1.5 h. Then, fermentations were conducted using G0, G10, G20,
G30, G40, and G50 under antibiotic pressure. Samples (2 ml) were drawn at 0 h and every
12 h until the fermentation ended and then, analyzed for cell growth, EPS and 2,3-BD production. The same experiment was conducted without antibiotic supplementation.
Generations G0, G10, G20, G30, G40, and G50 were obtained as described above and then used to conduct fermentations. The antibiotic resistance of all generations (with and without erythromycin supplementation) was determined by PCR and replica plating with erythromycin. For each generation, samples of P. polymyxa levansucrase null mutant
206 were drawn during the stationary growth phase (12-16 h of growth) and sub-cultured into fresh pre-culture medium and grown until another stationary growth phase (12-16 h of growth) was attained. The cells were diluted to a concentration of 108 cfu/ml and were plated on TSA plates without antibiotic (erythromycin) supplementation. The plates were incubated at 35 °C for 12 h. Colonies from each generation were picked and screened for the presence of erythromycin gene using PCR. Recombinant pGEM®7Zf(+) harboring the levansucrase inactivation construct was used as erythromycin gene control. Gel electrophoresis was performed using the colony PCR products. The presence of the erythromycin gene in the colonies confirmed the stability of P. polymyxa levansucrase null mutant. Colonies from the TSA plates (without erythromycin supplementation) were transferred by replica plating to fresh TSA plates supplemented with 35 µg/ml erythromycin. The antibiotic supplemented plates were incubated at 35 °C for 12 h and the numbers of colonies compared to the plates without antibiotic supplementation. The schematic representation of step-by-step procedure employed to evaluate the stability of the P. polymyxa levansucrase null mutant is shown in Fig 6.3.
6.3.15 Statistical analysis and calculations
General Linear Model of Minitab 17 (Minitab Inc., State College, PA) was used for all statistical analyses. Analysis of variance (ANOVA) using Tukey’s method for pairwise comparisons was employed to compare differences between treatments.
Differences in growth, sugar utilization, maximum product concentrations, 2,3-BD yields and productivities were compared at 95 % confidence interval. 2,3-BD yield was expressed as the gram of 2,3-BD produced from one gram of substrate (sucrose or
207 glucose). 2,3-BD productivity was expressed as the gram per liter of 2,3-BD produced per hour of fermentation.
6.4 Results
6.4.1 Inactivation of levansucrase gene in P. polymyxa DSM 365
The levansucrase gene of P. polymyxa was successfully inactivated by homologous recombination. Erythromycin gene was inserted between a 210 bp upstream fragment and a 213 bp downstream fragment of levansucrase gene creating a 1224 bp levansucrase inactivation construct (Fig 6.1). A stop codon was placed downstream of
LevFragA sequence followed with a ribosomal binding site and spacer sequence upstream of erythromycin gene. In addition, a transcription terminator sequence was added downstream of the erythromycin gene before the LevFragB sequence. The strategy and design used to generate the levansucrase inactivation construct is shown in Fig 6.1.
Inclusion of stop codon, ribosomal binding site and spacer sequence, and transcription terminator sequences was to ensure that only erythromycin gene is transcribed into mRNA without creating additional metabolic burden on P. polymyxa. The PCR-amplified levansucrase inactivation construct corresponding to 1224 bp (Fig 6.1) was digested and ligated to a pre-digested pGEM®7Zf(+), and was used to transform competent P. polymyxa protoplasts. Considering that pGEM®7Zf(+) exists as a single-stranded circular
DNA in P. polymyxa, the homologous regions of the levansucrase inactivation construct of the recombinant plasmid enabled successful exchange of DNA between P. polymyxa genome and recombinant pGEM®7Zf(+) via double-cross homologous recombination, thus conferring P. polymyxa resistance against erythromycin. Subsequently, erythromycin
208 was used to select recombinant P. polymyxa with disrupted levansucrase gene. PCR- screening of the levansucrase null mutant using LevFragA_fwd/Erm_rev2,
Erm_fwd/Erm_rev2 and Erm_fwd/LevFragB_rev primer pairs show that the mutant possesses LevFragA-ERM, ERM, and ERM-LevFragB genes corresponding to 1008,
800, and 1015 bp, respectively (Fig 6.2), thus confirming successful double-cross homologous recombination.
6.4.2 Effect of levansucrase disruption on EPS formation
Batch 2,3-BD fermentations were conducted on sucrose and glucose substrates to evaluate EPS formation by the levansucrase null mutant. The fermentation cultures were supplemented with 0, 0.2, 0.4 g/L CaCl2. Fermentation on sucrose showed that EPS formation by the mutant decreased 5.8-, 6.4- and 6.1-fold in the 0, 0.2 and 0.4 g/L CaCl2 treatments, respectively, when compared to the wildtype (Table 6.5, Fig. 6.2B). The levansucrase null mutant showed no measurable levansucrase activity whereas more than
0.6 units of levansucrase activity per milligram protein were detected in the wildtype
(Figs 6.4F, 6.5F, 6.6F). The absence of any measurable levansucrase activity in the mutant confirms successful inactivation of levansucrase gene in P. polymyxa. However, despite the fact that no measurable levansucrase activity was detected in the mutant, 2-3 g/L EPS was synthesized by the mutant in sucrose cultures (Table 6.5), thus suggesting that P. polymyxa produces other EPS forms other than levan.
EPS formation by the levansucrase null mutant in glucose cultures decreased 2.4-,
1.7- and 1.9-fold in the 0, 0.2 and 0.4 g/L CaCl2 treatments, respectively, when compared to the wildtype (Table 6.6). Interestingly, no levansucrase activity was observed in both
209 the wildtype and levansucrase null mutant grown on glucose. However, EPS produced by the wildtype cultures in the glucose medium decreased at least 4-fold when compared to the cultures grown on sucrose (Tables 6.5, 6.6). This observation suggests that sucrose is an important activator of levansucrase expression in P. polymyxa. Further, it is likely that the EPS synthesized by P. polymyxa during growth on glucose medium might not be levan given that levansucrase activity was not detected in cultures grown on glucose medium.
6.4.3 Effect of calcium supplementation on growth, sugar utilization, 2,3-BD yield and productivity
Initial fermentations with the levansucrase null mutant resulted in a sharp drop in pH, which adversely affected cell growth and product formation, particularly when the pH fell below 5.5 (Figs 6.4A, B and 6.7A, B;). Thus, CaCO3 and CaCl2 supplementation were adopted because of the ability of calcium to influence key cellular processes such as sugar transport, product formation and tolerance, and to mitigate drop in pH (Han et al.,
2013). While CaCO3 has been previously used to enhance cell growth, product formation and culture pH in a strict anaerobic Gram positive solvent-producing bacterium,
Clostridium beijerinckii NCIMB 8052 (Han et al., 2011), it has been used to increase cell growth and solvent production in facultative Gram negative bacteria, Zymomonas mobilis, and yeast, Scheffersomyces stipitis (Okonkwo et al., 2016; Zeng et al., 2010).
Supplementation of the fermentation broth with CaCO3 and CaCl2 to levansucrase null mutant did not improve culture pH (Figs B.1 and B.2). However, CaCl2 exerted a remarkable influence on growth and 2,3-BD production. Following CaCl2 addition, cell
210 biomass production increased in both the sucrose and glucose cultures for both the wildtype and the levansucrase null mutant with considerable increases in substrate consumption (Tables 6.5 and 6.6). Growth of the levansucrase null mutant on sucrose increased by 22% and 34%, respectively, in 0.2 and 0.4 g/L CaCl2-supplemented cultures when compared to the wildtype (Figs 6.5A and 6.6A) grown under same conditions. As shown in Figs 6.5A and 6.6A, CaCl2 supplementation stimulated production of higher biomass and perhaps stabilized the functions of growth-associated proteins, an effect that has been previously reported for calcium (Han et al., 2011; Han et al., 2012; Okonkwo et al., 2016), and this may have prolonged the stationary growth phase for both the levansucrase null mutant and the wildtype. Although CaCl2 did not affect pH of the levansucrase mutant cultures, the 2,3-BD yield on sucrose increased 27% and 21% with
0.2 and 0.4 g/L CaCl2 treatments, respectively, relative to the wildtype (Table 6.5). In addition, the 2,3-BD productivity of the mutant on sucrose increased by approximately
3% and 4% with 0.2 and 0.4 g/L CaCl2 treatments, respectively, when compared to the wildtype (Table 6.5). However, without CaCl2 supplementation the productivity of the levansucrase null mutant on sucrose decreased by 8.8% compared to the wildtype (Table
6.5). CaCl2 supplementations exceeding 0.4 g/L increased cell growth, but did not improve 2,3-BD yield and productivity on sucrose for both the wildtype and the levansucrase null mutant (data not shown). The stimulatory effects of CaCl2 on growth,
2,3-BD yield and productivity diminished at concentrations greater than 0.4 g/L.
In glucose cultures, addition of 0.2 and 0.4 g/L CaCl2 to levansucrase null mutant cultures increased growth by 27% and 34%, respectively, compared to the mutant grown
211 in cultures without CaCl2 addition (Table 6.6). Similarly, growth of the wildtype increased by 25% and 17% with 0.2 and 0.4 g/L CaCl2 supplementation (Table 6.6).
When compared to the wildtype grown on glucose alone, growth of the levansucrase null mutant with 0.2 and 0.4 g/L CaCl2 treatments increased by 38% and 54%, respectively
(Table 6.6). As observed in the sucrose cultures, glucose utilization by wildtype and levansucrase null mutant strains improved with the supplementation of the fermentation medium with 0.2 and 0.4 g/L CaCl2. Glucose utilization by the mutant increased by 20% and 22% with 0.2 and 0.4 g/L CaCl2 treatments, respectively, when compared to the cultures without CaCl2 addition (Table 6.6). Further, 0.2 and 0.4 g/L CaCl2 supplementation increased glucose consumption by 12% and 11%, respectively, for the wild type when compared to wildtype cultures without CaCl2 treatment (Table 6.6).
Glucose utilization in the levansucrase null mutant cultures increased by 9% and 13% with 0.2 and 0.4 g/L CaCl2 treatments, respectively, when compared to the wildtype grown under similar conditions (Table 6.6). The 2,3-BD yield and productivity of the levansucrase null mutant grown on glucose increased from 0.34 g/g and 0.57 g/L/h
(without CaCl2), respectively, to 0.37 g/g and 1.62 g/L/h (with 0.2 g/L CaCl2) and 0.38
g/g and 1.64 g/L/h (with 0.4 g/L CaCl2), respectively, whereas, 2,3-BD yield and
productivity of the wildtype increased from 0.36 g/g and 0.51 g/L/h (without CaCl2),
respectively, to 0.36 g/g and 0.71 g/L/h (with 0.2 g/L CaCl2) and 0.37 g/g and 0.72 g/L/h
(0.4 g/L CaCl2), respectively. However, the 2,3-BD yield and productivity of the levansucrase null mutant in glucose cultures increased by 3% and 4% (without CaCl2), and 128% and 127.7% in the 0.2 and 0.4 g/L CaCl2 cultures, respectively, relative to the
212 wildtype (Table 6.6). Efficient glucose utilization rate of the levansucrase null mutant may be responsible for the enhanced 2,3-BD titer, yield and productivity relative to the wildtype.
The levansucrase null mutant efficiently converted sucrose to 2,3-BD with diminished ability to produce EPS (Fig. 6.2B). However, the mutant utilized glucose much faster than sucrose resulting in higher 2,3-BD productivity in the glucose cultures relative to fermentations conducted in sucrose medium (Tables 6.5 and 6.6). The mutant achieved a maximum 2,3-BD yield of 0.42 g/g in the 0.2 g/L CaCl2 sucrose cultures which is 27% and 13.5% greater than that (0.33 g/g_sucrose and 0.37 g/g_glucose) achieved by the wildtype grown in sucrose- and glucose-based media (Tables 6.5 and
6.6). The mutant also produced 70% more ethanol on glucose relative to the cultures grown on sucrose, which accounts for the observed decrease in 2,3-BD yield in glucose medium (Tables 6.5 and 6.6).
6.4.4 Stability of levansucrase null mutant
The use of homologous recombination in levansucrase gene inactivation resulted in a P. polymyxa levansucrase null mutant that was found to be stable after growing for
50 generations with and without antibiotic supplementations. Fermentations were conducted with generations 0, 10, 20, 30, 40 and 50 in sucrose medium with and without erythromycin addition. As shown in Figs 6.10 and 6.11, maximum growth of levansucrase null mutant generations grown under antibiotic pressure were in the range of
11.0 to 13.9 OD600nm, whereas without antibiotic, growth ranged from 12.5 to 15.1.
Maximum growth of levansucrase null mutant without antibiotic increased at least 8%
213 relative to the growth observed in the mutant with 35 µg/ml erythromycin addition
(Table 6.7). The observed decrease in the mutant growth when erythromycin was added may be attributed to the presence of erythromycin which triggered expression of erythromycin gene that confer resistance to the antibiotic prior to growth thus, accounts for the reduced growth when compared to fermentation conducted without erythromycin addition.
EPS production in all the tested generations (G0-G50) of the levansucrase null mutant with or without erythromycin was relatively unchanged. Maximum EPS produced by levansucrase null mutant under antibiotic pressure was in the range of 3.1 to 4.1 g/L, whereas, without erythromycin supplementation, the EPS ranged from 2.7 to 3.6 g/L. In addition, no measurable levansucrase activity was obtained in the mutant grown with or without erythromycin addition (Table 6.7). The 2,3-BD production capacities of levansucrase null mutant grown over different generations (G0-G50) with or without antibiotic pressure were indistinguishable. The maximum 2,3-BD produced by the mutant grown under antibiotic pressure was in the range of 35.3 to 39.4 g/L, whereas without antibiotics, the 2,3-BD produced by the mutant was in the range of 36.1 to 39.0 g/L
(Table 6.7). Colony-PCR and replica plating techniques were further employed to characterize each levansucrase null mutant generation (G0-G50) for antibiotic resistances.
The results are shown in Figs 6.12 and 6.13. Notably, the levansucrase null mutant developed in this study retained antibiotic resistance to erythromycin after 50 generations of growth with/without erythromycin addition.
214
6.5 Discussion
EPS production during 2,3-BD fermentation constitutes a nuisance during fermentation and diverts substrate carbon away from the 2,3-BD pathway, thus decreasing 2,3-BD yield and productivity. Also, viscosity of the fermentation broth increases with EPS production, which impairs mixing during fermentation (Häßler et al.,
2012). More importantly, EPS negatively impacts 2,3-BD downstream processing thereby increasing the overall cost of production. Therefore, the aim of this study was to develop a mutant strain of P. polymyxa with diminished ability to synthesize EPS. We employed double cross homologous recombination to inactivate levansucrase gene in P. polymyxa. The following objectives were achieved: (i) disruption of levansucrase gene in
P. polymyxa by inserting erythromycin gene between upstream and downstream fragments of the levansucrase gene, and subsequently use erythromycin as a selection marker for the levansucrase null mutant, and (ii) phenotypic characterization of the levansucrase null mutant by determining its growth, EPS, 2,3-BD, acetoin, ethanol and acetic acid production parameters.
The genome of P. polymyxa DSM 365 has not been completely sequenced. Thus, due to the lack of sufficient genomic information on this microorganism, the nucleotide and protein sequences of the P. polymyxa DSM 365 levansucrase from the available shot- gun sequences were compared to other P. polymyxa strains whose complete genome sequences are available. As shown in Tables 6.1 and 6.2, the completely sequenced P. polymyxa strains have a single copy of levansucrase gene and shared 92-96% and 95-
97% similarities to the protein and nucleotide sequences of P. polymyxa DSM 365,
215 respectively. Levansucrase, which is the only characterized enzyme in P. polymyxa responsible for EPS formation (Rütering et al., 2016; Choi et al., 2004), was targeted for inactivation in this study. The fallouts from the present study are grouped under different attributes.
6.5.1 Effect of levansucrase inactivation on the growth of P. polymyxa levansucrase null mutant
The P. polymyxa levansucrase null mutant was characterized for growth by measuring its optical density (OD600nm) at different time points during 2,3-BD fermentation. Our results clearly suggest that levansucrase is not significantly involved in the growth of P. polymyxa, as inactivation did not result in observable reduction in growth, relative to the wildtype (Tables 6.5 and 6.6). In the natural environment, P. polymyxa has been reported to use EPS for attachment to plant roots (Bezzate et al.,
2001). The results obtained in this study indicate that even though attachment may influence survival in natural environment, EPS biosynthesis does not appear to have a direct impact on cellular growth. For the purpose of potential industrial fermentative application, knock out of levansucrase gene removes a major biochemical expense in terms of carbon diversion to EPS formation, which frees up more substrate carbon for
2,3-BD biosynthesis by the P. polymyxa levansucrase null mutant.
Typically, 2,3-BD is produced via a mixed fermentation acid pathway, which results in the accumulation of acetic, formic and lactic acids during fermentation.
However, acetic acid is re-assimilated during fermentation with concomitant increase in culture pH. Inactivation of P. polymyxa levansucrase gene resulted in acetic acid
216 accumulation, leading to a decrease in culture pH (Figs 6.4B and 6.7B). The acetic acid profile of the levansucrase null mutant relative to the wildtype suggests that levansucrase may be involved in acetic acid re-assimilation. In both the glucose and sucrose-based cultures, with and without CaCl2 supplementation, acetic acid accumulation was observed for the null mutant (Figs. 6.4E, 6.5E, 6.6E, 6.7E, 6.8E, 6.9E). There are no previous reports on any link between EPS biosynthesis and acetic acid assimilation in P. polymyxa or other 2,3-BD producers, hence, this finding warrants further examination. It is likely that this might be a secondary or cascade effect stemming from downstream effectors of levansucrase not directly involved in EPS biosynthesis. Solvent-producing, biphasic Gram positive bacteria typically produce acids and then, reabsorb them during solvent formation. Disruption of their native biology has been reported to engender acid accumulation, due to poor acid assimilation. A similar pattern has been previously reported for Clostridium beijerinckii NCIMB 8052 following knockdown of acetoacetate decarboxylase (Han et al., 2011) and Clostridium acetobutylicum ATCC 842 following inactivation of the carbon catabolite control protein, CcpA (Ren et al., 2010). Perhaps, a similar phenomenon exists in P. polymyxa, also a solvent-producing biphasic Gram positive bacterium. We expect that future molecular analyses might shed more light on this. It is interesting to note though that despite acetic acid accumulation and the attendant drop in culture pH, the levansucrase null mutant exhibited an overall better growth than the wildtype in all conditions tested. This observation perhaps points to relieve of biochemical burden that limits growth when the EPS biosynthesis pathway is fully operational. Alternatively, levansucrase inactivation might confer some form of
217 stress resistance to the mutant cells which mitigates acid-mediated stress. Overall,
CaCl2 supplementation enhanced growth for the mutant and the wildtype in both glucose- and sucrose-based media. This is attributable to previously reported global effect of calcium on cellular metabolism, sugar utilization and stress mitigation (Han et al., 2013).
6.5.2 Effect of levansucrase inactivation on EPS biosynthesis by P. polymyxa levansucrase null mutant
Successful knockout of levansucrase gene in P. polymyxa was confirmed by PCR, restriction digestion, levansucrase activity assay, antibiotic selection and genetic stability.
Knockout of levansucrase gene in P. polymyxa resulted in significant reduction in EPS formation by the levansucrase null mutant in both sucrose- and glucose-based media
(Figs. 6.4C, 6.5C, 6.6C, 6.7C, 6.8C, 6.9C). Clearly, reduction in the level of EPS accumulated by the levansucrase null mutant confirms that levansucrase is a key player in
EPS biosynthesis in P. polymyxa and that the targeted open reading frame (ORF) in the P. polymyxa DSM 365 shotgun sequence encodes a levansucrase. Reduction in EPS production by the levansucrase null mutant was more pronounced with sucrose cultures relative to the glucose-grown cultures. This is ascribable to the fact that EPS formation is more strongly favored by sucrose, which is hydrolyzed by levansucrase to release glucose and fructose (Yanase et al., 1992). The vast majority of the fructose molecules are then linked to form EPS by the same enzyme (levansucrase). Therefore, sucrose consumption by P. polymyxa results in significantly higher EPS production, which was almost completely abolished in the null mutant. In fact, levansucrase activity was not detected in the levansucrase null mutant grown in both glucose- and sucrose-based media, while the
218 wildtype exhibited levansucrase activity during growth in sucrose-based medium but not with glucose medium. The activity profile of levansucrase in the wildtype lends further support to earlier reports on the role of levansucrase in sucrose metabolism in P. polymyxa (Bezzate et al., 2001; Velazquez-Hernandez et al., 2009).
Notably, EPS was detected in both wildtype and the levansucrase null mutant cultures grown on glucose, albeit to greatly reduced concentrations. It is plausible that P. polymyxa produces different polysaccharides with different sugars, with sucrose favoring levan biosynthesis, while glucose supports the production of other uncharacterized polysaccharides. This is not unusual among EPS-producing microorganisms as Bacillus spp., Zymomonas mobilis, Leuconostoc mesenteriodes, Agrobacterium radiobacter,
Xanthamonas campestris, and Pseudomonas aeruginosa have been shown to produce alginate, xanthan, curdlan or dextran with different sugars (Koepsell et al., 1953; Esser et al., 2012; Colvin et al., 2012; Cooley et al., 2016; Papagianni et al., 2001; Saudagar and
Singhal, 2004; Zhang and Chen, 2010). Thus, it would be instructive to characterize the physicochemical properties and molecular signatures of the EPS obtained in glucose- and sucrose-grown cultures of P. polymyxa. However, it is worthy of note that levan is the predominant EPS synthesized by P. polymyxa – in the event that different polysaccharides are produced by this microorganism with different sugars - given the levels of EPS isolated from the cultures of the wildtype and levansucrase null mutant grown on glucose and sucrose, with the latter resulting in greater EPS levels for both strains, particularly the wildtype. Alternatively, a poorly expressed mechanism that is less dependent on levansucrase activity might account for the production of levan when P.
219 polymyxa is grown on glucose, and might explain far less reduction in EPS production in the levansucrase null mutant relative to the wildtype when both strains were grown on glucose medium.
Whereas EPS production was significantly reduced in the levansucrase mutant when compared to the wildtype, this did not translate to a significant increase in 2,3-BD production with sucrose as substrate. While sucrose utilization by the levansucrase null mutant was 1.3-fold lower than that of the wildtype, the amount of 2,3-BD produced was similar to that produced by the wildtype (Table 6.5). This implies that for considerably less substrate (sucrose), the levansucrase null mutant produced the same amount of product. This is an attractive trait from an economic standpoint for potential large-scale production. Further, reduced sucrose utilization by the levansucrase null mutant underscores disruption of sucrose utilization or processing following levansucrase inactivation in the null mutant. Comparatively, the levansucrase null mutant utilized 1.1- fold more glucose than the wildtype, which lends further weight to the role of levansucrase in sucrose utilization in P. polymyxa and successful inactivation of the encoding gene. Increased glucose utilization by the null mutant accounts for 1.1-fold and
1.7-fold increases in 2,3-BD and ethanol production, when compared to the wildtype.
Increased product accumulation (2,3-BD and ethanol) by the levansucrase mutant relative to the wildtype may stem from redirection of free carbon from EPS biosynthesis to the
2,3-BD and ethanol biosynthesis pathways. However, EPS accumulation was only slightly reduced in the levansucrase null mutant grown in glucose-based medium relative to the wildtype, so carbon redirection does not fully account for the increases in product
220 formation observed. Therefore, it is likely that a different mechanism might be at play.
Perhaps, levansucrase inactivation relieved a growth limiting machinery in the levansucrase null mutant leading the observed increase in growth and consequently, increased product formation. Furthermore, ethanol production was clearly enhanced in the levansucrase null mutant relative to the wildtype when grown on glucose medium; an effect that was not observed with sucrose. It is not clear why this pattern occurred, thus warranting further study. However, this result highlights ethanol biosynthesis as a veritable candidate for future inactivation towards developing a 2,3-BD over-producing strain.
6.5.3 Effect of CaCl2 supplementation on sugar utilization, 2,3-BD yield and productivity by P. polymyxa levansucrase null mutant
Initial experiments showed low culture pH stemming from the accumulation of acetic acid in cultures of the levansucrase null mutant (Fig. 6.4E). Despite likely pH- related stresses due to acetic acid accumulation, the levansucrase null mutant showed
39% higher optical density than the wildtype (Fig. 6.4A). In attempts to mitigate pH- related stresses and increase growth and product formation, CaCl2 was added to cultures of the levansucrase null mutant and wildtype P. polymyxa. Previously, Ca2+ has been shown to relieve pH-related stresses (by enhancing acid reassimilation), enhance sugar utilization (by triggering increased expression of sugar transporters), and upregulate expression of heat shock genes/proteins, which contribute to amelioration of solvent- and acid-mediated stresses in butanol-producing solventogenic clostridia (Han et al., 2013;
Han et al., 2011). In this study, CaCl2 produced similar effects on P. polymyxa, 221 particularly the levansucrase null mutant with regards to growth, 2,3-BD production, sugar utilization, and tolerance to pH stresses (Figs. 6.4 – 6.9). Without CaCl2, sucrose utilization was 17% lower by the levansucrase null mutant than the wildtype (Table 6.5).
Conversely, the levansucrase mutant used 3% more glucose than the wildtype, without
CaCl2 (Table 6.6). While this underscores disruption of sucrose metabolism in the levansucrase null mutant, the results obtained with CaCl2 supplementation underline the
2+ role of Ca in modulating stresses in solvent-producing microorganisms. With CaCl2
(0.4 g/L), sucrose utilization increased in cultures of the levansucrase null mutant of P. polymyxa by 27% (Table 6.5), compared to cultures of the levansucrase null mutant grown in the absence of CaCl2. Similar effects were also observed with the wildtype, where a residual sucrose concentration of 2.5 g/L was detected after fermentation without
CaCl2, whereas cultures supplemented with 0.2 and 0.4 g/L CaCl2 had 0 g/L sucrose post- fermentation. CaCl2 had minimal effect on glucose utilization by wildtype P. polymyxa.
With or without CaCl2, the levansucrase null mutant exhibited enhanced glucose utilization, when compared to the wildtype. The molecular profile of both the levansucrase null mutant and the wildtype may shed more light on whether or not proteins involved in sugar transport and catabolism were upregulated in the presence of
CaCl2.
Notably, acetic acid accumulation in cultures of the levansucrase null mutant did not appear to impede growth for both glucose- and sucrose-based media. When CaCl2 was added to cultures of the levansucrase null mutant, cell optical density increased 32% on average for both glucose- and sucrose-based cultures, relative to the cultures not
222 supplemented with CaCl2 (Figs. 6.4 – 6.9). Similarly, CaCl2 led to an average 33% increase in cell optical density for the wildtype, relative to CaCl2-unsupplemented cultures. Concomitantly, acetic acid concentrations increased in cultures of the levansucrase null mutant following CaCl2 supplementation (Figs 6.4 – 6.9). Hence, increased cell growth particularly for the levansucrase mutant upon CaCl2 supplementation was not as a result of improved acetic acid reassimilation. We speculate that Ca2+-mediated stress mitigating effects are responsible for the growth profile of the levansucrase null mutant despite acetic acid repletion in the culture. High acid concentrations trigger membrane, DNA and protein damaging effects in bacteria (Roe et al., 2002), which in the absence of a mitigating factor impede growth and product formation. Typically, P. polymyxa cultures require extensive buffering during fermentation to obviate acid-mediated premature termination of fermentation. On the other hand, Ca2+ has been shown to upregulate heat shock proteins (involved in the repair of damaged or aberrant proteins) and DnaK involved in DNA synthesis, transcription and repair in C. beijerinckii NCIMB 8052 (Han et al., 2013). Additionally, Ca2+ has been implicated in the stabilization of bacterial membrane, which reduces the effects of membrane-damaging factors such as acids (Hansen et al., 2001; Kotra et al., 1999).
Therefore, the effects observed with CaCl2 for both strains, albeit more pronounced in the levansucrase null mutant of P. polymyxa, in which acetic acid accumulation was evident, likely stemmed from Ca2+-mediated mitigation of pH stresses. Further, in addition to mitigating pH stresses, CaCl2 directly promotes cell growth through the activities of heat shock proteins and DnaK (Han et al., 2013), which may in part, account for high optical
223 densities observed with CaCl2. For instance, at 24 h of fermentation, the optical density of the levansucrase null mutant was at least 28% higher with CaCl2 supplementation, when compared to non-CaCl2 supplemented cultures.
With regards to product formation, CaCl2 supplementation led to marginal increase in 2,3-BD production on sucrose medium– both for the levansucrase null mutant and the wildtype. However, on glucose medium, CaCl2 supplementation resulted in 32% increase in 2,3-BD accumulation with concomitant 32% increase in ethanol production, when compared to cultures not supplemented with CaCl2. In addition, CaCl2 supplementation increased 2,3-BD and ethanol production by 14% and 74%, respectively, in the levansucrase mutant relative to the wildtype. It is plausible that enhanced cell accumulation due to increased growth in the CaCl2-supplementated cultures ultimately led to increased product formation vis-à-vis 2,3-BD and ethanol.
Collectively, Ca2+-mediated pleiotropic effects on cell growth and relief of pH stress likely account for the fermentation profiles observed with CaCl2.
6.5.4 Stability of P. polymyxa levansucrase null mutant
The stability of microbial strains intended for industrial bioprocesses is critical for uniform and consistent product generation. This is particularly important when genetically modified strains are used. Hence, stability of the levansucrase null mutant generated in this study was tested. Stability results clearly showed that this mutant is stable as demonstrated by similar fermentation profiles (growth, 2,3-BD concentration, acid and ethanol concentrations, and EPS production) for the levansucrase null mutant
224 grown to different generation times (up to 50 generations) in the presence and absence of antibiotic (Figs 6.10 and 6.11).
6.6 Conclusions
The levansucrase gene was successfully inactivated in P. polymyxa via homologous recombination, creating a stable and faster growing strain with significantly reduced EPS production, improving 2,3-BD fermentation and product recovery. The ability of the levansucrase null mutant to grow and produce higher concentrations of 2,3-
BD on glucose makes it attractive as a basis for generating a 2,3-BD overproducing strain for lignocellulosic biomass; of which glucose is the major sugar component. Inactivation of ethanol biosynthesis in this new strain may further improve 2,3-BD titer, yield and productivity. Addition of small amounts of CaCl2 has promise as a means of mitigating metabolic disruptions that might arise following metabolic engineering of P. polymyxa, which typically occur in solvent-producing Gram positive bacteria.
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Table 6.1. Comparison of Protein sequence between P. polymyxa DSM 365 levansucrase and other P. polymyxa strains with complete genome sequence using NCBI Blastp algorithm for alignment.
S/N Levansucrase protein identity (%) P. polymyxa DSM 365 1 P. polymyxa SC2 95 2 P. polymyxa E681 97 3 P. polymyxa M1 95 4 P. polymyxa CR1 97
233
Table 6.2. Comparison of nucleotide sequence between P. polymyxa DSM 365 levansucrase and other P. polymyxa strains with complete genome sequence using NCBI
Blastn algorithm for alignment.
S/N Levansucrase nucleotide P. polymyxa DSM 365 Accession number sequence identity (%) 1 P. polymyxa SC2 92 NC_014622.2 2 P. polymyxa E681 96 NC_024483.2 3 P. polymyxa M1 92 NC_017542.1 4 P. polymyxa CR1 96 NC_023037.2
234
Table 6.3. List of primers and PCR strategies used to generate levansucrase inactivation construct.
S/ Gene/gene Primers sequence (5’-3’) and PCR strategy N fragment 1 Levansucrase LevFragA_fwd: TGG GGA TCC TTG AAG TTT AAC AAA TGG TTC AGT AAA GC gene LevFragB_rev: GGG CTC GAG TTA TTT CTT TCC ATA CTC ATT TGG AG PCR strategy: Levansucrase gene was amplified from P. polymyxa using LevFragA_fwd and LevFragB_rev (AT1 = 59 ºC and AT2 = 61 ºC). NB: the underlined sequences in LevFragA_fwd and LevFragB_rev are BamHI and XhoI restriction sites, respectively 2 Erythromyci Erm_fwd: TAA CTG TTT AGG AGG ACT GAT AAT ATG AAC AAA AAT ATA AAA TAT TCT n gene CAA AAC (ERM) Erm_rev1: TAA AAA AAT AAG AGT TAC CAT TTA TTA TTT CCT CCC GTT AAA TAA TAG ATA AC Erm_rev2: GCC ACT ATG AAA CAA TAT TAA AAA AAT AAG AGT TAC CAT TTA TTA TTT CC PCR strategy: (1) Erythromycin gene was amplified from pMutin using Erm_fwd and Erm_rev1 primer
235 pair (AT1 = 58 ºC and AT2= 61ºC) (2) next, erythromycin gene from (1) was re-amplified using Erm_fwd and Erm_rev2 primer pair (AT1 = 69 ºC and AT2 = 60 ºC). NB: the use of Erm_rev2 brings
into the erythromycin gene the complete transcription terminator sequence. The underlined sequence in Erm_fwd represents the ribosomal binding site. 3 LevFragA LevFragA_fwd: TGG GGA TCC TTG AAG TTT AAC AAA TGG TTC AGT AAA GC LevFragA_rev: CCT CCT AAA CAG TTA GGA CGG AAC CTC ATA TTT CTC TTT GCC PCR strategy: Levansucrase fragment A (LevFragA) was amplified from levansucrase gene using LevFragA_fwd and LevFragA_rev primer pair (AT1 = 73 ºC and AT2 = 69 ºC). 4 LevFragB LevFragB_fwd: ACT CTT ATT TTT TTA ATA TTG TTT CAT AGT GGC AAT AAC GTA GTC G LevFragB_rev: GGG CTC GAG TTA TTT CTT TCC ATA CTC ATT TGG AG PCR strategy: Levansucrase fragment B (LevFragB) was amplified from levansucrase gene using LevFragB_fwd and LevFragB_rev primer pair (AT1 = 70 ºC and AT2 = 61 ºC).
Continued below table
235
Table 6.3 Continued above table.
S/ Gene/gene Primers sequence (5’-3’) and PCR strategy N fragment 5 LevFragA- LevFragA_fwd: TGG GGA TCC TTG AAG TTT AAC AAA TGG TTC AGT AAA GC ERM LevFragA_rev: CCT CCT AAA CAG TTA GGA CGG AAC CTC ATA TTT CTC TTT GCC Erm_fwd: TAA CTG TTT AGG AGG ACT GAT AAT ATG AAC AAA AAT ATA AAA TAT TCT CAA AAC Erm_rev1: TAA AAA AAT AAG AGT TAC CAT TTA TTA TTT CCT CCC GTT AAA TAA TAG ATA AC Erm_rev2: GCC ACT ATG AAA CAA TAT TAA AAA AAT AAG AGT TAC CAT TTA TTA TTT CC PCR strategy: (1) amplified LevFragA from levansucrase gene using using LevFragA_fwd and LevFragA_rev primer pair (AT1 = 73 ºC and AT2 = 69 ºC) (2) amplified ERM using Erm_fwd and Erm_rev1 primer pair (AT1 = 58 ºC and AT2= 61ºC). ERM was re-amplified using Erm_fwd and
236 Erm_rev2 primer pair (AT1 = 69 ºC and AT2 = 60 ºC). (3) LevFragA and ERM amplicons were spliced via One-Step SOE-PCR: step1- LevFragA and ERM served as both templates and primers (forward and
reverse templating fragments respectively) to generate LevFragA-ERM and the annealing temperature represents the annealing temperature of the overlapping region. Step 2- the generated LevFragA-ERM was amplified using LevFragA_fwd and Erm_rev2 (AT1 = 73 ºC and AT2 = 68 ºC). 6 LevFragA- LevFragA_fwd: TGG GGA TCC TTG AAG TTT AAC AAA TGG TTC AGT AAA GC ERM- Erm_rev2: GCC ACT ATG AAA CAA TAT TAA AAA AAT AAG AGT TAC CAT TTA TTA TTT CC LevFragB LevFragB_fwd: ACT CTT ATT TTT TTA ATA TTG TTT CAT AGT GGC AAT AAC GTA GTC G LevFragB_rev: GGG CTC GAG TTA TTT CTT TCC ATA CTC ATT TGG AG PCR strategy: (1) Re-amplified LevFragA-ERM using primer pair LevFragA_fwd and Erm_rev2 (AT1 =73 ºC and AT2 = 68 ºC). (2) Re-amplified LevFragB using LevFragB_fwd and LevFragB_rev primer pair (AT1 = 70 ºC and AT2 = 69 ºC). (3) Next, LevFragA-ERM and LevFragB amplicons from (1) and (2), respectively, were spliced via One-Step SOE-PCR: step1- LevFragA-ERM and LevFragA served as both templates and primers (forward and reverse templating fragments respectively) to generate LevFragA-ERM-LeveFragB and the annealing temperature represents the annealing temperature of the overlapping region. Step 2- the generated LevFragA-ERM-LevFragB was amplified using LevFragA_fwd and LevFragB_rev primer pair (AT1 = 73 ºC and AT2 = 69 ºC).
236
Table 6.4. List of microorganisms, vectors and enzymes used in this study and their respective characteristics and sources
Strain/vector/enzymes Characteristics Source Strains E.coli JM109 endA1, recA1, gyrA96, relA1 Promega Corporation P. polymyxa DSM 365 Wildtype DSMZ, Germany P. polymyxa DSM 365 Lev null ∆Lev; Ermr This study mutant Vectors pMutin Ermr Bacillus Genetic Stock Center, OH pGEM7Zf(+) Ampr, f1 oriC, lacZ Promega Corporation Enzymes GXL DNA polymerase High fidelity, amplifies GC-rich Takara Clontech templates BamHI - New England Biolabs XhoI - New England Biolabs T4 DNA ligase - New England Biolabs
237
Table 6.5. Substrate consumed, growth, maximum products, 2,3-BD yield and productivity during sucrose fermentation by P.
polymyxa DSM 365 wildtype and levansucrase null mutant.
Treatment WT+0 g/L M+0 g/L WT+0.2 g/L M+0.2 g/L WT+0.4 g/L M+0.4 g/L CaCl2 CaCl2 CaCl2 CaCl2 CaCl2 CaCl2
a b a a a b Sucrose consumed 101.9±0.3 84.7±3.0 110.1±0.5 84.5±1.2 107.9±0.4 92.8±1.3 (g/L) a b a b a b Residual sucrose (g/L) 2.49±0.0 25.4±3.4 ND 25.6±1.7 ND 17.3±1.8 a b a a a b Residual glucose (g/L) 5.7±0.1 ND Nd Nd 2.2±0.0 ND a b a b a b Max. growth 6.8±0.4 9.7±0.4 10.4±1.1 12.7±1.1 9.6±0.9 12.8±0.9 (OD600nm)
238 a a a a a a Max. 2,3-BD (g/L) 32.6±0.7 30.9±2.3 36.3±2.0 35.7±1.7 36.1±2.5 37.4±0.9
a b a b a b EPS (g/L) 17.4±2.2 3.0±0.6 18.4±2.7 2.9±0.5 17.6±0.36 2.9±0.2 a a a b a a 2,3-BD Yield (g/ g) 0.32±0.01 0.35±0.02 0.33±0.02 0.42±0.03 0.33±0.02 0.40±0.02 a a a a a a 2,3-BD Productivity 0.68±0.01 0.62±0.05 1.01±0.05 1.04±0.14 0.75±0.05 0.78±0.02 (g/L/h) a b a b b b Acetoin (g/L) 2.6±0.2 4.1±0.4 21.6±3.8 5.0±0.5 1.3±0.4 6.0±0.6 a b a a a a Ethanol (g/L) 5.7±0.3 6.8±0.1 5.8±0.2 5.1±0.4 5.8±1.1 5.5±0.5 a b a b a b Acetic acid (g/L) 1.5±0.3 2.8±0.1 1.8±0.1 3.9±1.2 1.2±0.3 5.5±0.2 Fisher’s LSD pairwise comparisons between wildtype and levansucrase mutant were conducted. Treatments with different
superscripts across a row are significant at p < 0.05. The maximum acetoin, ethanol and acetic acid generated during
fermentations are reported.
238
Table 6.6. Substrate consumed, growth, maximum products, 2,3-BD yield and productivity during glucose fermentation by P.
polymyxa DSM 365 wildtype and levansucrase null mutant.
Treatment WT+0 g/L M+0 g/L WT+0.2 g/L M+0 g/L WT+0.4 g/L M+0.4 g/L CaCl2 CaCl2 CaCl2 CaCl2 CaCl2 CaCl2
a a a a a b Glucose consumed 85.6±3.9 87.8±1.9 96.1±3.1 105.0±0.0 94.7±2.7 106.7±2.5 (g/L) a a a b a b Residual glucose (g/L) 23.7±3.2 21.45±1.2 13.2±2.3 4.3±0.7 14.5±2.0 2.5±1.8 a b a b a b Max. growth 8.9±0.2 12.1±1.3 11.2±0.2 15.4±1.2 10.5±0.3 16.1±0.7 (OD600nm) a a a a a a 239 Max. 2,3-BD (g/L) 30.7±1.2 29.8±2.0 34.2±2.7 39.0±1.3 34.5±3.4 39.4±4.4 a b a b a b EPS (g/L) 5.5±0.4 2.27±0.1 4.2±0.2 2.4±0.3 5.4±0.5 2.8±0.2 a a a a a a 2,3-BD Yield (g/ g) 0.36±0.01 0.34±0.02 0.36±0.02 0.37±0.01 0.37±0.04 0.38±0.04 a a a b a b 2,3-BD Productivity 0.51±0.02 0.57±0.09 0.71±0.06 1.62±0.05 0.72±0.07 1.64±0.19 (g/L/h) a b a b a b Acetoin (g/L) 3.3±0.0 2.4±0.4 10.1±2.1 2.8±0.4 8.8±1.1 2.6±0.3 a b a b a b Ethanol (g/L) 5.8±0.1 7.1±0.3 5.6±0.4 8.7±0.7 5.4±0.5 9.4±0.2 a b a b b b Acetic acid (g/L) 0.0±0.0 1.7±0.0 0.6±0.0 2.5±0.9 0.0±0.0 2.9±0.1 Fisher’s LSD pairwise comparisons between wildtype and levansucrase mutant were conducted. Treatments with different
superscripts across a row are significant at p < 0.05. The maximum acetoin, ethanol and acetic acid generated during
fermentations are reported.
239
Table 6.7. Comparison of fermentations with and without erythromycin supplementation during stability test of P. polymyxa
levansucrase null mutant.
Genera Growth 2,3-BD EPS Acetoin Ethanol Acetic tion (OD600nm) (g/L) (g/L) (g/L) (g/L) acid (g/L) a a a a a a G0 35 µg/ml erythromycin 11.6±0.9 35.3±3.7 3.7±0.4 5.0±0.5 7.0±0.8 2.9±0.1 a a a a a a 0 µg/ml erythromycin 12.5±0.5 36.4±0.1 3.6±0.4 4.8±0.2 8.1±0.1 2.9±0.2 a a a a a a G10 35 µg/ml erythromycin 11.8±0.6 38.4±4.1 3.1±0.6 5.7±1.2 5.9±0.4 3.5±0.6 a a a a a a 240 0 µg/ml erythromycin 13.2±0.5 39.0±2.3 3.3±0.2 4.6±1.2 6.7±0.6 2.5±0.2 a a a a a a G20 35 µg/ml erythromycin 13.3±0.9 38.7±3.0 3.9±0.5 8.0±1.8 7.5±0.5 3.1±0.5 a a a b a a 0 µg/ml erythromycin 14.6±0.6 39.0±1.4 2.7±0.3 5.7±0.2 6.8±0.0 3.2±0.2 a a a a a a G30 35 µg/ml erythromycin 11.9±0.5 37.5±1.9 3.8±0.3 7.4±1.2 9.6±0.8 3.2±0.1 b a a b b a 0 µg/ml erythromycin 15.0±0.3 35.6±1.3 3.4±0.1 5.3±0.9 6.3±0.3 2.7±0.3 a a a a a a G40 35 µg/ml erythromycin 11.0±1.0 39.0±0.4 4.1±0.1 7.0±0.9 9.2±0.6 3.1±0.1 a a b a b a 0 µg/ml erythromycin 13.2±1.2 36.2±1.4 3.4±0.1 5.6±1.0 7.5±0.8 3.1±0.3 a a a a a a G50 35 µg/ml erythromycin 13.9±0.7 39.4±2.3 3.3±0.4 9.4±0.7 9.0±0.0 3.1±0.1 a a a b b a 0 µg/ml erythromycin 15.1±0.0 37.6±0.9 3.5±0.2 6.1±0.2 7.3±0.5 3.1±0.0 Fisher’s LSD pairwise comparisons between 35 and 0 µg/ml erythromycin supplemented levansucrase mutant treatments were
conducted. Treatments with different superscripts down a column are significant at p < 0.05.
240
Figure 6.1. Levansucrase inactivation construct generation. A: levansucrase gene was amplified from the genome of P. polymyxa and was used to generate the inactivation construct with erythromycin gene placed between the upstream (210 bp) sequence
(LevFragA) and downstream (213 bp) sequence (LevFragB) of levansucrase gene. The construct was ligated into previously double digested pGEM7Zf(+) and used to inactivate levansucrase gene in the chromosome of P. polymyxa via double-cross homologous recombination. The red dots at the end of LevfragA and ERM represent stop codons, whereas the green and blue dots represent ribosome binding site and transcription termination sequence, respectively. B. The gene fragments showing levansucrase gene,
LevFragA, LevFragB, ERM, LevFragA-ERM and LevFragA-ERM-LevFragB gene fragments during generation of levansucrase inactivation construct.
241
242
Figure 6.2. A:Gel image showing colony PCR of P. polymyxa levansucrase null mutant. Lanes 1-3,5 -7 and 8-10 show bands
corresponding to LevFragA-ERM, ERM and ERM-LevfragB gene fragments, respectively. Lane 4 is 1kb DNA ladder. Lanes
1,5 and 8 represent bands from pGEM7Zf+ harboring the levansucrase inactivation construct; lanes 2,6 and 9 represent bands
from colony1 and lanes 3,7 and 10 represent bands from colony2. B: Precipitation of EPS in the fermentation broth of P.
polymyxa wildtype and levansucrase null mutant in sucrose medium.
242
243
Figure 6.3: The schematic representation of step-by-step procedure employed to evaluate the stability of the P. polymyxa
levansucrase null mutant.
243
244
Figure 6.4: Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in sucrose cultures without CaCl2
supplementation.
244
245
Figure 6.5: Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in sucrose cultures supplemented with
0.2 g/L CaCl2.
245
246
Figure 6.6: Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in sucrose cultures supplemented with
0.4 g/L CaCl2.
246
247
Figure 6.7: Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in glucose cultures without CaCl2
supplementation.
247
248
Figure 6.8: Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in glucose cultures supplemented with
0.2 g/L CaCl2.
248
249
Figure 6.9: Fermentation profile of P. polymyxa levansucrase null mutant and wildtype in glucose cultures supplemented with
0.4 g/L CaCl2.
249
250
Figure 6.10: Stability test: product profile in sucrose-based medium and 0. 4 g/L CaCl2 with 35µg/ml erythromycin
supplementation.
250
251
Figure 6.11: Stability test: product profile in sucrose-based medium and 0.4 g/L CaCl2 without erythromycin supplementation.
251
Figure 6.12: Agarose (1%) gel images of stability test colony-PCR of P. polymyxa levansucrase null mutant. All colonies from G0 (a and b) to G50 (a and d) showed that the mutant retained the erythromycin gene during several generations grown with (a and c) or without (b and d) erythromycin supplementations.
252
Figure 6.13: Replica plates showing genetic stability test of P. polymyxa levansucrase null mutant. Selected colonies from TSA plates without erythromycin (a and b) were transferred to fresh replica TSA plates containing 35 µg/ml erythromycin (a and d). All the selected colonies from G0 to G50 (cultivated with or without erythromycin) transferred from TSA plate with no erythromycin to a replica plate containing antibiotic all grew showing resistance to erythromycin.
253
Figure 6.14 Determination of generation (doubling time) of levansucrase null mutant
254
Chapter 7: Conclusions and recommendations
7.1 Conclusions
Low 2,3-BD yield, high cost of conventional sugar substrates, formation of 2,3-
BD competing products during fermentation, and high cost of 2,3-BD recovery from the fermentation broth due to the high boiling point of 2,3-BD (~180 ºC), are major factors impeding commercialization of biological production of 2,3-BD. Additionally, the majority of 2,3-BD producing microorganisms such as Klebsiella pneumoniae, K. oxytoca, Enterobacter aerogenes, E. cloacae and Serratia marcescens are pathogenic, which limits their use in industrial processes due to liability issues. Another major concern with pathogenic 2,3-BD producing microorganisms is their inability to produce levo-2,3-BD, the most desired 2,3-BD isomer. The levo-2,3-BD is desired because it can easily be dehydrated to 1,3-butadiene, the monomer from which synthetic rubber is produced. In light of these concerns, the non-pathogenic Paenibacillus polymyxa, which is capable of accumulating up to 98% levo-2,3-BD as opposed to meso-2,3-BD isomer during fermentation, was selected for this study for development into metabolic workhorse for 2,3-BD production.
The objectives of this study were to (1) identify key fermentation parameters that influence 2,3-BD fermentation by P. polymyxa and optimize them for maximum 2,3-BD production, (2) investigate 2,3-BD-mediated feedback inhibition during 2,3-BD
255 fermentation, (3) evaluate the feasibility of using readily available non-food lignocellulosic biomass (LB) as substrate for 2,3-BD fermentation, and (4) inactivate exopolysaccharide (EPS) production pathway of P. polymyxa and minimize EPS production during 2,3-BD fermentation. The chapters of this dissertation described series of experiments conducted to accomplish these objectives, and obtained results and their scientific significance are summarized in chapters 3 – 6.
Chapter 3 examined the impact of factors - yeast extract, tryptone, ammonium acetate, ammonium sulfate, crude glycerol concentration, inoculum size, and fermentation temperature on 2,3-BD production by P. polymyxa. One-factor-at-a-time experiments were conducted and results show that seven of these factors influenced, to various degrees, 2,3-BD production by P. polymyxa. Using Plackett-Burman experimental design for data analysis, results showed that only tryptone, temperature, and inoculum size had significant effect on 2,3-BD production by P. polymyxa. Using Box-
Behnken design and response surface methodology, fermentation parameters influencing
2,3-BD production by P. polymyxa was optimized, increasing 2,3-BD concentration from
~27 g/L to 51.1 g/L in batch bioreactor cultures, representing 89% increase in 2,3-BD production. In fed-batch cultures, 2,3-BD production increased from ~47 g/L to 68.5 g/L, representing ~46% increase in 2,3-BD production. Although glycerol addition in the fermentation broth did not significantly increase 2,3-BD production, the presence 7 g/L in the fermentation medium was found to improve overall 2,3-BD production because glycerol catabolism by fermenting microbes leads to NADH regeneration, which provides more electrons needed for the production of reduced fermentation end-products
256 like 2,3-BD and for detoxification of lignocellulose-derived microbial inhibitory compounds (LDMICs) (Lin, 1976; Neijssel et al., 1975; Palmqvist and Hahn-Hagerdal,
2000; Ujor et al., 2015). In addition to improvement in 2,3-BD production by P. polymyxa, ethanol and EPS formation were reduced by 11% and 19%, respectively, under optimized conditions. Through optimization design and experiments, the amount of expensive media components such as tryptone and yeast extract was cut down by 50% and 29%, respectively, which will reduce the operating cost of microbial 2,3-BD production, and ultimately, improve the economics of 2,3-BD production by P. polymyxa.
Feedback inhibition of 2,3-BD formation in fed-batch cultures occurred when 2,3-
BD concentration in the fermentation medium reached 6% with concomitant accumulation of acetoin. Therefore, Chapter 4 evaluated the response of P. polymyxa to high concentrations of levo-2,3-BD during growth and 2,3-BD fermentation. Levo-2,3-
BD inhibited the growth of P. polymyxa in a concentration dependent manner, exerting complete growth inhibition when concentration attained 60 g/L. However, although P. polymyxa achieved considerable growth at 60 g/L levo 2,3-BD when its exposure to it was incremental, 2,3-BD was reconverted to acetoin, possibly to alleviate 2,3-BD toxicity. High concentration of levo 2,3-BD in the bioreactor was found to stimulate the production of the meso-2,3-BD isomer by P. polymyxa. It was also found that meso-2,3-
BD exhibited lower growth inhibitory effect on P. polymyxa than levo-2,3-BD.
Therefore, we hypothesized that production of meso-2,3-BD by P. polymyxa during high levo-2,3-BD concentration could be one mechanism with which it uses to alleviate levo-
257
2,3-BD toxicity during fermentation. The results obtained in this study could shed light on why some pathogenic 2,3-BD producing microorganisms are capable of producing meso-2,3-BD at concentrations exceeding 100 g/L whereas P. polymyxa is unable to produce more than 6% levo-2,3-BD.
Due to the high cost of conventional sugar substrates currently used in fermentations and impact on the overall production cost of biofuels and chemicals,
Chapter 5 investigated the feasibility of using readily available non-food lignocellulosic biomass (LB) and the monomeric sugars (hexose and pentose) that make up the LB as alternative substrates for 2,3-BD fermentation by P. polymyxa. To release fermentable sugars, LB must first be pretreated to deconstruct the lignin matrix for easy access of hydrolytic enzymes to cellulose and hemicellulose components of LB, which leads to the generation of undesirable LDMICs that poses significant roadblock to sugar utilization by fermenting microorganisms. The ability of P. polymyxa to utilize the representative mixed sugars (glucose, xylose and arabinose) of LB was evaluated. Interestingly, P. polymyxa was found to ably co-metabolize and ferment the representative mixed sugars
(glucose, xylose and arabinose) components of LB to 2,3-BD without showing signs of carbon catabolite repression characteristics. This is a first report on mixed sugar fermentation by P. polymyxa. The growth of P. polymyxa in 60%, 80% and 100% WSH- based media showed 17%, 27% and 32% increases, respectively, relative to the glucose- based control medium. The level of growth observed with WSH when compared to the glucose control suggests that P. polymyxa may have sequestered additional carbons from the LDMICs present in WSH. In addition, some components of LDMICs, especially at
258 concentrations present in the WSH, may have stimulated the growth of P. polymyxa. 2,3-
BD production was 32, 31 and 23 g/L in 60%, 80% and 100% WSH, respectively, which was comparable to 32 g/L obtained in the glucose control. Although fermentations using hydroxymethylfurfural (HMF), a LDMIC, as sole carbon source showed at least a 2.4- fold increase in the growth of P. polymyxa relative to the control with no carbon source, suggesting that P. polymyxa might have utilized HMF in WSH for cell biomass accumulation. Further studies are necessary to confirm that P. polymyxa can use HMF as sole carbon sole for cell growth. In addition, P. polymyxa showed robust tolerance to furfural, coumaric acid, vanillic acid and vanillin during fermentation.
Although, the optimization study in Chapter 3 reduced EPS formation by 19%,
EPS production remained a problem during 2,3-BD fermentation. Carbons are diverted to
EPS during fermentation, thus decreasing 2,3-BD yield. Further, EPS formation constitutes nuisance by increasing medium viscosity and clogging bioreactor lines, which prevent proper mixing of medium during fermentation. More importantly, additional purification steps are required to remove EPS prior to 2,3-BD separation which add to the overall cost of 2,3-BD production. To address the problem of EPS formation, metabolic engineering strategy was explored to inactivate EPS biosynthesis pathway of P. polymyxa. Levansucrase gene (which encodes levansucrase enzyme responsible for EPS biosynthesis) of P. polymyxa was disrupted via double cross homologous recombination
(Chapter 6). The resulting P. polymyxa levansucrase null mutant showed 34% and 54% increase in growth in sucrose and glucose media, respectively, when compared to the P. polymyxa wildtype. Additionally, the levansucrase null mutant also exhibited 6.4- and
259
2.4-folds decrease in EPS formation in sucrose and glucose media, respectively, relative to the P. polymyxa wildtype. The reduction in EPS production by the levansucrase null mutant resulted in 27% and 4% increase in 2,3-BD yield, and 4% and 128% increase in
2,3-BD productivity in sucrose and glucose media, respectively, when compared to the P. polymyxa wildtype. Following evaluation of the stability of P. polymyxa levansucrase null mutant in the presence and absence of erythromycin antibiotic, the levels of 2,3-BD and EPS produced by the P. polymyxa levansucrase null mutant did not vary, even after fifty growth generations. Inactivation of P. polymyxa levansucrase gene led to the development of a stable and faster growing P. polymyxa strain with significantly reduced
EPS production.
Collectively, this study clearly showed use of novel strategies to enhance 2,3-BD production by P. polymyxa. This study also generated a novel strain of P. polymyxa with efficient 2,3-BD production and reduced EPS formation during fermentation, showing that P. polymyxa has the potential of being 2,3-BD fermentation workhorse and for improving the economics of commercial microbial 2,3-BD production.
7.2 Recommendations
This study focused on the development of process design and metabolic engineering to improve 2,3-BD production by P. polymyxa DSM 365 using mixed sugars and lignocellulosic biomass hydrolysates. While considerable success was achieved, some challenges still remain which should be addressed in future works. Additionally, some aspects of 2,3-butanediol fermentation research were not considered in this work
260 due to time constraints. Consequently, the following recommendations are suggested for further studies:
Since P. polymyxa produces meso-2,3-BD when the concentration of levo-2,3-BD is high in the bioreactor (see objective 2), and butanediol dehydrogenase (a single enzyme in P. polymyxa) catalyzes the biosynthesis of levo- and meso-2,3-BD, understanding the underlying mechanism(s) of meso-2,3-BD biosynthesis would provide insights for possible engineering of butanediol dehydrogenase protein for selective production of levo-2,3-BD.
The developed P. polymyxa levansucrase null mutant in this study produced
~74% more ethanol in the glucose medium than the wildtype. Hence, inactivation of ethanol biosynthesis pathway of P. polymyxa to further improve 2,3-BD yield and productivity is recommended. Bioinformatics search of the genome of P. polymyxa shows six genes annotated as alcohol dehydrogenase capable of ethanol biosynthesis from acetaldehyde. However, P. polymyxa possesses only a single copy of bifunctional acetaldehyde-CoA/alcohol dehydrogenase (BAD) gene that can convert acetyl CoA to acetaldehyde and then to ethanol. Targeting BAD for inactivation could reduce ethanol production in P. polymyxa. Considering also that BAD produces acetaldehyde, the substrate for the six alcohol dehydrogenases in P. polymyxa, it will be relevant to inactivate BAD. In addition, targeting all six alcohol dehydrogenases of P. polymyxa for inactivation may result in impaired P. polymyxa growth as some of the alcohol dehydrogenases may play crucial roles for cell survival.
261
During this study, some efforts were made to separate 2,3-BD from fermentation broth via vacuum-assisted 2,3-BD distillation. While 2,3-BD was successfully separated from the fermentation broth and up to 96% purity achieved, the 2,3-BD obtained from the spent broth was decolorated. Discoloration of 2,3-BD occurred during recovery from the fermentation broth due to the high temperatures (150-180 °C) used for the separation method that possibly caused caramelization of residual sugars, medium components, and other fermentation products. Also, the high temperatures used for the 2,3-BD recovery may have resulted in the formation of esters between the hydroxyl groups of 2,3-BD and acetic, lactic or other acids in the spent broth. Therefore, development of efficient salting out process for the removal of acids from the fermentation broth prior to 2,3-BD distillation is recommended. In addition, 2,3-BD distillation should be conducted under vacuum to drastically reduce the boiling point of 2,3-BD for efficient recovery from the fermentation broth.
Given that > 5% levo-2,3-BD in the bioreactor poses strong inhibitory effects on
P. polymyxa, it is crucial to develop integrated 2,3-BD fermentation and in situ 2,3-BD recovery to further increase production efficiency. However, it is necessary to identify or develop membrane filters that can selectively allow easy passage of 2,3-BD for this process to be feasible.
262
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Appendix A One factor-at-a-time experiments
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Figure A.1. One-factor at a time experiments showing effects of crude glycerol on 2,3-
BD production by P. polymyxa. Inoculum size, temperature, yeast extract, tryptone, ammonium sulfate and ammonium acetate were kept at 8%, 37 ºC, 7g/L, 7 g/L, 3g/L and
5 g/L, respectively. Error bars show standard deviations of means (n=3).
301
Figure A.2. One-factor at a time experiments showing effects of temperature on 2,3-BD production by P. polymyxa. Inoculum size, glycerol, yeast extract, tryptone, ammonium sulfate and ammonium acetate were kept at 8%, 10 g/L, 7g/L, 7 g/L, 3g/L and 5 g/L, respectively. Error bars show standard deviations of means (n=3).
302
Figure A.3. One-factor at a time experiments showing effects of inoculum size on 2,3-
BD production by P. polymyxa. Temperature, glycerol, yeast extract, tryptone, ammonium sulfate and ammonium acetate were kept at 37 ºC, 10 g/L, 7g/L, 7 g/L, 3g/L and 5 g/L, respectively. Error bars show standard deviations of means (n=3).
303
Figure A.4. One-factor at a time experiments showing effects of ammonium sulfate on
2,3-BD production by P. polymyxa. Inoculum size, temperature, glycerol, yeast extract, tryptone and ammonium acetate were kept at 8%, 37 ºC, 10 g/L, 7g/L, 7 g/L and 5 g/L, respectively. Error bars show standard deviations of means (n=3).
304
Figure A.5. One-factor at a time experiments showing effects of ammonium acetate on
2,3-BD production by P. polymyxa. Inoculum size, temperature, glycerol, yeast extract, tryptone and ammonium sulfate were kept at 8%, 37 ºC, 10 g/L, 7g/L, 7 g/L and 3 g/L, respectively. Error bars show standard deviations of means (n=3).
305
Figure A.6. One-factor at a time experiments showing effects of tryptone on 2,3-BD production by P. polymyxa. Fig. A5. One-factor at a time experiments showing effects of ammonium acetate on 2,3-BD production by P. polymyxa. Inoculum size, temperature, glycerol, yeast extract, ammonium sulfate and ammonium acetate were kept at 8%, 37 ºC,
10 g/L, 7g/L, 3 g/L and 5 g/L, respectively. Error bars show standard deviations of means
(n=3).
306
Figure A.7. One-factor at a time experiments showing effects of yeast extract on 2,3-BD production by P. polymyxa. Inoculum size, temperature, glycerol, tryptone, ammonium sulfate and ammonium acetate were kept at 8%, 37 ºC, 10 g/L, 7g/L, 3 g/L and 5 g/L, respectively. Error bars show standard deviations of means (n=3).
307
Figure A.8. Competing products generated during 2,3-BD fermentation with the un- optimized medium and conditions.
308
Appendix B CaCO3 supplementation in wildtype and levansucrase null mutant cultures
309
Figure B.1. Fermentation profile of levansucrase null mutant in glucose medium supplemented with 4 g/L CaCO3. A, cell growth (OD600nm); B, culture pH; C, 2,3-BD concentration; and D, EPS concentration. Levansucrase mutants A and B are the same but from different colonies.
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Figure B.2. Fermentation profile of levansucrase null mutant in glucose medium supplemented with 4 g/L CaCO3. A, acetoin; B, ethanol; and C, acetic acid concentrations. Levansucrase mutants A and B are the same but from different colonies.
311