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The marine, neutrophilic, and chemolithoautotrophic iron-oxidizing bacteria: insights into the physiology of Zetaproteobacteria and the discovery of novel iron-oxidizing Gammaproteobacteria

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The marine, neutrophilic, and chemolithoautotrophic iron-oxidizing bacteria: insights into the physiology of Zetaproteobacteria and the discovery of novel iron-oxidizing Gammaproteobacteria

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Content THE MARINE, NEUTROPHILIC, AND CHEMOLITHOAUTOTROPHIC IRON-OXIDIZING BACTERIA: INSIGHTS INTO THE PHYSIOLOGY OF ZETAPROTEOBACTERIAAND THE DISCOVERY OF NOVEL IRON-OXIDIZING GAMMAPROTEOBACTERIA. Copyright 2014 by Roman Alfredo Barco A Dissertation Presented to the FACULTY OF THE GRADUATE SCHOOL UNIVERSITY OF SOUTHERN CALIFORNIA In Partial Fulfilhnent of the Requirements for the Degree DOCTOR OF PHILOSOPHY (BIOLOGY) August 2014 Roman Alfredo Barco 11 DEDICATION To my wife, Sandra Marin, and my daughters Sofia and Ali whom I love with all my heart, for always believing in me and for putting up with my hectic graduate student life over these past six years. To my parents Martha Arrieta and Mario Barco for teaching me their work ethic and for giving me infinite encouragement. To my brothers Alejandro, Leonardo, Gustavo and Arnaldo for their support and inspiration. To my parents-in-law, Graciela Marin and Harvey Marin, for lovingly babysitting my daughters when I was on campus and for making the most awesome Colombian meals. To Katrina Edwards, my mentor, for believing in me since day one. To my friends on-campus and off-campus that have kept me sane during all these years. 111 ACKNOWLEDGEMENTS I will forever be grateful for the support that the university has given me in order to fulfill this part of my education and for awarding me a merit fellowship during my first year as a graduate student at USC. I am also grateful to be part of the C-DEBI community and for receiving support in the form of a graduate fellowship for two years. The summer fellowship received from the Wrigley Institute was also very important to me and critical in the new direction that my research took; therefore, it is properly acknowledged here. I would like to also acknowledge the people that during these past six years, and beyond, have inspired me to be a better scientist and pursue excellence: Dr. Katrina Edwards, all members of my quals/dissertation committee (Dr. Kenneth Nealson, Dr. Frank Corsetti, Dr. David Emerson, Dr. Jason Sylvan, Dr. Eric Webb, and Dr. Dave Caron), and current/past members of the Edwards laboratory that I had the pleasure to work with: Dr. Beth Orcutt, Dr. Sarah Bennett, Dr. Jean-Paul Baquiran, Dr. Brandi Reese, Dr. Amanda Haddad, Dr. Esther Singer, Gustavo Ramirez, Mike Lee, Andrew Gross, Juliane Finn, Colleen Hoffman, Jaime Waite, Arkadiy Garber, John Zhong, Ryan Lesniewski, and Greg Horn. I am also very thankful to Dr. Anand Patel and the WMSC staff including Lauren Czarnecki, Kellie Spafford, Trevor Oudin and Gerry Smith for technical advise on the bacterial trap deployments at Fisherman's Cove. TABLE OF CONTENTS DEDICATION 11 ACKNOWLEDGEMENTS 111 LIST OF TABLES v1 LIST OF FIGURES vu ABSTRACT x1 CHAPTER I: The relevance of neutrophilic, chemolithoautotrophic, iron-oxidizing bacteria in the marine environment. 1 INTRODUCTION 1 MAIN GOALS OF DISSERTATION 6 CHAPTER I REFERENCES 7 CHAPTER II: Interactions of proteins with biogenic iron oxyhydroxides and a new culturing technique to increase biomass yields of neutrophilic, iron-oxidizing bacteria. 11 CHAPTER II ABSTRACT 11 INTRODUCTION 11 MATERIALS AND METHODS 14 RESULTS 22 DISCUSSION 34 CHAPTER II REFERENCES 39 CHAPTER III: Proteomic profiling ofneutrophilic, Fe-oxidizing M ariprofundus ferrooxydans indicate an alternative-complex III, cytochrome c 553 and cytochrome cbb 3 oxidases involved in Fe oxidation. 44 CHAPTER III ABSTRACT 44 INTRODUCTION 45 MATERIALS AND METHODS 47 RESULTS AND DISCUSSION 52 CHAPTER III REFERENCES 72 CHAPTER IV: In-situ incubation of iron-sulfide mineral in seawater reveals colonization by iron-oxidizing Thiomicrospira and sheath-forming Zetaproteobacteria. 78 CHAPTER IV ABSTRACT 78 IV INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CHAPTER IV REFERENCES BIBLIOGRAPHY 78 80 88 106 116 121 V VI LIST OF TABLES Table 3-1. The 25 most abundant proteins identified in M. ferrooxydans' 53 proteomic profile. Table 3-2. List of cytochromes that were identified in M. ferrooxydans' 63 proteomic profile sorted by # of peptides identified. Table 4-1. Species richness estimators and diversity indices of 95 environmental samples. vu LIST OF FIGURES Figure 2-1. (A) Left: large batch culture of M. ferrooxydans, strain PV-1 23 after 1 day of growth, compared to a control. Top-Right: formation of biofilm along the wall of culture bottle after 1 day of growth. Bottom- Right: the biofilm dislodges and forms fluffiy flocculent material. (B) SEM image of the mats of PV-1 (bar~ 10 µm). (C) Confocal fluorescent and bright field composite image of the mats of PV-1. Inset: composite image of isolated PV-1 cell and stalk (bar ~ 4 µm). Figure 2-2. (A) Growth curve of M. ferrooxydans, strain PV-1, using a 24 large-batch culture. Error bars indicate standard deviation from the mean (n~3). (B) Overall cell yields obtained by using a method to separate cells from their stalks. Figure 2-3. Enrichment of cells without stalks after the sample goes 24 through the cell-separation method (bar~ 10 µm). Figure 2-4. (A) PV-1 cells in the mat tend to clump together as the iron in 25 the stalks dissolves with 0.2 M oxalic acid at pH 1. The cells stain red when live/dead stained ( dead cells) and stick to a white material that remains undissolved. (B) PV-1 cells treated with 0.2 M oxalic acid at a higher pH of 3. The majority of cells are no longer static (hard to focus image) and stain green when live/dead stained (live cells). Some of the white material still remains in the sample and a minority of cells stick to its surface. Bar~ 10 µm. Figure 2-5. SDS-PAGE gels of osmotic-shock fraction (left) and crude 27 extract (right) stained with Coomassie Blue. Figure 2-6. Left: Number of assigned spectra as a function of isoelectric 27 point of the identified protein (A: osmotic-shock sample; B: crude extract). Right: Number of proteins identified as a function of their isoelectric point (C: osmotic-shock sample; D: crude extract). Figure 2-7. Number of proteins as a function of isoelectric point. Left: 28 Theoretical distribution based on the proteome of PV-1. B. Theoretical distribution based on proteins with molecular weights between 33.6 and 51.1 kDa (common range of MW of the proteins identified between osmotic-shock and crude fractions). Figure 2-8. Venn diagram showing the number of proteins that were 29 uniquely identified in the osmotic-shock fraction (in red) and crude fraction (extracted with NaOH; in green). Shared proteins between fractions are shown in yellow. Venn diagram is based on proteins identified with at least 2 peptides. All proteins identified in each fraction are listed in Supplemental Table S2-l. Figure 2-9. (A) Interaction of cytochrome c with biogenic iron 30 oxyhydroxide. Fractions 1-4 were eluted in 20 mM sodium acetate buffer, pH 5.25. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M NaCl. Error bars indicate ± one standard deviation from the mean. (B) Interaction of BSA with biogenic iron oxyhydroxide. Fractions 1-4 were eluted in 20 mM Tris buffer, pH 8.00. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M Na Cl. Error bars indicate ± one standard deviation from the mean. Figure 2-10. X-ray diffraction pattern of the stalks of PV-1 compared to 31 synthetic 2LF, showing the development of slightly more crystallized iron oxyhydroxide with time. At 12 hours, the sample was growing at log- phase and the biofilm had an off-white/light yellow color. At 6 months, the sample was reddish. At 3 days of growth, the sample had an orange/reddish color which generated a similar spectrum as the 12 hour sample (data not shown). The d-spacing values correspond to 2LF for this synthetic material. Figure 2-11. (A) Interaction of cytochrome c with synthetic iron 33 oxyhydroxide. Fractions 1-4 were eluted in 20 mM sodium acetate buffer, pH 5.25. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M NaCl. Error bars indicate ± one standard deviation from the mean. (B) Interaction of BSA with synthetic iron oxyhydroxide. Fractions 1-4 were eluted in 20 mM Tris buffer, pH 8.00. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M Na Cl. Error bars indicate ± one standard deviation from the mean. Figure 3-1. M. ferrooxydans' proteomic profile sorted by cluster of 55 orthologous genes (COG) functional categories. Figure 3-2. Heme-stained SDS-PAGE gels of soluble and insoluble 56 fractions. (A) Soluble fraction in non-reducing conditions. (B) Soluble fraction in reduced conditions. (C) Membrane fraction in non-reducing conditions. The excised gel areas spanning regions 80 kDa to 120 kDa contain all the proteins encoded by the mob operon. Figure 3-3. Proposed electron transport chain ofM. ferrooxydans based 58 on proteomic analysis. OM, outer membrane; IM, inner membrane. vm Figure 3-4. Mob operon ofFeOB in comparison to the act operon in 58 heterotrophic bacteria (R. marinus) and phototrophic bacteria (C. aurantiacus). Figure 3-5. Amino acid alignment ofmobB and actB proteins. Sequences 60 1-4 are from known neutrophilic FeOB. The conserved cysteine amino acids at the N-terminal ofmobB in FeOB are marked by blue asterisks. Figure 3-6. Maximum-likelihood phylogenetic tree of proteins within the 60 molybdopterin-binding protein superfamily. Numbers on branches indicate bootstrap values based on 1000 replicates. Figure 3-7. Amino acid sequence of cytochrome c 553 gene in M. 63 ferrooxydans M34 (IMG locus tag A37KDRAFT _ 02147). Figure 3-8. Gene neighborhood of area with conserved synteny between 63 Mariprofundusferrooxydans strains M34 and PV-1. (x) indicates the fragmented cytochrome c553 gene in PV-1 (locus tag SPVl _ 07306) that is a homolog to the cytochrome c553 gene in M34 (y; IMG locus tag A37KDRAFT_02147). Figure 3-9. Amino acid alignment comparing cyc2-mfto two iron 65 oxidases: cyc2 and cyt572. Alignment was produced in Geneious by using the CLUSTAL aligner. Signal peptides were removed. Figure 3-10. Phosphate-specific transport (pst) operon of M. ferrooxydans 66 in comparison to other known pst operons in E. coli and B. subtilis. Figure 4-1. Relative abundances of bacterial communities in pyrrhotite 93 traps that were in-situ incubated in the water column (A) or resting on the sediment (B). The relative abundances of the bacterial community in Fisherman's Cove seawater is also shown (C). The abundances are color coded at the class level for Proteobacteria and phylum level for non-Proteobacteria. Taxonomic affiliation is based on full-length 16S rRNA gene sequence matches to sequences in the SILVA database ( similarity cutoff value of 97% ). Figure 4-2. Maximum-likelihood 16S rDNA phylogenetic tree (1000 94 bootstraps) of representative clones recovered from pyrrhotite (OTU defined at 97% similarity cutoff-value). Figure 4-3. Rarefaction curve of species richness (similarity cutoff value 95 of 97% for OTU designation). The bars indicate 95% confidence interval. IX Figure 4-4. A. Thiomicrospira SC-1 in direct contact with bulbous iron 96 oxides. B. Cellular structures connecting the cells to the bulbous iron oxides are typically seen. Figure 4-5. X-ray diffraction pattern of the iron oxide phase that 97 accumulates in a culture of Thiomicrospira SC-1. A reference pattern of lepidocrocite is included for comparison. Figure 4-6. Maximum-likelihood phylogenetic tree based on 16S rRNA 98 gene alignment ( 1000 bootstraps). Figure 4-7. Growth curves ofThiomicrospira SC-1 in artificial seawater 99 in the presence and absence ofreduced Fe. Error bars indicate one standard deviation from the mean. Figure 4-8. Sheath-forming Zetaproteobacteria enriched from a bacterial 100 trap containing pyrrhotite mineral. Figure 4-9. The structure of this particular sheath-Zetaproteobacteria is 101 heat sensitive. The laser from a confocal microscope decreases the stability of the sheath structure and reveals rod-shaped cells encased within it. A few more seconds oflaser exposure lyse the cells, resulting in round-shaped cells. Scale bar ~ 6 µm. Figure 4-10. Maximum-likelihood phylogenetic tree of aligned 16S rRNA 101 genes (1000 bootstraps). The 16S rRNA gene sequences of the sheath-Zetaproteobacteria enriched from pyrrhotite mineral and described in this manuscript are shown in red. Environmental 16S rRNA gene sequences from Zetaproteobacteria (not necessarily sheath-Zetaproteobacteria) obtained from iron-mats that were microscopically observed to exhibit sheath-formation are in blue (Fleming et al., 2013). Figure 4-11. Epifluorescent/phase contrast microscopy merged image of 102 M ariprofundus strain SC-1. Inset bar ~ 6 µm. Larger image bar~ 12 µm. Figure 4-12. Alignment ofmolybdopterin oxidoreductase (mobB) DNA 103 sequences from M. strain SC-1 and M. ferrooxydans PV-1. The resulting amino acid sequences are located below the DNA sequences. Figure 4-13. Venn diagram of abundant proteins identified in mild steel 104 (green), pyrrhotite (red), and surface seawater (blue). The NCBI Proteobacteria database was used for identification of proteins in this figure. X XI ABSTRACT Iron-oxidation performed by bacteria at or near neutral pH is a biological reaction that has been known since the early 1800's. Despite the number of years since its documentation and its global contribution to the biogeochemical cycle of iron, the biological mechanism of bacterial, neutrophilic, iron-oxidation has remained an enigma. M ariprofundus ferrooxydans is the first marine, neutrophilic, chemolithoautotrophic, iron-oxidizing bacteria (FeOB) that has been isolated. Its genome has been sequenced and insights about its physiology were inferred by identifying potential genes in the electron transport chain but no definite mechanism of iron-oxidation was proposed. Here, different approaches involving large-scale culturing and proteomics were combined in order to provide more definite answers about the iron-oxidation mechanism of M. ferrooxydans and FeOB in general. In-situ incubations and proteomics were also combined and applied in the marine environment to study FeOB communities colonizing iron-sulfide minerals. FeOB are historically difficult organisms to work with that usually produce low-biomass. In order to produce enough biomass for proteomic analysis, a large-scale culturing technique was developed for M. ferrooxydans. Proteins released from these cultures were found to interact strongly with the iron mats of M. ferrooxydans and thereby affect protein extractions. Therefore, a method to circumvent protein binding to the mats is described herein. These methods were used to produce a proteomic profile of actively- growing M. ferrooxydans. The resulting proteomic profile identified numerous components of the electron-transport chain, including an abundant periplasmic cytochrome c as well as cbb,-type cytochrome oxidases. As a result, a more specific pathway for electron transport in M. ferrooxydans is described. XU In order to test the developed methods in the marine environment, FeOB communities colonizing iron-sulfide minerals in shallow waters of Catalina Islands, CA were analyzed. In general the results indicate that in-situ enriched iron-sulfides host species-rich communities that are different from the background seawater and similar to inactive hydrothermal vent chimney sulfides. Many of the clones recovered from the iron-sulfide mineral were closely related to deep-sea clones, indicating that this in-situ incubation method is appropriate for the study of microorganisms that are usually seen in deep-sea habitats such as hydrothermal vents and exposed ocean crust. From these in-situ incubations a microorganism previously known only for sulfur oxidation, Thiomicrospira spp., was isolated and shown to be capable of performing Fe oxidation. It is also shown for the first time that the sheath-Zetaproteobacteria can be grown in the laboratory. The data herein presented reveals that environmental proteomics of hard substrates such as iron-sulfide mineral and mild steel can be successfully achieved, opening the door to similar analyses in the ocean floor. CHAPTER I: The relevance of neutrophilic, chemolithoautotrophic, iron-oxidizing bacteria in the marine environment. INTRODUCTION When it was discovered that organisms could be classified phylogenetically by sequencing the gene coding for the small subunit of the ribosomal RNA (16S rRNA) (Woese and Fox, 1977), the field of microbiology dramatically changed from one where microorganisms could not be identified at taxonomic rankings lower than Bacteria level to one where now identification is possible at the species and strain level. It lead to the discovery of one of the domains of life, the Archaea, which microscopically is indistinguishable from Bacteria (Fox et al., 1977). This finding in combination with vital technological advancements, including optimization of the polymerase chain reaction (PCR), lead microbiologists to start studying microbial diversity at an unprecedented level in all possible habitats, from the human body (Human Microbiome Project Consortium, 2012) to soil (Mendes et al., 2011) to river streams (Lyautey et al., 2005) to the oceans (Venter et al., 2004). Even in the most extreme of the cases, these microorganisms were always present and abundant. 1 It is not surprising then that iron (Fe), which is one of the most abundant metals on Earth and the most abundant redox metal in the biosphere, is used by these microorganisms (i.e. as cofactors for enzymes). However, it was not until relatively recently that it was 2 determined that bacteria can have a much more profound interaction with metals through respiration. The discovery by Charles Myers and Ken Nealson (1988), only 25 years ago, that Shewanella oneidensis can respire manganese (Mn) and Fe led to the birth of a field within microbiology studying direct interactions between microorganisms and metals. The increased interest and the ease of growing these organisms in the laboratory resulted in the purification of proteins that are associated with the biological reduction of Fe(III), specifically c-type cytochromes (Pitts et al, 2003). With time, Shewanella and bacteria in the genus Geobacter became well-characterized model organisms for the study of dissimilatory Fe reduction (Childers et al., 2002). Having established that Bacteria and Archaea (Vargas et al., 1998; Tor et al., 2001) have the ability to reduce metals such as Fe, there was a revival of interest in studying microbial Fe oxidation. The biological Fe (II) oxidation reaction is often coupled to the reduction of 0 2 and used by microbial populations of autotrophs to gain chemical energy in the form of ATP to eventually fix carbon. Since, the kinetics of Fe oxidation indicate that abiotic reaction is relatively fast (i.e. minutes) in oxygenic seawater at pH~ 8.1, a microbial population of Fe-oxidizers would not have a chance to oxidize much of it. Therefore, in order for FeOB to fully benefit from Fe(II) oxidation, the half-life ofFe(II) needs to increase to hours. For this to happen, their immediate environment has to be, at least, suboxic and/or slightly acidic (Ferris 2005; Nealson 1997). Suboxic, slightly acidic environments that will retard abiotic Fe (II) oxidation are known to exist in the ocean floor, more specifically in hydrothermal vent systems where suboxic levels ofO 2 (< 10 % ambient), and high levels of CO 2 or H 2 S are typical (Edwards et al. 2004; Emerson and Moyer 1997). 3 The members of the microbial communities of Fe-oxidizers are varied, ranging from acidophiles (acid-loving microorganisms) (i.e., Thiobacillus ferrooxidans) to neutrophiles (i.e. Gallionella ferruginea and M ariprofundus ferrooxidans) to thermophiles (i.e. Ferroglobus placidus) to phototrophs (Rhodopseudomonas palustris). To this day only a few marine microorganisms have been isolated directly from Fe-oxides from the ocean's floor: M. ferrooxydans of the zeta (t;)-class of Proteobacteria (Emerson and Moyer, 2002; Emerson et al. 2007) and Fe-oxidizing representatives from a and y Proteobacteria (i.e. related to heterotrophic Hyphomonas and Marinobacter, respectively (Edwards et al. 2003)). These Fe-oxidizers are autotrophic and microaerophilic; although, in the case of M ariprofundus, it appears to be an obligate chemolithoautotroph. One interesting fact about M. ferrooxidans is that it is the first isolate from the marine environment that has been shown to produce the microfilaments that are typically seen in Fe deposits in the ocean floor - in modern and geologically ancient deposits (Alt, 1988; Karl, 1989). These organic extracellular structures are extruded by the cell in conjunction with Fe oxyhydroxides and are very similar to the ones produced by G. ferruginosa in freshwater habitats (Emerson et al., 2010). Recently, it was reported that some Zetaproteobacteria can also form sheath structures similar to the ones produced by Leptothrix spp. (Fleming et al., 2013). Fe-oxidizing bacteria with these types of morphologies (i.e. twisted stalks and sheaths) are thought to contribute> 50% of the Fe deposits in hydrothermal vent systems such as Lo'ihi Seamount in the Central Pacific Ocean (Emerson and Moyer 2002; Rassa et al., 2009). These morphologies of Fe oxides have also been recently documented in several other hydrothermally active marine sites around the world, including Eolo Seamount in the Mediterranean Sea (Dekov et al., 2007) and Axial Volcano in the North-Eastern Pacific Ocean (Kennedy et al. 2004). Overall, the Zetaproteobacteria has been identified in deep-sea habitats associated with hydrothermal activity, including spreading centers and seamounts. However, there has been a recent report indicating that the Zetaproteobacteria can also be in-situ enriched in shallow marine waters (McBeth et al., 2011). McAllister et al. (2011) analyzed the biogeography of Zetaproteobacteria based on these limited available reports and indicated their cosmopolitan distribution in the world's oceans. 4 At the protein level, most of the research efforts so far have been done on acidophilic Fe oxidizers. Community proteomics of natural iron-oxidizing microbial mats from an acid mine drainage site at Richmond Mine in Iron Mountain, CA indicates that c-type cytochromes are abundant (Ram et al., 2005). A cytochrome is a type of protein that catalyzes redox reactions and forms part of the electron transport chain. In general, the common feature in cytochromes is the heme prosthetic group with an iron atom at its center. In particular, c-type cytochromes possess a covalently-bound heme prosthetic group. Furthermore, the cytochromes that have been isolated from biofilms ofFe oxidizing bacteria at Richmond Mine display Fe-oxidizing activity via spectrophotometric analysis (Singer et al., 2008; Jeans et al., 2008). These cytochromes were associated with acidophilic, chemolithoautotrophic Leptospirillum spp. that are known to leach sulfide ore. Alternatively, a study of an Fe-oxidizing thermophile originally isolated from another acid mine drainage site in Japan, determined that an a type cytochrome, containing a heme-group, is an Fe oxidase (Takai et al., 2001). Acidithiobacillusferrooxidans, another acidophilic Fe-oxidizer, was also determined to have two cytochromes involved in the electron transfer system associated with oxidation of Fe (II) (Yamanaka et al., 1995). 5 The only reports on iron-oxidizing proteins from neutrophilic FeOB appeared in the early 1990s and were based on the heterotrophic FeOB Leptothrix discophora. The two articles reported on partial evidence that Mn- and Fe-oxidation in spent medium of L. discophora were enzymatically catalyzed ( de Vrind-de Jong et al., 1990; Corstjens et al., 1992). In this particular case, L. discophora had lost its ability to form sheaths. Nonetheless, a colony-forming Leptothrix mutant that had retained Mn- and Fe-oxidation activities was studied. The Mn and Fe oxidases that were identified by the use of in-gel activity assays were never actually sequenced and, unfortunately, studies on these proteins were not continued (Corstjens, personal communication). The genome of the marine Fe-oxidizer M. ferrooxydans, strain PV-1 was sequenced recently, and potential functions of the different complexes in the electron transport chain are highlighted (Singer et al., 2011). Results from the annotation of M.ferrooxydans' genome indicated that it contains 32 c-type cytochromes, 5 b-type cytochromes, and 5 gene clusters for Fe uptake and transport. There are also 2 terminal oxidases: cbb 3 -type cytochrome oxidase and cytochrome d oxidase, both of which are associated with organisms living in microaerophilic conditions due to their high affinity to oxygen as the final electron acceptor. No Fe-oxidase has been definitively identified. The high number of c-type cytochrome genes present in the genome, along with results from other studies, makes this type of proteins clear candidates for the biological function of Fe-oxidation. But, experimental evidence is needed in order to narrow down the long list of cytochromes that can potentially be involved in Fe oxidation. The genomes of neutrophilic, freshwater FeOB Gallionella and Sideroxydans were also recently sequenced and annotated (Emerson et al., 2013). These genomes can also help gain further insights into the function of certain proteins in M. ferrooxydans. MAIN GOALS OF DISSERTATION 6 The main goals of this dissertation are: (1) to develop methods to study the physiology of neutrophilic FeOB, specifically Zetaproteobacteria, at the protein level (2) to use these methods to gain knowledge about the specific components of the electron transport chain of Zetaproteobacteria (3) to use these methods on environmental samples in conjunction with microbial ecology analysis and enrichment culturing in order to generate hypotheses about ecological functions of deep-sea microorganisms, especially marine neutrophilic chemolithoautotrophic FeOB. 7 CHAPTER I REFERENCES Alt, J.C. (1988). Hydrothermal oxide and nontronite deposits on seamounts in the eastern Pacific. Mar. Geo/. 81:227-239. Childers, S.E., Ciufo, S., Lovley, D.R. (2002). Geobacter metallireducens accesses insoluble Fe(III) oxide by chemotaxis. Nature. 416(6882):767-769. Corstjens, P.L., de Vrind, J.P., Westbroek, P., de Vrind-de Jong, E.W. (1992). 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Characterization of the Shewanella oneidensis MR-1 decaheme cytochrome MtrA: expression in Escherichia coli confers the ability to reduce soluble Fe(III) chelates. J. Biol. Chem. 278(30):27758-27765. Ram, R.J., VerBerkmoes, N.C., Thelen, M.P., Tyson, G.W., Baker, B.J., Blake, R.C., Shah, M., Hettich, R.L., Banfield, J.F. (2005). Community proteomics of a natural microbial biofilm. Science. 308:1915-1919. Rassa, A.C., McAllister, M.S., Safran, S.A., Moyer, C.L. (2009). Zeta-Proteobacteria dominate the colonization and formation of microbial mats in low-temperature hydrothermal vents at Lo'ihi Seamount, Hawaii. Gemicrobiol. J. 26:623-638. Singer, S.W., Chan, C.S., Zemla, A., VerBerkmoes, N.C., Hwang, M., Hetlich, R.L., Banfield, J.F., Thelen, M. (2008). Characterization of cytochrome 579, an unusual cytochrome isolated from an iron-oxidizing microbial community. Appl. Environ. Microbial. 74(14):4454-4462. Singer, E., Emerson, D., Webb, E.A., Barco, R.A., Kuenen, J.G., Nelson, W.C., Chan, C.S., Comolli, L.R., Ferriera, S., Johnson, J., Heidelberg, J.F., Edwards, K.J. (2011). M ariprofundus ferrooxydans PV-1 the first genome of a marine Fe(II) oxidizing Zetaproteobacterium. PLoS One. 6(9):e25386. Takai, M., Kamimura, K. and Sugio, T. (2001). Anew iron oxidase from a moderately thermophilic iron oxidizing bacterium strain TI-1. Eur. J. Biochem. 268(6):1653-1658. Tor, J.M., Kashefi, K., and Lovley, D.R. (2001). Acetate oxidation coupled to Fe(III) reduction in hyperthermophilic microorganisms. Appl. Environ. Microbial. 67: 1363- 1365. Vargas, M., Kashefi, K., Blunt-Harris, E., and Lovley, D.R. (1998). Microbiological evidence for Fe(III) reduction on early Earth. Nature. 395:65-67. 10 Venter, J.C., Remington, K., Heidelberg, J.F., Halpern, A.L., Rusch, D., Eisen, J.A., Wu, D., Paulsen, I., Nelson, K.E., Nelson, W., Fouts, D.E., Levy, S., Knap, A.H., Lomas, M.W., Nealson, K., White, 0., Peterson, J., Hoffman, J., Parsons, R., Baden-Tillson, H., Rogers, Y.H., Smith, H.O. (2004). Environmental genome shotgun sequencing of the Sargasso Sea. Science. 304(5667):66-74. Woese, C.R. and Fox, G.E. (1977). Phylogenetic structure of the prokaryotic domain: the primary kingdoms. PNAS. 74(11):5088-5090. Yamanaka, T. and Fukumori, Y. (1995). Molecular aspects of the electron transfer system which participates in the oxidation of ferrous ion by Thiobacillus ferrooxidans. FEMS Microbial. Rev. 17(4):401-413. CHAPTER II: Interactions of proteins with biogenic iron oxyhydroxides and a new culturing technique to increase biomass yields of neutrophilic, iron-oxidizing bacteria. ABSTRACT 11 Neutrophilic, bacterial iron-oxidation remains one of the least understood energy generating biological reactions to date. One of the reasons it remains under-studied is because there are inherent problems with working with iron-oxidizing bacteria (FeOB), including low biomass yields and interference from the iron oxides in the samples. In an effort to circumvent the problem of low biomass, a new large batch culturing technique was developed. Protein interactions with biogenic iron oxides were investigated confirming that such interactions are strong. Therefore, a protein extraction method is described to minimize binding of proteins to biogenic iron oxides. The combination of these two methods results in protein yields that are appropriate for activity assays in gels and for proteomic profiling. INTRODUCTION Neutrophilic microbial iron oxidation is a biological reaction that has been known since the early 19th century (Klitzing, 1833; Pringsheim, 1948). Yet it remains poorly understood when compared with other thoroughly investigated reactions such as sulfur oxidation, nitrogen fixation or iron reduction due to an array of difficulties including low biomass, lack of pure cultures, and interference of iron oxides with sample processing. The majority of the neutrophilic, iron-oxidizing bacteria (FeOB) that have been isolated so far are obligate iron oxidizers; therefore, they cannot grow on media enriched in organic carbon as other groups of bacteria can (i.e. the iron reducing bacteria). These historical difficulties have contributed to the dearth of data on the basic physiology of neutrophilic FeOB, despite important advances in the understanding of acidophilic and phototrophic FeOB. 12 There are several methods available to grow neutrophilic FeOB, as recently reviewed by Emerson and Floyd (2005). These methods include the use of gradient tubes, gradient plates, solid media and liquid media. From these methods, the most useful in isolating novel organisms from the environment has been the gradient tube method. This method consists of using small glass tubes that have a FeS plug at the bottom and a defined freshwater or marine medium stabilized with agarose overlaying the plug. Since the bottom FeS plug diffuses Fe(II) upwards and the 0, (g) diffuses downwards into the bottom of the tube, the two opposing gradients meet forming a microaerobic region where FeOB can grow and produce a sharp band of growth. The reddish band of growth containing oxidized iron and cells can then be carefully removed, serially diluted, and subcultured with the whole process repeated numerous times to obtain a pure FeOB culture. This method has been crucial in the isolation of many FeOB including Sideroxydans ES-1, Galli one/la ES-2, Mari profundus ferrooxydans, and recently Ferriphaselus amnicola (Emerson and Moyer, 1997; Emerson and Moyer, 2002; Kato et 13 al., 2013). But one of the problems is with this method is that it does not yield enough biomass for protein research, with total number of cells ranging 10 6 -10 7 per tube (Emerson and Moyer, 2002). A minimum of I 0 9 cells would be required to obtain approximately 100-150 µg of protein (Bremer and Dennis, 1996), which is enough protein yield for proteomic analysis. However, if activity assays (i.e. in-gel or based on spectroscopy) are planned, a minimum of 10 10 cells should be planned. This would indicate that if the gradient tube method is employed, there would have to be thousands of tubes inoculated per experiment; this is clearly an impractical approach. Alternatively, the gradient plate method (i.e. with and without agarose) provides for more volume (15 mLs) and larger total cell numbers, but the number of plates needed per experiment would still be in the hundreds. Solid media is often not an option to grow autotrophic FeOB, since they do not form classical bacterial colonies. The liquid medium method presented in Emerson and Floyd (2005) offers more volume than the gradient tube and gradient plate methods, but it is still relatively low in volume ( 40 mLs ). An attractive factor in using the liquid medium is that it is more realistic to the true environment than the semi-solid agarose medium, a factor that is important ifwe want to study the physiology of FeOB in a state that is comparable to their natural environments. In addition to the above challenges, there is an inherent problem to studying FeOB, which is the presence of iron oxyhydroxides (i.e. both biogenic and non-biogenic) in samples. This problem is known for the extraction of DNA in soil and rock samples, for example, where special commercial soil kits are recommended for consistent and appropriate 14 yields of DNA for Polymerase Chain Reaction (PCR) experiments (Wang and Edwards, 2009). In the case of protein analysis, such amplification is not possible. However, there have been several studies that have shown that consistent protein extraction from sediment, soil, rock and wastewater can be attainable (Ogunseitan, 1993; Benndorf et al., 2007; Benndorf et al., 2009; Keiblinger et al., 2012). Nonetheless, protein extraction remains the biggest bottleneck in environmental - or laboratory based - proteomics due to the technical challenges ranging mainly from low yields, complexity of matrix, and humic acid interference (Keller and Hettich, 2009; VerBerkmoes et al., 2009; Bastida et al., 2009). In this manuscript, a way to scale up culturing of the FeOB Mariprofundus ferrooxydans in liquid medium is presented for purposes of obtaining high-yields of biomass for subsequent proteomic experiments. We also present a method to separate cells from the stalks of biogenic iron oxides that can be useful for analysis of isolated cells. Both of these methods were shared and shortly described in Saini and Chan (2013) but we expand on the details and present new findings. Strong interactions of proteins with both synthesized and biogenic iron oxyhydroxide are also shown and proven to affect protein extractions. A method to circumvent this problem is presented. MATERIALS AND METHODS Large batch cultures: To generate enough biomass needed for proteomic analysis, a new 15 large-scale culturing method for M. ferrooxydans is described. M. ferrooxydans, strain PV-1 (hereinafter PV-1 ), is grown from a stock culture provided by David Emerson (Bigelow Laboratory for Ocean Sciences, East Boothbay, ME, USA). The liquid medium is adopted from Emerson and Floyd (2005) with modifications appropriate for scaling up to 100 mL and 800 mL cultures. The large batch cultures are grown in 1 L autoclavable polycarbonate bottles (VWR, Visalia, CA) each with 784 mL of medium. Immediately after autoclaving, the medium is sparged with N 2 (g) for 30 minutes. The bottle is then sealed with an autoclaved rubber stopper #6 coated in a thin film of silica gel and capped with an open-top cap. After the bottles cool to room temperature, 15.2 mLofsterile sodium bicarbonate solution and filter-sterilized CO 2 (g) are added to adjust pH to 6.2-6.5 (1.5 minutes at 15 psi; must check empirically). This is followed by addition of800 µL of vitamin cocktail (ATCC, Manassas, VA), 32 mL of filter-sterilized air and 3.2 mL of filter-sterilized 100 mM FeCizsolution. The culture vessel is inoculated with 40 mL of log-phase PV-1 ( approximately 10 7 cells grown in 100 mL batch culture) and incubated in the dark, horizontally at room temperature without agitation. Iron is added every 24 hours. The headspace is gas-exchanged every 24 hours with filter-sterilized N 2 :CO 2 (70:30 v/v) (g) and 32 mL of filtered-air. Active cultures in small bottles are maintained in log-phase by subculturing every 48 hours until the start of the experiments. Cell counts to determine cell density for these experiments were performed as described in Emerson and Moyer (2002). Briefly, duplicate samples were fixed with paraformaldehyde solution (0.8% final concentration), stored at 4 'C for 1 hour and then frozen at -20 'C until counted. Slides printed with 4 mm diameter circles (Electron Microscopy Sciences, Fort 16 Washington, PA) were coated with 1 % agarose solution and allowed to cool. A sample volume of 4 µL (2 µL of resuspended culture sample mixed with 2 µL of 1 mM propidium iodide solution (Life Technologies, Grand Island, NY)) was loaded within the boundaries of the circle and allowed to dry in the dark. Fifty fields per circle were counted at 1 OOX magnification using an epifluorescent Axiostar Plus microscope equipped with an HBO 50 mercury lamp and Cy3 filter for green light excitation. Alternatively, samples were imaged on a TCS SPE confocal microscope (Leica Mycrosystems, Buffalo Grove, IL) using a 488 nm solid-state laser for excitation and emission wavelenghts of 495-550 nm for SYTO 9 dye. Protein interaction with iron oxides: For each interaction experiment, the mats oflog phase PV-1 were directly harvested from 2 small batch cultures by pipetting and centrifuged at 10,000 x g for 5 minutes. The pelleted mats were washed with 50 volumes of milli-Q water to 1 volume of mats 3 times. Each washing step was followed by centrifugation at 10,000 x g for 5 minutes. The mats were then resuspended in either 50 mL of 20 mM sodium acetate buffer, pH 5.25 (for experiments with cytochrome c), or 20 mM Tris Base buffer, pH 8.0 (for experiments with bovine serum albumin, BSA) and loaded into sealed Econo-Pac® chromatography columns (1.5 x 12 cm) (Bio-Rad, Hercules, CA) in a minimal volume of buffer. The final volume of packed mats of PV-1 was 0. 8 mL. The mats were then spiked with 50 µg of either cytochrome c from horse's heart (1 mL of 50 µg/mL fresh stock in 20 mM sodium acetate buffer, pH 5.25)(Sigma Aldrich, St. Louis, MO) or BSA (1 mL of 50 µg/mL fresh stock in 20 mM Tris base 17 buffer, pH 8.0)(Bio-Rad). The sample was allowed to incubate for 15 minutes at room temperature with resuspension every 5 minutes. After this incubation period, 5 mL fractions are collected while sodium acetate buffer (for cytochrome c experiment) or Tris buffer (for BSA experiment) is continually added to the column. Starting with the 5th fraction, either 0.1 N NaOH solution (for both cytochrome c or BSA) or 1 M NaCl/20 mM sodium acetate buffer, pH 5.25 (for cytochrome c experiment) or 1 M NaCl/20 mM Tris base buffer, pH 8.0 (for BSA experiment) was added to the column. A total of 14 fractions were collected per experiment, which are performed in duplicates. Protein concentrations were measured by the Bradford Assay (Bio-Rad). Preparation of 2-line ferrihydrite: The method of Schwertmann and Cornell (2000) was used to produce 2-line ferrihydrite (2LF). Forty g ofFe(NO,),·9H,O was dissolved in 500 mL ofmilli-Q water. To this solution, 330 mL of lM KOH was slowly added to bring the pH to a value between 7-8. The mixture was then aliquoted into 50 mL tubes and washed three times in milli-Q water by centrifuging at 10,000 x g for 10 minutes at 4 'C. The pellets were frozen at -20 °C for 1 day and oven dried for another day at 50 °C. This freezing/drying procedure was repeated 3 times. The dried product was ground to particle size > 180 µm and <355 µm. Before the start of the experiments, 1 gram of synthetic 2LF was washed 5 additional times with 50 mL of milli-Q water. The synthetic 2LF was then resuspended in 50 mL of buffer (20 mM sodium acetate, pH 5.25 for cytochrome c experiments or 20 mM Tris Base, pH 8.0 for BSA experiments) and loaded into a sealed Econo-Pac® chromatography column in a minimal volume of buffer to cover 18 it. The synthetic 2LF had a volume of 0. 8 cm 3 in the column. Samples were spiked as indicated in the last section and were incubated for 15 minutes at room temperature with resuspension of materials every 5 minutes. The mineral was characterized at the Los Angeles National History Museum by X-ray diffraction (XRD) to confirm its identity. Data were recorded using a R-Axis Rapid II (Rigaku, The Woodlands, TX) curved imaging plate microdiffractometer with monochromatized MoKa radiation. Observed d spacings and intensities were derived by profile fitting using JADE 2010 software (Materials Data Inc., Livermore, CA). Protein extraction - osmotic-shock fraction: A total of 8 L oflate-log phase PV-1 culture was harvested by filtration on 0.22 µm mesh black Whatman-Nucleopore polycarbonate filters (GE Healthcare Life Sciences, Piscataway, NJ) with 10 µm support TCTP filter (EMD Millipore, Billerica, MA). PV-1 mats were treated for osmotic-shock by immersing the filters in 30 mL of 40 mM Tris-Base pH 8.5/20% sucrose solution followed by addition of 60 µL of0.5 M EDTA, pH 8.0 and stirring slowly at room temperature for 10 minutes (Neu and Heppel, 1965). The mixture was centrifuged at 14,000 x g for 10 minutes at 4 'C and the supernatant discarded. The pellet was resuspended in 30 mL of ice cold, sterile 5 mM MgClz and incubated on ice for 10 minutes with slow stirring for osmotic shock (Ausubel et al., 1989). The mixture was centrifuged again at 14,000 x g for 10 minutes. The supernatant was saved and stored at -20 'C. The osmotic-shock fraction was quantitated by using the Quick Start™ Bradford Assay (Bio-Rad), according to manufacturer's instructions using an UV-1601 spectrophotometer (Shimadzu Scientific Instruments, Carlsbad, CA). Protein extraction - crude extract: A total of 8 L oflate-log phase PV-1 culture was harvested as described above. The filters were incubated in 40 mL of 0.1 N NaOH 19 solution on ice for 15 minutes with vigorous vortexing for 30 seconds every 3 minutes to lyse cells and release proteins. The mixture was centrifuged at 14,000 x g for 10 minutes at 4 °C to pellet cellular debris and iron oxides. The clarified supernatant was saved and stored at -20 °C. The resulting pellet was immersed in another 40 mL of 0.1 N NaOH solution and processed as described above to maximize protein yields. The combined, clarified supernatants were concentrated and buffer-exchanged with 40 mM Tris-Base, pH 8.5 by using Macrosep 3K concentrators (Pall, Ann Arbor, MI) with a 3 kDa molecular weight cutoff (MWCO) membrane according to manufacturer's instructions. The above procedure resulted in a concentrated orange crude extract containing both insoluble and soluble proteins, which was then frozen at -20 °C until further analysis. Cell separation: A total of2.4 L (0.8 L triplicates) of log phase cultures were harvested as described above. The filters were immersed in total volume of 250 mL of cold, filter sterilized 0.2 M oxalic acid, pH 3.0 for two hours with mixing on a shaker table. The oxalic acid partially dissolves the stalks of iron oxides so that the cells are released. The mixture was centrifuged at 14,000 x g for 10 minutes at 4 'C to pellet the resulting white paste material and the cells. The supernatant was discarded and the pellet is resuspended in 8 mL of sterile artificial seawater medium (ASW). The resuspended material ( 4 mL x 20 2 tubes) was mixed with filter-sterilized 0.001 % resazurin (0.2 mL) and carefully layered over a filter-sterilized Nycodez® solution with 1.3 g/mL density (12 mL) (Axis-Shield, Oslo, Norway) in a sterile, polypropylene 50 mL conical tube. Resazurin does not readily mix with Nycodenz and helps visualize the top layer containing the cells. The tube was centrifuged at 10,000 rpm for 50 minutes at 4 °C to separate cells from the white paste/stalks. Being careful not to disturb the two layers, the top pink layer and some of the Nycodenz were transferred into a new tube. The pink solution was diluted 1 :2 with ASW to dilute any remaining Nycodenz and centrifuged at 14,000 x g for 10 minutes at 4 'C. The resulting pellet, containing cells, was resuspended in a minimal amount of ASW or buffer of choice and processed for cell counting and/or protein extraction. Sodium dodecyl sulfate - polyacrylamide gel electrophoresis (SDS-PAGE): Proteins were separated on gels at 90 V according to standard protocols (Laemmli, 1970). Soluble (n ~ 2) and solubilized membrane fractions (n ~ 2) were run on 12 % TGX™ polyacrylamide gels (Bio-Rad) under reduced and non-reduced conditions (i.e. no dithiothreitol (DTT) and no heating). Gels were stained with Bio-Safe Coomassie Stain (Bio-Rad) for 1 hour, destained with HPLC-grade water for 2 hours and visualized with a Gel Doc™ XR+ imaging system (Bio-Rad) equipped with a white light transilluminator. Images were analyzed with Image Lab™ software (Bio-Rad). Slices were excised as close as possible to the band of interest with a clean sterile scalpel, stored in HPLC-grade water at 4 °C and immediately submitted for proteomic analysis. 21 Proteomic analysis: Excised protein samples were submitted to the Children's Hospital of Los Angeles - Proteomic Core Facility for trypsin-digestion and LC-MS/MS on a Thermo LTQ-Orbitrap XL mass spectrometer (San Jose, CA) equipped with an Eksigent (Dublin, CA) Nanoliquid Chromatography 1-D plus system. The resulting MS/MS spectra were searched against the generated proteome of PV-1 in the Uniprot database ( accession numbers: NZ_ AATS0l 000001-AATS0 1000032; Singer et al., 2011) using the Proteome Discoverer SEQUEST Daemon search engine. Protein probabilities were assigned by the Protein Prophet algorithm (Nesvizhskii et al., 2003). The criteria for having a protein identified at> 95% probability is that there needs to be a minimum of two unique peptides matching to it. Each peptide was established at> 95% probability(> 5% probability that it is a false positive match to the spectra) by the Scaffold Local FDR algorithm. Datasets were normalized based on the number of assigned spectra (i.e. number of peptides identified). Theoretical isoelectric points and molecular weights were generated by the ExPASy computational tool Compute pI/Mw by using Uniprot Knowledge base accession numbers corresponding to the reference proteome of PV-1 (Gasteiger et al., 2005). Isoelectric point bias (b) were calculated as defined by Kiraga et al. (2007); b ~ 100 (Nbasic - Nacidic)/(Nb,<c + Nacidic), where Naci<lic and Nb,<c represent the numbers of acidic and basic proteins, respectively. Statistical analysis was performed by using JMP®, Version 11 (SAS Institute Inc., Cary, NC). The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium (Vizcaino et al., 2014) via the PRIDE partner repository with the dataset identifier PXD000937. 22 SEM: The sample was fixed in paraformaldehyde solution (0.8% final concentration), incubated at 4 'C for 1 hour and stored at -20 'C afterwards. It was dehydrated sequentially in 25%, 50%, 75%, 90% and 100% molecular grade ethanol with 10 minutes per dehydration step. Then it was critically-point-dried and gold coated. Images were taken with the scanning electron microscope JSM-7001F-LV (JEOL, Peabody, MA) at the Center for Electron Microscopy and Microanalysis at USC. RESULTS Growth in a large-batch culture was successfully achieved with massive production of biofilm during the first 24 hours of culture (Figure 2-la). As the biofilm is disrupted, it forms flocculent material that tends to clump together and sink. The biofilm consists of cells and networks of stalks, with mostly biological products of iron oxidation seen as opposed to amorphous products of chemical iron oxidation (Figure 2-lb, 2-lc). Figure 2- 2a shows a representative growth curve of PV-1 growing in a large batch culture. The doubling time is 10 hours. The stationary phase is reached at the third day of growth. With an initial concentration of 1.6 x 10 5 cells/mL, the cell density plateaus at an approximate average of 8.0 x 10 6 cells/mL. A total yield of 6.4 x 10 9 cells per single large batch culture is achieved. Figure 2-2b shows the number of cells that were released from the iron-oxide/stalks matrix following incubation on oxalic acid and centrifugation on a density gradient medium. The number ofreleased cells was approximately 58 % of the total number of cells in the large batch bottle (total yield of2.8 x 10' cells). This fraction 23 was more abundant than the fraction of cells that were still stuck in the iron-oxide/stalks matrix, which corresponded to 3 8 % of the total population. There was a small fraction of cells(< 5% of total) that were present in-between the top and bottom layers. Figure 2-3 shows the enrichment of released cells and the absence of stalks. Incubation with oxalic acid at a pH < 2, dissolves the networks of stalks but makes the cells clump together in a white paste material (Figure 2-4a, b ). Incubation with oxalic acid at pH 3, only partially dissolves the network of stalks so that the cells are released without clumping. Figure 2-1. (A) Left: large batch culture of M ferrooxydans, strain PV-1 after I day of growth, compared to a control. Top-Right: formation of biofilm along the wall of culture bottle after I day of growth. Bottom Right: the biofilm dislodges and forms fluffly flocculent material. (B) SEM image of the mats of PV-1 (bar = 10 µm). (C) Confocal fluorescent and bright field composite image of the mats of PV-1 (bar= 25 µm). Inset: composite image of isolated PV-1 cell and stalk (bar= 4 µm). l.00E+07 --- --'!'- - ::3' ~ I C 0 g ~ l.00E+06 - § u 'ii u l.00 E+0S A 20 40 60 80 100 120 140 Time (hours) l.00E+ IO , ~ ~ ,n ~ 1.00E+09 C. - -"' ] 0 ~ l.00E+ 08 ,n E - ~ C ] ,:; l.00 E+07 B r eleased cells cells with iron oxides between-layers Figure 2-2. (A) Growth curve of M ferrooxydans, strain PV-1, using a large-batch culture. Error bars indicate standard deviation from the mean (n=3). (B) Overall cell yields obtained by using a method to separate cells from their stalks. 24 Figure 2-3. Enrichment of cells without stalks after the sample goes through the cell-separation method (bar = 10 µm). 25 Figure 2-4. (A) PV-1 cells in the mat tend to clump together as the iron in the stalks dissolves with 0.2 M oxalic acid at pH 1. The cells stain red when live/dead stained (dead cells) and stick to a white material that remains undissolved. (B) PV-1 cells treated with 0.2 M oxalic acid at a higher pH of 3. The majority of cells are no longer static (hard to focus image) and stain green when live/dead stained (live cells). Some of the white material still remains in the sample and a minority of cells stick to its surface. Bar= IO µm. Extraction of proteins following physical disruption of the PV-1 mats (i.e. bead-beating or sonication) yielded either zero bands or only a few faint bands on a coomassie-stained polyacrylamide gel after SDS-PAGE, even when cocktails of various detergents were used (results not shown). Yields were< 1 µg of protein per L of culture. Similar results were obtained when PV-1 mats were partially dissolved in oxalic acid and the cells were sonicated in the presence of a white material that does not dissolve. Alternatively, a treatment for osmotic-shock that did not disrupt the iron oxides, resulted in more protein extracted ( ca. 3 µg of protein per L of culture) compared to the other disruptive methods including bead-beating, sonication, and acidification. After this sample is run via SDS PAGE, one of the strongest bands that stains with the coomassie stain was a band around 40 kDa (Figure 2-5). When a duplicate gel was stained with the heme-stain it showed a weak positive result in an area that matches the molecular mass of 40 kDa. Following these results, a third gel was run via SDS-PAGE and the 40 kDa band was excised and submitted for proteomic analysis. An analysis of the identified proteins indicated that 26 most of them were cytoplasmic; thus, PV-1 cells were lysed by the osmotic-shock treatment (Supplemental Table S2-l). Remarkably, most of the proteins were acidic or negatively-charged (97.5% of total proteins). Only 1 of 41 identified proteins were basic. In order to determine that the absence of basic proteins was due to a pH effect, the pH of the extraction solution was increased to pH 13 by using 0.1 N NaOH and without physically disrupting the mats. This approach yielded > 50 µg of protein per L of culture. The sample was run via SDS-PAGE and the 40 kDa band was excised and submitted for proteomic analysis (Figure 2-5). The results indicate that the protein extraction with NaOH circumvented the bias towards acidic proteins seen in the osmotic-shock treatment (Supplemental Table S2-l). From 121 proteins identified, 20 were basic (16.5% of total proteins) and with theoretical subcellular localizations including the outer membrane, periplasm and cytoplasm. Figure 2-6 shows that more basic and acidic proteins were identified following extraction in 0.1 N NaOH (weighted average pl~ 5.83; weighted a~ 1.42), while mainly acidic proteins were identified following the osmotic-shock treatment (weighted average pl ~ 5.19; weighted a~ 0.61 ). A non-parametric Kolmogorov Smirnov two-sample test found that the distributions are significantly different (p > 0.0001). 85 50- 40 30- 25- 20- 15 10 Figure 2-5. SDS-PAGE gels of osmotic-shock fraction (left) and crude extract (right) stained with Coomassie Blue. Molecular masses (in kDa) are indicated on the left. ~ l al .§l ~ A 350 300 250 200 150 100 50 0 60 50 ~ 40 ! -g 30 §, ] 20 10 0 3 B 3 ♦ ♦ ♦ ♦ ' ... . .. ....... · ♦ .... ♦ ♦ .. 1 0 I I 10 II pl ,2 C O 3 0 12 D 3 10 I I 10 I I pl 27 12 12 Figure 2-6. Left: Number of assigned spectra as a function of isoelectric point of the identified protein (A: osmotic-shock sample; B: crude extract). Right: Number of proteins identified as a function of their isoelectric point (C: osmotic-shock sample; D: crude extract). The theoretical distribution of the pl values of the proteins generated from the proteome of PV-1 follows a bimodal pattern (Figure 2-7) that is seen in all organisms so far 28 investigated, including prokaryotes (Schwartz et al., 2001; Knight et al., 2004; Kiraga et al., 2007). A bimodal distribution is also seen if the focus is on a subset of proteins with molecular weights similar to the proteins identified in the bands excised for proteomic analyses. The pl bias of the proteome of PV-1 is acidic (b = -36.0), even when only the subset of proteins is taken into account (b = -45.0). The pl bias calculated from the proteins extracted with 0.1 N NaOH remains acidic (b = -68.6) while the pl bias calculated from the proteins extracted via osmotic shock are extremely acidic (b = -95.1) indicating that basic proteins are mostly absent in this latter fraction. Figure 2-8 shows that the protein extraction with 0.1 N NaOH resulted in more proteins identified, including 85% of the proteins from the osmotic shock fraction. 20 - -~ 15 0 e; 10 0 j 5 - z 0 2 A ♦ ♦ ♦ ♦ ♦ .. ♦ ♦♦- -♦- ♦- ♦- ........ ♦♦♦ ... .... . ... ......... ♦ ♦ ♦ ... .......... .. . .... ...... .. ♦ ..... ------------ ------...... -♦--........ -----♦ --♦-----♦---- ... ♦ ........... 11111■11 ♦♦ ..... ♦♦ ...... ·--·-···-··· ........................................ ♦ ♦ ..... I II I 1■11 ■Ill■• ■N ♦ ♦ Ill I I -U· ♦ 8 - 7 6 ........ ....... ..... ♦ • -♦ ----♦♦ -- -•--• -.. --... --♦ -♦ --- I I ■- NUIIII ■IU-11■ 1 ♦ 0--------------- 6 7 9 10 11 12 13 2 4 9 10 11 12 13 pl B pl Figure 2-7. Number of proteins as a function of isoelectric point. (A) Theoretical distribution based on the proteome of PV-1. (B) Theoretical distribution based on proteins with molecular weights between 33.6 and 51.1 kDa ( common range of MW of the proteins identified between osmotic-shock and crude fractions). 86 • osmotic-shock fraction shared • crude fraction Figure 2-8. Venn diagram showing the number of proteins that were uniquely identified in the osmotic shock fraction (in red) and crude fraction (extracted with NaOH; in green). Shared proteins between fractions are shown in yellow. Venn diagram is based on proteins identified with at least 2 peptides. All proteins identified in each fraction are listed in Supplemental Table S2-1. 29 Protein interaction experiments were designed to corroborate the results obtained by the proteomic analysis and to test if proteins were actually binding to the PV-1 mats. Figure 2-9a shows that cytochrome c, which is positively-charged at pH 5.25, binds to the PV-1 mats. It remains in the mats even when elution with 1 M NaCl is attempted, suggesting that a force stronger than electrostatic interaction is taking place between the protein and the mats. A pH increase of the elution solution to pH 13 with 0.1 N NaOH, eluted effectively the spiked content of cytochrome c. At this pH, cytochrome c is above its isoelectric point (pl=I0.5); therefore, it would be negatively charged. The point zero charge (PZC) of the stalks of PV-1, the pH at which point the charge is neutral, is not known; however, it is a fair assumption that it will be negatively charged at pH 13. This is because the PZC of pure poorly crystalline 2LF has a value that is 8.11-8.3 (Appelo et al., 2002; Lafferty et al., 2005), which would indicate that above this pH the surface charge of the mineral is negative. While the composition of the stalk is not known in detail, 2LF is directly associated with it (Chan et al., 2011; Edwards et al., 2003b; Toner et al., 2009) 30 (Figure 2-10). Therefore, at pH 13, negative charge repulsion can explain that cytochrome c was eluted at this high pH. 70 □ PY! mats + NaOH ■ PY! mats+ cyt c + NaOH 60 ■ PY! mats + cytc + NaCl 50 'oil 2, 40 ~ ;;:: = ·~ 30 .. 20 10 0 ""' El A 10 11 12 13 14 Elution Fraction # 70 □ PV I mats + NaOH ■ PY! mats+ BSA + NaOH 60 ■ PVI mats + BSA + NaCl 50 'oil 2, 40 :s! ;;: = ·a:; 0 30 0: 20 I 10 10 12 13 14 B Elution Fraction # Figure 2-9. (A) Interaction of cytochrome c with biogenic iron oxyhydroxide. Fractions 1-4 were eluted in 20 mM sodium acetate buffer, pH 5.25. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M NaCl. Error bars indicate± one standard deviation from the mean. (B) Interaction of BSA with biogenic iron oxyhydroxide. Fractions 1-4 were eluted in 20 mM Tris buffer, pH 8.00. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M NaCl. Error bars indicate ± one standard deviation from the mean. 31 - 2-line ferrihydrite c=J PV-1 stalks - 6 months 25 - PV-1 stalks - 12 hours 20 d=1.497(7)A 10 o~~~~~~~~~~-~~~~~~~~~~---.------f :l 10 15 20 25 30 Two-Theta (deg) Figure 2-10. X-ray diffraction pattern of the stalks of PV-1 compared to synthetic 2LF, showing the development of slightly more crystallized iron oxyhydroxide with time. At 12 hours, the sample was growing at log-phase and the biofilm had an off-white/light yellow color. At 6 months, the sample was reddish. At 3 days of growth, the sample had an orange/reddish color which generated a similar spectrum as the 12 hour sample ( data not shown). The d-spacing values correspond to 2LF for this synthetic material. Figure 2-9b shows that BSA, which is negatively-charged at pH 8.00, also binds to the PV-1 mats. Similarly to cytochrome c, it is retained in the mats even when elution with 1 M NaCl is attempted. As mentioned above, this suggests that more direct interactions than electrostatic forces are taking place between the protein and the mats. This can include hydrophobic interactions. A pH increase to pH 13, eluted effectively the spiked content of BSA, which is still negatively-charged because it is above its pl value of 4.75. At pH 8.00; it is unclear what charge the stalks would have because its PZC is not known. At pH 13, because it is such an extreme pH, it is assumed that the stalks would be negatively-charged. Thus, negative charge repulsion can likely explain our observations that BSA was eluted at pH 13. 32 In comparison to the control using synthetic 2LF (Figure 2-10), PV-1 mats have a lower retention capacity of the proteins spiked. This indicates that it is easier to des orb proteins from PV-1 mats than from synthetic 2LF. At pH 5.25, positively-charged cytochrome c binds to positively-charged 2LF (PZC~S.11-8.33). In this case, repulsion is expected but the 15 minutes contact time, allows for the proteins to bind effectively to synthetic 2LF. The desorption of cytochrome c from synthetic 2LF was complete but took five fractions of 0.1 N NaOH to achieve it (fractions 5-9). In comparison, cytochrome c was desorbed from PV-1 mats effectively with the first fraction of 0.1 N NaOH added (Figure 2-lla). Figure 2-llb shows that at pH 8.00, negatively-charged BSA binds to positively-charged synthetic 2LF. The desorption of BSA from synthetic 2LF is only partially achieved, with synthetic 2LF retaining close to half of the original amount of BSA spiked after elution is attempted with 0.1 N N aOH. In this latter case, the higher protein retention capacity observed in synthetic 2LF can be explained by a combination of electrostatic forces as well as direct mineral-protein interaction (i.e. hydrophobic interaction), which is a type of interaction that can be irreversible (Dobrikova et al., 2007; Safi et al., 2011; Rocker et al., 2009). Spiked BSA was only minimally eluted with IM NaCl, which would indicate that the hydrophobic interactions between the protein and mineral play a major role than the electrostatic forces involved. 30 25 20 'oil 2, 32 " 15 >= .s " 2 ~ 10 0 A 20 15 'oil 2, 32 " 10 >= 0: ·;; 2 ~ 0 B ..... 2 t 2 • 4 6 Iii 4 6 33 ■ 2-line-ferrihydrite + cyt c + NaOH ■ 2-line-ferrihydrite + cyt c + NaCl l Iii I • ..... :E Iii .__ 7 8 9 10 II 12 13 14 Elution Fraction # ■ 2-line-ferrihydrite +BSA+ NaOH ■ 2-line-ferrihydrite +BSA+ NaCl llllil -- aa 7 9 10 I] 12 13 14 Elution Fraction # Figure 2-11. (A) Interaction of cytochrome c with synthetic iron oxyhydroxide. Fractions 1-4 were eluted in 20 mM sodium acetate buffer, pH 5.25. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M NaCl. Error bars indicate± one standard deviation from the mean. (B) Interaction of BSA with synthetic iron oxyhydroxide. Fractions 1-4 were eluted in 20 mM Tris buffer, pH 8.00. Fractions 5-14 were eluted in 0.1 N NaOH or 1 M NaCl. Error bars indicate ± one standard deviation from the mean. 34 DISCUSSION One of the goals of this study was to develop practical methods to increase biomass of FeOB so that proteomic studies could be realized. Past studies have shown that 10' cells of Escherichia coli and Streptomyces coelicolor yield 100-150 µg and 143-181 µg of protein respectively (Bremer and Dennis, 1996; Cox, 2004). Using these numbers as guidelines, and the minimum requirements of 30-50 µg of protein for a full proteomic profile via LC-MS/MS, the goal is to achieve a minimum of 10' M. ferrooxydans cells per experiment. This total cell number yields enough protein to use for proteomic analysis, with enough material for a possible technical replicate. However, if spectroscopic analysis or activity in-gel assays are planned ( e.g. staining with heme stain), then a minimum cell number of l0 10 needs to be targeted so that protein yields are closer to 1 mg. The growth curve obtained from the large batch cultures (Figure 2-1) shows a cell density of> 10 6 cells/mL and that a total number of cells of 10 9 can be achieved per bottle within 24 hours. This can be compared to total cell numbers of 10 7 within 24 hours using other types of cultivation methods such as the gradient plate method (Emerson et al., 2007). The growth curve also shows that the cell densities have intermediate values compared with other previously published cell densities of 10 6 cells/mL to 10 7 cells/mL (Emerson et al., 2007; Chan et al., 2011). Cultivation with 10 large bottles (i.e. total of 8 L of culture), following the method presented herein, produces enough biomass (10 10 cells) for proteomic analysis and in-gel activity assays. Alternatively, a more concentrated inoculum should also increase final cell yields. 35 The doubling time of 10 hours reported in this study was calculated for the first 45 hours of growth. If it is calculated during the first day of growth, as seen in a recent paper by Kikuchi et al. (2004), the doubling time is 7 hours which is the same value obtained with Kikuchi's diffusion chamber method. The longer than usual doubling times of PV-1 that were determined by Kikuchi et al. (2004) when testing the liquid batch culture method may have to do with the addition of a single -high- dose of Fe 2 + at t~0 h, a practice that is not consistent with the protocols in Emerson and Floyd (2005) for liquid batch cultures which call for daily addition of lower concentrations of Fe. Addition of excessive amounts of Fe at t~0 h sequesters more than usual levels of phosphate in the medium, which in turn increases the doubling time of PV-1. The identification of two proteins that are associated with ABC-phophate-transporters (NCBI GI numbers: 114777113 and 114777114) identified in this study, supports the explanation that PV-1 goes through phosphate limitation by the time of harvest (third day of growth). Therefore, even higher levels of Fe in the medium should exacerbate this effect. The other goal of this study was to present methods to facilitate protein extraction for proteomic analysis of cells that are in close contact with biogenic iron oxides. Experiments on the interactions of proteins with the mats of PV-1 originated with extractions using disruptive forces such as bead-beating, sonication, and even acidification of sample; all of these methods consistently resulted in low yields of proteins in the presence or absence of detergents. Another hint was given when the non disruptive osmotic-shock method was employed to enrich for cytochrome c; instead an 36 enrichment of acidic proteins was obtained. Only one basic protein (pl~ 9.4) was identified, which happened to be one of the proteins associated with ABC phosphate transporters (NCBI GI number: 114777113). By increasing the pH of the extraction solution to pH 13, all the proteins in PV-1 's proteome should in theory be above their pl, which would make them negatively charged if they are expressed. This is a different approach from proteomic studies involving acidophilic FeOB where the Fe(III) is already dissolved in a strongly acidic medium or in acidic mine water and sonication of washed cells is not a problem (Bouchal, et al., 2006; Ram et al., 2006). However, when this approach is taken for PV-1, a substantial amount of white paste remains, which seems to effectively bind to proteins in solution (i.e. no bands seen in gel stained with coomassie blue). This white paste has been mentioned by Emerson and Moyer (2002) as being organic matter originating from the stalks, but no characterization has yet been made. The use ofNaOH along with SDS has long been used in alkaline lysis of bacterial cells (Birnboim and Doly, 1979) and more recently in yeast cells (Kushnirov, 2000). There have been also recent studies on soil proteomics that have used NaOH and SDS as the protein extraction solution, followed by a phenol extraction protocol to remove humic acids (Benndorf et al., 2007). The use of SDS alone can be effective in lysis of marine microorganisms (Furhman et al., 1988). In light of the results from these past studies, it is recommended that the NaOH solution used in this study to desorb proteins from biogenic iron oxides be complemented with SDS to maximize cell lysis ofneutrophilic FeOB as well as to help solubilize membrane proteins in the crude extract. 37 The fact that the spiked proteins were easily adsorbed and desorbed from PV-1 mats, and the different behavior in synthetic 2LF, suggests that poorly crystalline iron oxyhydroxide is not the only substrate playing a role in adsorption/desorption of proteins. This can be explained by a close interaction of Fe and an organic polymer, which is consistent with different models that have been proposed for biomineral formation in PV-1 (Bennett et al., 2014; Chan et al., 2011). These results can also suggest that the proteins bound to PV- 1 mats have the potential to be more accessible or bio-available than proteins bound to synthetic 2LF. CONCLUSION A method that yields sufficient biomass for proteomic analysis involving neutrophilic FeOB has been developed. Our method circumvents the problem of protein adsorption onto biominerals and shows that both acidic and basic proteins can be identified. Using the methods herein presented, a proteomic profile of M. ferrooxydans was produced but a full analysis will be described elsewhere. Our results indicate that the twisted-stalks produced by M. ferrooxydans not only bind strongly to proteins but also can desorb them more easily than synthetic 2LF. This leaves open the possibility that fields of iron mats seen in the ocean floor, such as the recently identified FeMO Deep site at the base of Loi'hi Seamount, near Hawaii (Edwards et al., 2011 ), are natural banks of bioavailable adsorbed protein, not only from neutrophilic FeOB, but from the rest of the microbial community surrounding it. 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A quantitative fluorescence study of protein monolayer formation on colloidal nanoparticles. Nat Nanotechnol. 4, 577-580. 42 Safi, M., Courtois, J., Seigneuret, M., Conjeaud, H, Berret, J.F. (2011). The effects of aggregation and protein corona on the cellular internalization of iron oxide nanoparticles. Biomaterials. 32, 9353-9363. Saini, G. and Chan, S. (2013). Near-neutral surface charge and hydrophilicity prevent mineral encrustation of Fe-oxidizing micro-organisms. Geobiology. 11, 191-200. Schwartz, R., Ting, C.S., and King, J. (2001). Whole proteome pl values correlate with subcellular localizations of proteins for organisms within the three domains of life. Genome Res. 11, 703-709. Schwertmann, U. and Cornell, R.M. (2000). Iron oxides in the laboratory: Preparation and characterization. Hoboken, NJ:Wiley-VCH, 105. Singer, E., Emerson, D., Webb. E.A., Barco, R.A., Kuenen, J.G., Nelson, W.C., Chan, C.S., Comolli, L.R., Ferriera, S., Johnson, J., Heidelberg, J.F., Edwards, K.J. (2011). Mariprofundus ferrooxydans PV-1 the first genome of a marine F e(II) oxidizing Zetaproteobacterium. PLoS One. 6(9):e25386. Toner, B.M., Santelli, C.M., Marcus, M.A., Wirth, R., Chan, C.S., McCollom, T., Bach, W., Edwards, K.J. (2009). Biogenic iron oxyhydroxide formation at mid-ocean ridge hydrothermal vents: Juan de Fuca Ridge. Geochim. Cosmochim. Ac. 73, 388-403. VerBerkmoes, N.C., Denef, VJ., Hettich, R.L., and Banfield, J.F. (2009). Systems biology: functional analysis of natural microbial consortia using community proteomics. Nat Rev Microbial. 7, 196-205. Vizcaino, J.A., Deutsch, E.W., Wang, R., Csordas, A., Reisinger, F., Rios, D., Dianes, J.A., Sun, Z., Farrah, T., Bandeira, N., Binz, P.A., Xenarios, I., Eisenacher, M., Mayer, G., Gatto, L., Campos, A., Chalkley, R.J., Kraus, H.J., Albar, J.P., Martinez-Bartolome, S., Apweiler, R., Omenn, G.S., Martens, L., Jones, A.R., Hermjakob, H. (2014). ProteomeXchange provides globally co-ordinated proteomics data submission and dissemination. Nature Biotechnol. 30(3):223-226. doi is: 10.1038/nbt.2839. 43 Wang, H. and Edwards, K.J. 2009. Bacterial and archaeal DNA extracted from inoculated experiments: implication for the optimization of DNA extraction from deep-sea basalts. Geomicrobiol. J. 26, 463-469. CHAPTER III: Proteomic profiling ofneutrophilic, Fe-oxidizing Mariprofundus ferrooxydans indicate an alternative-complex III, cytochrome c 553 and cytochrome cbb 3 oxidases involved in Fe oxidation. ABSTRACT Neutrophilic-iron-oxidizing-bacteria (FeOB) oxidize Fe at or near neutral pH, making them relevant in numerous non-acidic environments with elevated reduced Fe concentrations. However, the biochemical mechanisms for Fe oxidation by neutrophilic FeOB are unknown, and biomarkers for this process are unavailable. In the ocean, microorganisms in the M ariprofundus genus are the only known organisms to chemolithoautotrophically oxidize Fe and concurrently biomineralize it in the form of twisted stalks. In order to identify active bacterial oxidation of Fe in the environment, biomarkers for this functionality are needed. The aim of this study was to identify the proteins in the electron transport chain and potential functional biomarkers that are diagnostic ofneutrophilic bacterial Fe oxidation. To this end, Mariprofundus ferrooxydans, strain PV-1 was cultivated, its proteins were extracted and assayed for redox-activity and analyzed via liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS) for identification of peptides. The results indicate that a molybdopterin oxidoreductase, a cytochrome c 553 and cbb,-type cytochrome oxidases were highly expressed and suggest involvement in the process of neutrophilic bacterial iron oxidation. Proteins associated with carbon fixation and biofilm formation were 44 45 abundant, consistent with the life style of M ariprofundus. Phosphate transport proteins were also abundant, indicating significant biochemical investment in phosphate uptake in an environment were phosphate is scavenged by reactive Fe oxides. This proteomic survey presents candidate proteins involved in neutrophilic Fe oxidation in the marine environment, providing targets for biomarkers for quantifying this process in environmental samples. INTRODUCTION While the microbial ecology and physiology of many types of marine prokaryotes have been studied in-depth, the biologically catalyzed reaction of iron (Fe) oxidation at or near neutral pH has remained largely unexplored even though iron is one of the most abundant elements on Earth and a major component of the oceanic crust. This lack of data is due, in part, to obstacles ranging from difficulties in obtaining samples of iron oxidizing bacteria (FeOB) mats from the deep sea, culturing fastidious FeOB, relative low cell densities in cultures, and interference of iron-oxides with sample preparation. Consequently, these challenges have impeded the ability to understand the mechanisms of neutrophilic Fe oxidation and inhibited the development of molecular diagnostics targeting genetic markers for such a biological function (i.e. molecular probes targeting genes, transcripts, or proteins indicative of activity). Thus quantitative activities of biological, neutrophilic Fe oxidation have yet to be assessed. Mariprofundusferrooxydans is a chemolithoautotrophic FeOB that belongs to the 46 Zetaproteobacteria class of Proteobacteria and is known for its ability to biomineralize Fe in the form of extracellular twisted filaments (Emerson et al., 2007; Chan et al., 2011 ). It is one of the few isolates from the marine environment that has been shown to chemolithoautotrophically oxidize iron at neutrophilic conditions, in direct competition with abiotic chemical oxidation (Emerson et al., 2007; Edwards et al., 2003a; McBeth et al., 2011). Where Zetaproteobacteria are abundant, the biomineralizing capacity is quite important, contributing significantly to total iron deposition in certain deep-sea hydrothermal vent systems (Emerson and Moyer, 2002; Kato et al., 2009). Their presence in environmental samples has also been documented in several deep sea sites including Juan de Fuca Ridge, Lo'ihi Seamount, Tonga Arc and Southern Mariana Trough (see McAllister et al., 2011 for review). Recent studies in coastal waters of Maine, USA, and China have shown evidence that this class of microorganisms is involved in microbiologically-influenced-corrosion (MIC) of steel (McBeth et al., 2011; Dang et al., 2011 ). In addition to its relevance, M. ferrooxydans is a useful organism to study because its genome has been recently sequenced and annotated (Singer et al., 2011); however, its genome contains 101 coding DNA sequences (CDS) for energy production and conversion and at least 37 CDS for different types of cytochromes, based on pfam ID search in Integrated Microbial Genomes (IMG)(Markowitz et al., 2012). Therefore, a proper model of the electron transport chain is difficult to elucidate without some information regarding expression of genes. In this study, a proteomic profile of M. ferrooxydans was generated to help answer 47 fundamental questions about its physiology, with a focus on the expressed proteins that are part of the electron-transport-chain (i.e. cytochromes). Recently developed methods to obtain enough FeOB biomass for proteomic analysis (Barco and Edwards, 2014) were applied and allowed us to identify both abundant and low-copy number proteins in active M. ferrooxydans' cultures. Potentially critical proteins involved in Fe oxidation are identified and discussed. MATERIALS AND METHODS Culturing: To generate enough biomass for protein extraction and characterization, a recently developed large-scale culturing method for M. ferrooxydans was used (Barco and Edwards, 2014). M. ferrooxydans, strain PV-1 was grown from a stock culture provided by David Emerson (Bigelow Laboratory for Ocean Sciences, East Boothbay, ME, USA). Final salinity was decreased from 37 ppt to 35 ppt (average seawater salinity) to prevent possible expression of salinity stress proteins. The large batch cultures were grown in ten replicate I L autoclavable polycarbonate bottles (VWR, Visalia, CA) per experiment with each bottle containing 800 mL of medium. The culture vessel was inoculated with 40 mL of log-phase M. ferrooxydans ( approximately I 0 7 cells grown in I 00 mL batch culture) and incubated in the dark horizontally at room temperature without agitation. Filter-sterilized Fe(II) and air were added every 24 hours. Cell counts to determine cell density were performed as described in Emerson and Moyer (2002). Briefly, duplicate samples were fixed with paraformaldehyde solution (0.8% final 48 concentration), stored at 4 °C for 1 hour and then frozen at -20 °C until counted. Slides printed with 4 mm diameter circles (Electron Microscopy Sciences, Fort Washington, PA) were coated with 1 % agarose solution and allowed to cool. A sample volume of 4 µL (2 µL ofresuspended culture sample mixed with 2 µL of 1 mM propidium iodide solution (Life Technologies, Grand Island, NY)) was loaded within the boundaries of the circle and allowed to dry. Fifty fields per circle were counted at 1 OOX magnification using an epifluorescent Axiostar Plus microscope equipped with an HBO 50 mercury lamp and Cy3 filter for green light excitation. Protein extraction - crude extract: To isolate proteins from the large-scale cultures of M. ferrooxydans, recently developed protein extraction protocols to overcome interference from iron oxides were used (Barco and Edwards, 2014). A total of 16 L (8 L duplicates) oflate-log phase cultures were harvested by filtration on 0.22 µm mesh black Whatman Nucleopore polycarbonate filters (GE Healthcare Life Sciences, Piscataway, NJ) with 10 µm support TCTP filter (EMD Millipore, Billerica, MA). The filters were incubated in 80 mL of 0.1 N NaOH/2% (w/v) sodium dodecyl sulfate (SDS) solution on ice for 10 minutes with vigorous vortexing for 30 seconds every 2 minutes to lyse cells and release proteins. The mixture was centrifuged twice at 14,000 x g for 10 minutes at 4 'C to pellet cellular debris and iron oxides. The clarified supernatant was concentrated and buffer exchanged with 40 mM Tris-Base, pH 8.5 by using Macrosep 3K concentrators (Pall, Ann Arbor, MI) with 3 kDa molecular weight cutoff (MWCO) membrane according to manufacturer's instructions. SDS was removed with SDS-OUT (Thermo Fisher 49 Scientific, Rockford, IL) according to manufacturer's instructions. The above procedure results in a concentrated slightly pink crude extract containing both insoluble and soluble proteins, which was then frozen at -20 °C until further analysis. Protein extraction - insoluble (membrane) fraction: A total of 16 L (8 L duplicates) of late-log phase cultures were harvested as described above, and then the filters were immersed in total volume of250 mL of cold, filter-sterilized 0.2 M oxalic acid, pH 3.0 for two hours with mixing on a shaker table. The cell-separation protocol is then followed per Barco and Edwards (2014) to remove stalks from cells. The pellet containing cells is resuspended in 5 mL of 0.5 M NaCl/40 mM Tris-Base buffer at pH 8.5 and sonicated for IO cycles of 30 seconds pulses followed by 30 seconds of cooling using a Branson 450 Digital Sonifier (Branson Ultrasonics, Danbury, CT) to lyse the cells. The resulting protein extract was centrifuged at 14,000 x g for IO minutes at 4 °C to pellet cell debris. The supernatant was loaded in 13 x 51 mm, thinwall, polyallomer tubes (Beckman Coulter, Brea, CA) and centrifuged at I 00,000 x g for two hours at 4 °C using an Optima Max XP ultracentrifuge (Beckman Coulter) to fractionate between soluble (supernatant) and membrane proteins (pellet). The reddish pellet containing the membrane fraction was washed with O. 5 M Na Cl/ 40 mM Tris-Base buffer for two additional ultracentrifugation cycles and was subsequently resuspended in I mL of0.5% Ultrol Grade, n-Dodecyl-beta D-maltoside (Calbiochem, San Diego, CA) to solubilize proteins. The solubilized membrane fraction was quantitated by using the RC DC™ Protein Assay (Bio-Rad) according to manufacturer's instructions using a UV-1601 spectrophotometer (Shimadzu 50 Scientific Instruments, Carlsbad, CA). All samples were stored at -20 °C. Protein extraction - soluble fraction: A total of 16 L (8 L duplicates) oflate-log phase cultures were harvested as described above and treated for osmotic-shock by immersing the filters in 30 mL of 40 mM Tris-Base pH 8.5/20% sucrose solution followed by addition of 60 µL of0.5 M EDTA, pH 8.0 and stirring slowly at room temperature for 10 minutes (Neu and Heppel, 1965). The mixture was centrifuged at 14,000 x g for 10 minutes at 4 °C and the supernatant discarded. The pellet was resuspended in 30 mL of ice cold, sterile 5 mM MgClz and incubated on ice for IO minutes with slow stirring for osmotic shock (Ausubel et al., 1989). Periplasmic and cytoplasmic proteins are released during this step but they bind strongly to the iron oxides (Barco and Edwards, 2014). The mixture was centrifuged again at 14,000 x g for IO minutes. The supernatant was removed, and the pellet was resuspended in cold, filter-sterilized 0.1 N NaOH to both lyse the cells and desorb proteins from the iron oxides. This mixture was incubated on ice for 10 minutes with vortexing every I minute, and centrifuged at 14,000 x g for 10 minutes to pellet any cellular debris and iron oxides. The supernatant containing the crude protein extract was transferred into a thinwall, polyallomer tube for subsequent ultracentrifugation at I 00,000 x g for 2 hours to separate soluble proteins from membrane proteins. The orange supernatant containing the soluble fraction was concentrated and buffer-exchanged with 40 mM Tris-Base, pH 8.5 by using 3K Microsep concentrators (Pall) with 3 kDa MWCO membrane. Soluble fractions were quantitated by using the Quick Start™ Bradford Assay (Bio-Rad, Hercules, CA), according to manufacturer's 51 instructions using a UV-1601 spectrophotometer (Shimadzu Scientific Instruments). Sodium dodecyl sulfate - polyacrylamide gel electrophoresis (SDS-PAGE) and gel assays: Proteins were separated on gels at 90 V according to standard protocols (Laemmli, 1970). Soluble (n ~ 2) and solubilized membrane fractions (n ~ 2) were run on 12% TGX™polyacrylamide gels (Bio-Rad) under reduced and non-reduced conditions (i.e. no dithiothreitol (DTT) and no heating). Gels were stained with Bio-Safe Coomassie Stain (Bio-Rad) for 1 hour, destained with HPLC-grade water for 2 hours and visualized with a Gel Doc™ XR+ imaging system (Bio-Rad) equipped with a white light transilluminator. Images were analyzed with Image Lab™ software (Bio-Rad). Slices were excised as close as possible to the band of interest with a clean sterile scalpel, stored in HPLC-grade water at 4 °C and immediately submitted for proteomic analysis. For the full proteomic profile, 30 µg of duplicate crude protein extracts were loaded on a NuPAGE® Novex® 4-12% Bis-Tris pre-cast polyacrylamide gel (Life Technologies) and run at 100 V in non-reducing conditions. The gel was stained and destained as described above. The duplicate gel lanes were cut into 11 slices per lane in same areas of molecular weight, stored in HP LC-grade water at 4 °C and immediately submitted for proteomic analysis. In-gel, protein redox activity was assayed according to Francis and Becker (1984). Peptide identification and protein analysis: Excised protein samples were submitted to the Children's Hospital of Los Angeles - Proteomic Core Facility for trypsin-digestion 52 and LC-MS/MS on a Thermo LTQ-Orbitrap XL mass spectrometer (San Jose, CA) equipped with an Eksigent (Dublin, CA) Nanoliquid Chromatography 1-D plus system. The resulting MS/MS spectra were searched against the proteomes of M. ferrooxydans strain PV-1 in the Uniprot database using the Proteome Discoverer SEQUEST Daemon search engine. Alternatively, the proteome of M. ferrooxydans strain M-34 in the JGI database was searched to cover gaps present in PV-1 's proteome. The criteria for having a protein identified is that there needs to be a minimum of two unique peptides matching to it, with each peptide established at> 95% probability. Predictions of subcellular localization were performed on two different platforms, CELLO and PSORTb (Yu et al., 2004, 2010). Protein signatures were searched on InterProScan (Hunter et al., 2012; Quevillon et al., 2005). Alignment and analysis of amino-acid sequences were performed in Geneious version R6 (Biomatters, Auckland, New Zealand). Maximum-likelihood phylogenetic trees were obtained from Geneious using the Phy ML plugin (Guindon and Gascuel, 2003) and based on 1000 bootstrap replications. The mass spectrometry proteomics will be deposited to the ProteomeXchange Consortium via the PRIDE partner repository. RESULTS AND DISCUSSION General proteomic profile results: Overall, 825 proteins from a total of2866 protein coding genes (28. 7 % ) were identified by using the generated amino acid database from the genome of PV-1 (Supplemental Table S3-l ). There were 200 gene products annotated 53 as hypothetical proteins (24.2 % of all identified proteins), including the most abundant protein (SPVl _ 07114). COG functional category codes were assigned to 709 proteins, including some proteins which were annotated as hypothetical (86 % of all identified proteins). The 25 most abundant proteins identified in the profile are listed in Table 3-1 and include phosphate transport proteins, proteins associated with the Calvin cycle, a soluble cytochrome c, a cytochrome cbb3 oxidase subunit, and a molybdopterin oxidoreductase. All of these gene products and their potential function in M ferrooxydans will be discussed below among other proteins of interest. Table 3-1. The 25 most abundant proteins identified in M. ferrooxydans' proteomic profile. Gene Product Name Locus Tag NCBI GI/ IMG Gene ID Molecular Weight COG # of Peetides Identified hypothetical protein SPV1_07114 GI 114777 11 3 40 kDa 1700 ABC transporter phosphate-binding protein SPV1_07119 GI 114777 11 4 37k0a COG0226===ABC-type phosphate transport system , periplasmic component 1174 chaperonin, 60 kDa subunit SPV1_00170 GI 114777816 57 k0a COG0459===Chaperonin GroEL (HSP60 family) 896 translation elongation factor Tu SPV1 _09894 GI 114778826 43 k0a COG00S0===GTPases - translation elongation factors 776 Ribulose-bisphosphate carboxylase SPV1 04963 GI 11477565 1 51 kDa COG1850===Ribulose 1,5-bisphosphate carboxylase , large subunit 555 hypothetical protein SPV1=06074 GI 114777966 22kOa COG3637===Opacity protein and related surface antigens 538 elongation factor Ts SPV1_05739 GI 114777899 31 kDa COG0264===Translation elongation factor Ts 435 F 0 F, ATP synthase subunit beta SPV1 13824 - GI 114777383 50 kOa COG0055=== F 0 F 1 -type ATP synthase, beta subunit 429 flagellar protein SPV1_01952 GI 114776444 37 kOa COG1344===Flagellin and related hook-associated proteins 414 F 0 F 1 ATP synthase subunit alpha SPV1 13814 GI 114777381 55kOa COG0056=== F 0 F 1 -type ATP synthase, alpha subunit 413 - transketolase SPV1 05909 GI 114777933 71 kDa COG0021 ===Transketolase 370 translation elongation factor G SPV(09899 GI 114778827 76 kOa COG0480===Translation elongation factors (GTPases) 345 Fructose-bisphosphate aldolase , class II SPV1_05929 GI 114777937 37kOa COG019 1===Fructose/tagatose bisphosphate aldolase 338 ribosomal protein L7/L 12 SPV1 - 12170 GI 114778902 13 kOa COG0222===Ribosomal protein L7/L 12 333 DNA-directed RNA polymerase beta' subunit SPV1 - 12180 GI 114778904 154 kDa COG0086===DNA-directed RNA polymerase, beta' subu niU160 kD subu nit 316 Cytochrome c~ A37KDRAFT_02147 Gene ID 2513995557 28kOa COG2863===Cytochrome c~ 304 cytochrome oxidase, cytochrome c subunit SPV1_0740 1 GI 114778975 33kOa COG2993===Cbb 3 -type cytochrome oxidase, cytochrome c subunit 301 hypothetical protein SPV1_07396 GI 114778974 25 kOa 295 Glutamine synthetase type I SPV1_00922 GI 114776238 45 kOa COG0174===Glutamine synthetase 293 F 0 F,-typeATP synthase, subun it b SPV1 13804 - GI 114777379 17kOa COG0711 === F 0 F 1 -type ATP synthase, subunit b 292 Molybdopterin oxidoreductase Fe, S 4 region SPV1_03948 GI 114775448 83 kOa COG0243===Anaerobic dehydrogenases, typically selenocysteine-containing 255 molecular chaperone DnaK SPV1_07796 GI 114777593 69 kOa COG0443===Molecular chaperone 242 DNA-directed RNA polymerase beta subunit SPV1 - 12175 GI 114778903 149 kDa COG0085===DNA-directed RNA polymerase, beta subuniU140 kD subunit 242 hypothetical protein SPV1 00467 GI 114778085 14 kDa 233 ribosomal protein S8 SPV(09814 GI 114778810 15 kOa COG0096===Ribosomal protein S8 231 hypothetical protein SPV1_08251 GI 114778394 17kOa COG0085===DNA-directed RNA polymerase, beta subuniU140 kD subunit 225 Phosphoglycerate kinase SPV1_05919 GI 114777935 4 1 kDa COG0126===3-phosphoglycerate kinase 218 50S ribosomal protein L 1 SPV1 12160 GI 114778900 23kOa COG0081 ===Ribosomal protein L 1 216 glyceraldehyde-3-phosphate dehydrogenase, typ{ SPV1=05914 GI 114777934 36 kOa COG0057===Glyceraldehyde-3-phosphate dehydrogenase/erythrose-4-phosphate del 21 1 Figure 3-1 illustrates the proportion of assigned COG functional categories based on the number of assigned spectra in the obtained proteomic profile. Translation, ribosomal structure and biogenesis (COG code J) was the functional category with the most assigned spectra (20.3 %). The three elongation factors required for translation in prokaryotes - Tu, Ts and G - were among the most abundant proteins expressed in 54 PV-1 (Table 3-1 ). Proteins associated with translational functions are also abundant as seen in other proteomic studies involving bacteria (Chao et al., 2010; Hansmeier et al., 2007; Rosen et al., 2004). As described above, there was a high proportion of proteins with no known function and no COG code assigned (NC in Figure 3-1). They made up the second largest group and constituted 12. 8 % of all the assigned peptide spectra. The third largest COG group represented proteins with function in energy production and conversion. Within this group, the most abundant proteins identified were associated with ATP synthase and electron-transport chain components such as cytochrome cbb 3 oxidase, cytochrome cm and molybodpterin oxidoreductase. The fourth largest group belonged to proteins with post-translational modification, protein turnover and chaperone function (COG code 0). The most abundant proteins within this group were the chaperone DnaK and the chaperonin GroEL that may have functions related to preventing aggregation of newly synthesized polypeptides as well as preventing protein misfolding (Hartl and Hayer-Hartl, 2002). In strain PV-1, the most abundant protein with 1700 identified peptides (SPVl _ 07114) was not assigned a COG code. However, it has to be taken into account that the gene encoding for SPVl _ 07114 is located in an operon comprised entirely of genes associated with ABC-phosphate transporters (COG code P; inorganic ion transport and metabolism). A hypothetical inclusion ofSPV1_07114 into COG P makes this group the fourth largest overall. This indicates that PV-1 places a high priority on ion-transporters and actively invests energy in producing them. E G 0 ■ J - Translation, ribosomal structure and biogenesis NC - no assigned COG ■ C - Energy production and conversion ■ 0 - Posttranslational modification, protein turnover, chaperones G - Carbohydrate transport and metabolism E -Amino acid transport and metabolism ■ M - Cell wall/membrane/envelope biogenesis ■ P - Inorganic ion transport and metabolism ■ K - Transcription ■ X -Assigned multiple COG ■ R - General function prediction only ■ H - Coenzyme transport and metabolism ■ N - Cell motility ■ F - Nucleotide transport and metabolism L - Replication , recombination and repair S - Function unknown ■ U - Intracellular trafficking, secretion, and vesicular transport T - Signal transduction mechanisms ■ I - Lipid transport and metabolism ■ Q - Secondary metabolites biosynthesis, transport and catabolism V - Defense mechanisms D - Cell cyde control, cell division, chromosome partitioning Figure 3-1. M f errooxydans' proteomic profile sorted by cluster of orthologous genes (COG) functional categories. Redox activity assay: When heme-staining of a gel is performed under non-reducing 55 conditions, redox active, heme and non-heme proteins can be identified. The non reduced, non-heated soluble fraction that was stained with the heme stain (Figure 3-2A) shows that there are in particular two regions in the gel that display high redox activity, an area between 25-27 kDa with possible multiple bands and an area with a doublet at > 200 kDa. The band(s) around 25-27 kDa were also seen in a sample that was reduced and heated (Figure 3-2B) indicating that protein(s) with covalently-bound heme prosthetic group could be present in this area. LC-MS/MS analysis of this area of the gel identified a cytochrome c 553 . This cytochrome c (locus tag: A37KDRAFT_02147) was identified by using an amino acid sequence database generated from the draft genome of Mferrooxydans strain M34. It was subsequently determined that there is a sequencing 56 gap (12,704 bp) in the draft genome of Mferrooxydans strain PV-1 that flanks the cytochrome c553 gene (locus tag: SPVI _ 07306) at the 269 bp position. Based on the fact that the genes upstream from cytochrome c553 in PV-1 share 100% identity with the corresponding genes in M34, we assume that the percent identity is equal or close to 100% for this gene. The theoretical molecular weight (MW) of cytochrome c553 in M34 is ca. 25.2 kDa if the signal peptide is not included in the mature protein, which agrees with the results observed in the PV-1 protein extract after an SDS-PAGE run (Figure 3- 2A-B). The doublet band> 200 kDa contains several redox proteins including a molybdopterin oxidoreductase. 200- 120 30 40 A B C Figure 3-2. Heme-stained SOS-PAGE gels of soluble and insoluble fractions. (A) Soluble fraction in non reducing conditions. (B) Soluble fraction in reduced conditions. (C) Membrane fraction in non-reducing conditions. The excised gel areas spanning regions 80 kDa to 120 kDa contain all the proteins encoded by the mob operon. The non-reduced, non-heated membrane fraction (Figure 3-2C) also underwent the redox activity assay after SDS-PAGE. Membrane fractions were run under non-reducing conditions to avoid precipitation of proteins. In this fraction, there was a region of high redox activity between ca. 80 kDa and 150 kDa. This region contained most of the gene products in the mob operon (mobABCDE and mobG; no mobF), with the iron-sulfur protein containing a molybdopterin oxidoreductase (SPVl _ 03948) being the most abundant protein based on assigned number of spectra (Supplemental Table S3-2). Cytochrome ebb, oxidases were also identified in this region. Since the sample was not reduced, nor heated, disulfide bonds would still be present; therefore, it is possible that these proteins migrated in complexes. 57 Fe oxidation and the electron transport chain: The schematic diagram in Figure 3-3 presents an emergent picture from the electron transport chain (ETC) in PV-1. Electrons removed from Fe(II) are transported by a periplasmic cytochrome Cm, which are then donated to a cbb,-type cytochrome oxidase, and ultimately accepted by oxygen, generating a proton motive force in the process that can drive the production of ATP through ATP synthase. Reverse electron transport is also a possibility, with the electron from cytochrome c 553 being donated to the cytochrome be 1 complex ( complex III) (SPVl _ 03843-SPVl _ 03873), then transported to the quinone pool and ultimately accepted by NADH dehydrogenase to produce reducing power in the form ofNADH. However, peptides for this bc 1 complex, which is linked by a short dehydrogenase, are present at relatively low levels. Instead, an Alternative Complex III (ACIII) that was recently discovered to be an analog of the bc1 complex in the respiratory chain (Yanyushin et al., 2005; Pereira et al., 2007; Refojo et al., 2013) is represented in the profile by an abundant number of peptides. A molybdopterin oxidoreductase associated with the AC III was identified in PV-1 through our preliminary proteomic work in Singer 58 et al. (2011) and was denominated the 'Mob' (Mo-binding) complex, a complex seen in other neutrophilic FeOB such as Gallionella, Sideroxydans and Leptothrix. The mob operon (mobABCDEFG) is located upstream from the genes that encode the putative bc1 complex and includes a penta-heme cytochrome c, a molybdopterin oxidoreductase, ferredoxin, polysulphide reductase, a hypothetical protein with putative quinol- cytochrome c oxidoreductase function, a mono-heme cytochrome c, and another polysulfide reductase (Figure 3-4). Fe 2 + Fe 3 + \ \,.,.__;,"' OM --------------------cyc2-mf ______________ _ NADH IM Dehydrogenase -=--=-Q 7"\. NADH NAD+ ) cyt cm 2W \ cyt cbb 3 1/2 O,+ 2W H,O ATP synthase "' ATP ADP Figure 3-3. Proposed electron transport chain ofM. ferrooxydans based on proteomic analysis. OM, outer membrane; IM, inner membrane. Mariprofundus feffooxydans - Cytochrome c mobA mobB mobC mobD mobE mobF mobG - Molybdopterin oxidoreductase Gaflionella capsiferriformans ~ - Ferredoxin FeOB - Polysulphide reductase mobA mobB mobC mobD mobE mobF mobG - Quinol:cytochrome c oxidoreductase Sideroxydans lithotrophicus - Periplasmic protein of unknown function mobA mobB mobC mobD mobE mobF mobG Rhodothermus marinus non-FeOB actA actB actC actD actE actF actG Chloroflexus aurantiacus actB actE actA actG Figure 3-4. Mob operon ofFeOB in comparison to the act operon in heterotrophic bacteria (R. marinus) and phototrophic bacteria (C. aurantiacus). 59 Due to the recent discovery of the ACIII encoded by the act operon, there are still many questions regarding its function despite important advancements in the model organisms Chlorojlexus aurantiacus, a filamentous, anoxygenic phototroph, and Rhodothermus marinus, a marine heterotroph. It was proposed that actB, which is the product of a fused gene encoding an iron-sulfur protein analog to mobB and mobC, contains neither molybdenum nor molybdopterin (Yanyushin et al., 2005; Pereira et al., 2007). The presence of the four conserved cysteines at the N-terminal of mobB differentiates it from actB in Chlorojlexus and Rhodothermus and is suggestive of the presence of a molybdopterin cofactor and molybdenum. Remarkably, these conserved cysteines are also seen in the other neutrophilic FeOB that contain the mob operon (Figure 3-5). Since actB and mobB belong to the superfamily of molybdpterin-binding proteins, a maximum likelihood (ML) phylogenetic tree was produced (Figure 3-6) based on an amino acid alignment using proteins from this family, including fdhH (formate dehydrogenase, subunit H) and nap A (nitrate reductase, subunit A). The ML phylogenetic tree shows that mobB forms a distinct clade from actB, fdhH and napA suggesting that it is functionally different. It remains important to elucidate the function of the mob complex in FeOB, and specifically determine if it is associated with downhill electron transport, uphill (reversed) electron transport or both. There is no evidence at this point that the ACIII is involved in reversed electron transport by donating electrons to the quinone pool as is the case with the bc 1 complex, to produce reducing power in the form ofNAD(P)H. In this proteomic profile reported here, all the gene products belonging to the mob operon were identified (Supplemental Table S3- l ). In addition to this, the gene product annotated as a 60 'molybdopterin oxidoreductase Fe 4 S 4 region' (mobB) is Mthin the top 25 most abundant proteins in the whole profile (fable 3-1 ), suggesting that it is functionally relevant to PV- 1. 1. mob8 • ~ ~ydMl:t P\,C,-1 2.mob8-~~sM3A 3, tl'l00B • G,l,l(OntRf ~IW E$-2 4, mobB·S~nslirhofroplrkusE.S-1 5. mob0 - lEIJ)IOlM)( ochr:acH l 12 6, mob8 • G~bo mid;.<1$1$ Bem 7. mob8 - G~Ut&"Mt86Uo&rlsRf & e.mooe-G~~nsGs-15 9 mob8 •G~.sp . M18 10. actB- CtJJomilexus&l#al1tlaCUs J-10-F\. 11. aicie • R~s mo,,nv.s ,o;o 11~ 12;0 1JO ,.-o ,so ,~ ,~o 1~ •~ .---1:~-l'- -;;;:-_ T m i e:, _,_ ..:,. I T cti 'I ,I M - vri? ""' mc:I =---Tw.... OYVRPG I£MYYAS': C IA C VHARIR£GRVLKLE:GNPVSDVNHGR C MGOAGLQS HYN9DRLTKPMLRKG----GKLV£ lSWD£.A£0VLJ\K DYVRPGIEMYYAS C C AC C VHARIREGRVLKLEGNPVSDVNHGR C MGQAGLOS H'fNFDRLTKPMLRKG----GKLVEISWDEAEDVLRK DYHVPG IGVHNS: C I: A I: VNGRVRoGRVl,Kl,~GNPPSA tNKGKl C 1,GQAGVQHHYN~DRVRoPl,1,RNG··•·NKGoA I SW DKAYALIAo SYVIPS ISHYFNS" C C AC C IMGRVREGRVLKAEGNPNSPINRGK C LGQSGVQA HYNPDRVRQPLL------- - KGEA I TWOKALGVIA O DFVVPGVGVYYAS C C S C VMGRVREGRVLKLEGSRSSVINAGK C LGQAAVQAHYHPDRLTVPMVREG----NAL KE TTWEKAMALLTE ENIRPGSWTVFAT. C C A C MHLSHROGRVTKA.EGNPi\HPVNRGA C RGQSAPQGLYOPDRLRQVLYRSG----GASRPSNWQOALSAIST EGVTPGKA OYYAS C C A C ILVRVSE.GRAKKIEGNPAHPVNRGK C MG0AVLQELYH1PDRVPQPLKRSGPRGSGAFTR1S \oi££$LELLAG EGITPGKGTYYAS~ C C A C ILVRVSEGRAKKIEGNPEHPIHRGK C RGQAVVQELYHFDRVPHPLKRNGPRGSGAFTRISWQEGIGLLTD t:.GITPGKAVYYAS C C A C lLVRVS&GRAKKIEGNP£HPVNRGK .C RGQALLQE:LYKf'DRVPQPLKRSGPRGSGQfTRISl'it&GLNLLTG EGRVPG I PQYFASTLT LG-GYCTGV L VRSNEGRPTK VEGN PRH PASLGGTOLFAQA El LTMYDPDRSTTV LRQGV -------- PS TWA& FTTTLGf l &EI I PG I PLYYA. TAMP f'R<iSVRPLL VESHEGRPTK IEGNPD!i PLSRG/1. TGVFEQA. SLLN L YDPDRSQQV LRKGE------- - PASWGO FVQ-FA R Figure 3-5. Amino acid alignment of mobB and actB proteins. Sequences 1-4 are from known neutrophilic FeOB. The conserved cysteine amino acids at the N-teiminal ofmobB in FeOB are marked by blue asterisks. 100 100 100 100 100 99 actB - Rhodothermus marinus (ABV55245) actB - Chloroflexus aurantiacus J-10-FL (YP _001634251) ~------- actB - Myxococcus xanlhus DK 1622 (ABF89402) 100 100 100 77 mobB - Mariprofundus ferrooxydans PV-1 (EAU55940) mobB - Mariprofundus ferrooxydans M34 (WP _018294696) mobB - Leptothrix ochracea L 12 (EIM31728) mobB - Sideroxydans fithotrophicus ES-1 (YP _ 003523268) mobB - Ga/fionella capsiferriformans ES-2 (YP _003846257) mobB - Geobacter sp. M18 (AOW15368) mobB - Geobacter uraniireducens Rf4 (YP _001231524) mobB - Geobacter metallireducens GS-15 (YP _006720773) ~---------- mobB - Geobacterbemidjiensis Bern (ACH37156) 75 FeOB ACIII napA- Shewanefla denitrificans 0S217 (ABE54782) napA - Shewanefla oneidensis MR-1 (AAN53924) napA- Klebsieffa pneumoniae 1S22 (CDK74678) napA - Salmonella enterica L T2 (AAL21161) Periplasmic Nitrate Reductase 0.3 100 fdhH - Desulfobacula toluofica Tol2 (CCK79330) fdhH - Clostridium carooxidivorans P7 (AD012080) fdhH - Escherichia coli K-12 (BAE78081) fdhH - Salmonella enJerica ST4/74 (AOX20056) Formate Dehyd rogenase Figure 3-6. Maximum-likelihood phylogenetic tree of proteins within the molybdopterin-binding protein superfamily. Numbers on branches indicate bootstrap values based on 1000 replicates. 61 The identification of cytochrome cbb 3 oxidase, which is a member of the class of respiratory haem-copper oxidases, is consistent with the microaerophilic nature of PV-1, since this type of terminal oxidase has higher affinity for oxygen than the typical complex IV (Pitcher and Watmough, 2010). Because the chemical kinetics ofFe(II) oxidation in seawater are sufficiently slow at reduced oxygen concentrations, marine FeOB can gain access to the soluble pool of reduced Fe (Singer and Stumm, 1970; Millero et al., 1987). Therefore, as a result of microaerobic conditions, the spontaneous chemical oxidation of Fe(II) can be retarded and PV-1 can oxidize it instead, eventually transferring the electrons from Fe to oxygen. There are two different cytochrome cbb3 oxidases that are expressed when PV-1 is grown in batch cultures. This may be reflective of differing concentrations of dissolved oxygen along the vertical length of the medium. The most abundant cytochrome cbb3 oxidase is a previously unidentified respiratory complex that is associated with the locus tags SPV1_07391, SPV1_07396, SPV1_07401 and SPV l _ 07406. This operon consists of a gene with a conserved domain for the catalytic subunit CcoN (SPVl _ 07406), a membrane bound cytochrome c, CcoO (SPVl _ 07401), and two genes that encode for membrane proteins with no conserved domains in known families of proteins. All components of this putative respiratory complex were identified in the proteomic profile, with the exception of SPVl _ 07391, which is a small (5 kDa) membrane protein. Interestingly, this putative operon is missing a gene for CcoP, which is a membrane di-haem cytochrome c. A different cytochrome cbb 3 oxidase that is associated with a ccoNOP operon (SPV1_10291, SPV1_10301, and SPV1_10306) also seems relevant and appears in the proteomic profile as well, although in more moderate numbers and without any peptide hits for CcoN (SPV1_10306). Although genes for cytochrome bd quinol oxidase are present in the genome of PV-1, there is no evidence that they are being expressed. 62 Based on its positioning within the most abundant proteins (Table 3-1 and 3-2), a cytochrome c 553 is suggestive of being one of the most important electron-transfer proteins in PV-1, transferring electrons from Fe(II) to the cytochrome cbb 3 oxidase (Figure 3-3). Its amino acid sequence reveals two conserved CXXCH motifs, which make this a di-haem cytochrome c (Figure 3-7). A protein signature search on InterProScan reveals that it is part of the cytochrome c 4 family of proteins (IPR024167), which are known to have high redox potentials. Analysis on CELLO and PSORTb indicate that this protein is periplasmic and contains a signal peptide. There is evidence originating from studies on Vibrio cholerae and Helicobacter pylori that cytochrome c 553 and cytochrome c,, in general, are the natural electron donors to cytochrome cbb3 oxidase (Koyanagi et al., 2000; Chang et al., 2010). This hypothesis seems to be supported by the proteomic dataset herein presented. An acidophilic FeOB, Acidithiobacillus ferrooxidans, uses a cytochrome c, (GI# 218665058) in order to transport electrons from Fe(II), but it lacks a cytochrome cbb 3 oxidase (Giudici-Orticoni et al., 2000). Interestingly, when a BLASTP search is performed against the M34 genome for a homo log to this cytochrome c 4 , there is only one protein that is classified as an ortholog by IMG: cytochrome cm (30% identity). The same result is obtained when the search is performed against the PV-1 genome, but the cytochrome c 553 is fragmented. Table 3-2. List of cytochromes that were identified in M jerrooxydans' proteomic profile sorted by# of peptides identified. 63 Gene Product Name Locus Tag COGIPFA.WINTERPRO NC81 GI/IMG GENE ID MW Proteome Queried Peptides ki8rllifl8d cytoehtomec., A37KDRAFT_02147 OOG2863u -cyioehrome ~ GENE ID 2513995557 28 kOa M34" 304 cytochrome oxle{ls,e, c:y1oc:hrome c: $1Jbvnil $PV1_07401 000299~,-~pe cy1ochroma w3cSaa . cytochrome c subunit GI 1U77897$ 33k0a PV-1' 301 ey.tO(:htOme c oxida5e heme band oower-bl~ 5lilunit SP\/1 07400 OOG3278::=.cbb 1 .typecylochrom&o.x1dase . &ubooll 1 cytoehromac SPV1: 121as COG3474••-cytochromo't Gl11'77/m1tl 53kCa Pv-1 1 94 GI 114778905 27 kDa PV-1' 50 hypothetical protein SPV1_00030 PFAM00034 • cytochrome c GI 114777788 22kDa PV-1' 50 cytochrome coxldase, mono-netne subunitJF1:(0 SPV1_10301 QOG2893:a::=cob ,-t)'pecywavom&oxldas& . cy1octirome csut:ulit GI 114773941 24 kDa PV-1' 42 l')'VU~t pruttrln SPv1_0»,J IP~0110J1 - m1,mft1aem cy1oct1run1e G11 14n'44s zo~Ca PV•I• ,, cytochrome coxidase, ebbj type, 5Ubunil Ill SPV1_ 10291 OOG201():,:.:=cytochrome c. mono- aAd<liheme variants GI 114n8939 27 kDa PV-1' 29 hypothetlcal prolein SPV1_03928 PFA~t13442 - Cy'l0ct'l,ome_c:bo 1 G I 114775444 2· 1 kDa PV-1' 28 cy:toehrome c, $PV1_08026 OOG3474="'=qtochrome c, GI 114778349 14 k08 PV-1• 21 ey.tOchrome c, da5s I SPV1_ 10264 OOG3474===q,toet,rome c., GI 114778525 12 kDa PV •1' 21 hypOthetieal protein SPV1_0t482 PFAM13442-c:y1ochrome_CC0 1 GI 114776350 11 kDa PV-1' 19 cytochrome c SPV1 03873 OOG3474.u~rome c., cytochrome c petwi:13$,tf•mlty prottin SPV1)3'628 0001858-•~rome c pe1'0Xi::l;UO GI 114775433 26kDa PV-1' 14 GI 11477538,4 36 k0a P\1 . 1• 13 cy.tOChrome b. p!Aa1MI SPV1_11001 OOG196~~~ Ni,F9-fty(kogenase lcytocwome bSllbiJnil G1m1n6738 27 kDa PV-1 1 11 hypothetical p,oteln $PV1 Ot3?6 PFAM00034 - cyloetltome c t,ypo;het'eal p,otelo SPV( 1:!036 PFAAIOOO(l3. eytoc1,ombN GI 1U778970 19k0a P\1 . 1• 9 GI 11 4776945 2A kOa PV-1 1 7 cy:toehromab'b6-4il<eprocaln SPV1_03863 OOG1290:==cytochromebaubuni1ofthebccompleJI GI 114775431 46kDa PV-1' hypothetk:al protein SPV1_07306 0002863-s•ceytochrome ~ GI 114778956 9kDa P\'-1c hypothetical proteln $PV1 01867 PFAM01322-cytochromec ~ ey1od'll'Ol'M c.,. A37KOAAfT _02122 0002863==.::eyioetlron"le ~ GI 114776427 16 k08 PV -t • GENE 10 251399$532 28 kOa M34" cytochrome e 1 • l'lflme prolera'I P'teutSOf SPV1 03858 OOG2857• n q,tochrome c, melhylamlne utilization protefnlcylochrome c pel'Oxida se SPV1= 11291 OOG 1858===cytochrome c pe,oxklase GI 114n5430 26k0a PV-1' GI 114776796 39 k0a PV-t• cy.tochrome c_ A37KORAFT _02123 OOG286J::=cytoehrome c:_ GENE 1 0 2$1399$$33 14 kOa M34' probablecytoctwomect l)t8Q"50f'Pf0C81,i SPV1_10541 C-OG1629::=outor momb rane reoeptor l)(Oteins, mosuyF e transo, GI 114776646 52k0a f'V.,-.. • Homolog lo SPV1_07306 . wtliCh is fragmented by gap ' Found in M34 genome as well ~Fr3gmented by g.ap ; 99'% simile, lOA37KORAFT_02147 • Not cwesont If\ PV1 Qenomo Clue 10 gap • More abundant in membrane fraction {see Tabaa S3) 1 10 20 ~ 40 ~ &0 )'~ 80 MKRMISTMGACALLAWFAAPTVASAEEAMGNVNNGRLIFQNGKGDAVPACQGCHGIDGNGSDEMGTPRLAYQVDTYILKQLTDFAE $ 1 GNA1.Pe.PTIOE ~ ~ HEME BINDING MOTIF " i" NKRTDDTMYQMNDIAKALSPQDRKDLAA YVHTLKSPYIGSNLDQLRRDGAEVGDPAQGKMIAEYGAPDHGVPACKSCHGFHGRSAG ~ HEME BINDIJI«} MOTIF . . ~ ~ ~ . ~ . . RMYPAIGGQNYVYLKHELESWRNGANTSAADIAAGKREDNARVNDYMGQMRAVASHMTDADIANVAAFLTLSKPNSPGNPRTPSHM Figure 3-7. Amino acid sequence of cytochrome C553 gene in M jerrooxydans M34 (IMG locus tag A37KDRAFT_02147). Mariprofundus fem,oxydans M34 Mariproftmdvs terrooxydans PV-1 H c::::I Hypothetical prate in - Truncated hemoglobins c::::::J Cytochrome c - Cbb, cytochrome oxidase subunit - Ferredoxin ~ Mullicopper oxidase - Thioredoxin - Spermine synthase Figure 3-8. Gene neighborhood of area with conserved synteny between Afariprofimdusferrooxydans strains M34 and PV-1. (x) indicates the fragmented cytochrome c553 gene in PV-1 (locus tag SPV1_07306) that is a homologto the cytochrome c553 gene in M34 (y; IMG locus tagA37KDRAFT_02147). A noteworthy outer membrane cytochrome c that was identified in the proteomic profile as a low copy number protein is annotated as "probable cytochrome c 1 precursor protein" (SPV1_10541, Table 3-2). This protein was also identified in the solubilized membrane fraction but with a higher spectral count than in the crude fraction (Supplemental Table S3-3). It has an assigned COG function for inorganic ion transport and metabolism (COG code P), specifically Fe transport as a membrane receptor protein. An interesting aspect to this cytochrome c is that it shares some similarities to other outer membrane cytochrome c such as cyc2 and cyt572, known Fe-oxidizing proteins from A. ferrooxidans and Leptospirillum, respectively (Jeans et al., 2008; Yarzabal et al., 2002, 2004). The amino acid alignment (Figure 3-9) reveals that there is a highly conserved N-terminal as well as a conserved heme-motif (CXXCH) among the three different proteins. As Jeans et al. (2008) pointed out with cyt572 and cyc2, the heme binding motif of cyc2-mf also starts 12 amino acids from the N-terminal. This is relevant because electron transfer takes place precisely at the heme active site of cytochrome c. A heme-stain of the SDS-PAGE gel containing the reduced membrane fraction did not reveal any bands (data not shown) but this could be due to the low abundance of this protein at late log-phase. Shared similarities to other Fe-oxidizing proteins and the putative localization to the outer membrane makes this cytochrome ca candidate for the first electron acceptor from Fe. 1. M . ferrooxydans PV-1 - cyc2-mf (EAU55163J 2. L. sp. Group 11 '5 way CG' -cyt572 (EOZ3951) 3. A. lerrooxydans-cyc2 (CAA07031) 1. M. ferrooxydans PV-1 - cyc2-mf (EAU55163) 2. l. sp. Group 11 '5 way CG' -cyt572 (EOZ3951) 3. A. lerrooxydans • cyc2 (CAA07031) 1. M. feo-ooxydans PV-1 - cyc2-mf (EAU55163) 2. L sp. Group 11 '5 way CG'· cyt572 (EOZ3951) 3. A. ferrooxydans - cyc2 (CAA07031) 1. M. ferrooxydans PV-1 - cyc2-mf (EAU55163) 2. L sp. Group 11 '5 way CG' -cyt572 (EOZ3951) 3. A. ferrooxydans - cyc2 (CAA07031) 1. M. ferrooxydans PV-1 - cyc2-mf (EAU55163) 2. L sp. Group 11 '5 way CG' -cyt572 (EOZ3951) 3. A. lerrooxydans • cyc2 (CAA07031) 1. M. ferrooxydans PV-1 . cyc2-mf (EAU55163) 2. L sp. Group 11 '5 way CG'. cyt572 (EOZ3951) 3. A. lerrooxydans • cyc2 (CAA07031) 1. M. ferrooxydans PV-1 - cyc2-mf (EAU55163] 2. L sp. Group 11 '5 way CG' - cyt572 (EDZ3951) 3. A ferrooxydans • cyc2 (CAA07031) 1. M. ferrooxydans PV-1 - cyc2-mf (EAU55163) 2. L sp. Group 11 '5 way CG' - cyt572 (EOZ3951) 3. A. ferrooxydans - cyc2 (CAA07031) 65 P F"i:VRSQlL FSG.AGP 11:DAAGQPVHPG■R--- - -- - - - - -FGMkDQA. TRF'.l:KGVQNGiL;QI\GEW S TNNK )',G,S}I V NGVN~!lVN SSl S KQ)llQKGTlJI. ~ G DY AGliGGW ANG VGAtlT PWO lESGG~ I NPWl $.FW VQPG Y f,LQI\S J:\TNVGG.1,Qi>,v FGS<"NSN ANAS l'IIN --- - -N VQl'PQQ VS L FYI\GE U'P Hl GS •·,t;HlJ"(SGGGS " 140 I i---, = I === == = === == = r, V■F■':J.GL:t;S SGS YON AF■1'1~DAN.QN G■G'1LE G G Y Y/iJ; IU,F KV RYG RVQN DVG YGMTMMS R RP!tlG f GA N■S■Gt.Dil..ViQFSNI AA I PGN FN DV ! AfT■LDR■ ,;L"W'G■DRA FNIINATY ~ YH DH YGI QGG- -- - ----YRN!IWGS■N)"GIJYTTTYTNSG SPDTSNEWIEAS■LPW ,.;■--- 480 490 500 510 520 530 540 . ., ,;.., . L . . '"'" i,, a===a...;, ------FLTITNSKNIVTSMGMNM!'■~STIVR----------------QQSrL■Tl:IAG:rD!llL DISLFAOYQIMINYGIAGTLSAFSA■sGGSYSSKTTYSDATTGAGGFFGNVEA■NFSVGrDF■'l - - -----TRISLRYVIYNKFNGVGS■SSNNUG--- - -- - -- ------YGASAYITLE[JLAWI■Y Figure 3-9. Amino acid aligrunent comparing cyc2-mfto two iron oxidases: cyc2 and cyt572. Alignment was produced in Geneious by using the CLUSTAL aligner. Signal peptides were removed. Phosphate transport: It has been well documented that in Escherichia coli, P seu domonas aeruginosa and Bacillus subtilis, expression of the phosphate-specific transport operon (pst) is induced when phosphate levels become low (Amemura et al., 1985; Surin et al., 1985; Nikata et al., 1996; Takemaru et al., 1996). Both gram-negative E. coli andP. aeruginosa have a pstSCAB-phoU operon, while gram-positive B. subtilis lacks phoU. The most abundant protein in the proteornic profile of PV-1 is annotated as a hypothetical protein that is encoded by a gene with locus tag SPVl _ 07114, which is the first gene of what we suggest is a putative pst operon (Figure 3-10). Amino-acid sequence analysis on CELLO and PSORTb predicted this protein to be located in the outer membrane. A protein signature search on InterProScan matched a signal peptide and a porin domain (IPR023614), adding support to the above prediction. Additionally, a BLASTP search indicated that this protein is quite unique and shares less than 31 % identity to other known porins, especially phosphate-selective porin O and P. The second most abundant protein in PV-1 is an ABC transporter phosphate-binding protein with predicted localization in the periplasm, after analysis on CELLO and PSORTb. Analysis on 66 Inter Pro classifies this protein within the PstS family of ABC-transporters (IPR005673 ). A blast search on Uniprot also indicates that there is homology to other PstS proteins. The other 4 genes downstream from pstS have domains that identify them in the following order as pstC, pstA, pstB and phoU. Therefore, these results suggest that SPVl _ 07114 is an additional component to the already known pst operon. The high abundance of SPY l _ 07114 and PstS may indicate that PV-1 was undergoing P limitation at the time of harvest, during the late-log phase. M. ferrooxydans SPV1 07114 pstS pstC pstA pstB phoU E. coli pstS pstC pstA pstB phoU B. subtilis pstS pstC pstA pstB pstB Figure 3-10. Phosphate-specific transport (pst) operon of M ferrooxydans in comparison to other known pst operons in E. coli and B. subtilis. 67 It is well known that sorption of inorganic phosphate (Pi) to pre-formed Fe(III) oxides can result in the removal of P, from solution (Yeoman et al., 1988). The amount of P, sorption increases when, instead of adding pre-formed Fe(III) oxides, Fe(II) is oxidized to Fe(III) oxyhydroxide in the presence of Pi (Mayer and Jarrell, 2000). Therefore, it is plausible that the pool of soluble P, (initial cone. of 542 µM) in the culture medium decreased as more Fe(II) was added daily and the products of biological iron oxidation accumulated with time. It is also reasonable to think that both marine and freshwater FeOB may encounter this problem in their natural habitat, since they are surrounded by iron oxides. But, in coastal marine sediments, a community of Fe-reducing bacteria (FeRB) can release significant amounts of P, as a result of remineralization of organic compounds, which can be beneficial to FeOB in terms of available P, and Fe(II) (Sundby et al., 1986; McManus et al., 1997; Noflke et al., 2012). Therefore, a closer look at P, uptake in FeOB in the presence of FeRB is warranted, in the laboratory and/or in the environment. In the absence of such constant sources of P,, obligate Fe(II) oxidizers such as PV-1, would need to use P, uptake mechanisms actively, for assimilation into a wide range of cellular material such as DNA, proteins and lipids. In agreement with the above, an acidophilic FeOB, Acidithiobacillus ferrooxidans, has been found to upregulate pstS when it is grown on Fe(II) and downregulate it when grown on sulfur (Ramirez et al., 2004). Carbon fixation: PV-1 has two different forms ofribulose-biphosphate carboxylase, or RuBisCO, the enzyme involved in the first step of the Calvin-Benson-Bassham (CBB) Cycle: Form IAq and Form II (Singer et al., 2011). The proteomic results indicate that 68 Form II is one of the most abundant proteins expressed in the large batch cultures, which in part is explained by the autotrophic lifestyle of PV-1 (Table 3-1 ). In contrast to this result, Form IAq was not identified in the proteomic profile. Activation proteins cbbO and cbbQ associated with Form II RuBisCO do show up in the profile but at lower abundance (Supplemental Table S3-l). In general, Form II is differentiated from Form IAq by having higher turnover numbers, lower affinity to CO, (g), and for being functional at lower concentrations ofO 2 (g)(Badger et al., 2008). Therefore, Form II is more prevalent in suboxic environments with high CO 2 (g) (> 1. 5 % ). The large bottles used for culturing PV-1 in this study had a headspace concentration of 30% CO, (g) and <2% 0 2 (g), which may explain the preference for Form II RuBisCO. Similar environments with high CO 2 (g) and low 0 2 (g) are seen in Lo'ihi Seamount, an underwater volcano near the Island of Hawai'i, where the summit is located in an oxygen minimum zone and the hydrothermal vents' fluids are the source of CO 2 and reduced Fe (Karl et al., 1988). It is possible that Form II is predominant in FeOB that colonize these CO, -rich deep sea environments. Other abundant proteins involved in carbon fixation were identified, including transketolase, phosphoglycerate kinase, glyceraldehyde-3- phosphate dehydrogenase, phosphoribulokinase and fructose-bisphosphate aldolase, indicating that PV-1 has a fully functional CBB cycle. Stalk Formation: The molecular mechanism for stalk formation is not yet known; however, there are two identified groups of genes in locus tags SPVl _ 00467 to SPVl _ 00517 and SPVl _ 07411 to SPVl _ 07616 that are particularly rich in permeases, 69 glycosyl transferases and capsular polysaccharide synthesis enzymes (Bennett et al., 2014). Some gene products from these two groups of genes are present in the proteomic profile; thus, these proteins could be good candidates for subsequent molecular approaches to studying stalk formation. It is also possible that stalk formation is a more complex process that includes proteins from the inner membrane, periplasm and outer membrane; therefore, other groups of proteins, including proteins associated with the electron transport chain, can potentially be targeted. Outer Membrane Proteins: In PV-1, there are two abundant proteins with similar structures to OmpA: SPV1_06074 and SPV1_00467 (Table S3-l). OmpAis a versatile integral outer membrane protein of high abundance in Escherichia coli, from which most of the knowledge about this protein comes from (Koebnik et al., 2000). It plays important roles in maintaining the outer membrane stability as well as in the formation ofbiofilm, among other functions (Orme et al., 2006; Smith et al., 2007). There are some authors that have proposed that OmpA can act as a porin in E. coli, but the debate is still ongoing (Smith et al., 2007). It is not clear what function(s) these OmpA-like proteins have in PV-1, but production ofbiofilm is evident within the first 24 hours of cultivation along the bottom of the bottles. Other hints can be provided by acidophilic FeOB, which express OmpA-like proteins abundantly as well. An OmpA-like protein, FopA, is one of three most abundant outer membrane proteins expressed in the extreme acidophilic FeOB, A. ferrooxidans (Castelle et al., 2008). A more focused study concluded that this OmpA-like protein is physically associated with peptidoglycan and suggests it can stabilize the outer membrane of A. ferrooxidans (Manchur et al., 2011 ). An investigation on proteins that are associated with extracellular polymeric substance (EPS) on biofilms developed in an acid mine drainage site revealed that an OmpA-like protein is within the twenty most abundant proteins (Jiao et al., 2011). The microbial community of these biofilms are known to be dominated by acidophilic iron-oxidizing Leptospirillum. CONCLUSION 70 The analysis of the proteome of an important marine FeOB, M. ferrooxydans, revealed new insights about its physiology that were not previously identified by analyzing the genome alone. Our results indicate that M. ferroxydans uses a periplasmic cytochrome cm for electron transport and in particular, an abundant terminal cbb,-type cytochrome oxidase. A molybdopterin-oxidoreductase complex may be functionally important to the organism based on its abundance and functionally different from other alternative complex-III seen in heterotrophic and phototrophic bacteria. An outer membrane cytochrome c, homologous to other Fe oxidases, is identified as a possible Fe oxidase. The profile also indicates that M. ferrooxydans has to deal with phosphate limitation probably due to the presence of Fe oxyhydroxides that bind Pi. 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Bioinformatics. 26(13):1608-1615. 78 CHAPTER IV: In-situ incubation of iron-sulfide mineral in seawater reveals colonization by iron-oxidizing Thiomicrospira and sheath-forming Zetaproteobacteria. ABSTRACT Sulfide mineral precipitation occurs at mid-ocean ridge (MOR) spreading centers, both in the form of plume particles and massive sulfide structures. A common constituent of MOR sulfide mineral is pyrrhotite (Fe 1 .xS). This mineral was chosen as a substrate for in situ incubation studies in the shallow waters of Catalina Islands, CA to investigate the colonization of iron-oxidizing bacteria. Gammaproteobacteria and Alphaproteobacteria largely dominate the bacterial community on pyrrhotite samples incubated in the water colunm. Pyrrhotite samples incubated at the sediment/water column interface show more even dominance by Gammaproteobacteria, Alphaproteobacteria, Deltaproteobacteria, specifically "cable bacteria" belonging to the family Desulfobulbaceae, and Bacteroidetes. Known iron-oxidizing bacteria such as M ariprofundus were identified and isolated from pyrrhotite samples incubated in the water column. An iron-oxidizing Thiomicrospira was enriched from pyrrhotite incubated in the surface sediment and subsequently isolated. This is the first time that any organism within the Thiomicrospira genus has been proven to grow autrotrophically on reduced iron. Additionally, the first 16S rRNA gene sequences specific to sheath-forming Zetaproteobacteria are presented. INTRODUCTION 79 The first Thiomicrospira isolate, T pelophila, was isolated from mud flats containing sulfides in an estuarine ecosystem in the Netherlands and was described as an obligate sulfur-oxidizer in 1972 (Kuenen and Veldkamp). Since then, all other autotrophic species in the Thiomicrospira genus have been associated only to sulfur oxidation and classified within the Gammaproteobacteria phylum with the known exception of T denitrificans which has been reclassified within the Epsilonproteobacteria and renamed Sulfurimonas denitrificans (Muyzer et al., 1995; Takai et al., 2006). The discovery of hydrothermal vents at the Galapagos Rift in 1977 led to increased sgnificant attention to the chemoautotrophic microbial communities inhabiting different marine habitats in the deep sea (Ballard, 1977; Corliss et al., 1979; Jannasch and Wirsen, 1979). In 1979, another expedition to the Galapagos Rift resulted in the identification of 12 strains of chemoautotrophic sulfur-oxidizing Thiomicrospira that were isolated from hydrothermal vent samples (Ruby et al., 1981). Subsequently, Thiomicrospira has been isolated from other hydrothermally active sites such as the East Pacific Rise and the Mid-Atlantic Ridge (Jannasch et al., 1985; Wirsen et al., 1998). Neutrophilic, autotrophic iron oxidizing Alphaproteobacteria and Gammaproteobacteria have also been isolated from the deep sea (Edwards et al., 2003a), but these pure cultures did not survive prolonged culturing in the laboratory. More recently, the neutrophilic, iron-oxidizing, chemolithoautotrophMariprofundusferrooxydans in the novel phylum of the Zetaproteobacteria was isolated from Loihi Seamount in the late 1990s, and its characterization published in 2007 (Emerson and Moyer, 1997; Emerson et al., 2007). This organism extrudes twisted-stalks containing iron oxyhydroxide. Edwards et al. 80 previously observed these morphological features in an in-situ incubated pyrrhotite sample from Juan de Fuca Ridge (2003b). The relationship of Thiomicrospira to a biological iron-oxidizing function was suggested, based on unpublished clone libraries of microbially colonized pyrrhotite from this site (Edwards et al., 2003b ). Here, we present the first report on an isolated putative novel, neutrophilic Thiomicrospira spp. with iron-oxidizing capabilities. The organism was isolated from bacterial traps containing pyrrhotite mineral that were exposed to shallow marine waters. The bacterial community that colonized these samples are also analyzed, giving insight into other possible iron-oxidizers. Additionally, a method is also presented to enrich for sheath-forming Zetaproteobacteria. Finally, environmental proteomic samples are analyzed giving us a closer look at the physiology and activity of the communities colonizing this mineral substrate as well as mild steel. MATERIALS AND METHODS Study site: In-situ enrichment experiments were conducted at Big Fisherman's Cove, in Santa Catalina Island, CA, USA (33.445124° N, 118.484147° W), a marine-protected area that is adjacent to the Wrigley Marine Science Center (WMSC). The experiments were deployed off the WMSC pier, facing south, in early September and retrieved in late December 2012. The water depth in the area near the pier ranged between 5 to 7 m. The salinity of the seawater ranged between 32 and 34 %0 and the surface seawater temperature ranged between 12.2 - 23.4 °C (buoy station 46222; data retrieved from http://www.ndbc.noaa.gov/). One pair of traps was deployed onto the sediment surface and another pair was suspended in the water column, 3 m above the sediment. 81 Microbial traps: Each trap set is composed of 2 chambers sealed with nylon mesh (190 µm opening)(McMaster-Carr, Santa Fe Springs, CA) using Gorilla Glue™ (The Gorilla Glue Company, Cincinatti, OH). Each chamber was constructed from 4" schedule (SCH) 40 acrylonitrile butadiene styrene (ABS) pipe and cut to 2" height with band saw. Each chamber contained 300 g ofpyrrhotite (Ward's Natural Science, San Luis Obispo, CA) wrapped in nylon mesh. Pyrrhotite was sliced into coupons by using the IsoMet® low speed saw (Buehler, Lake Bluff, IL). The coupons were around 2 cm in length x 1 cm in width x 2 mm in thickness and were polished and autoclaved. In addition to the mineral substrate, a layer of sterile glass wool(~ 2.5 g) was added above the wrapped pyrrhotite in order to prevent any other debris from making direct contact with it (in suspended traps, a layer of glass wool was also placed underneath the pyrrhotite ). Cable ties were used to secure a 5 lb coated lead weight to the sediment microbial trap. The weight was centered on top of the traps so that there was direct contact between sediment and the bottom of the traps. Polypropylene rope (1/4" x 50 ft) was tied to the weight and the trap set secured to the pier railing. For the suspended bacterial trap set, the 5 lb weight was used as an anchor, sitting on the sediment. Polypropylene rope was tied to the weight and the trap set was positioned 10 ft from it by using knots immediately below and above the trap set. The rope connecting the trap set was secured to the pier railing. Traps were 82 retrieved and placed in a container with cove seawater; thereby, minimizing the amount of seawater that can come out of the traps and preventing drying of samples. After retrieval, each trap set was transported to a laboratory at WMSC and processed for DNA and protein extractions. A few coupons were selected for enrichment purposes and stored on ice overnight using filter-sterilized cove seawater. Alternatively, a few coupons were processed for imaging purposes and fixed in 4% paraformaldehyde/lX phosphate buffered saline (PBS) solution at 4 °C. These samples were kept on ice and then stored at -20 °C in the mainland laboratory. Pyrrhotite coupons were handled by using autoclaved metal tweezers which were subsequently EtOH-cleaned and flame-sterilized at different steps throughout sample processing. DNA extraction: Four pyrrhotite coupons per trap (i.e. 4 for sediment trap and 4 for water colunm trap) were separated for DNA extraction at WMSC. The coupons were placed each in a sterile 15 mL tube, stored in dry ice, transported to the USC campus and stored at -80 °C until processing. Once thawed, 2 coupons were combined and processed for DNA extraction. DNA was extracted in a polypropylene 15 mL tube by using the FastPrep®-24 homogenizer equipped with a Teen-Prep adapter and by using components of the FastDNATM Spin Soil Kit following manufacturer's protocols (MP Biomedicals, Santa Ana, CA). Extracted DNA samples were stored at -80 °C until processing. For the seawater sample, 1 L of cove surface seawater was filtered on a 0.22 µm Whatman Nucleopore polycarbonate filter (GE Healthcare Life Sciences, Piscataway, NJ) supported by a 10 µm TCTP filter (EMD Millipore, Billerica, MA). The 0.22 µm filter 83 was then placed in a microcentrifuge tube, stored in dry-ice, transported to campus and stored at -80 °C until processing. DNA was extracted by using the FastDNA TM Spin Soil Kit. DNA concentrations were measured by Nanodrop 1000 (Thermo Fisher Scientific, Waltham, MA). PCR and cloning of J 6S rRNA and mobB genes: Small subunit ribosomal RNA (16S rRNA) genes present in the environmental samples were amplified by PCR on a Veriti® thermal cycler (Life Technologies, Grand Island, NY) as follows: 1 step of denaturation at 95 'C for 4 min; 30 cycles of denaturation, melting, and extension (95 'C for 30 sec, 55 °C for 30 sec, 72 °C for 90 sec respectively); 1 step of extension at 72 °C for 10 min; 1 final step of cooling at 4 'C for oo. Primers used were the universal 27F (5'- AGAGTTTGATCMTGGCTCAG-3') and 1492R (5'-TACGGHTACCTTGTTACGACTT- 3') for bacterial 16S rRNA (Lane, 1991). The amplicons were purified by using the QIAquick PCR Purification Kit (Qiagen Inc., Valencia, CA). The mobB gene was PCR amplified as follows: 1 step of denaturation at 95 'C for 4 min; 32 cycles of denaturation, melting, and extension (95 °C for 30 sec, 54 °C for 40 sec, 72 °C for 1 min respectively); 1 step of extension at 72 'C for 10 min; 1 final step of cooling at 4 'C for oo. For environmental sampes in specific, the purified amplicons served as template (1 µL) for an additional 10-cycle PCR run. Amplicons were again purified and used for cloning. Primers used for mobB amplification were 664F (5'-TCGTTTGGTGCGGATTTCCTG- 3') and 1123R (5'-GCAAAATCGTTTCACCGACATT-3'). MobB primers specific for Zetaproteobacteria were designed in Geneious by aligning and focusing on a semi- 84 conserved region of the mobB genes of Zetaproteobacteria and Betaproteobacteria. Designed primers were checked on Primer3Plus for melting temperature prediction and potential hairpin and dimer structures (Untergasser et al., 2012). Designed primers were also blasted in NCBI for specificity. DNA concentrations of purified amplicons were measured by Nanodrop 1000. The 16S rRNA and mobB genes were cloned by using the TOPO TA Cloning® kit (Life Technologies) and One Shot® TOPI0 chemically competent E. coli. Sanger sequencing of vector inserts were performed by Beckman Coulter (Danvers, MA). Community Analysis: Sequences obtained from 16S rRNAclone libraries were checked for quality, trimmed and assembled into contigs in Geneious version R6 (Biomatters, Auckland, New Zealand). The sequences were processed in Mothur version 1.28.0 (Schloss et al., 2009) to obtain rarefaction curves with OTUs defined at the 0.03 distance cutoff and to obtain abundance, evenness and diversity indeces. Taxonomic classification of sequences were mainly obtained from the SILVA database (Pruesse et al., 2012) by using the SILVA Incremental Aligner (SINA version 1.2.11). Alternatively, a few sequences were classified by using Classifier from the Ribosomal Database Project (RDP)(Wang et al., 2007). Chimeras were identified by U chime and ChimeraSlayer algorithms implemented from Mothur. Maximum-likelihood phylogenetic trees were obtained from Geneious based on 1000 bootstrap replications. Protein extraction: Coupons from the retrieved microbial traps were immersed in batches 85 of filter-sterilized 30 mL of 0.1 N NaOH/2% SDS in 50 mL sterile polypropylene tubes (i.e. approximately 30 coupons per tube; total of 16 tubes per trap set), vortexed for 5 minutes, stored in dry-ice, transported to the USC campus and stored at -80 °C until processing. After thawing samples on ice in the laboratory, the tubes were vortexed vigorously for 5 minutes and the mixtures (i.e. without coupons) were centrifuged at 10,000 x g for 10 minutes at 4 °C to precipitate any debris. The supernatants were transferred into Macrosep 3K concentrators (Pall, Ann Arbor, MI) with 3 kDa molecular weight cutoff (MWCO) membrane and centrifuged at 5,000 x g for 90 min at 4 °C. A concentrated crude protein extract with a yellow/slightly-green color is obtained. No protease inhibitors were used due to negative impact on identification of proteins. Concentrated protein extract was not buffer-exchanged to inhibit protease activity, which is kept at a minimum in 0.1 N NaOH/2% SDS solution. Excess SDS was removed by using SDS-OUT (Thermo Fisher Scientific, Rockford, IL) with no clarification step in order to minimize processing time and degradation of proteins. An aliquot was separated for subsequent measurement of protein concentration by using the Quick Start™ Bradford Assay (Bio-Rad, Hercules, CA), according to manufacturer's instructions using a UV-1601 spectrophotometer (Shimadzu Scientific Instruments, Carlsbad, CA). Samples were further concentrated by using the Nanosep 3K concentrators (Pall) according to manufacturer's instructions. Immediately after this step, samples were run in a gel or stored at -80 °C. Sodium dodecyl sulfate - polyacrylamide gel electrophoresis (SDS-PAGE): Proteins were 86 separated on gels according to standard protocols (Laennnli, 1970). Enviromental protein extracts were run on 12 % TGX™ polyacrylamide gels (Bio-Rad) under non-reduced conditions (i.e. no dithiothreitol (DTT) and no heating). A short-stack gel was run at 100 V for a short distance into the resolving gel (i.e. approximately 1.5 cm below the well). After electrophoresis, the gel was fixed in 50% HPLC-grade methanol/7% HPLC-grade acetic acid solution for 1 hr, stained with Bio-Safe Coomassie Stain (Bio-Rad) for 1 hr, destained with HPLC-grade water for 2 hours and visualized with a Gel Doc™ XR+ imaging system (Bio-Rad) equipped with a white light transilluminator. Images were analyzed with Image Lab™ software (Bio-Rad). The regions of the gel containing the proteins were excised with a clean sterile scalpel. The regions targeted were 10 kDa to <50 kDa and <50 kDa. Excised samples were each transferred into sterile microcentrifuge tubes, stored in dry ice and immediately submitted for proteomic analysis. Proteomic analysis: Excised protein samples were submitted to the University of California - Los Angeles (UCLA) - Molecular Instrumentation Center - Proteomic Center for trypsin-digestion and LC-MS/MS on a Thermo LTQ-Orbitrap XL mass spectrometer (San Jose, CA) equipped with an Eksigent (Dublin, CA) Nanoliquid Chromatography 1- D plus system. The resulting MS/MS spectra were searched against the proteomes of M. ferrooxydans strains PV-1 and M34 in the NCBI and JGI databases respectively. Additionally, the proteomes of all Proteobacteria as well as the non-redundant RefSeq amino acid database in NCBI were searched. Genomes were searched using the Matrix 87 Science MASCOT Daemon search engine (Boston, MA). The criteria for having a protein identified is: (1) there must be a minimum of two peptides matching to it that are ranked as number 1 and (2) they must have ion scores with p <0.05. The criteria for protein assignment to an organism is that at least one peptide (selected per the above criteria) must be best-matched to the organism. The mass spectrometry proteomics data will be deposited to the ProteomeXchange Consortium via the PRIDE partner repository. Enrichments: The medium was prepared per Bennett et al. (2014) using 100 mL serum bottles. The concentration of 0, (g) in heads pace was increased to 3 %. Biofilm present in a pyrrhotite coupon was scrapped off with a sterile needle in an empty 50 mL tube. The dislodged biofilm was mixed in a minimal volume (i.e. 0.2 mL) of sterile, artificial seawater (ASW) and injected into a serum bottle containing 100 mL of autoclaved ASW. Freshly-prepared and filter-sterilized Fe (II) was added after inoculation and daily thereafter. The coupon was handled by using a sterilized metal tweezer and by following aseptic techniques. Enrichment bottles were incubated statically in the dark, in a horizontal position and at room temperature. Microscopy: Scanning electron microscopy (SEM) samples were fixed in paraformaldehyde solution (0.8% final concentration), incubated at 4 °C for 1 hour and stored at -20 °C afterwards. They were dehydrated sequentially in 25%, 50%, 75%, 90% and 100% molecular grade ethanol with 10 minutes per dehydration step. They were critically-point-dried and gold coated. Images were taken with the scanning electron 88 microscope JSM-7001F-LV (JEOL, Peabody, MA) at the Center for Electron Microscopy and Microanalysis at USC. Alternatively, samples were imaged on a TCS SPE confocal microscope (Leica Mycrosystems, Buffalo Grove, IL) using a 488 nm solid-state laser for excitation and emission wavelenghts of 495-550 nm for SYTO 9 dye. Cell counts were performed as described by Emerson and Moyer (2002). Briefly, duplicate samples are fixed with paraformaldehyde solution (0.8% final cone.), stored at 4 'C for 1 hour and then frozen at -20 °C until counted. Slides printed with 4 mm diameter circles (Electron Microscopy Sciences, Fort Washington, PA) are coated with 1 % agarose solution and allowed to cool. A sample volume of 4 µL (2 µL of resuspended culture sample mixed with 2 µL of 1 mM propidium iodide solution (Life Technologies)) is loaded within the boundaries of the circle and allowed to dry in the dark. At least fifty fields per circle were counted at 1 00X magnification using an epifluorescent Axiostar Plus microscope equipped with an HBO 50 mercury lamp and Cy3 filter for green light excitation. X-ray diffraction: The biomineral produced by the iron-oxidizing Thiomicrospira isolate was characterized at the Los Angeles National History Museum by X-ray diffraction (XRD). Data were recorded using a R-Axis Rapid II (Rigaku, The Woodlands, TX) curved imaging plate microdiffractometer with monochromatized MoKa radiation. Observed d spacings and intensities were derived by profile fitting using JADE 2010 software (Materials Data Inc., Livermore, CA). RESULTS 89 Clone libraries: The clone library recovered from pyrrhotite samples incubated in the water column trap reveals that the bacterial community is co-dominated by Gammaproteobacteria ( 44.0%) and Alphaproteobacteria (34.5%) (Figure 4-IA). Other groups that were identified at relatively lower percentages included the phyla Planctomycetes (4.8%), Bacteroidetes (4.8%), and the classes Zetaproteobacteria (4.8%), Deltaproteobacteria (3.6%), Epsilonproteobacteria (1.2%) and Actinobacteria (1.2%). Within the Gammaproteobacteria (37 clones), the most abundant phylotype, clone 15 _ D09 (22 clones) was unclassified at order or lower taxonomic ranks (OTU defined at the similarity cut-off value of97%). Three relatively new genera were identified, Kangiella, Thiohalophilus andArenicella. Kangiella species so far described are facultative anaerobic, heterotrophic marine organisms (Yoon et al., 2004). The only described species in the Thiohalophilus genus is a known chemolithoautotrophic sulfur oxidizer: T thiocyanatoxydans (Sorokin, et al., 2007). The two Arenicella species so far described are aerobic, and heterotrophic (Romanenko, et al., 2010; Nedashkovskaya, et al., 2013). Within the Alphaproteobacteria class (29 clones), the family Rhodospirillaceae represents close to half of the clones identified (13 clones). Within the Rhodospirillaceae, the majority of the clones were classified as Magnetospira (11 clones). The clone 15 _ A04 (5 clones; OTU defined at the similarity cutoff value of97%) belonging to this group is overall the second most abundant phylotype in the whole library. M agnetospira is another recently proposed novel genus that is distantly related to M agnetospirillum (Williams et 90 al., 2012). It has the ability to grow autotrophically on reduced sulfur and heterotrophically on other substrates. The phylogenetic tree in Figure 4-2 indicates that these clones are also closely related to Magnetovibrio, a more recently isolated magnetotactic bacterium, which is not yet present in the Silva or RDP databases (Bazylinski et al., 2013). The other classes of Proteobacteria were all represented, with the exception of the Betaproteobacteria. In the case of the Zetaproteobacteria, the chemolithoautotrophic, iron-oxidizing M ariprofundus is overall the third most abundant phylotype ( 4 clones; OTU defined at the similarity cutoff value of 97%) in the whole library. The bacterial traps that were in direct contact with the surface sediment reveal that the bacterial community is also dominated by Gammaproteobacteria (36.9% )(Figure 4-lB). Other major groups that were identified include the Deltaproteobacteria (19.0%), Alphaproteobacteria (14.3%) and Bacteroidetes (14.3%). In comparison with the traps in the water colunm, the Alphaproteobacteria were less abundant and the Deltaproteobacteria and Bacteroidetes gained more representation. Zetaproteobacteria and Betaproteobacteria were not recovered. In common with the pyrrhotite water column traps, the Epsilonproteobacteria, Planctomycetes and Actinobacteria appear to be present in low percentages. Within the Gammaproteobacteria (29 clones), the order Insertae Sedis, JTB255 Marine Benthic Group, and the family Piscirickettsiaceae emerge as dominant taxa. Five clones 91 cannot be classified into lower taxonomic ranks below phylum level. Within the Deltaproteobacteria (16 clones), the family Desulfobulbaceae (including genus Desulfobulbus) represents the most abundant taxon with 10 clones. The Desulfobulbaceae is also the most abundant lineage (11. 8%) in the whole library of clones originating from bacterial traps in close contact with sediment. The identified Desulfobulbaceae shares <91 % similarity to other known and isolated Desulfobulbus. The representative clones in this study share 95-97% similarity to environmental filamentous Desulfobulbaceae clones from sediment of Aarhus, Denmark (Pfeffer et al., 2012). Also known as "cable bacteria", these organisms have been shown to oxidize sulfide in the sediment to reduce oxygen at the interface of water column/sediment. In comparison with the pyrrhotite traps that were incubated in the water column, the bacterial community belonging to the Bacteroidetes phylum tripled in abundance with the added representation of the family Cryomorphaceae and other members of the order Sphingobacteriales. The clone library obtained from Fisherman's Cove surface seawater sample (Figure 4- 1 C) reveals that the bacterial community is dominated by the phyla Actinobacteria (27.4%) and Alphaproteobacteria (26.2%) with Bacteroidetes (17.9%) and Cyanobacteria (16.7%) also constituting large groups. The most abundant phylotype is seen in the Actinobacteria phylum, specifically the OCS155 Marine Group of the order Acidimicrobiales, a commonly found taxa in surface seawater (Rusch et al., 2007). Other ubiquitous taxa include SARll, Synechococcus, and members of the families 92 Rhodobacteraceae, and Flavobacteriaceae. In contrast with the pyrrhotite samples, the Gammaproteobacteria was represented in low percentages (3.6%) and by different taxa such as Vibrio and members of the family Alteromonadaceae. Also in contrast to the pyrrhotite samples, the Betaproteobacteria was identified and represented by members of the Methylophilaceae family. Altogether, these results indicate that the bacterial consortia seen in Fisherman's Cove seawater is distinct from the bacterial consortia colonizing the pyrrhotite traps. Another 16S rRNA gene clone library was obtained preliminarily from mild steel that was exposed to the marine surface sediment for one month (Supplemental Figures S4-l,S4-2 and Supplemental Table S4-l). A) Pyrrhotite (suspended trap) Unclassified Gammaproteobacteria (29/37) lncertae Sedis (5/37) Kangiella (1 /37) Acidimicrobiales (1/1) Helicobacteriaceae (1 /1) Desulfobacterales (2/3) Flammeovirgaceae (2/4) Haliscomenobacter (214) Phycisphaeraceae (2/4) Mariprofundus (4/4) Sneathiella (2/29) Phyllobacteriaceae (2/29) 93 ■ Gammaproteobacteria (44.0%) ■Alphaproteobacteria (34.5%) ■Zetaproteobacteria (4.8%) ■ Planctomycetes (4.8%) ■ Bacteroidetes (4.8%) ■ Deltaproteobacteria (3.6%) ■ Unclassified Bacteria (1 .2%) ■ Epsilonproteobacteria (1 .2%) Actinobacteria (1 .2%) Chromatiales (1 /37) Oceanospirillales; Other (1/37) Rhodospirillaceae;Other (2/29) Rhodobacteraceae (5/29) B) Pyrrhotite (sediment trap) Unclassified Gammaproteobacteria (5/29) Piscirickettsiaceae (4/29) Chromatiales (2/29) Kangie//a(1/29) Thiotrichaceae (1 /29) Desulfobulbaceae (10/ 16) C) Seawater Desulfuromonadales (1 /1) OCS 155 Marine Group (22/23) Sva0996 Marine Group (1/23) Rhodobacteraceae (15/22) Phycisphaera (2/2) Su/furimonas (2/2) 155 Marine Group (4/4) acteroidetes;Other (2/12) Cryomorphaceae (2/12) Flavobacteriaceae (4/12) Sphingobacteriales (4/12) Magnetospira (1 /12) Parvularcu/a (2/12) Rhodobacteraceae (3/12) Alphaproteobacteria;Other (6/12) Verrucomicrobiaceae (2/2) Methylophilaceae (3/3) Alteromonadaceae ( 1 /3) Vibrio (1 /3) Oceanospirillales (1 /3) Synechococcus (14/14) NS9 Marine Group (1 /15) ■ Gammaproteobacteria (36.9%) ■ Deltaproteobacteria (19.0%) ■Alphaproteobacteria (14.3%) ■ Bacteroidetes (14.3%) Actinobacteria (4.8%) ■ Epsilonproteobacteria (2.4%) ■ Planctomycetes (2.4 % ) ■ Gemmatimonadetes (1 .2%) Chlorobi (1 .2%) ■Cand idate division WS3 (1 .2%) ■ Candidate division 0D1 (1 .2%) ■ Acidobacteria ( 1 .2%) Actinobacteria (27.4%) ■Alphaproteobacteria (26.2%) ■ Bacteroidetes (17.9%) ■Cyanobacter ia (16.7%) ■ Gammaproteobacteria (3.6%) ■ Betaproteobacteria (3.6%) ■Verrucomicrob ia (2.4%) ■ Unclassified Bacteria (1 .2%) ■ Deltaproteobacteria (1 .2%) Figure 4-1. Relative abundances of bacterial communities in pyrrhotite traps that were in-situ incubated in the water column (A) or resting on the sediment (B). The relative abundances of the bacterial community in Fisherman's Cove seawater is also shown (C). The abundances are color coded at the class level for Proteobacteria and phylum level for non-Proteobacteria. Taxonomic affiliation is based on full-length 16S rRNA gene sequence matches to sequences in the SIL VA database (similarity cutoff value of 97% ). 94 - Pyrrhotite/Sediment ~----- Vibrio fischeristrain ATCC 25918 - Pyrrhotite/Suspended Thiomicrospira crunogena XCL-2 l' Shared Thioafkalimicrobium cyclicum ALM 1 12_C05 Piscirickettsiaceae Uncultured bacterium clone 3M23_072, inactive sulfide chimney (JQ287101) Uncultured deep-sea bacterium clone 263-25, sediment near vent (FN554105) 100 Uncultured deep-sea bacterium BD3-6 (AB015548) I Uncu ltured deep-sea bacterium clone JTB255, cold-seep (JTB255) Marine Benthic Group Uncultured bacterium clone ARTE12_202, marine sediment (GU230336) JTB255 100 12_E10 Ectothiorhodospira mobifis I y-Proteobacteria 96 99 Thioalkalivibrio thiocyanodenitrificans ARhD 1 Ectothiorhodospiraceae Thioa/kalivibrio sulfidophi/us HL-EbGR? 15_009 Uncultured bacterium clone 2A3, sediment in brackish lake (HQ003527) Uncultured deep-sea bacterium clone crust20, basalt borehole (GQ903353) Uncultured deep-sea bacterium clone Ba2, sulfide chimney (FJ640811) 12_G05 l' Uncultured deep-sea bacterium P9X2b2C11 , seafloor lavas (EU491253) Uncultured marine sediment bacterium clone 88S-10 (EU652539) Uncultured marine sediment bacterium clone 1_76 (KC009910) Mariprofundus sp. GSB2 Mariprofundus ferrooxydans PV-1 15_F08 Uncu ltured deep-sea bacterium clone 3M34_066, inactive sulfide chimney (JQ287471) Uncultured deep-sea bacterium clone RESET_221C01 , sediment trap near vent (JN874291) ,---- Sneathiella chinensis CBMAI 737 Uncultured deep-sea bacterium clone SCB135 (JX227607) ~--- Magnetospirillum magneticum AMB-1 ~--- Magnetospira thiophila strain MMS-1 Magnetovibrio blakemorei MV-1 Rhodospirillaceae 15_A04 Uncultured deep-sea bacterium clone VS_CL-282, basaltic rock (FJ497534) Uncultured deep-sea bacterium clone VS_CL-279, basaltic rock (FJ497531) 100 ~--- Oesulfobulbus propionicus DSM 2032 .------1~--- Desulfobulbus sp. 1S6 SO Uncultured marine sediment bacterium clone Fe_B_ 110, methane seep (GQ356927) 12_C10 T ~---- Desu/furomusa ferrireducens strain 102 0,06 a-Proteobacteria I '-'""''°'""'"' Figure 4-2. Maximum-likelihood 16S rDNAphylogenetic tree (1000 bootstraps) ofrepresentative clones recovered from pyrrhotite (OTU defined at 97% similarity cutoff-value). Rarefaction curves indicate that the in-situ-incubated pyrrhotite samples are more diverse than the surrounding seawater (Figure 4-3). The pyrrhotite samples incubated at the water column/sediment interface display a slightly higher diversity than the pyrrhotite samples incubated in the water-column. Shannon's diversity index values are consistent with this trend (Table 4-1 ). The species richness estimators ACE and Chao 1 both estimate that the bacterial communities of the pyrrhotite samples in the water/sediment interface and in the water-column are statistically different from those of the surrounding seawater. The evenness index indicates that the pyrrhotite samples at the surface sediment have a more even distribution than the pyrrhotite samples at the water-column. Since diversity takes into account both richness and evenness, the pyrrhotite samples at the surface sediment 95 are the most diverse. 70 -- Pyrrhotite - Sediment 60 -- Pyrrhotite - Water Column -- Catalina Seawater 50 "' ::::> E-< 40 0 'c5 [> .r, 30 § z 20 10 10 20 30 40 50 60 70 80 90 Number of clones Figure 4-3. Rarefaction curve of species richness (similarity cutoff value of97% for OTU designation). The bars indicate 95% confidence interval. Table 4-1. Species richness estimators and diversity indices of environmental samples. Sample Pyrrhotite - Sediment Pyrrhotite - Water Column Seawater Number of Sequences 84 84 84 Observed OTUs 63 50 30 ACE (95% C.I.) 219 (196-242) 428 (308-547) 120 (98-141) Chaol (95% C.I.) Shannon Index Evenness 186 (166-206) 4.02 0.97 221 (185-258) 3.35 0.86 64 (54-74) 2.79 0.82 Enrichments and isolation of Thiomicrospira: Several enrichment cultures were obtained from the rusty biofilm developed on pyrrhotite. A series of dilution to extinction procedures with these enrichment cultures allowed for the isolation of an iron-oxidizing Thiomicrospira spp. from a sample of surface sediment pyrrhotite (Figure 4-4). This organism, Thiomicrospira SC-1, forms rusty round spots (1-2 mm diameter) on the walls 96 of the culture bottle when it is sufficiently diluted. The iron oxides associated with this organism are predominantly globular in shape (Figure 4-4A). The vibrio-shaped cells, which are typically 1 µm in length, are usually seen in direct contact with the iron oxides and also have the ability to form appendages for attachment (Figure 4-4B). X-ray diffraction analysis reveals that the iron oxide phase that accumulates in the culture is fairly crystalline with peaks matching to the mineral lepidocrocite (Figure 4-5). Figure 4-4. A. Thiomicrospira SC-1 in direct contact with bulbous iron oxides. B. Cellular structures connecting the cells to the bulbous iron oxides are typically seen. Lepidocrocite • FeO(OH) w c 35 - 30 25 i5 20 8 .:=- "iii C 2 C - 15 - 10 5- ---- _______ , ____ Jj ___ J __ 1 L . t1 5 ro m ~ ~ w Two-Theta (deg) Figure 4-5. X-ray diffraction pattern of the iron oxide phase that accumulates in a culture of Thiomicrospira SC-1. A reference pattern of lepidocrocite is included for comparison. ·-- ---- 97 The 16S rRNA gene of Thiomicrospira SC- I was analyzed and found to have a pairwise % identity of97.6% to both of its closest isolated relatives, the chemolithoautotrophic Thiomicrospira crunogena, an organism originally isolated from a hydrothermal vent field in the East Pacific Rise (Jannasch et al., 1985) and Thiomicrospira strain MA-3 (a T crunogena strain), an organism isolated from a Mid-Atlantic-Ridge hydrothermal vent system (Wirsen et al., 1998). The result of the maximum-likelihood phylogenetic tree corroborates the close relationship to the strains of T crunogena (Figure 4-6). It also indicates that strain SC-1 forms a clade that is separate from T crunogena and other isolated species. .------------------ Aquifex pyrophilus Thiomicrospira thermophila 178 71 Thiomicrospira L-12 Thiomicrospira crunogena ATCC35932 2 69 Thiomicrospira MA-3 00 Thiomicrospira crunogena XCL-2 78 Thiomicrospira SC-1 Thiomicrospira kuenenii I B-A 1 97 Thiomicrospira sp. Milos-T1 Thiomicrospira frisia JB-A2 Thiomicrospira chilensis CH-1 100 Thiomicrospira pelophila 99 Thiomicrospira thyasirae Thioalkalimicrobium cyclicum ALM1 0.04 Figure 4-6. Maximum-likelihood phylogenetic tree based on an alignment of 16S rRNA gene sequences from isolated Thiomicrospira species ( 1000 bootstraps). 98 In order to confirm that strain SC-1 is an iron-oxidizing organism, growth patterns were observed in cultures that were amended with Fe 2 + and with no Fe 2 + under microaerobic conditions (Figure 4-7). The results show that the presence of Fe 2 + in the medium causes strain SC- I to grow exponentially by approximately two orders of magnitude over a span of 7 days with a doubling time of 12 hours. The cultures with no added Fe 2 + do not grow exponentially, and the overall cell count numbers remain stable over the same period of time. These results indicate that strain SC-1 is able to grow chemoautotrophically with 99 -noFe -+- Fe l.00E+05 ---+- ,-----------------~ 0 20 40 60 80 100 120 140 160 Time (hours) Figure 4-7. Growth curves of Thiomicrospira SC-1 in artificial seawater in the presence and absence of reduced Fe. Error bars indicate one standard deviation from the mean. The batch of enrichments originating from the same pyrrhotite sample (i.e. bacterial trap in sediment), resulted in an enrichment of sheath-forming Zetaproteobacteria as well. This is the first time that a sheath-forming Zetaproteobacteria was successfully grown in the laboratory. This type of organism is characterized by forming long filaments (> 100 µm) and by forming large cream-colored spots on the wall of culture bottles (Figure 4-8). These enrichments survived only a few rounds of dilution-to-extinction procedures. The cells and the filament structures could not be preserved at room temperature nor at -80 °C. The fragility of the filament is documented in images that were captured on a confocal microscope (Figure 4-9). The rapid collapse of the filament appears to be caused by exposure to the heat of the laser of the confocal microscope. The collapse of the filament reveals a chain of cells which are rod-shaped and about 3 µm in length. The 100 released cells rapidly lyse and lose the rod shape, subsequently exhibiting a permanently round-shape. Analysis of the cloned 16S rRNA genes from this enrichment indicate that the culture contained strains that had between 94.0-94.3% pairwise identity to Mjerrooxydans PV-1, its closest relative in pure culture. The clones also shared between 97.9-98.2% pairwise identity to an environmental clone originally obtained from a surface iron mat that was microscopically observed to exhibit sheath-formation from Pele's Pit at Loihi Seamount (Accession Number JX468894 )(Fleming et al., 2013 ). The 16S rRNA phylogenetic tree shown in Figure 4-10 indicates that it is closely related to another environmental clone from the Lohiau hydrothermal vent in Loihi Seamount and to the Zetaproteobacteria OTU #6 (Accession Number JF320713)(McAllister et al., 2011 ). The enrichment clones identified in this manuscript share between 99. 5% and 99.9% pairwise identity to these latter environmental clones. Figure 4-8. Sheath-forming Zetaproteobacteria enriched from a bacterial trap containing pyrrhotite mineral. 101 Figure 4-9. The structure of this particular sheath-Zetaproteobacteria is heat sensitive. The laser from a confocal microscope decreases the stability of the sheath structure and reveals rod-shaped cells encased within it. A few more seconds of laser exposure lyse the cells, resulting in round-shaped cells. Scale bar= 6 µm. ------[================--;,~;;;;;;;:;;:;,;;;;--- Aquifex pyrophilus I Thermotoga maritima 100 100 ~---------- Nitratiruptor tergarcus ---~-'-----<L ___ _! 1_2<00 !..___j===========--;;, Su/furovum fithotrophicum Su!furimonas autotrophica 100 ------- Hyphomonasjannaschiana 97 -------""-'--i 100 Rhodobacterg/uconicum 72 Roseobacter denitrificans ~-------------- Thiomicrospira psychrophila ~------------ Marinobacter aquaeolei 100 [===-=-=-=--=-~--~ Gi:z:~~~e;;~rr~:~~ophicus L_ ____ _,,1 0"'0""1-- Le:e~::t~~;:~~::a J2_ 482_8S7 _clone_C08 (JQ287656) J2-481_ BS4_clone_C07 (JQ287647) J2_ 482_8S7 _clone_G05 (JQ287653) UNH_clone_95 (JF320785) J2_ 481 _BS4_clone_ D08 (JQ287649) Poh_clone_27 (JF320718) PVB_OTU_ 4_clone_PVB_ 13 (U15116) J2_ 481 _8S4_clone_F06 (JX468894) Sheath-Zetaproteobacteria enrichment clone 2 Sheath-Zetaproteobacteria enrichment clone 4 Zeta Sheath-Zetaproteobacteria enrichment clone 5 OTU 6 Loh_clone_ 49 (JF3207 13) Sheath-Zetaproteobacteria enrichment clone 1 Sheath-Zetaproteobacteria enrichment clone 3 Mariprofundus sp. GSB2 Mariprafundus ferrooxydans PV-1 Mariprofundus ferrooxydans M34 Mariprofundus SC-1 UNH_clone_ 122 (JF320787) Poh_clone_9 (JF320714) UNB_clone_ 44 (JF261518) J2_ 482_8S7 _clone_B02 (JQ287657) L ______ _111 00 ~=========~~: Desuffurella acetivorans Hippea maritima 0.04 Epsilonproteobacteria Alphaproteabacteria Gammaproteobacteria Betaproteobacteria Zetaproteobacteria I Deltaproteobacteria Figure 4-10. Maximum-likelihood phylogenetic tree of aligned 16S rRNA genes ( 1000 bootstraps). The 16S rRNA gene sequences of the sheath-Zetaproteobacteria enriched from pyrrhotite mineral and described in this manuscript are shown in red. Environmental 16S rRNA gene sequences from Zetaproteobacteria (not necessarily sheath-Zetaproteobacteria) obtained from iron-mats that were microscopically observed to exhibit sheath-formation are in blue (Fleming et al., 2013). 102 Another batch of enrichments originated from pyrrhotite that was incubated in the water column. These enrichments resulted in the isolation of Mariprofundus SC- I. An analysis of the 16S rRNA gene of this strain indicates that it shares 99.3%, 99.2% and 97% pairwise identity to Mferrooxydans M34,Mferrooxydans PV-1, andM GSB2 respectively. The cells have curved-rod shapes and are typically between 1-2 µmin length. They produce twisted-stalks similar to the other strains of M ferrooxydans (Figure 4-11). Phylogenetic analysis using 16S rRNAgenes corroborates the close relatedness to M ferrooxydans (Figure 4-10). This strain also contained the gene that encodes for a molybdopterin oxidoreductase, mobB, sharing 95.4% pairwise identity over 417 bases with mobB fromMferrooxydans PV-1 (Figure 4-12). Figure 4-11. Epifluorescent/phase contrast microscopy merged image of Mariprofundus strain SC-1. Inset bar = 6 µm. Larger image bar = 12 µm. Consensus Identity 1. rnobB - Mariprofundus ferrooxydans PV-1 - pos 664-1123 2. rnobB - Mariprofundus SC-1 - pos 664-1123 Consensus Identity 1.mobB MariprofundusferrooxydansPV-1-pos664-1123 2.mobB MariprofundusSC-7-pos664-1123 Consensus Identity 1.mobB MariprofundusferrooxydansPV-1-pos664-1123 2.mobB MariprofundusSC-1-pos664-1123 Consensus Identity 1.rnobB MariprofundusferrooxydansPV-1-pos664-1123 2.mobB MariprofundusSC-7-pos664-1123 Consensus Identity 1. mobB Mariprofundus ferrooxydans PV-1 - pos 664-1123 2.rnobB MariprofundusSC-1-pos664-1123 103 GGCACATGGAT GT CACCGGT GCAGT TT TCC CCCAAT AT GC GAGT T CCGCAAT GCACCGCGCGGT AC ACT GGT GCAGATT GAG T GGCACATGGAT GT CACCGGT GCAGT TT TCC GCCCAAT AT GC~GAGT T CCGCAAT GCACCGCGCGGT AC ACT GGT GCAGA TT GAG A "' P ~ , r~ G-,. 'BG " . T _ .. "'_ "' CCGAAGATGAC GCGGAT CGCT GGATTCCT GCACGT C CGGGT ACC GAAGG GCAT CT GGCT CTGGCACT G CCGAAGAT GAC GCTGACCGGCGC CAAIIGCGGAT CGCT GGA TTCCT GCACGT C CGGGT ACC GAAGG' CAT CT GGCT CTGGCACT G GCCT CACT GCT T GT ACAAAAAT CT GAAT AT GCCGAC AGGGTTCC CGCAGAT GT GGTT GCAT C CCT GAAGGAT GT ~ AACGTT GAC GCCT CACT GCT T GT ACAAAAAT CT GAA TAT GCCGAC AGGGTTCC CGCAGA T GT GGTT GCA TC CCT GAAGGAT GT AACGTT GAC GAAGT &jGCGAAGCTGT GT GAT AT CCCGGTT GAIICGAAT D Figure 4-12. Alignment of molybdopterin oxidoreductase (mobB) DNA sequences from M. strain SC-1 and M. ferrooxydans PV-1. The resulting amino acid sequences are located below the DNA sequences. Proteomic analysis of in-situ incubations: A bulk of the pyrrhotite samples that were in- situ incubated were processed for proteomic extractions. The focus was on FeOB, and more specifically, on samples where the Zetaproteobacteria were identified by clone libraries; therefore, the pyrrhotite samples in water column traps were selected for subsequent proteomic analysis. In addition to this, mild steel coupons that were in-situ incubated for 2 weeks in the water column were analyzed as well as background seawater from Fisherman's cove. The results show that 16 proteins were identified in pyrrhotite, 72 in mild steel, and 19 in seawater with best match hits represented by at least 2 peptides. There were approximately a similar number of proteins from each sample that were best- matched but only by 1 high-scoring peptide (p<0.05 for peptide and best match in 104 database for a given organism)( data not shown). Shared proteins among different samples were low in number indicating that the extracts originated from indigenous bacterial communities (Figure 4-13). This trend is extended when the proteins with only I high scoring peptide are included ( data not shown). Detailed lists of identified proteins are included in Supplementary Tables S2-4. - mild steel - pyrrhotite - seawater 68 Figure 4-13. Venn diagram of abundant proteins identified in mild steel (green), pyrrhotite (red), and surface seawater (blue). The NCBI Proteobacteria database was used for identification of proteins in this figure. The most abundant proteins in pyrrhotite have functions associated with transcriptional regulation (Gammaproteobacteria), carbon dioxide concentrating mechanism 105 (carboxysome shell protein)(Gammaproteobacteria), outer membrane porin (Alphaproteobacteria), carbohydrate transport and metabolism (Betaproteobacteria), and energy production and conversion (Betaproteobacteria). In mild steel, the most abundant proteins have functions associated with transcriptional regulation (Gammaproteobacteria), energy production and conversion (Epsilonproteobacteria), outer membrane porin (Alphaproteobacteria), carbon fixation via Calvin cycle (Betaproteobacteria), carbon fixation via reductive tricarboxylic acid cycle (Epsilonproteobacteria), chemotaxis (Gammaproteobacteria), antioxidant activity (Epsilonproteobacteria) and phosphate transport (Gammaproteobacteria). Epsilonproteobacteria are known to have cytoplasmic and periplasmic NiFe hydrogenases (Nakagawa et al., 2007; Sievert et al., 2008). The periplasmic hydrogenase from S. denitrificans that is identified in this study is the large subunit (HydB) that directly removes electrons from H 2 to eventually donate them to the quinone pool (Sievert et al., 2008). The identified proteins that are diagnostic of a functional reductive tricarboxylic acid (rTCA) cycle include the key enzymes pyruvate ferredoxin/flavodoxin oxidoreductase and 2-oxoglutarate ferredoxin oxidoreductase (Hiigler et al., 2005). These rTCA cycle proteins in the environmental samples were assigned to two genera within the Epsilonproteobacteria: Sulfurimonas and Wollinella. Carbon fixation proteins associated with the Calvin cycle were also identified predominantly from Betaproteobacteria (Ralstonia and Nitrosospira). Numerous peptide hits to chemotaxis proteins were recovered from Gammaproteobacteria (Vibrio), Alphaproteobacteria (Rhizobium) and Deltaproteobacteria (Geobacter) indicating a very active sensory response within the 106 community in mild steel. It is important to note that these results are matched to what is available in databases, which is currently limited in the number of available genomes especially from environmentally relevant strains; therefore, species assignments to certain proteins should be taken into account with available 16S rRNA clone libraries. DISCUSSION An initial report by McBeth et al. (2011) indicated that Zetaproteobacteria are involved in microbial induced corrosion (MIC) of mild steel in shallow marine waters, and therein confirming that marine neutrophilic FeOB could be successfully enriched from such sites, instead of only from deep-sea hydrothermal vent sites. Based on these results, a set of preliminary in-situ incubations were designed in order to investigate the microbial weathering of basaltic rock and minerals usually found in the ocean crust by exposing them to shallow waters of California ( data not shown). The preliminary results indicated that among the bacterial traps with basalt, basalt with olivine, pyrite and pyrrhotite, the latter was the most altered after a two-week exposure to seawater, having spots of localized weathering. Based on these preliminary results, the iron-sulfide mineral pyrrhotite was adopted for a subsequent three-month in-situ incubation, with the goals of performing an in-depth analysis of its bacterial community, enriching for FeOB and ultimately generating hypotheses applicable to the deep-sea. The bacterial traps containing substantially more pyrrhotite were suspended in the water column for a total of three months to increase the weathering of the mineral and the development ofbiofilm on its surface. Traps were also set on the sediment/water column interface for the same period of time. 107 FeOB in Water-Column Pyrrhotite Samples: Analysis of the 16S rRNA gene clone library indicates that clone 15 _F08, classified as Mariprofundus, represents the third most abundant phylotype in the whole library ( 4 clones). This phylotype was the only one identified from pyrrhotite in this study. Mariprofundus strain SC-1 was isolated from a pyrrhotite sample originating from a bacterial trap that was suspended in the water colunm. Growth in this enrichment culture containing SC-1 was evident within 24 hours of inoculation by the production of abundant fluffy yellow/orange iron-oxides. This is relatively fast growth, taking into account the 12 hr doubling time of Mariprofundus ferrooxydans PV-1 (Emerson et al., 2007). Comparison of the 16S rRNA gene sequences reveals that M ariprofundus strain SC-1 and clone 15 _F08 share pairwise % identity of 92%. McBeth et al. (2011) suggested that mild steel was colonized by single phylotypes per sample which is also seen with pyrrhotite in this study. However, analysis of the Zetaproteobacteria clones recovered from mild steel (Supplemental Figure S4-3) in our study reveal three different phylotypes (MS_ Hl0; MS_ 127; MS_ 120), indicating that at least mild steel can host different phylotypes of Zetaproteobacteria. FeOB in Surface Sediment Pyrrhotite Samples: It is not clear what factors play a role in the apparent absence of Mariprofundus in the surface sediment pyrrhotite but one of them could be the absence of oxygen. In the water-column bacterial trap, seawater with plenty 108 of oxygen diffuses from both the top and bottom sections of the trap that are sealed in mesh. The bacterial trap located on the surface sediment (with 5 lb weight on top) only had the top surface available for oxygen diffusion. Therefore, oxygen may be more limited in the sediment bacterial trap. Another explanation is that there was a possible succession of the microbial community within the three months of in-situ incubation. In support of this explanation, an SEM image taken from a preliminary two-week in-situ incubation at the sediment/water colunm interface shows twisted-stalk biosignatures associated with Zetaproteobacteria, in an area of mineral alteration (Supplemental Figure S4-4). The clone library results identify members of the Piscirickettsiaceae family, which Thiomicrospira is a member of. Since pyrrhotite is composed of both reduce sulfur and reduced iron, it is possible that certain members of the Piscirickettsiaceae family can do both iron and sulfur oxidation. By oxidizing the sulfur, the microorganism will have more access to the reduced iron that is entrapped in the crystal lattice of the iron-sulfide mineral. This iron can be biologically or chemically oxidized. The oxidized iron can in turn oxidize more sulfur. This is a similar process to acid mine drainage, but most likely without much acidification of the seawater due to its high buffering capacity. The identification of Thiomicrospira spp. in the 16S rRNA gene clone library of mild steel (99% reduced Fe in composition), as well as the pairwise identify of >99% of these clones to the 16S rRNA gene of strain SC-1 supports the claim that Thiomicrospira spp. are iron-oxidizers. 109 The isolation of an iron-oxidizing Thiomicrospira spp. is surprising if it is taken into account that this genus has always been linked to sulfur oxidation since its discovery in 1972 (Kuenen and Veldkamp, 1972). To our knowledge, there has been only one documented attempt to test Thiomicrospira spp. for autotrophic growth on Fe. This was tested by Wirsen et al. (1998) on Thiomicrospira strain MA-3, a T crunogena strain, eventually concluding that strain MA-3 did not grow on Fe 2 +. From this paper, it is noted the high concentration ofFe 2 + (10 mM) that is initially present in the medium and its pH of 7. The pH of the medium ideally should be at pH 6.5 or lower; a higher pH would favor the kinetics of chemical iron oxidation in the presence of 0, (g). In comparison, cultivation of strain SC-1 was conducted at pH 6.2-6.3 with initial Fe 2 + concentration of 400 µM, and daily Fe 2 + addition thereafter in microaerophilic conditions. It is now more evident that neutrophilic FeOB go through phosphate limitation at concentrations of 1 to 2 mM due to the binding of phosphate to iron oxyhydroxides and due to the abundant expression of phosphate-specific transporters (Pst) at late-log phase (Barco and Edwards, 2014 ). Therefore, an initial F e 2 + concentration of 10 mM will exacerbate this effect and would capture much more phosphate in the medium. Wirsen et al. (1998) also performed a series of Fe toxicity tests on strain MA-3 with Fe 2 + concentrations ranging from Oto 90 mM in anaerobic conditions. The results show that 10 mM Fe 2 + indeed decreases the number of viable cells of strain MA-3. No Fe 2 + concentrations were tested between 0 and 10 mM. Therefore, it is difficult to conclude iflower Fe concentrations would have been toxic to strain MA-3. Autotrophic growth on Fe should be revisited in other Thiomicrospira spp. taking into account our results and toxicity results from Wirsen et al. 110 (1998). Fleming et al. (2013) recently reported that sheath-forming microorganisms from Loihi Seamount are actually Zetaproteobacteria by using fluorescence-in-situ-hybridization (FISH) probes specific to this class. Clones recovered from this study could not be assigned specifically to sheath-Zetaproteobacteria due to the presence of other organisms from this class. In the study herein presented, a culture originating from sediment surface pyrrhotite enriched for sheath-forming Zetaproteobacteria. The clone sequences recovered from this enrichment are the first DNA sequences that can identify sheath Zetaproteobacteria. Implications for Epsilonproteobacteria: Since Thiomicrospira spp. has been reported to be a dominant community member of hydrothermal vent sites in the East Pacific Rise and Mid-Atlantic Ridge (Muyzer et al., 1995), the implication of the finding that it can perform neutrophilic Fe oxidation is important for interpreting the microbial ecology of deep-sea habitats. It also opens the possibility that other sulfur-oxidizing bacteria can perform neutrophilic Fe oxidation, at least within the genus Thiomicrospira. The results show that Thiomicrospira as well as Mariprofundus colonize mild steel in marine waters. However, mild steel colonization is dominated by members of the Epsilonproteobacteria, more specifically Sulfurimonas and Arcobacter. Thus, it is possible that Sulfurimonas and other members of the Epsilonproteobacteria can potentially perform neutrophilic Fe oxidation. The isolation of Sulfurimonas or Arcobacter from Fe mats would help 111 determine this ecological function. Alternatively, already isolated species of these Epsilonproteobacteria can be tested for autotrophic growth on Fe 2 +. Finally, the same implications can apply to members of the Alphaproteobacteria that were identified in mild steel, especially Phaeobacter which was particularly abundant in the clone library. P haeobacter is an organism of interest because it is able to produce different types of broad range antibiotics. The potential for close interactions (i.e. symbiosis) between P haeobacter spp. and Epsilonproteobacteria are evident. General assessment of bacterial communities colonizing pyrrhotite: Despite important advancements in the exploration of deep-sea habitats over the past few decades, the number ofreports on the microbial ecology of the ocean crust and associated minerals remains limited (Orcutt, et al., 201 la, 2013). In regards to pyrrhotite in the marine environment, there are two reports that originated from an in-situ incubation of a sulfide chimney from Juan de Fuca Ridge that was characterized to be rich in pyrrhotite (Edwards et al., 2003b; Toner et al., 2009). These two manuscripts focus on the mineralogy and alteration of the sulfide chimney after a colonization period of 2 months and also include elemental sulfur, pyrite, marcasite, spharelite and chalcopyrite. There were no clone libraries published from these colonization experiments. Three other colonization experiments at Juan de Fuca Ridge and Loihi Seamount focused on basaltic rock, harzburgit, pyrite/hematite, biotite, olivine, glass wool, forsterite, fayalite, hornblende, obsidian, augite, diopside, anorthite, bytownite, k-feldspar, and apatite (Orcutt et al., 2010, 2011b; Smith et al., 2011). Clone libraries are available for some of 112 these substrates (Orcutt et al., 2010, 2011b). In addition to in-situ colonization studies, microbial communities of environmental samples associated with basaltic rock, active and inactive hydrothermal vent chimneys have been analyzed from various sources (Santelli et al., 2008; Huber et al., 2006; Suzuki et al., 2004; Sylvan et al., 2012; Sylvan et al., 2013; Toner et al., 2013). Remarkably, the overall composition of the bacterial community ofpyrrhotite samples in-situ incubated on the marine surface sediment of Fisherman's Cove is similar to the bacterial composition of inactive sulfides from deep-sea environments. Hydrothermal vent chimneys when active, spew fluids rich in sulfide and reduced metals such as iron and manganese, leaving behind structures often composed of iron-sulfide, and zinc-sulfide minerals when they cease activity. A recent study on inactive sulfides of the Eastern Pacific Rise hydrothermal system indicated that the classes Garnmaproteobacteria, Deltaproteobacteria, Alphaproteobacteria and phylum Bacteroidetes are dominant in samples associated with inactive, nonventing chinmeys (Sylvan et al., 2012). Our results are consistent with this trend (Figure 4-1 ). Rarefaction curve analysis, Shannon diversity and evenness indexes show support to the claim that pyrrhotite incubated in the surface sediment is more diverse than pyrrhotite incubated in the water column. The Epsilonproteobacteria were present in both pyrrhotite samples incubated in the sediment and water column. In both cases, their relative abundance was low. Similar results were seen in other reports on microbial communities from inactive sulfides from 113 the deep-sea (Suzuki et al., 2004; Kato et al., 2010; Sylvan et al., 2012). These previous reports suggest that hydrothermal activity may be an important factor in regards to the abundance of Epsilonproteobacteria on sulfides since this class is abundant in active sulfides where the mineral is in contact with hydrothermal vent fluids (Schrenk et al., 2003; Zhou et al., 2009; Takai et al, 2009). Despite the trend seen with Epsilonproteobacteria and hydrothermal activity, there have been recent reports, that point to the specific mineralogy of the sulfide as the main driver for microbial community composition (Sylvan et al., 2013; Toner et al., 2013). The possibilities imply that slight differences in mineral composition can host different types of microbial communities. This is where in-situ incubations of fully characterized minerals and rocks can be beneficial in understanding microbial colonization of such complex structures as hydrothermal vent chinmeys. Most of the clone sequences obtained in this study match sequences from the deep-sea. The easy access to shallow marine waters makes it more practical and cost-effective to monitor a wide range of minerals and rocks nearby a fully equipped research laboratory, over defined periods of time (i.e. compared to once a year in the deep-sea). These minerals cannot replace actual deep-sea samples due to differences in physical factors (i.e. pressure and temperature), geochemical factors (i.e. heterogeneity of ocean crust and presence/absence of reduced ions) and microbial factors (i.e. effect of presence of different microbial community in adjacent mineral). At the microbial level, the minerals in-situ incubated in shallow waters can be intended to be used as tools to only approximate what is really going on in 114 the ocean floor. Magnetotactic bacteria were particularly abundant in the pyrrhotite sample that was suspended in the water column. The OTU (represented by 15_A04) identified in our study as magnetotactic bacteria belonging to the Magnetovibrio genus (Alphaproteobacteria), shares 98% similarity to clones recovered from basaltic rock of Vailulu'u Seamount, a hotspot volcano in American Samoa known to host Fe-rich mats (Sudek et al., 2009; origin of clones confirmed via personal communication). Since pyrrhotite is slightly magnetic and contains high levels of Fe relative to seawater, it is possible that M agnetovi brio was magnetically attracted to the mineral and eventually in situ enriched. This is not the first time that magnetotactic bacteria have been identified in marine sulfide samples. Suzuki et al (2004) identified abundant 16S rRNAgene sequences related to 'Candidatus Magnetobacterium bavaricum', a magnetotactic bacterium belonging to the Nitrospirae phylum, in inactive chimney sulfide samples containing sections with chalcopyrite, FeS and barite. Microbial lineages related to 'Candidatus M. bavaricum' were identified only in the chimney regions with chalcopyrite and FeS, but not in regions with barite, suggesting a correlation between metal sulfides and the presence of magnetotactic bacteria. These magnetotactic bacteria were also commonly identified in inactive sulfides from the Eastern Pacific Rise (Sylvan et al., 2012). Protein extraction from minerals and mild steel: One of the goals of the in-situ ll5 incubations in shallow marine waters was to test bacterial traps as prototypes for eventual deployments in deep-sea environments for the purpose of protein extractions. One of the main concerns was the amount of biomass needed for protein extraction and the amount of mineral or solid substrate needed to attain such a biomass. Our results indicate that about 75 g of sliced pyrrhotite can yield comparable biomass to 1 L of seawater. Protein identification was a challenge because the bacterial composition of pyrrhotite is very diverse, and the dominant phylotypes do not have genomes available in databases. However, the fact that proteins were recovered in relatively low amounts of minerals is promising for subsequent experiments in the deep-sea, especially, samples with known relative low diversity, such as sulfides from active hydrothermal vents that enrich for Epsilonproteobacteria. Proteomic results obtained from mild steel show the expression of proteins associated with the electron-transport chain and reductive tricarboxylic acid cycle specific to the Epsilonproteobacteria, a dominant class that colonizes this substrate as shown by the 16S rRNA gene clone library. These results show that the Epsilonproteobacteria (especially Sulfurimonas) are indeed active, a claim that cannot be made only with the support of clone library results. This would be important, especially in cases where not much is known about the ecological function of the microbial consortia colonizing basaltic rock below the seafloor. 116 CHAPTER IV REFERENCES Barco, R.A. and Edwards, K.J. (2014). Interactions of proteins with biogenic iron oxyhydroxides and a new culturing technique to increase biomass yields of neutrophilic, iron-oxidizing bacteria. Front. Microbiol. 5:259. doi:10.3389/fmicb.2014.00259. 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The results from this clone library indicate that the Epsilonproteobacteria (39.1%) and Alphaproteobacteria (36.2%) dominate the bacterial community in abundance. Other identified groups include the Zetaproteobacteria (8.7%), Gammaproteobacteria (7.2%), Deltaproteobacteria (4.3%), Bacteroidetes (2.9%) and Planctomycetes (1.4%). Within the Espilonproteobacteria, the most abundant phylotype (23 clones, OTU defined at 97% similarity cut-off value) belongs to the genus Sulfurimonas in the order Campylobacterales. Sulfurimonas is a known chemolithoautotrophic sulfur-oxidizing bacterium. Within the Alphaproteobacteria, the second most abundant phylotype belongs to the genus Phaeobacter (12 clones) in the order Rhizobiales. Phaeobacter is a heterotrophic marine bacterium that is known for producing the auxin phenylacetic acid and the antibiotics tropodithietic acid and thiotropocin (Seyedsayamdost et al., 2011). The Zetaproteobacteria constituted the third most abundant class in the clone library, with Mariprofundus being the only known FeOB identified in the whole clone library. Other noteworthy genera include the chemolithoautotrophs Thiomicrospira (Gammaproteobacteria) and Arcobacter (Epsilonproteobacteria), known sulfur-oxidizers. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 Figure S4-1. Relative abundances of bacterial groups colonizing mild steel after 1 month in-situ incubation at the sediment/water column interface (20 ft depth) in Fisherman's Cove, Catalina Island, CA. Figure S4-2. Rarefaction curve of mild steel in-situ incubated at Fisherman's Cove in comparison to the pyrrhotite and background seawater samples. 26 27 29 30 31 33 34 35 36 37 38 39 Table S4-1. Richness, diversity and evenness indexes of mild steel in-situ incubated in comparison with pyrrhotite samples and background seawater. Figure S4-3. Maximum likelihood phylogenetic tree based on 16S rRNA gene sequences comprising all FeOB identified in this study (1000 bootstraps). 40 41 42 44 45 46 47 48 49 Figure S4-4. SEM and energy-dispersive X-ray spectroscopy (EDS) analysis of a pyrrhotite sample that was in-situ incubated at the surface sediment for 2 weeks. A. Pyrrhotite sample with an area of significant alteration (round and raised rusty spot). B. Area at the edge of the rusty spot (box in A) showing twisted-stalks characteristic of Zetaproteobacteria. C. EDS elemental analysis of an area in pyrrhotite without significant alteration. D. EDS elemental analysis of the rusty spot in A (note the decrease in intensity of the sulfur (S) peak). 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 Table S4-2. Protein hits obtained from samples of mild steel. Results are based on best peptide matches to the Proteobacteria protein database from NCBI. 65 66 67 68 70 71 72 73 Table S4-3. Proteins hits obtained from samples of pyrrhotite. Results are based on best peptide matches to the Proteobacteria protein database from NCBI Table S4-4. Protein hits obtained from surface seawater. Results are based on best peptide matches to the Proteobacteria protein database from NCBI. 74 75 76 77 79 80 81 82 84 85 86 87

Abstract (if available)

Abstract Iron-oxidation performed by bacteria at or near neutral pH is a biological reaction that has been known since the early 1800's. Despite the number of years since its documentation and its global contribution to the biogeochemical cycle of iron, the biological mechanism of bacterial, neutrophilic, iron-oxidation has remained an enigma. Mariprofundus ferrooxydans is the first marine, neutrophilic, chemolithoautotrophic, iron-oxidizing bacteria (FeOB) that has been isolated. Its genome has been sequenced and insights about its physiology were inferred by identifying potential genes in the electron transport chain but no definite mechanism of iron-oxidation was proposed. Here, different approaches involving large-scale culturing and proteomics were combined in order to provide more definite answers about the iron-oxidation mechanism of M. ferrooxydans and FeOB in general. In-situ incubations and proteomics were also combined and applied in the marine environment to study FeOB communities colonizing iron-sulfide minerals.

FeOB are historically difficult organisms to work with that usually produce low-biomass. In order to produce enough biomass for proteomic analysis, a large-scale culturing technique was developed for M. ferrooxydans. Proteins released from these cultures were found to interact strongly with the iron mats of M. ferrooxydans and thereby affect protein extractions. Therefore, a method to circumvent protein binding to the mats is described herein. These methods were used to produce a proteomic profile of actively-growing M. ferrooxydans.The resulting proteomic profile identified numerous components of the electron-transport chain, including an abundant periplasmic cytochrome c as well as cbb₃-type cytochrome oxidases. As a result, a more specific pathway for electron transport in M. ferrooxydans is described.

In order to test the developed methods in the marine environment, FeOB communities colonizing iron?sulfide minerals in shallow waters of Catalina Islands, CA were analyzed. In general the results indicate that in-situ enriched iron-sulfides host species?rich communities that are different from the background seawater and similar to inactive hydrothermal vent chimney sulfides. Many of the clones recovered from the iron-sulfide mineral were closely related to deep-sea clones, indicating that this in-situ incubation method is appropriate for the study of microorganisms that are usually seen in deep-sea habitats such as hydrothermal vents and exposed ocean crust. From these in-situ incubations a microorganism previously known only for sulfur oxidation, Thiomicrospira spp., was isolated and shown to be capable of performing Fe oxidation. It is also shown for the first time that the sheath-Zetaproteobacteria can be grown in the laboratory. The data herein presented reveals that environmental proteomics of hard substrates such as iron-sulfide mineral and mild steel can be successfully achieved, opening the door to similar analyses in the ocean floor.

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Creator Barco, Roman Alfredo (author)

Core Title The marine, neutrophilic, and chemolithoautotrophic iron-oxidizing bacteria: insights into the physiology of Zetaproteobacteria and the discovery of novel iron-oxidizing Gammaproteobacteria

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Biology

Degree Conferral Date 2014-08

Publication Date 12/20/2014

Defense Date 04/21/2014

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag bacteria,Gammaproteobacteria,iron,iron oxyhydroxide,iron-oxidizing,Mariprofundus,OAI-PMH Harvest,oxidation,oxidizer,protein,proteomics,Zetaproteobacteria

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Edwards, Katrina J. (committee chair), Corsetti, Frank A. (committee member), Emerson, David (committee member), Nealson, Kenneth H. (committee member), Sylvan, Jason B. (committee member)

Creator Email barco@usc.edu,romanb777@yahoo.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-424514

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