NUTRIENT LIMITATION ALTERS METABOLISM, CR(VI) RESPONSE, AND BIOFILM MATRIX COMPOSITION IN DESULFOVIBRIO VULGARIS HILDENBOROUGH by Lauren Christine Franco A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Microbiology and Immunology MONTANA STATE UNIVERSITY Bozeman, Montana November 2017 ©COPYRIGHT by Lauren Christine Franco 2017 All Rights Reserved ii ACKNOWLEDGEMENTS Looking back on the last seven years of graduate school, I can’t help but think that I have been very fortunate to spend the better part of a decade living and studying in Montana surrounded by some of the best people I know. There were challenges, of course, but the support system that is the Center for Biofilm Engineering at MSU and all of the people that work there made it so that help was never hard to find. First, I must thank my advisor, Professor Matthew Fields, for being supportive, patient, and an all- around great mentor throughout this process. He has been a great example of leadership and professionalism, while still being approachable and kind. I would also like to thank my committee members, Christine Foreman, Brent Peyton, and Mike Franklin for their invaluable insight and input throughout my time here. After every meeting, I have felt grateful for their contributions and humbled by their intellect. I want to acknowledge that my experience as a graduate student would not have been nearly as successful and fulfilling without CBE friends and labmates past and present that have become like my family here -- Heidi, Kristen, James, Sara, Kara, Hannah, Luisa, Chiachi, Luke, Laura, Isaac, Katie, Anna, and Greg. Graduate school would not have been the same without all of your intellectual input and lessons in winter, outdoor activities, and life in general. Also, to the Molecular Biosciences Program and the Department of Energy for funding throughout the years. Lastly, I have to thank my family for being beacons of support from afar and letting me find my own path. iii TABLE OF CONTENTS 1. INTRODUCTION ...........................................................................................................1 Nutrient Limitation ..........................................................................................................1 Sulfate-Reducing Microorganisms ..................................................................................2 Applications of Sulfate Reducing Bacteria ..............................................................3 Desulfovibrio vulgaris Hildenborough ....................................................................6 Heavy Metal and Radionuclide Reduction in Desulfovibrio ...................................8 Cr(V) Reduction in D. vulgaris ...............................................................................9 Biofilm Physiology ........................................................................................................10 The Biofilm Matrix ................................................................................................11 Biofilm Resistance to Heavy Metal and Radionuclide Toxicity ...........................15 Physiology of D. vulgaris Biofilms .......................................................................17 Research Objectives ......................................................................................................18 References .....................................................................................................................21 2. Cr(VI) REDUCTION AND PHYSIOLOGICAL TOXICITY IS IMPACTED BY RESOURCE RATIO IN DESULFOVIBRIO VULGARIS HILDENBOROUGH ..................................................31 Contributions of Authors ...............................................................................................31 Manuscript Information .................................................................................................32 Abstract ..........................................................................................................................33 Introduction ....................................................................................................................34 Materials and Methods ...................................................................................................36 Strains and Growth Conditions ..............................................................................36 Experimental Design ..............................................................................................38 Cr(VI) Reduction Analysis ....................................................................................38 Analytical Techniques ...........................................................................................39 Mutant Generation .................................................................................................39 Results .............................................................................................................................41 Temperature Affects D. vulgaris Growth and Cr(VI) Reduction/Toxicity Under a Balanced Resource Ratio .........................................41 Resource Ratio Imbalance Affects D. vulgaris Cr(VI) Reduction and Tolerance ........................................................46 Electron Donor-Limitation (EDL) ....................................................................46 Electron Acceptor-Limitation (EAL)................................................................47 iv TABLE OF CONTENTS CONTINUED Viability .................................................................................................................48 Growth with Increased Sulfate ...............................................................................49 Growth with Normalized Sulfate Levels ...............................................................50 Sulfate Permease Mutants ......................................................................................51 Ascorbate ...............................................................................................................53 Discussion .......................................................................................................................54 References .......................................................................................................................61 Supplemental Figures .....................................................................................................66 3. NUTRIENT LIMITATION CAUSES DECLINE IN METABOLITES IMPORTANT FOR CELL CYCLE PROGRESSION IN BACTERIAL BIOFILM ...............................................................70 Contributions of Authors ................................................................................................70 Manuscript Information Page .........................................................................................71 Abstract ...........................................................................................................................72 Introduction .....................................................................................................................73 Materials and Methods ....................................................................................................75 Sample Collection ..................................................................................................76 Biomass Analyses ..................................................................................................76 Metabolomic Analysis of Biofilm .........................................................................76 Results .............................................................................................................................77 Biofilm Growth under Nutrient-Limited and Balanced Conditions ..............................................................................................77 Metabolomic Analysis of D. vulgaris Biofilm ......................................................79 Discussion .......................................................................................................................86 References .......................................................................................................................92 4. EXTRACELLULAR MEMBRANE STRUCTURES IN DESULFOVIBRIO VULGARIS HILDENBOROUGH BIOFILMS .............................95 Contributions of Authors ................................................................................................95 Manuscript Information ..................................................................................................97 Abstract ...........................................................................................................................98 Introduction .....................................................................................................................99 Materials and Methods ..................................................................................................101 v TABLE OF CONTENTS CONTINUED Bacterial Strains and Growth Conditions ............................................................101 Sample Collection ................................................................................................102 Biomass Analyses ................................................................................................102 Imaging Methods .................................................................................................103 Fluorescence Microscopy ..............................................................................103 High Pressure Freezing and Freeze Substitution ...........................................103 Transmission Electron Microscopy ...............................................................104 Volume Electron Microscopy ........................................................................104 Energy Dispersive Spectroscopy ...................................................................105 Biofilm Exposure to Cr(VI) .................................................................................105 Cytochrome Staining and Heme Quantification ..................................................106 Results ...........................................................................................................................107 2D TEM Imaging of Desulfovibrio vulgaris Biofilms ........................................107 Large Volume 3D SBF/SEM Imaging .................................................................111 Elemental Analysis of Extracellular Metal Deposits ...........................................113 Underlying Metal Deposition Sites are Biological Membranes ..........................115 The Effect of Nutrient Limitation on Cr(VI) Toxicity in D. vulgaris Biofilms ........................................................................................119 Discussion .....................................................................................................................122 References .....................................................................................................................129 5. OUTER MEMBRANE VESICLES AND ASSOCIATED PROTEINS PRODUCED BY DESULFOVIBRIO VULGARIS BIOFILMS ..............133 Contributions of Authors .............................................................................................133 Manuscript Information Page ......................................................................................134 Abstract .......................................................................................................................135 Introduction .................................................................................................................136 Materials and Methods ................................................................................................138 Bacterial Strains and Growth Conditions ............................................................138 Isolation of Outer Membrane Vesicles ................................................................139 Transmission Electron Microscopy .....................................................................140 Nucleic Acid Detection ........................................................................................140 Proteomic Analysis ..............................................................................................140 Results and Discussion ................................................................................................141 Outer Membrane Vesicles and Tubes ..................................................................141 Proteomic Analysis of OMVs ..............................................................................146 vi TABLE OF CONTENTS CONTINUED References ...................................................................................................................151 6. EPILOGUE ..................................................................................................................156 References ...................................................................................................................167 REFERENCES CITED ....................................................................................................170 APPENDICES .................................................................................................................197 APPENDIX A: Supplemental Figures ........................................................................198 Introduction ..........................................................................................................199 Materials and Methods .........................................................................................199 Results ..................................................................................................................200 APPENDIX B: Desulfovibrio carbinoliphilus Oakridgensis, Spp. Nov., an Organic Acid-Oxidizing, Sulfate-Reducing Bacterium Isolated from Uranium(VI)-Contaminated Groundwater ..........................202 Contributions of Authors .....................................................................................202 Manuscript Information Page ..............................................................................203 Abstract ................................................................................................................204 Introduction ..........................................................................................................205 Materials and Methods .........................................................................................206 Results and Discussion ........................................................................................211 References ............................................................................................................218 Supplemental Figures ...........................................................................................221 APPENDIX C: Autonomous Metabolomics for Rapid Metabolite Identification in Global Profiling .........................................................................227 Contributions of Authors .....................................................................................227 Manuscript Information Page ..............................................................................228 APPENDIX D: Comprehensive Bioimaging with Fluorinated Nanoparticles Using Breathable Liquids ......................................................................................................237 Contributions of Authors .....................................................................................237 Manuscript Information Page ..............................................................................238 vii LIST OF TABLES Table Page 1. Strains used in this study ...................................................................................67 2. Primers used in this study ..................................................................................67 3. Lactate and Sulfate concentrations for the different nutrient ratios and biomass yields based on lactate and sulfate consumption ...................................................................79 4. List of metabolites down-regulated under the EAL condition ...........................81 5. List of metabolites up-regulated under the EAL condition ...............................83 6. Proteins identified in purified OMV samples ..................................................144 7. Summary of substrate and electron acceptor utilization of FW-101 isolate and closely related strains ................................216 viii LIST OF FIGURES Figure Page 1. Growth of D. vulgaris at 20 and 30°C with 0 and 50µM Cr(VI) under EAL, BAL, and EDL ......................................................44 2. Cr(VI) reduction at 20 and 30°C with 100 and 50µM Cr(VI) under EAL, BAL, and EDL ......................................................45 3. Growth with 50µM Cr(VI) and increasing sulfate concentrations .......................................................................................50 4. Growth with 50µM Cr(VI) and normalized sulfate concentrations .......................................................................................51 5. Growth of sulfate permease mutants with 0 and 50 µM Cr(VI) .........................................................................................53 6. Growth with 50µM Cr(VI) and ascorbate ........................................................54 7. Schematic of EAL, BAL, and EDL scenarios .................................................59 8. Lactate, acetate, sulfate concentrations over growth with 0 and 50µM Cr(VI) ......................................................................66 9. Protein, carbohydrate, and sulfide concentrations in biofilms grown under EAL, BAL, and EDL ...............................................78 10. Cloud plot of dysregulated metabolites ...........................................................80 11. Pathways down-regulated under EAL .............................................................84 12. Large-scale microscopic analysis of biofilm Grown under EAL, BAL, and EDL ...............................................................108 13. TEM images of extracellular structures .........................................................110 14. 3D reconstruction of extracellular membrane structures ...............................112 ix LIST OF FIGURES CONTINUED Figure Page 15. 3D images of 40 x 40 x 100µm biofilm with and without cells ....................113 16. EDS elemental mapping of metal deposits ....................................................114 17. Differential staining of extracellular membrane structures ...........................116 18. FAME content of biofilm ...............................................................................118 19. Viability in EAL, BAL, EDL biofilm ............................................................118 20. Planktonic growth of scraped biofilm with Cr(VI) ........................................121 21. Protein gel with cytochrome stain ..................................................................122 22. TEM images of isolated OMVs .....................................................................143 23. SEM images of biofilm grown at different temperatures ..............................201 24. Phylogenetic tree of 16s small subunit RNA .................................................212 25. Phylogenetic tree of dsrAB gene ...................................................................213 26. Growth of D. carbinoliphilus Oakridgensis at different temperatures ................................................................................221 27. Growth of D. carbinoliphilus Oakridgensis at different pH values .....................................................................................222 28. Growth of D. carbinoliphilus Oakridgensis with different substrates .................................................................................223 29. Growth of D. carbinoliphilus Oakridgensis with different nitrogen sources ......................................................................224 30. Growth of D. carbinoliphilus Oakridgensis with different electron acceptors ....................................................................225 x LIST OF FIGURES CONTINUED Figure Page 31. Growth of D. carbinoliphilus Oakridgensis with Cr(VI) and U(VI) ...................................................................................226 xi ABSTRACT Sulfate-reducing bacteria (SRB) are a diverse group of anaerobic microorganisms that live in anoxic environments and play critical roles in biogechemical cycling, namely linkages between the carbon and sulfur cycles. Desulfovibrio vulgaris Hildenborough (DvH) is a model organism for SRB that has been studied for its ability to reduce toxic heavy metals to insoluble forms and its involvement in microbially induced corrosion in oil pipelines and other industrial settings. The described work investigated how the availability of electron donor/carbon sources and electron acceptors affected Cr(VI) reduction, metabolism, and biofilm growth and composition in DvH. DvH was grown planktonically at 20°C and 30°C in batch mode or as a biofilm under continuous flow at 20°C. In the second chapter of this dissertation, it is established that electron acceptor- limitation (EAL) predisposes cells to Cr(VI) toxicity compared to a balanced electron donor to electron acceptor (BAL) condition and electron donor-limited (EDL) condition. The effect of nutrient limitation on DvH biofilms is investigated, and microscopy revealed unique extracellular membranous structures that have not previously been observed. The extracellular structures were heterogeneously distributed, connected to cells, co-localized with metal precipitates, and more prevalent under EAL compared to BAL condition. Differential staining indicated that the structures were composed of lipid, consistent with the observation that these structures are membrane derived. Metabolomic analysis revealed an up-regulation of fatty acids under the EAL condition, which was confirmed and quantified via GC-MS. Down-regulated metabolites for biofilm grown under the EAL condition included those involved in DNA turnover, N-cycling, and peptidoglycan turnover, indicating that EAL may induce a switch from growth to fatty acid production that may coordinate with alternative electron transfer mechanisms. Outer membrane vesicles (OMVs) were purified from DvH biofilm and proteins detected in OMVs included porins, lipoproteins, hydrogenases, and oxidative stress response proteins. The results presented here show that nutrient limitation and resource ratio affect DvH physiology in both biofilm and planktonic growth modes. The analysis of the DvH biofilm matrix highlights the importance of investigating extracellular capabilities that are unique to the biofilm growth mode and has implications for activities and physiological states in the environment. 1 CHAPTER ONE INTRODUCTION Nutrient Limitation Across trophic levels, availability of nutrients determines growth rate, population size, and competition between organisms. In the microbial world, the effect of nutrient limitation on community structure and metabolism is an area of interest due to the desire to utilize and/or control microbial growth and metabolism. On a global scale, limitation of N, P, or Fe in oceans determines primary productivity, which subsequently affects global carbon flux and nutrient availability below the photic zone and to higher trophic levels (Moore et al. 2013). On a smaller scale, nutrient limitation can affect microbial community structure, with organisms that are capable of growth on low concentrations of substrates outcompeting other microbes, also known as the resource ratio theory (Mello et al. 2016). In a monoculture or coculture, nutrient limitation has been shown to affect microbial metabolism, physiology and interactions with other organisms. For example, some sulfate-reducing bacteria experiencing sulfate limitation can produce hydrogen as an electron sink. The production of hydrogen can attract hydrogenotrophic bacteria or archaea such as methanogens and initiate a syntrophic relationship (Brileya et al. 2013). The work presented here investigates the effect of nutrient limitation, in the form of electron donor/carbon limitation and electron acceptor/sulfate limitation on the sulfate- 2 reducing bacterium, Desulfovibrio vulgaris Hildenborough. Moreover, nutrient limitation is also examined in terms of a “resource ratio,” or how the ratio of concentration of carbon source/electron donor to sulfate/electron acceptor affects physiology. Previous work has shown that electron acceptor-limitation (oxygen limitation) in Shewanella oneidensis induces the production of extracellular nanowires and increases gene expression for certain cytochromes, while electron acceptor-limitation in Geobacter sulfurreducens also caused an increase in expression genes involved in the utilization of alternative terminal electron acceptors (Gorby et al. 2006; Bansal et al. 2013; Barchinger et al. 2016). Similarly, electron donor/carbon source limitation can result in an increase in abundance of transporter proteins, proteins involved in utilizing alternative carbon sources, and adjusted ribosome content to reflect slower growth rates (Gaal et al. 1997; Liu and Ferenci 1998; Marozava et al. 2014). Sulfate-Reducing Microorganisms Sulfate reducing microorganisms (SRM) are ubiquitous in nature and play an important role in the global sulfur cycle by reducing sulfate to sulfide. SRM are a phylogenetically diverse group of prokaryotes, consisting of gram-negative (belonging to δ-Proteobacteria) and gram-positive (belonging to Firmicutes and Nitrospira) bacteria, and archaea (Crenarchaeota, Euryarchaeota) (Castro 2000). SRM inhabit a variety of different environments including terrestrial subsurface environments, marine and freshwater sediments, and animal guts (Widdel 1988; Rey et al. 2013). SRM have been 3 isolated from cold environments, such as arctic sediments, hot environments such as hydrothermal vents and hot spring sediments, acidic, and hypersaline environments (Knoblauch et al. 1999; Fishbain et al. 2003; Foti et al. 2007; Alazard et al. 2010; Frank et al. 2013). Given the wide distribution of the sulfate reducing metabolism, it is apparent that these organisms play an essential role in ecosystem function and global nutrient cycling. SRM are responsible for an estimated 12-29% of organic carbon oxidation in anoxic seafloor sediments and up to 50% in wetland environments (Blodau et al. 2007; Keller and Bridgham 2007; Bowles et al. 2014), making them major drivers of carbon cycling in these environments. SRB are important microbial community members, consuming H2, organic acids, short chain and long chain fatty acids, and other carbon compounds, while also producing reduced sulfur compounds and influencing the redox state of the local environment (Muyzer and Stams 2008). SRB are also linked to microbial methane production through syntrophic relationships with methanogenic archaea in which methanogens consume hydrogen produced by SRB (Stams and Plugge 2009; McInerney et al. 2009). Applications of Sulfate-Reducing Bacteria Currently, SRB are recognized for both the beneficial role they play in the reduction of heavy metal pollutants to insoluble forms and the problems they cause in industrial settings such as corrosion of steel and oil souring in pipelines and reservoirs. 4 The role of SRBs in corrosion has been heavily researched because of the cost associated with repairing infrastructure that has been compromised (Koch and United States. Federal Highway Administration 2002). The current model for corrosion caused by SRB is bimodal in that the H2S produced by SRB is corrosive in itself and also some SRB are capable of using metallic iron as their sole electron donor (Enning et al. 2012; Enning and Garrelfs 2014). Oil souring is also caused by the production of H2S and therefore it is of keen interest to better understand how to inhibit the growth of SRB (Hubert and Voordouw 2007). SRB have also been extensively studied due to their ability to reduce heavy metals such as U(VI), Cr(VI), Se(VI), and Tc(VII) both chemically, by reacting with sulfide to produce insoluble metal-sulfides, and enzymatically (Lovley et al. 1993; Lovley and Phillips 1994; Lloyd et al. 1998; Hockin and Gadd 2003). Interest in using naturally occurring SRB for bioremediation of such heavy metal pollutants in contaminated sites has been explored as a more economical and environmentally friendly treatment than traditional chemical pump and treat methods (Tyagi et al. 2011). Pilot studies have analyzed the effectiveness of biostimulation strategies at contaminated field sites by injecting a carbon source and electron donor into the subsurface and tracking contaminant concentrations and microbial community composition over time. At one such site, the DOE Rifle site in Colorado, acetate was injected as the stimulant, which resulted in a substantial reduction in U(VI) levels (from 1-1.5µM to 0.05-0.1µM) (Williams et al. 2011). In situ incubations of sediment columns during acetate 5 amendment lead to the conclusion that U(VI) reduction occurs through multiple redox pathways, including enzymatic reduction followed by binding of U(IV) with biomass and abiotic reduction via FeS embedded in extracellular matrix materials in SRB biofilms. (Williams et al. 2011; Bargar et al. 2013) At another site, the DOE Oak Ridge site in Tennessee (FRC), contaminated groundwater is characterized by low pH, high levels of nitrate and U(VI) (Spain and Krumholz 2011). Previous studies have demonstrated reduction of nitrate and uranium levels in the subsurface upon bio-stimulation with ethanol at the FRC, and during bio- stimulation an increase was observed in DNA sequences corresponding to SRB (Hwang et al. 2009). Injections of emulsified vegetable oil (EVO) resulted in decreased U(VI) concentrations from between 9.1 and 3.8 µM to less than 1µM across all wells and resulted in an initial increase in SRB from the Desulforegula genus along with Veillonellaceae (belonging to Firmicutes). Post injection, after the EVO had been degraded to acetate, NO3-, Fe(III), SO42-, and U(VI) reduction decreased and was attributed to members of the Comamonadaceae, Geobacteriaceae, and Desulfobacterales (Gihring et al. 2011). However, further work is needed to fully develop and parameterize the proper constraints to model heavy metal bioremediation in both groundwater and sediments and better understand the physiology of important populations. Cr(VI) and U(VI) are also groundwater contaminants at the Hanford site in Washington that have been targeted for bioremediation. A 2004 injection of hydrogen release compound (HRC), a glycerol and polylactate mixture that also releases hydrogen 6 upon hydrolysis, into a Cr(VI) contaminated area resulted in a significant decrease in Cr(VI) and an increase in Pseudomonas, Desulfovibrio, and Geobacter in the groundwater (Faybishenko et al. 2008). The increase in relative abundance of dissimilatory metal-reducing bacteria such as SRB and Fe(III) reducers during these field-scale pilot studies is telling of the important role that they play in heavy metal and radionuclide reduction at contaminated sites. Desulfovibrio vulgaris Hildenborough Desulfovibrio vulgaris Hildenborough is a model organism for studying the sulfate-reducing metabolism and SRB-metal interactions. D. vulgaris is a gram-negative bacterium belonging to the δ-Proteobacteria that was isolated from clay soil in Hildenborough, England in 1946. The DvH genome was sequenced and published in 2004 and has aided in the elucidation of many metabolic pathways and gene functions (Heidelberg et al. 2004). D. vulgaris generates ATP through coupling the oxidation of carbon sources/electron donors with the reduction of sulfate to sulfide. Carbon, typically in the form of organic acids and alcohols, is incompletely oxidized to acetate, resulting in a nearly 1:1 stoichiometric output of acetate to carbon input (e.g., lactate). When lactate is provided as the carbon source, it is first oxidized to pyruvate, then to acetyl-CoA via lactate dehydrogenase and then pyruvate:ferredoxin dehydrogenase. Acetyl-CoA is then used for substrate-level phosphorylation to generate one ATP per lactate (Keller and Wall 2011a). 7 Sulfate reduction in D. vulgaris requires an initial input of energy (1 ATP) to activate sulfate and form adenosine phosphosulfate (APS). APS is then reduced by the APS reductase to bisulfite (HSO3-), which requires the hydrolysis of the pyrophosphate and two electrons. Bisulfite is reduced to sulfide (HS-) using six electrons via the dissimilatory sulfite reductase and another ATP molecule is required to convert the AMP produced during APS reduction to ADP. Two lactate molecules are required to reduce one sulfate molecule, resulting in a 2:1 ratio of lactate to sulfate needed for a “balanced” nutrient ratio. Because two ATP molecules are produced during lactate oxidation and two are consumed during sulfate reduction, resulting in a net gain of zero ATP, cells must have another way to conserve energy. Oxidative phosphorylation via ATP synthase is thought to be the mechanism for additional ATP production and to yield one net ATP, 6- 12 protons must be pumped for every molecule of sulfate reduced (Bargar et al. 2013). Membrane potential (Δp) can be generated via multiple proposed pathways, including hydrogen cycling, in which protons and electrons produced during the oxidation of lactate or pyruvate are used to produce H2 via a cytoplasmic hydrogenase and then the H2 can diffuse across the membrane where it can be re-oxidized by periplasmic hydrogenases. Electrons can then flow through the cytochrome c network in the periplasm and inner membrane to be used in the sulfate reduction pathway and the protons are pumped back across the membrane via ATP synthase (Odom 1981; Keller and Wall 2011). Additionally, the ferredoxin:NADH oxidoreductase Rnf is required when substrate-level phosphorylation is not possible and can pump ions across the 8 membrane, contributing to (Δp), and transfer electrons to NAD+ where they are donated to the sulfate reduction pathway via Hrd/flx protein complex that couples ferredoxin reduction with DsrC reduction (Ramos et al. 2015). Heavy Metal and Radionuclide Reduction in Desulfovibrio Another aspect of the D. vulgaris metabolism that has been subject to study is its ability to reduce heavy metals and radionuclides. Mechanisms for heavy metal/radionuclide reduction have been identified, but more study is necessary to fully understand responses to heavy metal toxicity and reduction. Cr(VI) was selected for study in this work due to its prevalence at the Department of Energy superfund site in Hanford, WA and Cr(VI) is also a prevalent contaminant at many DOE sites. Cr is the seventh most abundant element on earth and exists primarily as Cr(VI) or Cr(III). Cr(VI), which is typically associated with oxygen as chromate (CrO42-) or dichromate (Cr2O72-) ions, is considered the more toxic form of Cr due to its solubility in water. Cr(III) is usually associated with organic matter in the environment, exists as an oxide, hydroxide, or sulfur compound, and is generally less soluble and therefore less mobile than Cr(VI) (McGrath 1995). Cr(VI) Reduction in D. vulgaris Because sulfate and chromate are chemical analogues, it is hypothesized that chromate can enter the cell through sulfate permeases in D. vulgaris. Chromate uptake through sulfate transport systems has been shown in other organisms such as 9 Pseudomonas fluorescens, Escherichia coli, and Salmonella typhimurium (Pardee et al. 1966; Ota et al. 1971; Karbonowska et al. 1977; Ohtake et al. 1987; Sirko et al. 1990). Once inside the cell, multiple mechanisms for enzymatic Cr(VI) reduction by D. vulgaris are possible and this is probably because there is no specific chromate reductase enzyme, but rather enzymes that can use multiple substrates. Cytochrome c3 has been shown to function as both a Cr(VI) and U(VI) reductase and purified cytochrome c3 and hydrogenase in the presence of H2 as the electron donor can reduce Cr(VI) to Cr(III) (Lovley and Phillips 1994). A cytochrome c3 mutant in a different Desulfovibrio sp. (D. desulfuricans), had a slower U(VI) reduction rate than wild type and U(VI) reduction in the mutant was slower when H2 was the electron donor compared to lactate or pyruvate, further indicating that there is more than one Cr(VI) and/or U(VI) reduction pathway (Payne et al. 2002). Additionally, nitroreductase enzymes in Escherichia coli and Vibrio harveyi have been shown to also function as chromate reductases (Kwak et al. 2003a; Robins et al. 2013a), and unpublished microarray data shows that the nitroreductase in D. vulgaris is upregulated when Cr(VI) is present (Kwak et al. 2003b; Robins et al. 2013b)). Although D. vulgaris is capable of Cr(VI) reduction, energy production during Cr(VI) reduction is not linked to growth and Cr(VI) reduction intermediates such as Cr(V) and hydroxyl radicals have toxic and mutagenic effects on the cell (Chardin et al. 2002; Klonowska et al. 2008). One example of this is reduction of Cr(VI) to Cr(V) by NAD(P)H, FADH2, or glutathione can create hydroxyl radicals when H2O2 is present, causing intracellular damage to DNA (Kawanishi et al. 1986; Shi and Dalal 1990a; Shi 10 and Dalal 1990b; Kadiiska et al. 1994; Itoh et al. 1995). Intracellular Cr(III) can also have mutagenic effects by forming adducts with phosphate groups of DNA strands and preventing or causing errors in replication (Nishio and Uyeki 1985; Bridgewater et al. 1994). Because intracellular Cr(III) can cause DNA replication errors, it is advantageous to the organism to export Cr(III) out of the cell quickly. D. vulgaris has a few different potential mechanisms for defense against chromate toxicity, including chrA, a chromate transporter that is critical to chromate efflux in P. aeruginosa, merP, a mercury transporter, and acrA, a subunit of a multidrug efflux pump (Alvarez et al. 1999; Pimentel et al. 2002). These three genes were up-expressed in D. vulgaris when exposed to Cr(VI) (Fields et al., unpublished results). It has been shown that Cr(III) that has been reduced from Cr(VI) is located on the cell surface, in the cytoplasmic and outer membranes, and the periplasmic space (Goulhen et al. 2005). This is consistent with the evidence that periplasmic and membrane bound hydrogenases and cytochromes are involved in Cr(VI) reduction and then Cr(III) is excreted from the cytoplasm. Biofilm Physiology Microorganisms attached to a surface, known as a biofilm, have distinct characteristics and growth physiology compared to planktonic counterparts in the bulk aqueous phase. The biofilm growth mode is thought to be the primary growth mode for microorganisms in a wide range of environments and because of this it is important to understand the components of microbial biofilms that make them unique (Hall-Stoodley 11 et al. 2004). Moreover, microorganisms associated with sediments (i.e., biofilms) play an important role in nutrient and contaminant cycling and this unique physiology must be considered when trying to understand metal fate in the environment. One of the defining characteristics of the biofilm growth mode is the production of an extracellular matrix, also known as EPS (extracellular polymeric substances). The extracellular matrix can be composed of many different materials depending on the organism and the environment and can account for 50-90% of the total organic matter of a biofilm (Christensen and Characklis 1990; Nielsen et al. 1997). The biofilm matrix aids in microbial attachment to surfaces and other cells and determines the unique structure of biofilms, which affects nutrient and waste transport and the formation of microenvironments within the biofilm. A description of the major matrix components and examples of their proposed functions follows. The Biofilm Matrix Polysaccharides. Polysaccharides were originally assumed to be the majority of biofilm matrices, but now protein, nucleic acids, lipids, and other macromolecules are also recognized as important matrix components. Polysaccharide is a general term used for linked, repeating sugar molecules. Polysaccharides in biofilm matrices can exist as homopolysaccharides and/or heteropolysaccharides and contribute to attachment and morphology/structure in many organisms. Organisms often produce more than one type of polysaccharide, as is the case for P. aeruginosa that produces alginate, Pel, and Psl. P. 12 aeruginosa with a mutation in the σ-factor that regulates alginate production, results in an overproduction of alginate and a “mucoid” phenotype with increased resistance to antimicrobials (Hentzer et al. 2001). P. aeruginosa biofilms deficient in alginate production can still form biofilms due to the Pel and Psl polysaccharides. The Pel polysaccharide is a glucose-rich polysaccharide important for pellicle formation at the air-liquid interface and Psl is a mannose- and galactose-rich polysaccharide that is important for cell-cell and cell-surface attachment (Friedman and Kolter 2004; Ma et al. 2006). The exopolysaccharides produced by P. aeruginosa are examples of the diversity of polysaccharides that can be produced by one species/strain and the specific role that each can play in biofilm attachment, structure, and function; however, it appears that there is little conservation in exopolysaccharide production and function across microbial biofilms. Proteins. Proteins are also integral components of microbial biofilm matrices and the roles of specific proteins in biofilm formation, structure, and function have been elucidated in many microorganisms. Both enzymatic and structural proteins have been detected in biofilm matrices. Extracellular enzymes are involved in degradation of matrix materials during times of starvation or to enable dispersal (Vats and Lee 2000; Zhang and Bishop 2003). Structural proteins, such as pili and flagella, are also important parts of many biofilm matrices with flagellar and pili mutations in some organisms causing changes in biofilm morphology compared to wild-type (E. coli, P. aeruginosa, 13 Streptococcus pyogenes) (Pratt and Kolter 1998; Klausen et al. 2003; Kimura et al. 2012). A variety of other biofilm matrix proteins have distinct functions specific to a particular microorganism (Flemming and Wingender 2010), for example LapA in Pseudomonas species (Ivanov et al. 2012). DNA. Another biofilm matrix component that is important for the structure of some biofilms is extracellular DNA. Strategies for release/secretion of eDNA from cells differs from organism to organism with some using cell lysis as a release mechanism, while others package eDNA into membrane vesicles for release or have a yet unidentified secretion mechanism (Molin and Tolker-Nielsen 2003; Liao et al. 2014). Treatment of organisms with DNase has been shown to either prevent biofilm formation and/or disband established biofilms without having an effect of the organisms themselves, indicating the importance of eDNA for maintaining biofilm structure (Whitchurch 2002; Liao et al. 2014; Sena-Vélez et al. 2016). Further, visualization of eDNA in biofilms has shown DNA in the extracellular matrix can form extensive networks and connect cells across channels within the biofilm (Böckelmann et al. 2006; Jurcisek and Bakaletz 2007). In P. aeruginosa biofilms, eDNA has also been shown to protect against aminoglycosides, indicating that eDNA may play more than a structural role for some organisms (Chiang et al. 2013). The genomic content of the eDNA in biofilm matrices has also been investigated and, again, varies according to the organism studied. In a bacterial isolate from the Saskatchewan River, isolated eDNA from the biofilm matrix 14 was similar to genomic DNA, but treatment with restriction endonucleases and DNA fingerprinting resulted in slightly different banding patterns (Böckelmann et al. 2006). Alternatively, eDNA isolated from P. aeruginosa and subjected to the same fingerprinting analysis as above, revealed no differences between genomic and extracellular DNA (Steinberger and Holden 2005). Lipids. Lipids have also been reported as a component of biofilm matrices in some organisms, but have not been as well studied as polysaccharides, proteins, or eDNA. Serratia marcescens produces serrawettins, hydrophobic molecules composed of hydroxy fatty acids that aid in colony biofilm spreading by reducing surface tension at the interface of different phases (Matsuyama and Nakagawa 1996). More recently, it has been found that matrix in mixed species biofilms that form on membrane bioreactors for wastewater treatment is composed largely of lipid rather than protein or polysaccharide, however these free lipids may be the product of dead cell debris (Inaba et al. 2017). While literature often cites lipids as being a potential component of biofilm matrices, there are few detailed reports on their abundance, composition, and function and is therefore an area in which more study is needed. Outer Membrane Vesicles. The last component of biofilm matrices that will be highlighted here is outer membrane vesicles (OMVs). OMVs are small (20-250 nm) bilayer structures that bleb from the outer membrane of cells and carry cellular components in and on them. OMVs are thought to be produced by all Gram-negative 15 bacteria and also some Gram-positive bacteria and archaea (Ellen et al. 2009; Lee et al. 2009; Kulp and Kuehn 2010). The OMVs thus far studied can be a combination of proteins, DNA, and small molecules and have been implicated in many different processes from pathogenesis to horizontal gene transfer (Yaron et al. 2000; Kuehn 2005). Interestingly, OMVs produced by biofilm cells have distinct protein profiles compared to those produced by the same organism growing planktonically, suggesting that OMVs could play a unique role in biofilms (Schooling and Beveridge 2006; Park et al. 2015; Grande et al. 2015). Biofilm Resistance to Heavy Metal and Radionuclide Toxicity The biofilm growth mode is prevalent in many different environments and biofilm characteristics, such as increased resistance to antimicrobials or other external stressors, may be attributed to the presence of a heterogeneous matrix containing protective elements. Studies have shown that biofilms can have increased resistance to external stressors such as changes in water availability, UV radiation, or the presence of antibiotics or heavy metals compared to planktonic cells (Elasri and Miller 1999; Harrison et al. 2005; Chang et al. 2007), but it can be difficult to delineate the combined attributes of inherent physiology unique to the biofilm and intrinsic physical characteristics of being in biofilm itself. The effect of heavy metals on biofilms compared to planktonic cells is of interest due to the potential for indigenous microorganisms to interact with high concentrations of 16 heavy metals and/or radionuclides at contaminated sites and to better understand how corroding biofilms interact with metals. Microorganisms in these contaminated subsurface environments exist predominantly as biofilms, and therefore it is important to investigate how microbes interact with metals in the biofilm growth mode (Dar et al. 2013). In P. aeruginosa, cells growing as a biofilm were twice as resistant to lead compared to planktonic cells and 600 times as resistant to copper. The authors of this study propose that increased Cu and Pb resistance in P. aeruginosa biofilms could be due to metal complexation with EPS components, which has been demonstrated in other organisms (Kaplan et al. 1987; McLean et al. 1990; Kim et al. 1996; Teitzel and Parsek 2003). Pseudomonas putida biofilm exposed to Cr(VI) increases production of extracellular carbohydrate, protein, and DNA compared to biofilm not exposed to Cr(VI), further supporting the hypothesis that the extracellular matrix of a biofilm is involved in metal resistance (Priester et al. 2006). Interactions of sulfate reducing biofilms with metals has also been investigated and studies have observed that naturally occurring, multispecies SRB biofilms contain metals such as selenium and zinc immobilized in the matrix (Hockin and Gadd 2003; Labrenz and Banfield 2004). In monoculture biofilms of Desulfovibrio desulfuricans exposed to lead or U(VI), the amount of Pb(II) or U(VI) immobilized in the biofilm was dependent on sulfide production, as chemical reduction of Pb(II) or U(VI) by sulfide was the dominant mode of metal reduction in these biofilms. Biogenic U(VI) reduction also contributed to the overall immobilization of U(VI) in the biofilm, but the ratio of abiotic 17 to biotic reduction is unclear (Beyenal et al. 2004; Beyenal and Lewandowski 2004). The production of sulfide by SRB is advantageous for increasing metal reduction and immobilization through abiotic means; however, the presence of sulfide also makes it very challenging to investigate the biotic response of the organisms. Physiology of D. vulgaris Biofilms Limited research has been done to characterize the properties and characteristics of D. vulgaris biofilms or other SRB compared to other model organisms such as P. aeruginosa, E. coli, or S. aureus, but the work that has been done has laid a good foundation for understanding gene expression and protein regulation in the biofilm vs. planktonic growth mode. Clark et al. (2012) observed that biofilm cells have altered electron flow pathways, decreased amino acid and nucleotide biosynthesis, and decreased cell wall and membrane biosynthesis, which all point to an altered metabolic state compared to exponentially growing planktonic cells. Further, carbohydrate metabolism was altered, with a reported decrease in gluconeogenesis related genes and an increase in glycerol uptake and utilization related genes. Expression of transporter genes was also altered, with genes for ABC transporters, two other transporter proteins, and a type I secretion system protein being up-expressed under biofilm conditions along with a type III secretion system protein also that had increased abundance. In addition, recent results showed the important role of a type I secretion system for biofilm formation in D. vulgaris Hildenborough (De León et al. 2017). The increase in expression/abundance of 18 particular transporter/secretion systems for biofilm cells is perhaps indicative of an increase of small molecules, proteins, or other substrates entering and exiting the cell and/or biofilm matrix compared to planktonic cells (Clark et al. 2012). Previous results with D. vulgaris indicated that biofilms on glass did not contain an extensive exopolysaccharide matrix, used protein filaments to form biofilm between cells and the silica oxide surfaces, and the filaments appeared to be flagella (Clark et al., 2007). The data suggested that D. vulgaris used flagella for more than a means of locomotion to a surface, but also used flagella, or modified flagella, to establish and/or maintain biofilm structure (Clark et al. 2007). Proteomic analysis of the extracellular fraction of D. vulgaris biofilm revealed 188 proteins. Noteworthy is a highly abundant protein that contains a vonWillebrand factor domain, which can be involved in cell adhesion, pattern formation, and signal transduction. In addition, abundant in the extracellular fraction are two outer membrane porin proteins whose function in the biofilm matrix is unknown (Clark et al. 2012; Zeng et al. 2017). Research Objectives The overarching theme of this work is the effect of nutrient limitation, in the form of electron donor/carbon or electron acceptor limitation, on the growth and response of D. vulgaris Hildenborough to Cr(VI) stress in both planktonic and biofilm growth modes. Limitation of electron donors/carbon sources or electron acceptors is a scenario the SRB likely encounter in natural environments as well as engineered systems. For example, in 19 the context of bioremediation, electron donor/carbon sources are injected into the subsurface during biostimulation regimes that can create an electron acceptor-limited (EAL) environment. In offshore oil wells, ocean water is typically injected into the well to recover the remaining oil, but in doing so they are injecting ~30mM sulfate, creating an electron donor/carbon limited (EDL) system. Better understanding of how nutrient limitation affects D. vulgaris planktonic growth, biofilm production, and response to Cr(VI) could contribute to better microbial control in engineered systems and better understanding of how SRB interact with their environment and other organisms in natural settings. Further, most studies investigating the physiological response of dissimilatory metal reducing bacteria have cultured them at 30°C, an optimal growth temperature in terms of growth rates, but not always an environmentally relevant temperature. The research presented in Chapter 2 compared planktonic growth and Cr(VI) reduction at 20 and 30°C to determine how temperature affected rate of Cr(VI) reduction in addition to how nutrient limitation affected D. vulgaris tolerance to Cr(VI). Based on the findings from Chapter 2, we decided that further investigation into how nutrient limitation affected biofilm growth in D. vulgaris was warranted. For some of the more in-depth analysis and experiments, the EAL and BAL conditions were chosen for comparison with the BAL condition being a “control” in which lactate and sulfate concentrations are “balanced” and the EAL being the “stressed” or “limited” condition. To limit sample number, the EDL condition was omitted in a few instances. To understand how nutrient limitation affected the metabolism of D. vulgaris biofilm under 20 sulfate-limiting conditions (EAL), the metabolite profiles from biofilm grown under EAL and BAL conditions were compared. These results are presented in Chapter 3 and the data from this experiment was also used for metabolomic method and data analysis development, which is presented in Appendices C and D. Because of the inherent heterogeneity, gradient and micro-niche formation in biofilms, D. vulgaris biofilms grown under EAL and BAL conditions were also compared using high resolution microscopy and differential staining techniques to detect such differences within a biofilm. The observations made during these experiments regarding the extracellular matrix of the biofilm are presented in Chapter 4 and lead to the development of methods to isolate fractions of the extracellular matrix. Outer membrane vesicles were isolated from D. vulgaris biofilms grown under EAL and BAL conditions and analyzed via proteomic methods. The results of these experiments are detailed in Chapter 5. While studying model organisms such as D. vulgaris Hildenborough is useful and practical due to the body of literature available for reference, it is becoming increasingly apparent that model organisms are not always representative of the systems in which we are interested. The work presented here strives to apply some field-relevant parameters to the growth of a model organism, but moving forward, relevant field isolates should be selected for study. 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Chemosphere 50:63–69. 31 CHAPTER TWO Cr(VI) REDUCTION AND PHYSIOLOGICAL TOXICITY IS IMPACTED BY RESOURCE RATIO IN DESULFOVIBRIO VULGARIS HILDENBOROUGH Contribution of Authors and Co-Authors Manuscript in Chapter 2 Author: Lauren C. Franco Contributions: Developed experimental design, performed experiments, analyzed data, wrote and revised the manuscript. Co-Author: Grant Zane Contributions: Created mutant strains, wrote and revised the manuscript. Co-Author: Sadie Steinbeisser Contributions: Performed experiments. Co-Author: Judy Wall Contributions: Created mutant strains, wrote and revised the manuscript. Co-Author: Matthew W. Fields Contributions: Developed experimental design, analyzed data, wrote and revised the manuscript. 32 Manuscript Information Page Lauren C. Franco, Grant Zane, Sadie Steinbeisser, Judy Wall, Mathew W. Fields Applied Microbiology and Microbiology Status of Manuscript: __ _ Prepared for submission to a peer-reviewed journal __x_ Officially submitted to a peer-review journal ____ Accepted by a peer-reviewed journal ____ Published in a peer-reviewed journal Springer Berlin Heidelberg August 14, 2017 33 Abstract Desulfovibrio spp. are capable of heavy metal reduction and are well-studied systems for understanding metal fate and transport in anaerobic environments. Desulfovibrio vulgaris Hildenborough was grown under environmentally-relevant conditions (i.e., temperature and nutrient limitation) to elucidate the impacts of Cr(VI) reduction on cellular physiology. Growth at 20°C was slower than 30°C and the presence of 50 µM Cr(VI) caused extended lag times for all conditions, but once resumed the growth rate was similar to that without Cr(VI). Cr(VI) reduction rates were greatly diminished at 20°C for both 50 µM and 100 µM Cr(VI), particularly for the electron acceptor limited (EAL) condition in which Cr(VI) reduction was much slower, the growth lag much longer (200 h), and viability decreased compared to balanced (BAL) and electron donor limited (EDL) conditions. Similar results were observed between the different resource ratio conditions when the sulfate levels were normalized (10 mM), and these results indicated that resource ratio (donor:acceptor) impacted D. vulgaris response to Cr(VI) and not merely sulfate limitation. When sulfate levels were increased in the presence of Cr(VI), cellular responses improved via a shorter lag time to growth. The results suggest that temperature and resource ratios greatly impacted the extent of Cr(VI) toxicity, Cr(VI) reduction, and the subsequent cellular health via Cr(VI) influx and overall metabolic rate. The results also emphasized the need to perform experiments at lower temperatures with nutrient limitation to make accurate predictions of heavy metal reduction rates as well as physiological states in the environment. 34 Introduction Chromium is a naturally occurring trace element that is widely used for industrial purposes (i.e., metallurgy, leather tanning) and has become a major pollutant of soil and groundwater (Barnhart 1997; Zayed and Terry 2003; Dhal et al. 2013). Hexavalent chromium is soluble and can easily be transported across cell membranes whereas trivalent chromium is mostly insoluble and therefore not as toxic to cells. Cr(VI) is a known mutagen and carcinogen, although it appears that chromium toxicity is largely caused by reactive intermediates such as Cr(V) or hydroxyl radicals produced during Cr(VI) reduction and Cr(III) forming adducts with DNA or natural organic matter (Nriagu and Nieboer 1988; Aiyar et al. 1991; Zhitkovich et al. 1995; 2013; Gustafsson et al. 2014). Desulfovibrio species are model sulfate-reducing bacteria (SRB) and have been shown to reduce metals, metalloids, and radionuclides (Heidelberg et al. 2004; Klonowska et al. 2008). Previous work with purified enzymes and whole cells has shown that D. vulgaris is capable of reducing Cr(VI) via hydrogenases and cytochrome c3, but cells are unable to use Cr(VI) as a terminal electron acceptor linked to growth (Lovley and Phillips 1994; Chardin et al. 2002; Elias et al. 2004). One promising approach for treatment of contaminated environments is in situ biostimulation, the process of promoting indigenous microbial activity via the addition of carbon, nitrogen, phosphorus, and/or energy sources (Tyagi et al. 2011). Typically, microbial activity is promoted 35 through the addition of carbon/energy sources as soluble substrates and/or more recalcitrant, complex substrates that degrade more slowly (Faybishenko et al. 2008). For example, biostimulation has been used successfully to reduce contaminants at the Cr(VI) contaminated 100-H area of the Hanford Site in south-central Washington state. Injections of electron donors such as polylactate hydrogen-release compound (HRC) into the subsurface have been shown to stimulate Cr(VI)-reducing microorganisms such as Desulfovibrio spp. and other metal-reducing bacteria in situ (Zhang et al. 2015). Electron donors and electron acceptors are rarely balanced according to metabolic stoichiometry for a particular organism or guild. In fact, it is common practice to stimulate growth of existing microorganisms by injecting an excess electron donor into the subsurface, creating an electron acceptor limited environment. Previous studies have reported that substrate limitation can increase Cr(VI) susceptibility (Chardin et al. 2002), and injections of electron donor without a corresponding increase in electron acceptor creates unbalanced electron donor to acceptor ratios. However, little is known about the physiological responses of metal-reducing populations in the context of unbalanced ratios that occur as a consequence of stimulated conditions. In addition, metal-reducing bacteria are often studied at optimal growth temperatures (between 30 and 40°C) although much lower temperatures are experienced in situ. A previous study examined uranium reduction in a sulfate-reducing consortium at 10, 20, and 30°C, and observed that growth and reduction kinetics were affected by temperature (Boonchayaanant et al. 2008). Therefore, it is important to understand the growth characteristics of 36 microorganisms that are capable of heavy metal immobilization, such as SRB, under typical field conditions to accurately assess and predict ability and rate of contaminant immobilization. To our knowledge, previous studies have not evaluated Cr(VI) effects on SRB growth and reduction rates at a temperature lower than 30°C under unbalanced stoichiometries of electron donors and acceptors. Materials and Methods Strains and Growth Conditions The Desulfovibrio vulgaris Hildenborough culture was acquired from Dr. Romy Chakraborty (Lawrence Berkeley National Laboratory). LS4D medium (Borglin et al. 2009) (modified to 2.5 µM resazurin and 130 µM riboflavin in Thauer’s vitamins (Brandis et al. 1981)) was prepared anoxically by boiling water under oxygen-free N2 gas, adding medium components, and dispensing into N2 gassed tubes or serum bottles sealed with butyl stoppers and aluminum crimp seals as previously described (Klonowska et al. 2008). LS4D medium was modified to alter electron donor (lactate) to electron acceptor (sulfate) ratios: 60 mM to 30 mM (balanced, or 20:10), 50 mM to 10 mM (electron acceptor-limited), and 18 mM to 50 mM (electron donor-limited or 5:10). The medium was not prepared with a reducing agent to avoid abiotic Cr(VI) reduction. Inocula for all experiments unless otherwise noted were grown in the same electron donor to acceptor ratio (i.e., inoculum grown in balanced medium was 37 inoculated into balanced medium). Cultures were grown at ambient room temperature (approximately 20°C) unless otherwise stated. Inocula for all experiments were prepared by washing mid-exponential phase (approximately 0.4 OD600) cultures to remove sulfide as previously described (Klonowska et al. 2008). To do this, cultures (9 ml aliquots) were harvested by centrifugation at 5,750×g for 10 min at room temperature. The supernatant was removed anoxically and aseptically by needle attached to a vacuum flask under constant flow of N2 gas to maintain neutral pressure. Pellets were then re-suspended in 9 ml fresh LS4D medium that contained lactate and sulfate concentrations of the respective growth condition and washed once more, after which the pellets were re-suspended and concentrated in 1 ml fresh LS4D. Concentrated cell cultures were inoculated into 25 ml balch tubes with LS4D medium (15 ml) that contained Cr(VI) (potassium chromate) at 0, 50, and 100 µM to an OD600 between 0.06 and 0.07. Growth was monitored by optical density (600 nm) and samples of 0.2 to 0.5 ml were withdrawn throughout the growth cycle to monitor Cr(VI), lactate, acetate, and sulfate concentrations. All experiments were performed in triplicate. Ascorbate, which has been shown to reduce Cr(VI) to Cr(III) and form short-lived Cr(V) and Cr(IV) intermediates and Cr(III)-ascorbate complexes (Cieǎṡlak-Golonka et al. 1992; Stearns and Wetterhahn 1994), was added to cultures with and without Cr(VI) to determine the effect that complexed Cr(III) had on growth. Cells were prepared as described above and inoculated into electron acceptor- limited LS4D medium that contained either 50 µM sodium ascorbate, 50 µM potassium 38 chromate, 50 µM sodium ascorbate and 50 µM potassium chromate, or 50 µM potassium chromate added prior to inoculation and 50 µM sodium ascorbate added 3 hours post- inoculation. Experimental Design In order to determine the physiological responses and Cr(VI) reduction rates under field relevant conditions, D. vulgaris was grown under three conditions of resource ratios with and without Cr(VI) (50 and 100 µM) at 20°C. A ratio of 60:30 (or 20:10) mM (lactate:sulfate) was used for balanced (BAL), 50:10 mM for electron-acceptor limited (EAL), and 18:50 (5:10) mM for electron-donor limited (EDL). Cell cultures were washed of sulfides to observe cellular responses to field relevant levels of Cr(VI). Cr(VI) reduction rates, growth rates, biomass yields (Ysulfate), and viability were measured to assess physiological responses. Similar levels of lactate (electron-donor) were tested with increasing levels of sulfate (electron-acceptor) and showed the role of Cr(VI) influx, metabolism, and toxicity on cell viability and Cr(VI) reduction. Cr(VI) Reduction Analysis Cr(VI) reduction was measured by quantifying Cr(VI) concentrations over time using the diphenylcarbazide method with Hach ChromaVer 3 reagent (Hach Company, Loveland, CO) as previously described (Viamajala et al. 2002; Klonowska et al. 2008) except that absorbance was read in disposable cuvettes (Plastibrand, Germany) at 540 nm. Cells were removed by filtration (0.22 µm) so that only Cr(VI) in the filtrate was 39 measured. Primary and secondary Cr(VI) reduction rates were calculated from bi-phasic decline Cr(VI) levels and the culture density did not change significantly from inoculation during the time in which the rates were calculated. Cr(VI) in uninoculated medium and cultures containing heat-killed cells were also measured. Analytical Techniques Lactate and acetate concentrations were measured by HPLC (Dionex) with an Aminex HPX-87H ion exclusion column. Sulfate concentrations were measured by ion chromatography (Dionex) with an IonPac AS11 column (Dionex). All samples were measured in triplicate. Cell viability was measured via the most probable number (MPN) method at 3 and 48 hours. Washed cells were inoculated into fresh LS4D medium (~0.07 OD600) that was electron donor to acceptor balanced or electron acceptor-limited and contained 0 or 50 µM K2CrO4. Subsamples were removed and serially diluted with LS4D medium that contained sodium sulfide as a reducing agent. MPN cultures were incubated at 30°C for three weeks at which point the MPN for each condition was calculated (Jarvis et al. 2010). Mutant Generation Construction of the mutants lacking the annotated sulfate permease genes, sulP, was accomplished similarly to the generation of marker-less deletion strains in the uracil phosphoribosyltransferase (upp, pyrimidine salvage pathway enzyme) deletion mutant (JW710; resistant to 5-fluoro uracil, 5FU) of Desulfovibrio vulgaris Hildenborough 40 (Keller et al. 2009). In short, two plasmids were necessary for each gene deletion. The first (marker-exchange) plasmid, contained the antibiotic-resistance cassette encoding the aminoglycoside-3’-phosphotransferase (npt, Kmr) driven by the native promoter (Pnpt) followed by the upp gene (Pnpt-npt-upp) and was exchanged with the gene of interest (selected with the kanamycin analog G418). The second (marker-less deletion) plasmid was used for removing the cassette that conferred sensitivity to 5FU (selected with 5FU) resulting in the in-frame, marker-less deletion. Following each transformation, putative transformants were screened for phenotypes with spectinomycin, 5FU and G418 to determine the probability that a double homologous recombination event took place rather than a single recombination event resulting in plasmid insertion. Three of the isolates with correct phenotypes were further confirmed by Southern blot analysis and one selected as the confirmed deletion strain. Multiple deletions were obtained through an iterative process of transforming the appropriate parental strain with the marker- exchange/marker-less deletion plasmids and selecting for the appropriate resistances. A total of nine plasmids were constructed for generating the deletion strains of the three putative sulfate-permease genes (Table S1). Each marker-exchange plasmid was initially constructed with only the npt gene and no upp gene, as done in other studies (Parks et al. 2013; Lovley and Phillips 1994; Li and Elledge 2007). These plasmids were later modified to include the upp gene as the second gene in an artificial operon driven by Pnpt. This was accomplished by PCR amplifying the plasmid with a pair of primers that allowed for the addition of the counter-selectable feature by the SLIC (sequence- and 41 ligation- independent cloning) technique (Li and Elledge 2007). The construct for a marker-less, in-frame deletion plasmid was produced by PCR amplification of the original marker-exchange plasmid with primers that excluded the kanamycin-resistance marker and ligated closed using the SLIC procedure. The regions of each plasmid necessary for homologous recombination were sequenced (DNA core, University of Missouri, Columbia) to determine fidelity to the published sequence. These plasmids were transformed into JW710 for the single deletion strains or into one of the marker-less deletion strains to generate strains deleted of multiple genes. All primers are listed in Table S2. Results Temperature Affects D. vulgaris Growth and Cr(VI) Reduction/Toxicity Under A Balanced Resource Ratio D. vulgaris was grown at both 20°C and 30°C, with 0 and 50 µM Cr(VI), to assess differences in growth rate, biomass yield, and Cr(VI) tolerance at different resource ratios. Under balanced ratio conditions, D. vulgaris had a 3.6-fold slower growth rate (0.03 h-1) at 20°C compared to growth at 30°C (0.12 h-1) (Figure 1C and D). When exposed to Cr(VI) under balanced conditions, D. vulgaris could tolerate and reduce Cr(VI) faster at 30°C compared to at 20°C. At 30°C, only a small growth rate difference (10% lower) was observed when cells were exposed to 50 µM Cr(VI) (Figure 1C and D). At 20°C and BAL conditions, cells exposed to 50 µM Cr(VI) had an extended 42 lag time (~100 h), but the subsequent growth rate was similar to cultures without Cr(VI) (Figure 1D). In addition, the YSulfate values were similar between 0 and 50 µM Cr(VI) once growth was complete. Even under balanced conditions at 20°C, exposure to 100 µM Cr(VI) caused variable growth and greatly extended lags even though Cr(VI) levels declined (Figure 2D). For EDL and EAL conditions at 20°C, 100 µM Cr(VI) caused culture death and incomplete Cr(VI) reduction (Figure 2). Therefore, growth at 50 µM Cr(VI) was selected as a sub-toxic level to elucidate physiological responses across the tested resource ratios. At 20°C and 30°C, cultures with and without Cr(VI) under balanced conditions utilized lactate and produced equimolar levels of acetate until sulfate was depleted (Figure S1C). In the presence of 50 µM Cr(VI) under balanced conditions, lactate and sulfate were not utilized during the growth lag, and growth coincided with the utilization of lactate and sulfate (Figure S1D). Temperature also affected D. vulgaris ability to reduce Cr(VI) (50 or 100 µM) under balanced conditions. At 30°C, 50 µM Cr(VI) was reduced at a rate of 23.63±0.12 µM Cr(VI)/h whereas at 20°C, 50 µM Cr(VI) was reduced at an initial rate of 10.81±2.14 µM Cr(VI)/h and a secondary rate of 1.51±0.18 µM Cr(VI)/h (average O.D. 0.08)). At 30°C, 100 µM Cr(VI) was reduced at an initial rate of 22.53±0.78 µM Cr(VI)/h and a secondary rate of 0.45±0.22 µM Cr(VI)/h (average O.D. of 0.08)). At 20°C, 100 µM Cr(VI) was reduced at an initial rate of 14.21±0.58 µM Cr(VI)/h and a secondary rate of 0.42±0.15 µM Cr(VI)/h (average O.D. 0.07)) (Figure 2C, D). Ultimately, a decrease in 43 temperature caused significant decreases in D. vulgaris Cr(VI) reduction rates under the balanced condition. Previous work has highlighted many different enzymes that are capable of Cr(VI) reduction in D. vulgaris (Lovley and Phillips 1994; Chardin et al. 2003a; Li and Krumholz 2009a), but these studies have been conducted at temperatures selected for maximal growth. Temperature effects on D. vulgaris growth and Cr(VI) reduction were pronounced and emphasized the need to study microorganisms at temperatures that are field relevant in order to provide improved estimates for ecosystem function (e.g., Cr(VI) reduction). The difference in Cr(VI) reduction and growth at 20°C compared to 30°C is likely due to the combination of a slower growth rate (i.e., slower overall metabolism) at 20°C and the temperature dependence of enzyme reaction rates. When U(VI) reduction was measured at 20°C and 30°C in a sulfate-reducing consortium, pseudo second-order rate constants for uranium reduction tripled with the 10°C increase in temperature (Boonchayaanant et al. 2008), further emphasizing the role that temperature plays in microbial processes such as heavy metal reduction. Our results showed that the rate of Cr(VI) reduction during D. vulgaris growth was approximately 2-fold slower at 20°C compared to 30°C and the slower Cr(VI) reduction likely contributed to the increased Cr(VI) toxicity. 44 Figure 1. Growth of D. vulgaris at 30°C (a, c, and e) and 20°C (b, d and f) under EDL (a,b), BAL (c,d), and EAL (e,f) conditions with 0 (●) and 50 (◻) µM Cr(VI). Growth rates (h-1) and growth yields (grams of protein/moles sulfate consumed) are overlaid for each condition. 45 Figure 2. Cr(VI) reduction at 30°C (a, c, and e) and 20°C (b, d and f) under EDL (a,b), BAL (c,d), and EAL (e,f) conditions with 50 (○) and 100 (■) µM Cr(VI). Dashed lines indicate heat-killed (◆) and uninoculated controls (x) (C). Cr(VI) reduction rates (µM Cr(VI)/hr) (primary and secondary) are overlaid for each condition. C r( V I) (µ M ) C r( V I) (µ M ) C r( V I) (µ M ) Time (Hours) Time (Hours) E F C D A B 46 Resource Ratio Imbalance Affects D. vulgaris Cr(VI) Reduction and Tolerance Electron-Donor Limitation (EDL). D. vulgaris was grown at both 20°C and 30°C, with 0 and 50 µM Cr(VI) to assess differences in growth rate, biomass yield, and Cr(VI) tolerance under electron-donor limitation. Under EDL conditions, D. vulgaris had a 2.9-fold slower growth rate (0.04 h-1) at 20°C compared to growth at 30°C (0.12 h-1) (Figure 1A, B). When exposed to Cr(VI) under EDL conditions, D. vulgaris could tolerate and reduce Cr(VI) faster at 30°C compared to 20°C, but to a lesser extent than under balanced conditions. At 30°C and exposure to Cr(VI), the growth rate declined 25% (Figure 1A). At 20°C, cells exposed to 50 µM Cr(VI) had an extended lag time that was longer than the balanced condition (~125 h), but the subsequent growth rate was similar to the no Cr(VI) treatment (Figure 1B). At 20°C, cultures without Cr(VI) under EDL conditions utilized lactate and produced equimolar levels of acetate until lactate was depleted (Figure S1A). In the presence of 50 µM Cr(VI) under EDL conditions, lactate and sulfate were not utilized during the growth lag, and growth coincided with the utilization of lactate and sulfate (Figure S1B). Temperature also affected D. vulgaris ability to reduce Cr(VI) under EDL conditions. At 30°C, 50 µM Cr(VI) was reduced at an initial rate of 24.40±0.12 µM Cr(VI)/h (average O.D. of 0.08)) similar to the balanced condition, but at 20°C, 50 µM Cr(VI) was reduced at an initial rate of 9.12±3.45 µM Cr(VI)/h and a secondary rate of 1.32±0.22 µM Cr(VI)/h (average O.D. of 0.07)). At 30°C, 100 µM Cr(VI) was reduced at an initial rate of 22.74±0.66 µM Cr(VI)/h and a secondary rate of 0.53±0.36 µM Cr(VI)/h 47 (average OD 0.08). At 20°C, 100 µM Cr(VI) was reduced at a rate of 12.38±2.35 µM Cr(VI)/h that slowed to 1.96±0.20 µM Cr(VI)/h (average O.D. 0.08) (Figure 2A, B). Ultimately, the decrease in temperature caused a decrease in Cr(VI) reduction rates and the electron donor limitation with 50 µM Cr(VI) caused a 25 h increase in lag time compared to the balanced condition with 50 µM Cr(VI). Electron-Acceptor Limitation (EAL). D. vulgaris was grown at both 20°C and 30°C, with 0 and 50 µM Cr(VI) to assess differences in growth rate, biomass yield, and Cr(VI) tolerance under electron-acceptor limitation. Under EAL conditions, D. vulgaris had a 3.5-fold slower growth rate (0.04 h-1) at 20°C compared to growth at 30°C (0.13 h- 1) (Figure 1E and F). When exposed to Cr(VI) under EAL conditions, D. vulgaris could tolerate and reduce Cr(VI) faster at 30°C compared to at 20°C but to a much lesser extent than balanced conditions. At 30°C and exposure to Cr(VI), the growth rate declined 24% (Figure 1E). At 20°C, cells exposed to 50 µM Cr(VI) had an extended lag time that was longer than the balanced or EDL condition (~200 h), and the subsequent growth rate was 15% slower than no Cr(VI) treatment (Figure 1F). At 20°C, cultures without Cr(VI) under EAL conditions utilized lactate and produced equimolar levels of acetate until sulfate was depleted (Figure S1E). In the presence of 50 µM Cr(VI) under EAL conditions, lactate and sulfate were not utilized during the growth lag, and growth coincided with the utilization of lactate and sulfate after 200 h (Figure S1F). 48 Temperature also affected the ability of D. vulgaris to reduce Cr(VI) under EAL conditions. At 30°C, 50 µM Cr(VI) was reduced at an approximate rate of 21.92±0.07 µM Cr(VI)/hr (average O.D. of 0.08) and at 20°C 50 µM Cr(VI) was reduced at an initial rate of 10.15±3.40 µM Cr(VI)/h, which then slowed to 0.27±0.05 µM Cr(VI)/h (average O.D. of 0.08). At 30°C, 100 µM Cr(VI) was reduced at an initial rate of 26.28±1.12 µM Cr(VI)/h that slowed to 0.30±0.19 µM Cr(VI)/h (average OD 0.08) and at 20°C, 100 µM Cr(VI) was initially reduced at a rate of 7.64±2.47 µM Cr(VI)/h, that slowed to 0.45±0.15 µM Cr(VI)/h (average O.D. 0.07). Ultimately, under the EAL condition and 20°C, 50 µM Cr(VI) caused an extended growth lag (approximately 200 h) with prolonged exposure to higher Cr(VI) levels compared to the balanced and EDL conditions. Viability To assess if Cr(VI) caused more cell death in EAL grown D. vulgaris compared to cells under BAL conditions, the most probable number method was used to compare viability during the lag phases at 20°C. Viability after three hours of exposure to Cr(VI) was two orders of magnitude lower under EAL conditions compared to BAL conditions (4.7 x 103 cells/ml vs. 4.7 x 105 cells/ml). At 48 hours post-Cr(VI) exposure, the cells under both conditions were starting to recover, but the EAL cells still had decreased viability (2.1 x 104 cells/ml vs. 1.2 x 106 cells/ml). Viability for both EAL and BAL cells was similar under the respective condition without Cr(VI) (approximately 108 cells/ml). 49 Growth with Increasing Sulfate Concentrations To explain the observation that sulfate limitation (EAL condition) resulted in increased Cr(VI) susceptibility, it was hypothesized that chromate competed with sulfate through sulfate permeases/transporters in D. vulgaris under the tested growth conditions. Previous work in different organisms has shown that the cellular uptake of chromate oxyanions can occur through sulfate permeases (Pepi and Baldi 1992; Appenroth et al. 2008; Aguilar-Barajas et al. 2011) and chromate was previously shown to block sulfate accumulation and reduction completely in Desulfovibrio desulfuricans (Cypionka 1989). To determine if increased sulfate levels could alleviate intracellular stress and damage under the EAL condition, cells were inoculated into medium that contained 50 mM lactate, increasing sulfate concentrations (10, 20, 30, and 50 mM sulfate) and 50 µM Cr(VI). In addition, the inoculum cells were either grown in a BAL or EAL condition. Results showed that increasing sulfate concentration did not provide protection from Cr(VI) toxicity, but inoculum grown under EAL conditions (Figure 3A) showed an increased lag time when inoculated into EAL medium compared to inoculum grown under BAL conditions was inoculated into EAL medium (Figure 3B). These results indicated that the electron donor to acceptor ratio that D. vulgaris is grown in prior to Cr(VI) exposure can impact the cellular response in a way that predisposes the cells to increased Cr(VI) susceptibility. 50 Figure 3. Growth at 20°C with 50 µM Cr(VI) and increasing sulfate levels of 10 mM (●), 20 mM (◻), 30 mM (■), and 50 mM (○) inoculated with cells grown under EAL conditions (a) or BAL conditions (b). Growth with Normalized Sulfate Levels Because D. vulgaris was the most affected by Cr(VI) under the EAL condition (50:10 mM), Cr(VI) exposure was normalized to nutrient ratios with 10 mM sulfate in order to see if increased Cr(VI) susceptibility is caused by sulfate concentration or the ratio of lactate to sulfate (BAL -20:10 mM; EDL - 5:10 mM, and EAL -50:10 mM). The 20:10 balanced condition lagged for approximately 100 hours similar to the 60:30 mM ratio (Figure 4). These results indicate that it is not only the sulfate concentration that affects Cr (VI) toxicity, but also the ratio of electron acceptor to electron donor. 51 Figure 4. Growth at 20°C with 50 µM Cr(VI) and lactate to sulfate ratios normalized to 10 mM sulfate. Normalized ratios were EDL (5 mM: 10 mM - ●), BAL (20 mM:10 mM - X), and EAL (50 mM:10 mM - ■). Sulfate Permease Mutants To further understand the role of sulfate permeases in exposure to Cr(VI) in D. vulgaris, a search of the annotated genome revealed three genes designated as sulP that are presumptive sulfate permeases in D. vulgaris Hildenborough (microbesonline.org). Three individual sulfate permease mutants (∆DVU0053, ∆DVU0279, ∆DVU1999) and a triple mutant in which all three sulfate permease genes were knocked out were Time (h) O pt ic al D en si ty (6 00 nm ) 52 constructed. The wild-type, three individual mutants, and the triple mutant had similar growth rates and yields when grown via lactate-dependent sulfate respiration without Cr(VI) at 20°C under BAL conditions (Figure 5, data not shown for individual mutants). These results indicated that D. vulgaris Hildenborough has other mechanisms of sulfate influx (specific and/or non-specific) in addition to the presumptively annotated transporters. When grown in the presence of 50 µM Cr(VI), the triple mutant had a longer lag time by approximately 40 hours compared to wild-type cells grown under BAL conditions (Figure 5). These results indicated that the absence of the annotated sulfate permeases did not prevent Cr(VI) from entering and harming the cells and that triple mutant cells had decreased ability to regulate Cr(VI) influx and/or sulfate influx even under balanced conditions. The results suggested that the unidentified mechanism(s) by which Cr(VI) can enter the cell are not as tightly linked to Cr(VI) reduction and/or the alternative mechanisms for sulfate transport have a strong affinity for Cr(VI) that results in increased Cr(VI) sensitivity. 53 Figure 5. Growth of wild type D. vulgaris (○) and sulfate permease triple mutant (●) without Cr(VI) and with 50 µM Cr(VI) (◻,■) at 20°C under BAL conditions. Ascorbate D. vulgaris was also grown with Cr(VI) in the presence of ascorbate to determine if a reducing/complexing agent could prevent or alleviate Cr(VI) cellular toxicity. Cell growth was compared between cells grown with and without 50 µM Cr(VI), with 50 µM Cr(VI) and 50 µM ascorbate added concurrently, with 50 µM Cr(VI) and 50 µM ascorbate added 3 hours post inoculation, and with 50 µM ascorbate only. When Cr(VI) and ascorbate were added concurrently, D. vulgaris cells grew similar to the control without Cr(VI), indicating that the reduced/complexed Cr compounds were not toxic and cells were most likely able to maintain viability (Figure 6). If ascorbate was added 3 hours post-inoculation, the cells were still impacted, but not as drastically as without 54 ascorbate. Approximately 30 µM Cr(VI) remained in solution when the ascorbate was added at the 3 h time point, which might explain why the lag time for this condition was shorter than the condition in which 50 µM Cr(VI) was added without ascorbate. Figure 6. Growth of D. vulgaris during Cr(VI) and ascorbate exposure at 20°C under EAL conditions. Cells were exposed to 50 µM Cr(VI) and 50 µM ascorbate (■), 50 µM Cr(VI) added prior to inoculation and 50 µM ascorbate added at 3 h post-inoculation (●), 50 µM ascorbate (△), 50 µM Cr(VI) (○), or medium only (◻). Discussion The mechanism for Cr(VI) reduction in Desulfovibrio spp. is not well-understood, but it is hypothesized that Cr(VI) enters the cell and is reduced to Cr(III) by cytochrome c3, periplasmic hydrogenases, and possibly thioredoxin reductase (Chardin et al. 2003; Lovley and Phillips 1994; Li and Krumholz 2009). After the Cr(VI) is reduced to Cr(III), 55 the Cr(III) is either excreted from the cell or remains in the inner or outer membrane of the cell (Goulhen et al. 2005). It has also been shown that the presence of Cr(VI) alters the metabolism of Desulfovibrio spp. by transiently uncoupling lactate oxidation and sulfate reduction and causing lactate to be used for energy production or to lower the redox potential of the medium without concurrent growth at 30°C (Chardin et al. 2002; Klonowska et al. 2008). The redox couple for Cr(VI) reduction to Cr(III) is 1.41V (Nriagu and Nieboer 1988); however, D. vulgaris Hildenbourough is not known to conserve energy from Cr(VI) reduction and most data indicate that direct and indirect Cr(VI) reduction is linked to detoxification (Chardin et al. 2002; Klonowska et al. 2008). However, microbial fuel cells have recently been shown to reduce Cr(VI) with anode electrons (Habibul et al. 2016); and therefore, improved understanding of electron movement from the bulk phase to heavy metals under different conditions is needed. Waters contaminated with heavy metals can be deficient in electron donors, electron acceptors, and/or carbon sources related to desired microbial activities, and the exact relationship between growth, activity, and substrate utilization is more challenging to understand when the activity of interest is not directly linked to growth. Previous studies have compared different electron donors ranging from organic acids, alcohols, carbohydrates, and polysaccharides to stimulate Cr(VI) reduction with pure cultures, mixed consortia, and in situ (Liamleam and Annachhatre 2007; Zhang et al. 2015). Geets et al. (2006) tested the removal of Zn, Cd, Co, and Ni from contaminated groundwater in 56 column experiments and concluded that additional experiments were needed to better understand the interplay between sulfate-, metal-, and carbon source concentrations. The electron donor to electron acceptor ratio had a very strong effect on Cr(VI) reduction by D. vulgaris at both 20° and 30°C, with the EAL condition at 20°C being the most detrimentally affected. To explain this, we hypothesized that because chromate (CrO42-) and sulfate (SO42-) are structural analogues, the amount of sulfate in the environment affects the rate at which sulfate versus chromate enters the cell. It has been shown that Cr(VI) can enter cells through sulfate transport systems in numerous bacteria including, Pseudomonas fluorescens, Salmonella typhimurium, and Escherichia coli (Pardee et al. 1966; Ota et al. 1971; Karbonowska et al. 1977; Ohtake et al. 1987; Sirko et al. 1990) and the slightly longer lag time in the sulfate permease triple mutant in D. vulgaris compared to the wild type alludes to these three permeases playing a role in sulfate and/or Cr(VI) transport. However, the effect does not seem to be merely a sulfate concentration threshold, because the EDL condition has more sulfate compared to the BAL condition (50 mM vs 30 mM) and yet the EDL condition has a longer lag time (at 20°C) compared to the BAL condition. It is important to note that the three resource ratios have similar growth rates without Cr(VI) at 20° and 30°C. In addition, when the resource ratios were normalized to sulfate level (5:10 mM versus 20:10 mM versus 50:10 for EDL, BAL, and EAL respectively), similar results were observed between EAL and BAL, and EDL had even a longer lag time. These results suggested that the co-metabolic 57 rate of lactate and sulfate utilization influences Cr(VI) influx and reduction, and thus, the subsequent overall cellular toxicity. To further explore the hypothesis that limiting sulfate increases Cr(VI) toxicity in D. vulgaris due to competition between sulfate and chromate for transport across the cell membrane, growth experiments with increasing levels of sulfate and sulfate permease mutants were done. The results of the increasing sulfate experiment support our finding that cells grown under EAL conditions are more susceptible to Cr(VI) toxicity than those grown under BAL conditions. A previous transcriptomic study on D. vulgaris transitioning from exponential growth to stationary phase shows changes in expression of the annotated sulfate permease genes and other cell components for lactate oxidation as the cells transition from exponential to stationary phase (i.e., decreasing sulfate levels), indicating that perhaps different cell machinery is used to optimize the utilization of electron donors and acceptors (i.e., presumptive sulfate permeases have different affinities for sulfate) (Clark et al. 2006). Deletion of the three presumptive sulfate permease genes did not affect growth with lactate and sulfate, indicating that sulfate had additional ways to enter the cell. The triple mutant did have an increased lag time when cells were exposed to 50 µM Cr(VI), and this result suggested that the triple mutant had either a decreased affinity/capacity for sulfate influx and/or increased affinity/capacity for chromate influx. An alternative explanation could be an overall decreased metabolic flux due to limited transport, but the triple mutant had similar growth rates in the absence of 58 Cr(VI). Further work is needed to elucidate the additional mechanisms of sulfate transport in Desulfovibrio and the role of these transporters in Cr toxicity. The decrease in biomass yield based on sulfate consumption when cells were exposed to 50 µM Cr(VI) could be explained by the expenditure of energy toward Cr(VI) detoxification and cellular repair from damaging Cr(VI) reduction intermediates instead of allocation towards growth. To determine whether the increased lag time under the EAL condition compared to the BAL condition was due to Cr(VI) causing a greater decrease in cell viability, we measured cell numbers throughout the lag phase in both EAL and BAL conditions with 50 µM Cr(VI). Results indicate that the increased lag time under EAL conditions was due to decreased cell viability, which indicates that Cr(VI) toxicity was more extensive in cells grown in the EAL condition compared to the BAL condition. The interplay between chromate and sulfate levels is most likely a result of a balance between Cr(VI) influx and the ability of cells to reduce Cr(VI) to the less toxic Cr(III), and this detoxification requires adequate reducing potential. The cellular response to Cr(VI) at different resource ratios suggests that D. vulgaris modulates metabolic flow in response to the ratio of e-/C levels and e- acceptors, and further work is needed to elucidate the mechanisms of metabolic control and the implications for desired activities. The increase in Cr(VI) toxicity for sulfate/lactate limited cells has large implications for applications such as bioremediation where carbon and electron sources are injected into the subsurface, most likely creating unbalanced pairings of energy/C sources (Figure 7). 59 Figure 7. Three potential scenarios (EDL, BAL, EAL) in which nutrients are added to stimulate microbial activity and the ratio of resources are shifted. The different resource ratios can impact desired microbial activity. In the subsurface, sulfate-reducing bacteria can account for a majority of biotic metal reduction via direct and indirect mechanisms, but little work has investigated the implications of biostimulation practices on microbial physiology and activity with respect to changing resource ratios and other environmentally-relevant conditions (i.e., temperature). 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Name Sequence* Purpose DVU0053-1b CGGGAAAGACCTCCGCCTTG Cloning of upstream region of DVU0053 DVU0053-2 AAGACTGTAGCCGTACCTCGAATCTA CGTCCCGTGTTCCCTGTTTG Cloning of upstream region of DVU0053, with overhang for annealing to kanamycin-resistance gene DVU0053-3 AATCCGCTCACTAAGTTCATAGACCG CATATGGGTGCTGTCAGGTC Cloning of downstream region of DVU0053, with overhang for annealing to 68 kanamycin-resistance gene DVU0053-4 AGCAGCCCATGTTGAGGTCG Cloning of downstream region of DVU0053 bc0050f TAGATTCGAGGTACGGCTACAGTCTT ATCTCTGAAGAAGCCGACAC CCCCAGAGTCCCGCTCAG Amplification of kanamycin-resistance gene bc0050r CGGTCTATGAACTTAGTGAGCGGATT AGTAACAGTCGTGAACATCG GAGGTAGCTTGCAGTGGGCT Amplification of kanamycin-resistance gene DVU0279-1 GACTGCGGGAGCATCATGCG Cloning of upstream region of DVU0279 DVU0279-2 AAGACTGTAGCCGTACCTCGAATCTA ATGCGCCTCCTTTGCGATTT Cloning of upstream region of DVU0279, with overhang for annealing to kanamycin-resistance gene DVU0279-3 AATCCGCTCACTAAGTTCATAGACCG CGTTGATGACAGACGTGACG Cloning of downstream region of DVU0279, with overhang for annealing to kanamycin-resistance gene DVU0279-4 ATGAGATTCGCGCCCTGTAC Cloning of downstream region of DVU0279 bc0051-f TAGATTCGAGGTACGGCTACAGTCTT ATCGAACTAACGTACATGCC CCCCAGAGTCCCGCTCAG Amplification of kanamycin-resistance gene bc0051-r CGGTCTATGAACTTAGTGAGCGGATT ATCGCCTAACCTAGATACAG GAGGTAGCTTGCAGTGGGCT Amplification of kanamycin-resistance gene DVU1999-1 CCCCAAACCCCATCTCGATCGAG Cloning of upstream region of DVU1999 DVU1999-2 AAGACTGTAGCCGTACCTCGAATCTA CTGGGGAGACGTTGCGTCTT Cloning of upstream region of DVU1999, with overhang for annealing to kanamycin-resistance gene DVU1999-3 AATCCGCTCACTAAGTTCATAGACCG ACCCGACAGTGAGCCGCCAG Cloning of downstream region of DVU1999, with overhang for annealing to kanamycin-resistance gene DVU1999-4 ATGATTTGGGCGGCTTCGGC Cloning of downstream region of DVU1999 bc0052-f TAGATTCGAGGTACGGCTACAGTCTT GTGTGACATGCTGCTAGAAC CCCCAGAGTCCCGCTCAG Amplification of kanamycin-resistance gene bc0052-r CGGTCTATGAACTTAGTGAGCGGATT GCGTCGTAATAGTGGTTATC GAGGTAGCTTGCAGTGGGCT Amplification of kanamycin-resistance gene pMR- >pMLD-Km GAACACGGCGGCATCAGAG Amplification of marker-replacement plasmid pMR- >pMLD-Cm GCACCAAGTAAGACTGTAGCCGTACC TCGAATCTA Amplification of marker-replacement plasmid KanR-upp- pMR-F AACAGACAATCGGCTGCTCTGATG Amplification of kanr-upp cassette KanR-upp- pMR-R TAGATTCGAGGTACGGCTACAGTCTT ACTTGGTGCCGAATATCTTGTCGCC Amplification of kanr-upp cassette; contains overhang for common sequence DVU0053- GGAACACGGGACGCATATGGGTGCTG Amplification from marker-replacement 69 MLD-F TCAGGTCTTCG plasmid to construct the marker-less deletion plasmid for DVU0053 DVU0053- MLD-R CAGCACCCATATG CGTCCCGTGTTCCCTGTTTGC Amplification from marker-replacement plasmid to construct the marker-less deletion plasmid for DVU0053 DVU0279- MLD-F AAAGGAGGCGCATCGTTGATGACAGA CGTGACGTTCC Amplification from marker-replacement plasmid to construct the marker-less deletion plasmid for DVU0279 DVU0279- MLD-R TCTGTCATCAACG ATGCGCCTCCTTTGCGATTT TCC Amplification from marker-replacement plasmid to construct the marker-less deletion plasmid for DVU0279 DVU1999- MLD-F AACGTCTCCCCAG ACCCGACAGTGAGCCGCCAG Amplification from marker-replacement plasmid to construct the marker-less deletion plasmid for DVU1999 DVU1999- MLD-R CTCACTGTCGGGT CTGGGGAGACGTTGCGTCTT Amplification from marker-replacement plasmid to construct the marker-less deletion plasmid for DVU1999 * - underlined region represents overhang used for annealing to neighboring PCR product in SOE and SLIC reactions. Table S2. Primers used in this study. 70 CHAPTER THREE NUTRIENT LIMITATION CAUSES DECLINE IN METABOLITES IMPORTANT FOR CELL CYCLE PROGRESSION IN BACTERIAL BIOFILM Contribution of Authors and Co-Authors Author: Lauren Franco Contributions: Experimental design, performed experiments, data analysis, wrote and revised manuscript Co-Author: Julijana Ivanisevic Contributions: Experimental design, performed experiments, data analysis, wrote and revised manuscript Co-Author: Gary Siuzdak Contributions: Experimental design, data analysis, revised manuscript Co-Author: Matthew W. Fields Contributions: Experimental design, performed experiments, data analysis, wrote and revised manuscript 71 Manuscript Information Page Lauren C. Franco, Julijana Ivanisevic, Gary Siuzdak, Mathew W. Fields Applied Microbiology and Microbiology Status of Manuscript: __x_ Prepared for submission to a peer-reviewed journal ____ Officially submitted to a peer-review journal ____ Accepted by a peer-reviewed journal ____ Published in a peer-reviewed journal 72 Abstract The effect of nutrient limitation on Desulfovibrio vulgaris Hildenborough biofilm growth and metabolism is investigated in this chapter. D. vulgaris was grown in a CDC reactor under electron donor-limitation (EDL), electron acceptor-limitation (EAL) and electron donor to electron acceptor balanced (BAL) conditions. Biofilm growth was measured via protein and carbohydrate and were used to make biomass yield calculations based on lactate and sulfate consumption. Growth measurements normalized to protein and biomass yields did not differ between the three conditions. Although biofilm growth was not significantly affected by nutrient limitation, metabolomic analysis revealed significant dysregulation between D. vulgaris biofilm grown under the EAL condition compared to that grown under the BAL condition. Biofilm grown under the EAL condition had an overall down-regulation of metabolites compared to the BAL condition. Of those that could be confidently identified, metabolites involved in purine and pyrimidine synthesis, glutamine and glutamate synthesis, and peptidoglycan synthesis were down-regulated under the EAL condition, indicating an overall decrease in cell division and growth compared to the BAL condition. Interestingly, fatty acid metabolites were the majority of up-regulated metabolites that could be identified under the EAL condition compared to the BAL condition. Taken together, these results point to a metabolic switch from growth to biomass maintenance caused by electron acceptor- limitation. The results highlight the need to further explore the role of the fatty acids produced in D. vulgaris biofilm grown under the EAL condition and further emphasize the effects that nutrient limitation can have on microbial metabolism and physiology. 73 Introduction The recognition of changing environmental conditions, such as nutrient availability, is essential to organismal growth and survival, and microorganisms are acutely adept at maximizing available resources for biomass and energy conservation. However, most microbial studies have been conducted on planktonic cells grown in suspension due to utility for study and sampling, and not the more predominant life-style of attached growth (i.e., biofilm). Bacteria have the ability to sense extracellular signals, via two-component regulatory systems for example, and can then feed those signals into transcriptional regulatory cascades, resulting in a change in gene expression and subsequent adjustment of metabolism to better perform under given external constraints. Another layer of extracellular sensing is the cell’s ability to coordinate control of multiple inputs and adjust accordingly. For example, Escherichia coli carbon utilization can be coordinated with nitrogen availability, and carbohydrate transport can be coordinated with the regulation of carbohydrate-utilizing enzymes (i.e., glucose and the Lac operon). In the instance of coordinated N and C utilization, α-ketoglutarate inhibits enzyme I of the phosphotransferase system, and subsequently glucose uptake is adjusted in response to N availability (Doucette et al. 2011). Thereby, using the α-ketoglutarate/glutamate metabolic node of the TCA (tricarboxylic acid cycle) to help control influx of carbon (i.e., glucose) in light of available N in terms of α-ketoglutarate/glutamate/glutamine. In another example, carbon and sulfur metabolisms are linked by CysB, a transcriptional 74 regulator of the sulfur metabolism in E. coli and Salmonella that regulates an operon encoding genes required for uptake of sulfur sources and for the synthesis of cysteine. When cysB is mutated, the organism has 50% less activity of enzymes involved in carbon source uptake and utilization (Quan et al. 2002). In D. vulgaris, dissimilatory sulfate reduction and carbon uptake and utilization are presumably coordinated as well, although the mechanism has not yet been elucidated. One potential player in this coordination is RexB, a repressor of sulfate adenylyl transferase (sat), the gene that encodes the enzyme responsible for activating sulfate in the first step of sulfate reduction. RexB is regulated by cellular levels of NAD/NADH+H+ in that NADH+H+, but not NAD+, binds with RexB and prevents it from binding DNA. Elevated intracellular levels of NADH+H+ therefore prevent RexB from repressing sat transcription, allowing the first step of sulfate reduction to commence (Christensen et al. 2015). While RexB certainly links sulfate reduction to the redox state of the cell through the NAD/NADH+H+ ratio, it could also be a link to the central carbon metabolism, as NADH is a product of the TCA cycle. Although it has been shown in D. vulgaris that the type of electron donor/carbon source and electron acceptor affects physiological responses to Cr(VI) and electron flow within the cell, little work has been done on how altering the electron donor/carbon source concentration affects its metabolism (Klonowska et al. 2008; Zhou et al. 2017). Moreover, limited work has been done with diverse microorganisms to better understand the overall metabolic state of biofilms. The work presented in this chapter investigates 75 how nutrient limitation, in the form of sulfate limitation, affects the metabolism and growth of D. vulgaris Hildenborough in the biofilm growth mode. Materials and Methods Bacterial Strains and Growth Conditions D. vulgaris Hildenborough was grown in either batch or continuous mode. For batch growth, cells were inoculated into sterile N2 gassed balch tubes containing LS4D, a lactate and sulfate medium (modified to 2.5 µM resazurin and 130 µM riboflavin in Thauer’s vitamins) (Brandis and Thauer 1981; Borglin et al. 2009). The medium consisted of varying lactate and sulfate concentrations as follows: 60 mM sodium lactate and 30 mM sodium sulfate (balanced condition), 50 mM sodium lactate and 10 mM sodium sulfate (electron acceptor limited condition), and 18 mM sodium lactate and 50 mM sodium sulfate (electron donor limited condition). For continuous culture of D. vulgaris biofilm, cells were grown in a CDC reactor modified with continuous sparging of the headspace and medium with N2. Exponential phase cells were inoculated into a CDC reactor containing electron acceptor limited, balanced, or electron donor limited LS4D medium. Cells were grown in batch mode for 48 hours and then continuous mode with a dilution rate of 0.04-hr. Reactors were grown at room temperature (20-23°C), stirred at 60 rpm, and the headspace was continuously sparged with sterile N2 gas to maintain anaerobic conditions. Coupons of glass slides were submerged in the reactor 76 body as a surface for biofilm growth. Sample Collection Biofilm samples were collected for biomass analysis at 168 hours (7 days). For biomass analyses glass slide coupons were removed from the reactor, gently submerged in PBS to remove any loosely attached biomass, and then biomass was scraped from both sides of the slide with a razor blade into 1 ml degassed water. Sulfide content was measured immediately after sampling, samples for protein and carbohydrate were measured after one freeze-thaw cycle to lyse cells. Biomass Analyses Protein was measured using the Qubit protein assay (Life Technologies, Carlsbad, CA). Carbohydrate in the form of hexose was measured using a colorimetric L-cysteine sulphuric acid assay with glucose as the standard (Chaplin and Kennedy 1986). Dissolved and precipitated sulfide was measured as described (Cord-Ruwisch 1985). All assays were performed in triplicate and error bars represent standard deviation. Results were compared using the unpaired student’s t-test. Biomass yields were calculated by summing all of the biomass (biofilm and planktonic) in individual CDC reactors and dividing that by the amount of substrate consumed. Metabolomic Analysis of Biofilm Biofilm samples for metabolite analysis were harvested from reactors at 168 hrs, 77 quickly rinsed in ice cold 1:10 diluted phosphate buffered saline, flash frozen in liquid nitrogen, and stored at -80°C. Samples were extracted and analyzed at the Scripps Center for Metabolomics using an untargeted approach with a dual separation method (reversed- phase liquid chromatography and hydrophilic interaction chromatography) to maximize metabolite separation and recovery (Ivanisevic et al. 2013). Metabolite features were confirmed via tandem mass spectrometry (MS/MS) and identified by matching a standard in the METLIN database (Benton et al. 2015). Results Biofilm Growth under Nutrient-Limited and Balanced Conditions D. vulgaris growth under continuous flow conditions as a biofilm was affected by altering electron donor and electron acceptor concentrations. More biomass, as quantified by protein, was produced under the balanced condition (47.6 ± 2.9 µg/cm2) than under limiting conditions (22.1 ± 3.3 µg/cm2 for EAL and 21.9 ± 1.4 µg/cm2 for EDL) (Figure 1). This trend mimics the biomass production for D. vulgaris grown under batch planktonic conditions in which cells under the balanced condition reached a higher maximum O.D. compared to EAL and EDL (Chapter 2). Carbohydrate, as a measure for extracellular polymeric substances produced by the biofilm, also differed according to electron donor and acceptor limitation, but when normalized to protein, there was not a statistically significant difference between the three conditions (g carbohydrate/g protein 78 = 0.08±0.015 for EAL, 0.07±0.006 for BAL, and 0.06±0.005 for EDL; unpaired student’s t-test p-value < 0.05 for all comparisons). Sulfide concentration within the biofilm biomass varied according to electron donor and electron acceptor concentration as well, but when sulfide was normalized to protein, there was no statistically significant difference between the three conditions (g sulfide/ g protein = 4.56±0.6 for EAL, 3.32±0.5 for BAL, 4.21±0.1 for EDL unpaired student’s t-test p-value < 0.05 for all comparisons). Additionally, biomass yields for lactate and sulfate that included the biofilm and planktonic phase biomass, were not statistically different between the three conditions (Table 1) and the proportion of biomass in the biofilm phase compared to the planktonic phase was between 5 and 6% for all three conditions. Figure 1. Protein, carbohydrate, and sulfide concentrations in D. vulgaris biofilm grown under EAL, BAL, and EDL conditions. 79 Lactate: Sulfate Concentration (mM) Yield (g Protein/g Lactate) Yield (g Protein/g Sulfate) Electron acceptor limited (EAL) 50:10 0.059 ± 0.006 0.113 ± 0.014 Balanced (BAL) 60:30 0.057 ± 0.004 0.112 ± 0.008 Electron donor limited (EDL) 18:50 0.066 ± 0.002 0.125 ± 0.003 Table 1. Lactate and Sulfate concentrations for the different nutrient ratios and biomass yields based on lactate and sulfate consumption. Metabolomic Analysis of D. vulgaris Biofilm To understand how nutrient limitation affects the metabolism of D. vulgaris biofilm, an untargeted metabolomic method was implemented. Overall, there were 552 dysregulated features with a p-value less than 0.01 and a fold-change greater than 1.5 when D. vulgaris biofilm grown under EAL was compared to that grown under BAL (Figure 2). 80 Figure 2. Cloud plot representation of dysregulated metabolites with a greater than 1.5 fold-change and p-value less than 0.01. Size of dot represents fold-change, with larger diameters representing greater fold change, and intensity of color represents p-value, with darker intensities representing smaller p-values. Metabolites that were down-regulated under the EAL condition compared to the BAL condition fell into the categories of purine and pyrimidine metabolism, glutamine and glutamate metabolism, and peptidoglycan metabolism. Individual metabolites within these pathways are listed in Table 2 and pathways containing more than one down- regulated metabolite are depicted in Figure 3 (adapted from KEGG (Kanehisa 2000; Kanehisa et al. 2016; Kanehisa et al. 2017)). 81 Surprisingly, metabolites that were up-regulated under the EAL condition were almost entirely different types of fatty acids. These fatty acids are listed in Table 3. Taurine and possibly 3-deoxy-d-manno-octulosonate were also up-regulated under the EAL condition, but 3-deoxy-d-manno-octulosonate, a Lipid A precursor, could not be confirmed by MS/MS because there is no standard in the database. The up-expression of fatty acids in D. vulgaris biofilm is investigated further in Chapter 4. Metabolite ID Fold Change p-value Purine Metabolism Inosine 22.2 1.20E-03 Hypoxanthine 7.8 1.35E-05 Adenylylsulfate 6.5 1.10E-04 Guanosine 6.3 1.60E-02 Adenosine monophosphate (AMP) 5.5 3.70E-03 Guanine 4.3 9.00E-04 Adenine 2.7 6.00E-04 Pyrimidine Metabolism Uridine diphosphate (UDP) 13.3 4.00E-03 Glutamate 8.0 2.30E-02 Uracil 6.3 9.00E-05 Uridine monophosphate (UMP) 3.3 1.50E-03 UDP-glucose 3.2 2.00E-02 Cytidine monophosphate (CMP) 2.0 2.50E-02 Glutamine and Glutamate Metabolism 82 UDP-MurNAc-Ala-Glu 13.2 2.40E-03 D-Glutamine 8.0 2.30E-02 UDP-N-Acetyl-muramate 4.6 3.50E-03 L-Glutamate 3.7 2.20E-03 Peptidoglycan UDP-MurNAc-Ala-Glu 13.2 2.40E-03 Alanine 6.0 7.00E-06 UDP-N-Acetyl-muramate 4.6 3.50E-03 UMP 3.3 1.50E-03 Other Tryptophan 14.1 2.90E-03 2-C-Methyl-D-erythritol 2,4-cyclodiphosphate 10.7 8.50E-07 Lysophospholipid 9.4 3.00E-05 Methionine 7.8 2.70E-03 Methylthiouracil 6.6 4.00E-03 Lysophospholipid (17:1/0:0) 6.5 9.00E-04 Phosphoethanolamine (8:0/8:0) 6.0 2.00E-05 Aspartic acid 5.5 5.00E-06 Phosphatidylglycerol (15:1/0:0) 4.3 9.00E-04 NAD 4.0 1.00E-04 GDP-fucose 2.9 5.00E-03 Phenylalanine 2.5 5.80E-03 Phosphoethanolamine (8:0/8:0) 1.5 3.00E-02 Table 2. List of metabolites down-regulated under the EAL condition categorized by metabolic pathways they are linked to and associated fold change and p-value. 83 Metabolite ID Fold Change p-value Fatty Acids Methyl-heptanoic acid 9.3 3.0E-03 Methyl-hexanoic acid 6.7 6.0E-03 Methyl-octanoic acid 5.2 5.0E-03 Decanoic acid 4.0 5.0E-03 Dihydroxy-octadecadienoic acid 3.8 6.0E-03 Hydroxy-octadecadienoic acid 3.5 5.0E-03 Methyl-decanoic acid 2.3 8.0E-03 Dodecanoic acid 2.0 2.0E-03 Methyl-hexadecanoic acid 1.7 8.0E-03 Hydroxy-undecenoic acid 1.6 8.0E-03 Hydroxy-hexadecanoic acid 1.4 4.0E-03 Methyl-tetradecanoic acid 1.3 8.0E-03 Other Taurine 2.9 3.00E-03 3-Deoxy-D-manno-octulosonate 1.9 7.20E-03 Table 3. List of metabolites up-regulated under the EAL condition, with associated fold change and p-value. 84 85 Figure 3. Metabolic pathways with multiple down-regulated metabolites, highlighted in yellow, under EAL conditions. Green boxes represent genes/proteins that are annotated in the D. vulgaris Hildenborough genome. Figures adapted from KEGG. 86 Discussion Nutrient limitation, in the form of electron donor/carbon source or electron acceptor limitation, had notable effects on biofilm metabolism and growth. Growth, as measured by protein biomass, reached a higher density under the BAL condition, as was expected given that the BAL condition provides ~three times the amount electron flux compared to the EAL and EDL conditions (240, 80, and 72 electron equivalents, for BAL, EAL, and EDL, respectively). Although there was not three times the amount of biofilm biomass produced in accordance with the difference in lactate and sulfate concentration for the BAL condition, the biomass yields for the entire reactor (biofilm plus planktonic phase) for EAL, BAL, and EDL were similar to each other, indicating that the biomass produced, in either planktonic or biofilm phase, was consistent with how much lactate and sulfate was provided. Carbohydrate did not contribute a large portion of the total biofilm biomass under any of the three conditions, which is consistent with results from Clark et al., in which the authors analyzed D. vulgaris Hildenborough biofilm grown at 30°C (Clark et al. 2007; Clark et al. 2012). Despite similar biofilm biomass yields, there was a considerable amount of dysregulation in D. vulgaris biofilm grown under EAL compared to the BAL condition. Overall, the down-regulated metabolites for biofilm grown under EAL indicated that cellular growth was suppressed compared to biofilm under the BAL condition through the dysregulation of metabolites involved in purine and pyrimidine synthesis (for 87 nucleotide production), glutamine and glutamate synthesis (for amino acid production, purine and pyrimidine production, and NAD production), and peptidoglycan synthesis (for cell wall production). Under the condition of lower energy state (EAL), the data suggest that biofilm cells slow down “growth” related resource expenditure by decreasing the production of precursors for purine and pyrimidine biosynthesis and cell well biosynthesis. Because the comparison was between a nutrient limited biofilm and a nutrient balanced biofilm in reactors at the same dilution rate, the data suggest that the biofilm responded to energy limitation by decreasing metabolites important for new cell biosynthesis (i.e., purines, pyrimidines, cell wall precursors). This could be analogous to cell cycle check-points that stop or slow the cell cycle in eukaryotic cells under unfavorable conditions. Both glutamate and glutamine are crucial precursors to the de novo synthesis of both purines and pyrimidines as well as important amino acids for the coordination of N and C flux. Under the tested EAL condition, C and N are not limiting; however, electron acceptor is limiting and thus electron flux is lower. These results indicate that energy limitation causes a decline in precursors for purines, pyrimidines, and cell wall biosynthesis. This physiological biofilm state under energy limitation could be similar to a quiescent (Q0) or quiescent- like state in eukaryotic cells, and could possibly contribute to the increased tolerance of biofilms to stresses (e.g., chemicals, metals, antibiotics). It has become increasingly evident that metabolic pathways are integrated and coupled to cell cycle progression and work in higher eukaryotes has shown that cell cycle progression and metabolism are 88 linked to body fat content in mice (Naaz 2004) as well as type II diabetes in humans (Saxena et al. 2007). However, little is known of the mechanisms involved and if bacteria and archaea display similar controls. It is typically accepted that cells must sense the energetic status, and the energetic status is linked to the growth and cell cycle. Two commonly studied systems in eukaryotes are the AMP-activated protein kinase (AMPK) and mTOR (a serine/threonine kinase). It is not known if similar functions exist in bacteria and archaea, although interestingly, of the approximately 100 genes that were related to metabolic periodicity with the yeast cell cycle, two-thirds were involved in mitochrondrial function (Lee and Finkel 2013) Few studies have used comprehensive metabolomics to characterize biofilm physiology and responses to nutrient deprivation. Interestingly, the metabolomic data suggested important roles for nucleotides and cell wall precursors. For purines, guanosine, guanine, inosine, hypoxanthine, and adenine were at lower levels compared to BAL conditions. Typically, inosine is a precursor that can lead to xanthine, guanine, and adenine. For pyrimidines, uridine monophosphate (UMP) was lower in EAL biofilms, and typically UMP is the precursor from which cytosine and thymine are produced. Glutamine and glutamate were also both lower in EAL biofilms, and these amino acids link DNA/RNA and cell wall turnover in several ways. Glutamine and glutamate are important for de novo nucleotide biosynthesis (amino donors), and the amino acids (D and L forms) can be placed into the peptide linkages between the glycan strands of the 89 cell wall. In addition, uridine-5’-triphosphate (UTP) serves as a donor for urdylated precursors during cell wall biosynthesis, and both UDP-N-acetyl-muramate and UDP-N- acetyl-muramate-A-E-A were at lower levels in the EAL biofilms. Moreover, a key intermediate to the biosynthesis of undecaprenyl-P, 2-C-methyl-D-erythritol 2,4- cyclodiphosphate, was depleted in EAL biofilms. Undecaprenyl-P is a crucial lipid carrier for cell wall biosynthesis, and previous work in Escherichia coli and Bacillus subtilis has shown loss of the synthase responsible for synthesizing this metabolite results in abnormal cell walls (Campbell and Brown 2002). In E. coli, a filamentous phenotype was observed, and in B. subtilis irregular morphology and thickened cell walls was observed. Purine and pyrimidine biosynthesis has also been linked to cell wall synthesis/rigidity in Lactococcus lactis, with mutations in the either guaA or pyrB genes causing changes in cell wall rigidity due to an accumulation of L-aspartate, which is an essential component of peptidoglycan crosslinking proteins, over conversion of L-Asp to pyrimidine precursors (Solopova et al. 2016). Purine biosynthesis has also been observed to be dysregulated during meiotic differentiation of yeast spores (Walther et al. 2014), and little is known of metabolic re-programming in bacterial biofilms or the repercussions for eukaryotic cell cycles and differentiation. In the context of the presented data with D. vulgaris, multiple metabolites crucial to DNA/RNA and cell wall turnover were dysregulated, and the data suggest a linkage between nucleotide and cell wall biosynthesis that could help coordinate the biofilm cell cycle and nutrient limitation. 90 Interestingly, the majority of the metabolites that were up-regulated under the EAL condition were fatty acids. Fatty acids are a major component of cellular membranes, but given that pathways critical to cellular growth were down-regulated, it does not make sense that metabolites associated with cellular membranes are up- regulated. This suggests that the fatty acids are not being produced as part of cellular membranes, but for another purpose. Fatty acid biosynthesis requires inputs of ATP and reducing equivalent in the form of NADPH, further supporting the idea that energy and reducing equivalents may be being used for fatty acid production rather than cellular growth. The up-regulation of fatty acids and their role in D. vulgaris biofilms grown under the EAL condition will be further explored in Chapter 4. Of the other up-regulated metabolites under the EAL condition, only one could be positively identified in METLIN and that is taurine. Taurine is a sulfonic acid containing an amino group and its biosynthesis is widely distributed in eukaryotes. Taurine biosynthesis in bacteria is not as well understood, but a pathway has been elucidated in a Synechococcus species in and there may be similar pathways in other bacteria (Agnello et al. 2013). While certain Desulfovibrio strains can use sulfonates, including taurine, for electron acceptors, utilization of taurine as an electron acceptor by D. vulgaris Hildenborough has not been demonstrated (Laue et al. 1997; Lie et al. 1999). The production of taurine under sulfate-limiting conditions raises the question of whether taurine could be produced from cysteine as an electron acceptor when sulfate is limiting. This question is not addressed in this work, but could be a future direction for research. 91 While D. vulgaris biofilm growth under nutrient limited and balanced conditions was consistent with how much substrate was provided, and carbohydrate and sulfide concentrations were approximately the same when normalized to protein, the metabolomic analysis revealed significant dysregulation of metabolic features between the sulfate-limited and balanced conditions. The differences in metabolite regulation brings to light the ability of D. vulgaris to sense carbon/electron donor and sulfate/electron acceptor and adjust its metabolism accordingly. 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Franco Contributions: Developed experimental design, performed experiments, analyzed data, wrote and revised the manuscript. Co-Author: Siva Wu Contributions: Developed experimental design, performed experiments, analyzed data. Co-Author: Michael Joo Contributions: Developed experimental design, performed experiments, analyzed data. Co-Author: Joel Mancuso, Jonathan Remis, Amita Gorur, Ambrose Leung, Danielle M. Jorgens, Joaquin Correa Contributions: Developed experimental design, performed experiments, analyzed data. Co-Author: Manfred Auer Contributions: Developed experimental design, analyzed data, wrote and revised manuscript. Co-Author: Matthew W. Fields 96 Contributions: Developed experimental design, analyzed data, wrote and revised the manuscript 97 Manuscript Information Page Lauren C. Franco, Siva Wu, Michael Woo, Manfred Auer, Matthew W. Fields Status of Manuscript: __x_ Prepared for submission to a peer-reviewed journal ____ Officially submitted to a peer-review journal ____ Accepted by a peer-reviewed journal ____ Published in a peer-reviewed journal 98 Abstract Biofilm samples of Desulfovibrio vulgaris Hildenborough grown under electron acceptor-limited (EAL), electron donor-limited (EDL), and electron donor to electron acceptor-balanced (BAL) conditions were analyzed via high resolution electron microscopy to investigate the composition and potential function of extracellular features in the biofilm matrix. Microscopy revealed the presence of membrane vesicles, extracellular filaments, and extracellular membranous structures. Membranous structures varied in shape, with some being sheet-like in structure, whereas others take on a more geometrical shape and form pockets in the biofilm that are devoid of cells. Transmission electron microscopy (TEM) images show that these structures connect to intact cells suggesting a biological origin and function. TEM imaging showed that membranous structures were associated with electron dense precipitates and energy dispersive x-ray spectroscopy (EDS) imaging analysis showed that these precipitates were composed of Fe, O and P. Wide-field montage TEM allowed observation of the biofilm grown under the three different conditions on a larger scale. Qualitatively, membranous structures produced in biofilm grown under the EAL condition were larger and more extensive than those produced under the EDL or BAL condition. Differential staining revealed that the structures were lipid-based. Fatty acid content of whole biofilm grown under BAL and EAL conditions was quantified and the results showed that biofilm grown under the EAL condition had ~3 times more fatty acid methyl ester (FAME) content. Since biofilm grown under the EAL condition had more extensive extracellular membranous structures, and these structures were associated with metal precipitates, biofilms grown under EAL and BAL were exposed to Cr(VI). Results suggested that biofilm grown under the EAL condition were more protected from Cr(VI) toxicity than those grown under the BAL condition, but the finding was not statistically significant. Together, the results presented here highlight lipids as an important and previously unrecognized part of the sulfate-reducing biofilm matrix and suggest that they might be involved in microbe-metal interactions. These results also continue to highlight the effects of nutrient limitation on microbial metabolism and physiology. 99 Introduction The biofilm matrix is an all-inclusive term for the extracellular substances that allow cells to stick to surfaces and each other. These substances include, but are not limited to polysaccharides, extracellular DNA, membrane vesicles, cell debris from lysed cells, membrane vesicles, enzymes, small molecules, and lipids (Flemming and Wingender 2010). While some of these components have been identified and the function established for a particular species, there are still many matrix components that have not been identified or described. This chapter highlights extracellular membranes as an important constituent of sulfate reducing biofilm matrices. Microorganisms are predicted to comprise the largest amount of biomass on earth (Whitman et al. 1998), are known to play vital roles in ecosystem function (Coleman and Whitman 2005; Duffy and Stachowicz 2006; Danovaro et al. 2008), and most likely represent immense, as yet undiscovered, biochemical capacities (Castelle et al. 2013). Additionally, microorganisms are known to play important roles in global biogeochemical processes such as carbon cycling and bioremediation (Anderson et al. 2003; Karl et al. 2012). It is well accepted that microorganisms can exist attached to surfaces and each other, often surrounded by extracellular polymeric substances (EPS) (Hall-Stoodley et al. 2004; Gross et al. 2007; Stewart and Franklin 2008). It is becoming increasingly clear that a mode of attached growth more closely resembles in situ 100 conditions for many microorganisms in different environments and might likely be a universal feature that presents an important physiology to explore in addition to the typically conducted studies on planktonic cells (Dunne 2002; Kolter 2005). Further, cells growing as a biofilm are known to have different physiologies and properties such as increased resistance to external stresses such as antimicrobials, heavy metal exposure, and desiccation (McKew et al. 2011; Clark et al. 2012; Stylo et al. 2015). The presence of extracellular membranes in a biofilm matrix has not yet been reported, however structures of a similar appearance have been described in other metal- reducing organisms. Shewanella oneidensis MR-1 nanowires were originally thought to be proteinaceous filaments composed of pilin (Gorby et al. 2006), but have most recently been identified as extensions of the outer membrane and periplasm that contain the cytochromes MtrC and OmcA, similar to the previously described outer membrane vesicle chains and membrane tubes in Myxococcus xanthus planktonic cultures and biofilms (Remis et al. 2014; Pirbadian et al. 2014). S. oneidensis MR-1 biofilms have also been reported to produce extracellular polymeric substances (EPS) that play an active role in uranium reduction. The EPS-glycocalyx type structure was found to be lined with the MtrC and OmcA cytochromes that were in close association with UO2 nanoparticles, indicating extracellular uranium (IV) reduction (Marshall et al. 2006). Differences between planktonic and biofilm growth modes have been well documented in DvH; however, differences between biofilms grown under different conditions have not been as well studied (Clark et al. 2012). In the previous chapter the 101 metabolic effects of nutrient limitation on D. vulgaris biofilm were investigated. In the current chapter, the effect of nutrient limitation on biofilm structure and composition is explored. To do this, we employed a variety of advanced high-resolution imaging and chemical characterization techniques, including 2D transmission electron microscopy (TEM), 3D serial block face scanning electron microscopy (SBF/SEM) imaging, and elemental analysis via energy-dispersive x-ray spectroscopy. This work describes extracellular membranous structures that are produced by D. vulgaris biofilms grown under nutrient limited and balanced conditions. The prevalence of these structures in the biofilm suggests that they are an important, but understudied component of sulfate reducing biofilm matrices. Materials and Methods Bacterial Strains and Growth Conditions D. vulgaris Hildenborough was grown in either batch or continuous mode. For batch growth, cells were inoculated into sterile N2 gassed balch tubes containing LS4D, a lactate and sulfate medium (modified to 2.5 µM resazurin and 130 µM riboflavin in Thauer’s vitamins) (Brandis and Thauer 1981; Borglin et al. 2009). The medium consisted of varying lactate and sulfate concentrations as follows: 60 mM sodium lactate and 30 mM sodium sulfate (balanced condition), 50 mM sodium lactate and 10 mM sodium sulfate (electron acceptor limited condition), and 18 mM sodium lactate and 50 mM sodium sulfate (electron donor limited condition). For continuous culture of D. 102 vulgaris biofilm, cells were grown in a modified CDC reactor equipped with additional gas lines to maintain anoxic conditions. Exponential phase cells were inoculated into a CDC reactor containing electron acceptor limited, balanced, or electron donor limited LS4D medium. Cells were grown in batch mode for 48 hours and then continuous mode with a dilution rate of 0.04-hr. Reactors were grown at room temperature (20-23°C), stirred at 60 rpm, and the headspace was continuously sparged with sterile N2 gas to maintain anaerobic conditions. Coupons of glass slides or aclar (7.8 mil thickness) (Electron Microscopy Sciences, Hatfield, PA) were submerged in the reactor body as a surface for biofilm growth. Sample Collection Biofilm samples were collected for biomass analysis and microscopic analysis. For biomass analyses glass slide coupons were removed from the reactor and biomass was scraped from both sides of the slide with a razor blade. Samples on aclar for microscopic analyses were collected at 168 hours. Immediately after sampling, aclar with attached biofilm was rinsed three times in phosphate buffered saline and then fixed in a 2.5% gluturaldehyde, 0.05M sodium cacodylate solution. Biomass Analyses Cell viability was determined via the most probable number method (Jarvis et al. 2010). Biofilm was scraped from glass slides, homogenized by vortexing and normalized by optical density to account for differing amounts of biomass between the different conditions. Normalized biofilm samples were then diluted to extinction and incubated at 103 30°C. After 3 weeks, growth was assessed and used to calculate the number of viable cells/ml for each biofilm condition. Lipid content from scraped biofilm (in triplicate) was chloroform extracted, transesterified, and measured by GC-MS according to the method described in Lohman et al. (Lohman et al. 2013). Imaging Methods Fluorescence Microscopy. Biofilms were fixed in 4% paraformaldehyde for 4 hours at room temperature, followed by incubation with the lipophilic fluorescent dye FM1-43 (www.thermofisher.com) and extensive washes with 20 mM HEPES buffer. Stained biofilm samples were either imaged on a Zeiss LSM 710 confocal microscope with Zen software or a Zeiss Axioskop Imaging platform with SPOT Basic software or embedded in O.C.T. freezing medium. Blocks of biofilm samples in O.C.T. were sectioned on the Leica Cryostat CM3050S (Leica Microsystems, Wetzlar, Germany) and ~10 µm-thick sections were imaged by epifluorescence microscopy. High pressure freezing and freeze substitution. Biofilms, unfixed or fixed, were placed in 1 mm wide by 200 µm deep aluminum freezing hats, then surrounded with 10% glycerol-solution that acts as a cryo-protectant prior to freezing. The biofilms were then cryo-immobilized using a BAL-TEC HPM-010 high-pressure freezer (BAL-TEC, Inc., Carlsbad, CA). Alternatively, biofilms were subjected to routine bench-top processing. As discussed in the main text, different biofilm samples were subjected to different freeze-substitution/staining protocols, resulting in samples that were unstained (did not see any heavy metal staining during sample preparation), 0.1% uranyl acetate stained, or 104 stained by 1% osmium tetroxide plus 0.1% uranyl acetate. For all samples that were freeze-substituted we used either a Leica AFS2 (Leica Microsystems, Wetzlar, Germany) following a previously described protocol (McDonald et al. 2007) or adopted a method of super quick freeze substitution (McDonald and Webb 2011). Upon completion of freeze- substitution, the biofilm samples were rinsed five times in pure acetone and then progressively infiltrated with an epon-araldite resin (McDonald and Müller-Reichert 2002). Samples were flat embedded between two slides using two layers of parafilm as a spacer before being polymerized overnight in an oven at 60°C (McDonald and Müller- Reichert 2002; Müller-Reichert et al. 2003). Transmission electron microscopy. 70-100 nm sections were collected on formvar-coated grids using a Reichert Ultracut E ultramicrotome (Leica Microsystems, Germany). For serial section TEM, 100 nm sections were collected in ribbons of ~5 sections upon each grid to preserve the order and orientation of the sectioned material. Sections were post-stained using 2% uranyl acetate in 70% methanol followed by Reynold’s lead citrate. The sections were imaged in an FEI Tecnai 12 TEM (FEI, Hillsboro, OR) operated at 120 kV. Images were recorded using a Gatan CCD with Digital Micrograph software (Gatan Inc., Pleasanton, CA). ImageJ software (Abramoff et al. 2004) and Adobe Photoshop CS4 (Adobe Systems Inc., San Jose, CA) were used for further image processing and for the registration of the serial section images for 3D analysis. Volume Electron Microscopy Imaging. For SBF-SEM, biofilms, embedded in 105 resin, were mounted onto an aluminum pin with a cyanoacrylate adhesive. The pin, which takes the place of a normal SEM stub, was loaded into a sample holder for the Gatan 3View (Gatan Inc., Pleasanton, Ca). Serial block face scanning electron microscopy was carried out as previously described (Denk and Horstmann 2004). SBF-SEM data was collected using a 3View system mounted on a Zeiss Sigma FE-SEM; serial images were acquired at 5 kV at 4k x 4k at 10 nm XY pixel size, and 50 nm Z slice dimensions. Energy Dispersive X-ray Spectroscopy. High resolution scanning transmission electron microscopy (STEM) and energy-dispersive spectroscopy (EDS) were carried out on a Zeiss Sigma FE-SEM in backscatter or Scanning Transmission Electron Microscopy mode. Thin sections were mounted on continuous carbon-coated Formvar TEM grids. High-angle annual dark field (HAADF) scanning transmission electron microscopy (STEM) images and X-ray elemental line scans were acquired with a 1-nm probe at 120 or 200 kV. The specimens were tilted 10 degrees toward the X-ray detector to optimize the X-ray detection geometry. Collection times were 300 live seconds for each line scan. The Effect of Cr(VI) on Whole and Scraped Biofilm D. vulgaris biofilm grown under EAL or BAL conditions was harvested at 168 hours. For testing the effect of Cr(VI) on scraped and homogenized biofilm, biofilm CDC reactors were quickly moved into an anaerobic glove bag with gas, media input, and waste output lines clamped off. In the glove bag, biofilm slides were removed from the reactor, gently dipped in PBS three times, and then scraped into a microcentrifuge tube with 1 ml LS4D medium containing lactate and sulfate concentrations consistent with the 106 condition they were grown in. Scraped biomass was drawn up through a 16-gauge needle and transferred to a sterile, anaerobic balch tube containing 8 mls LS4D medium. Balch tubes containing the scraped biofilm grown under EAL or BAL conditions was homogenized via vortexing three times at 30 second intervals and then centrifuged at 5,750×g for 10 min at room temperature. To “wash” cells of sulfide that would abiotically reduce Cr(VI), the supernatant was removed anoxically and aseptically by needle attached to a vacuum flask under constant flow of N2 gas to maintain neutral pressure. Pellets were then re-suspended in 9 ml fresh LS4D medium and washed once more, after which the pellets were re-suspended and concentrated in 1 ml fresh LS4D. Concentrated cell cultures were inoculated into 25 ml balch tubes with LS4D medium (10 ml) that contained Cr(VI) (potassium chromate) at 0 or 50 µM to an OD600 between 0.06 and 0.07. Growth was monitored by optical density (600 nm). All experiments were performed in triplicate. To test how Cr(VI) affected viability of intact biofilm grown under EAL and BAL conditions, biofilm samples from the same CDC reactors as above in an anaerobic glove bag, were removed, gently dipped in anaerobic PBS and then transferred into tubes containing either EAL or BAL LS4D medium and 0 or 500 µM Cr(VI). The samples were incubated for 3 hours in the glove bag after which the slides were removed and biofilm was scraped into 1ml fresh LS4D medium. Scraped biofilm was then serially diluted in 96-well plates for MPN analysis as described above. Cytochrome Staining and Heme Quantification 107 For cytochrome staining, periplasmic and spheroplast cellular fractions from scraped biofilm grown under BAL and EAL conditions were separated according to Baumgarten et al. and Badziong and Thauer (Badzjong and Thauer 1980; Baumgarten et al. 2001). Periplasmic fractions were incubated in 3 molar urea and 90 mM sodium dodecyl sulfate at room temperature for 1 hour before the sample was separated on a polyacrylamide gel (4-20%) via electrophoresis. Cytochromes were visualized via a peroxidase and 3,3’,5,5’-tetramethylbenzadine stain (Thomas et al. 1976). Stained bands corresponding with the size of cytochrome c (~14 kDa) were qualitatively compared. Heme content was measured from scraped and homogenized biofilm grown under EAL of BAL conditions as a quantitative way to detect any differences between the two conditions. Heme was measured using the QuantiChrom Heme Assay Kit (BioAssay Systems, Hayward, CA), a colorimetric assay for heme. Results 2D TEM Imaging of Desulfovibrio vulgaris Biofilms Using high pressure cryogenic preservation, followed by freeze-substitution and resin-embedding to preserve native biofilm structure, large portions of D. vulgaris biofilm grown under EAL, EDL, and BAL conditions were examined via wide-field TEM. Given the very large size typically encountered in a bacterial biofilm, we examined approximately 1 µm semi-thin sections of resin-embedded biofilms that were toluene blue-stained and subjected to montage optical microscopy imaging (Figure 1). 108 Selected regions with metal deposits were re-imaged by wide-field montage TEM using 70-100 nm ultrathin sections, and increasingly higher resolution images reveal typical metal deposits under EDL, BAL and EAL conditions. Metal deposits were observed under all three conditions, but they differed significantly in the amount of metals that were deposited onto these membrane structures and the overall dimensions of the metal deposits. It should be noted metal deposits were a highly regional phenomenon under all three conditions with most of the biofilm not showing any significant metal deposits. Imaging of DvH biofilms grown under EAL, EDL, and BAL conditions revealed the presence of heterogeneously distributed extracellular structures that were connected to some cells and were closely associated with metal precipitates. Figure 1. Large-scale microscopic analysis of sections of biofilms grown under EAL (A), BAL (B), and EDL (C) conditions highlighting differences in the extracellular membrane appearance. Each image shows progressively higher magnification within a panel. 0.5 µm 0.5 µm 10 µm 1 µm 0.5 µm 1 µm 10 µm 10 µm 1 µm 109 It was initially, qualitatively, observed that biofilm grown under the EAL condition contained more, and larger, extracellular structures that those grown under the EDL or BAL conditions (Figure 1). Extracellular metal deposits, appearing as electron dense dark spots, were distributed in highly heterogeneous patterns throughout the biofilm (Figure 2). In areas of close proximity, metal deposition was varied, with metal deposits located in certain places between bacteria, whereas neighboring extracellular spaces were devoid of any metal deposits and appeared empty (Figure 2a). Metal deposits varied in proximity to cells, with some associated with cell surfaces (Figure 2a, b) and others being deposited on thin branched string-like structures (Figure 2c, d). On occasion, structures forming amorphous pockets devoid of cells were observed within the biofilm (Figure 2b). Close-up images of the connection between cells and the structures associated with metal deposits reveal that these structures appear to be connected to and continuous with the cellular outer membrane (Figure 2c, d). The fact that these objects of string-like appearance can be followed through large portions of approximately 70-100 nm ultrathin section of the biofilm suggests that these objects are not strings of metal deposits, but are sheet-like in structure, extending in three dimensions throughout the biofilm. In favorably oriented biofilm sections we observed extracellular metal deposits with underlying structure directly connected at multiple sites to the bacterial outer membrane (Figures 2c, d). Note that the samples are not counterstained, thus the bacteria appear as light and rather featureless, although the outer and inner membrane can still be 110 detected (Figure 2c). The metal deposits appear dark and in many instances composed of nano-scale crystals decorating a continuous underlying biological structure. Interestingly, these extracellular features form rather complex large structures with many branches originating what appears in 2D to be ring-like core objects whose interior is devoid of any bacteria or other material (Figure 2b). Given the role that Desulfovibrio spp. play in catalyzing processes involving metal transformations (i.e., biocorrosion, heavy metal reduction) and the observed physical connection between the bacterial outer membrane and extracellular structures decorated with metal deposits leads to the hypothesis that these structures are involved in microbe-metal interactions. 111 Figure 2. TEM images of D. vulgaris biofilm thin sections showing extracellular membrane structures. EMs vary in shape and size (A, B), connect with cells, and are associated with metal deposits (C, D). Large Volume 3D SBF/SEM Imaging Based on the 2D TEM imaging, it appeared that the extracellular structures extended three-dimensionally throughout the biofilm. To visualize the structures in 3D, we employed serial block face (SBF)-SEM. As observed in Figure 3a, which represents one of over 3,000 slices of the reconstructed 3D volume, metal deposits adopt lamellar, 112 spherical and/or barrel-like shapes (Figure 3a-c). The barrel-like shape of the metal deposit becomes clearer upon 3D rendering of approximately 3 µm of the respective volume (Figure 3d-f). For the sake of clarity only an approximately 1 µm thin central slab of the bacteria is rendered, illustrating their close proximity to the metal deposit (Figure 3f). Figure 3. Thin sections of biofilm show extracellular membrane structures within the biofilm (B and C are magnifications of features in A). 3D renderings of extracellular membrane structures within the biofilm (D, E, F). A 3D rendering of a 40µm x 40µm x 100µm sample of biofilm showed densely packed bacterial cells, extracellular structures, and metal deposits buried within the biofilm (Figure 4a). When the cells and extracellular structures are removed, leaving only the metal deposits, the heterogeneous distribution of the metal deposits throughout 113 the biofilm is apparent (Figure 4b). The extracellular structures shown in Figure 4b, extend through the entire 3D volume, thus extending for approximately 15 µm in the XY direction and at least 100 µm in the Z direction. Additionally, vesicular objects were observed nearby extracellular structures, with some vesicles apparently being in intimate contact or being in the process of and/or having fused with these objects. These presumptive vesicles are further investigated in Chapter 5. Figure 4. 3D reconstruction of 40 x 40 x 100 µm section of biofilm based on SBF imaging (A) and the same section with cells removed and only metal precipitates remaining (B). Elemental Analysis of Extracellular Metal Deposits Having determined that extracellular metal deposits are localized to thin lamellar sheets or stacks of sheets, which can connect to bacterial outer membranes and that can extend for at least 100 micrometers throughout a biofilm, the elemental composition of 10 µ m 114 the metal deposits were determined using line-profile dispersive X-ray spectroscopy scanning transmission electron microscopy (EDS STEM) imaging of non-osmicated, resin-embedded samples (Figure 5). The EDS STEM imaging line profiles (Figure 5c, e) showed very little background on the resin, but showed clear peaks that corresponded well with metal deposits (Figure 5d, f). Surprisingly, iron, oxygen and phosphorus, but no sulfur peak (Figures 5d, f) nor other commonly found elements such as nitrogen, carbon or chlorine was detected. This suggests, contrary to prior expectations, that metal deposits predominantly consist of iron oxides and possibly iron phosphates, but not iron sulfide, despite the presence of dissolved and precipitated sulfide detected within the biofilm during the experiment (Chapter 3) and abundance of H2S gas throughout the experiment. 115 Figure 5. EDS elemental of metal precipitates associated with two different instances extracellular membranes within D. vulgaris biofilm (A, B). Path of analysis follows the lines shown in C, E and elemental composition is shown along the path in D, F. Underlying Metal Deposition Sites are Biological Membranes The images showing the connection between bacterial cellular membranes and the extracellular structures lead to the hypothesis that the structures may be composed of biological lipid membranes. In order to test this hypothesis, biofilm samples were differentially stained prior to resin-embedding. Unstained or uranyl acetate-only-stained samples reveal a ~10 nm gap between adjacent metal deposits (Figure 6a), whereas 0.5 µm 0.5 µm 0 100 200 300 400 500 600 700 800 nm 0 100 200 300 400 500 600 700 800 nm 0 100 200 0 100 200 0 100 200 nm nm nm In te ns ity (C ou nt s) In te ns ity (C ou nt s) In te ns ity (C ou nt s) In te ns ity (C ou nt s) In te ns ity (C ou nt s) 116 osmicated samples did not display such a gap but instead a continuous filamentous structure onto which metal nanocrystals are deposited (Figure 6b). Additionally, ~5µm serial thick cryo-stat sections of unstained, frozen biofilm samples that were stained with the lipophilic dye, FM1-43, exhibited patterns of fluorescent structures (Figure 6d) that appeared to constitute a large portion of the biofilm matrix, consistent with the hypothesis that these structures are of biological lipid membrane origin. Ruthenium Red staining of biofilm samples (Figure 6e) showed very little to no carbohydrate associated with the extracellular structures under investigation compared to the control stained with osmium tetroxide and uranyl acetate only (Figure 6c). 117 Figure 6. Differential staining of extracellular membranes. Uranyl acetate only stained structure (A), followed by osmium tetroxide and uranyl acetate stained structure (B). Ruthenium red stained thin section (E), compared to control (osmium tetroxide and uranyl acetate) (C). Staining with lipophilic fluorescent dye FM1-43 (D). Because the differential staining evidence supports the hypothesis that the structures are lipid based, and because we qualitatively observed that there were more and larger structures produced under the EAL condition, total fatty acid methyl ester (FAME) content of biofilm grown under EAL and balanced conditions was quantified to 118 determine if there was a difference in fatty acid content. FAME content in DvH biofilms grown under the EAL condition was three times higher as compared to biofilm grown under balanced conditions when normalized to dry weight (18.9±0.36% vs. 6.38±0.59% dry weight) (Figure 7). The increase in fatty acid production under electron acceptor limitation is also biofilm specific, with planktonic cultures grown under balanced and electron acceptor limitation both yielding approximately 5-6% FAME per dry weight. Due to the concern that the higher FAME content and presence of larger extracellular membranous structures for biofilm grown under the EAL condition could be due to a greater number of aggregated membrane material from dead cells under this condition compared to the BAL condition, cell viability between the three conditions was also tested. There was no statistically significant difference in viability between the EAL, BAL, and EDL conditions (Figure 8), indicating that the higher FAME content and larger, more extensive, membranous structures are not merely a consequence of decreased cell viability under the EAL condition. 119 Figure 7. Weight percent of FAME content in dry weight of D. vulgaris biofilm grown under EAL and BAL conditions. Figure 8. Viability (cells/ml) of scraped and homogenized biofilm grown under EAL, BAL, and EDL conditions normalized by optical density. 120 The Effect of Nutrient Limitation on Cr(VI) Toxicity in D. vulgaris Biofilm Because D. vulgaris grown planktonically under EAL conditions was more susceptible to Cr(VI) toxicity compared to BAL conditions, as was shown in Chapter 2, the effect of Cr(VI) on D. vulgaris biofilm was investigated here. Measuring the biotic effect of heavy metal toxicity on sulfate-reducing biofilms is inherently difficult because the concentration of sulfide within the biofilm is so high that at field relevant levels of Cr(VI), most if not all of the Cr(VI) reduction will be abiotic. In a natural system, growth and therefore sulfide production may be slower, and other organisms may be utilizing the sulfide produced by the SRB, so understanding biological Cr(VI) reduction and toxicity is still warranted. The effect of Cr(VI) on D. vulgaris biofilm was investigated in two ways: the first with scraped, homogenized biofilm, and the second with intact biofilm. Scraping and homogenizing the biofilm was advantageous because more of the sulfide could be removed via washing of the cell pellet. Removing sulfide from the intact biofilm was not possible, and therefore the Cr(VI) concentration was 10-fold higher than that for the scraped and homogenized cells. The lag time for scraped, homogenized D. vulgaris biofilm cells was ~75 hours for cells grown under both EAL or BAL conditions (Figure 9). This differed from the results presented in Chapter 2 in that there was no difference in lag time between the two conditions and the lag time was much shorter (approximately half) than the data presented in Chapter 2. The intact biofilm had a log reduction of 0.9±0.19 for cells 121 grown under the BAL condition and 0.5±0.37 for cells grown under the EAL condition, suggesting that cells growing under the BAL condition were slightly more susceptible to Cr(VI) toxicity than cells grown under the EAL condition (mixed effects ANOVA, p- value=0.06). It should also be noted that the biofilm grown under EAL and BAL conditions was exposed to the same concentration (500 µM) of Cr(VI) even though the biofilm grown under BAL contained approximately twice as many cells compared to biofilm grown under the EAL condition, so EAL biofilm had a higher concentration of Cr(VI) per cell, but a smaller log reduction compared to the BAL biofilm. Biofilm grown under the BAL condition also has a higher concentration of sulfide (Chapter 3), meaning that there was a higher potential for abiotic Cr(VI) reduction under the BAL condition. The results suggest that nutrient limitation/resource ratio affect Cr(VI) toxicity in the biofilm growth mode and this finding should be further investigated, especially given that this result was opposite of the results presented in Chapter 2 in which planktonic cells grown under the EAL condition were more susceptible to Cr(VI) compared to those grown under the BAL condition. 122 Figure 9. Planktonic growth of scraped and homogenized D. vulgaris biofilm grown under EAL (squares) and BAL (circles) conditions with 0 (filled) or 50 (empty) µM Cr(VI). To determine if there was a difference in cytochrome quantity for biofilm grown under EAL compared to BAL that could help explain the difference in Cr(VI) toxicity between biofilm grown under EAL and BAL, cytochromes from purified periplasmic fractions was stained on a protein gel and visually compared and heme content was measured. There was a slight difference in band intensity of the two bands corresponding to cytochrome c at ~14 kDa (Figure 10), with the band associated with the BAL conditions being slightly darker. To determine if there was a quantifiable difference 123 between the two conditions, heme concentration was measured and compared in biofilm grown under EAL and BAL conditions. There was no difference in heme concentration between the two conditions (0.013 ± 0.004 µg heme/µg protein for BAL and 0.015 ± 0.005 µg heme/µg protein for EAL). Figure 10. Protein gel showing ladder in kilodaltons (kDa) on right and stained cytochrome c at ~14-15 kDa from biofilm grown under BAL and EAL conditions. Discussion Hardiness of biofilm cells can be attributed to the distinct physiological state of cells existing and growing in the biofilm growth mode and also to the secreted matrix components that interact with surfaces and the external environment. The biofilm EPS is an all-inclusive term for the extracellular macromolecules that enable cells to adhere to surfaces and each other, and the understanding of the biofilm matrix has progressed to include, but are not limited to polysaccharides, extracellular DNA, membrane vesicles, 124 cell debris from lysed cells, enzymes, and structural proteins (reviewed in Flemming and Wingender, 2010). While some of these components have been identified and a function established for a particular species, there are still many matrix components that have not been identified or described, including membrane vesicles and related structures, particularly for environmental microorganisms. Membrane vesicles (MVs) in Bacteria and Archaea studied to date can play roles in nutrient acquisition, biofilm development, and pathogenesis dependent upon the organism (Schooling and Beveridge, 2006; Kulp and Kuehn, 2010; Manning and Kuehn, 2013). Moreover, the biofilm matrix is increasingly recognized to contain a variety of intra- and inter-matrix interactions that contribute and may control biofilm behavior (Schooling and Beveridge, 2006; Payne and Boles, 2016), and Manning and Kuehn (2013) likened the activity of MVs to that of intracellular membrane-bound processes. More recently, Hooper and Burstein posited that the minimization of extracellular space in prokaryotic (i.e., Bacteria and Archaea) biofilms promoted cellular associations that impacted metabolism and may have contributed to the evolution of Eukarya (Hooper and Burstein 2014). Identification of extracellular matrix components in biofilms is integral to our understanding of how biofilms form, persist, and evolve. The results of this study show that membranous structures contribute a previously unrecognized portion of the extracellular matrix of D. vulgaris biofilms. These structures connect to and are continuous with cellular membranes and the use of lipophilic dyes, metabolomic analyses, and fatty acid quantification support the hypothesis that these structures are 125 lipid-like in composition. The connection of these structures to the outer membrane of structurally intact, and hence presumably viable bacteria is highly significant as it suggests a metabolic connection to membrane structures and possibly neighboring bacteria. Metal precipitates co-localize with the membrane structures, and these observations suggest the structures may be involved in microbe-metal interactions. One possible function that these structures may fulfill is a mechanism to sequester metal precipitates away from the cell. A mechanism to remove or reduce metals away from the cell body would be beneficial given that planktonic D. vulgaris cells have been shown to accumulate reduced, precipitated Cr(III) in the periplasmic space along membranes, potentially encasing themselves (Goulhen et al. 2006). Given that the reduction of certain metals, such as Cr(VI), is not linked to growth for D. vulgaris, but is rather an energy-costing detoxification strategy (Chardin et al. 2002), the ability to reduce metals away from the cell body may be advantageous. Isolated Shewanella EPS has been shown to reduce U(VI), suggesting that this may be a strategy of metal-reducing bacteria (Cao et al. 2011). Based on the idea that the extracellular structures could play a role in metal reduction, the effect on Cr(VI) toxicity on biofilm grown under EAL and BAL conditions was investigated. Given that there were more extensive extracellular membranes, three times more measured FAME content, and upregulated fatty acid metabolites observed in biofilms grown under the EAL condition, we hypothesized that biofilm grown under the EAL condition would be less susceptible to Cr(VI) toxicity compared to the BAL 126 condition. Scraped and homogenized biofilm cells grown under EAL and BAL conditions had similar lag-times (~75 hours) when exposed to 50 µM Cr(VI), which contradicted the results from Chapter 2 in which planktonic cells grown under the EAL condition were more susceptible to Cr(VI) toxicity. An overall shorter lag-time for biofilm cells is in agreement with the many reports of biofilm cells being more resistant to heavy metal stresses compared to planktonic cells. It is interesting to note that even when the structure of the biofilm is removed, the cells were still somewhat protected from Cr(VI) toxicity compared to planktonic cells. Previous results have shown that biofilm growth is a unique physiological state compared to planktonic (Clark et al., 2012), and the current results show that biofilms grown under different energy conditions (BAL vs. EAL) have different physiological responses to Cr(VI) even when intact biofilm structure is removed. When intact D. vulgaris biofilm grown under EAL and BAL conditions were exposed to high concentrations of Cr(VI) to overcome abiotic Cr(VI) reduction, the cells grown under the BAL condition had a greater log-reduction of viable cells than those grown under the EAL condition. While the difference in log-reduction between the two conditions was not statistically significant at a p-value of 0.05, the finding is suggestive that in the biofilm growth mode, the EAL condition might confer increased resistance to Cr(VI) toxicity and warrants further investigation. It should be noted that BAL biofilms typically contain more sulfide than EAL biofilms and thus, EAL biofilms were likely more tolerant to Cr(VI) exposure. 127 Elemental analysis of extracellular membrane associated metal precipitates surprisingly did not show the presence of sulfur compounds, but rather iron oxides and iron phosphates. Interestingly, iron and phosphorus were also prevalent in the extracellular glycocalyx-like structures that Shewanella oneidensis MR-1 produces and the iron and phosphorus were closely associated with reduced UO2 particles. The authors suggest that the iron was present due to heme containing proteins that were involved in the uranium reduction (Marshall et al. 2006). Additionally, intracellular membrane encapsulated iron-phosphorus granules have been reported in Desulfovibrio magneticus RS-1. These iron-phosphorus granules are distinct from the magnetite crystals but possible functions are unknown (Byrne et al. 2010). While quite different in lineage and growth conditions, these three findings together support a possible biological role of iron- phosphates in dissimilatory metal reducing bacteria that call for additional research. Other components of D. vulgaris biofilm matrix have been analyzed in previous work as well as this one. In agreement with Clark et al. (Clark et al. 2007a; Clark et al. 2012), we observed that extracellular carbohydrate was not a significant part of the biofilm matrix under any of the tested nutrient ratios. Although carbohydrate is detected in whole biofilm samples (cells and extracellular materials), little carbohydrate is detected extracellularly by carbohydrate-sensitive staining. Extracellular proteins have been shown to be integral to biofilm formation and maintenance, and a proteomic analysis of D. vulgaris biofilm grown under different conditions from this study (60mM lactate, 50mM sulfate, 30°C) detected 188 extracellular proteins (Clark et al. 2007b; 128 Clark et al. 2012). Extracellular DNA has been shown to be important for biofilm formation and structure in other organisms, but DNase treatment of D. vulgaris biofilms did not affect structural integrity (data not shown). Membrane vesicles have also been observed in close association with cells and extracellular membranes in D. vulgaris biofilms and are the subject of Chapter 5. In addition to the presence of extracellular membranes in the matrix, this study also highlighted how SRB biofilms respond to nutrient limitation. The lipid analysis revealed that the EAL biofilm produces approximately three times more fatty acids than the BAL biofilm. Also, metabolomic analysis from Chapter 3 showed an upregulation of fatty acid metabolites under the EAL condition. This indicates that sulfate limitation causes cells to shift to a lipid storing/excreting metabolism. Further, metabolic analysis shows that limiting sulfate causes dysregulation of 3303 metabolites (fold change ≥ 1.5, p-value ≤ 0.01), suggesting that many metabolic pathways are altered to cope with the limitation of sulfate. Previous studies have shown Desulfovibrio spp. response to electron acceptor limitation (Okabe et al. 1992; Villanueva et al. 2008), but none have investigated effects of electron acceptor limitation in a biofilm growth mode. One of the key distinctions of this study is that it is by far the largest 3D volume of a bacterial biofilm ever studied at high resolution. This was possible by using a new 3D EM technology known as Serial Block Face Scanning Electron Microscopy, where 40 microns by 40 microns in XY have been recorded at ~10 nm stepsize, and ~100 microns 129 has been recorded at ~30 nm stepsize, using iterative SEM imaging of the block surface and subsequent removal of the block surface with a diamond knife. Likewise, correlative optical microscopy and 2D TEM was utilized to image toluene blue stained ~1 micron or unstained ~70 nm ultrathin sections, which allowed the study of biofilm on a larger scale. Identifying the frequency and locations of significant extracellular metal deposits by optical microscopy enabled qualitative observations to be made and then studied in much more detail by TEM analysis and other quantitative methods. The multiscale imaging analysis was key to this study, as metal deposits are highly heterogeneous; therefore, it was key to first gain an overview image of the entire biofilm, before examining certain areas at higher resolution. This result is also significant for bulk omic/chemical/analytical analysis, which is widely employed for many different environments and systems, but may be insufficient to detect highly significant local differences that only manifest themselves in a small fraction of sampled volume. 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Proc Natl Acad Sci U S A 95:6578–6583. 134 CHAPTER FIVE OUTER MEMBRANE VESICLES AND ASSOCIATED PROTEINS PRODUCED BY DESULFOVIBRIO VULGARIS HILDENBOROUGH BIOFILMS Contribution of Authors and Co-Authors Author: Lauren Franco Contributions: Developed experimental design, performed experiments, analyzed results, wrote and revised manuscript. Co-Author: Chris Petzold Contributions: Proteomic data acquisition, data analysis Co-Author: Matthew W. Fields Contributions: Developed experimental design, analyzed results, wrote and revised manuscript. 135 Manuscript Information Page Lauren C. Franco, Chris Petzold, Mathew W. Fields Applied Microbiology and Microbiology Status of Manuscript: __x_ Prepared for submission to a peer-reviewed journal ____ Officially submitted to a peer-review journal ____ Accepted by a peer-reviewed journal ____ Published in a peer-reviewed journal 136 Abstract Outer membrane vesicles (OMVs) have been observed in Desulfovibrio vulgaris Hildenborough cultures, but their function is unknown. OMVs produced by other bacteria and archaea have a wide variety of functions and have been well studied in pathogenic bacteria, but are understudied in non-pathogenic organisms. Further, OMVs produced by biofilms have been shown to be distinct in protein content compared to their planktonic counterparts. In this study, OMVs were isolated from D. vulgaris biofilms grown under electron acceptor-limitation (EAL) and electron donor/electron acceptor balanced (BAL) conditions. Isolated OMVs were between 20-60 nm in diameter and OMV fractions also contained tube structures that appeared to be precursors to individual OMVs. Proteins in the isolated OMV fractions were identified by untargeted shotgun proteomics and revealed expected outer membrane and periplasmic components such as lipoproteins and porins. Proteins involved in oxidative stress and two hydrogenases were also detected in OMVs, suggesting a potential function for OMVs in D. vulgaris. Differences in abundance of the proteins detected between OMVs isolated from biofilm grown under EAL vs. BAL conditions were not detectable, but more biological replicates in the future may resolve this issue. Future studies might also incorporate metabolomic analysis to identify and unique metabolites that are contained within OMVs. 137 Introduction Sulfate reducing bacteria are ubiquitous in the environment and play pivotal roles in natural and industrial processes such as biogeochemical cycling, biocorrosion, and bioremediation. Many of these processes involve the formation of biofilms by these microorganisms (Truper 1984; Neria-González et al. 2006; Liu et al. 2007; Marsili et al. 2007). Desulfovibrio vulgaris Hildenborough is a well-characterized model organism for sulfate reducing bacteria that has been studied in biofilm growth mode. It has been shown that D. vulgaris exhibits a distinct gene expression pattern when growing as a biofilm as opposed to the different phases of planktonic growth and some of these differences are due to the cellular production of biofilm matrix components (Clark et al. 2007; Clark et al. 2012). The biofilm matrix can consist of many different components such as carbohydrates, proteins, lipids, pili and flagella, DNA, and outer membrane vesicles (OMVs) (Flemming and Wingender 2010). A variety of functions have been deduced from proteomic analyses of purified OMVs of many different organisms. These include, but are not limited to, DNA exchange, cell-cell communication, antibiotic resistance, heavy metal resistance, phage resistance, toxin release, and biofilm formation (Ciofu et al. 2000; Klieve et al. 2005; Mashburn and Whiteley 2005; Gorby et al. 2008; Yonezawa et al. 2009; Manning and Kuehn 2011; Guidi et al. 2013). Many of these studies have focused on pathogenic organisms and the role that OMVs play in pathogenesis and fewer 138 studies have examined the role that OMVs play in non-pathogenic bacteria. Further, there are very limited studies on how environmental changes or stress affects OMV production and composition, and the need for such work was emphasized in a recent review article (Orench-Rivera and Kuehn 2016). While many studies have isolated and analyzed OMVs from planktonic cultures, relatively few studies have examined the function that OMVs have in biofilms. OMVs have been observed in naturally occurring biofilms in environments such as in a domestic sink drain, water treatment plants, and in a riverbed (Schooling and Beveridge 2006). Additionally, studies on cultured biofilms have shown that OMVs in biofilms are distinct compared to their planktonic counterparts. Schooling and Beveridge (2006) were the first to report on the differences between Pseudomonas aeruginosa OMVs from a biofilm versus planktonic cells: biofilms produced approximately 16 times more MVs according to dry weight, biofilm OMVs had smaller average diameters than planktonic OMVs, and SDS-PAGE analysis of biofilm and planktonic OMVs revealed different banding profiles, indicating potential differences in vesicle protein content. Further, protein composition in OMVs from P. aeruginosa biofilms differs from planktonic counterparts, pointing to differences in OMV function in biofilm versus planktonic produced OMVs (Park et al. 2015). Helicobacter pylori OMVs from biofilms have also been observed to be distinct from planktonic OMVs with biofilm produced OMVs containing double stranded DNA that is thought to contribute to biofilm structure and aggregation (Grande et al. 2015). 139 OMVs in Desulfovibrio species have been documented in the literature, but have yet to be purified and analyzed (Nanninga and Gottschal 1987). Given the role that sulfate reducing biofilms play in the natural environment and industrial processes, it is important to understand possible OMV functions in biofilms. Based upon observations of highly localized, extracellular membrane structures in D. vulgaris biofilms, we developed a method to determine if vesicles could be enriched from biofilm samples and potential protein content. Materials and Methods Bacterial Strains and Growth Conditions Desulfovibrio vulgaris Hildenborough was obtained from Dr. Romy Chakraborty (Lawrence Berkeley National Lab). D. vulgaris was grown in LS4D medium, which contains lactate as the carbon source and electron donor and sulfate as the electron acceptor (Clark et al. 2006). Lactate and sulfate concentrations were altered to create balanced and electron acceptor limited conditions. The balanced condition was defined as 60 mM sodium lactate and 30 mM sodium sulfate and the electron acceptor limited condition was defined as 50 mM sodium lactate and 10 mM sodium sulfate. D. vulgaris was grown as a biofilm under continuous flow conditions in a modified CDC reactor. Exponential phase cells were inoculated into a reactor containing balanced or electron acceptor limited LS4D medium and were grown in batch mode for 48 hours. Reactors were grown at room temperature (20-23°C), with a dilution rate of 0.04- 140 hr, stirred at 60 rpm, and the headspace was continuously sparged with sterile N2 gas to maintain anaerobic conditions. Coupons of glass slides were submerged in the reactor body as a surface for biofilm growth. Isolation of outer membrane vesicles Glass slides were removed from the reactor at 168 hours and biomass from each slide was aseptically scraped into a tube containing 1 ml 100mM HEPES buffer, pH 7.4. Scraped biomass was homogenized by vortexing for 5 minutes and then centrifuged at 12,000 x g for 20 minutes at 4°C. Supernatant was collected and kept on ice while the biomass pellet was resuspended in another 1 ml of HEPES buffer and the vortexing, centrifugation, and supernatant collection were repeated 2X. Supernatants were pooled, filtered through a 0.45 µm filter, then a 0.2 µm filter. Filtered supernatant was centrifuged at 125,000 x g for 1.5 hours to pellet OMVs and any other extracellular biofilm components. Pellets were resuspended in 45% optiprep (Sigma) in 100 mM HEPES buffer, pH 7.4. The samples were aliquoted into 5 ml ultracentrifuge tubes and optiprep in decreasing concentrations was layered on top (0.6 ml 45% with sample, 0.6 ml 40%, 0.8 ml 35%, 0.8 ml 30%, 0.8 ml 25%, 0.8 ml 20%, 0.4 ml 15%). Density gradients were centrifuged at 100,000 x g for 16 h at 4°C in a 30° fixed angle rotor (Thermo Scientific S110-AT rotor). Fractions were collected in 0.2 ml aliquots and screened for the presence of OMVs by transmission electron microscopy (TEM). Fractions containing isolated OMVs were concentrated via ultracentrifugation at 141 125,000 x g for 1.5 h, resuspended in 50 mM ammonium bicarbonate, flash frozen in liquid nitrogen, and stored at -80°C. Transmission Electron Microscopy Aliquots of the density gradient fractions (20 µl) was spotted on a 40nm silicon oxide membrane TEM grid (Ted Pella, Inc.), allowed to sit for 3 minutes, excess liquid wicked away, and then washed with sterile deionized water. After washing, samples were stained with 10 µl 2% w/v uranyl acetate for 1 min and then wicked dry. Samples were imaged on a Zeiss Leo 912AB transmission electron microscope. Nucleic Acid Detection The Qbit high sensitivity DNA assay kit (Life Technologies) was used according to the manufacturer's directions to measure DNA associated with OMVs. OMV samples were boiled at 95°C for 5 minutes prior to measuring to access any DNA held within the OMV. Proteomic Analysis An untargeted shotgun proteomics approach was taken for identifying peptides in isolated OMV samples. Proteins were extracted and precipitated from samples via a chloroform/methanol extraction protocol, digested with trypsin, and desalted according to established protocols (Parsons et al. 2013; González Fernández-Niño et al. 2015). Purified samples were analyzed on on an Agilent 6550 iFunnel Q-TOF mass 142 spectrometer (Agilent Technologies) coupled to an Agilent 1290 UHPLC system with the parameters described in (González Fernández-Niño et al. 2015) et al. (González Fernández-Niño et al. 2015). The 20 most intense ions within the 300-1,400 m/z mass range and above a threshold of 1,500 counts were selected for MS/MS analysis. MS/MS data was analyzed using Mascot (Matrix Biosciences) with the parameters listed in González Fernández-Niño et al. (2015) and matched with the annotated genome sequence of D. vulgaris Hildenborough (GenBank accession number AE017285 (chromosome) and AE017286 (plasmid)) (Heidelberg et al. 2004). Identified peptides were further analyzed with Scaffold (Proteome Software Inc.) and peptide identifications were accepted if they reached a >95% probability according to the Peptide Prophet algorithm (Keller et al. 2002). Protein identifications were accepted if the reached a >95% confidence according to the Protein Prophet algorithm and contained at least 1 identified peptide at >95% confidence (Nesvizhskii et al. 2003). To quantify protein abundance, total spectral counts were normalized to the amino acid length to account for total protein length, yielding a normalized spectral abundance factor (NSAF) (Zybailov et al. 2007). Results and Discussion Outer Membrane Vesicles and Tubes Screening of density gradient fractions revealed that OMVs were found between 1.12 and 1.15 g/ml in the gradient. Surprisingly, density gradient fractions containing 143 OMVs also contained tubular structures that appeared to be OMV precursor structures, and observed OMV-tubes are presumed to be different stages of OMV development (Figure 1). In addition, numerous individual OMVs and tube structures with what look like OMVs forming at each end were observed (Figure 1). Further, OMVs in chain patterns can be observed, possibly individual OMVs in the process of forming from tube structures or tubes in the process of forming individual OMVs (Figure 1c). OMV-tube structures have been recently observed in other bacteria such as Myxococcus xanthus, Francisella novicida, and Shewanella oneidensis and more organisms may produce similar structures, but OMV isolation methods typically exclude the tube form from the enrichments (Remis et al. 2013; McCaig et al. 2013; Pirbadian et al. 2014; Subramanian et al. 2016). Production of OMV-tubes instead of individual OMVs could be a “power in numbers” strategy that would guarantee a higher OMV concentration at a particular location rather than sending off individual OMVs with no control over localization or co- localization. OMVs varied in size between 20 to 60 nm in diameter and tubes were typically 20-30 nm in diameter. OMVs and OMV-tubes from D. vulgaris biofilms grown under EAL vs. BAL conditions did not differ in appearance. Due to some reports of OMVs containing DNA and being involved in horizontal gene transfer, isolated D. vulgaris OMV fractions were tested for the presence of DNA (Pérez-Cruz et al. 2013; Fulsundar et al. 2014). DNA was measured before and after a boiling step (5 min) and detectable DNA was not measured in either instance, leading to 144 the conclusion that OMVs produced by D. vulgaris grown under the EAL and BAL condition were not vehicles for DNA transfer. Figure 1. TEM images of isolated OMVs and OMV-tubes. Arrows in B and C point to OMV-tubes in different stages of vesicle differentiation. 145 Protein Gene ID NSAF BAL EAL Outer Membrane Porin DVU0799 0.114±0.082 0.109±0.023 Uncharacterized protein DVU0797 0.045±0.045 0.039±0.015 Lipoprotein, putative DVU1573 0.068±0.080 0.046±0.027 OmpA family protein DVU1422 0.034±0.045 0.029±0.024 Lipoprotein, putative DVU0761 0.069±0.088 0.036±0.028 Lipoprotein, putative DVU3249 0.018±0.022 0.010±0.011 Transporter, putative DVU0766 0.034±0.042 0.019±0.016 Lipoprotein, putative DVU2428 0.049±0.031 0.022±0.004 Lipoprotein, putative DVU3042 0.038±0.015 0.038±0.013 Peptidoglycan-associated lipoprotein, putative DVU3104 0.028±0.035 0.006±0.006 146 Lipoprotein, putative DVU3352 0.009±0.013 0.004±0.003 Uncharacterized protein DVU0255 0.012±0.016 0.004±0.003 Periplasm Periplasmic [NiFeSe] hydrogenase, large subunit DVU1918 0.067±0.067 0.078±0.033 Periplasmic [NiFe] hydrogenase, large subunit DVU1922 0.021±0.036 0 Periplasmic [NiFeSe] hydrogenase, small subunit DVU1917 0.024±0.030 0.021±0.010 TPR domain protein DVU1930 0.014±0.017 0.009±0.008 Cytoplasm Sulfite reductase, dissimilatory-type subunit beta DVU0403 0.030±0.008 0.035±0.005 Adenylyl-sulphate reductase, alpha subunit DVU0847 0.017±0.002 0.011±0.005 Sulfite reductase, dissimilatory-type subunit alpha DVU0402 0.016±0.003 0.012±0.005 Glyceraldehyde-3-phosphate dehydrogenase DVU0565 0 0.033±0.018 Inorganic pyrophosphatase, manganese- dependent DVU1636 0.021±0.039 0.007±0.007 Zinc resistance-associated protein homolog DVU3384 0.039±0.047 0.010±0.012 50S ribosomal protein L21 DVU0927 0 0.016±0.021 147 50S ribosomal protein L23 DVU1305 0.029±0.045 0.021±0.021 Rubrerythrin DVU3094 0.012±0.003 0.005±0.000 Rubredoxin-oxygen oxidoreductase DVU3185 0.004±0.001 0.002±0.002 Inner Membrane Amino acid ABC transporter DVU0966 0.025±0.027 0 Table 1. Proteins identified in purified OMV samples from D. vulgaris biofilm grown under BAL and EAL conditions. Protein abundance is estimated by normalized spectral abundance factor (NSAF). Proteomic Analysis of OMVs To understand the potential function of the OMVs in D. vulgaris biofilms grown under EAL and BAL conditions, protein content was analyzed by untargeted shotgun proteomics. The proteins detected for both the EAL and BAL conditions are presented in Table 1, organized based on cellular location. It is expected that OMVs contain proteins whose origin is the outer membrane or periplasmic space, but the detection of proteins belonging to the cytoplasm or inner membrane would indicate a specific sorting event or contamination of samples with cell lysis products during the OMV separation protocol. It is rare that proteomic studies of OMVs are free of cytoplasmic proteins, even when extremely stringent isolation protocols are used to avoid cell lysis (Schwechheimer and Kuehn 2015). In attempt to differentiate between cytoplasmic proteins detected due to contamination from lysed cells compared to intentional sorting, the Protein Abundance 148 Database (pax-db.org) was used to identify proteins of high abundance in D. vulgaris. Six out of ten cytoplasmic proteins detected in the OMV fractions were among the top 5% of proteins present in D. vulgaris based on a compilation of proteomic data, indicating that these are likely a contamination (Wang et al. 2012). Cytoplasmic associated proteins detected in OMV fractions that are typically less abundant in D. vulgaris cells are two ribosomal proteins (50S ribosomal proteins L21 and L23), rubrerythrin (provides protection from oxidative stress), and a zinc resistance-associated protein that has been shown to increase in abundance when D. vulgaris is exposed to oxygen (Mukhopadhyay et al. 2007). Interestingly, three out of the ten proteins of cytoplasmic origin that are associated with oxygen stress (rubrerthyrin, zinc resistance- associated protein, and rubredoxin-oxygen oxidoreductase), pointing toward a possible role for OMVs in oxidative stress protection. The most abundant protein detected in OMVs was a porin that was recently characterized and found to form a homotrimeric structure, have a slight preference for anionic sugars over uncharged sugars, and is similar in permeability to the major porin of Escherichia coli (Zeng et al. 2017). This porin (encoded by DVU0799) is one of the most abundant outer membrane proteins, indicating its importance in controlling outer membrane permeability. Given that this porin is highly abundant in the outer membrane, it is not surprising that it is also highly abundant on OMVs. Another highly abundant, but uncharacterized, protein in the OMV fractions is encoded by DVU0797, which is separated from DVU0799 in the genome by a small (31 AA) hypothetical 149 protein. DVU0797 is similar in size to DVU0799 and has 60% sequence identity, making it another candidate for a porin (Walian et al. 2012). Also present in D. vulgaris OMVs were eight different lipoproteins. Lipoproteins are typically associated with the cytoplasmic membrane, the inner leaflet of the outer membrane and the outer cell surface, as was recently demonstrated in Gram-negative cells (Wilson and Bernstein 2016). Lipoproteins have a wide range of functions, including nutrient acquisition, pathogenesis, sporulation, transport and folding of proteins, and membrane integrity (Mathiopoulos et al. 1991; Alloing et al. 1994; Berry and Paton 1996; Schmaler et al. 2010). Defects in lipoproteins have been shown to result in a decrease in membrane integrity and hyper-vesiculation, leading some researchers to propose that lipoproteins, or spacing of lipoproteins in the outer membrane, may be involved in OMV formation (Bernadac et al. 1998a; Cascales et al. 2002). D. vulgaris has 45 annotated lipoproteins in the genome, most of which are of unknown function. One of the eight that were identified in the isolated OMVs, DVU3104, has significant homology with Pal in E. coli (Walian et al. 2012), which is anchored in the outer membrane and binds with peptidoglycan, and when mutated causes hyper- vesiculation (Bernadac et al. 1998b). Several genes encoding lipoproteins in D. vulgaris were down-expressed after changing from syntrophic growth with a methanogen to a sulfidogenic metabolism, indicating that lipoproteins in D. vulgaris may also play an important role in interspecies interactions (Plugge et al. 2010). 150 Two periplasmic hydrogenases were identified in the OMV fractions as well. Both the [NiFeSe] hydrogenase and [NiFe] hydrogenase mature in the cytoplasm and are then transferred across the cytoplasmic membrane via the Tat system (Valente et al. 2005). The [NiFeSe] hydrogenase has been shown to also be a lipoprotein and it is hypothesized that the [NiFe] hydrogenase maybe is as well (Valente et al. 2007). Hydrogenases have an essential role in energy production in SRB as they oxidize H2, donate the electrons to the electron transport chain and produce protons for generation of a proton motive force that is used for ATP production (Peck 1993). Additionally, isolated hydrogenases from different Desulfovibrio sp. are capable of Cr(VI) reduction (Chardin et al. 2003). Whether or not these hydrogenases function in an OMV or if they happen to be packaged in OMVs due to their location in the periplasm is not clear. However, vesicles that contained hydrogenase activity could be supplied with an electron source via H2, and therefore, could feed electron transfer mechanisms away from the cell. Interspecies H2 transfer is an important component of syntrophic relationships, with microbes forming aggregates and biofilms structured to maximize the availability of substrates (Ishii et al. 2005). In the example of D. vulgaris and Methanococcus maripaludis, hydrogen production by the SRB and consumption by the methanogen is critical to the syntrophy and extracellular hydrogenase activity could contribute to such relationships (Brileya et al. 2014). Further work is needed to ascertain long-distance, intra-species H2 transfer in biofilms. 151 Another protein present in isolated OMVs contained a tetratricopeptide repeat domain (encoded by DVU1930) and has homology with CpoB, a cell division coordinator associated with regulating peptidoglycan synthesis and outer membrane invagination (Gray et al. 2015). A possible role for CpoB in isolated OMVs could be to regulate division of OMV-tube structures to individual OMVs, but it cannot be ruled out that its presence is a consequence of being an outer membrane associated protein. However, in a study that established an outer membrane protein dataset for D. vulgaris, this protein was not detected, further hinting at its involvement in OMV division (Walian et al. 2012). 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Mol Biosyst 3:354–360. 157 CHAPTER SIX EPILOGUE The motivation behind this work and the theme throughout this dissertation is understanding how nutrient limitation and resource ratio affect D. vulgaris growth, metabolism, and response to Cr(VI) toxicity in both planktonic and biofilm growth modes. While there have been many studies that have elucidated sensory systems in bacteria that control gene expression and therefore physiological responses to environmental changes, relatively few studies have investigated how nutrient limitation in terms of resource ratio affects microbial physiology, especially in response to heavy metal toxicity in both planktonic and biofilm growth modes. Investigating how nutrient limitation and resource ratio affected growth and metabolism of D. vulgaris Hildenborough in the biofilm growth mode led to the observation of complex extracellular structures and vesicles that warranted further investigation. The characterization of these structures and vesicles is the subject of Chapters 4 and 5. The response of D. vulgaris to Cr(VI) toxicity is a secondary theme throughout this dissertation. Chapter 2 presents the response of D. vulgaris to Cr(VI) when grown planktonically in batch mode. Under all conditions tested (20 and 30°C, and EAL, BAL, and EDL resource ratios), D. vulgaris responded to Cr(VI) toxicity with an increased lag time compared to growth without Cr(VI). The length of the lag time increased with increased Cr(VI) concentration, but surprisingly lag time and Cr(VI) reduction rate was 158 also linked to resource ratio, with cells grown under the EAL condition being much more susceptible to Cr(VI) toxicity compared to cells grown under the BAL condition. In attempt to explain this, sulfate concentration was tested as a possible factor that could affect Cr(VI) toxicity. Increased sulfate concentration did provide some protection from Cr(VI) toxicity, but when sulfate levels were normalized, resource ratio still affected Cr(VI) toxicity, measured by lag time. These results indicate that there may be a dual effect of sulfate concentration and resource ratio that dictates Cr(VI) susceptibility in D. vulgaris. The effect of Cr(VI) on D. vulgaris biofilm grown under EAL and BAL conditions was also measured to determine if growth under the EAL condition in the biofilm growth mode had a similar effect on D. vulgaris Cr(VI) susceptibility as in planktonic growth. Surprisingly, in the biofilm growth mode, D. vulgaris grown under the EAL condition was equally susceptible or more resistant to Cr(VI) toxicity compared to the BAL condition. This finding, opposite of the results for planktonic growth with Cr(VI), is surprising, but is consistent with the plethora of literature documenting the physiological differences between biofilm and planktonic growth modes (Xu et al. 2000; Stewart 2002; Harrison et al. 2007). Although it is difficult to directly compare Cr(VI) susceptibility between biofilm and planktonic cells for SRB due to abiotic Cr(VI) reduction by sulfide, it seems that regardless of resource ratio, the biofilm growth mode confers increased Cr(VI) resistance for D. vulgaris, which is consistent with other organisms, such as Pseudomonas sp. (Teitzel and Parsek 2003; Chien et al. 2013). 159 Although there is not as much information available on the topic of increased resistance to heavy metals in biofilms compared to planktonic growth, the information that is available suggests similar mechanisms that contribute to biofilm resistance to antibiotics may be at play. These include the presence of slow growing cells within the biofilm due to heterogeneity in nutrient availability, dead cells/cell debris within the biofilm that may react with or sorb metal ions, and the production of extracellular matrix materials that might chelate or reduce metal ions (Rani et al. 2007; Harrison et al. 2007). The presence of slow growing cells and the role that extracellular matrix components play in biofilm resistance to heavy metals is particularly relevant to the work presented in Chapters 3 and 4. Chapter 3 describes differences in the metabolism of D. vulgaris biofilms grown under EAL compared to BAL conditions. Biofilm grown under the EAL condition had many metabolites belonging to pathways essential for growth and cell division division down-regulated, indicating that D. vulgaris growing under the EAL condition may be growing more slowly, or in a biomass retention state. One theory for increased resistance of biofilms to external stressors (i.e. antibiotics, heavy metals), is that cells that are limited for an essential nutrient are forced to enter a very slow-growing or dormant state. Upon exposure to an external stress or antimicrobial agent, those cells within the biofilm that have nutrients available and therefore have a faster growth rate are more susceptible because they are prone to take up and metabolize the antimicrobial agent. The active cells also then serve to protect the less active cells by metabolizing the agent at their own 160 expense (Walters et al. 2003; Borriello et al. 2004). Further, linking purine/pyrimidine and glutamine/glutamate down-regulation to microbial resilience is a study in which removal of nucleobases and amino acids from medium supporting E. coli growth resulted in increased antibiotic resistance (Fung et al. 2010). The authors attribute increased resistance to induction of a stringent response, and/or additional unidentified mechanisms. Under the EAL condition, D. vulgaris is not limited for C or N, but sulfate limitation induces cellular regulation of C and N possibly through RexB or other unidentified mechanisms. The changes in metabolic state induced by electron acceptor limitation may therefore actually be protective in the biofilm growth mode whereas in the planktonic growth mode it is disadvantageous. In Chapter 4 extracellular membranous structures are characterized and found to be closely associated with metal precipitants that were determined to be iron oxides and iron phosphates. The presence of larger and more extensive extracellular membrane structures produced under the EAL condition could also explain the opposing results for the effect of resource ratio on Cr(VI) toxicity for planktonic compared to biofilm cells if these structures have Cr(VI) reduction/sorption/chelating properties. Future studies should attempt to visualize such extracellular membrane structures within the biofilm after exposure to Cr(VI) and locate Cr(III) deposition within the biofilm. While the exact function of the extracellular membranous structures still eludes us, there are multiple lines evidence supporting the hypothesis that they are involved in microbe-metal interactions. The first, mentioned above, is the association with iron 161 oxides/iron phosphates with the structures. There are metal deposits throughout the biofilm, with some seemingly far from cells and any extracellular structures and others adjacent to cells, but the association of metal precipitates with the extracellular membranous structures is quite notable. The identification of these precipitates as iron oxides and iron phosphates was surprising, as we would have otherwise assumed that they were iron sulfide precipitates since sulfide is abundant in the biofilm and readily reacts with ferrous iron (which is in the medium at a concentration of 0.063 mM as FeCl2) to form mackinawite (FeS) or greigite (Fe3S4) (Haaningnielsen et al. 2005; Gramp et al. 2010). Because samples were processed and stored aerobically, it is possible that iron sulfides were oxidized to iron oxides in the time between sampling and analysis (Rickard and III Luther 2007). Still, it is surprising that no sulfur was detected and so we cannot rule out the possibility that these iron phosphates are not artifacts from sample preparation. The second piece of evidence supporting the hypothesis that the extracellular membranous structures are involved in microbe-metal interactions is the presence of hydrogenases in OMVs. While it would be difficult to isolate the extracellular membranous structures away from the rest of the biofilm for further analysis, outer membrane vesicles were sometimes associated with and connecting to extracellular membranous structures and OMVs could be isolated and analyzed in an indirect attempt to learn more about the function of the extracellular membranous structures. Isolated OMVs contained [NiFe] and [NiFeSe] hydrogenases, which are critical to electron 162 transfer and hydrogen production and oxidation in cells. Extracellular hydrogenase activity in a biofilm has many implications for electron and hydrogen transfer over long distances. Additionally, hydrogenases isolated from SRB have been observed to reduce Cr(VI) (Michel et al. 2001; Chardin et al. 2003). Whether or not hydrogenases located within OMVs are capable of Cr(VI), or other metal, reduction if H2 is present has not been tested, but these results suggest that OMVs could potentially contribute to metal reduction and resistance in both biofilm and planktonic growth modes. Further, the presence of hydrogenases in OMVs and the observation that OMVs are associated with extracellular membranous structures suggests that these two extracellular matrix components may be linked in structure and/or function. Third, the overproduction of these structures under the EAL condition compared to BAL and EDL, suggests that their production may be in attempt to locate an alternative electron acceptor. Other dissimilatory metal-reducing organisms such as Shewanella sp. and Geobacter sp. produce extracellular nanowires in response to electron acceptor limitation (Gorby et al. 2006; Bond et al. 2012). While the topic of nanowires is one of intense debate and current hypotheses and models are continually changing, the production of extracellular structures in response to electron acceptor limitation in these other dissimilatory metal-reducing bacteria is well established. Extracellular electron transport has not been demonstrated and outer membrane cytochromes have not been detected in Desulfovibrio sp., making use of extracellular insoluble electron acceptors unlikely; however, there are parallels between the extracellular membrane structures and 163 nanowires produced by S. oneidensis. The most recent description of S. oneidensis nanowires shows that they are actually extensions of the outer membrane and periplasmic space that then differentiate into OMVs, which is seemingly very similar to the OMV- tube structures that we observed with the isolated OMVs (Pirbadian et al. 2014; Subramanian et al. 2016). Further, direct extracellular electron transfer via extracellular cytochromes is not the only mechanism for extracellular electron transfer. Transport of electrons via hydrogen (interspecies hydrogen transfer) is the foundation for many syntrophic microbial relationships, such as those between SRB, methanogens, or other fermentative microorganisms (Iannotti et al. 1973; Nakamura et al. 2010) and extracellular hydrogenases could facilitate the transfer of electrons from H2 to an exogenous electron acceptor. While all of this taken together suggests that extracellular membranous structures play a role in microbe-metal interactions, other possible functions of these structures, such as waste extrusion and intercellular communication cannot be ruled out. A unique aspect of the work presented in this dissertation is that all of the experiments were carried out at a room temperature (20-23°C), which is lower than the optimal growth temperature of 30°C for D. vulgaris. The experiments in the majority of published studies on dissimilatory metal-reducing bacteria are performed at 30°C, probably to be consistent with the body of literature available for comparison and to increase growth rate, and therefore reduce experiment length. With the rationale of perturbing resource ratio to better mimic “real world” scenarios, we thought it also 164 important to culture D. vulgaris at a temperature that was closer to subsurface temperatures (~5-17°C at Hanford Site, WA) (Truex et al. 2012). Decreased temperature resulted in a decreased growth rate for D. vulgaris, which is expected. Cells grown at 20°C were also more susceptible to Cr(VI) toxicity as measured by increased lag time and had slower Cr(VI) reduction rates compared to 30°C. A temperature increase of 10°C in a sulfate-reducing consortium resulted in a uranium reduction rate with three times the rate constant, indicating that temperature, growth rate, and enzyme reaction rates are linked (Boonchayaanant et al. 2008). The Cr(VI) reduction rate at 20°C was ~2 times slower than at 30°C, which most likely contributed to increased Cr(VI) toxicity at the lower temperature. Additionally, temperature also affected production of extracellular filaments in D. vulgaris biofilms regardless of electron donor/electron acceptor ratio. Relatively few filaments were observed in biofilms grown at 30°C compared to those grown at 20°C and filament production in biofilms grown at 25°C was more than that of 30°C, but less than that of 20°C, suggesting that extracellular filament production is temperature dependent (Appendix A). To rule out that filament production was not based on growth rate, biofilms were also grown at 30°C, but at the dilution rate appropriate for a reactor grown at 20°C. This thermoregulation of filament production is not unique to D. vulgaris, as Geobacter sulfurreducens also produces pili at 25°C, but not 30°C and E. coli produces fimbrae at 37°C, but not 25°C (Goransson et al. 1989; Cologgi et al. 2011). Although we cannot determine what these filaments are in relation to the extracellular membranous 165 structures described in Chapter 4 due to possible sample distortion from ethanol dehydration and critical point drying, the presence of these extracellular structures at lower temperatures further emphasizes the importance of culturing organisms at temperatures that may be sub-optimal but field-relevant. The work presented here investigated the effects of nutrient limitation and resource ratio on D. vulgaris growth, metabolism, and Cr(VI) susceptibility in the planktonic and biofilm growth modes. Measuring growth alone, whether planktonically or as a biofilm, revealed that D. vulgaris maximum biomass production is dictated by sulfate limitation in the EAL condition and lactate limitation in the EDL condition, which was expected. Cells grown under the EAL condition could have a growth advantage because they are able to produce H2 for a short time after sulfate is depleted. This was measured in planktonic batch tube growth, but could not be detected in a CDC biofilm reactor (possibly due to H2 consumption within the biofilm or the limit of detection on the gas chromatograph) and biomass yields were not significantly higher under the EAL condition, indicating that if H2 production was occurring, it was not at a high enough level to contribute bo biomass production. While measuring microbial growth in response to a stress and with different substrates or varying nutrient concentration can be an elegant way of discerning microbial physiology, complementing growth experiments with other analytical and “omics” approaches was instrumental to understanding how nutrient limitation and resource ratio affected D. vulgaris metabolism and biofilm matrix composition. The 166 combination of microscopy, differential staining, metabolomics, and fatty acid quantification in D. vulgaris biofilms was essential to the discovery and characterization of lipids as an important component of the biofilm matrix. Based on the findings presented here, we conclude that nutrient limitation and resource ratio have notable effects on planktonic D. vulgaris Cr(VI) susceptibility and reduction, with electron acceptor limitation causing decreased Cr(VI) reduction rates and increased cell death in response to Cr(VI). In the biofilm growth mode, nutrient limitation altered the flow of carbon and energy in D. vulgaris, with electron acceptor limitation causing a switch to a biomass retention state with increased fatty acid production. Further, fatty acid content of biofilms was visualized and quantified and corresponded with extensive extracellular structures that connected to cells and were associated with extracellular metal deposits. The function of these structures is undetermined, but we hypothesize that they are involved in microbe-metal interactions and may confer protection against external stressors or antimicrobial agents. The identification of extracellular membranes as an important constituent of the biofilm matrix is significant because they have not been previously recognized, and because they may also be important to consider from an evolutionary perspective. Since containment of organelles within a membrane is a distinguishing factor between eukaryotic and prokaryotic organisms, the presence of extracellular membranes in biofilms where organisms from different domains of life are potentially coexisting and cooperating, has significant implications for the evolution of more complex life forms. 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Agency for Toxic Substances and Disease Registry (US), Atlanta (GA) 198 APPENDICES 199 APPENDIX A PRODUCTION OF EXTRACELLULAR FILAMENTS IN DESULFOVIBRIO VULGARIS BIOFILMS IS TEMPERATURE-DEPENDENT 200 Introduction Initial experiments to characterize Desulfovibrio vulgaris Hildenborough biofilm were carried out at 30°C (optimal growth temperature) and room temperature (20-23°C) to determine if culturing biofilm at room temperature was feasible. Biofilms grown at 30, 25, and 20°C were examined for extracellular matrix structure via SEM. Materials and Methods Biofilm Growth Conditions Biofilm samples were grown in a CDC reactor (Biosurface Tech.) under electron acceptor limitation at 30°C, 25°C or 20°C in LS4D media described in Chapter 3. Dilution rates (D) were dependent on growth rate at said temperature with the exception of the sample shown in Figure 1b. Biofilm grown on glass slides were sampled at 72 hours for samples grown at 30°C and 168 hours for samples grown at 25 and 20°C. Scanning Electron Microscopy and Sample Preparation Biofilm coated slides were gently washed with phosphate buffered saline to remove loosely attached cells and then fixed overnight in a 2% paraformaldehyde, 2.5% gluturaldehyde, 0.05M sodium cacodylate solution. Samples were then dehydrated with increasing amounts of ethanol (25%, 50%, 75%, 95%, 100%x3 v/v EtOH, 20 min each treatment) and then critical point dried in a Tousimis Samdri-795 drier to preserve bacterial cell structure. Samples were mounted on aluminum stubs, coated with iridium, 201 and viewed on a Zeiss SUPRA 55VP field emission scanning electron microscope at 1.0 kV. Results Extracellular filament production in D. vulgaris biofilms grown at 30, 25, and 20°C varied according to temperature with biofilms grown at 30°C producing very few extracellular filaments (Figure 1a). Biofilm grown at room temperature (~20°C) produced many extracellular filaments that connected cells and cells to the surface (Figure 1d). Biofilm grown at a middle temperature of 25°C produced an intermediate number of extracellular filaments (Figure 1c). To determine if filament production was actually dependent on temperature or if it could be linked to growth rate/dilution rate, biofilm grown in a third reactor at 30°C, but with the same dilution rate of those grown at 20°C (D=0.04-hr) was examined. These samples showed minimal filament production (Figure 1b), similar to those grown at 30°C with a higher dilution rate, indicating that filament production is temperature-dependent, not growth rate-dependent. 202 Figure 1. FE-SEM images of D. vulgaris biofilm grown at different temperatures and dilution rates. 203 APPENDIX B DESULFOVIBRIO CARBINOLIPHILUS OAKRIDGENSIS, SPP. NOV., AN ORGANIC ACID-OXIDIZING, SULFATE REDUCING BACTERIUM ISOLATED FROM URANIUM(VI)-CONTAMINATED GROUNDWATER Contribution of Authors and Co-Authors Manuscript in Appendix B Author: Bradley D. Ramsey Contributions: Developed experimental design, performed experiments, analyzed data, wrote manuscript. Co-Author: Lauren C. Franco Contributions: Developed experimental design, performed experiments, analyzed data, wrote and revised manuscript. Co-Author: Matthew W. Fields Contributions: Developed experimental design, analyzed data, wrote and revised manuscript. 204 Manuscript Information Page Bradley D. Ramsay, Lauren C. Franco, Matthew W. Fields International Journal of Systematic and Evolutionary Microbiology Status of Manuscript: __x_ Prepared for submission to a peer-reviewed journal ____ Officially submitted to a peer-review journal ____ Accepted by a peer-reviewed journal ____ Published in a peer-reviewed journal The Microbiology Society 205 Abstract A novel sulfate-reducing bacterium was isolated from a uranium and nitrate contaminated well (FW-101) at the Oak Ridge Field Research Center. Phylogenetic analysis revealed that it is closely related to Desulfovibrio carbinoliphilus D41T, although physiological studies indicate distinguish the two strains. The isolate could utilize pyruvate, lactate, formate, malate, maleate, methanol, ethanol, 1,2-propanediol, 1,3- propanediol, and fumarate and substrates and sulfate, sulfite, and thiosulfate as electron acceptors. Nitrate could be used as an N-source if ammonium was absent, but could not be used as an electron acceptor. The isolate grew in the presence of 100µM U(VI) or Cr(VI), but could not use U(VI) or C(VI) as a sole electron acceptor linked to growth. Based on the similarities in phylogeny, but distinctions in growth characteristics between this isolate and D. carbinoliphilus D41T, the novel strain Desulfovibrio carbinoliphilus Oakridgensis is proposed. 206 Introduction The Field Research Center (FRC) at Oak Ridge National Lab (ORNL) offers a unique research opportunity to study bioremediation in situ. Part of the Y-12 security complex, the FRC is located in the Bear Creek drainage near Oak Ridge, Tennessee. Beginning in the 1950s, and ending in 1983, an estimated 300 million liters of mixed acid, organics, and heavy metal wastes were deposited in four unlined waste disposal ponds. Capped with an asphalt parking lot in 1988, wastes from this area have seeped into the underlying bedrock and groundwater table. Some of the many mobile subsurface contaminants at this site include nitrate, uranium, thorium, technetium, and several volatile organics (Wu et al. 2006). ICP-MS of samples collected in late 2001 from well FW-101 has yielded uranium concentrations between 20 and 250ppm, and chromium concentrations between 35 and 85ppm. For comparison, US EPA drinking water standards in April 2017 for uranium and total chromium are 30ppb and 100ppb, respectively (www.epa.gov). Because of these highly contaminated zones within the FRC, it is hypothesized that novel denitrifying and metal-reducing microorganisms may live in these areas and therefore attempts to culture them could lead to discoveries of new metabolisms or lineages. Dissimilatory metal-reducing bacteria are of particular interest at sites such as the ORNL FRC due to their ability to transform contaminants such as U(VI) and Cr(VI) from soluble to insoluble forms. One group capable of such metal reduction is the sulfate- 207 reducing bacteria (SRB) which contains the genus Desulfovibrio. Belonging to the ẟ- proteobacteria class, the Desulfovibrio genus consists of flagellated, Gram-negative curved-rods, that live in a variety of anoxic environments. Representatives from Desulfovibrio have been found in environments from cold (Vandieken et al. 2006; Sasi Jyothsna et al. 2008) to hot (Alazard 2003); from acidic (Karnachuk et al. 2015) to basic (Abildgaard 2006); and from freshwater to hypersaline (Magot et al. 2004). While sulfate is the preferred electron acceptor, electron donors for Desulfovibrio include alcohols (Ouattara et al. 1992; Allen et al. 2008), sugars (Ollivier et al. 1988), amino acids (Baena et al. 1998), and a number of different organic acids (Sun et al. 2000). Previous studies have demonstrated reduction of nitrate and uranium levels in the subsurface upon bio-stimulation with ethanol (Hwang et al. 2009). During bio- stimulation, an increase was observed in DNA sequences corresponding to SRB. Here we present physiological and phylogenetic characterization of one novel SRB isolated from the ORNL FRC that belongs to the genus Desulfovibrio. Materials and Methods Groundwater from well FW-101 at the FRC site was collected during the uranium-reduction phase of a previously described biostimulation experiment (Hwang et al. 2009) and used as inoculum for a culture enriching for sulfate-reducing bacteria. The enrichment was grown at room temperature anaerobically in ES4D media with the 208 following composition: 40mM ethanol (E), 50mM sodium sulfate (S4), 8mM magnesium chloride, 20mM ammonium chloride, 2.2mM orthophosphate, 600µM calcium chloride, 1% (w/v) PIPES, 0.1% (v/v) 1000X Thauer’s vitamins (Brandis and Thauer 1981), and 1.25% (v/v) 80X trace minerals solution. pH of the media was not adjusted, and was approximately 6.7. The trace mineral solution consisted of 50mM nitrilotriacetic acid, 5mM iron(II) chloride, 2.5mM manganese(II) chloride, 1.3mM cobalt(II) chloride, 1.5mM zinc chloride, 380M nickel(II) sulfate, 320M boric acid, 210M sodium molybdate, 30M sodium selenite, 20M sodium tungstate and 10M copper(II) chloride, pH adjusted to 6.5. All growth conditions were performed anaerobically under an atmosphere of 100% nitrogen. The grown enrichment culture was transferred for increased growth rates of SRBs into liquid LS4D media: ES4D where ethanol was replaced with 40mM lactate (L), and 2.2mM orthophosphate replaced with 2.2mM potassium phosphate. Cultures grown in LS4D cultures were plated on solid LS4D agar: LS4D with 1.5% (w/v) agar added. Plates were incubated at 25ºC in a glove-bag with an atmosphere of nitrogen with ~2.5% hydrogen. Several colony morphologies arose; each was re-streaked and subsequently transferred into liquid LS4D media and grown at room temperature. For sequencing of the small-subunit (SSU) ribosomal RNA gene of the isolate FW-101-2B, standard PCR methods were used to amplify the gene with previously described primers (Ye et al. 2004): FD1 (5’-AGAGTTTGATCCTGGCTCAG-3’) and 1540R (5’-AAGGAGGTGATCCAGCC-3’). The resulting ~1500bp band was submitted 209 for capillary Sanger sequencing using each of the following primers, previously described (Ye et al. 2004): FD1, 350R (5’-CTGCTGCSYCCCGTAG-3’), 519F (5’- CAGCAGCCGCGGTAA-3’), 529R (5’-CGCGGCTGCTGGCAC-3’), 788F (5’- ATTAGATACCCTGGTA-3’), 925R (5’-CCGTCAATTCMTTTRAGTTT-3’), 1099F (5’-GCAACGAGCGCAACCC-3’), and 1540R. Resulting sequences were spliced to form one contiguous sequence representing the entire gene which was entered into NCBI’s nucleotide BLAST engine (http://www.ncbi.nlm.nih.gov/blast/Blast.cgi). Whole genome sequencing of isolate FW-101-2B was performed by the Joint Genome Institute. The sequence for the organism’s dsrAB gene was obtained from the genome sequence. The dsrAB gene from D. carbinoliphilus D41T was amplified using standard PCR methods, and the sequence was determined by capillary Sanger sequencing, using previously described primers in both cases (Wagner et al. 1998; Giloteaux et al. 2010). 16s small subunit ribosomal RNA sequences were aligned with the SILVA Incremental Aligner (Pruesse et al. 2012) and a phylogenetic tree was constructed using the RAxML maximum likelihood algorithm with an inverse gamma distribution (Ludwig et al. 2004). dsrAB phylogenetic trees were constructed using the same method but with a dsrAB sequence database (Müller et al. 2015). Batch cultures of isolate FW-101-2B were used to determine growth characteristics in differing conditions. To test for utilized phosphate sources, cells grown in LS4D were transferred into tubes containing either potassium phosphate, 210 triethylphosphate, sodium metaphosphate, or sodium trimetaphosphate in place of 2.2mM orthophosphate, each added so that the total phosphorus input was 2.2mM. Each condition was performed once. Metabolism of different electron donors was tested in duplicates in LS4D media with 40mM of each of the following added: methanol; ethanol; 1-propanol; 2-propanol; 1,2-propanediol; 1,3-propanediol; glycerol (1,2,3-propanetriol); 1-butanol; 2-butanol; 1,4-butanediol; 2,3-butanediol; isoamyl alcohol (3-methyl-1-butanol); benzyl alcohol; 2- phenylethanol; sodium formate; sodium acetate; propionic acid; sodium butyrate; sodium lactate; sodium pyruvate; sodium malate; sodium maleate; sodium fumarate; D-fructose; and triethylphosphate. The poly-lactate mixtures Metals Remediation Compound and Hydrogen Releasing Compound (Regenesis, San Clemente, CA) were also tested at an overall lactate concentration of 40mM. Those cultures which grew with both lactate and the additional carbon source were transferred in duplicates into fresh tubes with only 40mM of the substrate in question. Positives were scored by an increase in turbidity (OD600) and successful growth after a transfer into fresh media. As pyruvate was the preferred carbon source and potassium phosphate was utilized as a phosphorus source, the remaining growth tests were carried out using pyruvate (P) and potassium phosphate (PS4D). To determine optimum growth temperature, isolate FW-101-2B was grown in PS4D at 4, 10, 16, 25, 30, 37, 42, and 60ºC in duplicates while monitoring the increase in optical density at 600nm. To determine optimum pH for growth initiation, PS4D was 211 prepared without PIPES but with 30mM potassium phosphate in its stead, then was adjusted with sodium hydroxide or hydrochloric acid to pH 4.5, 5, 6, 6.5, 7, and 8.5. Media above pH 6.5 required the magnesium concentration to be lowered from 8mM to 4mM due to precipitation with dibasic phosphate ions. Growth was performed in triplicates at each pH. PS4D and LS4D media lacking ammonium chloride was prepared to test the isolate for nitrate reduction and assimilation. A culture of FW-101-2B grown in normal PS4D or LS4D was pelleted and rinsed in nitrogen-free PS4D/LS4D, re-pelleted and inoculated into fresh nitrogen-free media to an OD600 of ~0.1. Nitrate was added to media containing sulfate at 0, 1, or 10 mM concentrations and with or without ammonium added. Nitrate was measured with ion chromatography over the growth period. Ammonium was measured with a Hach Ammonium Ion Selective Probe according to manufacturer protocols (Hach, CO). Four electron acceptors were tested with the isolate: sulfate, sulfite, thiosulfate, and fumarate. PS4D media was prepared without sulfate (P--D), and electron acceptors were added individually for each experiment. Cells grown in normal PS4D were pelleted and rinsed in P--D, re-pelleted and inoculated into fresh media to an OD600 of ~0.1. Growth was tested on sulfite, thiosulfate, and fumarate at 50mM concentration with pyruvate as the carbon source and electron donor. Uranium(VI) and chromium(VI) tolerance tests were carried out in PS4D with the sulfate concentration reduced to 25mM. The isolate was grown in duplicates in varying 212 concentrations of uranyl chloride, UO2Cl2: 1M, 10M, and 100M. Chromium(VI) toxicity was assessed in duplicates by growth in varying concentrations of potassium chromate, K2CrO4, with 25mM sulfate: 20mM, 50mM, and 100mM. Results and Discussion One colony chosen for study, isolate FW-101-2B formed a very dark brown glossy colony on solid LS4D agar approximately 3mm in diameter and 4mm tall. Viewed from underneath, the colony was light tan colored. A BLAST search of the SSU rRNA gene sequence revealed the isolate’s closest known relative is Desulfovibrio carbinoliphilus D41T at 99.0% identity. Although the SSU rRNA sequence showed remarkable similarities, whole genome sequencing of the isolate has revealed a G+C DNA content of 66.5%, compared to 63% in D. carbinoliphilus (Allen et al. 2008; Ramsay et al. 2015). Based upon SSU rRNA sequences, isolate FW-101-2B belongs to a clade of Desulfovibrio species including D. fructosovorans, D. marrakechenisis, D. alcoholvorans, D. carbinoliphilus, D. aerotolerans, D. burkinensis, D. carbinolicus, and D. magneticus. The tree of these species placed in the context of other Desulfovibrio spp. is shown below (Figure 1). Based upon dsrAB sequences, isolate FW-101-2B belongs to a clade of the same organisms. The dsrAB gene of the isolate matches the sequence from Desulfovibrio carbinoliphilus D41T at 91.64% identity. The tree of dsrAB Desulfovibrio sequences is shown in Figure 2. 213 Figure 1. Maximum likelihood phylogenetic tree of 16s small subunit RNA sequences from representatives of the Desulfovibrio genus. Branch length indicates number of nucleotide substitutions per site. 214 Figure 2. Maximum likelihood tree of the dissimilatory sulfite reductase alpha and beta subunit (dsrAB) genes from species belonging to the Desulfovibrio genus. Branch lengths represent nucleotide substitutions per site. Isolate FW-101-2B was found to thrive in many different growth conditions. No difference in growth was observed between each of the phosphate sources tested. Substrates that supported growth were methanol; ethanol; 1,2-propanediol; 1,3- propanediol; formate; fumarate; maleate; malate; lactate; and pyruvate. Presence of 40mM benzyl alcohol, propionic acid, hydrogen releasing compound, and metals remediation compound were inhibitory for growth. A trend towards relative non-toxicity of secondary alcohols and diols compared to primary alcohols is noticeable. The maximum growth rate was determined to lie between 30 and 37ºC with no measured growth occurring at 42ºC or warmer. Growth at 10ºC and colder was extremely slow. Growth rates of cultures at pH values greater than 7 were skewed due to vast production of iron sulfide. No difference in growth rate was detected by changing 215 the magnesium concentration at pH 6.5. Interestingly, final pH values were measured after growth and showed a sharp trend of neutralization towards pH 6.5, ranging from 5.75 to7.1. Only pH 6.5 remained unchanged. Only miniscule growth was detected below pH 6. Although optical growth determination is not reliable in the basic pH conditions due to mass production of sulfides, it is believed that the organism’s preferred pH is at or very near 6.5. Cultures provided with nitrate as their only electron acceptor did not grow and there was no decrease in nitrate concentration over time, indicating that this organism cannot use nitrate as an electron acceptor. Cultures provided with sulfate and nitrate, but no ammonium could grow and they consumed ~1.5mmol nitrate, indicating that the isolate is capable of assimilatory nitrate reduction. While no growth occurred when nitrate was the only nitrogen source, but the isolate grew with sulfate in the presence of nitrate, the organism appears to be nitrate tolerant, but will not reduce nitrate to meet nitrogen demands. Nitrate tolerance is expected considering the high nitrate levels present at the organism’s source of isolation. FW-101-2B cultures grew with sulfite and thiosulfate as electron acceptors. Growth on sulfite was similar to that on sulfate, but with thiosulfate growth occurred at a diminished rate (0.019-hr compared to 0.029-hr) and reached a lower maximum optical density (0.99 compared to 1.26). Two likely fates of thiosulfate are reduction and disproportionation of thiosulfate (Jackson and McInerney 2000). Both lead to double the HS- production generated during sulfate reduction, but thiosulfate reduction also 216 generates sulfite. The isolate could not use fumarate as an electron acceptor. P--D media with no electron acceptor supported very slow growth representative of pyruvate fermentation. Growth was not affected by addition of uranium(VI) at tested concentrations up to 100µM. Although the isolate experienced significant lag phases in 100µM chromate, otherwise normal growth occurred in each concentration. No growth occurred in the presence of U(VI) or Cr(VI) without sulfate. Considering the high level of uranium and chromium toxicity in this organism’s source environment, these findings are not unexpected. 217 Table 1. Summary of substrate and electron acceptor utilization of FW-101 isolate and closely related strains. 1, D. carbinoliphilus sp. nov. Oakridgensis; 2, D. carbinoliphilus D41T (Allen et al. 2008); 3, D. alcoholovorans (Qatibi and Garcia 1990); 4, D. fructosovorans (Ollivier et al. 1988); 5, D. aerotolerans (Mogensen et al. 2005); 6, D. magneticus (Sakaguchi et al. 2002); 7, D. carbinolicus (Nanninga and Gottschal 1987). ND - not determined, i - incomplete, w - weak. 218 Description of Desulfovibrio carbinoliphilus Oakridgensis spp. nov. Desulfovibrio carbinoliphilus Oakridgensis (oak · ridg · en´ · sis M.L. fem. adj. Oakridgensis coming from Oak Ridge, the city in the state of Tennessee whence the organism was isolated). Isolate FW-101-2B will utilize inorganic phosphate, metaphosphate, orthometaphosphate, and triethylphosphate has phosphorus sources. Carbon sources and electron donors utilized are methanol, ethanol, 1,2-propanediol, 1,3-propanediol, formate, fumarate, malate, maleate, pyruvate, and lactate. D. carbinoliphilus Oakridgensis will reduce sulfate, sulfite, and thiosulfate and will ferment pyruvate in the lack of electron acceptor. The organism has a maximum growth temperature of 30-37ºC and an optimum pH of 6.5. D. carbinoliphilus Oakridgensis cannot use nitrate as a dissimilatory electron acceptor, but it can assimilate nitrate when ammonium is not available. Finally, D. carbinoliphilus Oakridgensis is tolerant to soluble uranium and chromate at tested levels up to 100µM. Although physiological evidence would support classification of a new species, isolate FW-101-2B is phylogenetically very similar to D. carbinoliphilus D41T based upon small-subunit rRNA and dsrAB sequences, thus we propose its classification as new strain Desulfovibrio carbinoliphilus strain Oakridgensis. The GenBank accession number for the small-subunit ribosomal RNA gene of D. carbinoliphilus Oakridgensis is GU176294. 219 References Abildgaard L (2006) Desulfovibrio alkalitolerans sp. nov., a novel alkalitolerant, sulphate-reducing bacterium isolated from district heating water. Int J Syst Evol Microbiol 56:1019–1024. Alazard D (2003) Desulfovibrio hydrothermalis sp. nov., a novel sulfate-reducing bacterium isolated from hydrothermal vents. Int J Syst Evol Microbiol 53:173–178. 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Wu W-M, Carley J, Fienen M, Mehlhorn T, Lowe K, Nyman J, Luo J, Gentile ME, Rajan R, Wagner D, Hickey RF, Gu B, Watson D, Cirpka OA, Kitanidis PK, Jardine PM, Criddle CS (2006) Pilot-Scale in Situ Bioremediation of Uranium in a Highly Contaminated Aquifer. 1. Conditioning of a Treatment Zone. Environ Sci Technol 40:3978–3985. 222 Supplemental Figures Figure S1. Growth of D. carbinoliphilus Oakridgensis at 10, 16, 25, 30, and 37°C. 223 Figure S2. Growth of D. carbinoliphilus Oakridgensis at pH 4, 5, 6, 6.5, 7, and 8.5. 224 Figure S3. Growth of D. carbinoliphilus Oakridgensis with different substrates and sulfate as the electron acceptor. 225 Figure S4. Growth of D. carbinoliphilus Oakridgensis with nitrate and/or ammonium as the nitrogen source. 226 Figure S5. Growth of D. carbinoliphilus Oakridgensis with different electron acceptors. 227 Figure S6. Growth of D. carbinoliphilus Oakridgensis with Cr(VI) or U(VI). 228 APPENDIX C AUTONOMOUS METABOLOMICS FOR RAPID METABOLITE IDENTIFICATION IN GLOBAL PROFILING Contribution of Authors and Co-Authors Manuscript in Appendix C Author: Paul H. Benton Contributions: Experimental design, performed experiments, data acquisition and analysis, prepared and revised manuscript. Co-Authors: Julijana Ivanisevic, Nathaniel G. Mahieu, Michael E. Kurczy, Caroline H. Johnson, Lauren Franco, Duane Rinehart, Elizabeth Valentine, Harsha Gowda, Baljit K. Ubhi, Ralf Tautenhahn, Andrew Gieschen, Matthew W. Fields, Gary J. Patti Contributions: Performed experiments, revised manuscript. Co-Author: Gary Siuzdak Contributions: Experimental design, data analysis, prepared and revised manuscript. 229 Manuscript Information Page H. Paul Benton, Julijana Ivanisevic, Nathaniel G. Mahieu, Michael E. Kurczy, Caroline H. Johnson, Lauren Franco, Duane Rinehart, Elizabeth Valentine, Harsha Gowda, Baljit K. Ubhi, Ralf Tautenhahn, Andrew Gieschen, Matthew W. Fields, Gary J. Patti, and Gary Siuzdak Analytical Chemistry Status of Manuscript: ____ Prepared for submission to a peer-reviewed journal ____ Officially submitted to a peer-review journal ____ Accepted by a peer-reviewed journal __x_ Published in a peer-reviewed journal ACS Publications 2015, 87 (2), pp 884–891 230 Autonomous Metabolomics for Rapid Metabolite Identification in Global Profiling H. Paul Benton,† Julijana Ivanisevic,† Nathaniel G. Mahieu,‡ Michael E. Kurczy,† Caroline H. Johnson,† Lauren Franco,§ Duane Rinehart,† Elizabeth Valentine,# Harsha Gowda,†,¶ Baljit K. Ubhi,∫ Ralf Tautenhahn,†,∥ Andrew Gieschen,⊥ Matthew W. Fields,§ Gary J. Patti,*,‡ and Gary Siuzdak* †Scripps Center for Metabolomics and Mass Spectrometry, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, California 92037, United States ‡Departments of Chemistry, Genetics and Medicine, Washington University, One Brookings Drive, St. Louis, Missouri 63130, United States §Department of Microbiology and Immunology and Center for Biofilm Engineering, Montana State University, 109 Lewis Hall, Bozeman, Montana 59717, United States ∫ AB SCIEX, 1201 Radio Road, Redwood City, California 94065, United States ⊥Agilent Technologies, 11011 North Torrey Pines Road, La Jolla, California 92037, United States #The Skaggs Institute for Chemical Biology, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, California 92037, United States *S Supporting Information ABSTRACT: An autonomous metabolomic workflow com- bining mass spectrometry analysis with tandem mass spectrometry data acquisition was designed to allow for simultaneous data processing and metabolite characterization. Although previously tandem mass spectrometry data have been generated on the fly, the experiments described herein combine this technology with the bioinformatic resources of XCMS and METLIN. As a result of this unique integration, we can analyze large profiling datasets and simultaneously obtain structural identifications. Validation of the workflow on bacterial samples allowed the profiling on the order of a thousand metabolite features with simultaneous tandem mass spectra data acquisition. The tandem mass spectrometry data acquisition enabled automatic search and matching against the METLIN tandem mass spectrometry database, shortening the current workflow from days to hours. Overall, the autonomous approach to untargeted metabolomics provides an efficient means of metabolomic profiling, and will ultimately allow the more rapid integration of comparative analyses, metabolite identification, and data analysis at a systems biology level. Untargeted metabolomic experiments performed oncomplex biological matrices with high-resolution, high- throughput, and sensitive mass spectrometry (MS) technology have enabled the detection of thousands of metabolite features from a single experiment.1−4 However, the identification of these features presents a major bottleneck in the metabolomics workflow. It is not only a time-consuming process, taking weeks to carry out, but often results in a low yield of correctly identified metabolites. This is partly due to the manual interpretation required by the investigator and also the number of metabolites currently characterized in metabolite databases.5 This process can be potentially shortened by integrating metabolite profiling and identification into a single autonomous workflow. Current metabolomic studies typically adopt a multistep workflow (Figure 1). Comparative profiling is first carried out in MS mode; the data is then processed using bioinformatics software such as XCMS6,7 to reveal features of interest that show statistically significant differences. The features are then subjected to tandem mass spectrometry (MS/MS) acquisition and identified through MS/MS matching to standards in metabolite databases such as METLIN.8,9 This conventional approach typically involves the generation of a list of dysregulated metabolite features from an initial set of experiments followed by statistical analysis and manual selection of precursor ions, which are then fragmented to obtain mass spectra used for metabolite characterization.10 Received: July 11, 2014 Accepted: December 12, 2014 Published: December 12, 2014 Article pubs.acs.org/ac © 2014 American Chemical Society 884 DOI: 10.1021/ac5025649 Anal. Chem. 2015, 87, 884−891 This is an open access article published under an ACS AuthorChoice License, which permits copying and redistribution of the article or any adaptations for non-commercial purposes. 231 232 233 234 235 236 237 238 APPENDIX D COMPREHENSIVE BIOIMAGING WITH FLUORINATED NANOPARTICLES USING BREATHABLE LIQUIDS Contribution of Authors and Co-Authors Manuscript in Appendix D Author: Michael Kurczy Contributions: Experimental design, performed experiments, data acquisition and analysis, prepared and revised manuscript. Co-Authors: Zheng-Jiang Zhu, Julijana Ivanisevic, Adam M. Schuyler, Kush Lalwani, Antonio F. Santidrian, John M. David, Anand Giddabasappa, Amanda J. Roberts, Hernando J. Olivos, Peter J. O’Brien, Lauren Franco, Matthew W. Fields, Liliana P. Paris, Martin Friedlander, Caroline H. Johnson, Adrian A. Epstein, Howard E. Gendelman, Malcolm R. Wood, Brunhilde H. Felding, Gary J. Patti, Mary E. Spilker Contributions: Performed experiments, revised manuscript. Co-Author: Gary Siuzdak Contributions: Experimental design, data analysis, prepared and revised manuscript. 239 Manuscript Information Page Michael Kurczy, Zheng-Jiang Zhu, Julijana Ivanisevic, Adam M. Schuyler, Kush Lalwani, Antonio F. Santidrian, John M. David, Anand Giddabasappa, Amanda J. Roberts, Hernando J. Olivos, Peter J. O’Brien, Lauren Franco, Matthew W. Fields, Liliana P. Paris, Martin Friedlander, Caroline H. Johnson, Adrian A. Epstein, Howard E. Gendelman, Malcolm R. Wood, Brunhilde H. Felding, Gary J. Patti, Mary E. Spilker, and Gary Siuzdak Nature Communications Status of Manuscript: ____ Prepared for submission to a peer-reviewed journal ____ Officially submitted to a peer-review journal ____ Accepted by a peer-reviewed journal __x_ Published in a peer-reviewed journal Nature Publishing Group 6: 5998 (2015) 240 241 242 243 244 245 246 247