MICROBIAL ELECTROSYNTHESIS IN SHEWANELLA ONEIDENSIS MR-1 By Kathryne Caldwell Ford A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of Microbiology and Molecular Genetics – Doctor of Philosophy 2023 ABSTRACT Shewanella oneidensis MR-1 is a model electroactive bacterium that has been extensively studied for both foundational mechanisms and biotechnology. Despite this, the extent and complexity of the extracellular electron transport (EET) pathway is still being unraveled. This field of study is further complicated because cells modulate the flow of electrons based on rate of flux, redox partner, and whether electrons are being exported or imported into the cell. In this work, I explore the EET pathway of S. oneidensis in the context of microbial electrosynthesis (MES) optimization. MES technology aims to use microorganisms as biocatalysts to drive the formation of useful chemical products in a bioelectrochemical system (BES), ideally using carbon dioxide (CO2) as the substrate. Here, I will examine the native EET systems and synthetic biology efforts to engineer a strain of S. oneidensis capable of such electroautrophy for bioproduction. In Chapter 2, I look at the influence of oxygen (O2) on MES efficiency for 2,3- butanediol generation in S. oneidensis. To do this, butanediol dehydrogenase is expressed in wild-type (WT) S. oneidensis cells to catalyze the NADH-dependent reduction of acetoin to 2,3-butanediol. Our research group previously showed that electron uptake from a cathode to form NADH is an energetically unfavorable reaction and overcame this thermodynamic barrier through expression of the proton pump proteorhodopsin (PR). In the new design, the reaction is coupled to the energetically favorable reduction of O2 by native oxidase; during this bidirectional electron transfer, electrons from the cathode power both reactions. In Chapter 3, I use this same system to reassess the contribution of major cytochrome proteins in the EET pathway during MES. I demonstrate that the outer membrane MtrCAB complex is essential for this process, while other components like CymA and FccA have a more flexible role. Importantly, I show that exogenous flavins are unable to compensate for the loss of natively produced flavins for 2,3-butanediol production, despite their apparent influence on cathodic current. Finally, I reexamine the role of hydrogenases in this process, demonstrating their importance for cell survival on the electrode. Chapters 4 and 5 of this dissertation focus on the use of synthetic biology techniques to install a CO2 fixation pathway in the heterotrophic S. oneidensis. To achieve this goal, I combine in silico metabolic modeling with a CRISPRi knockdown system to create a strain in which multiple substrates are required for biomass synthesis and energy acquisition. By then expressing RuBisCO and PrkA in this strain (∆gpmA pCBB), I then devise a laboratory evolution experiment to generate a strain that will use CO2 to build biomass. In summary, there is still much to be understood about EET in S. oneidensis and an increasing array of bioengineering tools that can be used to this end. This work does just this by exploring the energetics and physiology of S. oneidensis’s EET network, as well as laying the groundwork for a functional electroautotrophic chassis for carbon- neutral bioproduction. This work is dedicated to my friends and loved ones. Those who made me food when I was too busy to cook, spent long days working alongside me, traveled across the world with me, and had social distance hangouts when we couldn’t be together. Though we continue to go our separate ways as we grow older, I will always carry your love with me. This is also dedicated to my oldest companion, Buttons, and my newest family members who arrived during my graduate school, Zachary, Abigail, and Gabriel. iv ACKNOWLEDGEMENTS There are many people I would like to thank for their support, mentorship, and guidance during my nine years at Michigan State University. I would like to thank the Michigan State University Honors College for offering me a spot in their Professorial Assistantship program, allowing me to begin my scientific journey in my first year of undergraduate studies. Similarly, the Bailey Scholars program and Clean Water Action, two communities that nurtured my interest in environmental protection and provided a space to both live and grow. These opportunities put me on my path towards pursuing applicable solutions to the environmental issues we currently and will continue to face. I am grateful to the Microbiology and Molecular Genetics, and Biochemistry and Molecular Biology departments that have been a welcoming environment to foster collaboration and growth during my tenure. Their financial support has enabled me to travel to research conferences around the world and maintain connections with exceptional researchers. Thank you to the National Science Foundation for the Graduate Student Research Fellowship that gave me the freedom to pursue work that inspired me, as well as their trust in my scientific future. I express my deepest gratitude to those who mentored me, fostering my scientific skills, nurturing integrity, and cultivating critical thinking abilities. My senior undergraduate work with Dr. Matthew Schrenk and the members of his research team inspired me to remain at Michigan State for graduate school. As well as his continued mentorship as a member of my thesis committee, alongside two inspirational scientists, Dr. Gemma Reguera and Dr. Robert Hausinger. Thank you for pushing my research abilities forward and help me create a scientific body of work of which I am immensely v proud. Additional thanks to those in the department who kept me sane through many a coffee break and commiserating chat, especially Dr. Kazem Kashefi, (soon-to-be-Dr.) Osama Alian, and Dr. Kody Duhl. There are not enough words to express my appreciation and gratitude for my mentor Dr. Michaela TerAvest and the members of our research team; Shaylynn Miller, Megan Gruenberg, Nicholas Tefft, Dr. Magda Felczak, and everyone who passed through during my time. Though I was hesitant to join a biochemistry and electromicrobiology lab, I am so glad that I did. Michaela’s mentorship has been invaluable to my growth as a scientist, collaborator, and person. She helped me maintain focus on my project while giving me the freedom to explore my crazy ideas. I am grateful to her for being an inspiration to what effective, successful, and thoughtful academic mentorship can be. To my lab mates: thank you for being there to bounce ideas off of, help me finish an experiment, and moral support for every time a bioreactor leaked or the HPLC errored. I know that without your help and company, my time in graduate school would have been difficult and lonely. Finally, I want to thank my loved ones. For my family that pushed me to excel and always believed in me. For my friends who kept me sane through food, movie nights, and great company. I will hold your friendships close to my heart as we grow and move away, change careers, experience loss, and find others we care for. Thank you to my partner John Shook, who has been my rock and my light at the end of the tunnel. You are one of the most naturally gifted scientists I have ever known, your laughter is infectious and always brings me joy, and I am excited to start the next chapter of my life with you. Last but not least, thank you to my little man and Zoom companion, Buttons. vi TABLE OF CONTENTS LIST OF ABBREVIATIONS .............................................................................................ix Chapter 1: Microbial Electrosynthesis in Shewanella oneidensis MR-1 .......................... 1 1.1 Abstract .................................................................................................................. 2 1.2 Introduction to Microbial Electrosynthesis .............................................................. 2 1.3 Technology Meets Microbiology ............................................................................. 4 1.4 EET Pathways ....................................................................................................... 7 1.4.1 Cytochrome-rich pathway .............................................................................. 10 1.4.2 Flavins ........................................................................................................... 12 1.4.3 Biofilms .......................................................................................................... 15 1.4.4 Nanowires ...................................................................................................... 18 1.5 Bioengineering Techniques.................................................................................. 20 1.5.1 Native Optimization ........................................................................................ 21 1.5.2 Tool Development .......................................................................................... 22 1.6 MES for Bioproduction ......................................................................................... 24 1.7 Research Overview .............................................................................................. 26 REFERENCES .......................................................................................................... 29 Chapter 2: The electron transport chain of Shewanella oneidensis MR-1 can operate bidirectionally to enable microbial electrosynthesis ....................................................... 45 2.1 Abstract ................................................................................................................ 46 2.2 Introduction .......................................................................................................... 46 2.3 Results and Discussion ........................................................................................ 51 2.3.1 Eliminating electrode-independent 2,3-BDO Production ................................ 51 2.3.2 Bidirectional Electron Transfer to Oxygen and NAD+..................................... 53 2.3.3 Reactive Oxygen Species Formation ............................................................. 57 2.4 Conclusion ........................................................................................................... 61 2.5 Materials and Methods ......................................................................................... 62 2.6 Author Contributions ............................................................................................ 66 2.7 Acknowledgements .............................................................................................. 66 REFERENCES .......................................................................................................... 67 APPENDIX A: Supplementary Figures for Chapter 2 ................................................. 73 Chapter 3: Flexibility of Inward Electron Transfer Pathway in Shewanella oneidensis MR-1 ............................................................................................................................. 76 3.1 Abstract ................................................................................................................ 77 3.2 Introduction .......................................................................................................... 77 3.3 Results and Discussion ........................................................................................ 80 3.3.1 Inward Electron Transport uses Mtr and CymA ............................................. 80 3.2.2 Flavins Have Complex Contributions to Electron Transport .......................... 83 3.3.3 Hydrogenases Enhance Cell Survival on Anode .......................................... 87 3.4 Conclusions ......................................................................................................... 89 3.5 Materials and Methods ......................................................................................... 91 3.6 Author Contributions ............................................................................................ 94 3.7 Acknowledgements .............................................................................................. 94 vii REFERENCES .......................................................................................................... 95 APPENDIX B: Supplementary Figures for Chapter 3 ............................................... 102 Chapter 4: Flux Balance Analysis and Mobile CRISPRi guided deletion of a conditionally essential gene in Shewanella oneidensis MR-1 .......................................................... 103 4.1 Abstract .............................................................................................................. 104 4.2 Introduction ........................................................................................................ 105 4.3 Results ............................................................................................................... 108 4.4 Discussion.......................................................................................................... 114 4.5 Materials and Methods ....................................................................................... 117 4.6 Acknowledgements ............................................................................................ 121 4.7 Author Contributions .......................................................................................... 121 REFERENCES ........................................................................................................ 122 Chapter 5: Engineering Strategy for a CO2 Fixation Pathway in S. oneidensis ........... 128 5.1 Introduction ........................................................................................................ 129 5.2 Results ............................................................................................................... 132 5.2.1 Determining Optimal Gene Knockout Strategy ............................................ 132 5.2.2 Investigating Growth Potential of ∆gpmA pCBB .......................................... 135 5.2.3 Designing a Directed Evolution Experiment ................................................. 139 5.3 Future Directions ................................................................................................ 140 5.4 Materials and Methods ....................................................................................... 142 REFERENCES ........................................................................................................ 147 Chapter 6: Conclusions and Future Directions ............................................................ 149 6.1 Conclusions and Significance ............................................................................ 150 6.1.1 Bidirectional Electron Transfer to Oxygen and NADH ................................. 150 6.1.2 Flexibility of the EET Pathway ..................................................................... 152 6.1.3 Synthetic Biology Strategies ........................................................................ 154 6.1.4 Engineering Carbon Fixation in S. oneidensis ............................................. 155 6.2 Future directions ................................................................................................ 155 REFERENCES ........................................................................................................ 159 viii LIST OF ABBREVIATIONS 2,3-BDO 2,3-butanediol apo- ‘without cofactor’ ATP Adenonine triphosphate BES Bioelectrochemical system BDH (Bdh) Butanediol dehydrogenase CB(B)C Calvin-Benson(-Bassham) Cycle CCCP Carbonyl cyanide m-chlorophenylhydrazone CMF Carbon microfiber CNF Carbon nanofiber CRISPRi Clustered regularly interspaced short palindromic repeats – interference CRISPRa Clustered regularly interspaced short palindromic repeats - activation DMSO Dimethyl sulfoxide DO Dissolved oxygen eDNA Extracellular DNA FAD Flavin adenine dinucleotide FBA Flux balance analysis FMN Flavin mononucleotide EAB Electroactive bacteria EET Extracellular electron transport EOC Exported organic carbon EPS Exopolymeric substances ETC Electron transport chain ix holo- ‘with cofactor’ HPLC High performance liquid chromotagraphy IPTG Isopropylthio-β-galactoside MES Microbial electrosynthesis MEC Microbial electrosynthesis cell MFC Microbial fuel cell NAD+ Nicotinamide adenine dinucleotide (oxidized form) NADH Nicotinamide adenine dinucleotide (reduced form) NAG N-acetylglucosamine NDH NADH dehydrogenase OMC Outer membrane cytochrome OME Outer membrane extension OMV Outer membrane vesicle PMF Proton motive force PR Proteorhodopsin ROS Reactive oxygen species RF Riboflavin sgRNA Small guide RNA SMF Sodium motive force SNP Small nucleotide polymorphism TCA Tricarboxcylic acid TMAO Trimethylamine N-oxide WT Wildtype x Chapter 1: Microbial Electrosynthesis in Shewanella oneidensis MR-1 Kathryne C. Ford1,2, Michaela A. TerAvest1* 1 Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA 2 Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI, 48824, USA 1 1.1 Abstract Human reliance on fossil fuels is unsustainable. As alternative technologies such as wind and solar electricity generation develop, researchers are simultaneously designing systems to store the electricity as chemical energy. Microbial electrosynthesis (MES) is a burgeoning field that offers a platform to do just this. In MES, electrons are taken up by microorganisms to drive targeted chemical reactions to produce useful products such as acetic acid and ethanol. This work will discuss the history and future of MES, specifically through the use of the exoelectrogenic bacterium Shewanella oneidensis MR-1. We will examine the physiology of electron transport in S. oneidensis, as well as how synthetic biology tools are being leveraged to optimize these pathways. 1.2 Introduction to Microbial Electrosynthesis The exchange of electrons is a fundamental biochemical process, ubiquitous and essential for life to exist. Cells rely on the uptake of electrons from feedstocks during catabolism, and the deposition of electrons onto readily available compounds. Understanding this flow of electrons, and how different species facilitate this flow, is fundamental to our knowledge of physiology, community interactions, and developing biotechnology. In natural environments, microorganisms evolve specialized mechanisms to fill specific ecosystem niches. This specificity means that the physiological capabilities of different bacterial species are closely tied to where they live. One of the strongest determinants is availability of terminal electron acceptors for respiration. Humans, most animals, and many microorganisms rely on aerobic respiration, utilizing oxygen as the terminal electron acceptor. In the absence of oxygen, organisms can use other molecules like nitrate or sulfate for anaerobic respiration, or internal redox 2 reactions to gain energy in a process called fermentation1. Some microorganisms have also evolved to use extracellular electron acceptors. This process requires specialized protein machinery to transport electrons from internal energy pools, across cellular membranes onto the cell surface to reduce available acceptors. This mechanism, termed extracellular electron transport (EET), allows microbes to use solid metal oxides like Fe(III) and Mn(IV) for anaerobic respiration. Researchers discovered that EET- capable microbes can extend this ability to use electrodes as electron acceptors and donors. As understanding of EET pathways expanded, so did the potential to use these electroactive bacteria (EAB) for biotechnology. A common technology that employs EAB is a process called microbial electrosynthesis (MES). In MES, EAB are used as biocatalysts to convert chemical energy into electrical energy, and vice versa. With increased interest in using renewable sources of energy such as wind and solar, being able to transform the electrical energy produced to chemical energy will be invaluable. Additionally, many MES systems aim to use autotrophs as the biocatalysts to capture and store atmospheric carbon dioxide (CO2)2–4. This diverse field has leveraged native and engineered microbes for bioremediation, carbon capture, biofuel production, and more. This work will discuss the energetic considerations for using a MES platform for the carbon-neutral generation of industrially relevant chemical products by the bacterium Shewanella oneidensis MR-1. This strain contains a complex and diverse network of EET proteins, so we will also look at the role of key players during electron transport to generate NADH. Additionally, we will discuss a synthetic biology pipeline for enacting difficult genetic modifications. Lastly, we will demonstrate how this 3 bioengineering pipeline can be applied to S. oneidensis for future work towards integrating this system with carbon capture (Figure 1.1). Figure 1.1 Goal of MES in S. oneidensis. An overarching goal of the TerAvest research group is to engineer a strain of S. oneidensis MR-1 that can be used in MES to produce industrially relevant chemical products. This organism is naturally capable of using a cathode as an electron donor. Therefore, research aims for this project include: (1) Understanding and optimizing electron uptake, (2) Introducing exogenous pathways for bioproduction, (3) Installing a CO2 fixation pathway in this heterotroph. Figure adapted from Tefft and TerAvest 20195. 1.3 Technology Meets Microbiology While the term ‘microbial electrosynthesis’ was first coined in 2010 by Nevin et al.3, the basis for this technology goes back over a hundred years6. During that time, many research developments employed bioelectrochemical systems (BES) as microbial fuel cells (MFCs). In an MFC, microorganisms are used as biocatalysts to convert chemical energy into electricity on an electrode7–9. This system offers an alternative means for energy generation from a wide range of sources outside of fossil fuels, including undesirable feedstocks like wastewater10,11. In principle, as bacteria liberate electrons from carbon substrates during metabolism, they generate reducing 4 equivalents (NADH, NADPH) or reduced cytochromes, which in turn reduce the soluble mediators that shuttle the electrons to the anode, while O2 and H+ react at the cathode to form H2O12,13. Beyond this core structure, the design of MFCs can vary greatly based on electrode material, vessel dimensions, bacterial species or consortia, and feedstock14. Often, early iterations of MFCs would rely on redox-active mediators to facilitate electron movement between the cells and the electron accepting electrode (anode)15,16. This was because the mechanisms of electron transfer by bacteria were poorly understood. Commonly used mediators include neutral red, methylene blue, and anthraquinone-2,6-disulfonate (AQDS). Eventually, interest grew in using this process in reverse, using electricity to power biochemical reactions. To do this, a BES is used as a microbial electrosynthesis cell (MEC), where bacteria on the cathode take up electrons to create reducing power such as NADH that can be used to drive a specific chemical reaction (Figure 1.2). This approach was enticing as a potential means of storing the electricity generated from renewable sources like wind and solar, a sort of biochemical battery. Additionally, depending on the physiological capabilities of the microbes, MECs are used to produce other useful chemicals, including acetic acid, ethanol, and even pharmaceuticals2,17–20. Since then, this process known as microbial electrosynthesis (MES) has undergone significant expansion, driven by researchers' efforts to improve the design and efficiency of MECs. This work encompasses three primary developmental approaches: material and structural design of BES, broadening the range of potential products, and characterizing and improving the electron transport pathways of the microbes on the cathode21. 5 Figure 1.2 S. oneidensis on a Cathode in a BES. In a 2-chamber MEC, S. oneidensis cells are inoculated into the working chamber with the cathode, while the anode is in the counter chamber, separated by an ion exchange membrane. In the counter chamber, water is split into O2 and H+ by energy provided by the anode. The working chamber, often sparged with inert gases such as N2 to prevent O2 intrusion, has cells in suspension and attached to the cathode. S. oneidensis will take up electrons either through direct contact with the cathode, or via redox-reversible mediators (inset). In the depicted design, the anode is a carbon rod, the cathode is a carbon felt sheet attached to a Pt wire, and the reference electrode, which acts as a reference point to maintain a constant potential in the MEC, is a silver wire saturated by KCl (Ag/AgCl). Though many early MFC designs had a limited understanding of how bacteria were capable of electron exchange with electrodes, the innate link between EET and anaerobic respiration was ultimately uncovered. In parallel with MFC research, it was discovered that there are bacteria capable of anaerobic respiration using extracellular materials such as U(VI), Mn(IV) and Fe(III)22–26. Microbes capable of this extracellular electron transport (EET) are referred to as dissimilatory metal reducing bacteria (DMRB), as they do not assimilate the reduced metals into cell components or protein structures. As the physiological mechanisms behind EET were elucidated, it was shown 6 that DMRB have the capacity to use an anode as a terminal electron acceptor and produce electrical energy27,28. Two of the first well-studied genera capable of this process were Geobacter (first described as strain GS-15) and Shewanella (previously Alteromonas)22,25,28–30. While there has since been found a wide variety of electroactive bacteria (EAB) capable of directly or indirectly interacting with electrodes (Desulfovibrio, Sporomusa, Clostridia, Pseudomonads, etc.), a vast majority of our understanding of the physiology of EET comes from these two foundational organisms9,31,32. Additionally, as a γ-proteobacterium, S. oneidensis is genetically tractable. Due to this, much work has utilized this organism as the basis for work on electrode-driven bioproduction. Since its inception, there has been interest in exploring the potential for combining autotrophy with EET for MES. Organisms with ‘electroautotropic’ metabolism could be fed carbon dioxide (CO2) and energy from electrodes as the sole inputs for bioproduction systems. This approach is the end goal for many MES system designs; consumption of CO2 that would otherwise be emitted while generating high-value chemical products using energy derived from renewable sources. In this dissertation, both EET and carbon fixation will be addressed, along with bioengineering strategies that have been developed to improve these systems in S. oneidensis for application in MES. First, we will examine the current understanding of EET in S. oneidensis, including the major components, how they are regulated, and how researchers have improved upon the native pathway. 1.4 EET Pathways As the ability to generate electricity by bacteria is a byproduct of native bio- reduction (respiration) and bio-oxidation pathways, there is not a single consensus 7 mechanism for performing reversible EET among EAB. Although the EET systems in S. oneidensis and Geobacter sp. are the most comprehensively studied, research has revealed a diverse field of microorganisms capable of EET. For instance, over a dozen genera have been identified to be capable of the bio-reduction of aqueous U(VI) to the insoluble form U(IV), a process which could be employed for removal of the metal from contaminated soils33. While there is overlap in general strategies and shared homologs of different pathways, the diversity in this ability allows for a deeper understanding of the reasons behind their evolution and the mechanisms involved. A significant portion of our knowledge of microbe-electrode interactions stems from research done under anodic potentials, where the electrode functions as an electron acceptor. Due to interest in using these organisms for MES, more work is being done towards understanding these pathways under cathodic, electron-uptake conditions5,34–40. In general, S. oneidensis employs two primary mechanisms of EET that have been differentiated: direct and indirect9,16,31,41. Direct EET involves outer membrane cytochrome (OMC) proteins on the cell surface directly interacting with an electrode to exchange electrons, whereas indirect utilizes extracellular mediators. While endogenous excreted flavins are the dominant mediators in S. oneidensis cultures, artificial electron shuttles like methyl viologen, neutral red, and methylene blue have been used to enhance EET16. S. oneidensis grows as biofilms on the electrode surface, allowing for a combination of direct and indirect EET through its conductive exopolymeric substances matrix (EPS)42–45. Additionally, the role of ‘nanowires’ in EET has garnered interest because it allows cells to deposit electrons over long distances. To optimize EET, understanding flavin chemistry, biofilm formation, nanowires, and 8 flexibility of proteome and regulatory pathways is essential (Figure 1.3). Figure 1.3 Considerations for Improving EET in S. oneidensis. When examining potential avenues for improving EET in S. oneidensis, four major areas of research are: streamlining the cytochrome network, clarifying the contribution of flavins, improving biofilm formation and conductivity, and understanding electron transport along nanowires. The primarily discussed electron pathway in S. oneidensis is electrons being passed from quinol-linked CymA to MtrCAB (+ OmcA, or the less efficient homologous MtrDEF) via periplasmic electron carriers (FccA, CctA). However, S. oneidensis has many more cytochromes, and recent work has suggested alterative pathways between the electrode and cell, within the periplasm, and quinol-linked proteins besides CymA34,46–49. In S. oneidensis, flavin adenosine dinucleotide (FAD) is created in the cytoplasm and transported into the periplasm via Bfe to be cleaved by UshA to form flavin mononucleotide (FMN) and adenosine monophosphate (AMP). FMN is exported through an unknown transporter, where it will act as cofactor for outer membrane cytochromes (OMCs) or abiotically convert to riboflavin (RF) to associate with OMCs or freely shuttle electrons50–52. While flavins have been known as important for EET for years, recent data have suggested they contribute more as OMC cofactors for OmcA and MtrC than as free flavins53–55. These flavins are also found within the EPS of electrode-attached biofilms42. Similarly, it has also been shown that the EPS matrix of S. oneidensis biofilms is rich in redox-active cytochrome proteins56. This creates a conductive environment that facilitates direct or mediated electron transfer (DET, MET). Over longer distances, cells will build so-called nanowires to transport electrons. In S. oneidensis, these are chains of outer membrane vesicles (OMVs) that are rich in cytochromes like MtrCAB57,58. 9 1.4.1 Cytochrome-rich pathway S. oneidensis contains over 40 cytochrome rich proteins. Approximately 80% of these proteins are localized to the outer membrane and can potentially aid in EET; the most well studied EET cytochromes are those in the Mtr pathway16,46,59,60. During anaerobic respiration, electrons enter the quinol pool, where they can be passed from the reduced quinols to the inner-membrane bound electron hub CymA49,61–65. From here, the electrons can then be passed directly to periplasmic terminal reductases (FccA, NapAB, SirA, NrfA, TorA, etc.) or shuttling cytochrome proteins (FccA, CctA)66–68. These shuttling proteins will chauffeur the electrons from the inner membrane to the Mtr complex in the outer membrane. Mtr is a 3-protein complex that spans the outer membrane, including a beta-barrel porin (MtrB) and two heme-rich cytochromes (MtrA, MtrC), and is often found associating with the surface attached protein OmcA69–73. Together these OMCs form an EET wire, with OmcA situated to the outside of the cell and MtrC reaching up to ~90 Å above the surface of the cell74. MtrC has a trifurcated heme structure, optimizing the distribution of electrons to potential electron acceptors. S. oneidensis also excretes the redox reversible flavin compounds riboflavin (Rf) and flavin mononucleotide (FMN) which shuttle electrons from the cell surface to electron acceptors during indirect EET50,52. Additionally, OmcA and MtrC associate with Rf and FMN to form flavocytochrome complexes, aiding in direct EET53–55,75,76. One important characteristic of the EET cytochrome network in S. oneidensis is its redundancy and versatility77,78. Though it would be intuitive that the cell uses the same pathway for reduction of similar compounds (e.g., Fe(II)-oxide and ferric citrate), this is not the case. In addition to the cytochrome proteins already mentioned, S. 10 oneidensis also contains two homologs of MtrCAB (MtrDEF, DmsEFABGH) which can complement the primary complex to varying degrees of success 47,48. Loss of either major periplasmic electron carrier, FccA or CctA, shows no phenotype on an anode and each is likely able to compensate for loss of the other68,79,80. CymA is essential for reduction of Fe(III), nitrate, nitrite, fumarate, DMSO, arsenate (V), and manganese (IV), but not for trimethylamine N-oxide (TMAO), sulfite, or thiosulfate63. There is also increasing evidence that the inward EET pathway diverges from the outward EET pathway34,39. S. oneidensis also expresses two hydrogenase complexes, which could potentially facilitate hydrogen-mediated EET. The flexibility of the Shewanella EET system has also been demonstrated and optimized through informed genetic manipulations. Delgado et al. showed that replacing several redox active periplasmic protein encoding genes (nrfA, ccpA, napA, napB) with cctA, thereby simplifying the network, led to a ~1.7-fold increase in Fe (III) reduction, and ~1.5-fold increase in current after 24 h80. More recently, Sun et al. showed that it is the specific tailoring of the network, and not merely excess cytochromes, that aids in EET; overexpression of cytochrome c maturation machinery (ccm) was slightly inhibitory compared to WT. However, they similarly saw improvements through the deletion of fccA, napB, and tsdB, and over expression of cctA66. This work highlights a promising avenue of research for improving EET by optimizing the native proteome. The reversibility of the MtrCAB pathway in S. oneidensis enables it to generate reducing power and drive formation of ATP5,81,82. Interestingly, recent work by Rowe et al. showed that deletion of five previously uncharacterized genes (SO0841, SO0181, SO0400, SO3660, SO3662) caused a significant defect in electron uptake from a 11 cathode34. These genes were predicted to encode proteins involved in cell signaling, regulation of a putative oxidoreductase, a quinol-monooxygenase, a transcriptional regulator, and an inner membrane ferredoxin, respectively. Of these, all but SO0841 showed a defect solely in inward EET, with no defect in outward transfer. This highlights that despite all the understanding of EET in S. oneidensis, there is still much to learn about this complex cytochrome network. One aspect that further obscures our study of EET proteins is their association with the redox active molecules Rf, FMN, and FAD. 1.4.2 Flavins Some iron oxide-reducing organisms were capable of doing so at a distance, and this was first demonstrated in S. oneidensis cells by Lies et al83. However, at the time it was unknown by what mechanism this was occurring, and researchers inferred that it was likely due to soluble redox-active molecules, specifically within a biofilm matrix. These redox active molecules were identified as flavins by Marsili et al. in 2008. Since then, there has been extensive work done to understand the role these redox molecules play in EET50,84. Researchers noted that unlike in experiments previously done in Geobacter spp., replacement of BES medium with fresh electrolyte decreased oxidation currents by ~73%. To determine if this reduction was due to soluble redox compounds, spent medium was filter sterilized and returned to the BES, restoring ~95% of anodic current. This redox compound was identified as Rf. This work was quickly followed by work describing the flavin profile of Rf, FMN, and FAD in S. oneidensis cells and supernatant, as well as their impact on extracellular ET51,72,84. Together, this tells us that FAD is manufactured in the cytoplasm, exported to the periplasm via a flavin transporter (Bfe) before being cleaved by UshA to FMN and AMP51,85. FMN is actively secreted 12 outside the cell and is abiotically transformed into Rf, while FAD remains inside the cell, acting as an essential cofactor for enzymes like fumarate reductase (FccA). The level of flavin secretion is closely associated with the redox state of the environment, as well as availability of carbon substrate and electron acceptor species86,87. Following these discoveries, researchers sought to understand the mechanism behind flavins’ role in ET. All three species of flavins mentioned here are freely redox reversible, so the prevailing theory for many years was that flavins acted as mediators, shuttling electrons between the cell and electron acceptor across long distances50,51,88– 90 . This was supported by data such as that described in Marsili et al. 2008, that when medium containing soluble flavins was removed, the current decreased, and recovered upon their return. Similarly, deletion of the flavin transporter (∆bfe) resulted in a ~75% decrease in anodic current versus WT S. oneidensis. However, there is increasing evidence suggesting that a more prominent role for flavins during EET is as bound cofactors for outer membrane cytochromes (OMC)38,39,53,54,56,75,76,88,91–93. Much of the work done to determine the role of specific compounds or proteins on EET has been interpretation of current generated on an anode; very often these working electrodes are also made of carbon materials. This design ignores a variety of confounding variables that can greatly influence data interpretation. One such example is abiotic interactions with the electrode by soluble molecules. Notably, flavins adsorb onto carbon electrode surfaces53,56. This leads to conflicting results between work done with carbon electrodes versus electrodes made from indium tin oxide, gold, or other materials. Additionally, because many conclusions are drawn from electrochemical data, it can be difficult to separate abiotic and biotic effects without thorough controls. To this 13 end, researchers have sought to resolve the binding relationship between OMCs (MtrC, OmcA) and flavins53,75,93,94. FMN and Rf associates with MtrC and OmcA, respectively, and these associations can either be transient or stable. A factor influencing this bond is the formation of a disulfide bond in the presence of oxygen, preventing flavin binding to MtrC75,93. Therefore, affinity for flavins to bind MtrC is tightly controlled by the redox state of the protein, flavins, and environment. To this end, later work parsed apart the influence of free versus bound flavins to ET. By using a carbon felt electrode, researchers were able to, for the first time, demonstrate the presence of free flavins, flavocytochromes, and OMCs in a single system. Because of this, they could demonstrate that while secreted flavins show a strong electrochemical response, they do not significantly contribute to the biotic current. Instead, it is cytochromes and flavocytochromes that contribute to a majority of the ET53. In experiments similar to those that first identified soluble mediators as important to ET, it was shown that in addition to free flavins, released flavin-bound OMCs are found in the bulk medium and capable of electron shuttling95. Importantly, while this body of work demonstrates the overestimation of the role of free flavins in the system, it does not negate the fact that flavins are vital for ET; they are simply more important as OMC-bound cofactors. Flavins also stimulate increased biofilm formation38,42,91,96,97. While the physiological mechanism has not been determined, evidence suggests that as the concentration of extracellular Rf increases (>15 nM), it initiates a regulatory shift affecting biofilm-related genes42. Biofilms act as an electroactive network, enabling increased EET using redox active molecules trapped within the EPS matrix92. Notably, quantification of flavins trapped within biofilms show ~10-fold increase in concentration 14 compared to the bulk medium. The amount of Rf in the biofilm (2.21 × 1015 molecules) falls within the range of potential OMC binding sites (3.49 × 1014 - 1.05 × 1016). Taken together, it can be inferred that even if a flavin is not tightly bound to an OMC, it can act as an electron shuttle to a much smaller degree, remaining within the biofilm rather than diffusing over long distances. Work characterizing the role of flavins during EET to an acceptor behind a barrier (cells cannot attach) supported this, showing that a loss of MtrC/OmcA cannot be complemented by exogenous flavin addition. This work reiterated that flavins can be recycled for shuttling but rely on OMCs and operate over short distances98. Keeping this in mind, we will next look at the importance of biofilm formation during EET. 1.4.3 Biofilms In natural environments, it is crucial for bacteria to form biofilms for protection, resource sharing and access, and horizontal gene transfer99. S. oneidensis forms biofilms on mineral surfaces, such as Fe-oxide, to promote anaerobic respiration using the insoluble minerals100. A biofilm consists of whole cells trapped within a matrix of EPS, DNA, proteins, and other secreted molecules. Cells can shift between motile and attached lifestyles via regulatory changes triggered by environmental stimuli. While motile, cells chemotactically sense their environment, and when encountering a favorable environment, such as Fe-oxide, adhere to the surface and initiate biofilm formation101. When grown in a BES using an anode as an electron acceptor, cells similarly form biofilms on the electrode surface. Cell attachment and subsequent biofilms are vital to the efficiency of EET between the cell and electrode102. Therefore, 15 much work has been done to understand both the structure and composition of the EPS, as well as the regulatory mechanisms that control them. Under dynamic conditions with flow, S. oneidensis cells develop biofilms, while in static aerobic cultures, they form surface pellicles103. Formation is stimulated by the presence of oxygen, and dispersal occurs upon oxygen depletion104,105. Genes identified as vital to biofilm formation include agglutination proteins (AggA), extracellular adhesion proteins (BpfA), pili proteins (MshA/PilD, Mxd operon), and the transformation and export systems for them106–109. Additionally, as with many microorganisms, the regulation of the small molecule messenger c-di-GMP impacts biofilm formation99,110,111. While EPS composition varies from organism to organism, S. oneidensis EPS is dominated by proteins, but also contains extracellular DNA (eDNA)103,112–114. The biofilm also transforms over time, resulting in changes in composition, thickness, conductivity, shape, and stability96,111–113,115,116. During growth on an anode, more current is generated when more cells are attached to the electrode. In nutrient rich, aerobic environments, S. oneidensis biofilms can grow relatively thick (100+ μM) and form 3-dimensional, mushroom like structures106,117,118. On electrodes in anoxic conditions however, the cells tend to form much thinner biofilm, only monolayers in many cases. This behavior is explained by the need for cells to be in physical contact with the electron acceptor to best survive. Attempts to increase EET by biofilm modulation have seen the most success by increasing the area of attachment and conductivity of the EPS matrix102,109,116,119–121. This increase was demonstrated as biofilms grown on electrodes at 0.0 VSHE or aerobically with no current applied will form the mushroom structures, while those grown 16 at 0.4 VSHE were flat and even, covering more of the electrode118. Additionally, even in thicker, electrode attached biofilms, a vast majority of the current (95%) was attributable to the tightly attached bottom layer of cells56. A unique example of this was reported by Zhao et al., who showed that overexpression of SulA, which inhibits FtsZ ring formation during cell division, resulted in biofilms with larger surface area coverage and EET. These increases were attributed to the morphological changes that resulted in bigger and elongated cells, therefore increasing the area covered per cell91. Another approach focuses instead on optimizing electrode material. Sanchez et al. demonstrated that by using carbon nanofiber (CNF) electrodes over carbon microfiber (CMF), there was increased surface area, creating more cell attachment locations, leading to thicker biofilms and more current generation119. Regulation of biofilm formation also impacts the structure and conductivity. Internally, many of the regulatory systems for attachment are controlled by the small messenger c-di-GMP. This signaling molecule can act as both a transcription and post- translational controller in a majority of well-studied bacteria99,111,122. It is formed by diguanylate cyclases and degraded by phosphodiesterases; high concentrations increase biofilm formation, and low concentrations signal for detachment123,124. Attempts to alter the internal concentration, through expression or deletion of diguanylate cyclases or phosphodiesterases, have shown that more c-di-GMP increases biofilms and current110,111. Interestingly, Ng et al. also showed that c-di-GMP increases expression of the Mtr pathway, resulting in thicker and more conductive biofilms99. This can also be achieved through external signaling cues. In the work showing biofilm formation at different anodic potentials, those grown at 0.4 VSHE had a lower EPS 17 content but higher concentrations of electroactive components118. Regulation of exonucleases is also important, as there is a required balance between having enough to degrade eDNA that could impact conductivity in older biofilms, but not so much as to abolish its role in attachment for young ones112. Another interesting demonstration of regulatory impact is the gene bolA. Overexpression of this transcriptional regulator increases biofilm and current generation, but a deletion of bolA did not have a phenotype compared to WT. The authors suggest that bolA is only expressed under harsh growth conditions, aiding in attachment under dire environmental situations116. The electroactive components of biofilms consist of flavins, cytochromes, and flavocytochromes. Work by Edel et al. showed that there was roughly ~10-fold more flavin compounds trapped within the biofilm matrix than in the effluent42. As previously mentioned, the amounts of flavins measured falls within the feasible concentration range of cytochrome proteins. Edel et al. suggest that this could corroborate the assertion that even in biofilms, flavins are bound to OMCs, facilitating ‘electron hopping’ between the redox centers. Whether the cells are using direct or mediated transfer is also significantly affected by regulation. Investigations into biofilm conductivity over time by Choi et al. suggest that younger biofilms mainly use direct EET, while the role of flavin mediated transfer increases as the biofilm ages and increases in thickness115. When considering regulatory impact on EET processes, a growing area of interest is the role of ‘nanowires’, both in their formation and their electroactive composition. 1.4.4 Nanowires Geobacter spp. are well known for their long electroactive pili structures capable of transporting electrons over long distances. These structures, named nanowires, were 18 believed to also be formed by S. oneidensis cells for EET. S. oneidensis can form type IV pili structures with high homology to those of Geobacter spp125. However, early evidence demonstrated that while S. oneidensis forms ‘nanowires’ capable of EET, these are not pili, and therefore not nanowires126,127. It was later demonstrated that outer membrane extensions (OMEs) form the observed conductive filaments128. This has led to some contention regarding nomenclature, with other researchers instead suggesting the names ‘nanopods’ or ‘nanocables’129,130; for consistency with literature, they will be referred to here as nanowires. Pirbadian et al. provided the first evidence that the nanowires are OMEs, made of chains of outer membrane vesicles (OMVs), containing periplasm (but not cytoplasm) and do not involve pili131. The nanowires form under anaerobic conditions and production is stimulated by surface attachment, regardless of substrate availability, surface type, and mixing conditions132,133. The conductive nature of the extensions is attributed to OMCs located along the cell surface, including OmcA and MtrC57,134. During nanowire formation, expression of Mtr pathway cytochromes, and a variety of periplasmic cytochrome proteins are greatly up regulated, highlighting the correlation between cell surface area and cytochrome usage133. While inclusion of OMCs is widely accepted as the physiological structure of nanowires, the mechanism behind the EET properties is more elusive. The prevailing theory supported by both the composition of the nanowires and energetics calculations, is that they facilitate ‘electron hopping’. Electrons are passed between the redox active centers of cytochromes embedded in the OM, with a calculated conductivity akin to artificial silicon nanowires (0.01-1 S/cm)129,130,134. 19 As much of the work on S. oneidensis nanowires is comparatively recent, OME contributions to EET, compared to flavin mediated and non-OME surface cytochromes, have not been thoroughly investigated. Some considerations when determining the relative importance should include the apparent discrepancies between these two established mechanisms. On one side, long distance EET has been attributed to secreted flavin molecules, both due to the distance traversed, and the ability to reduce compounds behind a physical barrier50–52,72,83,98. Conversely, evidence suggesting that a majority of flavins are bound to cytochromes as cofactors implies that they may not be as effective at long distance EET as previously thought39,42,53,56. Nanowires may fill this gap as they use cytochromes and flavocytochromes to transfer electrons and have been shown to reach up to 9 µm in length131. Additionally, if OMVs become detached from the larger nanowire structure, they could facilitate electron shuttling over distances similar to those proposed by free flavins. Work by Liu et al. potentially demonstrates this process, showing that large exported organic components (EOCs), meaning ‘excreted’ cytochromes and not flavins, are responsible for more than half (~56%) of extracellular reduction of Cr(VI)95. These components could potentially represent OMVs that were formed from nanowires. As this system diverges so greatly from our understanding of Geobacter spp. nanowires, it will be imperative to understand the contribution of S. oneidensis nanowires to EET. 1.5 Bioengineering Techniques The fields of synthetic biology and biotechnology are inextricably linked as researchers aim to harness biological tools to tackle challenges such as climate change. Synthetic biology incorporates tools from engineering, computer science, and 20 chemistry to redesign biological processes. S. oneidensis, as a model electroactive organism,135 has garnered significant interest as researchers aim to develop tools for its use during growth with electrodes. Being a γ-proteobacterium and closely related to Escherichia coli, S. oneidensis offers the advantage of genetic manipulability and the utilization of tools developed for E. coli. In this section, we will discuss different bioengineering approaches ranging from small genetic modifications like single nucleotide point mutations (SNP) to entire libraries of novel DNA constructs. Specifically, we will look at bioengineering approaches aimed at increasing EET between cell and electrode through optimization of native processes or genetic toolkit development. 1.5.1 Native Optimization As discussed, S. oneidensis contains many genes that play a role in EET. These encode conductive cytochromes, extracellular components, and metabolic enzymes, many of which have multiple homologous copies within the genome (e.g., MtrCAB & MtrDEF). We have already discussed a variety of approaches researchers have used to improve EET, including simplifying the cytochrome network, increasing biofilm formation or flavin excretion, and modulating the metabolome66,91,108,110,111,116,136–138. Other approaches have focused on increasing NADH concentration through enhanced synthesis pathways or substrate utilization. In this way, more electrons (in the form of more NADH) will lead to an increase in reduced quinols and increased electron export. Li et al. aimed to increase the pool of NAD+/NADH through expression of five non-native genes aimed at increasing import and synthesis of NADH precursors139. This modification led to a 4.4-fold increase in power output compared to WT cells. Work by Ding et al. aimed to increase NADH concentrations by increased expression of native 21 proteins found in central carbon metabolism140. Their targets include lactate importers, NADH-generating steps (GapA, Mdh), and NADH dehydrogenases (Ndh), leading to a 62% increase in power density when all four are overexpressed. A recent study implemented genetic alterations from a variety of research groups that have demonstrated various levels of improved EET141. A combination of gene knockouts (∆SO3171108, ∆exeS114, ∆SO194299,142, ∆SO3491), expression of Geobacter OMCs and B. subtilis flavin synthesis genes38, and use of an ‘artificial biofilm electrode’ increased the power density to ~39-fold more than WT S. oneidensis. This study illustrates that by combining various engineering techniques, including the utilization of established approaches, it is possible to achieve significantly enhanced output compared to using any of these techniques individually. 1.5.2 Tool Development While many tools that were first developed in E. coli can be adapted to S. oneidensis, the development of these systems to optimally operate in S. oneidensis can save time and effort that would otherwise be used for troubleshooting. Researchers have addressed this goal through thoughtful plasmid design and employing a wide variety of CRISPR-based editing systems. When constructing plasmids for expression, or overexpression, of different genes, considerations include induction system, copy number, plasmid compatibility, and more. Two commonly used induction systems include isopropyl ß-D-1-thiogalactopyranoside (IPTG )induction using the lac promoter, or arabinose induction. However, there is an interest in expanding these options as these systems can be poorly repressed (i.e., ‘leaky’) or have off-target metabolic effects. This effort has included leveraging native TMAO respiration enzymes for TMAO-based 22 induction and adapting a rhamnose-inducible system from E. coli143,144. While the TMAO based approach has the benefit of using native proteins, it is inefficient because it is leaky and inducer concentration dwindles with time. Conversely, the rhamnose-inducible system is beneficial as rhamnose is non-toxic and cannot be metabolized in S. oneidensis. However, many researchers prefer constitutive systems to prevent off target effects of inducers, and potentially poorly controlled expression. Yi et al. addressed this concern through introduction of T7 RNA polymerase into S. oneidensis, which enables the use of the widely-used constitutive T7 promoter system145. Meanwhile, Cao et al. comprehensively tested expression and control of a library of vectors, including 9 promoters of different strengths, 4 origins of replication, 2 shuttle genes for genetic transfer, and various antibiotic selection markers146. This knowledge allows researchers to choose which plasmid characteristics they need. Understanding of vector design is crucial when designing more complex systems, such as those using CRISPR. Many strategies for increasing EET involve gene deletions. Sucrose counter selection is a commonly used technique but can take weeks to result in a single knockout due to time needed for plasmid construction, conjugation, and screening147. Depending on the gene target, thousands of colonies may need to be screened to generate the desired mutant. CRISPR-based systems have been engineered to simplify screening, vector construction, and range of potential modifications. Corts et al. developed a clever design in which a Cas9 enzyme targets un-modified genomes, therefore killing any WT revertants and ensuring that only the desired mutant can survive148. An alternative to gene deletion can be single base conversions to generate premature stop codons, thereby preventing translation149,150. In 2022, Chen et al. 23 published two research papers that expanded the coverage of the editable genome (from 89% to 100%) by increasing recognized protospacer adjacent motif (PAM) sites151,152. This expansion is important because Cas9 enzymes require PAM sites (e.g. 5’-NGG-3’) to associate with the genome, and this development allows binding in AT- rich regions that were previously unsuitable. They also engineered a vector to enable simultaneous activation (CRISPRa) and inactivation (CRISPRi) of two different genes. The use of CRISPRi, where a sgRNA binds to a gene target and associates with an inactive Cas9 to block transcription, is also desirable as it can allow for inducible repression of genes as opposed to outright deletion. Such systems have been successfully developed in both S. oneidensis and E. coli153,154. Ford et al. adapted the E. coli-based CRISPRi system in conjunction with in silico metabolic modeling to generate a difficult knockout in a gene previously considered essential155. This result demonstrates the flexibility with which these different genetic tools can be operated. Many of these tools are developed in the context of increasing EET between S. oneidensis cells and the electrode. It is important to remember that the greater goal behind this is to improve MES systems to produce biofuels, bioplastics, and more. 1.6 MES for Bioproduction Development of effective, scalable MES platforms requires an intimate understanding of the biological processes involved. There are a variety of tactics researchers have used to approach this goal, including using mixed-cultures, engineering electron uptake in organisms that naturally generate valuable products, and engineering production pathways in electroactive organisms156,157. While each approach 24 has merit, we believe that the approach of engineering bioproduction in electroactive bacteria, and specifically S. oneidensis, is the best path forward158. A key challenge when using microbial communities for MES is the lack of specificity in product generation. Mixed-cultures typically rely on methanogens and acetogens as the bio-producers. This requirement presents a problem because though these designs use CO2 as the initial feedstock, poor electron uptake and subsequent energy loss as methane make them ineffective. Any attempts to improve these systems requires engineering of not only multiple, potentially uncharacterized, organisms, but of the interactions between organisms in the community. This requires additional time and effort to troubleshoot complex community systems. For these reasons, we find the flexible and rapid ability to adapt pure cultures to be a better alternative. Engineering EET pathways in pure cultures of organisms already capable of producing a specific product can face equally complex problems. Namely, understanding the intricacies of EET in their native host is already challenging. Deciding which pathway to use, how it will be regulated, and its impact on native processes are just some of the considerations needed. For example, researchers have implemented the MtrCAB pathway in the model organism E. coli, which has the benefits of being well characterized and genetically tractable89,159,160. Though this strain could achieve extracellular EET, it was slow, reducing extracellular iron at 2% the rate of S. oneidensis89. While other organisms have been modified to interact with electrodes, none are as efficient as native systems, and the only chemical produced in many instances is acetic acid. As highlighted in Prèvoteau et al., to make these designs economically feasible there cannot be ‘wasted electrons’ in the form of off-target 25 products or poor EET rates2. While it is tempting to use model organisms like E. coli, the growing field for bioengineering in S. oneidensis, as detailed above, demonstrates a diminishing disparity in terms of genetic manipulation. To this end, there has been an increase in publications illustrating S. oneidensis’s use for bioproduction and even bioremediation161. Flynn et al. showed electrode-dependent conversion of glycerol to ethanol at 82% efficiency162,163. Other work has shown production of n-butanol and methane, two other useful fuels164,165. More complex engineering has capitalized on the intricate redox network within S. oneidensis to overproduce the heme precursor and cancer drug 5-aminolevulinic acid (ALA) and recycle electron shuttles for redox-based polymerization166,167. This wide range of research exemplifies the diversity of potential products possible when developing bioproduction, rather than optimizing non- electroactive organisms to produce acetic acid. To this end, the work discussed in this dissertation addresses many of the questions broached above. My efforts include investigating the physiological properties of S. oneidensis during electron uptake (Chapter 2), the flexibility of the inward electron pathway (Chapter 3), a pipeline for characterizing and deleting conditionally essential genes (Chapter 4), and a look at work towards implementing a functional Calvin-Benson Cycle in S. oneidensis (Chapter 5). 1.7 Research Overview Previously published work by the TerAvest lab has demonstrated the ability for S. oneidensis to take up electrons from a cathode to drive 2,3-butanediol production from the precursor acetoin. Chapter 2 of this dissertation builds on this research by characterizing native processes. This investigation is essential because as noted above, understanding the roles and interactions of the EET network within an organism will be 26 instrumental in future work aimed at increasing product output. Specifically, I have demonstrated that proton motive force (PMF) is imperative for NADH generation; production increases with high levels of PMF and ceases when the proton gradient is collapsed 5,81. In this work, I expand on this result by ensuring that all 2,3-butanediol produced is electrode-dependent, and showing that the native oxidases can sustain PMF. This work demonstrates a feasible application for bidirectional EET, coupling unfavorable NADH production to the energetically downhill reduction of oxygen. Additionally, it details potential hurdles related to the presence of oxygen in a BES and how to overcome them. Similarly, Chapter 3 investigates the contribution of known EET pathway components in the context of MES. Often, conclusions on the contribution of different proteins, shuttles, or structures are drawn solely from electrochemical data. This can be misleading due to the abiotic interactions these redox molecules can have with electrodes. For this reason, I investigated how the loss of certain essential EET proteins impacts 2,3-butanediol generation. Using this model, I can more accurately assess how much of the current is biotic and going towards creating reducing power such as NADH during electron uptake from a cathode. From the components I examined, I determined that MtrCAB is essential for electron uptake. Interestingly, I found CymA and endogenous flavins to be important, while loss of FccA and exogenous flavins had no impact. I also demonstrated the value of the native hydrogenases, not in the context of hydrogen mediated EET, but for increased cell viability on the electrode. Together, this work both supports many established claims but also demonstrates the impracticality of using current data alone to interpret biological processes. 27 Chapter 4 discusses a pipeline that combines in silico metabolic modeling with CRISPRi based gene knockdown to quickly generate difficult knockouts155. Using OptKnock with a previously designed model for S. oneidensis central metabolism, I identified a genetic strategy for evolving a carbon dioxide fixing module within S. oneidensis168–173. This strategy relied on the deletion of the gene gpmA to sever the flow of carbon between gluconeogenesis and the TCA cycle. However, this gene has been characterized as expected essential and numerous attempts to generate a knockout were unsuccessful. Generating this mutant failed because there was not sufficient substrate to power these two modules (gluconeogenesis and TCA) separately. To address this, I used a CRISPRi knockdown system to characterize the potential phenotype of a ∆gpmA strain growing on different carbon substrates. I found that supplementing the medium with inosine and lactate restored a gpmA knockdown growth defect to WT levels. The gene deletion protocol was amended with the additional substrates, allowing for the isolation of a mutant. The rate of mutate formation during this was approximately half of all colonies screened (46%) further showing that this method worked and it was not merely chance that the mutant was made. The research in this chapter focuses on the development of this pipeline, while Chapter 5 will briefly cover the current stage of using this strain to evolve a carbon fixing pathway. Overall, this work will address the current challenges in using S. oneidensis for practical development of MES and how I are working to overcome them. I have demonstrated bidirectional EET to power unfavorable reactions, characterized the EET pathway during MES, and developed genetic engineering strategies to continuously push this field forward. 28 REFERENCES (1) Müller, V. (2009) Bacterial Fermentation. Encyclopedia of Life Sciences 1–7. (2) Prévoteau, A., Carvajal-Arroyo, J. M., Ganigué, R., and Rabaey, K. (2020, April 1) Microbial electrosynthesis from CO2: forever a promise? Curr Opin Biotechnol. Elsevier Ltd. 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TerAvest1 1 Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA 2 Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI, 48824, USA 45 2.1 Abstract Extracellular electron transfer (EET) by S. oneidensis can be used to drive intracellular biochemical reactions. Previous work used EET to generate 2,3-butanediol (2,3-BDO) via exogenous butanediol dehydrogenase reducing acetoin in an NADH dependent reaction. However, generating NADH via electron uptake from a cathode is energetically unfavorable, so NADH dehydrogenases couple the reaction to proton motive force. We therefore need to maintain the proton gradient across the membrane to sustain NADH production. This work explores accomplishing this task by bidirectional electron transfer, where electrons provided by the cathode go to both NADH formation and O2 reduction by oxidases. We show that oxidases use trace dissolved oxygen in a microaerobic bioelectrical chemical systems (BES), and the translocation of protons across the membrane during O2 reduction supports 2,3-butanediol generation. Additionally, at high levels of dissolved oxygen in an aerated system, cytotoxic reactive oxygen species form and result in cell death. 2.2 Introduction As reliance on fossil fuels becomes increasingly unsustainable from an economic and environmental perspective, researchers work toward alternative energy sources. One solution is microbial electrosynthesis, which is the microbially catalyzed transfer of electrons from an electrode to cells to drive a biochemical reduction reaction1–3. Microbial electrosynthesis can be catalyzed by electroactive bacteria capable of interfacing with an electrode surface in a BES or by using the electrode to generate electron carriers, such as H2 or formate, that can be taken up by bacteria4–6. When using electroactive bacteria, a potential is applied to the system to drive oxidation of an 46 electron donor (typically H2O) at the anode, and the electrons liberated are taken up by bacteria at the cathode surface. These electrons are used for reduction of a feedstock, such as CO2, to the desired product. When CO2 is the reactant, microbial electrosynthesis becomes a carbon sink, acting as a carbon-neutral platform to produce biofuels, bioplastics, or specialty chemicals7,8. Because CO2 is the ideal feedstock for microbial electrosynthesis, much of the existing research focused on bacterial strains or communities with the capacity for autotrophic growth. However, these microbes are inefficient due to slow growth rates and poor interaction with electrodes9–11. Recent advances in engineering autotrophy raise the possibility of expanding the applications of microbial electrosynthesis beyond the need for native autotrophy or mixed microbial populations12–15. Therefore, we have focused on a well-understood electroactive bacterial chassis, Shewanella oneidensis MR-1, and optimizing product generation16,17. S. oneidensis MR-1 is a metal-reducing bacterium with a well-characterized extracellular electron transfer pathway using MtrCAB16–20. The Mtr pathway allows S. oneidensis to respire anaerobically using extracellular, insoluble electron acceptors such as Fe(III) oxides, Mn(IV) oxides, and electrodes21. As with aerobic respiration, electrons are passed into the quinol pool (menaquinol and ubiquinol) by dehydrogenases in the electron transport chain that oxidizes metabolites such as lactate, formate, and NADH. For extracellular electron transfer, the reduced quinones (quinols) are oxidized by the inner membrane bound cytochrome CymA22–25. This protein acts as an electron hub, depositing electrons onto periplasmic electron carriers, such as fumarate reductase (FccA) and a small tetraheme cytochrome protein (CctA), to be shuttled onto terminal oxidoreductases. During respiration with an extracellular 47 electron acceptor such as an anode, electrons are transferred to the Mtr pathway, a three-protein complex that spans the outer membrane and extends into the extracellular space26,27. Figure 2.1 Overview of Bidirectional Electron Transfer to 2,3-BDO. S. oneidensis cells are incubated on a cathode poised at -500 mVAg/AgCl. Electrons are taken up by the cell via outer membrane MtrCAB pathway and passed to the inner membrane quinols via various periplasmic electron carriers (FccA, CctA) and CymA. Electrons from the quinol pool are used to reduce NAD+ to NADH, catalyzed by NADH dehydrogenases (NDHs) coupling the reaction to proton movement across the inner membrane. The electrode produced NADH is used to reduce exogenous acetoin to 2,3-BDO via butanediol dehydrogenase (BDH). Importantly, the Mtr pathway is reversible, allowing inward electron transfer from a cathode into the cell2,26,28–32. Electrons that are taken up via the Mtr pathway reduce respiratory quinones, and the cell can use the quinols as the electron donor for NAD+ reduction by reversing NADH dehydrogenases. NADH can be used to drive a wide 48 variety of intracellular reduction reactions. As a proof-of-concept, we previously demonstrated that reducing power from the electrode can be used to drive the NADH- dependent reduction of acetoin to 2,3-BDO via the heterologous enzyme butanediol dehydrogenase (Bdh) (Figure 2.1). By measuring the accumulation of 2,3-BDO in the system, we can assess the rate and efficiency of electron uptake28,33. Inward electron transfer requires a continuous supply of electrons and an electron sink. In this system, acetoin is provided as the electron sink and electrons are supplied by a cathode poised at -0.5 VAg/AgCl. At this electrode potential, electron transfer from the cathode to the MtrCAB complex is thermodynamically favorable. All subsequent reactions in the pathway from electrode to menaquinol are also freely reversible. However, there is a significant energy barrier for electron transfer from menaquinol (-80 mVSHE) to form NADH (-320 mVSHE) due to the large difference in reduction potential. To overcome the energy barrier, the reaction is catalyzed by ion- coupled NADH dehydrogenases working in reverse. S. oneidensis uses both H+- coupled and Na+-coupled NADH dehydrogenases. In the forward direction, these enzymes couple NAD+ reduction to the movement of ions down the electrochemical (∆ψ) and proton or sodium gradient (∆pH or ∆[Na+]) known as proton or sodium motive force (PMF or SMF) across the inner membrane34. The movement of ions down these gradients into the cytoplasm provides the energy needed to power unfavorable chemical reactions; examples of this include formation of ATP via FOF1-ATP synthase or, as in this case, NAD+ reduction by NADH dehydrogenases. In nature, reverse NADH dehydrogenase activity is a means to prevent the potentially lethal overreduction of the quinol pool by generating NADH25,32,34–36. 49 To enable electron uptake from the electrode, S. oneidensis cells are incubated in a BES in the absence of an organic substrate or native terminal electron acceptor. Under these conditions, the cells do not generate PMF via NADH dehydrogenases (Nuo or Nqr) or succinate dehydrogenase (Sdh) because no substrate for these complexes is available34,35. Similarly, the absence of a terminal electron acceptor prevents forward electron transport chain flux. Because the reduction of NAD+ to NADH requires the free energy provided by PMF, continuous PMF regeneration is necessary for the sustained production of 2,3-BDO. This concept is supported by the prior observation that addition of CCCP (carbonyl cyanide m-chlorophenyl hydrazone), an ionophore that dissipates PMF, results in the cessation of 2,3-BDO production33. This result underscores an important consideration for microbial electrosynthesis design; how the cell will maintain PMF to continuously drive the reduction reaction forward. To maintain PMF in previous experiments, we introduced proteorhodopsin (PR). PR is a light-driven proton pump that moves protons against the proton gradient into the periplasm, sustaining PMF. Active PR and illumination resulted in an increase of both 2,3-BDO production and cathodic current28. However, relying on PR as a source of PMF is not a viable solution for scale- up due to well-known issues with light penetration in industrial photobioreactors, and the additional energy cost associated with continuous illumination28. Understanding this, we sought to utilize bidirectional electron transfer so electrons from the electrode are used for generating both NADH and PMF. In bidirectional electron transfer, electrons taken up by the cell go towards both the generation of NADH and the reduction of a terminal electron acceptor, e.g., oxygen. This coupling of electron uptake to oxygen reduction by terminal oxidases for PMF 50 generation was first described in S. oneidensis by Rowe et al.32. They found that under carbon-starvation and aerobic conditions, S. oneidensis on a cathode generated "non- growth-linked energy" in the form of PMF via terminal oxidase activity. Evidence indicated that PMF was used for production of ATP and reduced cytoplasmic electron carriers (FMNH2, NAD(P)H). However, it is still unknown whether this process could continuously generate reducing power for use in MES as there was no sink for NADH. By implementing the Bdh-based system, we sought to determine if bidirectional electron transfer could sustain PMF generation to drive 2,3-BDO generation. 2.3 Results and Discussion 2.3.1 Eliminating electrode-independent 2,3-BDO Production We previously demonstrated that the combination of an electrode and active PR led to higher levels of 2,3-BDO production than without either of these components. Before exploring the possibility of using bidirectional electron transfer instead of PR to drive PMF generation, we reexamined 2,3-BDO production in wild-type (WT) S. oneidensis MR-1 with and without active PR. Importantly, this experiment was done using an updated version of the previously described experiment protocol. In previous experiments, 35-50% of 2,3-BDO production was independent of the electrode, likely generated using NADH from organic carbon oxidation. To address the high background, we altered the protocol to promote residual organic carbon in the presence of an electron acceptor (anode) to reduce the availability of alternative sources of NADH. Briefly, in the updated protocol S. oneidensis is pre-grown in minimal medium (M5) with 20 mM lactate aerobically (or anaerobically with 40 mM fumarate as described later) for 18 hours, followed by inoculation into the BES under aerobic and anodic conditions 51 (+0.2 VAg/AgCl). After six hours, N2 sparging is started to switch the cells from using oxygen to the anode as the electron acceptor. In the current study, this anaerobic, anodic phase continued for 40 hours (versus 18 hours in the previous protocol) before the potential is switched to cathodic (-0.5 VAg/AgCl). After this modification, we observed elimination of electrode-independent butanediol production (Figure 2.2, No Potential). Figure 2.2 All 2,3-Butanediol Production is Electrode Dependent. Measurement of 2,3-butanediol in BES experiment with modified protocol. WT cells with pBDH or pBDH- PR with (holo-) or without (apo-) retinal as a cofactor, were pre-grown aerobically, washed, and inoculated into anodic BES. After 40 hours potential was switched to cathodic, and acetoin was added to a final concentration of 1 mM (T=0). Samples were collected for HPLC analysis every 24 hours. Lines and error bars represent averages and standard error (n=3). 52 We tested the amended protocol using a strain expressing PR, with and without the essential cofactor all-trans-retinal. Cells with active PR (holo-PR) produced more 2,3-BDO than those with inactive PR (apo-PR). Interestingly, cells with active PR produced approximately the same amount of 2,3-BDO as cells not expressing PR, while cells with inactive PR showed a decrease in 2,3-BDO production (Figure 2.2). This finding suggests that while PMF generation by PR supports an increase in 2,3-BDO production, the metabolic burden or membrane occupancy constraints of expressing PR outweigh the benefits. Moreover, this result suggests that there is an unaccounted-for source of PMF in the absence of PR. Another source of PMF appears more likely than the possibility that PMF is unnecessary, based on experimental evidence and thermodynamic calculations. Our prior work demonstrated that 2,3-BDO production is halted when PMF is dissipated by CCCP33. Additionally, electron transfer from quinols to form NADH cannot occur at an appreciable rate without the energy gained from proton translocation across the membrane. To sustain the NADH dehydrogenase- catalyzed reaction, which utilizes PMF, there must be a mechanism to replenish the proton gradient. We considered formate dehydrogenase and FoF1 ATP synthase as possible PMF sources, but found them unlikely due to the lack of a formate or ATP source. Therefore, we speculated that trace amounts of oxygen entering the BES could be sufficient to enable bidirectional electron transfer. 2.3.2 Bidirectional Electron Transfer to Oxygen and NAD+ We investigated the possibility of bidirectional electron transfer as the unknown source of PMF because this reaction could be powered by the electrode, and the substrate (O2) is readily accessible. Although the working chamber was continuously 53 degassed by N2 bubbling (99.999% N2), we suspected that the environment was microaerobic. The BESs may not be completely airtight due to the use of neoprene tubing and plastic connectors, and the possibility of oxygen diffusion from the anode through the ion exchange membrane (Figure S2.1). Additionally, the N2 tank used can contain up to 1 ppm O2 contamination, per the product specifications (Airgas). To ascertain if oxygen was present, we inserted an optical dissolved oxygen (DO) probe into the BES and conducted an experiment as normal. The DO in the working chamber was at ~100% saturation before inoculation, decreased to ~60% saturation upon cell addition, and dropped to ~1% upon N2 bubbling (Figure 2.3A). This single experiment produced 0.046 mM 2,3-BDO over 3 days, which is consistent with previous experiments (Figure 2.3C). Figure 2.3 Presence of Oxygen in BES. Time course of the experiment from setup (Day -3) to final timepoint (Day 4), showing the current (A) and dissolved oxygen (B). Samples collected daily from Day 0 to Day 3 to quantify 2,3-BDO production (C) (n=1). Production of one 2,3-BDO molecule from acetoin requires oxidation of one NADH, which in turn depletes four H+ from available PMF via Nuo. One molecule of oxygen (O2) allows translocation of four H+ across the membrane if it is reduced by either of the proton-pumping terminal oxidases, Cco and Cox37–39. Therefore, the reduction of one O2 molecule can sustain production of one molecule of 2,3-BDO from 54 the perspective of PMF balance. The 1% saturation DO concentration observed is equivalent to 0.073 mg/L (30°C, 856` elevation), or ~2 µM of oxygen available throughout the experiment40. Considering this concentration, there is more than enough oxygen available to support 0.046 mM 2,3-BDO production over 72 hours (0.638 µM 2,3-BDO/hour) as the sole source of PMF. To verify the contribution of terminal oxidases in generating PMF during electron uptake, we compared current and 2,3-BDO production between WT MR-1 and a strain lacking all 3 terminal oxidases (∆cyd∆cco∆cox, here named ∆oxidase)32. This strain cannot use O2 as a terminal electron acceptor meaning that even with trace amounts of oxygen, the mutant cells would not be able to use it.41 This mutant cannot grow in aerobic conditions, so further comparisons were performed using anaerobic pre-growth of all strains. To ensure consistency, we compared anaerobic growth of WT versus ∆oxidase cells in M5 minimal medium (20 mM lactate, 40 mM fumarate). We observed similar growth rates (Figure S2.2), so for MES experiments, these strains were pre- grown anaerobically. WT cells pre-grown in an anoxic environment produced 0.047 ± 0.002 mM butanediol, consistent with previous work, and exhibited similar current profiles and magnitudes (Figure 2.4). Conversely, the ∆oxidase strain produced minimal butanediol, with only a small amount accumulating by day 6, and less than half the current of WT cells. This result highlights that the cells were unable to sustain electrode-dependent acetoin reduction in the absence of aerobic terminal oxidases. To confirm that the observed phenotype was due to the loss of PMF from proton- pumping oxidase activity, the ∆oxidase strain was functionally complemented by PR expression. If 2,3-BDO production is rescued by PR, it indicates that the loss of 2,3- 55 BDO generation in ∆oxidase is caused by a loss of PMF as opposed to off target effects, such as changes in gene regulation. When PR was expressed in ∆oxidase, we observed a restoration of 2,3-BDO production and partial rescue of current (Figure 2.4). In this instance, electron transfer to form NADH but not O2 is restored, and as one O2 is required to produce one 2,3-BDO molecule, a 50% rescue of current is consistent with our model. Taken together, these results support the hypothesis that PMF is a limiting resource, and the proton pumping activity of oxidases in this microaerobic environment is essential to continuous electron transfer to form NADH. Figure 2.4 2,3-BDO and Current Production in Microaerobic BES. (A) 2,3- Butanediol accumulation in BES with WT pBDH, ∆oxidase pBDH, and ∆oxidase pBDH- PR with nitrogen bubbling. (B) Cathodic current from BES. Noisy current marked by (*) was due to stirring issues with one of the BES replicates. Points (A) and lines (B) represent averages with standard error bars, n=3. 56 2.3.3 Reactive Oxygen Species Formation We next investigated whether increasing DO in the BES, and by extension oxidase activity, would result in an increase in 2,3-BDO. To do this, BESs were not sparged with N2 to allow passive aeration. DO measurements indicated a highly oxygenated environment (300 μM) in the BES (Figure S2.3). This condition resulted in a severe decrease in 2,3-BDO production relative to the N2-bubbling microaerobic condition (Figure 2.5). Current was greatly inflated by O2 intrusion (data not shown). Figure 2.5 2,3-BDO Production in Aerobic and Microaerobic BES. (A) 2,3- Butanediol accumulation in BES with WT pBDH with N2 bubbling (Microaerobic), and passive aeration (Aerobic) with or without the addition of 0.3 U/mL catalase. Points represent averages with standard error bars, n=3. 57 The failure of increased DO to translate to an increase in 2,3-BDO accumulation could be attributed to three factors: decreased mtrCAB expression, formation of reactive oxygen species (ROS), or a shift in electron flow to favor oxygen reduction over NADH generation. In the presence of oxygen, S. oneidensis MR-1 decreases expression of anaerobic respiration pathways such as Mtr in favor of aerobic respiration and a decrease in Mtr expression will likely result in a decrease in inward ET42. However, the cells are not actively growing under the experimental conditions, and it is improbable that a significant shift in the proteome occurred under the carbon starvation conditions of the experiment. Similarly, while loss of all electron flux in favor of oxygen reduction is possible, the small amount of 2,3-BDO produced during passive aeration suggests there are still electrons going towards NADH formation. Additionally, it has been shown that bidirectional electron transfer to NAD+ and oxygen occurs under active aeration; if all electrons were being lost to oxygen, the effect would likely have been more pronounced under those conditions32. Further research done to optimize DO concentration should explore this possibility. To assess the possibility of ROS formation, we measured hydrogen peroxide (H2O2) in the BESs. In the presence of oxygen and a strong reductant, such as a cathode or reduced flavin, O2 can be reduced to form H2O242–45. H2O2 accumulation in the BESs may result in cell death and a decrease in the ability to produce 2,3-BDO. H2O2 can also react directly with 2,3-BDO, possibly leading to reduced accumulation because of abiotic degradation46,47. To investigate if ROS accumulated in the passive aerobic condition, experiments with WT pBDH were performed with and without N2 bubbling, with and without potential, and samples were taken for colony forming unit 58 (CFU) and H2O2 measurements. When the potential was swapped from anodic to cathodic, we observed an immediate drop in CFUs/mL and generation of H2O2 in BES with passive aeration, while the microaerobic BES maintained the same levels of both (Figure 2.6). There was no detectable peroxide formation in the no potential controls (data not shown). This result demonstrates that the formation of H2O2 is dependent on the presence of oxygen and a cathode. The formation of H2O2 was correlated with ~2.5 log10 cell death in the first 3 hours. We next explored whether addition of catalase (an H2O2 degrading enzyme) could reduce H2O2 accumulation. This approach has the potential to harness the benefits of oxygen inclusion, such as PMF generation, while minimizing the production of harmful by-products42. Aerobic BESs were run with the addition of 0.3 U/mL catalase added immediately before the potential was switched to -0.5 VAg/AgCl. Aerobic BESs with catalase did not result in the same rapid decrease in CFUs/mL is increase in H2O2 as those without, as well as a less prominent spike in peroxide formation (Figure 2.6). Catalase addition also resulted in a partial rescue of 2,3-BDO production (Figure 2.5). These results show that one of the challenges with oxygen inclusion in a BES is the cytotoxic formation of H2O2, resulting in cell death and decrease in product yield. Future experimentation with aerated BES should focus on optimizing DO concentration to maximize reverse electron flux to oxygen reduction relative to forward electron flux to NAD+; having high oxidase activity to generate PMF without losing electron flow to NADH formation. 59 Figure 2.6 CFUs and [H2O2] in Aerobic and Microaerobic BES. (A) CFUs/mL and (B) [H2O2] µM in bulk medium of working chamber. Dashed line represents the potential change from 0.2 VAg/AgCl to -0.5 VAg/AgCl. Points represent averages of n=3 with standard error bars. Lines are included in CFU/mL data to guide the eye. CFUs/mL at inoculation (~46 hrs. prior to T=0) for all conditions were ~1.8 × 108. 60 2.4 Conclusion Effective microbial electrosynthesis requires attention to detail in both BES design and bacterial physiology. In the system discussed here, understanding the thermodynamic factors involved in driving inward electron transfer is crucial. The reversible nature of the electron transport pathways, which enables cells to use electrodes as electron acceptors and donors, depends on the reduction potential of each step22,48–52. Electron transfer reactions from electrode to quinol pool are freely reversible, but the final transfer from menaquinol to NADH formation has a much larger shift in potential between donor (-80 mV) and acceptor (-330 mV). This barrier is overcome by NADH dehydrogenases catalyzing the reaction and coupling the reduction to PMF utilization. In this work, we show that during electron transfer from an electrode to NADH, PMF can be regenerated by bidirectional electron transfer. Importantly, S. oneidensis’ native aerobic terminal oxidases (Cco, Cox, Cyd) can sustain PMF via oxygen reduction without fully redirecting the flow of electrons away from NADH. This ability was best demonstrated in microaerobic conditions, where the DO concentration struck the balance between electron flow to oxygen and NAD+. Higher levels of oxygen had the off-target effect of generating H2O2 that resulted in cell death. While the conditions tested here were limited to microaerobic and passively aerobic, future work should focus on fine tuning the DO in BES. This could be done through a combination of oxygen scavengers, gas mixing/modulating inflow, inclusion of other ROS neutralizing enzymes such as superoxide dismutase, or selective deletion of native oxidases as they have varying oxygen affinities and proton pumping efficiencies. The goal should be to balance the redox state of the quinone pool to maximize the flow of electrons ‘uphill’ to 61 NAD+ relative to the energetically favorable reduction of oxygen. Taken together, this work shows the strong influence even trace oxygen has on the energetics of inward electron transport. 2.5 Materials and Methods Table 2.1 Strains and Plasmids Used Strain or Description Source Plasmid S. oneidensis MR-1 Wild type S. oneidensis Meyers and Nealson, 1988 ∆oxidase Mutant with gene deletion of cco, cyd, cox, Rowe et al. (SO2361–SO2364, SO3285–SO3286, SO4606– 2018 SO4609) Plasmids pBDH pBBR1MCS2 bearing butanediol dehydrogenase Tefft and gene from Enterobacter cloacae, kanR TerAvest, 2019 pBDH-PR pBBR1MCS2 bearing butanediol dehydrogenase Tefft and from Enterobacter cloacae and proteorhodopsin TerAvest, 2019 (uncultured marine gamma proteobacterium EBAC31A08), kanR Strains and Plasmids Strains and plasmids used are listed in Table 2.1. S. oneidensis MR-1 strains were grown at 30 °C and shaking at 275 rpm for aerobic growth, and no shaking for anaerobic growth (~5% H2, balanced with N2). For BES experiments, MR-1 was pre- grown aerobically in 5 mL of lysogeny broth (LB) supplemented with 50 μg/mL kanamycin for strains with pBBR1-BDH, for inoculating minimal medium. For pre- growth, cells were grown in M5 minimal medium containing: 1.29 mM K2HPO4, 1.65 mM KH2PO4, 7.87 mM NaCl, 1.70 mM NH4SO4, 475 μM MgSO4·7 H2O, 10 mM HEPES, 0.01% (w/v) casamino acids, 1× Wolfe’s vitamin solution, and 1× Wolfe’s mineral solution, then the pH adjusted to 7.2 with 5 M NaOH. After autoclaving, D,L-lactate was 62 added to a final concentration of 20 mM. During anaerobic pre-growth, fumarate was added to a final concentration of 40 mM and 400 mL of medium was used per repeat. During bioelectrochemical experiments, the M5 medium recipe was amended to 100 mM HEPES, 0.2 μM riboflavin, and no D,L-lactate, fumarate, or casamino acids. Growth Curves For anaerobic growth experiments, cells were pre-grown in 5 mL LB supplemented with 40 mM fumarate and 20 mM D,L-lactate. Cells from the overnight culture were washed with M5 medium and resuspended to an OD600 of 0.05 in 2 mL M5 medium in a 24-well plate. OD600 was measured every 15 minutes for 35 hours in an anaerobic plate reader (BioTek, HTX). This protocol was repeated 3 times for replication. Bioelectrochemical System Experiments BES experiments were conducted in custom made two-chamber bioreactors kept at 30 °C as described in previous work (Tefft and TerAvest 2019)28, and a similar set up to work described in (Tefft et al. 2022)29. The working chamber was filled with 144 mL amended M5 medium, with 0.2 μM riboflavin being added an hour before inoculation, and the counter chamber contained ~150 mL of 1x PBS. For experiments run with PR, green LED lights were attached to the reactors. Bioreactors were autoclaved for 45 minutes, then connected to a potentiostat (VMP, BioLogic USA) and current data was collected every 1 s for the course of the experiment. After the initial setup, the working electrode poised at an anodic potential of +0.2 VAg/AgCl for ~16 hours. For aerobic pre- growth experiments, cells were grown in two 50-mL cultures of M5 in 250-mL flasks for each bioreactor (6 total for 3 replicates) for 18 hours. For anaerobic pre-growth experiments, cells were grown in 400-mL cultures of M5 in 1-L flasks for each bioreactor 63 (3 total for 3 replicates) for 18 hours. For experiments with PR, 400 μL 20 mM all-trans- retinal was added after 17 hours of growth as the essential cofactor for PR. Cultures were transferred to a 50-mL conical tube and centrifuged at 8000 rpm (Thermo Scientific ST8R; Rotor: 75005709) for 5 minutes. Pellets were washed twice in 30 mL M5 (100 mM HEPES, no carbon) and then resuspended in M5 (100 mM HEPES, no carbon), to a final OD600 of 3.6 in 10 mL. Then, 9 mL of this normalized resuspension was inoculated into the working chamber of the bioreactor using a sterile 10 mL syringe with an 18 g needle. Six hours after inoculation, N2 gas (99.999%, AirGas) was bubbled into reactors through a 0.2 μM filter, and a bubbler attached to a 0.2 μM filter connected to the gas outlet. For 40 hours after N2 bubbling, reactors were maintained at an anodic potential of +0.2 VAg/AgCl, before being changed to a cathodic potential of -0.5 VAg/AgCl. After three hours at cathodic potential, 17 mL of a sterile, de-gassed 10 mM acetoin solution was added to a final concentration of 1 mM in the bioreactor (Final volume in working chamber = 170 mL). The bioreactors were sampled (2 mL) immediately after acetoin addition for OD600 and HPLC analysis every 24 hours for 144 hours. DO Measurements DO measurements shown in Figure 3 were collected using a Hamilton VisiFerm DO sensor and ArcAir Software. The probe was calibrated before each experiment as described in the manual. The probe was inserted into the BES prior to autoclaving and secured with a rubber gasket. DO measurements were recorded every 5 s during the experiment. To ensure that the inclusion of the DO probe did not interfere with oxygen intrusion into the system, we also utilized a smaller fiber optic DO probe and collected data every 30 s using a NeoFox Fluorimeter and Software (Ocean Insight). The probe 64 consists of a patch made from 5% mixture of polymer (poly(2,2,2-trifluoroethyl methacrylate), Scientific Polymer Products Inc.) and 5 mM porphyrin (Pt(II) meso- tetra(pentafluorophenyl)porphine, Frontier Scientific) dissolved in a 50/50 mixture of 1,4- dioxane and 1,2-dichloroethane (Sigma Aldrich). This patch is deposited onto the end of a fiber optic probe53. Results from this probe corroborated observations made with the Hamilton Probe (Figure S2.3A). Data from passively aerobic BES (Figure S3B) was collected using the smaller fiber optic probe. CFU Plating During BES experiments, samples were taken every ~24 hours starting at inoculation, with additional time points in the three hours following potential change from anodic to cathodic. These samples were used for CFU plating, H2O2 measurements, and HPLC analysis. Samples were serially diluted in a 96-well plate and 10 μL of each of 8 dilutions (100-10-7) was plated on LB + Kan. Dilutions with between ~101-102 CFUs were counted and back calculated to determine CFUs/mL in bulk solution. Mean and standard error were calculated for biological replicates (n=3). H2O2 Measurements At each sampled time point, H2O2 formation was measured using the Pierce™ Quantitative Peroxide Assay Kit (ThermoFisher, Cat: 23280) according to the kit instructions. In brief, 20 μL of sample was mixed with 200 μL of reagent mixture in a 96- well plate, and absorbance was read at 595 nm. Sample values were compared to a standard curve with background subtraction of cell-only controls in 1xPBS to exclude any interference from cell OD600. Mean and standard error were calculated for biological replicates (n=3). 65 HPLC analysis HPLC analysis was performed as previously described (Tefft and TerAvest, 2019) with the amendments described in (Tefft et al., 2022)28,29. Sample analysis was performed on a Shimadzu 20A HPLC, using an Aminex HPX-87H (BioRad, Hercules, CA) column with a Microguard Cation H+ guard column (BioRad, Hercules, CA) at 65 °C with a 0.5 ml/min flow rate. 2,3-butanediol concentration in samples was calculated by comparing sample value to an external standard curve. Data analysis Analysis of HPLC data, DO %, OD, current data, and growth curve data was done using RStudio using the following packages: ggplot2, dplyr, ggpubr, plyr, data.table, stringr, and growthcurver54–60. 2.6 Author Contributions K.C.F. and M.T. conceptualized the project. K.C.F. lead the investigation and data visualization under the supervision of M.T. K.C.F. wrote the original draft of the manuscript, with review and edits by M.T. 2.7 Acknowledgements The authors would like to thank Nathan Frantz (Michigan State University - Department of Chemistry) and Dr. Denis A. Proshlyakov (Michigan State University - Department of Physiology) for their assistance with DO measurements. This research was supported by the National Science Foundation Graduate Research Fellowship Grant No. (DGE-1848739) to K.C.F., and a National Science Foundation CAREER Award (1750785) and 2018 Beckman Young Investigator Award to M.T. 66 REFERENCES (1) Rabaey, K., and Rozendal, R. A. (2010) Microbial electrosynthesis — revisiting the electrical route for microbial production. Nat Rev Microbiol 8, 706–716. (2) Karthikeyan, R., Singh, R., and Bose, A. (2019) Microbial electron uptake in microbial electrosynthesis: a mini-review. J Ind Microbiol Biotechnol 46, 1419–1426. 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(2013) Rapid electron exchange between surface-exposed bacterial cytochromes and Fe(III) minerals. Proc Natl Acad Sci U S A 110, 6346–6351. (52) Firer-Sherwood, M., Pulcu, G. S., and Elliott, S. J. (2008) Electrochemical interrogations of the Mtr cytochromes from Shewanella: Opening a potential window. Journal of Biological Inorganic Chemistry 13, 849–854. (53) Frantz, N. L., Brakoniecki, G., Chen, D., and Proshlyakov, D. A. (2021) Assessment of the maximal activity of complex IV in the inner mitochondrial membrane by tandem electrochemistry and respirometry. Anal Chem 93, 1360–1368. (54) Wickham, H. (2016) ggplot2: Elegant Graphics for Data Analysis. Springer-Verlag New York. ISBN 978-3-319-24277-4, https://ggplot2.tidyverse.org. (55) Wickham, H., François, R., and Henry, L. (2020) dplyr: A Grammar of Data Manipulation. https://dplyr.tidyverse.org, https://github.com/tidyverse/dplyr. (56) Kassambara, A. (2023) ggpubr:“ggplot2” based publication ready plots. https://CRAN.R-project.org/package=ggpubr 438. (57) Wickham, H., and Wickham, M. H. (2011) The Split-Apply-Combine Strategy for Data Analysis. J Stat Softw 40, 1–29. (58) Dowle, M., Srinivasan, A., Gorecki, J., Chirico, M., Stetsenko, P., Short, T., Lianoglou, S., Antonyan, E., Bonsch, M., and Parsonage, H. (2019) data.table: Extension of “data.frame.” https://CRAN.R-project.org/package=data.table 596. (59) Wickham, H. (2022) stringr: Simple, Consistent Wrappers for Common String Operations. http://stringr. tidyverse. org, https://github. com/tidyverse/stringr. 71 (60) Sprouffske, K., and Wagner, A. (2016) Growthcurver: an R package for obtaining interpretable metrics from microbial growth curves. BMC Bioinformatics 17, 1–4. 72 APPENDIX A: Supplementary Figures for Chapter 2 Figure S2.1 Bioelectrochemical System Design. Setup of our 2-chamber BES during an experiment with cells expressing PR, hence the inclusion of the green lights. Chambers are sealed with blue stoppers and N2 is bubbled in from a neoprene tube attached to a 0.2 µm filter (foreground). 73 Figure S2.2 Growth curves of WT and ∆oxidase. Each strain was struck on LB + 20 mM D,L-lactate + 40 mM fumarate + Kan plates and incubated anaerobically at 30°C. Three single colonies were used to start 5 mL anaerobic overnight cultures in LB + 20 mM D,L-lactate + 40 mM fumarate + Kan. Overnight cultures were spun down and pellets resuspended in M5 + 20 mM D,L-lactate + 40 mM fumarate + Kan to an OD600 of 1.0. These samples were used to inoculate 2 mL of M5 + 20 mM D,L-lactate + 40 mM fumarate + Kan in a 24-well plate at an initial OD600 of 0.05. OD600 was measured every 15 minutes for 35 hours. Lines and shaded region represent the mean and standard error, respectively, for n=3 replication. 74 Figure S2.3 Dissolved Oxygen in Microaerobic and Passively Aerobic BES. Measurements of DO in BES under N2 bubbling (A) and passive aeration (B) with the small fiber optic probe. T=0 corresponds to addition of acetoin. DO concentration for N2 bubbled reactors ~0.5-1.0 μM O2, and ~300 μM O2 after inoculation. 75 Chapter 3: Flexibility of Inward Electron Transfer Pathway in Shewanella oneidensis MR-1 Kathryne C. Ford1,2, Michaela A. TerAvest1 1 Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA 2 Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI, 48824, USA 76 3.1 Abstract The extracellular electron transport chain of S. oneidensis has been well characterized in the context of current generation on an anode. However, work to understand electron uptake from the cathode has largely relied on current data as opposed to biological outputs. To employ this organism as a biocatalyst for microbial electrosynthesis, we must have a clear picture of the path of electrons into the cell to ensure high coulombic efficiency and mitigate undesirable off-target reactions. We aim to do this by assessing the contribution of well-known electron transport pathway components to cathode-driven NADH production. In this work, we confirm that MtrCAB is essential for this, while CymA is important but can be slightly compensated for by other quinol-linked proteins. Additionally, we show that endogenous flavins are important for electron uptake, but their absence cannot be complemented by exogenous flavins. Finally, the hydrogenase HyaB plays an important role during cell survival during stationary phase on the electrode. 3.2 Introduction As a model exoelectrogenic bacterium, Shewanella oneidensis MR-1 has been extensively studied as a chassis for microbial electrosynthesis (MES). MES is a process by which microorganisms act as biocatalysts on a cathode to drive synthesis of valuable chemical products. MES systems have expanded their potential applications to include production of valuable chemicals like cancer therapeutics, as well as serving as a tool for biosensors and bioremediation1–11. As such, there are many different approaches for optimization of these platforms. One such push is the development of new tools for genetic engineering. 77 S. oneidensis MR-1 is a genetically tractable organism, with a thoroughly characterized genome and proteome12–15, making it a good target for rapid development of new tools for gene deletions, knockdowns, and precise regulation of gene expression from vectors16–21. Being able to rapidly generate different genetic modifications, and studying their resulting phenotypes, has been instrumental in growing our understanding of S. oneidensis physiology. In the context of MES, these genetic tools are employed to optimize electrons transfer between S. oneidensis and electrodes. To make such improvements, two major fields of research have investigated biofilm development and flavin export. Compared to another well-known exoelectrogen, Geobacter spp., S. oneidensis forms thinner, less conductive biofilms. Biofilms are essential for facilitating direct electron transfer to the electrode and creating an enclosed environment within which electron transfer mediators like flavins can be recycled. To this end, bioengineering work has aimed to increase biofilm thickness, conductivity, and surface area coverage22–30. These efforts have met with moderate success, with some of the greatest gains resulting from increased electrode coverage and improved conductivity of biofilm Exopolymeric substances (EPS)26,29,31,32. Excreted flavins play a role in shuttling electrons between the cell surface and the extracellular electron acceptor in S. oneidensis33–36. They also associate with outer membrane cytochromes (OMCs), such as MtrC and OmcA, to form flavocytochrome complexes37. The contribution of flavin-mediated electron transport has been debated in the context of long range versus short range transfer, as well as inward versus outward tranfer38,39. Genetic engineering allows exploration of these intricate relationships though gene overexpression or deletions. Increasing flavin export consistently increases 78 current generation40. There is also evidence that under cathodic conditions, flavins only aid electron transfer in the presence of OMCs41. Relying solely on electrical output as a benchmark for improved biotic reactions with the electrode may overlook other important factors. Assessing the impact of different genetic modifications on electron uptake should consider additional quantifiable outputs, depending on the research goal. Solely focusing on increasing electrical output may not always provide accurate insights into bacterial physiology. Electrochemistry experiments are influenced by electrode material, medium components, oxygen intrusion, and abiotic interactions with free flavins. This is further complicated when looking at differences between anodic and cathodic interactions. One way to address this is by calculating the coulombic efficiency. This is a measure of how effectively electrons are used in a specific reaction; ideally you want 100% efficiency where all the electrons are going solely to the desired reaction, or at least can be accounted for42. While this calculation has been done extensively with S. oneidensis on an anode, it has been less common for electron uptake with a cathode. There are a range of potential off target reactions during inward electron transfer. Therefore, we suggest that evaluation of genetic modifications for their impact on electron exchange should incorporate additional measurable outcomes, namely metabolite generation. One way this has already been implemented is by adding fumarate to BES to measure its reduction to succinate by FccA10,43–46. This measurement has been used to characterize the EET abilities of S. oneidensis mutants on cathodes and anodes. However, a caveat to this practice is that FccA is in the periplasm and can accept electrons directly from MtrA, therefore using fumarate as the metabolite may obscure the contribution of downstream 79 players like CymA. When examining the role of flavins, this is further complicated by the requirement of flavin mononucleotide (FMN) as a cofactor for FccA35. To address this problem, we have utilized a butanediol dehydrogenase (Bdh) based assay9,10. Within this system, electrons taken up by S. oneidensis enter the inner membrane to generate a pool of reduced menaquinol. NADH dehydrogenases use the menaquinols as redox partners to generate NADH from available NAD+ by coupling the reaction to proton motive force (PMF). NADH can be used as a redox partner for many metabolic reactions and is therefore an important intermediate for MES. In our experimental design, S. oneidensis expresses Bdh from a vector (pBDH) to convert exogenous acetoin to 2,3-butanediol through an NADH dependent reduction reaction. Because S. oneidensis does not naturally metabolize acetoin and 2,3-butanediol is stable under these conditions, we can measure the production of 2,3-butanediol as a proxy for NADH generation. We believe that this system provides a more accurate reflection of the influence that genetic modifications have on inward electron transfer in the context of MES, rather than solely relying on comparative current data. It allows for quick and straightforward quantification of product output. This system was employed to examine the role of prominent electron transport pathway components to examine their roles in inward electron transport. 3.3 Results and Discussion 3.3.1 Inward Electron Transport uses Mtr and CymA Traditionally, the electron transport pathway of S. oneidensis is described as electrons being passed from CymA to MtrCAB via periplasmic carriers, where they are then deposited onto extracellular electron acceptors though direct contact or mediated 80 by flavins. Recent developments have shown that not only is electron transfer more of a network than a pathway, but that it may be different when electrons are being taken into the cell than compared to when electrons are transported outward. We employed the Bdh system to investigate the major components of the electron transfer pathway in the organism S. oneidensis MR-1. We first examined the contribution of MtrA to electrode-dependent NADH generation during electron uptake to acetoin in a bioelectrochemical system (BES). MtrA is the decaheme cytochrome present in the periplasmic portion of the Mtr complex47–49. Of the single Mtr complex knockouts, MtrA consistently shows the strongest phenotype, and is poorly compensated for by homologs compared to MtrB and MtrC43,50. Loss of MtrA results in a severe loss of current on an anode, and a 97% decrease in current during fumarate reduction on a cathode, so we expect to see a similar decrease43,51. Additionally, determining the contribution of Mtr also sheds light on the role of hydrogen-mediated electron transport. Hydrogen formation is possible at the cathodic potential used in our experiments (-0.5 v VAg/AgCl) and could potentially act as a bypass of the Mtr pathway. MR-1 contains two hydrogenases, one of which (Hya) could potentially oxidize electrode-produced hydrogen H2 and transfer electrons into the quinol pool. Testing a ∆mtrA pBDH strain will aid in determining the flux of electrons through Mtr vs. hydrogen mediated electron transfer. In line with previous studies that looked at cathodic current in this knockout, we saw a ~95% decrease in 2,3-butanediol generation relative to WT (Figure 3.1). This supports the hypothesis that inward electron transport to form NADH relies on the Mtr pathway. Additionally, because this strain contained active hydrogenases, this result implies that hydrogen-mediated 81 electron transport does not compensate for the role of Mtr. A possible explanation for the limited accumulation of 2,3-butanediol by day 3 could be linked to cell death on the cathode creating a source of organic carbon for NADH formation. Figure 3.1 Inward Electron Transfer to Acetoin Reduction. Measurement of 2,3- butanediol accumulation and current generation in BES experiments with either WT, ∆mtrA, or ∆cymA. Cells were pre-grown in minimal medium aerobically, washed, and inoculated into anodic BES. 40 hours later, potential was changed to cathodic and acetoin was added to a final concentration of 1 mM (T=0) and samples were collected every 24 hours for HPLC analysis. Dotted line representative of abiotic current. Lines and error bars represent mean and standard error, respectively (n=3). All figures showing “WT” are the same set of experiments. The other major component of the electron transport pathway is CymA, an inner membrane tetraheme protein cytochrome c responsible for connecting electron carriers to the quinol pool. Consistent with the findings of Ross et al., we observed a significant reduction in both current and 2,3-butanediol production in the ∆cymA pBDH strain (~70%)43. This suggests that there is an alternative pathway for electrons to enter the quinol pool from MtrA. Recent work has shown the potential for such a pathway; a CymA-independent periplasmic electron transport pathway51,52. This alternative pathway 82 could account for the remaining fraction of electrode-dependent 2,3-butanediol production. This could be an uncharacterized feature of FccA or CctA or be the result of some yet unidentified shuttling protein as has been previously suggested51–53. Alternatively, Xiao et al. suggested a role for other quinol-linked oxidases like TorC or PsrC, compensating for a loss of CymA52. Relative to our results, the loss of CymA could relegate the cell to using these slower pathways. 3.2.2 Flavins Have Complex Contributions to Electron Transport The ratio of indirect to direct electron transfer (with and without soluble redox shuttles, respectively) is an important consideration for the efficiency of electron transfer. Flavins such as riboflavin (RF), flavin mononucleotide (FMN), and flavin adenine dinucleotide (FAD) are known to play a major role in electron transfer to the electrode suRFace33,34,37,40,54,55. In MR-1, FAD is transported across the inner membrane via Bfe, and cleaved in the periplasm by UshA to FMN and AMP. The FMN diffuses out of the cell, where a fraction of the extracellular FMN gradually hydrolyzes to form RF40. Extracellular RF has been shown to contribute to 70-95% of electron transfer in MR-133,34,36,56. Therefore, to probe the effects of flavins during inward electron transfer, we tested inward electron transfer capability in a strain incapable of exporting flavins to produce RF (∆bfe). The ∆bfe strain showed a significant decrease in BDO production of ~72%, which aligns almost exactly with the work by Marsili et al. for flavin impact during outward ET (Figure 3.2). We then aimed to compensate for this phenotype through the addition of exogenous RF at 1 μM. Surprisingly, while the addition of exogenous flavins resulted in a slight increase in current (Figure 3.2) it did not result in an increase in BDO production. The increase in current is likely be due to 83 abiotic interactions of photo-oxidized flavins at the electrode surface. Importantly, this did not result in increased BDO production. To determine whether the increased current was caused by abiotic reactions, sterile reactors were set up as normal with or without the addition of RF (0.2 µM). They demonstrated a similar pattern, with a slight increase in current in the reactors with RF versus those without (Figure S3.1). Figure 3.2 Endogenous flavins contribute to inward electron transfer. Measurement of 2,3-butanediol accumulation and current generation in BES experiments with either WT with exogenous RF or ∆bfe with exogenous RF or FMN. Cells were pre-grown in minimal medium aerobically, washed, and inoculated into anodic BES. 40 hours later, potential was changed to cathodic and acetoin was added to a final concentration of 1 mM (T=0) and samples were collected every 24 hours for HPLC analysis. Dashed line representative of abiotic current. Lines and error bars represent mean and standard error, respectively (n=3). We also tested electron uptake with 2 different concentrations of FMN (0.2 µM and 1 µM), because FMN associates with OMCs and increases conductivity. Similarly to results with RF, we saw a slight increase in current compared to ∆bfe with no flavin addition, but no increase in BDO production. Overall, addition of exogenous RF and FMN did not improve inward electron transfer in cells that lacked Bfe. Flavins also play 84 a regulatory role and increase expression of cytochromes and biofilm components29,57. Because cells are not actively growing in the BES, the observed phenotype could be due to regulatory changes occurring during the pre-growth in the absence of extracellular flavins. To address this, experiments with ∆bfe were repeated with either 0.2 µM RF or FMN added to the pre-growth medium and BES. This also failed to restore 2,3-butanediol accumulation to WT levels (data not shown). However, S. oneidensis cells undergo regulatory changes upon electrode attachment and during different stages of biofilm formation; cells are in stationary phase under our conditions, there may still be a regulatory role for flavins at these later time points and should be further explored57–60. Understanding the impact flavins have on inward electron transfer is particularly vital as there is much debate regarding their mechanism of action and importance under different conditions. Initial research proposed that a majority of electron exchange is mediated by free flavins in extracellular space33,34. Later evidence suggested that flavins are more important as cofactors associated with OMCs37,57,61. Additionally, flavins can stimulate transcriptional changes that impact the cytochrome profile, biofilm formation, and EPS conductivity, further obfuscating their influence41,58,59,62. Our work aims to clarify this uncertainty by directly characterizing the contribution of flavins to cathode- driven NADH production, as it does not rely solely on current data and is less susceptible to abiotic influence. To accomplish this goal, BES were inoculated with ∆bfe, with and without supplemental flavins (RF or FMN). Bfe transports FAD from the cytoplasm to the periplasm to then be cleaved by UshA to FMN and AMP. Deletion of this transporter prevents the excretion of flavins to both the periplasm and the extracellular space. This strain has a proportional decrease in both 2,3-butanediol 85 production and cathodic current (~60%*) compared to WT. While this doesn’t suggest a specific mechanism, it does demonstrate that flavins aid electron transfer. Interestingly, addition of exogenous RF or FMN does not rescue the ∆bfe phenotype for 2,3- butanediol production but does result in an increase in cathodic current. This was somewhat surprising, because it suggests that free flavins do not contribute to inward electron transfer to form NADH under these conditions. Additionally, if flavin’s main role is as cofactors for OMCs, they either cannot associate with OMCs that are already anchored in the membrane or have a stronger influence in a different capacity such as gene regulation or biofilm formation. We investigated the latter by pre-growing ∆bfe with either RF or FMN as well as supplementing the media in the BES but saw similar results. As FccA is a known periplasmic carrier that uses FMN as a cofactor, we tested a strain with a knockout of fccA to determine if this was the source of this observed phenotype, but observed results comparable to WT. Future work should investigate this phenomenon to determine why exogenous flavins appear unable to fulfill the role of endogenous flavins for inward electron transfer to NAD+. Also of note, the current in the ∆bfe BES increased with either RF or FMN in the medium. This result exemplifies why relying solely on current data may obscure our understanding of biological versus abiotic processes. During electron transfer, periplasmic c-type cytochromes facilitate the movement of electrons between CymA and MtrA, such as the small tetraheme cytochrome CctA and the fumarate reductase FccA. FccA uses FMN as an essential cofactor, which is unavailable in the ∆bfe strain35. To confirm that the phenotype of the Δbfe strain was due to flavin availability and not an off-target effect of an inactive FccA, we tested the 86 effect of a fccA deletion on 2,3-butanediol production. There was no significant difference in current or BDO production for ΔfccA compared to WT (Figure 3.3). There is evidence that FccA and CctA are functionally redundant, so the loss of FccA is likely compensated by CctA or other periplasmic electron carriers63–65.This result confirms that the phenotype seen in the flavin experiments is not attributable to FccA lacking its cofactor. Figure 3.3 FccA is dispensable for Inward Electron Transfer. Measurement of 2,3- butanediol accumulation and current generation in BES experiments with either WT or ∆fccA. Cells were pre-grown in minimal media aerobically, washed, and inoculated into anodic BES. 40 hours later, potential was changed to cathodic and acetoin was added to a final concentration of 1 mM (T=0) and samples were collected every 24 hours for HPLC analysis. Dashed line representative of abiotic current. Lines and error bars represent mean and standard error, respectively (n=3). 3.3.3 Hydrogenases Enhance Cell Survival on Anode In previously published work using this system, we noted that deletion of the two hydrogenases (∆hyaBhydA) resulted in an increase in BDO accumulation9. This increase was attributed to hydrogen generation acting as an electron sink, siphoning off 87 electrons that would otherwise have gone to form NADH. However, the experimental protocol has since been modified by extending the anodic phase prior to acetoin addition. This delay was done for the purpose of removing residual organic carbon to prevent electrode-independent NADH formation. When ∆hyaBhydA was used with this new protocol, we observed a decrease in both current and BDO accumulation relative to WT (Figure 3.4A,B). Single hydrogenase deletion mutants (∆hyaB, ∆hydA) indicated that this phenotype is largely attributable to HyaB; ∆hyaB showed the same 2,3- butanediol production as the double knockout, while ∆hydA behaved like WT. However, both single knockouts had current increase compared to ∆hyaBhydA. Figure 3.4 Hydrogenases Contribute to Cell Survival. Measurement of 2,3- butanediol accumulation (A) and current generation in BES experiments with either WT, ∆hyaBhydA, ∆hydA or ∆hyaB (B). Cells were pre-grown in minimal medium aerobically, washed, inoculated into anodic BES. 40 hours after inoculation, acetoin was added to a final concentration of 1 mM (T=0) and samples were collected every 24 hours for HPLC analysis. WT and ∆hyaBhydA BES samples were plated for CFUs every 24 hours starting at inoculation (T=-2) (C). Dotted line in (B) representative of abiotic current. Lines and error bars represent mean and standard error, respectively (n=3). HyaB is predicted to be a bidirectional hydrogenase, capable of either generating or oxidizing H2, while HydA only generates H266–68. Therefore, one potential explanation for this observation is that hydrogen mediates electron transfer between the cathode and the inner membrane quinols. However, as previously stated, most (~95%) of the 88 electrons being transported rely on the Mtr complex, making it unlikely that hydrogen is a major contributor to electron shuttling. A hypothesis that we find more likely is that the hydrogenases, primarily HyaB, act as a release valve to prevent overreduction of the redox pool in the cell. This is supported by Joshi et al. who showed greater survival of cells during stationary phase when hydrogenases were present68. To test this, WT and ∆hyaBhydA BES experiments were performed again and samples were taken every 24 hours starting from inoculation and plated for CFUs (Figure 3.4C). Overall, the CFUs/mL in the bulk medium decreased over time for both strains, which is not surprising as cells become attached to the electrode over time. However, while the CFUs/mL for the two strains were similar for the first 24 hours, by 48 hours ∆hyaBhydA decreased by a log10 more than WT. This finding is similar to the pattern of survival seen by Joshi et al. for cells during stationary phase. Therefore, we can attribute the decrease in 2,3-butanediol production to cell death during the first 48 hours of the experiment. This result also coincides with the time before acetoin addition (T=0) being extended compared to earlier iterations. 3.4 Conclusions MES is a technology that often utilizes the extracellular electron transfer capabilities of exoelectrogenic microorganisms to produce a variety of industrial chemicals, including biofuels and bioplastics. Recent developments in this field have focused on optimizing microbes in the system. In S. oneidensis, this approach has included streamlining the cytochrome network, increasing cell attachment to electrodes, and overproduction of electron shuttles40,61,69. A variety of creative strategies have greatly increased the electrical output in these systems. However, the increase in 89 current may not always translate to increased biotic activity. This inconsistency is a problem for research dedicated to the generation of industrially relevant products powered by electricity from a cathode. Our solution to this is the use of a Bdh-based system in which we can measure 2,3-butanediol accumulation as a proxy for NADH generation (Figure 3.5). For the purposes of optimizing inward electron transfer to NAD+ via genetic manipulation, this system allows for accurate more assessment of efficiency than just current. Figure 3.5 Inward Electron Transfer Path to Acetoin in S. oneidensis. During inward electron transfer to form NADH for acetoin reduction to 2,3-butanediol, electrons are taken up from a cathode via MtrCAB. Periplasmic electron carriers (PECs) shuttle electrons to CymA, or other quinol-linked reductases to a lesser extent. Endogenous exported flavins aid in inward electron transfer via mediation as free flavins, as OMC cofactors, or in a regulatory capacity. HyaB aids in cell survival during stationary phase on the electrode. 90 3.5 Materials and Methods Table 3.1 Strains and Plasmids Used Strain or Description Source Plasmid S. oneidensis MR-1 Wild type S. oneidensis Myers and Nealson, 198870 ∆mtrA Mutant with gene deletion of SO_1777 Rowe et al. 201871 ∆cymA Mutant with gene deletion of SO_4591 This work ∆bfe Mutant with gene deletion of SO_0702 Kotloski and Gralnick, 201333 ∆fccA Mutant with gene deletion of SO_0970 Gao et al., 201072 ∆hydA Mutant with gene deletion of SO_ 3920 This work ∆hyaB Mutant with gene deletion of SO_2098 This work ∆hyaBhydA Mutant with gene deletion of SO_3920 and Tefft and TerAvest, SO_2098 20199 Plasmids pBDH pBBR1MCS2 bearing butanediol Tefft and TerAvest, dehydrogenase gene from Enterobacter 20199 cloacae, kanR Strains and Plasmids Strains and plasmids used are listed in Table 1. S. oneidensis MR-1 strains were grown at 30 °C and shaking at 275 rpm for aerobic growth. For bioelectrochemical system experiments, MR-1 was pre-grown aerobically in 5 mL of lysogeny broth (LB) supplemented with 50 μg/mL kanamycin for strains with pBBR1-BDH, for inoculating minimal medium. For pre-growth, cells were grown in M5 minimal medium containing: 1.29 mM K2HPO4, 1.65 mM KH2PO4, 7.87 mM NaCl, 1.70 mM NH4SO4, 475 μM MgSO4·7 H2O, 10 mM HEPES, 0.01% (w/v) casamino acids, 1× Wolfe’s vitamin 91 solution, and 1× Wolfe’s mineral solution, then the pH adjusted to 7.2 with 5 M NaOH. After autoclaving, D,L-lactate was added to a final concentration of 20 mM. During bioelectrochemical experiments, M5 medium recipe was amended to 100 mM HEPES, 0.2 μM riboflavin, and no D,L-lactate, or casamino acids. Bioelectrochemical System Experiments Bioelectrochemical system experiments were conducted in custom made two-chamber bioreactors kept at 30 °C as described in previous work (Tefft and TerAvest 2019)9, and a similar set up to work described in (Tefft et al. 2022)10. The working chamber was filled with 144 mL amended M5 media, with 0.2 μM riboflavin being added an hour before inoculation, and the counter chamber contained ~150 mL of 1x PBS. Bioreactors were autoclaved for 45 minutes, then hooked up to a potentiostat (VMP, BioLogic USA) and current data was collected every 1 s for the course of the experiment. After the initial setup, the working electrode poised at an anodic potential of +0.2 VAg/AgCl for ~16 hours. For aerobic pre-growth experiments, cells were grown in two 50-mL cultures of M5 in 250-mL flasks for each bioreactor (6 total for 3 replicates) for 18 hours. Cultures were transferred to a 50-mL conical tube and centrifuged at 8000 rpm (Thermo Scientific ST8R; Rotor: 75005709) for 5 minutes. Pellets were washed twice in 30 mL M5 (100 mM HEPES, no carbon) and then resuspended in M5 (100 mM HEPES, no carbon), to a final OD600 of 3.6 in 10 mL. Then, 9 mL of this normalized resuspension was inoculated into the working chamber of the bioreactor using a sterile 10 mL syringe with an 18 g needle. 6 hours after inoculation, N2 gas (99.999%, AirGas) was bubbled into reactors through a 0.2 μM filter, and a bubbler attached to a 0.2 μM filter connected to the gas outlet. For 40 hours after N2 bubbling, reactors were maintained at an anodic 92 potential of +0.2 VAg/AgCl, before being changed to a cathodic potential of -0.5 VAg/AgCl. After 3 hours at cathodic potential, 17 mL of a sterile, de-gassed 10 mM acetoin solution was added to a final concentration of 1 mM in the bioreactor (Final volume in working chamber = 170 mL). The bioreactors were sampled (2 mL) immediately after acetoin addition for OD600 and HPLC analysis every 24 hours after that for 72 hours. CFU Plating During BES experiments, samples were taken every ~24 hours starting at inoculation, with additional time points in the three hours following potential swap from anodic to cathodic. These samples were used for CFU plating, H2O2 measurements, and HPLC analysis. Samples were serially diluted in a 96-well plate and 10 μL of each of 8 dilutions (100-10-7) was plated on LB + Kan. Dilutions with between ~101-102 CFUs were counted and back calculated to determine CFUs/mL in bulk solution. Mean and standard error were calculated for biological replicates (n=3). HPLC analysis HPLC analysis was performed as previously described (Tefft and TerAvest, 2019) with the amendments described in (Tefft et al., 2022)9,10. Sample analysis was performed on a Shimadzu 20A HPLC, using an Aminex HPX-87H (BioRad, Hercules, CA) column with a Microguard Cation H+ guard column (BioRad, Hercules, CA) at 65 °C with a 0.5 ml/min flow rate. 2,3-butanediol concentration in samples was calculated by comparing sample value to an external standard curve. 93 Data analysis Analysis of HPLC data, DO %, OD, current data, and growth curve data was done using RStudio using the following packages: ggplot2, dplyr, ggpubr, plyr, data.table, stringr, , and growthcurver73–79. 3.6 Author Contributions K.C.F. and M.T. conceptualized the project. K.C.F. lead the investigation and data visualization under the supervision of M.T. K.C.F. wrote the original draft of the manuscript, with review and edits by M.T. 3.7 Acknowledgements This research was supported by the National Science Foundation Graduate Research Fellowship Grant No. (DGE-1848739) to K.C.F., and a National Science Foundation CAREER Award (1750785) and 2018 Beckman Young Investigator Award to M.T. 94 REFERENCES (1) Yi, Y. C., and Ng, I. S. (2021) Redirection of metabolic flux in Shewanella oneidensis MR-1 by CRISPRi and modular design for 5-aminolevulinic acid production. Bioresour Bioprocess 8, 1–11. (2) Wang, J., Wu, M., Lu, G., and Si, Y. (2016) Biotransformation and biomethylation of arsenic by Shewanella oneidensis MR-1. Chemosphere 145, 329–335. (3) Webster, D. P., TerAvest, M. A., Doud, D. F. R., Chakravorty, A., Holmes, E. C., Radens, C. M., Sureka, S., Gralnick, J. A., and Angenent, L. T. (2014) An arsenic- specific biosensor with genetically engineered Shewanella oneidensis in a bioelectrochemical system. Biosens Bioelectron 62, 320–324. 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Current generation in abiotic BES experiments with (purple) or without (black) the addition of 1 µM RF. Lines and error bars represent mean and standard error, respectively (n=3). 102 Chapter 4: Flux Balance Analysis and Mobile CRISPRi guided deletion of a conditionally essential gene in Shewanella oneidensis MR-1 Kathryne C. Ford1,2, Joshua A.M. Kaste1,3, Yair Shachar-Hill3, Michaela A. TerAvest1* 1 Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA 2 Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI, 48824, USA 3 Department of Plant Biology, Michigan State University, East Lansing, MI, 48824, USA This chapter is adapted from a publication in ASC Synthetic Biology 103 4.1 Abstract Carbon neutral production of valuable bioproducts is critical to sustainable development but remains limited by the slow engineering of photosynthetic organisms. Improving existing synthetic biology tools to engineer model organisms to fix carbon dioxide is one route to overcoming the limitations of photosynthetic organisms. In this work, we describe a pipeline that enabled deletion of a conditionally essential gene from the Shewanella oneidensis MR-1 genome. S. oneidensis is a simple bacterial host that could be used for electricity-driven conversion of carbon dioxide in the future with further genetic engineering. We used Flux Balance Analysis (FBA) to model carbon and energy flows in central metabolism and assess the effects of single and double gene deletions. We modeled the growth of deletion strains under several alternative conditions to identify substrates that restore viability to an otherwise lethal gene knockout. These predictions were tested in vivo using a Mobile-CRISPRi gene knockdown system. The information learned from FBA and knockdown experiments informed our strategy for gene deletion, allowing us to successfully delete an ‘expected essential’ gene, gpmA. FBA predicted, knockdown experiments supported, and deletion confirmed that the ‘essential’ gene gpmA is not needed for survival, dependent on the medium used. Removal of gpmA is a first step towards driving electrode powered CO2 fixation via RuBisCO. This work demonstrates the potential for broadening the scope of genetic engineering in S. oneidensis as a synthetic biology chassis. By combining computational analysis with a CRISPRi knockdown system in this way, one can systematically assess the impact of conditionally essential genes and use this knowledge to generate mutations previously thought unachievable. 104 4.2 Introduction Mounting challenges associated with anthropogenic carbon emissions have led researchers to investigate microbial engineering solutions to capture carbon dioxide (CO2). Additionally, bacterial products made from fixed CO2 represent a carbon-neutral platform for generation of biofuels and chemicals. To achieve these goals, researchers employ an ever-expanding list of genetic engineering techniques to rewire microbial central carbon metabolism. These include broad approaches such as expression vectors and inhibitors, to specific genetic modifications like gene deletions and site- directed mutagenesis1,2. The latter can be an arduous task, depending on the organism and target site. Synthetic biology workhorse organisms like Escherichia coli and Saccharomyces cerevisiae have been studied extensively in the context of rewiring central carbon metabolism for the purposes of carbon capture and bioproduction3–12. As such, they come with a substantial genetic toolkit and mutant libraries (e.g., the Keio Collection)13,14 that allow quick and easy genome modifications. A prominent example relevant to this work is the development of a complete CO2 fixation pathway in E. coli15– 17 . In their work, Antonovsky et al. utilized a combination of Flux Balance Analysis (FBA), phage transduction of mutations in the Keio Collection, heterologous protein expression, and laboratory evolution to develop a CO2-fixing strain. This highlights the fact that even in well-understood bacterial species, bioengineering is typically much harder in vivo than on paper. When applying these strategies to non-model organisms with a less extensive investigative history, it can be more difficult to create the desired mutations. Here, we combined flux-balance analysis (FBA) and a CRISPRi gene knockdown system as a pipeline for troubleshooting difficult mutations and enabling 105 deletion of conditionally essential genes. Specifically, this was done in the electroactive bacteria Shewanella oneidensis MR-1 for the deletion of the gene gpmA. Creation of this mutant demonstrates the capacity of this approach to expand the scope of feasible genetic alterations. S. oneidensis MR-1 is a γ-proteobacterium commonly used in biotechnology research due to its genetic tractability and diverse metabolism18. It is a Gram-negative, facultative anaerobe that contains an extracellular electron transfer pathway (the Mtr pathway) that enables it to use extracellular metal oxides and electrodes as electron acceptors for respiration19–21. The Mtr pathway can also be reversed, expanding the potential for catalyzing electrode-driven carbon reduction22–24. A major benefit of using an electrode as an electron donor during bioproduction is the ability to use renewable energy, such as wind or solar, to power the reduction of CO2, eliminating the need for exogenous chemical electron donors25,26. Another benefit of using S. oneidensis is that previous research has demonstrated that S. oneidensis’ central carbon metabolism can be altered to grow on carbon substrates it cannot natively use, such as glucose27–29. S. oneidensis’ preferred carbon source is D,L-lactate, which is converted to pyruvate via lactate dehydrogenases (Ldh, Dld, Lld)30–32. In wild-type (WT) S. oneidensis, this pyruvate is used both to generate reducing power and ATP via oxidation by the TCA cycle and for gluconeogenesis and other biomass-building processes. A key reaction that connects these processes is catalyzed by the enzyme phosphoglycerate mutase (Gpm)33 (Figure 4.1). In this work, we sought to separate energy production and biomass synthesis in S. oneidensis as a first step toward rewiring it for carbon fixation through deletion of 106 gpmA. This would enable future repurposing of the existing gluconeogenesis machinery into a carbon-fixing Calvin-Benson Cycle (CBC), as was previously done in a ∆gpmAM mutant of E. coli15–17. However, before any carbon fixing pathway could be constructed, there were significant challenges associated with gpmA deletion in S. oneidensis MR-1. Deutschbauer et al. previously classified gpmA as ‘essential’ based on Flux Balance Analysis (FBA) and a lack of transposon insertion mutants in gpmA when the library was generated in LB medium34,35. However, Gpm (gpmAM) is not essential in E. coli, and E. coli ∆gpmAM mutants can grow in LB or minimal medium when two carbon sources are provided, one entering metabolism ‘above’ the Gpm reaction and one entering ‘below.’ This led us to hypothesize that gpmA was only conditionally essential in S. oneidensis. However, initial attempts to create a knockout using a common sucrose counter selection protocol were unsuccessful21,36. To overcome this pitfall, we adapted existing technologies to enable systematic modeling and testing of different growth media. Because the target gene participates in carbon metabolism, we hypothesized that this pipeline would reveal optimized conditions that allow the survival of gpmA mutants. We used FBA to predict conditions that would enable an S. oneidensis ΔgpmA strain to grow and implemented a CRISPRi gene knockdown system for characterization of gpmA in S. oneidensis37,38. In combination, in silico modeling and systematic testing of phenotypic responses of gene knockdown in different media enabled deletion of a gene thought to be essential. 107 Figure 4.1 Central carbon metabolism of S. oneidensis MR-1. Metabolites are shown in green boxes, and the genes encoding the proteins which catalyze enzymatic reactions (black arrows) are shown in red. This schematic comprises components of glycolysis, the pentose phosphate pathway, and the TCA cycle. The targeted reaction catalyzed by the product of gpmA is in the yellow box. Gene names listed in red are as annotated in NCBI for S. oneidensis genome (NCBI:txid211586). 4.3 Results We initially attempted to delete gpmA from the S. oneidensis genome using a homologous recombination method that is widely used for this organism21,36,39–41. Briefly, sequences homologous to regions upstream and downstream of gpmA were cloned into a non-replicating vector containing an antibiotic selection gene and sacB, then conjugated into S. oneidensis where it inserted into the genome at the gpmA locus to create primary integrants. A second homologous recombination event was enabled 108 by growth without antibiotic, and we selected for cells that had resolved the vector out of the genome by plating on 10% sucrose. The resulting colonies were screened for gpmA deletion or reversion to WT. Initial efforts were not successful, and all colonies (~300) screened after plasmid resolution were WT revertants. However, because gpmA could be deleted from E. coli and the mutant grew well when provided with separate energy and sugar backbone sources, we explored whether a two-substrate strategy would enable gpmA deletion in S. oneidensis. We used FBA to model the maximum theoretical specific growth rate (i.e., the growth potential [h-1]) of a WT strain and a ΔgpmA strain under a range of single- and double-carbon source conditions. To do this, we used FBA optimization functions built into the COBRA Toolbox42 and applied them to an existing model of S. oneidensis metabolism43. Based on previous work15,16, we hypothesized that a ΔgpmA strain would require two carbon sources, with one entering metabolism ‘above’ and one entering ‘below’ the gpmA deletion (Figure 4.1). We chose nucleosides (adenosine, inosine, and uridine) as three ‘upper’ carbon sources and lactate as the model ‘lower’ carbon source to model a strain grown in M5 Minimal Medium44. These carbon sources were chosen because they are known substrates for S. oneidensis39,45,46, which does not typically grow using sugars such as glucose, but is capable of robust growth on nucleosides47–51. As expected, we observed that a WT strain had a non-zero predicted growth potential in all the tested conditions, including single substrates (Table 4.1). In contrast, FBA analysis of the ΔgpmA strain showed zero growth potential when provided only lactate. Notably, the ΔgpmA strain showed non-zero growth potential only when supplemented with a carbon source entering ‘above’ gpmA, alone or in addition to lactate. 109 Table 4.1 Calculated growth potential of S. oneidensis MR-1 WT and ∆gpmA strains with different substrates Flux balance analysis was used to determine the effect of gpmA deletion on the growth potential of S. oneidensis. The predicted specific growth rate was calculated for WT and ∆gpmA cells grown in M5 medium with fixed uptake rates for either solely lactate, adenosine, inosine, uridine, or in combination. Strain Carbon Growth Potential (h-1) WT Lactate 0.533 Adenosine 0.543 Lactate + Adenosine 1.093 Inosine 0.547 Lactate + Inosine 1.103 Uridine 0.549 Lactate + Uridine 1.105 ΔgpmA Lactate 0 Adenosine 0.526 Lactate + Adenosine 1.079 Inosine 0.530 Lactate + Inosine 1.089 Uridine 0.534 Lactate + Uridine 1.092 We next used a CRISPRi-based gene knockdown method to experimentally evaluate the predictions of the FBA. In this system, a catalytically inactive dCas9 protein and a small guide RNA (sgRNA) targeting the gene of interest are chromosomally expressed under an isopropyl ß-D-1-thiogalactopyranoside (IPTG)-inducible promoter. The 20 bp tag on the sgRNA binds immediately downstream of the start codon on the gene of interest, and the dCas9 then binds to the sgRNA to prevent transcription37,38. Strains of S. oneidensis expressing the inducible dCas9 and gene-targeting sgRNA were pre-grown in Lysogeny Broth (LB), then plated on LB + 10 mM IPTG. We constructed three strains with this system, with sgRNA targeting either gpmA, rpoC (essential gene control), or no target (negative control). During growth on LB with 110 inducer, the strain with a gpmA-targeting sgRNA showed a severe growth defect compared to growth on LB without IPTG, similar to a strain with a rpoC-targeting sgRNA (Figure 4.2). In contrast, when the growth medium was supplemented with uridine and lactate, the gpmA knockdown strain survived similarly to the non-targeting sgRNA control, although it took longer for colonies to develop (24 hours) compared to WT (16 hours) (Figure 4.3). Figure 4.2 CRISPRi gene knockdown in S. oneidensis MR-1. Strains of S. oneidensis MR-1 were constructed to chromosomally express dCas9 and sgRNA targeting (A) nothing (non-essential gene control), (B) rpoC (essential gene control), or (C) gpmA. Both the dCas9 and sgRNA are under ITPG induction. Strains were pre- grown in LB + selection at 30oC. 3ul of serial ten-fold dilutions of a 1.0 OD600 cell suspension were plated on LB +/- inducer (10 mM IPTG) as described in Materials and Methods. Each panel shows replicate (n=3) plating of each strain. Figure 4.3 CRISPRi knockdown of genes in S. oneidensis MR-1 with supplemented media. Previously constructed strains of MR-1, with sgRNA targeting either nothing (A) or gpmA (B), were pre-grown as before. This time, strains were plated on LB + 10mM IPTG supplemented with 10mM uridine and 20mM lactate. The growth rate of the gpmA knockdown was restored to WT-levels under these conditions. Each panel shows replicate (n=3) plating of each strain. 111 Based on FBA and knockdown results suggesting that gpmA is non-essential when an upper carbon source and a lower carbon source are provided simultaneously, we repeated the gene deletion protocol and included 10 mM uridine and 20 mM lactate in the LB. In this iteration, a ΔgpmA strain was obtained (Figure 4.4A). Of the 50 colonies screened, 23 of them were ΔgpmA, 13 were WT revertants, and 14 failed to resolve out the vector. We grew the resulting ΔgpmA strain and WT cells in LB in a 96- well plate at 30°C with shaking and observed that the mutant had a severe growth defect (WT = 0.17 h-1, ∆gpmA = 0.08 h-1). Figure 4.4 Identification and Aerobic growth of S. oneidensis WT vs. ∆gpmA cells in LB. (A) PCR amplification of the gpmA locus from WT S. oneidensis MR-1 and a gpmA deletion strain obtained by homologous recombination under modified conditions. Primers were located approximately 500 bp upstream and downstream of the coding region of the 1.7 kb gene. (B) Overnight cultures were used to inoculate 500 µl of LB in a 48-well plate for a starting OD600 of 0.1 in the wells. The plate was incubated with constant shaking at 30°C in an aerobic plate reader, reading the OD600 every 15 mins. Lines represent the simple moving average of three biological replicates for WT cells (green, solid) and ∆gpmA cells (black, dashed), and grey ribbons represent standard deviation. 112 Figure 4.5 Experimental confirmation of growth capabilities of ∆gpmA cells grown in minimal medium with various substrates. M5 medium was prepared as described in the Materials and Methods. Growth of (A) ΔgpmA, and (B) WT with lactate (blue), the designated nucleoside (yellow), or a combination of the two (red) (lactate results are repeated in each graph). The ∆gpmA cultures were grown overnight aerobically in LB at 30°C. Cells were normalized to an OD600 of 1.0 and 500 µl of each was inoculated into the M5 medium in a 48-well plate. The plate was incubated with constant shaking at 30°C in a plate reader, measuring the OD600 every 15 mins. Lines represent the simple moving average of three biological replicates, and grey ribbons represent the standard deviation. We next measured growth of the mutant strain with the substrate combinations that were assessed by FBA. WT and ΔgpmA cultures were pre-grown in LB and washed with minimal medium with no carbon substrate or casamino acids. Cells were resuspended to an OD600 of 0.1 in M5 with either a single substrate or a combination of two substrates and incubated with shaking at 30°C. The ΔgpmA cells did not grow with any single substrate but did grow when provided two substrates (Figure 4.5A) while the WT culture grew under all conditions (Figure 4.5B). The highest final cell density of the ΔgpmA culture was observed with the combination of D,L-lactate and inosine. Cultures 113 provided solely nucleosides saw a decrease in OD600 while those provided D,L-lactate remained relatively stable at the starting density. This may be due to a small of amount of D,L-lactate being metabolized for energy but not for biomass, while nucleosides could not support any metabolic activity. 4.4 Discussion Using computationally guided methods, we deleted gpmA from the S. oneidensis genome although it was previously classified as essential. We hypothesized that two carbon sources would be required to enable the ∆gpmA strain to grow based on previous results with E. coli and the FBA for S. oneidensis (Figure 4.6)34. While our experiments agreed with most FBA predictions, the model predicted that the ∆gpmA cells would grow solely on nucleoside substrates that enter metabolism ‘above’ GpmA. However, we observed experimentally that nucleosides alone did not support growth of the ∆gpmA strain. We hypothesize that this discrepancy was caused by unrealistic pathway flexibility in the model. Specifically, the predicted growth of ΔgpmA on nucleosides alone relied on conversion of 3-phosphoglycerate to serine, which was broken down to pyruvate that could be used in the TCA cycle. However, the required flux through this workaround pathway was ~10-fold higher than in the WT model and is likely infeasible in vivo without overexpression of the enzymes involved. Thus, gpmA deletion functionally blocks nucleosides from being fully oxidized although there were possible workarounds in the metabolic model. Therefore, even though the model predicts growth on nucleosides, we did not observe this outcome due to the energy limitations imposed by the cutoff of the TCA cycle and by extension, ATP generation by oxidative phosphorylation. In future work, this issue could be addressed through 114 refinement of the model through the inclusion of proteomic and transcriptomic data to better represent energy acquisition in cells during growth. The inspiration for the project was to develop of a CO2 fixing strain of S. oneidensis using a similar method to Antonovsky et a15l., however, crucial differences from E. coli created significant challenges. Specifically, gpmA deletion was difficult in S. oneidensis and this gene was classified as essential34. We utilized a gene knockdown system called ‘mobile CRISPRi’ to evaluate FBA predictions of conditions that would enable growth of a gpmA knockout strain. On LB, the phenotype for a gpmA-targeting sgRNA was comparable to that of an sgRNA targeting the essential gene rpoC (105-fold decrease in fitness), supporting the hypothesis that gpmA is essential for growth on LB. This explains why the transposon library used by Deutschbauer et al., did not contain any gpmA mutants, because the library was generated on LB52. Indeed, even an improved library preparation protocol starting with over 39,000 transposon mutants did not yield a gpmA disruption, likely because this method also generated the library on LB53. While transposon library sequencing is an immensely informative technique for genome annotation and fitness analysis, our results highlight the biases introduced by the library-generation conditions. The CRISPRi knockdown system utilized in this work could be a powerful tool towards expanding our knowledge of gene functionality built on the foundations created by transposon libraries. Namely, complementation of transposon mutagenesis libraries with CRISPRi knockdown libraries will enable high- throughput characterization of conditionally essential genes38. 115 Figure 4.6 Metabolic strategy of MR-1 ∆gpmA cells. With the deletion of gpmA, metabolites entering the cell as a TCA cycle intermediate are cutoff from entering glycolysis to build biomass, and vice versa. To satisfy the cell requirements for growth, MR-1 ∆gpmA requires at least two carbon sources to grow; one entering ‘above’ gpmA to build biomass (nucleoside), and one entering ‘below’ gpmA to acquire energy and reducing equivalents (lactate). While use of various existing genetic tools is a staple of synthetic biology work, there are gaps in the prospective scope of their implementation, such as using them for deletion of genes classified as essential. Through this work, we have devised a means to generate such a novel deletion. This both expands the bounds of potential synthetic biology applications in S. oneidensis and paves the way for development of a CO2 fixing strain. Because the ΔgpmA mutant requires a sugar source, it can form the basis for laboratory evolution experiments to activate a heterologous pCBB pathway during growth with limiting sugar, as previously demonstrated by Antonovsky et al15. The pipeline developed here can be used to predict and characterize the potential phenotypes associated with deletion of conditionally essential genes, thereby informing strategies to construct the deletions. The use of FBA and CRISPRi analysis is a modular system that uses existing models and genetic tools and has high adaptability to 116 work in other common laboratory microorganisms. To our knowledge, this is the first instance of using these specific systems to generate difficult or otherwise infeasible gene deletions. This capability is especially useful for probing the effects of genes that are essential under typical laboratory conditions and opens the door to exploring the phenotypes of gene deletions previously thought to be lethal. Biotechnology research relies on the ability to manipulate microorganisms’ genomes to perform a specific task, such as engineering carbon fixation in heterotrophs. Expanding our toolkit to include deletion of conditionally essential genes is a crucial step to achieving these goals. 4.5 Materials and Methods Strains and Culture Media Table 4.2 Strains and plasmids used in this study Strain or Plasmid Description Source E. coli WM3064 Cloning and conjugation strain for S. oneidensis, dap- WM6026 sJMP2644 – Tn7 Transposase donor strain, Peters 2019 contains pJMP1039, dap- BW25141 sJMP2846 – CRISPRi transposon donor strain used Peters 2019 for cloning sgRNA, contains pJMP2846 S. oneidensis MR-1 Wild type Meyers and Nealson, 1988 ∆gpmA Mutant with gene deletion of gpmA This study sgRNA-gpmA MR-1 strain containing the CRISPRi knockdown This study vector targeting gpmA (pJMP2846-gpmA) sgRNA-rpoC MR-1 strain containing the CRISPRi knockdown This study vector targeting rpoC (pJMP2846-rpoC) sgRNA-EV MR-1 strain containing the CRISPRi knockdown This study vector targeting no genes (pJMP2846) Plasmids pJMP1039 Tn7 transposase donor vector, ampR Peters 2019 pJMP2846 CRISPRi transposon donor plasmid, used for Peters 2019 cloning sgRNA, kanR pJMP2846- CRISPRi transposon donor plasmid, knocks down This study gpmA gpmA 117 Table 4.2 (cont’d) pJMP2846-rpoC CRISPRi transposon donor plasmid, knocks down This study rpoC pSMV3.0 Deletion vector, KanR, sacB Saltikov and Newman 2003 Table 4.3 Primers used for making in-frame deletion mutants and CRISPRi knockdowns Primer Primer Sequence Deletions gpmA FO ACGAGGTCATGCCAGCATTGCA RO GCACTTGTTGCTCGGCCATCAA CRISPRi - knockdown gpmA FWD TAGTTCGAGGATCAACAACGCGAT REV AAACATCGCGTTGTTGATCCTCGA rpoC FWD TAGTGACCAAGAACGGATCAGATC REV AAACGATCTGATCCGTTCTTGGTC Strains and plasmids used are listed in Table 4.2 and primers in Table 4.3. This work was done in Shewanella oneidensis MR-1 and using Escherichia coli WM3064, WM6026, and BW25141 for plasmid construction and conjugation. Strains were grown using LB for growth curves and supplemented with 30 mM diaminopimelic acid (DAP), 50 μg/mL kanamycin (Kan) or 100 μg/mL ampicillin (Amp) as needed for construction of CRISPRi knockdown strains. M5 Minimal Medium (1.29 mM K2HPO4, 1.65 mM KH2PO4, 7.87 mM NaCl, 1.70 mM NH4SO4, 475 μM MgSO4·7H2O, 10 mM HEPES, 1× Wolfe’s mineral solution, and 1× Wolfe’s vitamin solution), with pH adjusted to 7.2 with NaOH. Casamino acids and riboflavin were not included in the media. M5 medium was supplemented with various carbon sources to final concentrations of: 20mM D,L-lactate and 10mM nucleosides (adenosine, uridine, or inosine). Growth experiments were conducted in either 96- or 48-well plates and OD600 was read using an H1M BioTek 118 Plate Reader. E. coli strains were grown at 37°C with shaking, and MR-1 strains were grown at 30°C, with shaking, with a starting OD600 of 0.1. Flux Balance Analysis The genome-scale model of S. oneidensis MR-1’s metabolic network was taken from Ong et al54. The ATP growth-associated maintenance (GAM) cost was removed from this version of the MR-1 model due to a lack of GAM measurements for other S. oneidensis strains that were being compared with MR-1. The GAM was added back in by referring to the information from an earlier iteration of the model43. The objective function for all optimizations was the maximization of flux through the model’s biomass equation, followed by the minimization of the sum of all fluxes. All optimizations were performed in MATLAB (version 2019a) using the COBRA Toolbox55 and version 8.1.1 of the Gurobi optimizer42. To model carbon utilization, specific uptake rates were set for each substrate, with lactate provided at two times the value (-20 mmol/gDW/hr) of the nucleosides (-10 mmol/gDW/hr) as done in vivo. Design and Assembly of CRISPRi knockdown vectors A complete list of knockdown vectors assembled are listed in Table 2. pJMP2846 was used as is for an ‘empty vector’ (EV) control, as it contains the CRISPRi knockdown system, but the sgRNA does not contain the gene-targeting tag. Gene-targeting plasmids were constructed as described in Peters et al. 2019. In short, to target either gpmA or rpoC, a 20bp region located near the 5’ end of the ORF with BsaI generated sticky ends was constructed using annealed oligonucleotides synthesized by Integrated DNA Technologies and cloned into pJMP284638. Target sites were selected using a publicly available script found at https://github.com/traeki/sgrna_design downloaded on 119 October 9th, 2020. Constructed plasmids were then transferred into MR-1 using a Tn7- based tri-parental mating protocol along with pJMP1039, which expresses the transposase transfer system (pJMP1039)38. Matings were re-streaked on LB with Kan plates without DAP to select for growth of MR-1 containing either pJMP2846, pJMP2846-gpmA, or pJMP2846-rpoC. CRISPRi Knockdown Analysis Strains carrying the knockdown vectors were pre-grown in LB with Kan and incubated with shaking for 16 h at 30°C. Overnight cultures were serially diluted in LB from 10-1 to 10-8 in a 96-well plate. 3 μL of each dilution was plated on LB with Kan or LB with Kan and 10mM IPTG in triplicate. Plates were incubated at 30°C for between 18 and 24 h, colonies were then counted, and plates imaged. When testing growth media to overcome the fitness defect of the sgRNA-gpmA knockdown strain, LB plates were additionally supplemented with either 20mM D,L-lactate, 10 mM uridine, or a combination of D,L-lactate and uridine. Gene Deletion Gene deletion was conducted as described in Blomfield et al. (1991). In short, two approximately 500bp regions of homology located immediately upstream and downstream of gpmA were cloned into the nonreplicating vector pSMV3.056. This vector, containing the sucrose sensitivity gene sacB, was inserted into the chromosome at the site of gpmA. Cells were pre-grown in LB with Kan, then subcultured into LB without NaCl. After growing for 16 hours, cells were plated on LB without NaCl and with 10% sucrose was used to select cells that had resolved the vector. Resulting colonies were screened via patch plating onto LB with Kan plates then onto LB plates to screen 120 for colonies that lost resistance associated with pSMV3.0. Colonies that were kanamycin sensitive were screened via colony PCR for sizes corresponding to ∆gpmA (1 kB) or WT revertants (2.7 kB) using primers flanking the gpmA locus. Suspected mutants were then confirmed via Sanger Sequencing. 4.6 Acknowledgements We thank Dr. Jason Peters and Dr. Amy Banta for providing the Mobile-CRISPRi knockdown system and helpful experimental suggestions. This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant No. (DGE-1848739) to K.C.F. and the 2018 Beckman Young Investigator Award to M.T. This work was supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences, under Award Number DE-SC0018269 (J.A.M.K, Y.S.-H.). This work was supported, in part, by the NSF Research Traineeship Program (Grant DGE-1828149) to J.A.M.K. This publication was also made possible by a predoctoral training award to J.A.M.K. from Grant T32-GM110523 from National Institute of General Medical Sciences (NIGMS) of the NIH. Its contents are solely the responsibility of the authors and do not necessarily represent the official views of the NIGMS or NIH. 4.7 Author Contributions K.C.F. and M.T. devised the project. K.C.F. planned and carried out the wet lab experiments. J.A.M.K. designed the computational framework for Flux Balance Analysis with conceptual ideas from Y.S.-H., and the modeling was carried out by K.C.F. K.C.F. and M.T. wrote the manuscript in consultation with J.A.M.K. and Y.S.-H. 121 REFERENCES (1) Xu, J.-Z., and Zhang, W.-G. (2016) Strategies used for genetically modifying bacterial genome: site-directed mutagenesis, gene inactivation, and gene over- expression *. J Zhejiang Univ-Sci B (Biomed Biotechnol) 17, 83–99. (2) Zeaiter, Z., Mapelli, F., Crotti, E., and Borin, S. (2018) Methods for the genetic manipulation of marine bacteria. Electron. J. Biotechnol. 33, 17–28. (3) Bertels, L. K., Murillo, L. F., and Heinisch, J. J. 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TerAvest1* 1 Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI, 48824, USA 2 Department of Microbiology and Molecular Genetics, Michigan State University, East Lansing, MI, 48824, USA 3 Department of Plant Biology, Michigan State University, East Lansing, MI, 48824, USA 128 5.1 Introduction The global effects of anthropogenic carbon emissions include damage to human and animal health, increased economic burden, and ecosystem destruction1–5. As efforts to phase out fossil fuels entirely becomes increasingly polarizing, researchers aim to develop more sustainable designs for existing infrastructure. One such effort relies on carbon capture, which would remove greenhouse gases like carbon dioxide (CO2) from the atmosphere. Other methods rely on biotic generation of fuel products like ethanol. Biofuel usage would eliminate the need for fossil fuel extraction, itself an extremely pollutive process, while producing fuels that can be used in existing infrastructure. Early developments towards biofuels used ethanol derived from plants like switchgrass and corn. While plant-based biofuels have demonstrated success and already been deployed for commercial use, they can present problems related to land management and food resource availability. Therefore, our research aims to circumvent this problem by using the electroactive bacterium Shewanella oneidensis MR-1 to develop a platform for microbial electrosynthesis (MES). MES is a burgeoning biotechnology that uses microorganisms to convert electricity and CO2 into useful chemical products like biofuels; this process can also be expanded to produce bioplastics, pharmaceuticals, and other specialty chemicals (Figure 5.1)6–9. However, to create this carbon-neutral platform using S. oneidensis MR-1, we must optimize the electron uptake pathway and install a carbon fixation module in this heterotrophic microbe. Work described in Chapters 2 and 3 focus on the former, while Chapter 4 lays out the initial steps taken towards engineering a strain of S. oneidensis MR-1 to be autotrophic. Here, I will describe the rationale behind our design strategy, what steps have been taken, and the future direction of this research. 129 Figure 5.1 Electroautotrophic Strategy. To engineer an electroautotrophic metabolism in S. oneidensis, we created a strain that requires separate carbon sources to build biomass and gain energy and reducing power (∆gpmA). This strain will be used to evolve a CO2 fixation module for building biomass and be grown on a cathode to use electron uptake for NADH production. This strain will use solely electricity and CO2 for growth. We can then implement an engineered bioproduction pathway, to produce useful chemicals from these minimal feedstocks. Our design for engineering autotrophy in S. oneidensis was inspired by work from Antonovsky et al. and the subsequent related publications10–15. They expressed the genes encoding ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO), phosphoribulokinase (prkA), and a carbonic anhydrase (CA) in Escherichia coli from a vector (pCBB). These, together with the native E. coli enzymes, created an artificial Calvin-Benson-Bassham (CBB) cycle. RuBisCO and PrkA bridge the gap between ribulose 5-phosphate and 3-phosphoglycerate, while CA, an enzyme commonly found in carboxysomes alongside RuBisCO, could concentrate CO2 in the cytosol (Figure 5.2). 130 Overexpression of these three genes was not sufficient to allow growth using solely CO2 to build biomass, therefore the team utilized metabolic modeling to generate a strategy for deleting endogenous genes to force reliance on RuBisCO for growth. Results from modeling led them to generate a knockout of the two genes that each encode a 3- phosphoglycerate mutase (gpmAM). This deletion severed the flow of carbon between the artificial CBB cycle and the tricarboxylic acid (TCA) cycle, requiring this strain to be supplied with two carbon substrates. The ∆gpmAM pCBB strain was evolved in a chemostat, in medium supplemented with excess carbon to feed the TCA cycle (pyruvate) and limiting substrate for biomass synthesis (xylose) in a 10% CO2 atmosphere. This way, strains that develop the ability to use RuBisCO and CO2 to build biomass will have a fitness advantage over those that do not, and become the dominant phenotype in the community. This approach was successful in generating a strain of E. coli capable of growth with CO2 as the sole substrate for biomass assimilation. We adapted this strategy to engineering autotrophic S. oneidensis. We were provided with the pCBB plasmid from the Milo Research Group and introduced it into WT S. oneidensis. Similar to results in E. coli, adding pCBB did not initially create an autotrophic phenotype in S. oneidensis. Proceeding forward, we applied the OptKnock program to an existing model of S. oneidensis metabolism to determine the best gene deletion strategy for our organism and the optimal mutant was generated16. The next steps of this research will focus on further developing this strain, including characterizing substrate usage, directed evolution to carbon fixation, and analysis of the acquired mutations in these lineages. 131 Figure 5.2 S. oneidensis central carbon metabolism amended with RuBisCO and PrkA. A metabolic map of central carbon metabolism, including the TCA cycle and an engineered Calvin-Benson Cycle. Solid lines represent enzymatic reactions native to S. oneidensis, dashed represent exogenous proteins, RuBisCO and PrkA, that are encoded by genes on pCBB. Adapted from Chapter 4 Figure 4.1. 5.2 Results 5.2.1 Determining Optimal Gene Knockout Strategy Despite S. oneidensis being a γ-proteobacteria and close relative to E. coli, our aim was to explore novel gene deletion strategies in silico for S. oneidensis, rather than 132 duplicating what has been done in E. coli. To do this analysis, we utilized a MATLAB program called OptKnock17. As previously described in Chapter 4, this program allows us to examine the flux of carbon within central metabolism and how loss of different enzymatic reactions affects this flow. We applied this to a previously published model of S. oneidensis and set the parameters to optimize flux though the RuBisCO reaction, allow up to two deletions in a limited set of target reactions, and growth on nine different substrates and CO218. Figure 5.3 Example for Determining Flux Coupling Slope. For each gene deletion strategy, single and double knockouts, a graph was generated that compared the growth potential of the mutant to WT when growing on different substrates. This is an example for ∆gapA growing on inosine. When WT is growing on inosine, it can reach its maximum growth rate potential (0.4 1/h) without no carbon flux through (and therefore use of) RuBisCO (0.4, 0). Conversely, the ∆gapA mutant can only have a maximum growth rate up to 0.15 1/h without RuBisCO; any higher growth rate will require the cell to use RuBisCO (green line = Flux Coupling Slope). When deciding which gene knockouts were best suited for our needs, we looked for mutations that showed a > 0 value for RuBisCO flux for any given growth rate. 133 This output determined whether or not a specific mutant could feasibly grow on a specific substrate with CO2, whether that growth would be dependent on the use of RuBisCO, and how much carbon would be fixed by RuBisCO, calculated as moles of CO2 fixed per gram of cell weight per hour (mmol CO2/gCWh) (Figure 5.3). This analysis was used to calculate the flux coupling slope (mmol CO2/gCW) for each potential mutant, and strains capable of growth without RuBisCO were ruled out (Figure 5.4). From the strains that were dependent on RuBisCO, the mutant that had the highest growth potential for the greatest number of carbon sources was a gpmA knockout (annotated in program as PGM). This meant that the optimal strategy for generating a strain of S. oneidensis that was capable of building biomass via carbon fixation was ∆gpmA expressing pCBB, the same strategy used for E. coli. However, unlike in E. coli, the enzymatic reaction carried out by GpmA was considered essential for S. oneidensis. This was based on both computational modeling and failure to generate a transposon insertion mutant during library preparation18. Though initial attempts to delete this gene were unsuccessful, we developed a genetic engineering pipeline to eventually generate ∆gpmA pCBB. Details for this are outlined in Chapter 4. 134 Figure 5.4 Results of Flux Balance Analysis. To identify a gene knockout strategy for optimizing flux of carbon through the RuBisCO reaction, we applied the OptKnock program to an existing metabolic map of S. oneidensis. We limited the scope of the program to only allow up to two knockouts of 27 central metabolic reactions. Intersections represent the two potential knockouts. Some reactions represent multiple genes that encode homologous proteins. The gradient represents the flux coupling slope in (mM CO2)/gCW (grams of cell weight), which means the amount of carbon that will flow through RuBisCO in a mutant that will only grow if RuBisCO is present. Within each box, are the flux coupling slope for a given knockout strategy grown on one of 9 carbon substrates 9 (inset) with 10% atmospheric CO2. See Abbreviations Table for the enzymes associated with these Enzymatic Reactions. 5.2.2 Investigating Growth Potential of ∆gpmA pCBB Once we created the ∆gpmA pCBB, we characterized its growth potential on various carbon substrates. Because this mutation severs the flow of carbon between the energy module (TCA) from the biomass module (gluconeogenesis/CBB), we hypothesized that this strain would need a minimum of two carbon substrates to grow. We compared growth of this strain in LB and M5 minimal medium lacking casamino acids supplemented with various combinations of carbon sources that either enter metabolism though the TCA cycle (pyruvate, D,L-lactate) or through glycolysis (adenosine, inosine, 135 uridine)19. These were compared to WT and WT expressing pCBB. Indeed, we observed that the OD600 of ∆gpmA pCBB only increased during growth in LB, or with two carbon sources, while WT and WT pCBB grew under all conditions (Figure 5.5). The most robust growth of ∆gpmA pCBB was observed using a combination of uridine and lactate, so these substrates were used in subsequent evolution experiments. The next step was to confirm that growth was due to consumption of both carbon substrates, as opposed to regulatory changes or other unintended effects. WT, WT pCBB, and ∆gpmA pCBB cells were grown in medium with either 20 mM D,L-lactate, 10 mM uridine, or both, and samples were taken for HPLC analysis at inoculation and when cultures reached maximum OD600. When both substrates are provided, WT and WT pCBB consumed all provided D,L-lactate and uridine, while ∆gpmA pCBB consumed about ~25% and ~35% respectively (Figure 5.6). This confirmed that both substrates were consumed by ∆gpmA pCBB, and the decreased assimilation accounts for the lower maximum cell density. In the single carbon medium, both WT strains consumed all of the provided carbon, while ∆gpmA pCBB consumed none of the D,L-lactate, and about ~25% of the uridine. It is likely that the small amount of uridine consumed went towards cellular maintenance and not biomass because there was no appreciable increase in OD600. 136 Figure 5.5 Growth of S. oneidensis Variants on Different Substrates. WT MR-1, WT pCBB, and ∆gpmA pCBB was grown overnight in LB (+Chl for strains with pCBB). Cells were washed and resuspended in 200 µL fresh LB, or M5 with one or two carbon substrates in a 96-well plate in triplicate (n=3). Medi key: LB = Lysogeny Broth, A = 10 mM adenosine, I = 10 mM inosine, U = 10 mM uridine, L = 20 mM D,L-lactate, and P = 20 mM pyruvate. Dashed line represents OD600 at inoculation. ∆gpmA pCBB only increased in OD in LB, or when 2 carbon substrates were provided. 137 Figure 5.6 Carbon Consumption by Strain. WT, WT pCBB, and ∆gpmA pCBB were pre-grown in LB (+Chl for pCBB), washed, and resuspended in M5 minimal medium with no casamino acids, and either 20 mM lactate, 10 mM uridine, or both. Samples were taken at inoculation, and after cultures reached a max OD for HPLC analysis. Bars and error bars represent mean and standard error, respectively (n=3). When provided with both carbon sources, all strains consumed both substrates, in their entirety in the case of WT strains (A,B). When provided only lactate, ∆gpmA pCBB did not consume any, while WT and WT pCBB consumed it all (C). When provided only uridine, ∆gpmA pCBB consumed ~25% and WT and WT pCBB consumed it all (D). ∆gpmA pCBB may have consumed this small amount of uridine for cellular maintenance, but was unable to grow to an appreciable OD600 (Figure 5.5). 138 5.2.3 Designing a Directed Evolution Experiment For chemostat evolution experiments, the medium is designed to ensure there is a fitness advantage for cells containing mutations that increase their use of RuBisCO. In ∆gpmA pCBB, D,L-lactate is taken up into the cell and converted into pyruvate and subsequently enters the TCA cycle. Providing D,L-lactate in excess ensures that the cells have ample reserves of NADH and ATP. Uridine is used to build biomass, and therefore will be the limiting factor for growth. This design ensures that ∆gpmA pCBB cannot reach its maximum growth rate, and that there is a selective pressure to utilize the other available carbon source, CO2. Cells that use CO2 will increase in growth rate and become the dominant phenotype in the culture. The amount of uridine needed to sustain growth under limiting conditions was determined by calculating the substrate concentration constant (Ks). This value represents the concentration of the limiting substrate, in this case uridine, at half the maximum growth rate (µmax) of a given strain when the limiting carbon is in excess. To determine this, ∆gpmA pCBB was grown in M5 minimal medium with no casamino acids, 20 mM D,L-lactate, and varying concentrations of uridine. The maximum growth rate for each version of the medium was calculated and plotted as a factor of uridine concentration (Figure 5.7). Applying the Monod equation to these points, µmax is calculated to be 0.1386 1/h, µmax/2 is 0.069 1/h, and Ks is 64.95 mg/L of uridine. Knowing these values, chemostat evolution experiments to develop a strain of ∆gpmA pCBB that grows by fixing CO2 will use medium with 20 mM D,L-lactate and 65 mg/L of uridine (~0.26 mM). 139 Figure 5.7 Growth Rate of ∆gpmA pCBB on Varying [uridine]. ∆gpmA pCBB cells were pre-grown in LB+Chl, washed, and resuspended in 2 mL of M5 without casamino acids, with 20 mM lactate and various concentrations of uridine, in a 24-well plate. OD600 was measured every 15 minutes until a maximum OD600 was reached for all wells. The max growth rate (1/h) was calculated and plotted against [uridine] g/L. Points and error bars represent average and standard error, respectively (n=3). Red line represents the calculated Monod Curve for this data, with µmax being the maximum potential growth rate, µmax/2 being half the maximum potential growth rate, and Ks being the [uridine] at µmax/2. 5.3 Future Directions From this work, we have everything needed to begin directed evolution to create a strain of S. oneidensis capable of fixing CO2 for biomass. Evolution experiments with ∆gpmA pCBB will be carried out using a BIOSTAT-A Fermenter, with an excess of D,L- lactate (20 mM) and limiting uridine (65 mg/L). The culture vessel will be regularly sampled to track substrate usage via HPLC, cell growth (OD600), and ensure no 140 contamination. At the onset of the experiment, we expect the culture to grow denser with time before reaching a steady state OD600 where all uridine is being consumed, but there is still excess D,L-lactate in the medium. Mutations will naturally occur within the cells, and any mutations that aid in fitness will be retained in subsequent generations. One such path for increasing fitness is being able to use the other available carbon source (CO2) to build biomass. Consequently, genotypes that express and use RuBisCO and PrkA to fix carbon will outcompete others that do not. Because these cells use the CO2 and uridine, they will simultaneously increase their consumption of D,L- lactate. Therefore, we can infer the evolution of the desired phenotype when there is an increase in OD600 and decrease in residual D,L-lactate from steady state. When the culture has reached a new, higher steady state OD600 where all D,L-lactate is consumed, samples will be taken to identify isolates capable of growth using solely CO2 and D,L- lactate. If multiple isolates with the CO2 fixing phenotype are identified, it is unlikely that they will all have the same acquired mutations (genotype). Therefore, the next step is to perform whole genome sequencing on various isolates. This genomic data will provide insight regarding how the metabolome was fine-tuned to redirect the flow of carbon through RuBisCO. Additionally, we will observe any overlap in mutations between isolates, indicative of that specific mutation being a greater influence. The information can also be used, as in Herz et al, to build a version of this strain with the smallest number of required mutations11. To do this engineering, the acquired mutations of different isolates will be pooled and ranked based on frequency and proximity to carbon 141 metabolism. They would then be systematically reintroduced into an unevolved ∆gpmA pCBB until the phenotype is restored. As this engineered strain will still require D,L-lactate to grow, an important follow up to this work would be to ensure that the D,L-lactate is being solely used for energy acquisition and not to build biomass. This assessment could be done using isotopically labeled CO2 and D,L-lactate to track which substrate is the building block for the synthesis of metabolites or cell components. This analysis will be crucial to using this strain for electroautotrophy. The intent with this design is to eventually replace the secondary carbon source (D,L-lactate) with the cathode. Therefore, we must ensure that the cell is capable of growth using solely CO2 for biomass synthesis, as the cathode will only facilitate NADH and ATP formation. Once autotrophy is confirmed, the bioengineered strain can be applied as a biocatalyst for a wide variety of MES applications. 5.4 Materials and Methods Strains and Plasmids Strains and plasmids used are listed in Table 5.1. S. oneidensis strains were grown at 30 °C and shaking at 275 rpm for aerobic growth. For growth curves, cells were pre- grown aerobically in 5 mL of lysogeny broth (LB) supplemented with 50 μg/mL chloramphenicol (Chl) for strains with pCBB, for inoculating minimal medium. For growth curves, cells were grown in M5 minimal medium containing: 1.29 mM K2HPO4, 1.65 mM KH2PO4, 7.87 mM NaCl, 1.70 mM NH4SO4, 475 μM MgSO4·7 H2O, 10 mM HEPES, 0.01% (w/v) casamino acids, 1× Wolfe’s vitamin solution, and 1× Wolfe’s mineral solution, then the pH adjusted to 7.2 with 5 M NaOH. After autoclaving, carbon 142 substrates were added depending on the experiment to final concentrations of: 20 mM D,L-lactate, 20 mM pyruvate, 10 mM uridine, 10 mM adenosine, or 10 mM inosine. Growth curves were performed in an aerobic plate reader (BioTek), grown in 2 mL of M5 (with appropriate carbon) in a 24-well plate. These were done in triplicate. Table 5.1 Strains and Plasmids used in this study Strain or Description Source Plasmid S. oneidensis MR-1 Wild type S. oneidensis Meyers and Nealson, 198820 ∆gpmA Mutant with gene deletion of SO_0049 This work Plasmids pCBB pZA11 vector expressing RuBisCO, prkA, and Antonovsky et al. CA, ChlR 201710 HPLC analysis HPLC analysis was performed as previously described (Tefft and TerAvest, 2019) with the amendments described in (Tefft et al., 2022)9,10. Sample analysis was performed on a Shimadzu 20A HPLC, using an Aminex HPX-87H (BioRad, Hercules, CA) column with a Microguard Cation H+ guard column (BioRad, Hercules, CA) at 65 °C with a 0.5 ml/min flow rate. Data analysis Analysis of HPLC data, OD, current data, and growth curve data was done using RStudio using the following packages: ggplot2, dplyr, ggpubr, plyr, data.table, stringr, , and growthcurver61–67. Flux Balance Analysis We used the genome-scale model of the metabolic network of S. oneidensis MR-1 from Ong et al54. The ATP growth-associated maintenance (GAM) cost was removed from 143 this version of the MR-1 model due to a lack of GAM measurements for other S. oneidensis strains that were being compared with MR-1. The GAM was added back in by referring to the information from an earlier iteration of the model41. All optimizations were performed in MATLAB (version 2019a) using the COBRA Toolbox55 and version 8.1.1 of the Gurobi optimizer40. To model carbon utilization, specific uptake rates were set for each substrate (inosine, L-lactate, D-lactate, adenosine, uridine, acetate, N- acetyl glucosamine (NAG), propionate, pyruvate) in equal amounts. Using the OptKnock program, the number of potential mutations was limited to a maximum of two from a designated pool of enzymatic reactions, see Table 5.2. Reactions for RuBisCO and PrkA were added to the model, and RuBisCO was the reaction targeted for optimizing carbon flux. For each potential mutant and substrate combination, it was determined whether the growth was possible, and if so if it was non-linked (non-growth coupled), linked (growth coupled no unique), or dependent (growth coupled) on flux through RuBisCO. Additionally, we calculated the flux couple slope (mmol CO2/gDW) for each mutant and substrate combination from the output, which is represented in the heatmap (Figure 5.4). Table 5.2 Reactions Targeted as Potential Deletions18 Enzymatic Reaction Enzyme Reaction Locus acetaldehyde acetaldehyde + CoA+ NAD+ -> ACALDI dehydrogenase acetoacetyl-CoA + H+ + NADH (SO2136) (SO0432) or ACONT aconitase citrate <=> isocitrate (SO0343) alcohol dehydrogenase EtOH + NAD+ <=> ALCD2X (ethanol: NAD+) acetaldehyde + H+ + NADH (SO2136) acetoacetyl-CoA + H2O + oxaloacetate -> citrate + CoA+ CS citrate synthase H+ (SO1926) 144 Table 5.2 (cont’d) 2-dehydro-3-deoxy- 2-dehydro-3-deoxy-D-gluconate phosphogluconate 6-phosphate -> glyceraldehyde EDA aldolase 3-phosphate + pyruvate (SO2486) D-glycerate 2-phosphate <=> ENO enolase H2O + phosphoenolpyruvate (SO3440) D-fructose 1,6-bisphosphate fructose- <=> dihydroxyacetone bisphosphate phosphate + FBA aldolase glyceraldehyde 3-phosphate (SO0933) D-fructose 1,6-bisphosphate + fructose- H2O -> D-fructose 6-phosphate FBP bisphosphatase + Pi (SO3991) D-glucose 6-phosphate + NADP+ <=> 6-phospho-D- glucose 6-phosphate glucono-1,5-lactone + H+ + G6PDHY dehydrogenase NADPH (SO2489) glyceraldehyde-3- glyceraldehyde 3-phosphate + phosphate NAD+ + Pi <=> 3-Phospho-D- (SO0538) or dehydrogenase glyceroyl phosphate + H+ + (SO2345) or GAPD (NAD+) NADH (SO2347) ATP + D-glucose -> ADP + D- HEX1 hexokinase glucose 6-phosphate + H+ (SO1656) (SO0424, pyruvate CoA+ NAD+ + pyruvate-> SO0425, PDH dehydrogenase acetoacetyl-CoA+ CO2 + NADH SO0426) glucose-6-phosphate D-glucose-6-phospate <=> D- PGI isomerase fructose-6-phospate (SO3547) 3-phospho-D-glyceroyl phosphoglycerate phosphate + ADP <=> 3- PGK kinase phospho-D-glycerate + ATP (SO0932) phosphoglycerate 3-phospho-D-glycerate <=> PGM mutase D-glycerate 2-phosphate (SO0049) 6-phospho-D-gluconate + phosphogluconate NADP+ -> CO2 + NADPH + PGDH dehydrogenase D-ribulose 5-phosphate (SO1902) Formate C- CoA + pyruvate <=> (SO2912, PFL acetyltransferase acetoacetyl-CoA + formate SO2913) ADP + H+ + phosphoenolpyruvate -> ATP + PYK pyruvate kinase pyruvate (SO2491) ATP + H2O + pyruvate -> AMP phosphoenolpyruvate + 2 H+ + phosphoenolpyruvate PPS synthase + Pi (SO2644) 145 Table 5.2 (cont’d) CO2 + H2O + phosphoenolpyruvate phosphoenolpyruvate -> H+ + PPC carboxylase oxaloacetate + Pi (SO0274) phosphoenolpyruvate ATP + oxaloacetate -> ADP + PPCK carboxykinase CO2 + phosphoenolpyruvate (SO0162) ribulose 5-phosphate D-ribulose 5-phosphate <=> RPE 3-epimerase D-xylulose 5-phosphate (SO0292) ribose-5-phosphate α-D-ribose 5-phosphate <=> RPI isomerase D-ribulose 5-phosphate (SO1150) glyceraldehyde 3-phosphate + sedoheptulose 7-phosphate <=> D-erythrose 4-phosphate + D- TAL transaldolase fructose 6-phosphate (SO3546) α-D-ribose 5-phosphate + D- xylulose 5-phosphate <=> glyceraldehyde 3-phosphate + TKT1 transketolase sedoheptulose 7-phosphate (SO0930) D-erythrose 4-phosphate + D- xylulose 5-phosphate <=> D-fructose 6-phosphate + TKT2 transketolase glyceraldehyde 3-phosphate (SO0930) dihydroxyacetone phosphate triose-phosphate <=> glyceraldehyde 3- TPI isomerase phosphate (SO1200) 146 REFERENCES (1) Lenoir, J., and Svenning, J. 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The work presented here contributes to this effort by employing an enzymatic system (Bdh) for reassessing the physiology energetics of inward electron transfer (Chapter 2 and 3),and using novel synthetic biology designs to implement an artificial carbon fixation pathway in S. oneidensis (Chapter 4 and 5). 6.1.1 Bidirectional Electron Transfer to Oxygen and NADH Previous work published by our research group demonstrated that a proton and electrochemical gradient must be maintained across the inner membrane of S. oneidensis to sustain NADH generation from a cathode 1,2. Through our investigation into sustained 2,3-butanediol production in S. oneidensis cells lacking the exogenous proton-pump proteorhodopsin (PR), we determined this was the result of coupling NADH generation to O2 reduction. This bidirectional electron transfer (to O2 and NAD+) maintained the energetic balance needed to restore PMF via oxidases as it was being consumed by NADH dehydrogenases (NDHs). We first established this by confirming that S. oneidensis expressing butanediol dehydrogenase (Bdh) could reduce acetoin to 2,3-butanediol at a steady rate in a bioelectrochemical system (BES) being sparged with N2. To investigate if bidirectional electron transfer was occurring under these conditions, we used a dissolved oxygen (DO) probe to determine the concentration of DO in the working chamber inoculated with S. oneidensis. We showed that despite N2 150 sparging, there was ~ 1% DO (2 µM) present throughout the course of the experiment; the environment is microaerobic and not anaerobic. Reduction of one molecule of O2 will translocate four protons across the inner membrane into the periplasm via S. oneidensis’s native oxidase (Cco, Cox). Similarly, four protons moving down the gradient into the cytoplasm are required to produce one NADH. Therefore, ~1% DO is more than sufficient to sustain the rate of 2,3-butanediol production that we observed. To confirm that the cells were using DO, we used a strain that is unable to reduce O2 (∆cco∆cyd∆cox pBDH). This strain showed a ~90% decrease in 2,3-butanediol accumulation compared to WT, and could be functionally complemented by expressing PR. Similarly, the current in ∆cco∆cyd∆cox pBDH decreased to approximately the same current observed in sterile BES, and was partially rescued by PR. Together, these results showed that S. oneidensis cells were using cathode-derived electrons to couple NADH generation to O2 reduction in the microaerobic BES. Interestingly, increasing DO via passive aeration negatively impacted 2,3- butanediol production. We determined this effect was due to the formation of reactive oxygen species (ROS) via interactions of O2 with the cathode or reduced flavins. This ROS formation killed ~2.5 log10 cells within three hours of the onset of cathodic potential. Addition of catalase prior to cathodic potential reduced formation of H2O2 and mitigated the severity of cell death. We concluded that a microaerobic environment is optimal for bidirectional electron transfer for NADH generation under these conditions, as high O2 levels have cytotoxic effects. There is room for continued optimization via modulation of DO %, as well as addition of other ROS-neutralizing enzymes such as superoxide dismutase. This will also be important for MES work that aims to operate in 151 an anaerobic environment. Careful attention must be paid to BES design to prevent oxygen intrusion as simply sparging with N2 is insufficient to maintain true anaerobicity. 6.1.2 Flexibility of the EET Pathway The EET pathway of S. oneidensis is a complex network of cytochromes, flavins and flavin associated proteins, that is heavily influenced by the environment.3,4 Attempts to improve MES in S. oneidensis by increasing EET have heavily relied on electrical output to determine what is helpful or detrimental to this cause. However, this approach is susceptible to abiotic interference that can obscure what is happening biotically. Therefore, we reassessed the role of major EET pathway components during inward electron transfer. By expressing Bdh and adding acetoin, we can measure the 2,3- butanediol produced via acetoin reduction with NADH as a proxy for electron uptake to form NADH. This is less likely to be impacted by abiotic reactions than electrochemical measurements. Additionally, this approach will allow us to determine if there are any potential electron sinks that are diverting the flow of electrons away from the targeted reaction. We first assessed EET via the Mtr pathway by comparing 2,3-butanediol accumulation between WT and ∆mtrA. This mutant showed a ~95% decrease in production and had cathodic current as low as abiotic BESs, confirming previous reports showing MtrCAB as essential for EET3. Similarly, results from a cymA deletion mutant were in alignment with the literature. The ∆cymA strain showed a decrease but not complete loss of both current and 2,3-butanediol. Recent reports have shown evidence for a CymA-independent pathway for electrons to enter the quinol pool from Mtr/periplasmic shuttles, and this work supports those findings; CymA is likely the 152 primary inner membrane electron hub, but other proteins can partially compensate its role5–7. We next examined the role of exogenous and endogenous flavins during EET. A strain unable to export flavins (∆bfe) showed a significant decrease in current and 2,3- butanediol accumulation. Somewhat surprisingly, the phenotype for 2,3-butanediol production could not be complemented with exogenous flavin mononucleotide (FMN) or riboflavin (RF). However, we showed that exogenous flavins did increase the cathodic current. We hypothesize that this was due to abiotic reactions of free flavins with the cathode and photooxidation of the flavins. The discrepancy between current generation and bioproduction highlights the need to use quantifiable, biological outputs and not only current to assess the efficiency of EET, especially because the impact of flavins on EET has been debated. There is evidence suggesting flavins mediate EET over long distances as free flavins, aid in direct electron transfer as outer membrane cytochrome (OMC) cofactors, and influence regulation of biofilms and pili8–10. We investigated the latter by pre-growing ∆bfe cells with exogenous flavins, but saw similar results as without, suggesting they do not greatly alter the proteome under these conditions. Because our BES operates with stationary phase cells with no substrate, we cannot entirely rule out regulation being important. Additionally, this environment may also impact OMC binding or flavin diffusion through the biofilm. Finally, we showed evidence that the native hydrogenases (Hyd, Hya) are important for increasing cell survival during stationary phase, in agreement with the literature11. We previously showed that deletion of the hydrogenases improved inward electron transfer to 2,3-butanediol. However, after modifying the experimental protocol 153 to remove residual organic matter, we saw their absence caused a decrease in production. In the initial design, the hydrogenases acted as an electron sink, siphoning electrons away from NADH, whereas in the absence of carbon they aid in survival, potentially by modulating the redox state of the quinol pool. 6.1.3 Synthetic Biology Strategies S. oneidensis is a genetically tractable organism, so there are a wide range of bioengineering techniques that can be applied for improving performance in MES. We developed a pipeline that combines existing techniques from E. coli and in silico modeling to generate a novel mutation in a conditionally essential gene, gpmA12–14. This mutant was selected as a starting point for evolution of a carbon fixation pathway in S. oneidensis as outlined in Chapter 515–17. The gpmA gene was considered essential based on metabolic modeling and failure to isolate a transposon insertion at this site during library preparation18. This was believed because GpmA connects the flow of carbon between gluconeogenesis and the tricarboxylic acid (TCA) cycle, and without GpmA cells either cannot build biomass or make ATP when given a single carbon source. To overcome this lethal phenotype, we used flux balance analysis to examine the growth potential of ∆gpmA with different carbon substrates. The results of the modeling were compared to phenotypes of a gpmA knockdown strain grown with different substrates. We determined that providing the cells with two substrates, one to build biomass and one to gain energy, restores growth phenotype of a gpmA knockdown to WT. Therefore, including additional substrate in the medium during the gene deletion protocol allowed strains that retain the mutation to compete with WT 154 revertants, resulting in a 44% rate of success in isolating a ∆gpmA mutant. This design can be implemented to generate other difficult genetic modifications. 6.1.4 Engineering Carbon Fixation in S. oneidensis Finally, this dissertation outlines the initial steps we have taken towards using the ∆gpmA mutant as a basis for carbon fixation in S. oneidensis. Chapter 5 lays out how we used flux balance analysis to identify this mutant, as it showed the highest reliance on using CO2-fixing RuBisCO to grow. We then further characterized this strain in relation to WT, and WT expressing the carbon fixation genes (pCBB). This strain (∆gpmA pCBB) was unable to grow without two substrates, whereas WT with and without pCBB showed robust growth regardless of the carbon source. The mutant only consumed D,L-lactate as a substrate and increased in OD600 when uridine was also available. This information will be the basis for directed evolution to generate a CO2 fixing strain of S. oneidensis. 6.2 Future directions This dissertation lays out the framework for building a chassis for carbon-neutral biofuel production using S. oneidensis. However, there is still a long road from where we are now to industrial application. As discussed in Chapter 2, the presence of oxygen, even at low levels, can have a large impact on the efficiency of MES systems. We showed that bubbling the BES working chamber with N2 creates a microaerobic atmosphere, with enough oxygen to sustain bidirectional electron transport without generating ROS. However, as the DO level measured in the BES was a by-product of our design, there is room to improve this model. From an engineering perspective, more control over the DO concentrations can be achieved by using copper tubing instead of 155 Figure 6.1 Comparing NADH and NADPH as Redox Partners with Bdh. Measurements of 2,3-butanediol accumulation in WT MR-1 expressing either a Bdh from Enterobacter cloacae (pBDH-NADH) or Clostridium ljungdahlii DSM 13528 (pBDH- NADPH) from an IPTG inducible promoter. Strains were pre-grown in LB, washed and resuspended in M5 medium with 20mM lactate, 10 mM acetoin, and 100 µM IPTG at an OD600 of 0.01. Samples were taken for OD600 readings (data not shown) and HPLC analysis every hour for twelve hours, then a final time point at 23 hours. The 2,3- butanediol accumulation in MR-1 pBDH-NADPH rapidly outpaced MR-1 pBDH-NADH, converting almost 80% of the acetoin in 23 hours, compared to less than 25%. neoprene, crimping caps to tighten the stopper seals, and using specialized gas mixes (e.g., 1% O2: 99% N2). Additionally, the medium could be amended with oxygen scavengers (e.g., sodium sulfite, ascorbic acid) or other antioxidants (e.g., superoxide dismutase, N-acetyl cysteine). The S. oneidensis genome encodes three oxidases (Cyd, Cco, Cox) that are expressed under different conditions, have varying affinities for O2, and different mechanisms for proton translocation. To determine if there is a specific oxidase, or combination of two oxidases, that work best for bidirectional electron 156 transfer, we would compare the current and 2,3-butanediol formation of single (∆cyd, ∆cco, ∆cox) and double (∆cyd∆cco, ∆cyd∆cox, ∆cco∆cox) oxidase deletion mutants. Additionally, there is interest in using bidirectional electron transfer to generate NADPH, so we have made a vector that expresses a Bdh from Clostridium ljungdahlii DSM 13528 which uses NADPH instead of NADH. Preliminary data show that this version is functional in S. oneidensis and is more effective than the NADH version (Figure 6.1). If S. oneidensis can drive cathode-dependent 2,3-butanediol production with NADPH, the higher rate of activity of this enzyme could make it easier to discern smaller differences between experimental conditions. The EET network in S. oneidensis has been well characterized in the context of power density and current generation. In few cases has this information been directly translated into impacts in MES. Therefore, expanding on the work in Chapter 3 would entail examining how other EET proteins contribute to 2,3-butanediol production. This approach could be done through a combination of gene deletion or overexpression, such as the mutant designed by Sun et al. that showed significantly higher current (∆fccA∆napB∆tsdB pHGEN-Ptac-cctA)19. The Bdh system could be used to confirm the conclusions drawn for any mutant or engineered strain. Similarly, as we failed to rescue the phenotype of ∆bfe through exogenous flavins, there are still questions as to the scope of flavin influence on EET. Our data indicate that the origin of the flavins is crucial, which could point to a regulatory role. We ruled out a role for flavins to impact regulation during pre-growth, but their influence may not be enacted until contact with the cathode and by this time the cells have entered stationary phase and fail to shift their proteome. To test this hypothesis, we can compare the proteomes of cell 157 population at different stages of the experiment to look at differences in proteins known to be regulated by flavins, such as biofilm formation in WT versus ∆bfe and cytochrome expression profiles. Finally, to better understand the role of hydrogenases in EET, experiments should compare hydrogen formation during the experiment between WT and single and double hydrogenase knockouts (∆hydA, ∆hyaB, ∆hydA∆hyaB). The work described in Chapter 4 and Chapter 5 lays the foundation for engineering an electroautotrophic S. oneidensis. As previously described, the next steps will be to design and implement laboratory directed evolution using ∆gpmA pCBB, a 10% CO2 atmosphere, and medium supplemented with excess D,L-lactate and limiting uridine. This will be done in a BIOSTAT-A fermenter operating as a chemostat, with samples taken daily to track metabolite concentration, OD600, and check for contamination. Once the culture reaches a steady state, we expect the onset of a CO2- fixing phenotype to be indicated by a rise in OD600 and a decrease in D,L-lactate concentration. This outcome indicates that there is a population of cells that is using the other available carbon source (CO2) and will therefore be consuming more D,L-lactate and have a fitness advantage over cells that are not. Then, CO2 fixing CFUs can be isolated and characterized. The next steps would be to replace the reliance on D,L- lactate for ATP and NADH with a cathode. 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