EXPERIMENTAL STUDIES on FERMENTATIVE FIRMICUTES from ANOXIC ENVIRONMENTS: ISOLATION, EVOLUTION, and THEIR GEOCHEMICAL IMPACTS By

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EXPERIMENTAL STUDIES ON FERMENTATIVE FIRMICUTES FROM ANOXIC

ENVIRONMENTS: ISOLATION, EVOLUTION, AND THEIR GEOCHEMICAL IMPACTS

JESSICA KEE EUN CHOI

A dissertation submitted to the

School of Graduate Studies

Rutgers, The State University of New Jersey

In partial fulfillment of the requirements

For the degree of

Doctor of Philosophy

Graduate Program in Microbial Biology

Written under the direction of

Nathan Yee

And approved by

______

New Brunswick, New Jersey

October 2017

ABSTRACT OF THE DISSERTATION

Experimental studies on fermentative Firmicutes from anoxic environments:

isolation, evolution and their geochemical impacts

by JESSICA KEE EUN CHOI

Dissertation director:

Nathan Yee

Fermentative microorganisms from the bacterial phylum Firmicutes are quite ubiquitous in subsurface environments and play an important biogeochemical role.

For instance, fermenters have the ability to take complex molecules and break them into simpler compounds that serve as growth substrates for other organisms. The research presented here focuses on two groups of fermentative Firmicutes, one from the genus Clostridium and the other from the class Negativicutes. Clostridium species are well-known fermenters. Laboratory studies done so far have also displayed the capability to reduce Fe(III), yet the mechanism of this activity has not been investigated further. We studied three clostridial organisms, Clostridium sp. FGH that was previously enriched from Oak Ridge, TN; Clostridium beijerinckii; and Clostridium acetobutylicum to determine how Fe(III) oxide reduction occurs. The results indicate that Fe(III) oxide reduction by fermentative clostridial species does not require direct cell-to-mineral contact, meaning that it is an extracellular process mediated by a soluble molecule. Fermentation oft leads to the accumulation of acidic by-products,

ii which can act as Fe(III)-chelating ligands. But during glucose fermentation, soluble

Fe(III) accumulation did not occur, nor did supplementation of organic acids to fermenting cultures stimulate either the rate or extent of Fe(III) reduction even though rapid Fe(III) solubilization was observed. This extracellular process may be mediated by flavins, and supplementation of flavin adenine dinucleotide (FAD), a flavin molecule that serves as an electron carrier intracellularly, stimulated Fe(III) oxide reduction. These results indicate the potential for Clostridium species to perform Fe(III) reduction in contaminated environments and may be relevant towards bioremediation.

Clostridum sp. FGH was enriched as a co-culture with another spore-forming

Firmicutes organism that we isolated and named Anaerosporomusa subterranea strain

RU4. This organism is likewise an obligate fermenter but with a very narrow metabolic niche. Surprisingly, this organism in pure culture showed no ability to reduce Fe(III), despite the fact that it was isolated from an Fe(III)-reducing enrichment culture. Furthermore, the addition of typical terminal electron acceptors were not utilized by A. subterranea. However, this organism was interesting because it was a Gram-negative (or didermic, since they have two membranes in their cell wall structure) organism within a phylum that was previously thought to be only Gram- positive (or monodermic, since these microorganisms have only one membrane as part of their cell wall). We investigated how this organism could have evolved through comparative phylogenomics of outer membrane biosynthesis genes.

We identified 46 potential protein sequences involved in lipopolysaccharide and outer membrane biosynthesis in the genome of A. subterranea. Specifically out of

iii these proteins, we focused on the proteins LptACD that are responsible for transporting mature lipopolysaccharide from the inner membrane, across the periplasmic space, and to the outer membrane. This is because most Gram-negative or didermic bacteria require these proteins to synthesize an outer membrane. Our results demonstrated that A. subterranea shared approximately 50% sequence similarity for the LptACD proteins with other Negativicutes. When the alignment was expanded to other Gram-negative or didermic bacterial clades, on average 25% similarity was reported. This suggested significant homology within the Negativicutes and with other microorganisms, such that these genes in A. subterranea were ancestrally derived. Furthermore, phylogenetic comparisons using the LptACD protein sequences showed comparable tree topology to the widely-accepted 16S phylogenetic tree, illustrating vertical gene transfer and that recent horizontal gene transfer did not occur. In conclusion, our data suggests that the ancestor to the

Firmicutes phylum was originally didermic and that A. subterranea may represent an ancestral lineage.

iv ACKNOWLEDGEMENTS AND DEDICATION

I would like to start by expressing my gratitude towards my advisor, Dr.

Nathan Yee. I respect him in being able to guide me through this whole process because it has been a tough 5 years. His guidance and counsel over time has developed me as a scientific researcher, forced me to adopt new thought processes, and changed the way I think and behave as a microbiologist and scientist. Additionally, I would like to extend this gratitude towards my thesis committee members Dr. Lily Young, Dr.

Costatino Vetriani, and Dr. Katherine Dawson for their time and commitment.

This research was also made possible through various sources of funding. I would like to thank the Department of Biochemistry and Microbiology here at Rutgers for funding me for multiple years as a Teaching Assistant and for the Woodruff

Fellowship I received in my first year. Lastly, I’d like to thank the U.S. Department of

Energy Office of Science (Grant Nos. DE-FG02-08ER64544 and DE-764 SC0007051) for funding my research.

During my time here at Rutgers, I’ve met and received help from many people.

Of them, the most important person would be Dr. Madhavi Shah who helped me to get acquainted with and started in the lab as well as with my research. Even after she graduated, she continued to give me advice and help when I needed, and I cannot thank her enough. Additionally, she helped tremendously with the preparation of

Chapters 2 and 3. My time in the lab was also enjoyable thanks to the friends I made in the lab, specifically Dr. Sarah Janssen, Cindy Wang, and Thomas Wang. I had the pleasure of working with numerous undergraduate students who helped me with

v various aspects of my project, so I would like to acknowledge Jennifer Marin, Amanda

Steitz, Chioma Ekedede, Beverly Chiu, and in particular Yelyzaveta Orlovetska for her contribution to the research outlined in Chapter 4. Dr. Ines Rauschenbach and Dr.

Ramaydalis Keddis gave me a lot of direction and advice during my time as a teaching assistant here at Rutgers, and helped me to improve myself as a mentor and instructor of higher education. Lastly, I would like to acknowledge the staff of the Department of

Biochemistry and Microbiology and of the Department of Environmental Sciences for their help in dealing with a plethora of technicalities and logistics. Specifically, I’d like to thank Ms. Maria Rivera because she was such a huge help with numerous things and became such a wonderful friend and support system; thank you so much for everything.

I would like to thank my friends Ananya Agarwal, Ashley Grosche, and

Sushmita Patwardhan, the girls whom I’ve had the pleasure of sharing this entire experience with since day 1 of my PhD. We’ve gone through the good, the bad, and the ugly altogether, and I couldn’t have asked for a better support group. Lastly, I would like to thank my absolutely fantastic parents, Dr. Sung Hee Choi and Sungsook Choi for their unconditional support and love. I don’t know how they did it, but they were ultimately the ones responsible for bringing me up to this point and I wouldn’t be the person I am today with them. They are my rock and my world.

Lastly, I would like to dedicate this work to two people. First, my grandfather,

Dr. Sangup Choi, the most intelligent man I’ve known. For reasons that I still am confounded by, I have thought of him very frequently during my PhD years and miss him dearly. He left the world a much better place for me, and I didn’t really realize that

vi until after his passing. Therefore, through this dedication I would like to show him my utmost gratitude, appreciation, and love. Thank you Grandpa. I would also like to dedicate my work to my nephew, Kane Ko, in hopes that I have made and will continue to make this world a better place for him, much like what my grandfather did for me.

I also hope through this dedication that he grows up and chooses to pursue science for his future career!

vii PREFACE

Chapter 2 on clostridial Fe(III) oxide reduction is under preparation for publication as Jessica K. Choi, Annette R. Rowe, Lori Zacharoff, Nathan Yee. 2017.

Jessica K. Choi participated in writing the manuscript and performed experiments involving growth of the Clostridium species with ferrihydrite, Fe-beads, and supplementation using either citrate or FAD.

Chapter 3 has been published as Jessica K. Choi, Madhavi Shah, Nathan Yee.

2016. Anaerosporomusa subterranea gen. nov., sp. nov., a spore-forming anaerobe belonging to the class Negativicutes isolated from saprolite. IJSEM 66:3848-3854. DOI

10.1099/ijsem.0.001275. Jessica K. Choi was responsible for writing the manuscript and for characterization of A. subterranea, including its metabolism experiments,

FAME analysis, genomic deposition, organism deposition into culture collections, and the phylogenetic analyses using various house-keeping genes.

Chapter 4 on the evolution of A. subterranea as a Gram-negative Firmicutes is being prepared for publication as Jessica K. Choi, Yelyzaveta Orlovetska, Nathan Yee.

2017. Jessica K. Choi participated in writing the manuscript and is responsible for the bioinformatics analyses on the LptACD proteins.

viii TABLE OF CONTENTS

Abstract of dissertation …………………………………………………………………………………… ii

Acknowledgements and Dedication …………………………….…………………………………... v

Preface ………………………………………………………………………………………………………….. viii

List of Figures ………………………………………………………………………………………………….. xi

List of Tables …………………………………………………………………………………………………. xiii

Chapter 1: Introduction …………………………………………………………………………………… 1

The Firmicutes phylum and their general characteristics …………………….. 1

Microbial fermentation ……………………………………………………………………... 2

The Negativicutes ……………………………………………………………………………… 3

Significance of fermenting Firmicutes ………………………………………………… 4

Fe(III) reduction by the Firmicutes …………………………………………………….. 6

Oak Ridge, Tennessee ………………………………………………………………………... 8

Goal and objectives of this study ………………………………………………………... 9

Chapter 2: Extracellular Fe(III) reduction by fermentative Clostridium species involves flavins ………………………………………………………………………………….. 17

Abstract ………………………………………………………………………………………….. 17

Introduction …………………………………………………………………………………… 18

Methods …………………………………………………………………………………………. 20

Results ……………………………………………………………………………………………. 24

Discussion ………………………………………………………………………………………. 26

Chapter 3: Anaerospormusa subterranea gen. nov., sp. nov., a spore-forming anaerobe belonging to the class Negativicutes isolated from saprolite ………... 42

ix Abstract ………………………………………………………………………………………….. 42

Introduction …………………………………………………………………………………… 43

Methods …………………………………………………………………………………………. 44

Results ……………………………………………………………………………………………. 48

Discussion ………………………………………………………………………………………. 50

Description of Anaerosporomusa gen. nov. ………………………………………... 52

Description of Anaerospoomusa subterranea sp. nov. ………………………… 53

Acknowledgements …………………………………………………………………………. 54

Chapter 4: Evolution of lipopolysaccharide biosynthesis genes in

Anaerosporomusa subterranea and implications for the origin of Gram- negative Firmicutes ………………………………………………………………………………………... 64

Abstract …………………………………………………………………………………………. 64

Introduction …………………………………………………………………………………… 66

Methods …………………………………………………………………………………………. 69

Results and Discussion ……………………………………………………………………. 70

Chapter 5: Conclusions ………………………………………………………………………………….. 98

References …………………………………………………………………………………………………… 103

x LIST OF FIGURES

Figure 1.1. Phylogenetic structure of the class Negativicutes. ……………………………… 11

Figure 2.1. Fe(III) reduction by C. beijerinckii and C. acetobutylicum does not

require direct contact and occurs during stationary phase of

growth. …………………………………………………………………………………………... 34

Figure 2.2. Organic acid metabolomic profile of C. beijerinckii. ……………………………. 35

Figure 2.3. Citrate supplementation under fermenting conditions. ……………………... 37

Figure 2.4. Flavin effect on Fe(III) reduction. ……………………………………………………... 39

Figure 2.S1. The effect of FAD on Clostridium sp. FGH cultures. …………………………… 41

Fig. 3.1. Transmission electron micrograph of cells of strain RU4T harvested

at exponential phase. …………………………………………………...... 55

Figure 3.2. Phylogenetic analysis of strain RU4T. ………………………………………………... 56

Figure 3.S1. TEM micrographs of RU4T vegetative cells and spores. …………………….. 58

Figure 3.S2. Phylogenetic analysis of the atpD gene product. ……………………………… 59

Figure 3.S3. Phylogenetic analysis of the GyrB protein. ………………………………………. 60

Figure 3.S4. Phylogenetic analysis of the InfB protein. ………………………………………... 61

Figure 3.S5. Phylogenetic analysis of the RecA protein. ………………………………………. 62

Figure 3.S6. Phylogenetic analysis of the rpoB gene product. ………………………………. 63

Figure 4.1. LPS transport pathway in didermic microorganisms. ………………………… 85

Figure 4.2. Phylogenetic placement of A. subterranea within the Negativicutes

class based on LptACD protein sequences. …………………………………………. 86

Figure 4.3. Phylogenetic analysis of the LptA protein from didermic clades. ………... 88

Figure 4.4. Phylogenetic analysis of the LptC protein from didermic clades. ………… 90

xi Figure 4.5. Phylogenetic analysis of the LptD protein involved in

lipopolysaccharide transport in didermic clades. ………………………………. 92

Figure 4.6. 16S ribosomal RNA gene sequence based phylogeny of didermic

bacteria. ……………………………………………………………………………………….... 94

Figure 4.7. Sporulation process in didermic and monodermic Firmicutes. …………... 96

xii LIST OF TABLES

Table 1.1. Fe(III)-reducing organisms from the phylum Firmicutes ……………………. 12

Table 2.S1. Number of single- and multi-heme c-type cytochromes present

in the complete, annotated genomes of various microorganisms. ………. 40

Table 3.1. Physiological properties of strain RU4T and related taxa. …………………... 57

Table 4.1. List of organisms used in this study and their respective accession

numbers. ………………………………………………………………………………………… 77

Table 4.2. List of enzymes required for outer membrane biosynthesis and genes

present in A. subterranea. ………………………………………………………………… 82

xiii 1

CHAPTER 1

Introduction

The Firmicutes phylum and their general characteristics

In the Bacteria domain, the Firmicutes phylum is comprised of more than 26 families and over 223 genera (Schleifer, 2009), making it a very large bacterial clade that diverged relatively early from other bacterial phyla (Galperin, 2013). The

Firmicutes phylum constitutes one of the two bacterial phyla with Gram-positive members, and was named such by Gibbons and Murray (1978) because of their Gram- positive cell type wall. Gram-positive microorganisms have one layer exterior to the bacterial cytoplasmic membrane, comprised of mainly a thick peptidoglycan layer, which has embedded (lipo)teichoic acids and wall-associated proteins (Navarre and

Schneewind, 1999; Madigan et al., 2009). The teichoic acids seem to play a plethora of roles, including protection against cationic antimicrobial compounds, resistance against lysozyme activity, maintenance of an ion gradient, cell division, and adherence/attachment to abiotic surfaces/biofilm formation (Weidenmaier and

Peschel, 2008).

Additionally, members of this phylum share relatively low G+C %mol DNA content, typically less than 50%. However, this and the Gram-positive staining reaction are phenotypically the only unifying characteristics amongst the members of this phylum (Galperin, 2013). Otherwise, members encompass a diverse range of phenotypes. Some members are bacilli-shaped (i.e. Bacillus species) whereas others are cocci-shaped (i.e. Streptococcus species); they can be motile or non-motile; and

2 some form endospores (i.e. Bacillus and Clostridium species) whereas other members do not. The Firmicutes also differ extensively in their metabolic capabilities, ranging from aerobes (i.e. Microccus and Staphylococcus species) to obligate anaerobes, obligate fermenters (i.e. lactic acid bacteria), and more (Madigan et al., 2009; Schleifer,

2009). The wide array of characteristics makes this phylum interesting for scientific research and applications.

Microbial fermentation

Fermentation is an anaerobic process where energy in the form of ATP is produced through substrate-level phosphorylation (Madigan et al., 2009). Substrate- level phosphorylation is the transfer of a phosphate group from metabolic intermediates to an ADP molecule to form ATP. This is in contrast to oxidative phosphorylation typical of microbial respiratory pathways, which synthesizes ATP through a proton gradient established across a membrane, which requires an electron transport chain embedded in the membrane. Furthermore, fermentation occurs in the absence of an exogenous terminal electron acceptor, so the growth substrate serves as both the electron donor and electron acceptor. Again, this is in contrast to respiration, where the electron donor and electron acceptor are two different compounds.

Many members of the Firmicutes phylum are capable of fermentation or are obligate fermenters. For example, lactic acid bacteria are well-known for producing lactic acid during fermentation. Other genera, such as Clostridium or Leuconostoc, are also obligate fermenters and are capable of producing different end-products from

3 fermentation (Madigan et al., 2009). In particular, Clostridium species can utilize a variety of substrates for fermentation, for instance glucose fermentation by C. butyricum and many other clostridia, amino acid fermentation by C. sticklandii and others, purine fermentation by C. acidurici, cellulose fermentation by C. cellulolyticum, and more (Stadtman and McClung, 1956; Dorn et al., 1978; Vasconcelos et al., 1994;

Girbal et al., 1995; Leschine, 1995; Guedon et al., 2002; Hetzel et al., 2003; Sousa et al.,

2007; Fonknechten et al., 2010; Kopke et al., 2010; Masset et al., 2010; Hartwich et al.,

2012; Yarlagadda et al., 2012; Wang et al., 2013).

The Negativicutes

The Negativicutes is a class within the Firmicutes phylum that also consists of fermentative microorganisms. Out of the 5 classes within the Firmicutes (Bacilli,

Clostridia, Erysipelotrichia, Negativicutes, and Thermolithobacteria), the Negativicutes is the only one that stains Gram-negative through the classic Gram-staining technique and, along with an order called the Halanaerobiales, is one of the two groups of bacteria within the Firmicutes that stains Gram-negative. When the first of these organisms were described in 1933 by Prévot before classification via 16S rRNA gene sequencing and other molecular markers, these organisms were first either placed under the family Neisseriaceae or not allocated to any family at all. In 1971, a new family was made called Veillonellaceae (Rogosa, 1971) that included these Gram- negative microorganisms. This family remained until a new class was developed containing Gram-negative Firmicutes called the Negativicutes (Marchandin et al.,

2010) based off of 16S rRNA sequence phylogenetic comparisons. Though the

4 branching of the Negativicutes was debated due to similarities in ribosomal protein sequences with the class Clostridia (Yutin and Galperin, 2013), support for the

Negativicutes as a class separate from the Clostridia is now widely accepted due to similarities in the genome and proteome with some Gram-negatives (Vesth et al., 2013;

Campbell et al., 2015).

The Negativicutes is currently split into three different orders,

Acidaminococcales, Selenomonadales, and Veillonellales (Fig. 1.1). There is one family each under the order Acidaminococcales, named Acidaminococcaceae, and the order

Veillonellales, called the Veillonellaceae. The order Selenomonadales has two families,

Selenomonadaceae and Sporomusaceae. Out of all four families identified, the

Sporomusaceae has the largest number of accepted genera with 21 isolates in pure culture. An interesting feature of the Negativicutes is their prevalence in a number of anoxic environments, including the skin/soft tissue (Marchandin et al., 2010), termite gut (Kane and Breznak, 1991), plaque or gingival crevices (Delwiche et al., 1985), subsurface sediments or soils (Shelobolina et al., 2007; Gihring et al., 2011; Choi et al.,

2016), and more. Many of the organisms in this class are fermenters, like many other members of the Firmicutes phylum (Dimroth and Schink, 1998).

Significance of fermentative Firmicutes

Fermentative microbes have been historically important for the production of fermented foods, such as beer and bread, making them economically relevant (Giraffa,

2004). Recently, these microbes have been more widely studied for their applications in biofuel production (Antoni et al., 2007; Yazdani and Gonzalez, 2007; Kopke et al.,

2010), alternative or renewable energy sources (Lee et al., 2008; Guo et al., 2010;

Dumitrache et al., 2017), oil recovery (Banat, 1995), wastewater treatment (Shu et al.,

2015), microbial fuel cells (Park et al., 2001; Wrighton et al., 2008), and bioremediation (Baba et al., 2007; Gilmour et al., 2013; Carpio et al., 2016; Xiao et al.,

2016). In fact due to their ubiquity, diversity, and versatility, Firmicutes often dominate in environments where fermentation occurs (Turnbaugh et al., 2006; Sousa et al., 2007; Gihring et al., 2011; Sharmin et al., 2013; Lee et al., 2014; Saujet et al.,

2014).

Furthermore, fermentative Firmicutes are often detected in studies from contaminated subsurface environments. For example, fermentative Clostridium were found in abundance from enrichments using polluted estuary sediments (Lin et al.,

2007) and could perform Fe(III) reduction; similarly, fermentative Clostridium capable of U(VI) reduction were enriched from uranium mine sediment (Suzuki et al.,

2002; Suzuki et al., 2003; Akob et al., 2008). Pelosinus species, another genus of fermentative Firmicutes, was isolated from Cr(VI)-contaminated groundwater upon lactate amendment and displayed Cr(VI) reduction activity (Mosher et al., 2012).

Other fermentative Firmicutes were also detected at Oak Ridge after amendment of emulsified vegetable oil (Gihring et al., 2011). These studies plus more all indicate that Firmicutes are quite ubiquitous in these types of environments, and suggest that they may play a critical biogeochemical role in contaminant transformation, immobilization, and sequestration (Akob et al., 2006; Burkhardt et al., 2011; Carpio et al., 2016; Xiao et al., 2016).

Fe(III) reduction by the Firmicutes

One of the first microbes shown to be capable of Fe(III) reduction were fermentative Clostridia (Starkey and Halvorson, 1927). Despite the fact that most current research on microbial dissimilatory Fe(III) oxide reduction have been done on Geobacter and Shewanella species, many members of the Firmicutes are also capable of Fe(III) reduction (Table 1.1). The Fe(III)-reducing Firmicutes are immensely diverse, ranging from pathogens, such as Listeria species and

Staphylococcus aureus that perform assimilatory Fe(III) reduction to acquire biogenic iron, to fermentative Firmicutes that do not require the exogenous Fe(III) for growth but yet somehow still perform this activity.

Understanding the mechanism of Fe(III) reduction by Firmicutes is critical for geochemical applications. Numerous studies have demonstrated the variability in direct and indirect mechanisms of Fe(III) reduction utilized by members of the

Firmicutes through a combination of different techniques. For example, physical or biochemical experiments indicated the presence of enzymes that require direct cell contact with the Fe(III) mineral. With Listeria monocytogenes, separation of the cells from the Fe(III) mineral by a dialysis membrane displayed a stark decrease in Fe(III) activity (Deneer et al., 1995). In a study on Carboxydothermus ferrireducens, a membrane-bound Fe(III) reductase was purified and characterized, consisting of 2 polypeptides and c-type cytochromes (Gavrilov et al., 2007). Similarly, soluble Fe(III) reductases were purified from Thermoanaerobacter indiensis strain BSB-33 that was first extracted but then identified through the microorganism’s annotated genome, ultimately enabling amino acid sequence and protein 3D structure comparisons (Pal,

2014). Alternatively, the involvement of enzymes was resolved by a loss of Fe(III) reduction activity through the addition of specific ferric reductase inhibitors in

Thermoanaerobacter ethanolicus cultures (Roh et al., 2002). Lastly, genomic information was provided as evidence of Fe(III) reduction. Genes that may be involved as Fe(III) reductases were determined from the genome of Pelosinus sp. strain HCF1

(Beller et al., 2013). Genes encoding for multi-heme c-type cytochromes were found in Thermincola potens (Carlson et al., 2012), which are the hallmark proteins for

Geobacter and Shewanella species for direct Fe(III) oxide reduction. All these studies collectively show that members of the Firmicutes phylum are able to employ direct contact for Fe(III) reduction activity.

However, not all Firmicutes employ direct mechanisms; other studies have described indirect mechanisms that are responsible for Fe(III) reduction by the

Firmicutes. Some produce chelators, such as Bacillus cereus that secrete siderophores

(Fukushima et al., 2014), that solubilize Fe(III). Others utilize different soluble electron carriers or shuttles to help transfer electrons to Fe(III) for reduction. For example, Moorella humiferra was isolated as a Fe(III)-reducing bacterium from enrichment cultures with AQDS as an electron shuttle (Nepomnyashchaya et al.,

2012). Desulfotomaculum reducens MI-1 however may use riboflavin to mediate Fe(III) reduction (Dalla Vecchia et al., 2014), although it is possible that D. reducens employs an enzymatic mechanism as well (Otwell et al., 2015). All in all, there are a diverse number of Firmicutes that can reduce Fe(III) to Fe(II) using a wide array of mechanisms to mediate this activity.

Oak Ridge, Tennessee

One site where fermentative Firmicutes have been discovered is in the subsurface at Oak Ridge, Tennessee. Oak Ridge is currently DOE-mandated site and was historically part of the secret government project called the Manhattan Project involved in designing and developing the atomic bomb. The site housed 4 different facilities that worked on radioactive plutonium production, uranium isotope separation, and lithium isotope purification using liquid mercury (Brooks and

Southworth, 2011). These processes altogether generated massive amounts of waste containing nitric acid, radionuclides, heavy metals, and various organic contaminants.

Back in 1951, waste-disposal ponds were built to dispose this hazardous waste at an approximate rate of 10 million gallons per year until 1983 (Spain and Krumholz,

2011). Eventually, the hazardous compounds have seeped out into the groundwater and into the air, creating a contaminated plume surrounding the Oak Ridge area.

Currently, research towards bioremediation and immobilization of the radionuclide waste is underway.

Previously, a co-culture was isolated from saprolite obtained from Oak Ridge,

Tennessee (Shah et al., 2014). This co-culture was obtained from Fe(III)-reducing enrichment cultures, and contained two microbes, Clostridium sp. FGH and another named now named Anaerosporomusa subterranea strain RU4. Together in the co- culture, these microbes seemed to reduce various solid-phase Fe(III) oxides, which has potential applications for other heavy metal transformations, such as with uranium or mercury that are problematic contaminants at Oak Ridge. These two organisms in separate pure cultures served as the model organisms for this study.

Goal and objectives of this study

The goal of this study was to investigate the importance of fermentative

Firmicutes microbes from a subsurface sediment.

Chapter 2 investigates the activity of Fe(III) oxide reducing clostridia, one of which was isolated from Oak Ridge, TN. These microbes are obligate fermenters, but reduced Fe(III) when available. The objectives were to (1) investigate whether direct contact was necessary for Fe(III) reduction; (2) show if products from glucose fermentation solubilized Fe(III) oxide; and (3) determine if extracellular flavins played a role.

Chapter 3 describes the isolation and characterization of another fermentative Firmicutes, Anaerosporomusa subterranea strain RU4 originating from

Oak Ridge, TN. The objectives were to (1) isolate A. subterranea from a co-culture containing Clostridium sp. FGH; (2) to characterize the microorganism’s morphology, optimal growth conditions, metabolic capabilities, and other phenotypic qualities; and (3) use sequences available from the A. subterranea draft genome to phylogenetically place the isolated organism and classify it.

Chapter 4 studies the outer membrane biosynthesis and sporulation pathways to determine the evolution of the Firmicutes phylum. The objectives of this study were to (1) identify genes from the A. subterranea genome that encoded for proteins involved in outer membrane biosynthesis; (2) determine whether these proteins share homology to those from other members of the Negativicutes; and (3) perform phylogenetic comparisons of these protein sequences with those from other

10 microorganisms to hint towards the cell wall structure of the ancestor to the

Firmicutes phylum.

Figure 1.1. Phylogenetic structure of the class Negativicutes. The Negativicutes, as described most recently by Campbell et al. (2015), comprises 3 orders that encompass 4 families total. Overall, the class contains approximately 30 genera of

Gram-negative organisms. Boxes outlined in yellow represented the class taxonomic level, blue boxes orders, orange boxes families, and green boxes genera.

Table 1.1. Fe(III)-reducing organisms from the phylum Firmicutes.

Organism Family, Order, Class Environment Reference Bacillus Bacilliaceae, Bacillales, Bacilli Mono Lake, CA Blum et al. (1998) arsenicoselenatis Salt flat sediments, Soap Bacillus agaradhaerens Bacilliaceae, Bacillales, Bacilli Pollock et al. (2007) Lake, WA Bacillus cereus Bacilliaceae, Bacillales, Bacilli Gley soils Ottow and Glathe (1971) Taylorsville Triassic Basin, Bacillus infernus Bacilliaceae, Bacillales, Bacilli Boone et al. (1995) VA Bacillus selenitireducens Bacilliaceae, Bacillales, Bacilli Mono Lake, CA Blum et al. (1998)

Bacillus subtilis Bacilliaceae, Bacillales, Bacilli Gley soils Ottow and Glathe (1971) Great Artesian Basin, Bacillus subterraneus Bacilliaceae, Bacillales, Bacilli Kanso et al. (2002) Australia Paenibacillus (Bacillus) Paenibacillaceae, Bacillales, Bacilli Silt subsoil, Miami, FL Roberts (1947) polymyxa Abandoned coal mine, Calculibacillus koreensis n/a Min et al. (2016) Taebaek, S. Korea Tepidibacillus Mt. Simon sandstone, n/a Dong et al. (2016c) decaturensis Illinois Basin, IL ATCC and Royal Univ. Listeria monocytogenes Listeriaceae, Bacillales, Bacilli Hospital (Canada) clinical Deneer et al. (1995) and other spp. isolates Lascelles and Burke

Staphylococcus aureus Staphylococcaceae, Bacillales, Bacilli n/a (1978); Torres et al. 12

(2006)

Organism Family, Order, Class Environment References Enterococcus gallinarum Enterococcaceae, Lactobacillales, Bacilli Port Dickson, Malaysia Kim et al. (2005) MG25 Alkaliphilus Verkhnee Beloe, Buryatia Clostridiaceae, Clostridiales, Clostridia Zhilina et al. (2009) peptidofermentans (soda lake) Great Artesian Basin, Caloramator australicus Clostridiaceae, Clostridiales, Clostridia Ogg and Patel (2009a) Australia Caloramator Great Artesian Basin, Clostridiaceae, Clostridiales, Clostridia Ogg and Patel (2011) mitchellensis Australia Clostridium sp. BC1 Clostridiaceae, Clostridiales, Clostridia Coal-cleaning waste Francis and Dodge (1988) Marine sediment, Halifex Clostridium sp. EDB2 Clostridiaceae, Clostridiales, Clostridia Bhushan et al. (2006) Harbor, Canada Clostridium sp. FGH Clostridiaceae, Clostridiales, Clostridia Saprolite, Oak Ridge, TN Shah et al. (2014) Clostridium Gessenbach creek soils, Clostridiaceae, Clostridiales, Clostridia Burkhardt et al. (2011) aminobutyricum Germany Freshwater lake sediment, Clostridium beijerinckii Clostridiaceae, Clostridiales, Clostridia University of East Anglia, Dobbin et al. (1999) UK Clostridium butyricum Clostridiaceae, Clostridiales, Clostridia Microbial fuel cell Park et al. (2001) Clostridium Garcia-Balboa et al. Clostridiaceae, Clostridiales, Clostridia “Brunita”, Murcia, Spain celerecrescens (2010) Clostridium Fossil laterite, Vogelsberg, Clostridiaceae, Clostridiales, Clostridia Ottow (1971) pasteurianum Germany Clostridium Clostridiaceae, Clostridiales, Clostridia Lake Pavin, France Lehours et al. (2010)

saccharabutylicum 13

Organism Family, Order, Class Environment References

Clostridium scatologenes Clostridiaceae, Clostridiales, Clostridia ATCC 25775 Doerner et al. (2008) Starkey and Halvorson Clostridium sporogenes Clostridiaceae, Clostridiales, Clostridia n/a (1927) Thermotalea Great Artesian Basin, Clostridiaceae, Clostridiales, Clostridia Ogg and Patel (2009b) metallivorans Australia Desulfitobacterium Gessenbach creek soils, Peptococcaceae, Clostridiales, Clostridia Burkhardt et al. (2011) dichloroeliminans Germany Desulfitobacterium Peptococcaceae, Clostridiales, Clostridia Lake Coeur d’Alene, ID Niggemeyer et al. (2001) hafniense Desulfitobacterium San Juan River, Shiprock, Peptococcaceae, Clostridiales, Clostridia Finneran et al. (2002) metallireducens NM Desulfosporosinus lacus Gessenbach creek soils, Peptococcaceae, Clostridiales, Clostridia Burkhardt et al. (2011) STP12T Germany Tebo and Obraztsova Desulfotomaculum Mare Island Naval Shipyard, (1998); Dalla Vecchia et Peptococcaceae, Clostridiales, Clostridia reducens MI-1 San Francisco, CA al. (2014); Otwell et al. (2015) Thermophilic microbial fuel Thermincola potens Peptococcaceae, Clostridiales, Clostridia Carlson et al. (2012) cell Ochre deposits and water, Thermincola ferriacetica Peptococcaceae, Clostridiales, Clostridia Zavarzina et al. (2007) Kunashir Island Syntrophomonadaceae, Clostridiales, Great Artesian Basin, Fervidicola ferrireducens Ogg and Patel (2009c) Clostridia Australia Carboxydocella Tokarev crater, Lake Unassigned, Clostridiales, Clostridia Slepova et al. (2006) sporoproducens Karymskoe, Russia Gessenbach creek soils, Sedimentibacter spp. Unassigned, Clostridiales, Clostridia Burkhardt et al. (2011) 14

Germany

Organism Family, Order, Class Environment References Halobacteroidaceae, Halanaerobiales, Briny groundwater, Illinois Dong et al. (2016a); Dong Orenia metallireducens Clostridia Basin, IL et al. (2016b) Caldanaerobacter Thermoanaerobacteraceae, Terrestrial hydrothermal Nepomnyashchaya et al. hydrothermalis Thermoanaerobacterales, Clostridia spring, Kamchatka, Russia (2010) Carboxydothermus Slobodkin et al. (1997); Thermoanaerobacteraceae, Calcite springs, Yellowstone (Thermoterrabacterium) Gavrilov et al. (2007); Thermoanaerobacterales, Clostridia National Park, WY ferrireducens Gavrilov et al. (2012) Carboxydothermus Thermoanaerobacteraceae, Geyser Valley, Kamchatka, Slepova et al. (2009) siderophilus Thermoanaerobacterales, Clostridia Russia Carboxydothermus Thermoanaerobacteraceae, Acidic hot spring, Kyushu, Yoneda et al. (2012) pertinax Thermoanaerobacterales, Clostridia Japan Thermoanaerobacteraceae, Terrestrial hydrothermal Nepomnyashchaya et al. Gelria glutamicus Thermoanaerobacterales, Clostridia spring, Kamchatka, Russia (2010) Thermoanaerobactericeae, Terrestrial hydrothermal Nepomnyashchaya et al. Moorella glycerini Thermoanaerobiales, Clostrida spring, Kamchatka, Russia (2010) Thermoanaerobactericeae, Terrestrial hydrothermal Nepomnyashchaya et al. Moorella humiferra Thermoanaerobiales, Clostrida spring, Kamchatka, Russia (2012) Thermoanaerobactericeae, Terrestrial hydrothermal Nepomnyashchaya et al. Mahella australiensis Thermoanaerobiales, Clostrida spring, Kamchatka, Russia (2010) Thermoanaerobacter Thermoanaerobacteraceae, Thar Jath oil field, Unity Schouw et al. (2013) spp. Thermoanaerobacterales, Clostridia State, South Sudan Thermoanaerobacter Thermoanaerobacteraceae, Piceance Basin, CO Roh et al. (2002) ethanolicus Thermoanaerobacterales, Clostridia Thermoanaerobacter Thermoanaerobacteraceae, Bakreshwar hot spring, Bhowmick et al. (2009); indiensis BSB-33 Thermoanaerobacterales, Clostridia West Bengal, India Pal (2014) Thermoanaerobacter Thermoanaerobacteraceae, Hydrothermal vents, Slobodkin et al. (1999)

siderophilus Thermoanaerobacterales, Clostridia Karymsky volcano, Russia 15

Organism Family, Order, Class Environment References Thermoanaerobacter Thermoanaerobacteraceae, Geothermal spring, Ayas, Balk et al. (2009) thermohydrosulfuricus Thermoanaerobacterales, Clostridia Turkey Caldicellulosiruptor Unassigned, Thermoanaerobacterales, Terrestrial hydrothermal Nepomnyashchaya et al. saccharoyticus Clostridia spring, Kamchatka, Russia (2010) Thermovenabulum Unassigned, Thermoanaerobacterales, Great Artesian Basin, Ogg et al. (2010) gondwanense Clostridia Australia Zhelezistyi hydrothermal Thermovenabulum Unassigned, Thermoanaerobacterales, source, Udon caldera, Zavarzina et al. (2002) ferriorganovorum Clostridia Russia Desulfosporomusa n/a Lake Stehlin, Germany Sass et al. (2004) polytropa Sporomusaceae, Selenomonadales, Pelosinus sp. HCF1 DOE 100H site, Hanford, WA Beller et al. (2013) Negativicutes Sporomusaceae, Selenomonadales, Pelosinus fermentans Russian kaolin Shelobolina et al. (2007) Negativicutes Thermosinus Sporomusaceae, Selenomonadales, Norris Basin, Yellowstone Sokolova et al. (2004) carboxydivorans Negativicutes National Park, WY Veillonella alcalescens Veillonellaceae, Veillonellales, Woolfolk and Whiteley (Micrococcus n/a Negativicutes (1962) lactilyticus) Thermolithobacteraceae, Thermolithobacter Calcite Spring, Yellowstone, Thermolithobacterales, Sokolova et al. (2007) ferrireducens WY Thermolithobacteria n/a = not available

CHAPTER 2

Extracellular Fe(III) oxide reduction by Clostridium species involves flavin

adenine dinucleotide

Abstract

Clostridium species are ubiquitous in subsurface environments and have been shown to reduce Fe(III). However, few studies have been done to investigate the mechanism. In this study, we examined the mechanism of Fe(III) oxide reduction by three different fermenting Clostridium species: Clostridium sp. FGH that was isolated from Oak Ridge, TN; Clostridium beijerinckii, which has been studied mostly for its solvent-producing capabilities; and Clostridium acetobutylicum, another solvent- generating microbe. An experiment was first conducted with Fe(III) in beads to determine direct contact requirement for Fe(III) reduction. Subsequently, experiments investigating Fe(III)-chelating compounds or the effect of flavins on

Fe(III) oxide reduction were performed. The results displayed that direct cell-to-

17 mineral contact was not required for electron transfer to solid-phase Fe(III) oxides.

Extracellular metabolite analysis showed production of organic acids, such as citrate that can chelate Fe(III). But the addition of citrate did not stimulate Fe(III) reduction.

Fe(III) reduction, however, was stimulated upon addition of exogenous flavin adenine dinucleotide (FAD) to glucose fermenting cultures. These results suggest that FAD is involved in extracellular Fe(III) oxide reduction.

Introduction

In contaminated soils and sediments, Fe(III) oxide minerals are commonly found in association with heavy metals and organic compounds (Kappler and Straub,

2005). Fe(III) oxide reduction is an important geochemical process (Nealson and

Saffarini, 1994; Lovley, 1997; Straub et al., 2001) that influences the fate of chemical species adsorbed and co-precipitated with Fe(III) oxides in anaerobic subsurface environments (Lovley, 1991; Weber et al., 2006). For instance, Fe(III) oxide reduction can release inorganic and organic compounds – such as CO2 due to organic matter decomposition – and affect the distribution and transport of heavy metal contaminants (Lovley, 1991; Lovley, 1997). Therefore, it is important to understand

Fe(III) oxide reduction in anoxic subsurface environments for applications in contaminant mobility and/or removal.

The process of Fe(III) oxide reduction is mediated by a diverse number of anaerobic microorganisms (Jones et al., 1984; Lovley, 1987; Lovley, 1991; Lovley and

Chapelle, 1995; Weber et al., 2006; Lin et al., 2007), including fermentative microbes

18 from the genus Clostridium. These microorganisms are found ubiquitously and in high abundance in Fe(III) reducing environments (Ottow, 1971; Ottow and Glathe, 1971;

Kostka et al., 2002; Petrie et al., 2003; Scala et al., 2006; Weber et al., 2006; Lin et al.,

2007; Lehours et al., 2009; Wang et al., 2009). Furthermore, clostridial species were among the first to be identified as Fe(III)-reducing bacteria (Starkey and Halvorson,

1927; Ottow, 1971; Hammann and Ottow, 1974). Thus, these microorganisms play an important role in the reductive dissolution of Fe(III)-bearing minerals and are thought to be the primary agents for the reduction of Fe(III) oxides in certain

19 sediments (Lovley et al., 1991; Kostka et al., 2002; Scala et al., 2006; Lin et al., 2007;

Lehours et al., 2009). To date, the mechanism of Fe(III) reduction by Clostridium spp. remains poorly understood.

Four strategies for Fe(III) reduction have been proposed in Fe(III)-respiring bacteria (Gralnick and Newman, 2007). Two of these strategies require direct contact of the solid-phase Fe(III) with either the cell surface or pili called “nanowires” and involves multi-heme c-type cytochromes (Lovley et al., 1993; Caccavo et al., 1994;

Gorby et al., 2006; von Canstein et al., 2008; Shi et al., 2009; Bucking et al., 2010;

Coursolle and Gralnick, 2010; El-Naggar et al., 2010; Coursolle and Gralnick, 2012; Shi et al., 2012; Pirbadian et al., 2014). The remaining two are indirect mechanisms of

Fe(III) reduction that involves either ligands that solubilize Fe(III) or involves electron shuttles, such as flavins utilized by Shewanella oneidensis MR-1 (Lies et al., 2005;

Marsili et al., 2008; von Canstein et al., 2008; Covington et al., 2010; Brutinel and

Gralnick, 2012a; Brutinel and Gralnick, 2012b; Kotloski and Gralnick, 2013),

Desulfotomaculum reducens MI-1 (Dalla Vecchia et al., 2014), and Geothrix fermentans

(Nevin and Lovley, 2002; Bond and Lovley, 2005; Mehta-Kolte and Bond, 2012) or phenazines by Pseudomonas spp. (Hernandez et al., 2004; Wang et al., 2010; Wang et al., 2011). Thus far, all Clostridium species tested in the laboratory have been shown to reduce Fe(III) and accumulate Fe(II) extracellularly during fermentation. These species include to C. pasteurianum, C. butyricum, C. saccharolyticum, C. sporogenes, C. beijerinckii, and C. celerecrescens (Ottow, 1971; Hammann and Ottow, 1974; Lovley et al., 1991; Dobbin et al., 1999; Park et al., 2001; Garcia-Balboa et al., 2010; Lehours et al., 2010; Yarlagadda et al., 2012). Currently, experimental studies on the mechanism

20 of clostridial Fe(III) reduction are lacking and the reaction pathway of how the Fe(III)- reduction process occurs is unknown. This represents a gap in knowledge and hence requires further examination.

In this study, we investigated the Fe(III) reduction activity of fermentative organisms Clostridium sp. FGH that was isolated in our lab from a co-culture using

Oak Ridge saprolite, Clostridium beijerinckii, and Clostridium acetobutylicum. The three objectives of this study are to: (1) investigate whether direct contact is necessary for Fe(III) reduction; (2) examine if products from glucose fermentation solubilized Fe(III) oxide by acting as chelating ligands; and (3) determine if extracellular flavins play a role. We show that these Clostridium species exhibit indirect Fe(III) reduction and that FAD mediates this process. The results presented through this research provide new insights on how Fe(III) reduction occurs by anaerobic fermentative bacteria.

Methods

Bacterial strains, culture media, and growth conditions

Clostridium sp. FGH, Clostridium beijerinckii strain ATCC 25752 and

Clostridium acetobutylicum strain ATCC 824 were used as the model organisms. All species were initially grown in anaerobic reinforced clostridial medium (RCM), composed of (g/L): 10 peptone, 10 beef extract, 5 dextrose, 5 NaCl, 3 yeast extract, 3 sodium acetate, 1 soluble starch, and 0.38 L-cysteine. C. acetobutylicum cultures were maintained in RCM. Cultures of Clostridium sp. FGH and C. beijerinckii were subsequently maintained in anaerobic defined media 2 (M2) supplemented with 2%

(w/v) peptone. The basal M2 medium contained (mM): 0.75 NH4Cl, 0.1 NaCl, 0.34

KH2PO4, 0.38 MgCl2·6H2O, 0.34 CaCl2·2H2O, and 100 1,4-piperazinediethanesulfonic acid (PIPES) as the buffer, adjusted to pH 6.8-7.0 by addition of NaOH. Additionally,

M2 contained a modified Wolfe’s mineral solution (10 mL/L), composed of (g/L): 1.5 nitrilotriacetic acid, 0.5 MnCl2·4H2O, 0.1 FeCl2·4H2O, 0.1 CoCl2·6H2O, 0.1 ZnCl2, 0.01

CuCl2·2H2O, 0.01 H3BO3, 0.01 NaMoO4·2H2O, 0.01 NaSeO3, 0.01 NiCl2·6H2O, and 0.01

Na2WO4·2H2O (Wolin et al., 1963). After autoclaving, a filter-sterilized Wolfe’s vitamin mix (Wolin et al., 1963) was added to the M2 medium. All cultures were grown under strict anaerobic conditions at 30°C with shaking at 200 rpm.

Iron oxide synthesis and reduction assays

Synthetic ferrihydrite was aseptically prepared in the laboratory using the protocol described previously by Schwertmann and Cornell (1991). Briefly, a solution of ferric nitrate was buffered to approximately pH 7 by drop-wise addition of a KOH solution, resulting in precipitation of ferrihydrite. This Fe(III) oxyhydroxide mineral

21 suspension was then washed with sterile milliQ water 2-3 times, and then purged with 100% N2 gas in a sealed serum bottle to remove dissolved oxygen. Nanoporous glass beads containing ferrihydrite were synthesized using the procedure described previously by Lies et al. (2005), where beads were added to a solution of ferric chloride and then the pH adjusted to 6.5-7 by drop-wise addition of KOH. The Fe- beads were washed 10 times with a KCl solution, then placed on a 0.105 mm polypropylene mesh filter and washed again. The resulting beads were examined by bright-field microscopy at 100X total magnification to check for Fe(III) particles

22 outside of the beads by comparing the beads before and after washing. Dithionite extraction was carried out (Mehra and Jackson, 1960) on both the ferrihydrite suspension and Fe-beads and the extracted/reducible iron concentration was quantified using a modified ferrozine assay (Viollier et al., 2000).

Iron reduction experiments were conducted with washed Clostridium cells in defined M2 with glucose as the carbon source. Clostridium sp. FGH and C. beijerinckii inoculum was grown in M2 + peptone, while C. acetobutylicum was grown in RCM.

Cells were harvested at mid-exponential phase and washed with basal M2 medium twice anaerobically. Then, washed Clostridium sp. FGH and C. beijerinckii cells were resuspended in fresh M2 supplemented with 100 mM glucose. On the other hand, C. acetobutylicum cells were resuspended in M2 without PIPES buffer, the pH adjusted to 4.5, and 100 mM glucose as the growth substrate. Approximately 1 mM of ferric iron as either ferrihydrite or Fe-beads was added to cultures before inoculation. Both unfiltered and filtered culture samples were collected at periodic time intervals.

Unfiltered samples were reacted with 0.5 N trace-metal grade HCl for 1 h before

22 quantifying the concentration of Fe2+ using the ferrozine assay, whereas filtered samples were analyzed immediately for both dissolved Fe2+ and Fe3+. Absorbance measurements were done using a Shimadzu Biospec Mini.

Metabolomics

Extracellular metabolomics was conducted through Metabolon, Inc. (Durham,

NC). Samples were prepped following Metabolon Sample Prep guidelines, with minor modifications. C. beijerinckii was grown and washed as described above, then

23 inoculated in M2 medium with 100 mM glucose. 0.5 mL samples of culture were first centrifuged anaerobically in a Coy chamber for 10 min at 10 krpm. Subsequently, the supernatant was syringe-filtered (0.22 μm pore size) into a cryogenic tube and immediately flash-frozen with liquid nitrogen. Samples were stored at -80°C until analysis by Metabolon. Briefly, after an initial protein precipitation and removal, each sample was divided into 5 aliquots. 2 of these aliquots were analyzed using reverse- phase ultra-high performance liquid chromatography-mass spectrometry coupled to mass spectrometry (UHPLC-MS/MS) with positive ion mode electrospray ionization

(ESI), 1 aliquot using reverse-phase UHPLC-MS/MS with negative ion mode ESI, and

1 aliquot using hydrophilic interaction liquid chromatography (HILIC) coupled to

UHPLC-MS/MS with negative ion mode ESI. The last aliquot was saved for back-up.

The raw data was analyzed and processed using Metabolon software (Evans et al.,

2012; Evans et al., 2014).

To test for chelation by organic acids, cultures with ferrihydrite were supplemented with 5 mM of sodium citrate. Inoculum of C. beijerinckii and C.

23 acetobutylicum was grown and washed according to the procedure above, and inoculated in respective assay media. Samples were taken at various time points to quantify total Fe(II) and soluble Fe(III) concentrations using the ferrozine assay.

Flavin iron reduction assays

Flavin mononucleotide (FMN) and FAD were tested for its effect on iron reduction. Inoculum of all three microorganisms was grown and washed according to the procedure described above, and then inoculated into M2 medium with 100 mM

24 glucose, 1 mM ferrihydrite, and 10 μM of FMN. Samples were taken at different time intervals for Fe2+ and Fe3+ quantification using the ferrozine assay.

Results

Fe(III) reduction does not require direct contact

C. beijerinckii and C. acetobutylicum were able to reduce Fe(III) confined inside nanoporous glass beads, thus indicating that direct contact for Fe(III) reduction is not required. Enclosing the Fe(III) oxide inside the beads had little to no effect on

Fe(III) reduction rate by C. beijerinckii, but only 25% of the Fe(III) given was reduced

(Fig. 2.1A). This was in stark contrast with C. acetobutylicum, which displayed almost

100% Fe(III) reduction (Fig. 2.1B). However, Fe(III) reduction in nanoporous beads by this organism had a longer lag phase compared to reduction of particulate Fe(III) oxide. Regardless, Fe(III) reduction from the beads was observed for both microbes during fermentative growth.

Clostridium species produces ligands but Fe(III) chelation via organic acids does not occur

We performed extracellular global metabolomics on C. beijerinckii cultures during glucose fermentation to examine the accumulation of compounds that might be involved in Fe(III) reduction. Organic acids, such as α-ketoglutarate and citrate, constituted the majority of the extracellular metabolites (Fig. 2.2), suggesting the possibility of chelation by ligands. Metabolomics done on fermenting C. acetobutylicum cultures (Amador-Noguez et al., 2011) similarly showed production

25 of organic acids.

Since citrate is a very well-known chelator, we then supplemented both C. beijerinckii and C. acetobutylicum cultures with citrate to explore potential effects of

Fe(III) chelation. There was rapid Fe(III) oxide solubilization by citrate when supplemented in the culture medium (Fig. 2.3A and C). On the other hand, the addition of this ligand to C. beijerinckii cultures did not stimulate Fe(II) production when compared to that of cultures without any supplemental citrate (Fig. 2.3A and B).

However, adding citrate inhibited Fe(III) reduction activity in C. acetobutylicum, whereas cultures without supplemental citrate seemed to reduce 100% of the provided Fe(III) oxide within 3 days (Fig. 2.3C and D).

Cultures of C. beijerinckii and C. acetobutylicum were both analyzed for soluble

Fe(III) accumulation without the addition of any citrate. Organic acids naturally produced by C. beijerinckii and C. acetobutylicum did not solubilize Fe(III) (Fig. 2.3B and D). This indicates that solubilization of solid-phase Fe(III) by organic acids during glucose fermentation is negligible and organic acids most likely do not play a role in

Fe(III) oxide reduction.

Flavins plays a role in Fe(III) reduction

In a preliminary experiment, the role of FAD was tested using Clostridium sp.

FGH (Fig. 2.S1). Because FAD was detected as one of the extracellular metabolites of

C. acetobutylicum, we supplemented FAD to Fe(III)-reducing cultures of C. beijerinckii and C. acetobutylicum (Fig. 2.4). For all three microbes, FAD supplementation stimulated Fe(III) reduction. The rate of Fe(III) reduction was initially enhanced

26 through the addition of FAD compared to control experiments without FAD addition.

Over time, the rate of Fe(III) reduction in the control experiments eventually reached similar values to experiments with FAD. We also supplemented FMN to C. beijerinckii and C. acetobutylicum cultures and observed no difference in Fe(III) reduction (data not shown).

Discussion

Clostridium species do not have the capacity to reduce Fe(III) by establishing direct cell-to-mineral contact via multi-heme c-type cytochromes. These unique proteins are the hallmark characteristic for direct Fe(III) oxide reduction by both

Geobacter and Shewanella spp. that are responsible for transferring electrons from the cell to the Fe(III) mineral surface (Shi et al., 2009; Bucking et al., 2010; El-Naggar et al., 2010; Pirbadian et al., 2014). As such, the annotated genomes of these two genera contain multiple c-type cytochromes that contain multiple heme-binding motifs (CXXCH) (Heidelberg et al., 2002; Butler et al., 2010; Wrighton et al., 2011).

This is very different from the information available in genomes of multiple

Clostridium species (Table 2.S1), including the three studied here. These Clostridium genomes show an absence of multi-heme binding motifs, and in fact have a maximum of one heme binding motif per c-type cytochrome annotated.

One experimental method that can establish direct contact necessity for microbial Fe(III) oxide reduction is through experiments with Fe(III)-containing beads as described previously (Lies et al., 2005), where the nano-sized pores of the beads prevent cells from contacting the iron mineral surfaces. These experiments

27 showed little to no effect on Fe(III) reduction rates by Clostridium sp. FGH during glucose fermentation (Shah, unpublished). Fe-beads experiments using C. beijerinckii and C. acetobutylicum showed a similar trend especially when compared to experiments with particulate Fe(III) (Fig. 2.1). This signifies that direct contact is not a requirement for Fe(III) reduction. However, the extent of Fe(III) reduction in beads was significantly different between C. beijerinckii and C. acetobutylicum. This could be due to the fact that visually C. acetobutylicum cultures become more turbid than C. beijerinckii cultures, regardless of whether the Fe(III) oxides were contained in beads or not. The 100% vs. 25% difference in amount of Fe(III) reduced could be attributed to a difference in cell numbers. Regardless, for all three Clostridium species, the small nano-sized pores of these beads do not allow large structures to pass through.

Therefore, we concluded that a small molecule must be produced or released that mediates Fe(III) reduction by an indirect mechanism.

Although indirect Fe(III) reduction can be achieved through Fe(III)-chelating agents, we eliminated the possibility of Fe(III) oxide solubilization via organic acids

27 produced during glucose fermentation by Clostridium species. Glucose fermentation is a very well-characterized process, especially in fermenting organisms like C. acetobutylicum (Vasconcelos et al., 1994). Under glucose fermenting conditions,

Clostridium species produce mainly butyrate, acetate, lactate, formate, and H2 gas

(Vasconcelos et al., 1994; Lin et al., 2007; Jeong et al., 2008; Kim et al., 2008; Pan et al.,

2008; Milne et al., 2011; Zhao et al., 2011; Masset et al., 2012; Tanasupawat et al.,

2014). These reactions are exergonic, with standard Gibb’s free energy values approximately -117 kJ (in the case that lactate is the sole end product) per mol of

glucose at pH 7 (Lee et al., 2008). Removal of any fermentation product, such as H2, would shift the equilibrium dynamics so that further glucose fermentation is still a favorable, exergonic process. Additionally, changes in the environment can lead to significant changes in fermentation products (Wang et al., 2012), switching from organic acids to solvents, such as butanol, acetone, and more significant amounts of ethanol. In some studies, adding multiple growth substrates other than glucose alone, such as glycerol or acetate, displayed metabolic shifts in fermentation products

(Vasconcelos et al., 1994; Girbal et al., 1995; Wang et al., 2011), whereas others have reported a difference in fermentative H2 gas production based off of temperature or pH (Kim et al., 2008; Masset et al., 2010; Zhao et al., 2011), addition of methyl viologen

(Yarlagadda et al., 2012), or exogenous electron shuttles (Hatch and Finneran, 2008;

Ye et al., 2012).

Extracellular metabolomics previously performed on C. acetobutylicum

(Amador-Noguez et al., 2011), detection of fermentation products by Clostridium sp.

FGH (M. Shah, unpublished), and global metabolite analysis examined here on

28 fermenting C. beijerinckii revealed the accumulation of numerous organic acids over time as predicted, including the well-known Fe(III)-chelating compound citrate (Fig.

2.2). We tested the possibility that Fe(III) reduction does not occur until the culture reaches stationary phase because it takes time for the cells to accumulate these chelating ligands outside the cell. In this case, the addition of these ligands at the time of inoculation should remove the time lag in Fe(III) reduction consistently observed thus far. Yet, cultures supplemented with citrate showed little to no effect on Fe(III) reduction (Fig. 2.3A and C) and cultures without supplemental citrate did not

29 accumulate any dissolved Fe(III) (Fig. 2.3B and D), therefore ruling out a chelation mechanism.

Our data indicates that indirect Fe(III) reduction by Clostridium species can be achieved through the release of electron carriers, a process that has also been observed in other dissimilatory iron-reducing bacteria (DIRBs). For instance,

Shewanella oneidensis MR-1 uses FMN as its electron carrier (Marsili et al., 2008; von

Canstein et al., 2008; Brutinel and Gralnick, 2012a; Kotloski and Gralnick, 2013) while

Geothrix fermentans may use riboflavin (Nevin and Lovley, 2002; Bond and Lovley,

2005; Mehta-Kolte and Bond, 2012). Because extracellular metabolomics on C. acetobutylicum (Amador-Noguez et al., 2011) revealed the accumulation of FAD over time, we supplemented Fe(III)-reducing cultures with FAD. These cultures consistently showed that FAD stimulated Fe(III) reduction significantly compared to cultures without any exogenous FAD added (Fig. 2.4, Fig. 2.S1), whereas supplementing cultures with FMN had no effect (data not shown). This suggests a role for FAD in mediating Fe(III) reduction by multiple fermentative clostridial species,

29 where the cells donate excess electrons to extracellular FAD and reduce it to FADH2.

The FADH2 then donates those electrons to Fe(III) oxides and reduces it to Fe(II).

Theoretically, FAD/FADH2 pair has a standard redox potential (ΔE˚’) of approximately -200 mV, whereas Fe(III) oxides used in this study have a redox potential ranging from -100 to +100 mV (Bird et al., 2011; Kracke et al., 2015).

Therefore, in the absence of bacterial cells, the reduced form of FAD can potentially transfer electrons to Fe(III) oxides. As a result, it is uncertain whether the terminal electron sink is FAD or the Fe(III) oxides. One experiment that could help resolve this

question is an abiotic reaction between reduced FAD (or FADH2) and Fe(III) oxides to look for Fe(III) oxide reduction by FADH2 in the absence of bacterial cells. Additionally, determining the necessity of FAD for Fe(III) oxide reduction, perhaps by creating genetic mutants unable to synthesize FAD, would clarify the role of FAD in Fe(III) oxide reduction by clostridia. It could also be interesting, if the technology is available, to do an experiment where the concentrations of extracellular FAD and FADH2 are individually and continuously monitored in Fe(III) oxide reducing cultures, which would also determine whether we can observe a constant cycle of FAD reduction/FADH2 oxidation that could serve as support for FAD as an electron shuttle for Fe(III) oxide reduction.

FAD is an important micronutrient that is synthesized from riboflavin (RF), or vitamin B2. FAD is synthesized from FMN by FAD synthetase or FMN adenylyltransferase that also functions as RF kinase, which makes FMN from RF. FAD is synthesized from FMN by addition of an adenine mononucleotide to the phosphate end of FMN. In turn, FMN is synthesized as an irreversible reaction by

30 phosphorylation of the 5’ position of the ribose chain of RF (Abbas and Sibirny, 2011).

FAD and other flavins are important for cellular function because they serve as cofactors for a variety of enzymes, and is estimated that approximately 1-3% of genes encode flavin-binding proteins (Abbas and Sibirny, 2011). FAD is useful as enzyme cofactors because it exists in three different redox states, including a fully oxidized or quinone state, one-electron reduced or semiquinone state, and a fully reduced or hydroquinone state. This makes them useful in enzymes with redox activity that require electron transport, such as electron-bifurcating dehydrogenases (Wang et al.,

2013), NADH-dependent flavin oxidoreductases (Morrison and John, 2015), and other enzymes involved in carbon metabolism (Hetzel et al., 2003; Kopke et al., 2010;

Hartwich et al., 2012; Dumitrache et al., 2017). It is not fully understood why

Clostridium species accumulate flavins extracellularly. One possibility is that the accumulation is not an active secretion process, but rather a result of cell lysis. Since

Clostridium are spore-forming organisms, it is possible that these flavins (and all other cell components) are released simultaneously with mature spore formation.

Accumulation of Fe(II) occurred during fermentation in the three species studied here as well as in other Clostridium tested in the laboratory thus far (Ottow,

1971; Hammann and Ottow, 1974; Lovley et al., 1991; Dobbin et al., 1999; Park et al.,

2001; Garcia-Balboa et al., 2010; Lehours et al., 2010; Yarlagadda et al., 2012; Shah et al., 2014). This is distinct compared to anaerobic Fe(III) respiration, as fermenters do not reduce Fe(III) for energy generation via an electron transport chain and grow perfectly without the exogenous Fe(III). Therefore, an important question that still remains unanswered is the reason why Clostridium species reduce Fe(III) oxides.

Some studies reported a difference in growth or fermentation products in the presence of Fe(III) (Lehours et al., 2010; Yarlagadda et al., 2012) compared to without, leading to the hypothesis that the Fe(III) serves as an electron sink during fermentation and thus allows the microorganism to consume and ferment more substrate. Usually during fermentation, excess electrons are transferred to H+ to make

H2 that accumulates in cultures and stops fermentation. But if these electrons are instead diverted to Fe(III) oxides or other mineral surfaces, then fermentation can continue and generate more energy for the organism. This may have been the case for

C. acetobutylicum, where cultures with Fe(III) oxides grew more turbid visually than cultures without Fe(III) oxides (data not shown). Further experiments would have to be done with this organism in order to determine why it reduces Fe(III) oxides.

However, other studies have demonstrated an insignificant difference in growth regardless of the presence of Fe(III) (Dobbin et al., 1999; Park et al., 2001).

This may be the case with C. beijerinckii and Clostridium sp. FGH, where growth with and without Fe(III) oxides seemed to have no effect on the extent of growth or their growth rate (data not shown). It is possible that Fe(III) oxides are reduced abiotically by FADH2, as noted above, and that this reaction just happens to occur as a result of

FAD reduction. Lastly, another theory is that the Fe(III) oxide acts as a natural pH buffer. Fe(III) oxides have a lot of anionic binding/adsorption sites, and so may buffer the acidic end-products from fermentation, allowing the microorganism to consume and ferment more of the substrate (Dong et al., 2016b).

This data presented here on the mechanism of fermentative Fe(III) oxide reduction by Clostridium species have biogeochemical implications for

32 bioremediation of contaminated subsurface environments, such as Oak Ridge, TN.

This site is heavily contaminated with uranium and mercury, but bioremediation studies at this site have detected Clostridium and other fermentative bacteria in the soil (Gihring et al., 2011). Fermentative organisms are commonly found in such environments, and if these organisms are capable of reducing Fe(III) or other heavy metals, they may be able to contribute to heavy metal contaminant removal. Although laboratory experiments have shown that DIRBs grow faster and reduce more Fe(III) than fermentative Firmicutes in the same time-frame, in natural environments

33 fermentative iron-reducers may be present in larger numbers and persist in the environment longer. We provide direct evidence that a fermenting organism isolated from Oak Ridge, Clostridium sp. FGH, is capable of Fe(III) oxide reduction via FAD.

Furthermore, this mechanism is shared by two other related organisms, C. beijerinckii and C. acetobutylicum. This suggests that Clostridium species may be capable or reducing and removing other heavy metals through the same process. Fe(III) reducing bacteria tend to utilize the same mechanism for reduction of other heavy metals, and therefore have been widely studied in context of bioremediation. Likewise,

Clostridium species are capable of reducing soluble U(VI) to insoluble U(IV) (Gao and

Francis, 2008; Dalla Vecchia et al., 2010) though the mechanism was not determined at the time. It is possible that FAD may be involved in this process as well and so may be important for future bioremediation studies.

Figure 2.1. Fe(III) reduction by C. beijerinckii and C. acetobutylicum does not

34 require direct contact and occurs during stationary phase of growth. (A) C. beijerinckii and (B) C. acetobutylicum reduced Fe(III) contained within nanoporous glass beads, as observed by the increase in percent of iron reduced. Black circles () are cultures grown in the presence of Fe-beads, white circles () are Fe-beads without cells, and the black triangles (▲) are cultures grown in the presence of particulate ferrihydrite (FH). Data points with error bars represents the average of triplicates, with the error bars depicting one standard deviation.

Figure 2.2. Organic acid metabolomic profile of C. beijerinckii. Different organic acids detected in the extracellular spent medium of C. beijerinckii cultures are depicted in order of most to least concentrated at the end of day 11 from time of inoculation compared to day 1. Black squares represent minimal to no fold changes, whereas bright yellow squares represent 103-fold changes in concentration.

Figure 2.3. Citrate supplementation under fermenting conditions. C. beijerinckii cultures at pH 6 (A) with and (B) without citrate, and C. acetobutylicum cultures at pH

4.7 (C) with and (D) without citrate supplemented during glucose fermentation.

While there was rapid solubilization of Fe(III) due to the presence of citrate, the availability of the Fe(III) in aqueous phase had no effect on Fe(II) production. Black circles () represent Fe(II) concentrations, while white circles () for soluble Fe(III) concentrations. Each data point represents the average of three replicates, and the error bars for 1 standard deviation.

Figure 2.4. Flavin effect on Fe(III) reduction. Rates of Fe(III) reduction for (A) C. beijerinckii and (B) C. acetobutylicum were calculated when FAD was added to cultures

and compared to the rate without any flavin. The data here shows evidence that FAD stimulates Fe(III) reduction. Each bar represents the average of triplicate cultures, and the error bars 1 standard deviation.

Table 2.S1. Number of single- and multi-heme c-type cytochromes present in the complete, annotated genomes of various microorganisms. Organisms shaded in grey indicate Gram-positive bacteria, and white Gram-negative bacteria.

Multiheme c-type Average heme Organism c-type cytochromes /cytochrome cytochromes Clostridium sp. FGH 0 0 - Clostridium acetobutylicum 1 0 1 Clostridium beijerinckii NCIMB 2 0 1 8052 Clostridium butyricum 6 0 1 Clostridium pasteurianum BC1 0 0 - Clostridium saccharobutylicum 0 0 - Clostridium scatologenes 1 0 1 Clostridium sporogenes 0 0 - Paenibacillus polymyxa SC2 1 0 - Bacillus megaterium ATCC 3 0 1 14581 Pseudomonas chlororaphis 15 3 1 PCL1606 Pseudomonas aeruginosa PA01 16 9 2

Rahnella aquatilis ATCC 33071 5 4 3 Aeromonas salmonicida 10 8 3 Shewanella oneidensis MR-1c 39 23 6 Geobacter metallireducensb 76 66 7 Geobacter sulfurreducens PCAb 89 78 8 Thermincola potensa 41 32 12 aData taken from Wrighton et al. (2011)and Carlson et al. (2012) bData taken from Butler et al. (2010) cData taken from Heidelberg et al. (2002)and Wrighton et al. (2011)

Figure 2.S1. The effect of FAD on Clostridium sp. FGH cultures. Fe(III) reduction by Clostrdium sp. FGH over time when supplemented with FAD was faster initially than compared to without any FAD. Each data point represents the average of three replicates, and error bars for one standard deviation.

CHAPTER 3

Anaerosporomusa subterranea gen. nov., sp. nov., a spore-forming anaerobe

belonging to the class Negativicutes isolated from saprolite

Published in the International Journal of Systematic and Evolutionary Microbiology

(IJSEM 2016, 66:3848-3854)

Abstract

A Gram-stain-negative, spore-forming, anaerobic bacterium designated strain

RU4T was isolated from a saprolite core collected from Oak Ridge, Tennessee, USA.

Cells were slightly curved rods and exhibited an outer membrane exterior to a thin cell wall. Strain RU4T formed heat-resistant endospores in late-log phase and stationary phase cultures. Under anaerobic conditions, strain RU4T grew by fermenting fumarate and maleate, but did not grow on glucose, glycerol, pyruvate, lactate, succinate, citrate, formate, acetate, propionate, butyrate or valerate. Strain

RU4T did not reduce sulfate or ferric iron. The main cellular fatty acids were C17:0 cyclo,

C16:0 and C15:0. The DNA G+C content was 52 mol%. Analysis of the 16S rRNA, rpoB, recA, infB, gyrB and atpD gene sequences indicated that the isolate is related to members of the family Sporomusaceae. Based on 92% sequence similarity of the 16S rRNA gene to its closest relatives in the family Sporomusaceae and divergent physiological traits, the newly-cultivated isolate was assigned to a novel species of a new genus, Anaerosporomusa subterranea gen. nov., sp. nov. The type strain of

Anaerosporomusa subterranea is RU4T (=DSM 29728T=ATCC BAA-2723T).

Introduction

The class Negativicutes is a group of anaerobic Gram-stain-negative bacteria in the phylum Firmicutes previously classified as Clostridia (e.g. Yutin and Galperin,

2013). Recently, Campbell et al. (2015) proposed a reorganization of this group with the division of the class into the three orders Selenomonadales, Veillonellales and

Acidaminococcales. Notably, members of the class Negativicutes display a Gram-stain- negative cell wall structure in which cells are surrounded by an outer membrane that is exterior to a thin layer of peptidoglycan. This is unique because the phylum

Firmicutes was once thought to comprise exclusively Gram-stain-positive bacteria with low DNA G+C content (Yutin and Galperin, 2013). As a result, microorganisms from this group were frequently misplaced into other bacterial lineages due to their

Gram-stain-negative phenotype or remained unclassified. Phylogenetic analyses based on 16S rRNA gene sequences and other molecular markers now place these

Gram-stain-negative bacteria firmly within the phylum Firmicutes (Jumas-Bilak et al.,

2004; Sass et al., 2004; Shelobolina et al., 2007; Sattley et al., 2008; Marchandin et al.,

2010; Church et al., 2011; Moe et al., 2012; Yutin and Galperin, 2013; Ueki et al., 2014;

Campbell et al., 2015; Wang et al., 2015).

Previously, an enrichment culture was established using unconsolidated saprolite (weathered bedrock) from a sediment core collected at the United States

Department of Energy (DOE) Oak Ridge Field Research Center (ORFRC) in Tennessee,

USA (Shah et al., 2014). This culture was composed of two fermenting Firmicutes species. Here, we report the isolation and characterization of one of these organisms from this enrichment culture designated as strain RU4T. Phylogenetic analysis using

44 the 16S rRNA gene indicates that strain RU4T belongs to the class Negativicutes.

Methods

Enrichment, isolation, and cultivation

The ORFRC enrichment culture was established using 4.5-6.0 m deep sediment cores as the initial inoculum and then maintained by transferring the culture every week to fresh liquid medium. The medium A (MA) contained 20 mM

NH4Cl, 2.2 mM KH2PO4, 0.6 mM CaCl2·2H2O, 8 mM MgCl2·6H2O, 10 mM PIPES buffer and 10 ml mineral solution l-1 (Wolin et al., 1963) adjusted to pH 7 using 1 M NaOH.

Prior to inoculation, 10 ml Wolfe’s vitamin mix l-1 (Wolin et al., 1963), 0.05 g yeast extract l-1, 1 mM cysteine and 30 mM fumarate as the carbon source were added to the medium. To isolate strain RU4T, the culture was transferred to inverted, deoxygenated MA tubes amended with 1.5% (w/v) Noble agar. Incubation at 30°C for

3 days resulted in formation of bubbles embedded in the agar. Under anoxic conditions, the cells in each bubble were then transferred using a needle and syringe

44 to collapse the bubble and subsequently injected into fresh inverted soft agar. After three consecutive transfers, individual bubbles were inoculated into liquid MA containing 10 g yeast extract l-1 to promote a higher density of growth. The pure culture was designated as strain RU4T and preserved at -80°C with 10% (v/v) DMSO.

Morphology and metabolic capabilities

The morphology, cell wall structure and sporulation of strain RU4T were characterized. First, cells of strain RU4T were examined using transmission electron

45 microscopy. To prepare thin sections, cells were fixed in 4% (v/v) formaldehyde and

1% (v/v) glutaraldehyde in 0.1M Millonig’s phosphate buffer at pH 7.3 for 3 h, incubated in 1% osmium tetroxide for 1 h, and then dehydrated in a graded ethanol series. Cells were then embedded in Epon-Araldite, sectioned with a diamond knife and stained with a 5% (w/v) uranyl acetate solution in 50% ethanol and 0.5% (w/v) lead citrate solution. Electron micrographs were taken using a model JEM 100 CX transmission electron microscope (JEOL).

Optimum growth for RU4T was tested anaerobically in defined MA with 30 mM fumarate, and determined based on the optical density (OD) at 600 nm after 7 days. The utilization of alternative electron donors was tested under anoxic conditions at 30°C. Growth by fermentation on different carbon substrates was examined in MA with 100% N2 headspace and supplemented with the following carbon sources (30mM): fumarate, maleate, glucose, glycerol, pyruvate, lactate, succinate, citrate, formate, acetate, propionate, butyrate, and valerate, as well as

H2/CO2 (2:1, v/v) and H2/30mM Na2SO4. Growth was measured by OD at 600 nm over

28 days. Positive growth was designated as a change in optical density at 600 nm

(ΔOD600) ≥0.1, while ΔOD600 <0.1 was considered as negative for growth.

The utilization of sulfate and ferric iron [Fe(III)] as electron acceptors was tested to investigate growth by anaerobic respiration. Sulfate reduction was examined using all carbon sources listed above. Cultures amended with 30 mM Na2SO4 were sampled periodically and sulfate concentrations were measured by ion chromatography on an ICS-1000 chromatograph (Dionex) with an anion column

(ASRS 250 column). Sulfide concentrations were also measured using the assay

46 method of (Cline, 1969). To test for reduction of ferric iron, 10 mM synthetic ferrihydrite was supplemented with 30 mM fumarate as the electron donor in MA.

Production of reduced ferrous iron was quantified using a modified ferrozine assay

(Viollier et al., 2000). Media samples were added to ferrozine solutions and absorbance was measured at 562 nm using a Biospec-mini spectrophotometer

(Shimadzu).

Fatty acid methyl ester (FAME) analysis

Fatty acid methyl esters (FAMEs) were obtained from 30 mg of freeze-dried cells grown in MA under fumarate-fermenting conditions. Fatty acid analysis was carried out by the Identification Service of DSMZ using the methods described by

Miller (1982) and Kuykendall et al. (1988). Briefly, FAME mixtures were separated using a SHERLOCK Microbial Identification System (MIDI, Microbial ID). The

Microbial Identification System Standard Software (Microbial ID) automatically integrated and identified peaks, then calculated respective percentages.

Genome sequencing and genome-based analyses.

The draft genome of strain RU4T was sequenced using an Illumina Genome

Analyzer IIX. Briefly, genomic DNA was extracted from a culture using the PowerSoil

DNA Isolation kit (MoBio), purified with the UltraClean DNA Purification kit (MoBio), and a paired-end library was prepared using an Illumina Nextera kit. DNA sequencing reads from the Illumina Genome Analyzer IIX were assembled using the CLC

Genomics Workbench 5.1. The G+C content was calculated using GeeCee and the

47 tetranucleotide frequency of both strands was calculated using TETRA (Teeling et al.,

2004), clustered with Cluster 3.0 and visualized in JavaTreeView (Saldanha, 2004).

This was used to verify all contiguous sequences (contigs) belonged to a single organism.

The G+C content of the DNA was alternatively determined by the

Identification Service of DSMZ using the protocols described by Cashion et al. (1977) and Mesbah et al. (1989). Briefly, individual deoxyribonucleotides were quantified by

HPLC calibrated with lambda DNA (Tamaoka and Komagata, 1984). Three DNA sequences – Bacillus subtilis DSM 402, DNA G+C content 43mol%; Xanthomonas campestris DSM 3586T, 65mol%; and Streptomyces violaceoruber DSM 40783, 72mol%

– were used as standards. G+C values were calculated according to the method of

Mesbah et al. (1989).

The 16S rRNA gene of strain RU4T was sequenced and analyzed. PCR amplification of the gene was carried out using universal primers GM3F (5’-

AGAGTTTGATCMTGGC-3’) and GM4R (5’-TACCTTGTTACGACTT-3’) and the product

47 was sequenced by Genewiz. The 1509 bp sequence was deposited in the National

Center for Biotechnology Information (NCBI) database under accession number

KX268498. Other 16S sequences from the Negativicutes were obtained from the NCBI database, and together aligned through the Ribosomal Database Project (Cole et al.,

2014). A maximum-parsimony tree was constructed with 1000 bootstrap replicates using MEGA version 4 (Tamura et al., 2007).

Results

Transmission electron micrographs of strain RU4T showed slightly curved rods 2-4 μm long and 0.3-0.5 μm wide, and enveloped by an outer membrane exterior to a thin cell wall (Figs. 3.1 and 3.S1a), indicating the strain is Gram-stain-negative.

Gram staining of RU4T cells using standard procedures (Gerhardt et al., 1994) confirmed this. A lateral flagellum located on the convex side of the rod was observed

(Fig. 3.S1b), indicating strain RU4T is motile. At late exponential phase, vegetative cells formed terminal endospores at the end of vegetative cells (Fig. 3.S1c). In stationary phase, the spores measured 1.0x0.6 μm and were covered by a wavy, loosely-fitting exosporium located exterior to the cortex and outer spore coat (Fig. 3.S1d). Spores remained viable even after heat pasteurization at 75°C for 15 min.

Growth occurred under neutrophilic and mesophilic conditions between 24 and 30°C, with optimum growth occurring at 28°C. The pH range for growth was 5-

7.5 with an optimum at pH 6.8. The presence of vitamins or yeast extract to the medium was not required, but if added promoted a higher density of growth. No

48 growth was observed when RU4T was incubated aerobically. The isolate grew anaerobically by fermentation on fumarate and maleate only, but not on any of the other substrates (Table 3.1). Strain RU4T was also unable to grow autotrophically using H2 as the electron donor and CO2 as the carbon source, both with and without sulfate. Furthermore, the addition of sulfate to the media did not stimulate growth on any of the carbon substrates, and neither loss of sulfate nor formation of sulfide was detected in any of the cultures. Growth of strain RU4T in the presence of iron oxides was observed but no iron reduction occurred.

FAME analysis indicated that the majority of fatty acids present in strain RU4T were saturated hydrocarbon chains. The most abundant fatty acids were C17:0 cyclo

(15.9%), C16:0 (15.6%), and C15:0 (13.0%). Other saturated fatty acids detected were

C17:0 (7.1%), C14:0 (3.7%), C11:0 (1.4%), C13:0 (1.3%), and C12:0 (1.7%). Fatty acids with one double bond in the hydrocarbon chain, such as C17:1 ω8c (8.0%), C15:1 ω8c (7.1%),

C18:1 ω8c (4.2%) and C16:1 ω9c (1.3%) were also present. The FAME analysis also resulted in several summed features representing two or more fatty acids that could not be separated by the Microbial Identification System. Summed feature 1 (6.1%) consisted of C13:0 3-OH and/or iso-C15:1, summed feature 2 (4.2%) consisted of C14:0 3-

OH and/or iso-C16:1, and summed feature 3 (3.8%) consisted of C15:0 2-OH and/or C16:1

ω7c.

Ultimately, 28 contigs were obtained for strain RU4T. These contigs were large in size and had coverage over 600x. Using the HPLC method, the G+C content of the

DNA from strain RU4T was determined to be 52mol%, whereas the average DNA G+C content was slightly lower at 46mol%. A total of 3411 open reading frames were

49 identified. Genes were annotated using BLAST, KEGG Automatic Annotation Server

(KAAS) and Rapid Annotation using Subsystem Technology (RAST) (Moriya et al.,

2007; Aziz et al., 2008). This Whole Genome Shotgun project has been deposited at

DDBJ/ENA/GenBank under accession LSGP00000000. The version described in this paper is version LSGP01000000.

Using the EzTaxon server (http://www.ezbiocloud.net/eztaxon; Kim et al.,

2012), the 16S rRNA gene sequence of RU4T showed the highest similarity to members of the family Sporomusaceae, order Selenomonadales (Fig. 3.2). Strain RU4T

50 was most closely related to Dendrosporobacter quercicolus DSM 1736T (accession number AJ010962, 91.7% sequence similarity), Propionispora vibrioides FKSB1T

(AJ279802, 91.4%), Acetonema longum DSM 6540T (AFGF01000106, 91.3%),

Sporomusa aerivorans TmAO3T (AJ506191, 91.2%), Sporomusa sphaeroides DSM

2875T (AJ279801, 91.1%), and Pelosinus fermentans DSM 17108T (AVKN01000045,

90.3%).

Additionally, housekeeping genes were identified in the draft genome sequence using the TBLASTX tool. These included the RNA polymerase ß-subunit

(RU4_gene_3630), ATP synthase F1 ß-subunit (RU4_gene_1585), translation initiation factor IF-2 (RU4_gene_749), DNA gyrase subunit B (RU4_gene_2192), and

DNA recombination and repair protein (RU4_gene_625) genes. Nucleotide sequences of these genes from related organisms were obtained from the NCBI database.

Alignment of translated housekeeping genes was performed using ClustalW and phylogenetic trees were reconstructed using the neighbor-joining method and 1000 bootstrap replicates in MEGA 4. The translated protein sequences of the

50 housekeeping genes were most closely related to the RpoB, AtpD, GyrB, InfB, and RecA sequences (Figs 3.S2-S6) in the genera Sporomusa and Propionispora. Sequence similarity of these housekeeping genes to those of related organisms ranged from 81 to 90%. Phylogenetic analysis of the housekeeping genes confirmed the taxonomic placement of strain RU4T in the family Sporomusaceae.

Discussion

Strain RU4T represents a distinct lineage within the family Sporomusaceae.

Phylogenetic analyses of the 16S rRNA, rpoB, atpD, gyrB, infB, and recA genes consistently place RU4T in its own group (Fig. 3.2, Figs 3.S2-S6). The relatively low sequence similarities with other known Sporomusaceae members lie below the taxonomic threshold at the genus level as reported by Yarza et al. (2014), which signifies that RU4T represents a novel species of the novel genus. Furthermore, the fatty acid profile of strain RU4T is composed primarily of saturated hydrocarbons and differs from the most closely related taxa. The most abundant fatty acids in strain

RU4T (C17:0 cyclo and C16:0) are absent or occur at very low levels in D. quercicolus DSM

1736T (Strompl et al., 2000). Similarly, the most dominant fatty acids in P. fermentans

(C14:0 DMA and C15:1 ω8c and/or C15:2) were not detected in strain RU4T (Moe et al.,

2012). Fatty acid profiles for D. quercicolus and P. fermentans were obtained using cultures grown in peptone and yeast extract supplemented with either fructose or glucose (Strompl et al., 2000; Moe et al., 2012). In contrast, strain RU4T cannot grow on glucose, and fatty acids were determined in cultures grown on fumarate. Based on the data presented, strain RU4T is considered to represent a novel species of a new

51 genus in the family Sporomusaceae, for which the name Anaerosporomusa subterranea gen. nov., sp. nov. is proposed.

A particularly unique characteristic of strain RU4T is that its growth is limited to fermentation on short-chain dicarboxylic acids (Table 3.1). While the fermentative metabolism of strain RU4T is a common trait among the Sporomusaceae, other members of this family typically ferment a wide range of carbon sources. Our study suggests that strain RU4T is limited to fermentation on short-chain dicarboxylic acids.

This stark contrast indicates that strain RU4T occupies a narrow metabolic niche in

52 its natural habitat and suggests that this organism is reliant on other members of the surrounding ecological community to produce its growth substrates. If RU4T relies on the metabolic products of other fermenters that breakdown complex molecules, then consumption of those products by strain RU4T may alleviate buildup of short-chain dicarboxylic acids and enhance further carbon metabolism by primary fermenters.

This may explain why members of this family (formerly known as Veillonellaceae) are highly enriched during carbon injection experiments into anoxic groundwater aquifers (Gihring et al., 2011).

The data presented in this study describe a newly isolated microorganism representing a novel species of a new genus, Anaerosporomusa subterranea RU4T, as part of the newly created family Sporomusaceae in the class Negativicutes. A recent study by Campbell et al. (2015) has rearranged the phylogeny of this group based on the identification of conserved molecular markers, and proposed a revised taxonomic framework for the class Negativicutes that includes two other orders in addition to

Selenomonadales. In this new organization, strain RU4T is placed under the family

Sporomusaceae within the emended order Selenomonadales based on 16S rRNA gene sequence similarities reported in this study.

Description of Anaerosporomusa gen. nov.

Anaerosporomusa (An.ae.ro.spo.ro.mu’sa. Gr. prefix an without; Gr. masc. n. aer air; Gr. n. spora seed; L. n. musa banana; N.L. fem. n. Anaerosporomusa an anaerobic spore- bearing banana).

Cells are slightly curved rods, Gram-stain-negative and motile. Spore-forming.

Mesophilic and neutrophilic. Anaerobic. Limited to growth by fermentation on short- chain carboxylic acids. Unable to reduce sulfate or Fe(III). The cellular fatty acid profile is primarily composed of saturated hydrocarbon chains.

A member of the family Sporomusaceae, class Negativicutes, according to 16S rRNA gene sequence analysis. The type species is Anaerosporomusa subterranea.

Description of Anaerosporomusa subterranea sp. nov.

Anaerosporomusa subterranea (sub.ter.ra’nea L. fem. adj. subterranea subsurface).

Exhibits the following features additional to those given in the genus description. Cells are 2-4 μm in length and 0.3-0.5 μm in diameter. Possesses a flagellum, placed laterally on the convex side of the rod. Terminal endospores are formed at late exponential phase. Endospores are heat-resistant. Growth occurs optimally at pH 6.8 and 28°C,

53 but can also occur at a pH range of 5.0-7.5 and a temperature range of 24-30°C. Can only use fumarate and maleate as fermentation substrates. The fatty acid profile is dominated by C17:0 cyclo, C16:0, and C15:0.

The type strain, RU4T (=DSM 29728T=ATCC BAA-2723T), was isolated from saprolite collected at the United States Department of Energy (DOE) Oak Ridge Field Research

Center (ORFRC) site located in Oak Ridge, Tennessee. The DNA G+C content is

52mol%.

Acknowledgements

This research was supported by the Office of Science (BER), U.S. Department of Energy

Grant No. DE-FG02-08ER64544. We thank Chu-Ching Lin for establishing the primary enrichment culture and Udi Zelzion for assembling the RU4T genome. Tamar Barkay and two anonymous reviewers provided insightful comments that greatly improved the manuscript.

IM OM 0.5μm

Figure 3.1. Transmission electron micrograph of cells of strain RU4T harvested at exponential phase. Bar, 0.5 μm. The inset shows the inner membrane (IM) and outer membrane (OM). Cells are curved rods approximately 2-4 μm in length and 0.3-0.5

μm in width. Additional images are displayed in Fig. 2-S1.

Figure 3.2. Phylogenetic analysis of strain RU4T. Phylogenetic tree reconstructed

56 T using the 16S rRNA gene sequence from strain RU4 and other organisms in the class

Negativicutes. Shown are representatives from the three orders Selenomonadales,

Veillonellales and Acidaminococcales. Sequences were aligned using the Ribosomal

Database Project aligner and a maximum-parsimony tree was reconstructed with

1000 bootstrap replicates through MEGA 4, with bootstrap values over 50% indicated.

Scale bar represents number of changes over the whole sequence. Shewanella oneidensis MR-1 served as the outgroup. Phylogenetic trees of the housekeeping genes rpoB, atpD, gyrB, infB and recA are displayed in Figs 2-S2-S6.

Table 3.1. Physiological properties of strain RU4T and related taxa.

Strains: 1, RU4T; 2, Propionispora vibrioides FKSB1T (data from Biebl et al., 2000); 3, Acetonema longum DSM 6540T

(Kane & Breznak, 1991); 4, Sporomusa aerivorans TmAO3T (Boga et al., 2003); 5, Sporomusa sphaeroides DSM

2875T (Möller et al., 1984); 6, Pelosinus fermentans R7T (Shelobolina et al., 2007). ND, Not determined.

1 2 3 4 5 6

Cell size (μm) 0.3-0.5x2-4 0.6x2.2-6 0.3-0.4x6-60 0.6-0.7x3-7 0.5-0.8x2-4 0.6x2-6 Straight Cell shape Curved rods Curved rods Curved rods Curved rods Curved rods rods Gram-stain Negative Negative Negative Negative Negative Negative DNA G+C content 52 48.5±0.2 51.5 ND 46.7-47.4 41.0 (mol%) Temperature 24-30 30-40 19-40 19-35 15-45 4-36 range (°C) pH range 5-7.5 5.0-8.5 6.4-8.6 6.2-8.2 5.7-8.7 5.5-8

Motility Yes Yes Yes Yes Yes Yes Spore Yes Yes Yes Yes Yes Yes formation Growth on

substrates*

H2/CO2 – – + + + –

Fumarate + – + + – + Maleate + ND ND ND ND ND

Glucose – – + – – + Glycerol – + – – + –

57 Pyruvate – – + + + +

Lactate – – – + + + Succinate – – – + – + Citrate – ND + + – + Formate – – – + + ND Acetate – ND – ND – ND *Electron donors tested under fermenting conditions. +, Visible growth observed; –, no growth.

(a) (b) F

0.5μm

0.5μm 0.1μm

Figure 3.S1. TEM micrographs of RU4T vegetative cells and spores. (a) Image of a single RU4T cell at exponential phase showing the curved rod morphology. (b) TEM

whole-mount image of RU4T showing a laterally-placed flagellum (F). (c) RU4T cells at late exponential phase showing the swelling at one end of the cell. (d) Spores of RU4T at stationary phase.

Figure 3.S2. Phylogenetic analysis of the atpD gene product. The β-subunit of the

F1 ATP synthase protein sequence from RU4T was compared to the AtpD protein

sequences from other members of the Sporomusaceae and Veillonellaceae families. 59

Sequences were aligned using ClustalW and a neighbor-joining tree with 1000 bootstrap replicates was generated. Bootstrap values (%) above 50% are indicated, and accession numbers are shown in parentheses. The atpD gene product from strain

RU4T did not group specifically with any genera and shared at most 90% sequence identity to AtpD protein from other organisms in the family.

Figure 3.S3. Phylogenetic analysis of the GyrB protein. The translated sequence of

60 the gyrB gene from strain RU4T was compared to the GyrB protein sequences from other members of the Sporomusaceae and Veillonellaceae families. The neighbor- joining tree was constructed using methods described for the rpoB gene sequence analysis. The gyrB gene product from strain RU4T shared a low sequence identity

(84%) to GyrB proteins from the most closely related organisms.

Figure 3.S4. Phylogenetic analysis of the InfB protein. The neighbor-joining tree of the infB gene product was constructed using methods described for the atpD gene 61

sequence comparison. InfB from strain RU4T was placed in the Sporomusaceae family but did not cluster with any of the known genera. A blastp comparison of InfB displayed at most 88% sequence similarity to the most closely related protein sequence from the Sporomusaceae family.

Figure 3.S5. Phylogenetic analysis of the RecA protein. The protein sequence of 62

the DNA recombination and repair protein encoded by the recA gene was compared to RecA sequences found in other organisms of the Sporomusaceae and Veillonellaceae family. The neighbor-joining tree was generated using methods described for the atpD gene sequence analysis. The recA gene product from strain RU4T showed only 81% sequence similarity to RecA proteins from closely related organisms.

Figure 3.S6. Phylogenetic analysis of the rpoB gene product. The β-subunit of the

RNA polymerase protein sequence from RU4T was compared to the RpoB protein

sequences from other members of the Sporomusaceae and Veillonellaceae families.

The neighbor-joining tree was generated using the same method as for the atpD gene analysis, with bootstrap values (%) from 1000 replicates indicated. The rpoB gene product from strain RU4T did not group specifically with any genera and shared at most 83% sequence identity to RpoB protein sequences from other organisms in the

Sporomusaceae family.

CHAPTER 4

Evolution of lipopolysaccharide biosynthesis genes in Anaerosporomusa

subterranea and implications for the origin of Gram-negative Firmicutes

Abstract

The Firmicutes phylum was once thought to consist of Gram-positive

(monodermic) members only. It was not until the advance of 16S rRNA gene sequencing and other molecular techniques that it was discovered some members of this phylum were Gram-negative (didermic). Here, we explored the evolution of the outer membrane in didermic Firmicutes, using Anaerosporomusa subterranea as our model organism. The objective of the study was to determine if lipopolysaccharide biosynthesis genes from A. subterranea were inherited from a common Firmicutes ancestor or if they were acquired by horizontal gene transfer. We identified 46 different proteins with various roles in LPS biosynthesis, including those involved in the production of individual LPS components and the export of LPS to the outer

64 membrane. These proteins, such as LptACD, which are responsible for exporting LPS to the outer membrane, shared approximately 50% amino acid similarity with the homologous proteins from other members of the class Negativicutes. Our results indicate that LPS of A. subterranea and other didermic Firmicutes share a common origin, and are thus evolutionarily related. Furthermore, phylogenetic analyses of these same proteins illustrated tree topology similar to that of the 16S-rRNA based taxonomic tree. These analyses support the hypothesis that the Firmicutes ancestor was a diderm and that modern monodermic Firmicutes evolved through a

65 simplification event involving the loss of the outer-membrane.

Introduction

In microbiology, bacterial cell wall structures are traditionally classified as

Gram-postive (monodermic) or Gram-negative (didermic). Monodermic bacteria are classically known as Gram-positives based on the Gram staining procedure, where these bacteria possess only one cytoplasmic membrane that is surrounded by a thick layer of peptidoglycan. Embedded in the Gram positive peptidoglycan layer are teichoic or lipoteichoic acids that face the outside environment. Didermic bacteria (or

Gram-negatives) also have a cytoplasmic membrane surrounded by a peptidoglycan layer that is much thinner, but also has a second membrane exterior to these two layers called the outer membrane. The outer membrane of diderms consists of lipopolysaccharide (LPS) that is important for bacterial viability, cell wall stability, endotoxin properties, interaction of the cell with its environment, and serotype classification (Luderitz et al., 1982; Raetz and Whitfield, 2002; Caroff and Karibian,

2003; Knirel, 2011; Silipo and Molinaro, 2011). This outer membrane also serves as a molecular sieve for some compounds to enter the periplasmic space (Costerton et al.,

1974; Decad and Nikaido, 1976), and acts as an impermeable layer to harmful compounds such as detergents (Sperandeo and Polissi, 2016). Although monoderms and diderms have very different structures, not many studies have been done investigating their evolutionary history (Sutcliffe, 2010; Tocheva et al., 2011; Antunes et al., 2016; Tocheva et al., 2016).

An important part of LPS biosynthesis in didermic bacteria is the LPS export machinery. Individual components of LPS are first synthesized in the cytoplasm and then exported to the outer membrane by the Lpt system (Fig. 4.1). LPS facing the

67 periplasmic space is extracted by an ABC transporter LptBFG (Okuda et al., 2016).

This energy-supplying unit then transfers LPS to LptC, which complexes with LptA, a soluble periplasmic protein (Ruiz et al., 2009; Sperandeo et al., 2011; Villa et al., 2013).

LPS is afterwards passed from LptC to LptA to LptDE sequentially (Dong et al., 2014).

LptDE is a transmembrane β-barrel protein complex that ultimately inserts LPS into the outer leaflet of the outer membrane in didermic bacteria (Gu et al., 2015; Botos et al., 2016; Sperandeo and Polissi, 2016). Most diderms require this system for outer membrane biogenesis.

The Firmicutes phylum is a large group of bacteria, including members from genera such as Bacillus, Staphylococcus, and Clostridium. Originally, this phylum was classified as monodermic (or Gram-positive) only. However, recent sequencing and genomic analyses have proven that this phylum also includes two didermic clades, named Halanaerobiales and Negativicutes (Sutcliffe, 2010; Antunes et al., 2016).

Recently, we isolated and characterized a novel bacterium, named Anaerosporomusa subterranean strain RU4, which is phylogenetically related to members of the

Negativicutes (Choi et al., 2016). Because this phylum now encompasses both dermic types of bacteria and is regarded as one of the early-diverging branches in the bacterial domain (Galperin, 2013), we were interested in studying the evolutionary history of this phylum using A. subterranea as our model organism.

Previous studies investigating the evolution of the didermic outer membrane have considered two scenarios concerning the earliest Firmicutes microorganism

(Sutcliffe, 2010; Tocheva et al., 2011; Antunes et al., 2016; Tocheva et al., 2016). One possibility is that the ancestor to the Firmicutes phylum is a monoderm, meaning that

68 didermic Firmicutes either evolved through convergent evolution or acquired the outer membrane biosynthetic machinery via horizontal gene transfer. The other possibility is that the ancestor to the phylum is a diderm, so that modern monodermic

Firmicutes arose through some sort of simplification event. Phylogenetic analyses support the latter scenerio, e.g. that the didermic structure is more ancient and much more prevalent across the bacterial domain, even when 16S sequence comparisons robustly place some didermic bacteria in a monodermic phylum (Sutcliffe, 2010;

Tocheva et al., 2011). For instance, an imaging study of the sporulation process in

Acetonema longum, a didermic Firmicutes, hints at sporulation being a key process for the development of an outer membrane (Tocheva et al., 2011). Moreover, comparisons using outer membrane proteins did not indicate a recent divergence, meaning that both sporulation and outer membrane biosynthetic machinery are primitive and widely shared (Tocheva et al., 2011; Tocheva et al., 2016).

Anaerosporomusa subterranea strain RU4 belongs to the Firmicutes phylum and has a didermic cell wall structure that was confirmed through both electron

68 microscopy and the Gram-staining technique (Choi et al., 2016). Through 16S rRNA gene sequencing, the organisms most closely related to A. subterranea are from the

Negativicutes. As a result, we looked into whether this organism represents an ancestral lineage to earliest didermic Firmicutes. The objectives of this study were: (1) to identify genes from the genome of A. subterranea encoding for proteins involved with LPS biosynthesis; (2) to determine if LPS genes shared homology to those in other didermic Firmicutes; and (3) to examine whether LPS genes were acquired by horizontal gene transfer. Our analyses indicate that A. subterranea inherited LPS

69 biosynthesis genes from a Firmicutes ancestor and that the earliest Firmicutes was most likely a didermic bacterium.

Methods

Sequence curation

The methods for genome sequencing of Anaerosporomusa subterranea strain

RU4 were described in Choi et al. (2016). Sequences of proteins involved in outer membrane biosynthesis were first curated from A. subterranea initially through the genome annotation available on NCBI. To confirm, genes were also identified by performing a BLASTp comparison of the protein sequences from E. coli. The protein sequences found in the A. subterranea genome through either method were subsequently obtained from the Negativicutes, Halanaerobiales, each of the 5 orders of the Proteobacteria, the Bacteriodetes, and the Aquificae by searching the UniProt database (http://www.uniprot.org/). When possible, sequences from 5 bacterial members from each clade were obtained. A list of all organisms and gene/protein

69 sequences used in this study with their respective accession numbers are provided in

Table 4.1.

Phylogenetic comparisons

We chose the three LptACD protein sequences to align and compare.

Alignment was done using three different methods: MUSCLE (Edgar, 2004) and

ClustalW (Larkin et al., 2007) through the MEGA 7 software program (Kumar et al.,

2016); and Kalign through the online multiple sequence alignment tool

(http://www.ebi.ac.uk/Tools/msa/kalign/, Lassmann and Sonnhammer, 2005). The subsequent alignments were individually used to create phylogenies in MEGA 7 using the neighbor-joining method with 1000 bootstrap replicates or in Seaview using the parsimony method with 100 bootstrap replicates.

Additionally, the 16S ribosomal RNA gene sequences of all organisms used were compared as a comparison to the Lpt proteins. The 16S sequences were aligned through the Ribosomal Database Project (Cole et al., 2014), and the aligned sequences were then compared phylogenetically in MEGA 7 using the neighbor-joining method with 1000 bootstrap replicates.

Results and Discussion

Identification of genes required for lipopolysaccharide biosynthesis/transport

Out of the over 100 genes encoding for proteins required for biosynthesis, assembly, and export of LPS, 46 genes were identified in the genome of A. subterranea

(Table 4.2). The genome contained genes encoding for LpxABCDIKLP, KdtA/WaaA,

FtsH, and KdsABCD involved in lipid A biosynthesis; GmhAB, HldACDE, WaaC/F, WaaV,

RmlAB, VioA, DdhA, CdgA, ManBC, Gmd, Fcl, Tld, Gna, FnlAC, MnaA, WbaP, WblG, and

WecA for biosynthesis of various oligosaccharides; MsbA which is important for LPS translocation across the cytoplasmic membrane; and LptABCDG for LPS export to the outer membrane. Some of these proteins were derived from annotation on NCBI (i.e.

LptCDG, MsbA, and LpxBL), whereas the rest were determined based on BLASTp sequence identity of approximately 25% or more and a query cover of 70% or greater.

A. subterranea inherited LPS biosynthesis genes from a Firmicutes ancestor

The LptACD protein sequences from A. subterranea were closely related to other didermic Firmicutes species, including Acetonema longum, Anaerovibrio lipolyticus, Propionispora vibrioides, Propionispora hippei, Selenomonas ruminantium,

Megasphaera elsdenii, Acetohalobium arabaticum, Halothermothrix orenii,

Halanaerobium congolense, Halanaerobium kushneri, Halobacteroides halobius, and

Orenia metallireducens. BLASTp results showed up to 51% sequence similarity for

LptA, 46% for LptC, and 47% for LptD between A. subterranea and the most closely related members of the Negativicutes. Thus, the high sequence similarity of these proteins in A. subterranea compared to the Negativicutes suggest definitive homology, as indicated by previous studies that correlated protein homology to sequence similarities (Rost, 1999; Addou et al., 2009). On the other hand, the sequence similarities between A. subterranea and the Halanaerobiales, Proteobacteria, Aquificae, and Bacteroidetes were approximately 30-35% sequence similarity for LptA, 25-30% for LptC, and 22-25% for LptD. These percentages are much lower than within the

Negatvicutes class, corresponding to a decrease in confidence of protein homology among other other didermic bacterial clades.

Phylogenetic comparisons of LptACD

We performed phylogenetic comparisons on the LptACD sequences from didermic Firmicutes to the corresponding sequences in other didermic bacteria.

Initially for all three proteins, alignment was achieved using MUSCLE and the neighbor-joining method. For these analyses, we included as many sequences from

72 the Negativicutes as was available on NCBI to confirm that this group is truly monophyletic as suggested by Fig. 4.2. The resulting trees displayed clustering by taxonomic groups with A. subterranea consistently branching and clustering with the

Negativicutes (Fig. 4.3 to 4.5). In the LptA tree, didermic Firmicutes from the

Negativicutes class all grouped together separately from the Proteobacteria, Aquificae, and Bacteroidetes, a few other didermic clades (Fig. 4.3). In the LptC and LptD trees, the Negativicutes and Halanaerobiales formed a separated group, and A. subterranea again grouped with members from the Negativicutes (Fig. 4.4 and 4.5). The didermic

Firmicutes (Negativicutes and Halanaerobiales) branched separately from other diderms in the Proteobacteria, Aquificae, and Bacteroidetes phyla.

One possibility for the evolution of a didermic cell wall structure in A. subterranea is acquisition of the LPS biosynthesis and export machinery through horizontal gene transfer. Certain genes, such as house-keeping genes, are transferred vertically from mother to daughter cells, but other genes can be transferred between species. In this particular scenario, if didermic Firmicutes like A. subterranea recruited

2 genes from didermic bacteria to produce LPS, then the Firmicutes ancestor would be monodermic. To examine this, we performed numerous amino acid sequence alignments and phylogeny determinations on the LptACD sequences to infer the evolutionary history of these proteins (Fig. 4.3 to 4.5). Analyses of the LptACD sequences done using different combinations of alignment and evolutionary distance computation algorithms all showed similar tree topologies as with the MUSCLE and neighbor-joining method (data not shown). The branching patterns of the Lpt trees resembled that of the 16S rRNA tree, where members from different clades clustered

73 with each other according to their taxonomic grouping (Fig. 4.6). No matter which alignment or phylogenetic algorithm was used, organisms branched and clustered according to their 16S-based taxonomic groups and A. subterranea always clustered with the Negativicutes. Furthermore, the two didermic Firmicutes groups,

Negativicutes and Halanerobiales, consistently grouped together, branching separately from the Proteobacteria, Aquificae, and Bacteroidetes. If horizontal gene transfer had recently occurred, then phylogenetic placement of the didermic microbes based off of the Lpt sequences would not look like a 16S-based phylogenetic tree. This is in contrast to genes that are laterally passed on, such as with antibiotic resistance genes that do not reflect the branching order shown in Fig. 4.6. Only vertical gene transfer from mother to daughter cells would ultimately result in a tree topology similar to that of a 16S tree. Hence, inter-phylum horizontal gene transfers was not detected in our analysis.

The ancestor to the Firmicutes phylum was a didermic, spore-forming bacterium

We examined the hypothesis that the Firmicutes ancestor was a diderm. The question remains as to how the modern monodermic Firmicutes would have evolved from a didermic ancestor. One possibility is that loss of that outer membrane occurred through simplification event, perhaps as an error during sporulation (Cavalier-Smith,

2005; Errington, 2013; Antunes et al., 2016; Munoz-Gomez and Roger, 2016). Like many organisms in the class Negativicutes, A. subterranea is a spore-forming diderm.

However, sporulation is a wide-spread characteristic among members of the

Firmicutes, including the genera Bacillus and Clostridium. Regardless of the cell wall

74 structure of the mother cell, A. subterranea spores and other spores made by members from this phylum go through a shared didermic state when the spores are engulfed (Fig. 4.6). This means that spores formed by members of the Negativicutes or Bacillus species have both an inner and outer spore membrane before the spores fully mature (Tocheva et al., 2011; Tocheva et al., 2016). During spore maturation, the outer spore membrane is either retained if the mother cell was didermic or lost if the mother cell was monodermic, which then translates into a didermic or monodermic cell when the spore germinates at a later point in time. Because of this common characteristic, we believe that A. subterranea and other spore-forming didermic

Firmicutes represent the ancestral lineage.

Another piece of evidence supporting this hypothesis is the relationship between the two didermic Firmicutes groups, the Negativicutes and Halanaerobiales.

If the ancestor was not didermic but monodermic, then these two classes should be each other’s closest relative. Yet, this is not the case; Halanaerobiales is an order of didermic Firmicutes that is most closely related to another monodermic order

Natranaerobiales, whereas the Negativicutes stands as its own class within the phylum and most closely related to the monodermic Clostridiales order (Antunes et al., 2016; Munoz-Gomez and Roger, 2016). Furthermore, our phylogenetic analyses show that these two groups are distinct from each other, despite the fact that they branch together but separately from other didermic clades (Fig. 4.2, 4.S1-2). This indicates that they did not share these genes recently, meaning the genes have ancient origins.

Sequence homology also rules out convergent evolution of LPS biosynthesis

75 in the Firmicutes. An alternate explanation for the presence of didermic cell wall in A. subterranea is convergent evolution from a monodermic ancestor. In this scenario, the machinery necessary for synthesis and export of components of the outer membrane developed from a monodermic Firmicutes to generate a diderm and evolved separately from that of other didermic bacteria. If this is the case, then no sequence homology would be observed and only protein functions would be conserved.

Because the LPS genes of A. subterranea is homologous to other didermic bacteria, we can conclude LPS machinery was inherited from a common ancestor and was not reinvented via convergent evolution.

The results of our study indicate the LPS biosynthesis genes in A. subterranea were inherited from a didermic spore-forming ancestor in the Firmicutes phylum. Our analysis found no evidence of recent horizontal gene transfer or convergent evolution.

We suggest that a simplification event involving the loss of the outer membrane, loss of sporulation, or both gave rise to modern monodermic Firmicutes. Although this model for the origin of didermic Firmicutes has previously been suggested (Tocheva

75 et al., 2011; Antunes et al., 2016) the inclusion of A. subterranea in the analysis strengthens and extends this evolution theory. However, our analysis here was limited to one small subset of proteins involved in the complex process of synthesizing LPS and establishing an outer membrane. Future work could delve into expanding the analyses done here to other LPS components or possibly even other components of the didermic cell wall structure, such as flagellar proteins, porins, or even the peptidoglycan layer. Another interesting research avenue from hereon could be looking at this problem from the monodermic side. In the bacterial domain, there are

76 two monodermic phyla, the Firmicutes and the Actinobacteria. Investigating the evolution of the teichoic acid biosynthesis machinery of the monodermic cell wall in these to clades may also enhance our understanding on the cell wall structure of the last common universal ancestor (LUCA).

Table 4.1. List of organisms used in this study and their respective accession numbers.

Organism Accession numbers Gene/protein analyzed AB122068.2 16S rRNA gene Bradyrhizobium japonicum WP_071917040.1 LptA WP_94184241.1 LptD AB688113.1 16S rRNA gene Brevundimonas abyssalis WP_021697870.1 LptA L26168.1 16S rRNA gene WP_005977802.1 LptA Brucella ovis WP_005977786.1 LptC WP_011950150.1 LptD NR_148263.1 16S rRNA gene Pelagibaca abyssi APZ52785.1 LptA APZ52784.1 LptC NR_043611.1 16S rRNA gene Pelagibaca bermudensis WP_007800295.1 LptD Pleomorphomonas NR_109585.1 16S rRNA gene diazotrophica SFM66571.1 LptC D16428.1 16S rRNA gene WP_013065863.1 LptA Rhodobacter capsulatus SDE65354.1 LptC WP_013068401.1 LptD NR_043150.1 16S rRNA gene SCB34730.1 LptA Rhizobium lusitanum SCB34722.1 LptC

WP_092572809.1 LptD 77

AB787214.1 16S rRNA gene WP_042626035.1 LptA Burkholderia plantarii WP_055139271.1 LptC WP_042635976.1 LptD NR_074658.1 16S rRNA gene Gallionella WP_013292266.1 LptA capsiferriformans WP_013292265.1 LptC WP_013292133.1 LptD NR_074178.1 16S rRNA gene WP_011478508.1 LptA Methylobacillus flagellatus WP_011478507.1 LptC WP_011480370.1 LptD M96399.1 16S rRNA gene WP_011110716.1 LptA Nitrosomonas europaea WP_011110717.1 LptC WP_011111492.1 LptD

Organism Accession numbers Gene/protein analyzed NR_025242.1 16S rRNA gene WP_064801482.1 LptA Ralstonia insidiosa WP_064801489.1 LptC WP_064801677.1 LptD Desulfobacterium NR_041853.1 16S rRNA gene autotrophicum WP_015904533.1 LptD NR_042971.1 16S rRNA gene Desulfobulbus propionicus WP_015725973.1 LptC HQ693571.1 16S rRNA gene Desulfovibrio desulfuricans WP_014321187.1 LptA ND132 WP_014321188.1 LptC WP_014322381.1 LptD NR_121678.1 16S rRNA gene Desulfuromonas WP_005998134.1 LptA acetoxidans WP_005998135.1 LptC NR_042769.1 16S rRNA gene WP_012529301.1 LptA Geobacter bemidjiensis WP_012529300.1 LptC WP_012531775.1 LptD NR_074975.1 16S rRNA gene Pelobacter propionicus WP_011735084.1 LptD NR_075002.1 16S rRNA gene Syntrophobacter WP_015723240.1 LptA fumaroxidans WP_011698924.1 LptC WP_011698527.1 LptD Caminibacter NR_043116.1 16S rRNA gene mediatlanticus TB-2 WP_007474423.1 LptA

L14630.1 16S rRNA gene 78

Campylobacter jejuni WP_002852252.1 LptA WP_044780114.1 LptD NR_115081.1 16S rRNA gene Campylobacter WP_039663423.1 LptA subantarcticus WP_052242975.1 LptD AF357197.1 16S rRNA gene Nautilia profundicola WP_015902489.1 LptA WP_041361562.1 LptD NR_074398.1 16S rRNA gene Sulfuricurvum kujiense WP_013459199.1 LptD Sulfurimonas WP_021286403.1 LptA hongkongensis WP_021287562.1 LptD LT745986.1 16S rRNA gene NP_417667.1 LptA Escherichia coli K12 WP_000030537.1 LptC WP_000746151.1 LptD

Organism Accession numbers Gene/protein analyzed AY360336.1 16S rRNA gene NP_439307.1 LptA Haemophilus influenza NP_439308.1 LptC NP_43889.1 LptD JN166982.1 16S rRNA gene WP_010960073.1 LptA Methylococcus capsulatus WP_010960074.1 LptC WP_050738231.1 LptD LN874213.1 16S rRNA gene NP_253150.1 LptA Pseudomonas aeruginosa NP_253149.1 LptC WP_003113209.1 LptD MF417385.1 16S rRNA gene AKB07198.1 LptA Vibrio cholerae AKB08296.1 LptC KKP19321.1 LptD AJ309733.1 16S rRNA gene Aquifex aeolicus WP_010880598.1 LptD NR_075040.1 16S rRNA gene Desulfurobacterium WP_013638756.1 LptA thermolithotrophum WP_013638300.1 LptD AF188332.1 16S rRNA gene Persephonella marina WP_015898884.1 LptA WP_012675990.1 LptC Sulfurihydrogenibium AF528192.1 16S rRNA gene azorense WP_012673740.1 LptA Sulfurihydrogenibium sp. WP_012458942.1 LptC YO3AOP1 79

NR_075037.1 16S rRNA gene Thermocrinis albus WP_012991902.1 LptA NR_075050.1 16S rRNA gene Thermovibrio WP_013537902.1 LptA ammonificans WP_013538096.1 LptD NR_148855.1 16S rRNA gene Apibacter mensalis WP_055425615.1 LptA AB682428.1 16S rRNA gene Chitinophaga niabensis SIN97345.1 LptA WP_012788181.1 LptC NR_043903.1 16S rRNA gene Cyclobacterium lianum WP_073091835.1 LptA WP_084096949.1 LptC M58768.1 16S rRNA gene Cytophaga hutchinsonii WP_072355958.1 LptD

Organism Accession numbers Gene/protein analyzed Flavisolibacter ginsengisoli NR_041500.1 16S rRNA gene DSM 18119 WP_072834467.1 LptD NR_115084.1 16S rRNA gene Flavobacterium rivuli WP_020212231.1 LptC Flavobacterium NR_115481.1 16S rRNA gene succinicans WP_064714819.1 LptD NR_116978.1 16S rRNA gene Mucilaginibacter mallensis WP_091372263.1 LptD NR_146018.1 16S rRNA gene Paludibacter jiangxiensis GAT62110.1 LptD NR_116376.1 16S rRNA gene Pedobacter glucosidilyticus KHJ39012.1 LptA AB003401.1 16S rRNA gene Prevotella ruminicola SHK68905.1 LptA NR_113879.1 16S rRNA gene Pseudopedobacter saltans WP_013634238.1 LptC NR_074672.1 16S rRNA gene Acetohalobium arabaticum WP_013279107.1 LptC NR_026044.1 16S rRNA gene Halanaerobium congolense SDH27345.1 LptD NR_044807.2 16S rRNA gene Halanaerobium kushneri SIR33398.1 LptA WP_076545691.1 LptC NR_102480.1 16S rRNA gene Halobacteroides halobius WP_015328002.1 LptC WP_015327990.1 LptD NR_074915.1 16S rRNA gene

Halothermothrix orenii WP_015923471.1 LptC 80 WP_015923462.1 LptD KP898732.2 16S rRNA gene Orenia metallireducens WP_068719515.1 LptC WP_068719502.1 LptD NR_041951.1 16S rRNA gene Acetonema longum WP_004097296.1 LptD KX268498.1 16S rRNA gene Anaerosporomusa WP_066241767.1 LptA subterranea RU4 WP_066241764.1 LptC WP_066241828.1 LptD AB034191.1 16S rRNA gene Anaerovibrio lipolyticus SHI90709.1 LptA DSM 3074 SHI90729.1 LptC NR_025418.1 16S rRNA gene Propionispora vibrioides SEO72936.1 LptD

Organism Accession numbers Gene/protein analyzed LC036319.1 16S rRNA gene WP_071884200.1 LptA Megasphaera elsdenii WP_014015052.1 LptC SF108572.1 LptD NR_036875.1 16S rRNA gene Propionispora hippei DSM SHI87561.1 LptA 15287 SHI87529.1 LptC AF161581.1 16S rRNA gene WP_073089097.1 LptA Selenomonas ruminantium WP_072305824.1 LptC WP_051598467.1 LptD

Table 4.2. List of enzymes required for outer membrane biosynthesis and genes present in A. subterranea.

Gene Protein function References Present in RU4?

LpxABCDHKLM Lipid A biosynthesis LpxABCDKL

LpxI Homolog of LpxH Yes

LpxP Homologous to LpxL Stead et al. Yes (2011) KdtA/WaaA Lipid A biosynthesis Yes Post-transcriptional FtsH regulation of Lipid A Yes biosynthesis Lipid core biosynthesis KdsABCD and attachment to lipid Yes A (Mamat et al., 2011) Oligosaccharide GmhAB Yes biosynthesis Samuel and Oligosaccharide HldACE, Reeves (2003); biosynthesis for core Yes HldD/GmhD Mamat et al. and O-antigen (2011) Oligosaccharide Mamat et al. One (WaaC or

WaaCF incorporation onto lipid 82 (2011) WaaF)

A Biosynthesis and Raetz and WaaQGPOTYWVL modification of outer Whitfield Potential WaaV core (2002) Positive regulation of Mamat et al. RfaH No core gene expression (2011) O-antigen sugar RmlABCD biosynthesis (E. coli, S. RmlAB enterica, and others) O-antigen sugar biosynthesis (E. coli, P. Samuel and Tll No aeruginosa, A. Reeves (2003) actinomycetemcomitans) O-antigen sugar VioAB VioA biosynthesis in E. coli

Gene Protein function Reference Present in RU4? O-antigen sugar Fcd biosynthesis in A. No actinomycetemcomitans Synthesis of FdfAB enterobacterial common No antigen O-antigen sugar biosynthesis (S. enterica, DdhABCD DdhA Y. pseudotuberculosis, V. anguillarum) O-antigen sugar CdgAB biosynthesis in Y. CdgA enterocolitica O8 O-antigen sugar ManABC biosynthesis (E. coli, S. ManBC enterica, and others) O-antigen sugar Gmd and Gmm biosynthesis (E. coli, S. Gmd enterica, and others) O-antigen sugar biosynthesis (E. coli Fcl and Per Fcl O157, V. cholera O1, B. Samuel and melitensis) Reeves (2003) O-antigen sugar ColAB biosynthesis (S. enterica No O35, E. coli O111)

O-antigen sugar 83

Rmd biosynthesis in P. No aeruginosa O-antigen sugar Tld biosynthesis in A. Yes actinomycetemcomitans O-antigen sugar GalE and Gne No biosynthesis in E. coli O-antigen sugar Gna biosynthesis in P. Yes aeruginosa O6 O-antigen sugar FnlABC biosynthesis (E. coli O26 FnlAC and P. aeruginosa O11) O-antigen sugar WbpKM biosynthesis in P. No aeruginosa

Gene Protein function References Present in RU4? O-antigen sugar MnaAB biosynthesis (E. coli, S. MnaA enterica, and more) O-antigen sugar NnaABCD biosynthesis (E. coli and No N. meningitis) O-antigen sugar biosynthesis (Y. DmhAB No pseudotuberculosis and B. mallei) O-antigen synthesis (S. WbaPZ WbaP enterica and E. coli) O-antigen synthesis (P. WbgY No shigelloides and E. coli) Samuel and O-antigen synthesis in E. WcaJ Reeves (2003) No coli O-antigen synthesis in B. WblG Yes pertussis O-antigen synthesis in WecA Yes Enterobacteriaceae O-antigen synthesis in P. WbpL No aeruginosa O-antigen synthesis in E. WbpP No coli O-antigen processing

Wzx, Wzy, Wzz No 84 and translocation

O-antigen assembly and Wzm, Wzt No export Ruiz et al. MsbA LPS translocation Yes (2009) LPS transport and Dong et al. LptABCDEFG LptABCDG export (2014)

Figure 4.1. LPS transport pathway in didermic microorganisms. Figure modified from Dong et al. (2014). The Lpt system in didermic bacteria is responsible for

85 transporting mature lipopolysaccharide (LPS) from the inner membrane (IM, or cytoplasmic membrane) across the periplasm to the outer membrane (OM). Genes encoding for LptABCDG were found in the A. subterranea genome.

Figure 4.2. Phylogenetic placement of A. subterranea within the Negativicutes class based on LptACD protein sequences. Available sequences from the NCBI database for the (a) LptA, (b) LptC, and (c) LptD proteins were curated for A. subterranea (highlighted in red boxes) and other members from the Negativicutes class. Sequences shared approximately 50% similarity with the closest relatives, and phylogenetic analyses were consistent for all three protein sequences.

Figure 4.3. Phylogenetic analysis of the LptA protein from didermic clades. The sequences of the LptA protein from representatives of didermic phyla were compared and analyzed. The scale bar represents the number of amino acid changes for the allotted length, and numbers at branch nodes the bootstrap percentages. Organisms from different clades are boxed according to color, with A. subterranea in a black- outlined box.

Figure 4.4. Phylogenetic analysis of the LptC protein from didermic clades. The sequences of the LptC protein from representatives of didermic phyla were compared and analyzed. The scale bar represents the number of amino acid changes for the allotted length, and numbers at branch nodes the bootstrap percentages. Organisms from different clades are boxed according to color, with A. subterranea shown in the black-outlined box.

Figure 4.5. Phylogenetic analysis of the LptD protein involved in lipopolysaccharide transport in didermic clades. The LptD protein sequences from representatives of didermic phyla were compared. The scale bar represents the number of amino acid changes for the allotted length, and numbers at branch nodes the bootstrap percentages. Organisms from different clades are boxed according to color, and A. subterranea shown in the black-outlined box.

Figure 4.6. 16S ribosomal RNA gene sequence based phylogeny of didermic bacteria. The widely-accepted phylogeny based off of the 16S small ribosomal subunit sequence was performed in MEGA 7 using the neighbor-joining method with

1000 bootstrap replicates. Organisms are boxed according to their respective clades.

Figure 4.7. Sporulation process in didermic and monodermic Firmicutes. Figure modified from Errington (2013). The process of spore formation and maturation in diderm and monoderms are shown. All spores in this phylum pass through a common didermic state before spore maturation, upon which spores formed by monodermic bacteria somehow lose the outer spore membrane. The monodermic or didermic mature spores then give rise to new vegetative cells with the respective cell wall structures.

CHAPTER 5

Conclusions and future directions

In Chapter 2 of this thesis, a mechanism for Fe(III) oxide reduction by three

Clostridium species – Clostridium sp. FGH, Clostridium beijerinckii, and Clostridium acetobutylicum – was described. These fermentative Firmicutes conduct this activity in a different way from the canonical dissimilatory iron reducing bacteria Shewanella and Geobacter. They do not require direct contact for Fe(III) oxide reduction, with genomic investigation portraying a lack of the characteristic multi-heme c-type cytochromes from multiple Clostridium species supporting this statement. Though these fermenters produce lots of organic acids from fermentation, the organic acids naturally do not solubilize Fe(III), and supplementation of cultures with additional organic acids did not seem to stimulate Fe(III) oxide reduction. However, addition of exogenous FAD, a flavin molecule, to fermenting and Fe(III)-reducing cultures showed an increase in Fe(III) reduction with all three Clostridium species. This suggests a role

98 for this molecule in mediating Fe(III)-reducing activity.

These results are significant because the findings highlight one process

Clostridium species use to reduce Fe(III) available in the environment during fermentation. This has numerous geochemical implications. In sediments and soils, the presence of Fe(III) may bolster growth of fermentative Clostridium species, which could result in syntrophic events in a complex microbial environment such as sediments and soils. Organisms such as C. beijerinckii and C. acetobutylicm are widely studied for their solventogenic properties that are dependent on their growth. If

99 growth can be enhanced when Fe(III) is provided, then microbial solvent production can be significantly and concurrently increased. Clostridium species also have applications towards bioremediation of contaminated environments. Clostridium sp.

FGH, for instance, was isolated from Oak Ridge, TN, that is heavily polluted with heavy metals. Because Clostridium sp. FGH demonstrated an ability to reduce Fe(III) oxides, it also has the potential capability to reduce other heavy metals possibly via FAD.

Because Clostridium species are generally found ubiquitously in subsurface environments, they could be utilized toward bioremediation efforts in contaminated environments such as Oak Ridge. To the best of our knowledge, this is the first study showing evidence of an indirect mechanism of Fe(III) reduction in Clostridium species involving FAD.

There are a considerable number of scientific research questions that could be studied further in the future. For instance, it would be critical to determine when

FAD is released into the environment, how much FAD is released, and the genetic controls underlying FAD accumulation if it is actively secreted extracellularly by

Clostridium cells. Another question that could be interesting is to see if Fe(III)- reducing Clostridium species exhibit any electrical activity. Organisms such as

Shewanella were able to transfer electrons to an electrode, and so it would be intriguing to determine if these microorganisms are able to do the same. Currently, we are collaborating with USC to quantify FAD production by Clostridium species and looking for electrochemical activity. Lastly, our study was limited to just three organisms from this genus. However, there are many other species that have been

100 shown capable of reducing Fe(III). It would be fascinating to examine whether all

Clostridium species manifest the same characteristics regarding Fe(III) reduction.

In Chapter 3, the isolation, characterization, and identification of an organism representing a novel species and genus, named Anaerosporomusa subterranea was reported. This organism was first found in co-culture with Clostridium sp. FGH. As an isolate in pure culture, A. subterranea were motile curved rods that displayed a didermic cell wall structure. The organism is a mesophile, a neutrophile, and an obligate fermenter. It was interesting to observe a very narrow metabolic niche for this organism, as it could grow by fermentation on two carbon substrates, fumarate and maleate. A. subterranea could not utilize any terminal electron acceptor, which was unusual considering that the organism first originated from a Fe(III)-reducing enrichment culture.

An interesting aspect about this organism is its meticulousness regarding growth substrate utilization. It is possible that this study did not test enough carbon sources. To truly examine whether the organism can metabolize only two carbon

0 10

sources, a wider variety of carbon substrates could be tested. Additionally, metabolism using fumarate as the electron donor is quite unique. Fumarate is commonly used as a TCA cycle intermediate or as an electron acceptor. It would be fascinating to map out the pathway of fermentation using fumarate as the growth substrate, including what products are formed from this pathway.

Phylogenetic comparisons using the 16S rRNA and other housekeeping gene sequencings robustly placed A. subterranea within the Negativicutes class of the

Firmicutes phylum, despite the fact that it is a didermic microorganism. Because of

101 this, we considered the evolution of A. subterranea as a didermic member of a monodermic phylum in Chapter 4. Proteins involved in the biosynthesis of lipopolysaccharide (LPS), a major component of the didermic outer membrane, were identified in the genome of A. subterranea. Specifically, the proteins involved in transport of LPS from the inner to the outer membrane LptACD shared between 25-

50% amino acid sequence similarities with other members of the Negativicutes. This high sequence similarity denotes protein homology, meaning that these proteins in A. subterranea are ancestrally derived. In addition, when the phylogenetic comparisons were expanded to include didermic bacteria from other phyla, such as the

Proteobacteria, Aquificae, and Bacteriodetes, organisms clustered together according to their 16S-based classification. Tree topologies using the LptACD protein sequences were comparable to that of the 16S gene sequence tree. This demonstrates that horizontal gene transfer between these different microbial groups did not occur, meaning that A. subterranea did not acquire the LPS export machinery from other organisms. Therefore, we predict that the ancestor to the Firmicutes phylum was a

1 10

didermic organism and that A. subterranea represents this lineage.

Our study and investigation into the evolution of A. subterranea was limited in that out of 46 LPS biosynthesis related proteins identified, only 3 were analyzed. It would be beneficial to be able to analyze other proteins involved, and possibly other proteins related to the cell wall such as peptidoglycan biosynthesis, outer membrane porins, or flagellar proteins. Analysis of additional proteins would provide a more comprehensive and in-depth survey into the evolution of A. subterranea and other didermic Firmicutes.

102

Like Clostridium species, A. subterranea and other members of the

Negativicutes are prevalent in the environment. Since A. subterranea was isolated from Oak Ridge, Tennessee, we can most likely infer that A. subterranea has some sort of biogeochemical role. Since it is an obligate fermenter, it could be responsible for fermenting and breaking down more complex carbon sources into simpler molecules that could serve as the growth substrates for other microbes in the same community.

For instance, a common fermentation product is acetate, which Geobacter species use as their primary growth substrates. If organisms such as A. subterranea are able to boost growth of Geobacter species, there could be significant bioremediation effects in contaminated environments.

The results of this thesis have provided new insights into the geochemical importance of fermentative organisms from the Firmicutes phylum. These organisms not only have biogeochemical impacts that are environmentally relevant, but can also help probe questions about the evolution of bacterial species and inquire as to the characteristics of the last common universal ancestor. These results have hopefully

2 10

opened up more avenues for future research regarding fermentative Firmicutes.

103

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