Degrading Enrichment Cultures

Community Diversity and Functional Capabilities of Benzene-Degrading Enrichment Cultures

Sarah Elizabeth McRae

A thesis submitted in conformity with the requirements for the degree of Master of Applied Science Graduate Department of Chemical Engineering and Applied Chemistry University of Toronto

Community Diversity and Functional Capabilities of Benzene- Degrading Enrichment Cultures

Sarah McRae

Master of Applied Science

Department of Chemical Engineering and Applied Chemistry University of Toronto

2015 Abstract

Benzene is prevalent, toxic, and persistent in anaerobic environments. Anaerobic benzene degradation has been proven, but is not well understood. The goals of this research were to determine the community composition of anaerobic nitrate-reducing benzene-degrading enrichments and propose likely metabolic roles of organisms present. Highly diverse communities were present in the enrichments, with multiple organisms capable of carrying out similar roles. A metagenome assembly was generated from a nitrate-reducing, benzene- degrading enrichment culture. Further evidence for fermenting Peptococcaceae as the initiator of benzene degradation was established. The potential for benzene degradation to be linked to other terminal electron acceptors (i.e., CO2) was confirmed, further supporting that the initial benzene degradation steps are carried out in a fermenting syntroph. An understanding of the functions of different organisms can inform culturing techniques to increase culture growth and benzene degradation rates. Genes present in the metagenome could be used as biomarkers for benzene degradation.

Acknowledgments

I would like to thank the many people who have encouraged, helped and taught me so much during this project. My advisor, Elizabeth Edwards has provided crucial advice, support and taught me so much over the past two and half years. Lab mates in Edwards’ Laboratory have been essential sources of information, technical support, and enthusiasm. Cheryl Devine and Fei Luo were especially helpful throughout and gave advice on nearly every aspect of this project, and I want to thank them for it. This project builds on their work, and the work of other former EdLab students who created and maintained these cultures over the years, and whose research guided me. I also want to thank Shuiquan Tang and Olivia Molenda, from whom I learned sequence analysis and metagenome assembly techniques. Xiaoming Liang taught me key techniques at the beginning. Other members of EdLab, both past and current, have given me advice, shared discussions and been generally wonderful. I thank Luz Puentas Jacome, Po Hsiang (Tommy) Wang, Nigel Guilford, Kirill Krivushin, Line Lomheim, Ivy Yang, Mabel Ting Wong, Torsten Meyer, Sam Huang, Mahbod Hajighasemi, Wendy Han Zhou, and Chris Tran. Endang Susilawati, our wonderful BioZone manager always provided support. BioZone has been a fantastic environment, full of people who are full of bright ideas and always willing to help, whether with advice, technical support, or reinforcement. I especially want to thank Kayla Nemr for her support and editing of this thesis. Financial support for this project was provided by BEEM (Bioproducts + Enzymes from Environmental Metagenomes) and the University of Toronto.

As always, I could not have done this without the love and encouragement of my wonderful family: Marie-Thérèse and Malcolm, Daniel, Jeremy and Michelle, Clare, and Rebecca. You have cheered me on from all corners of the globe, given me confidence to challenge myself and kept telling me I could do this. Thank you for everything.

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Table of Contents

ACKNOWLEDGMENTS III

TABLE OF CONTENTS IV

LIST OF TABLES VII

LIST OF FIGURES VIII

LIST OF ABBREVIATIONS X

LIST OF APPENDICES XI

CHAPTER 1 LITERATURE REVIEW 1

1.1 INTRODUCTION 1

1.2 REMEDIATION OPTIONS 2

1.3 ANAEROBIC BIOREMEDIATION OF BENZENE 4 1.3.1 Methylation 5 1.3.2 Hydroxylation 6 1.3.3 Carboxylation 7 1.3.4 Central benzoate metabolism pathway 9 1.3.5 Syntrophy 10 1.3.6 Organisms implicated in benzene degradation 10 1.3.7 Metagenome assembly 11

CHAPTER 2 BACKGROUND 12

2.1 RATIONALE FOR RESEARCH 12

2.2 HYPOTHESES 12

2.3 OBJECTIVES 13

2.4 APPROACH 13

2.5 ENRICHMENT CULTURES USED IN THIS STUDY 13

CHAPTER 3 METHODS 17

3.1 CULTURE MAINTENANCE 17

3.2 CULTURE SET-UP 17

3.3 GAS CHROMATOGRAPHY 18

3.4 ION CHROMATOGRAPHY 19

3.5 DNA EXTRACTION AND SEQUENCING FOR METAGENOMIC ANALYSIS 19

3.6 PYROTAG SEQUENCING 20 iv

CHAPTER 4 PYROTAG SEQUENCING TO DETERMINE COMMUNITY COMPOSITION 21

4.1 INTRODUCTION 21

4.2 RESULTS AND DISCUSSION 23 4.2.1 Alpha diversity of samples 23 4.2.2 Relative abundances of various OTUs 24 4.2.3 Relatedness of samples 29 4.2.4 Phylogenetic grouping of OTUs 30 4.2.5 Comparison to other benzene-degrading enrichment cultures 33 4.2.6 Comparison of OTUs to culture parameters 34 4.2.7 Multiple OTUs of a single phylogeny 37

4.3 KEY POINTS 39

CHAPTER 5 METAGENOMIC ANALYSIS OF A NITRATE-REDUCING BENZENE-DEGRADING CULTURE 40

5.1 INTRODUCTION 40

5.2 RESULTS AND DISCUSSION 41 5.2.1 ABySS Assembly 42 5.2.2 ALLPATHS-LS Assembly 45 5.2.3 Assessment of assemblies 50 5.2.4 Multiple Peptococcaceae present in the enrichment? 51 5.2.5 Identification of putative anaerobic benzene carboxylase operon 52 5.2.6 Identification of other enzymes in the anaerobic benzene degradation pathway 55

5.3 KEY POINTS 56

CHAPTER 6 METHANOGENIC CAPABILITY OF NITRATE-REDUCING BENZENE-DEGRADING CULTURE 57

6.1 INTRODUCTION 57

6.2 RESULTS 59

6.3 DISCUSSION 67

6.4 KEY POINTS 67

CHAPTER 7 CONCLUSIONS AND ENGINEERING SIGNIFICANCE 70

7.1 A HIGHLY DIVERSE COMMUNITY WITH FUNCTIONAL REDUNDANCY IS PRESENT 70

7.2 THE PUTATIVE ANAEROBIC BENZENE CARBOXYLASE “ABC” IS RESPONSIBLE FOR THE INITIATION OF

BENZENE DEGRADATION 71

7.3 NITRATE-REDUCING BENZENE DEGRADING ENRICHMENTS CAN BE SWITCHED TO METHANOGENESIS 71

7.4 FUTURE DIRECTIONS SUGGESTED 72

7.5 ENGINEERING SIGNIFICANCE AND CONTRIBUTIONS OF THIS RESEARCH 73

REFERENCES 74

APPENDIX 1: ASSESSMENT OF GENOME COMPLETENESS 90

APPENDIX 2: DEGRADATION OF MONOCHLOROBENZENE 96

APPENDIX 3: ATTEMPTED ANAMMOX ENRICHMENT 105

APPENDIX 4: MICROSCOPIC TECHNIQUES TO EXAMINE COMMUNITY 109

APPENDIX 5: LOCATION OF SEQUENCE DATA FILES 119

List of Tables

Table 3.1: Experimental set-up for testing the potential of CO as a terminal electron 2 18 acceptor in nitrate-reducing benzene-degrading enrichments Table 4.1: 18 OTUs common to all nitrate-reducing benzene-degrading enrichments 28 tested, with percent abundances in all samples

Table 4.2: Groups present during benzene degradation with different electron acceptors, compared to the presence or absence at greater than one percent 33 abundance in pyrotag sequencing data

Table 4.3: Percent abundances of methanogenic OTUs in consortia 34 Table 4.4: Metadata used in analysis of Pyrotag sequencing data 35 Table 5.1: Scaffolds that contain a 16S rRNA sequence identified from 46 ALLPATHS-LG assembly using digitally normalized data Table 5.2: Functions ascribed to genera present in assembly 48 Table 5.3: Proportions of genes putatively identified by myRAST 50

Table 5.4: Proportions of genes from single-copy gene set putatively identified by 50 myRAST

Table 5.5: Putative annotation of protein encoding genes on Scaffold 559. The four 53 subunits of the abc operon are present Table 5.6: Presence of genes encoding enzymes responsible for different stages of 55 anaerobic benzene degradation in selected scaffolds

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List of Figures

Figure 1.1: Anaerobic toluene degradation pathway, with metabolite and genes 6 involved shown Figure 1.2: Anaerobic phenol degradation pathway 7 Figure 1.3: Anaerobic carboxylation of benzene to benzoate 8 Figure 1.4: Anaerobic benzoyl-CoA degradation pathway 9 Figure 2.1: Approximate lineage of Cartwright enrichment cultures 15 Figure 2.2: Approximate lineage of Swamp enrichment cultures 16 Figure 4.1: Approximate lineage and benzene degradation rates of cultures submitted 22 for pyrotag sequencing Figure 4.2: Alpha diversity of pyrotag samples 23 Figure 4.3: Pyrotag sequencing results showing percent abundances of different OTUs 24 in a selection of cultures Figure 4.4: Venn diagram showing the number of OTUs at greater than one percent 27 abundance that are shared between, or unique to, different cultures sampled Figure 4.5: Bootstrap tree using UPGMA (Unweighted Pair Group Method with 29 Arithmetic mean) values to show clustering of samples Figure 4.6: Phylogenetic tree of Beta- and Gammaproteobacteria showing pyrotag sequences in blue, earlier sequences from these cultures in red, and root sequence in 31 purple Figure 4.7: Phylogenetic tree of all groups outside Beta- and Gammaproteobacteria showing pyrotag sequences in blue, earlier sequences from these cultures in red, and 32 root sequence in purple Figure 4.8: NMDS showing clustering of all pyrotag OTUs and the correlation of 36 benzene degradation rate with OTUs Figure 4.9: NMDS showing clustering of pyrotag OTUs present at greater than one 37 percent abundance in any of the enrichments, and the correlation with generation Figure 4.10: Geneious alignment of Peptococcaceae OTUs 38 Figure 5.1:a) ABySS assembly using four paired end libraries (k96, c20) 43 Figure 5.1:b) ABySS assembly using one paired end library (k96, c20) 43 Figure 5.1:c) ABySS assembly using one paired end library (k96, c10) 44

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Figure 5.1:d) ABySS assembly using one paired end library (k96, c100) 44 Figure 5.2a) Scaffold 29 shows 69% pairwise identity to Thermincola potens genome 51 Figure 5.2b) Scaffold 39 shows 26% pairwise identity to Thermincola potens genome 51 Figure 6.1: Nitrate-reducing benzene degrading culture only (without nitrate). Replicate 59 A Figure 6.2: Nitrate-reducing benzene degrading culture only (without nitrate). Replicate 60 B Figure 6.3: Nitrate-reducing benzene degrading culture only (without nitrate). Replicate 60 C Figure 6.4: Nitrate-reducing benzene culture blended with methanogenic toluene- 61 degrading culture. Amended with nitrate on Day 325. Replicate A Figure 6.5: Nitrate-reducing benzene culture blended with methanogenic toluene- 61 degrading culture. Amended with nitrate on Day 325. Replicate B Figure 6.6: Nitrate-reducing benzene culture blended with methanogenic toluene- 62 degrading culture. Amended with nitrate on Day 325. Replicate C Figure 6.7: Methanogenic toluene-degrading culture only. Amended with toluene on 62 Day 242. Replicate A Figure 6.8: Methanogenic toluene-degrading culture only. Amended with toluene on 63 Day 242. Replicate B Figure 6.9: Methanogenic toluene-degrading culture only. Amended with toluene on 63 Day 242. Replicate C Figure 6.10: Autoclaved control. Replicate A 64 Figure 6.11: Autoclaved control. Replicate B 64 Figure 6.12: Autoclaved control. Replicate C 65

List of Abbreviations

ABC - anaerobic benzene carboxylase BLAST - basic local alignment search tool BP – base pairs FID - flame ionization detector GC - gas chromatography GMO - genetically modified organism IC - ion chromatography NCBI - National Center for Biotechnology Information NKD - normalized kmer depth NMDS - non-metric multidimensional scaling MP – mate pair MCL - maximum contaminant level OTU - operational taxonomic unit ORF – open reading frame PE – paired end PCR - polymerase chain reaction PEG - protein-encoding gene PPC - phenylphosphate carboxylase PPS - phenylphosphate synthase qPCR - quantitative PCR rRNA - ribosomal ribonucleic acid

List of Appendices

Appendix 1: Assessment of genome completeness 90

Appendix 2: Degradation of monochlorobenzene 96

Appendix 3: Attempted Anammox enrichment 105

Appendix 4: Microscopic techniques to examine microbial communities 109

Appendix 5: Location of sequence data files 119

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Chapter 1 Literature Review

1.1 Introduction

Benzene is an environmental pollutant of great concern. It has been recognized as a human carcinogen and cause of leukemia and reproductive concerns since the 1940s, and yet remains a widespread contaminant (Clinton, 1948, Eckardt, 1973, Schwarzenbach et al., 2006). It is one of the most common contaminants at Superfund sites (U.S. Environmental Protection Agency, 2011), and its maximum contaminant level in groundwater (MCL) is 0.005mg/L in Canada and the U.S. (Health Canada, 2009, U.S. Environmental Protection Agency, 2009). Benzene is a component of crude oil, and as such is present in petroleum. Due to the huge volumes of petroleum transported and stored throughout industrialized nations, significant amounts of benzene are released to the environment through fuel leakages and spillages, despite its concentration in petroleum being limited at 1.3% by volume in the U.S. (U.S. Environmental Protection Agency, 2007) and at 1.5% by volume in Canada (Government of Canada, 1997). Leakage from underground storage tanks is a significant source of contaminants to groundwater (U.S. Environmental Protection Agency, 2008). Benzene is also produced industrially and used as a chemical precursor and as a solvent, which contributes another source of benzene to the environment (U.S. Environmental Protection Agency, 1993). Benzene may naturally occur in groundwater due to leakage from petroleum reservoirs, though oil extraction may increase benzene movement (Landon & Belitz, 2012).

Benzene leakage to groundwater from underground storage tanks is a significant and recalcitrant problem. Benzene is soluble in water (1780mg/L (MacLeod & Mackay, 1999)), and so spreads through groundwater from the source of contamination, causing large contaminant plumes. In the presence of oxygen, benzene is rapidly degraded by ubiquitous aerobic bacteria (Salanitro, Wisniewski, Byers, Neaville, & Schroder, 1997). Consumption of oxygen by aerobic bacteria during benzene degradation can deplete dissolved oxygen resulting in anaerobic

conditions in subsurface environments. Recharge of groundwater with fresh oxygenated water from above ground sources is slow, causing anoxic conditions to remain in place for decades after contaminant sources are removed in some circumstances (Smith, Repert, Barber, & LeBlanc, 2013). This causes benzene attenuation to slow or stall, as anaerobic benzene degradation has a slow rate and is less ubiquitous compared to aerobic benzene degradation. 30% of Canadians (8.9 million people) rely on groundwater as their sole source of drinking water (Government of Canada, 2013), making water contamination a key issue for public health.

1.2 Remediation options

A variety of in situ and ex situ remediation technologies are currently used for near-surface benzene remediation of various contaminants. Unsaturated soils in the vadose zone can be treated by soil vapour extraction, where gas flow is induced in the sub-surface by vacuum pumps, and contaminated vapour is collected and treated above-ground (Soares et al., 2010). In a similar treatment, thermal desorption can be carried out, where the subsurface is heated to cause desorption of contaminants from sediment particles, and vapour then collected and treated. Chemical oxidants can be applied to contaminated soils and water by pumping them into the contaminated zone to remediate many organic contaminants (Ojinnaka et al., 2012). Groundwater and soils can also be sparged to provide oxygen for aerobic benzene remediation (Zhang et al., 2012). An ex situ remediation technique is "pump and treat", where contaminated groundwater is pumped to the surface and through treatments such as granulated activated carbon filters, chemical oxidation or photo catalytic systems (Farhadian et al., 2008). Physical removal of the contaminated soils, “dig and dump”, can also be carried out. Removed soils can be stored in sealed landfills or treated aerobically, or incinerated. All these methods are energy and resource intensive, generally require sizeable infrastructure, and result in further disturbances of the sites. Furthermore, deeper or more widespread contamination is more difficult to treat as it is expensive and energy-intensive to access contamination further underground, or to oxygenate deep groundwater.

Bioremediation could provide an inexpensive, environmentally-safe method for benzene remediation. Bioremediation harnesses the abilities of organisms to degrade environmental contaminants. Different approaches are taken to this. In sites where organisms exist that are capable of degrading the contaminants present, and they have all the resources they require, natural attenuation may be observed. No intervention is required where natural attenuation is proceeding at an acceptable rate. Anthropogenic influence is limited to monitoring groundwater to ensure that contaminant levels are decreasing, and that this decrease is due to degradation rather than transport of the contaminants into other phases or down gradient. Bioattenuation can be observed directly though the reduction of contaminants, observation of degradation products, or indirectly through monitoring the growth of the organisms responsible via the abundances of genes known as biomarkers (Patel & Tolley, 2014, Key et al., 2014).

For some contaminants, the microorganisms required for biodegradation may be so specialized that they are not ubiquitous in the environment. In these cases, bioaugmentation, the addition of cultured degradation-specialized microorganisms into the subsurface may be required (Bento, Camargo, Okeke, & Frankenberger, 2005). Wild-type organisms are preferred to genetically modified organisms (GMOs), because there is more regulatory and public acceptance of non-GMOs.

In other cases, the microorganisms involved in degradation may deplete a required resource before contaminant removal is complete. Limiting resources may be electron acceptors such as oxygen, sulphate, ferric iron and nitrate or electron donors such as an organic carbon source. Biostimulation is the process in which the limiting nutrient is added to the contaminated zone to stimulate the growth of the required microorganisms (eg. Ramos, da Silva, Chiaranda, Alvarez, & Corseuil, 2013, Ponsin, Coulomb, Guelorget, Maier, & Höhener, 2014). In the case of biostimulation by oxygenation, it requires a great deal of energy input, and the resultant aerobic growth can quickly clod aquifers and the oxygen injection ports. Anaerobic processes yield less biomass, so are preferable both for this reason and because of the difficulties of oxygenating the subsurface environment.

Benzene biodegrades readily under aerobic conditions, and the pathway/genes involved in this process have been determined (reviewed in Jindrova, Chocova, Demnerova, & Brenner, 2002). Key genes are mono- or dioxygenases that add oxygen atoms across carbon double bonds. Anaerobic benzene degradation is much more difficult. The pathways, organisms and genes involved in anaerobic benzene degradation are consequently less well-known. Due to the previously mentioned difficulties involved in providing oxygen to the subsurface environment when oxygen is depleted, the anaerobic process has relevance to contaminated sites. A better understanding of anaerobic benzene degradation pathways, genes and organisms could lead to methods to readily quantify anaerobic benzene degradation in the environment. Protocols for stimulating anaerobic benzene degradation could be developed, and enrichments or isolates capable of anaerobic benzene degradation could be produced that can be used for bioaugmentation. Section 1.3 covers the current knowledge of potential anaerobic benzene degradation pathways and the organisms and communities implicated in this process.

1.3 Anaerobic bioremediation of benzene

Benzene is a relatively stable molecule, consisting of a single aromatic ring with no substituents to provide a point of attack. The resonance stability of benzene causes it to have a large activation energy barrier to overcome. Despite this, several cultures have been proven capable of anaerobic benzene degradation since the first confirmation of methanogenic benzene degradation in 1986 (Vogel & Grbić-Galić, 1986). Living cells gain energy through redox reactions that release energy they can capture and use to fuel growth and cell processes. Benzene provides a source of electrons, but an electron acceptor is also required for biological benzene degradation. Enrichments have been established which degrade benzene with the alternative electron acceptors nitrate, iron, sulfate and methanogenesis (Burland & Edwards, 1999, Lovley, Woodward, & Chapelle, 1994, Lovley, Coates, Woodward, & Phillips, 1995, and Kazumi, Caldwell, Suflita, Lovley, & Young, 1997). End products of degradation are methane and carbon dioxide, where no external terminal electron acceptors are provided. Where external electron acceptors are used, reduced forms of these electron acceptors will be produced. A number of

laboratories currently maintain anaerobic benzene degrading enrichments, and isolates capable of anaerobic benzene degradation have been reported under nitrate-reducing conditions (Coates et al., 2001, Kasai, Takahata, Manefield, & Watanabe, 2006) and iron-reducing conditions (Holmes, Risso, Smith, & Lovley, 2011, T. Zhang, Bain, Nevin, Barlett, & Lovley, 2012). The focus of this thesis is a selection of nitrate-reducing benzene-degrading enrichments, which will be further discussed in chapter 2.

Different potential intermediates and pathways have been proposed over the years, in light of developing evidence. Possible initial intermediates of benzene degradation such as toluene, phenol and benzoate have been detected under different conditions (Ulrich, Beller, & Edwards, 2005, Chakraborty & Coates, 2005, Vogel & Grbić-Galić, 1986, Caldwell & Suflita, 2000, and Mancini et al., 2008). The presence of these metabolites suggest mechanisms of methylation, hydroxylation and carboxylation respectively. Each of these possible mechanisms will be addressed, and the central benzoate metabolic pathway reviewed. Groups of organisms implicated in benzene degradation under different conditions will also be covered.

1.3.1 Methylation

Methylation involves the addition of a methyl group to the benzene ring to form toluene. Toluene has been detected under nitrate-reducing conditions (Ulrich et al., 2005). 13C-labelled toluene was detected when 13C-labelled benzene was provided to the culture. 13C-labelled benzoate was also detected, suggesting that benzene → toluene → benzoate was a potential path in this culture. No enzyme/gene was proposed for the initial reaction. The downstream degradation pathway of toluene is well established, and shown with genes involved in Figure 1.1 from (Heider, Spormann, Beller, & Widdel, 1999). The initial step of toluene degradation is the addition of the methyl carbon across the double bond of fumarate. This step is catalyzed by benzylsuccinate synthase (BSS). If the toluene →→ benzoate pathway is followed, BSS is anticipated in the genome of the organism(s) responsible.

Figure 1.1: Anaerobic toluene degradation pathway, with metabolite and genes involved shown (Figure from (Heider et al., 1999))

1.3.2 Hydroxylation

Hydroxylation involves the addition of a hydroxyl group to form phenol. Phenol has been detected in numerous studies over the years, suggesting an initial hydroxylation step (Vogel & Grbić-Galić, 1986, Caldwell & Suflita, 2000, Ulrich et al., 2005, Chakraborty & Coates, 2005). In some studies where phenol was detected, benzoate or toluene were also detected, suggesting that there could be dual pathways for benzene degradation in these cultures (Caldwell & Suflita, 2000, Chakraborty & Coates, 2005, Ulrich et al., 2005).

Phenol production from benzene hydroxylation in abiotic and autoclaved controls has however been detected, throwing in to doubt its role in anaerobic benzene degradation (Kunapuli, Griebler, Beller, & Meckenstock, 2008, Abu Laban, Selesi, Jobelius, & Meckenstock, 2009). In the first study, previously reduced medium produced phenol when exposed to air for sample analysis, and autoclaved sulfate-reducing culture produced phenol in the second study. Medium and sample preparation differences may have led to abiotic phenol production being a greater issue in these two studies, as they found phenol in all controls, and at high concentrations. Previous studies had not detected phenol in controls, and had detected ten-fold lower concentrations of phenol where it was present (Ulrich et al., 2005), so abiotic phenol formation may not be important in all samples.

The pathway for anaerobic phenol degradation is well studied (Breinig, Schiltz, & Fuchs, 2000), review in (Carmona et al., 2009). Genes and intermediates involved are shown in Figure 1.2. The key initial step is a phosphorylation of phenol to phenylphosphate, carried out by phenylphosphate synthase (PSS). Also of note is the phenylphosphate carboxylase (PPC) enzyme, which carries out a subsequent dephosphorylation/ carboxylation. ppcA and ppcD subunits of the ppc operon also show similarity to the known decarboxylase/ carboxylase enzyme UbiD. This may hint at carboxylation enzymes of interest in other cultures (Abu Laban, Selesi, Rattei, Tischler, & Meckenstock, 2010). No gene/enzyme has been proposed for the initial step of benzene → phenol.

OH# COO%# COSCoA# COSCoA#

PpsABC' PpcABCD' PpcABCD' HbaA' HbaBCD' HcrL' HcrCAB' PcmRST' Phenol# O# O# OH# OH#

O# P# O%# 4%hydroxybenzoate# 4%hydroxybenzoyl%CoA# Benzoyl%CoA# OH#

Phenylphosphate#

Figure 1.2: Anaerobic phenol degradation pathway (adapted from (Abu Laban et al., 2010))

1.3.3 Carboxylation

Carboxylation involves the addition of a carbon dioxide molecule to form benzoate directly. Benzoate has been detected in all metabolite studies to date (Caldwell & Suflita, 2000, Phelps, Zhang, & Young, 2001, Ulrich et al., 2005, Kunapuli et al., 2008, Abu Laban et al., 2009). Abu Laban et al. found that two proteins were specifically expressed in the presence of benzene, but not benzoate or phenol. These proteins showed sequence homology to PpcA and PpcD, mentioned in the previous section. A putative anaerobic benzene carboxylase (Abc) is proposed,

consisting of several subunits, some of which show similarity to carboxylases Ppc/UbiD/UbiX (Abu Laban et al., 2010). This work was carried out in an iron-reducing benzene-degrading enrichment dominated by Peptococcaceae. The putative benzene degradation gene cluster abcD, abcA, bzlA, ubiX has also been found in Edwards Laboratory nitrate-reducing benzene-degrading enrichments, and these genes were more highly transcribed in the presence of benzene than benzoate (Luo et al., 2014). This pathway is shown in Figure 1.3. Further evidence for direct carboxylation in benzene is provided by the demonstration of direct carboxylation in naphthalene, another unsubstituted aromatic molecule (Mouttaki, Johannes, & Meckenstock, 2012).

Another recently proposed possibility in nitrate-reducing cultures is that molecular oxygen is produced from the intracellular disproportionation of nitrate to nitrogen and oxygen gas. The oxygen could be used as a reactant by microorganisms for the initial attack on the benzene ring. Monooxygenase genes have been found in perchlorate-reducing Dechloromonas aromatica strain RCB which degrades benzene with perchlorate as the electron acceptor (Salinero et al., 2009). Monooxygenases typically split an oxygen molecule and add a hydroxyl group to a substrate. Putative anaerobic benzene degradation genes were not found in D. aromatica.

COOH# COSCoA#

CO2#

AbcDA' BzlA'

Benzene# Benzoate# Benzoyl%CoA#

Figure 1.3: Anaerobic carboxylation of benzene to benzoate (adapted from (Abu Laban et al., 2010))

1.3.4 Central benzoate metabolism pathway

Subsequently, all three proposed degradation pathways produce benzoyl Co-A as a downstream central metabolite (Ulrich et al., 2005, Chakraborty & Coates, 2005, Caldwell & Suflita, 2000). Benzoyl-CoA is then degraded via a well-established anaerobic pathway summarized in Figure 1.4, adapted from (Gibson & Harwood, 2002). Different enzymes have been found to carry out the different steps in anaerobic organisms. Genes encoding known enzymes involved in the benzoate/benzoyl-CoA anaerobic degradation pathway fall into different families according to the anaerobe they were first identified in. Families include bad, bcl, bcr, bzd, bam and bzl (Carmona et al., 2009). Genes belonging to these families are anticipated in the genomes of organisms that consume intermediates of benzene degradation.

2 3 1

72 3 6 5 1

Figure 1.4: Anaerobic benzoyl-CoA degradation pathway. (1) Benzoyl-CoA (2) Cyclohex-1,5-diene-1-carbonyl-CoA (3) Cyclohex-1-ene-1-carbonyl-CoA (4) 2- hydroxycyclohexane-1-carbonyl7 -CoA (5) 2-ketocyclohexane6 -1-carbonyl-CoA5 (6) Pimelyl-CoA (7) 3-hydroxy-pimelyl-CoA. Slight variations are found in the pathway in different organisms. Adapted from (Gibson & Harwood, 2002)

1.3.5 Syntrophy

Syntrophy is a specific form of mutualism where two groups of organisms are required to completely utilize an electron donor due to thermodynamic constraints. Without the consumption of degradation products, the earlier product-forming steps would be less energetically favourable. In the case of benzene degradation, the metabolites of benzene degradation must be consumed in order for the concentration of the metabolites to be kept low enough in the system that the initial benzene-cleavage reaction is favourable. Syntrophic relationships are commonly found in methanogenic consortia, where one or more fermentative organisms break down organic precursors to acetate and hydrogen, which are then utilized by methanogens (Sieber, McInerney, & Gunsalus, 2012). Nitrate-reducing benzene degradation can also involve a syntrophic process, where one organism initiates benzene degradation, and hands off degradation products to nitrate-reducing bacteria (van der Zaan et al., 2012). Syntrophic interactions mean that benzene-degrading organisms are very difficult to isolate, as in the absence of partner organisms, intermediates in the degradation of benzene accumulate and prevent the initial benzene degradation step occurring. Terminal electron acceptor variability is possible in syntrophic processes because the intermediates produced by the initiator of benzene degradation can be consumed by a variety of other organisms, which may each utilize a different electron acceptor.

1.3.6 Organisms implicated in benzene degradation

A variety of phylogenetic groups have been implicated in benzene degradation in different studies. Two Dechloromonas and two Azoarcus strains have each been grown as isolates under nitrate-reducing benzene-degrading conditions (Coates et al., 2001, Kasai et al., 2006). Rhodocyclaceae, Chlorobi/Ignavi, Acidobacteria/Holophaga, Decholoromonas, Peptococcaceae, Aquaspirillum, Comamonas, Zoogloea, Hylemonella, Novosphignobium, Xanthobacter, Burkholderiaceae and Gemmatimonadetes have been found in nitrate-reducing benzene-degrading enrichment cultures (Ulrich & Edwards, 2003 and van der Zaan et al., 2012). Under sulfate-reducing conditions, Thiomicrospira, Epsilonproteobacteria, Cytophagales and

Desulfobacteraceae have been found (Phelps, Kerkhof, & Young, 1998). Geobacter and Variovorax are implicated under iron-reducing conditions (Rooney-Varga, Anderson, Fraga, & Lovley, 1999). Under methanogenic conditions, Desulfosporosinus, Desulfobacterium, Dehalospirillum, Chloroflexi, Methanosaeta, Methanomicrobiales, Methanosarcinales and Methanobacteriales are the dominant phylotypes (Ulrich & Edwards, 2003). The roles of some of these organisms have been deduced, while the roles of other organisms is uncertain. Peptococcaceae is the proposed initiator of benzene degradation in nitrate-reducing consortia (Abu Laban et al., 2010, Luo et al., 2014). Azoarcus and Burkholderiaceae are potentially responsible for degradation of downstream metabolites and nitrate reduction (Ulrich & Edwards, 2003, Luo et al., 2014).

1.3.7 Metagenome assembly

Recent developments in sequencing capabilities have increased the availability of high-quality sequence data dramatically (Solomon, Haitjema, Thompson, & O’Malley, 2014). It is relatively easy to generate vast amounts of sequencing data, and cover metagenomes to a read depth sufficient to assemble near-complete genomes of relatively un-enriched organisms from metagenome sequence data (Tang, Gong, & Edwards, 2012). Assembly of such genomes, however, is not trivial. Full genomes of organisms within the same phylogenetic groups as some of the organisms consistently found in nitrate-reducing benzene-degrading cultures have been assembled. These genomes include two Peptococcaceae (Pelotomaculum thermopropionicum and Thermincola potens (Kosaka et al., 2008, Byrne-Bailey et al., 2010)), a member of the Azoarcus/Thauera clade (Aromatoleum aromaticum EbN1) (Rabus et al., 2005), a Burkholderiales (Burkholderia multivorans) (Varga et al., 2012) and an Ignavibacteriales (Melioribacter roseus) (Kadnikov et al., 2013). These genomes were not produced from organisms growing on benzene or intermediates of its degradation. Assembly of genomes of organisms from nitrate-reducing benzene-degrading cultures is likely to produce novel insights into variations between members of these clades that are grown on other sources compared to those grown on benzene, and thereby some of the adaptations required for growth on benzene.

Chapter 2 Background

2.1 Rationale for research

The focus of this research was on the benzene-degrading nitrate-reducing cultures first developed in 1995 and maintained under anaerobic conditions with nitrate and benzene since then. The background of these cultures is presented in Section 2.5. Many different bacterial and archaeal species are present in the cultures studied (Ulrich & Edwards, 2003). These cultures were found to harbour a syntrophic community of a Peptococcaceae and nitrate reducing partners using products from the fermentation of benzene (Gitiafroz, 2012). The diversity both within and among cultures is unexplored. Potential roles of different organisms in the cultures are also not fully understood. Therefore the overall objective of this work was to determine the functional capabilities of the different bacteria in specific benzene-degrading enrichment cultures. Understanding the functions of the different community members may enable us to enhance benzene degradation rates, and knowledge of this may be used to improve growth.

2.2 Hypotheses

• A core set of OTUs are present among different anaerobic nitrate-reducing benzene- degrading cultures.

• Peptococcaceae is responsible for the first step of benzene degradation and has genes coding for initial steps in benzene degradation.

• Organisms are capable of exploiting electron donors and acceptors that are present in culture besides benzene/nitrate.

2.3 Objectives

1. Compare and contrast the nitrate-reducing benzene-degrading community in different microcosms and track changes in one microcosm over time to try to identify factors that

result in increased benzene degradation rates.

2. Analyze metagenome sequence data to acquire further evidence for the hypothesized benzene degradation pathway in an anaerobic nitrate-reducing benzene-degrading

culture.

3. Understand alternative electron donor/acceptor reactions that could be carried out by the microbial community present in the nitrate-reducing cultures. In particular, explore if the syntrophic partner could be a methanogenic community rather than a nitrate-reducing

organism, thereby removing the need to continually supply nitrate.

2.4 Approach

Pyrotag 16S rRNA sequencing of a selection of cultures was used to determine the community composition (Chapter 4). High-throughput shotgun sequencing was used to generate sequence data for a nitrate-reducing benzene-degrading enrichment. Sequence data was then assembled and analysed (Chapter 5). Enrichment bottles were established to test whether nitrate-reducing benzene-degrading culture could degrade benzene with methanogenesis (Chapter 6). Appendices 2 – 4 detail additional projects carried out that were related to the main objectives.

2.5 Enrichment cultures used in this study

Enrichment cultures of anaerobic benzene-degrading bacteria used in this study have been maintained in the Edwards Laboratory for 19 years (Nales, Butler, & Edwards, 1998, Burland & Edwards, 1999, Ulrich & Edwards, 2003). Cultures originated from soil material collected at a decommissioned gas station (Cartwright Avenue, Toronto, Ontario), and a "pristine" site with no

history of benzene contamination (a swamp in Perth, Ontario). Benzene and nitrate are re-fed to the enrichments when depleted. Iron sulfide is added to the medium when new cultures are inoculated, or fresh medium added to existing enrichments. Ferrous iron could drive the abiotic reduction of nitrate to nitrite or nitrogen gas. Biotic or abiotic reactions could also lead to the production of ammonium, and iron and sulfur cycle reactions could also be taking place in the cultures. While these reactions may be taking place, the stoichiometry of benzene oxidation and nitrate consumption has consistently shown that benzene oxidation is most likely coupled to nitrate reduction to nitrite, with nitrite then being denitrified to nitrogen gas (Burland and Edwards, 1999, Ulrich and Edwards, 2003, Gitiafroz, 2012). No ammonium production has been detected, and ammonium consumption linked to the Anammox process (anaerobic ammonium + - oxidation, NH4 + NO2 → N2 + 2H2O) has been found in these enrichments in the past (Nandi, 2006). The only substrates continually added to the cultures are benzene and nitrate, and these are the main electron donor and acceptor.

Phylogenies of cultures made from material from each site are shown in Figures 2.1 (cultures inoculated with material from the Cartwright Ave gasoline station site) and 2.2 (cultures inoculated with material from the pristine “swamp” site). Transfer cultures prepared in 2003 or earlier were made with 50% inoculum into 50% of anaerobic defined mineral medium. Transfer cultures established in 2006 contained a maximum of 1% inoculum and were transferred into medium lacking FeS and resazurin. While these cultures have been maintained in the laboratory for a long time, they still grow slowly compared to growth on toluene or benzoate and show a long lag period before resuming activity after transfers. They often degrade benzene faster when more sediment is present in enrichments.

cultures

s (2012)% *CartT3f%(2012)% *CartT3g%(2012)% Cart%Cons2006%revival% Cart%Consolidated%FL1% Cart%Cons%FL2%*CartT3h% indicate

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*SwampT2a%(2008)% *SwampT3e%(2012)% Swamp%cons%2007%revival%

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Figure 2.2: Approximate lineage of Swamp cultures. lineage enrichment Approximate 2.2: Figure established are were in which The years cultures this thesis. names used throughout Labels with an asterisk are background green A and archived. that have been frozen cultures font indicate in brackets. Labels red listed “consolidated” to for sequencing. “Cons” is used as an abbreviation pyrotag submitted for cultures indicates bottles enrichment different biomass from by formed pooling together cultures indicate

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Chapter 3 Methods 3 3.1 Culture maintenance

Cultures were maintained in the dark at room temperature (approximately 20°C) in glass bottles ranging from 2L to 100mL in volume, sealed with either Mininert caps (VICI precision sampling, Baton Rouge, LA, USA) or black butyl rubber stoppers. Cultures were manipulated and stored in an anaerobic glovebox (Coy Lab Products, MI) with an atmosphere of 80% N2,

10% CO2, 10% H2. Cultures were supported in anaerobic mineral medium, reduced with amorphous iron sulfide (Burland & Edwards, 1999). Cultures were maintained with benzene as the sole electron donor, and nitrate as the sole electron acceptor. They were re-amended with neat benzene to 10-30mg/L benzene when less than 5mg/L benzene remained in culture.

Cultures were amended with nitrate to 2mM from a 500mM NaNO3 stock when nitrate dropped to below 0.5mM.

3.2 Culture set-up

Cultures were set up to test whether the benzene-degrading organisms in the nitrate-reducing cultures could also donate electrons to methanogens instead of nitrate-reducing organisms (Chapter 6). The nitrate-reducing benzene-degrading culture Cart T3 was tested for this capability in one treatment. In another treatment, an additional source of methanogens was provided by mixing toluene-degrading methanogenic culture (T3L) with Cart T3. Methanogens in T3L were adapted to the intermediates of toluene degradation, which are likely to be similar to the intermediates of benzene degradation. New cultures were set up by adding 40mL active culture to 100mL serum bottles with butyl rubber stoppers. The experimental set-up is shown in Table 3.1. All bottles were allowed to deplete the remaining nitrate in the medium and then were re-amended with 10mg/L benzene so that no electron acceptor was present apart from carbon

dioxide after 30 days. Methane and benzene concentrations were measured periodically using gas chromatography.

Table 3.1: Experimental set-up for testing the potential of CO2 as a terminal electron acceptor in nitrate-reducing benzene-degrading enrichments. Inoculum source Name Replicate Replicate Replicate 40mL nitrate-reducing benzene- Benzene culture A B C degrading Cart T3 culture Only 20mL nitrate-reducing benzene- Benzene culture A B C degrading Cart T3 culture, 20mL plus Toluene toluene degrading culture T3L culture 40mL toluene degrading culture T3L Toluene culture A B C Only 20mL nitrate-reducing benzene- Autoclaved A B C degrading Cart T3 culture, 20mL control toluene degrading culture T3L (autoclaved)

3.3 Gas chromatography

Benzene, toluene and methane concentration in all cultures was monitored by gas chromatography (GC). Headspace samples (400µL) were taken from culture bottles through butyl rubber stoppers or mininert caps using Pressure-Lok gas-tight syringes (Precision Sampling, Baton Rouge, LA). Syringe plungers were depressed to 300uL and the needle valve quickly opened and shut immediately to equalize pressure between different samples. The samples were introduced via an injection port to a Hewlett Packard 5890 Series II Gas Chromatograph equipped with a GSQ PLOT I.D. 30m x 0.53mm column (J&W Scientific, Folsom, CA) and a flame ionization detector (FID). The time of detection and peak areas were recorded, and compared to standards to determine the concentrations of compounds of interest. All volatile organic compounds measured in this study were analysed using the protocol as follows: Injector temperature: 200°C; Oven temperature (isothermal): 190°C; FID temperature: 250°C. Helium was used as a mobile phase, at a flow rate of 11mL/min. Under these conditions, the retention time for methane was 0.3 minutes, benzene was 1.3 minutes, and toluene was 2.6

minutes. Calibration of benzene and toluene was carried out using headspace samples from standards prepared gravimetrically from concentrated methanolic stocks, and stored in Mininert- sealed glass bottles. Methane standards were prepared from varying volumes of 99% methane stock independently.

3.4 Ion chromatography

Nitrate and nitrite concentrations were measured by ion chromatography (IC). Liquid samples were taken from cultures and filtered through 0.2 micron filters (EMD Millipore, MA) to remove particles. Samples were then diluted 1:20 by volume in deionized MilliQ water. Samples were analysed on a Dionex Ion Chromatograph ICS-2100 equipped with an AS14 column and electrolytic hydroxide eluent generation, with eluent concentration 23mM and suppressor 57mA. Retention time for nitrite was 5.3 minutes under these conditions, and 8.5 minutes for nitrate. Results were analysed using the integrated Chromeleon software package.

3.5 DNA extraction and sequencing for metagenomic analysis

Nitrate-reducing benzene-degrading Cart T3 culture (200mL) was taken anaerobically in falcon tubes and sealed with anaerobic tape in the glove box before being centrifuged at 8,000 rpm for 30 minutes. Supernatant was removed in the glovebox, and DNA extraction carried out immediately to prevent excess oxygen exposure before cell debris removal. The PowerSoil DNA extraction kit (MoBio Laboratories, Inc) was used to extract high quality, high molecular weight DNA from the cell pellet following the manufacturer’s protocol. Library preparation and sequencing were carried out by Genome Quebec. Both mate pair and paired end libraries were prepared, and DNA was sequenced by an Illumina Genome Analyzer II system. Low quality reads were trimmed by Trimmomatic (Bolger, Lohse, & Usadel, 2014). Assembly was carried out using ABySS (Simpson et al., 2009) and Allpaths LG (Gnerre et al., 2011).

3.6 Pyrotag sequencing

Total DNA was extracted from samples and a variable section of the 16S rRNA gene was amplified by PCR and labeled with a barcode to identify which sample the DNA was amplified from. Different samples were pooled, and sequenced by 454 pyrosequencing. Ten mL of each culture was taken anaerobically, and cells pelleted by centrifugation (8,000 rpm for 30 minutes) before DNA was extracted using the Powersoil DNA extraction kit (MoBio Laboratories, Inc) according to the manufacturer's protocol. DNA was amplified using forward primer 926f (5’- AAACTYAAAKGAATTGACGG-3’) and reverse primer 1392r (5’-ACGGGCGGTGTGTRC- 3’) (Moran, Hansen, Powell and Sabree, 2012). Different 10 base-pair barcodes incorporated before the forward primer were used for each sample. 25uL PCR reactions were carried out, containing 12.5uL 2x PCR master mix (Fermentas/ Thermo Fisher Scientific, MA), 0.5uL forward primer, 0.5uL reverse primer, 10.5uL water (nuclease free) and 1uL DNA extract. Amplification was carried out with an initial 3 minute 95°C denaturation step, followed by 25 cycles of 95°C for 30 seconds, 55°C for 45 seconds, and 72°C for 90 seconds, followed by a final extension phase of 72°C for 10 minutes on a PTC-200 DNA Engine thermocycler (MJ Research Inc., Waltham, MA). PCR product was purified by QIAquick PCR Purfication kit (Qiagen, Valencia, CA) and quantified by nanodrop and agarose gel electrophoresis. PCR products were then sent to Genome Quebec and sequenced by a GS FLX Titanium Series pyrosequencer. Results were analysed by the program QIIME (Caporaso et al., 2010). Alignments were carried out in Geneious, using MUSCLE (Edgar, 2004). Phylogenetic trees were created using a maximum-likelihood algorithim in PHYML with 1,000 bootstraps (Guindon et al., 2010).

Chapter 4 Pyrotag sequencing to determine community composition

4.1 Introduction

While organisms have been identified that are responsible for the initial step of benzene degradation and consumption of downstream metabolites, many other organisms are also present in benzene degrading consortia. These other organisms may be decomposing cell debris, gaining energy through photosynthesis or scavenging any oxygen present in the bottles. The impact of these groups on benzene degradation overall are unknown.

Pyrotag DNA sequencing can be used to obtain a snapshot of the community composition in a culture at a given point in time. The relative abundances of different OTUs within different cultures can be easily obtained. Pyrotag data can be compared between different consortia to see which groups are more or less abundant under different culture conditions. Comparisons to nitrate-reducing benzene-degrading groundwater samples and consortia grown from different inoculation sources from other laboratories provide an interesting point of comparison (Coates et al., 2001, Kasai et al., 2006, and van der Zaan et al., 2012). Both nitrate-reducing consortia and those utilizing alternative electron acceptors can be compared to pyrotag data (Phelps et al., 1998, Rooney-Varga et al., 1999, and Ulrich & Edwards, 2003). Previous clones libraries and qPCR studies on the consortia used also provide a point of reference (Ulrich & Edwards, 2003) .

Five established nitrate-reducing benzene-degrading consortia were submitted for pyrotag sequencing in 2012: Swamp T3, Swamp T2a, Cart T2d, Cart T3c, and Cart T3. Two further cultures were sequenced in 2013 (Cart T3g, Cart T3h) and Cart T3 was re-sequenced. In 2014, Cart T3 was re-sequenced again, and Cart T3f was also sequenced. These consortia were chosen to represent a variety of culture origins and histories (see Figure 4.1). Cart T3 was re-sequenced each year because there is evidence from culture maintenance data to suggest that over the years

of maintenance, the degradation abilities of the cultures change, and community composition changes may be a factor in the changes in degradation performance.

Total DNA was extracted from samples and a variable section of the 16S rRNA gene was amplified by PCR and labeled with a barcode to identify which sample the DNA was amplified from. Different samples were pooled, and sequenced by 454 pyrosequencing (Sogin et al., 2006). The location of raw data is reported in Appendix 5. Results were analysed using QIIME (Caporaso et al., 2010). A 99% identity threshold was used to group pyrotag reads into OTUs for downstream analysis.

Site%1% Site%2% Cartwright%Gasoline%sta2on% Uncontaminated%Swamp% (Toronto,%ON)% (Perth,%ON)%

Various%transfers% Various%transfers%

2006% 2002% 1998% 2002%

Cartwright%PW1%10A1%% Cartwright%Consolidated%% 0.2mg/L/day% 0.3A0.2mg/L/day% Swamp%NO3%1aB% Swamp%Consolidated% *Cart%T2d% *Cart%T3% 0.4mg/L/day% 0.3mg/L/day% *Swamp%T2a% *Swamp%T3%

2006%

Cartwright%Consolidated% 2012% 10A2% 0.2mg/L/day%% *Cart%T3c% Cartwright%Consolidated% Cartwright% Cartwright% 2006%revival%% Consolidated%FL1%% Consolidated%FL2%% 0.7mg/L/day% *Cart%T3g% *Cart%T3h% *CartT3f%

Figure 4.1: Approximate lineage and benzene degradation rates of cultures submitted for 1" pyrotag sequencing

4.2 Results and discussion

4.2.1 Alpha diversity of samples

The within-sample diversity (Alpha diversity) for these samples can be seen by charting the number of unique OTUs added per read sequenced for each sample (Figure 4.2). These rarefaction curves indicate how likely it is that the sequencing data covers all OTUs present. Where a curve flattens out, it indicates that further sequencing is unlikely to reveal more OTUs, as in Cart T3f and 2014Cart T3. Where a curve is still trending upwards, as for Swamp T3 and 2012Cart T3 here, it shows that not all of the OTUs present in the culture are likely to be represented in the sequence data. Despite not capturing all OTUs in this data, the most abundant OTUs are represented in the data. In general, cultures that have gone through fewer dilution and transfer events are more diverse. As pyrotag data reveals relative abundances of OTUs but not

800 2012SwampT3

700

2012CartT3 600

2012SwampT2a 500

2013CartT3 400 2013CartT3h 2012CartT3c 2013CartT3g 2013CartT2d 300 Number of observed OTUs 200 2014CartT3f 2014CartT3 100

0 0 1000 2000 3000 4000 5000 6000 7000 8000 Number of reads per sample Figure 4.2: Alpha diversity of pyrotag samples.

absolute numbers, an increase in the abundance of a particular OTU at the point in time that a sample was taken could skew these numbers.

4.2.2 Relative abundances of various OTUs

Figure 4.3 shows the percent abundances of different OTUs in the cultures studied, and the phylogeny assigned to the OTUs by QIIME. A large number of different OTUs are present, at varying abundances, and the cultures differ in their composition considerably. Selected groups are coloured, and these are the focus of the following discussion.

Cart#T3#(2014)# Cart Consolidated Cart#T3#(2013)# culture sequenced in 3 sequential Cart#T3#(2012)# years

Cart#T3h# Sub cultures of Cart T3 Cart#T3g#

Cart#T3f# Frozen Cart T3

Cart#T3c# Deep dilutions of Cart#T2d# Cart cultures

Swamp#T2a# Swamp cultures Swamp#T3#

0%# 10%# 20%# 30%# 40%# 50%# 60%# 70%# 80%# 90%# 100%#

Ignavibacteriales Peptococcaceae Burkholderiaceae Thiobacillus

Azoarcus Other Sum of OTUs individually less Rhodocyclaceae than 1% in all cultures

Figure 4.3: Pyrotag sequencing results showing percent abundances of different OTUs in a selection of cultures.

The first highlighted group, Ignavibacteriales (orange), is a facultative anaerobic heterotroph (Podosokorskaya et al., 2013). It has no proposed role in benzene degradation, but it, or the closely related Chlorobi clade is found in multiple nitrate-reducing benzene degrading enrichments. It is present in higher abundances in Cart T2d and Cart T3c, the only cultures sequenced which have undergone thousand-fold or higher dilutions. This may indicate it is opportunistic and grows when other OTUs are at lower abundance, or could be gaining energy and carbon from dead biomass.

The next highlighted group is the Peptococcaceae, in purple. This group contains the proposed initial benzene degrader. However, they are at fairly low abundances in the cultures. The abundance of Peptococcaceae was between 20% and 2%.There are some possible reasons for their low abundances. They may have a larger cell size than other organisms present, and so represent a higher proportion of the culture biomass than their abundances would suggest. There may also be multiple copies of the 16S gene in other organisms, increasing their relative abundances. Another reason for their low abundance could be that they are only harvesting a small proportion of the energy available in benzene. If they are only carrying out the initial steps of benzene degradation, a lot of the energy present in the electron donor will be handed off to other consortium partners. This could lead to their low growth.

The Burkholderiaceae (highlighted yellow) are a denitrifying organism (Gumaelius, Magnusson, Pettersson, & Dalhammar, 2001). They could be scavenging small organic molecules produced at some point along the degradation pathway of benzene. Their role is similar to the Azoarcus (light blue), a member of the Rhodocyclaceae family, and other Rhodocyclaceae (dark blue). Azoarcus however have been shown to degrade aromatic compounds, and an isolate of Azoarcus degrades benzene with nitrate reduction (Kasai et al., 2006). The abundance of Azoarcus was between 50% and 2% in the consortia studied.

Thiobacillus, shown in green, is a denitrifying and sulfide oxidizing bacteria also capable of carbon dioxide fixation (Beller et al., 2006). It may be carrying out these functions in the enrichments. It is at very high abundance in Cart T3f, a revival of a previously frozen culture.

Freezing and thawing may have promoted conditions where this OTU was able to increase in abundance.

The red bars show the sum total of OTUs that were present at less than 1% abundance in all cultures sequenced. These bars show that up to 40% of the cultures were composed of highly diverse very low abundance OTUs. This pool of diversity may provide more redundancy for the cultures, so that if conditions change, a different organism capable of carrying out the same functions may gain an advantage over the organisms currently occupying those roles, and increase in abundance while community function overall is unchanged.

Cart T3 culture shows great variation in relative abundances of multiple OTUs over the three years it was tracked. Changes may be temporary, related to a factor affecting the community at the time of sampling, or indicative of long-term trends. Transitory factors that may have affected the results include at which point in the degradation cycle DNA was taken. This varied from year to year. Another time-dependent factor that could affect the community is the length of time since fresh medium was added to the culture. More fresh medium was added to Cart T3 in the two months prior to sampling in 2013 than in 2012 or 2014. Measuring changes in the community over a shorter time scale, such as a single degradation cycle, would provide some clarification of this.

OTUs common to multiple samples were identified in pyrotag data for five samples, which were used as representatives of groups of similar samples. Presence in all cultures could indicate that OTUs represent the key groups in the community and should be considered more closely. Out of 51 OTUs present at greater than one percent abundance in any of the cultures used, 18 are common to all of them (Figure 4.4). Cultures that had previously been diluted had lower numbers of unique OTUs, likely due to the loss of OTUs during transfers. Very few OTUs are present in only one culture. The OTUs found only in one culture may be present at very low abundances and not affect culture performance, but may have a role in nutrient cycling, or provide alternative options for carrying out functions to increase the resilience of the cultures under changing environmental conditions. The common OTUs are presented in Table 4.1. The

potential roles of most of these organisms has been discussed previously. Previously discussed groups are shaded grey.

SwampT3

CartT3(2012) CartT2d

CartT3h CartT3f

Figure 4.4: Venn diagram showing the number of OTUs at greater than one percent abundance that are shared between, or unique to, different cultures sampled. A core set of 18 OTUs is shared between all cultures sampled.

( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $

4.46 0.13 0.03 0.27 0.51 0.04 0.01 2.88 0.02 0.47 0.31 2.10 0.13 8.97 1.30 0.04 0.96 57.40 (2014) Cart$T3$ ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( (

$ 2.93 0.11 0.90 0.45 0.53 0.23 0.24 3.41 0.15 0.87 0.19 1.40 0.11 0.73 5.23 0.88 12.78 20.69 (2013) Cart$T3$

( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 4.71 0.18 0.10 2.64 1.32 0.64 1.08 5.41 0.42 5.14 0.19 5.59 0.56 3.74 1.93 0.56 1.16 1.97 (2012) Cart$T3$ ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 0.88 0.24 0.23 0.34 2.94 1.25 1.08 0.04 0.58 0.07 0.70 0.60 9.45 1.04 6.40 0.63

15.40 32.74 Cart$T3h ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 0.12 0.11 1.45 0.12 0.27 0.24 0.24 2.40 0.00 0.53 0.09 1.30 2.51 1.47 2.81 0.39 14.87 43.87 Cart$T3g

( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 2.11 0.22 0.01 3.73 0.10 0.09 0.01 2.23 0.19 0.86 0.92 3.39 0.22 1.06 0.01 3.96 2.69 56.03

Cart$T3f ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 0.13 0.00 0.52 0.00 0.07 0.31 1.56 0.52 0.07 0.00 4.36 0.18 0.00 0.10 0.10 6.88 1.09

49.80 Cart$T3c ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 1.62 0.22 0.13 0.60 0.37 0.48 2.76 0.35 0.04 0.06 2.35 0.06 0.09 1.25 0.53 0.51 54.01 11.07 degrading enrichments tested, with percent abundances in all - Cart$T2d

( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 0.18 0.52 1.23 1.34 1.27 0.59 1.25 6.57 4.10 0.06 1.18 0.07 0.91 6.09 2.02 0.35 2.80 16.42 T2a

Swamp$ ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( $ 0.67 2.00 5.68 1.43 0.92 0.64 0.77 3.53 0.10 0.67 0.03 0.03 1.58 4.35 0.52 0.14 2.39 10.60 T3 Swamp$ reducing benzene - ( $ ( ( ( ( ( ( ( ( ( ( ( ( 31 4 ( ( ( 36 4 ( ( 8:SJA 4 Phylogeny$assigned$by$QIIME Acidobacteria:iii1 Bacteroidales:Porphyromonadaceae Ignavibacteriales:Ignavibacteriaceae Anaerolineae:SBR1031:SHA Clostridiales:Peptococcaceae Clostridiales:Peptococcaceae Clostridiales:Peptococcaceae Clostridiales:Peptococcaceae Gemmatimonadetes Rhizobiales:Hyphomicrobiaceae:Rhodoplanes Rhodospirillales:Rhodospirillaceae Burkholderiales:Burkholderiaceae Burkholderiales:Comamonadaceae Hydrogenophilaceae:Thiobacillus Rhodocyclaceae:Azoarcus Rhodocyclaceae:Azoarcus Rhodocyclaceae:Dok59 Deinococcaceae:Deinococcus ! ( ( ( ( ( ( ( ( ( $ . Shaded discussed .Shaded section groups inearlier 4.2. are those ( ( ( ( ( ( ( ( ( 4.1: 4.1: 18 OTUs common to all nitrate

OTU$

Number OTU#1757 OTU#895 OTU#1516 OTU#1530 OTU#293 OTU#1409 OTU#788 OTU#1322 OTU#1164 OTU#1282 OTU#263 OTU#853 OTU#145 OTU#736 OTU#827 OTU#1067 OTU#287 OTU#1254 Table samples

4.2.3 Relatedness of samples

Figure 4.5 shows the relatedness of different samples. 2014 Cart T3, which originated from Cartwright cultures biomass grown during various experiments and later pooled together and 2014 Cart T3f, which was set up in 2012, using frozen material from Cart T3 in 2006, cluster together. These cultures have the lowest diversity. They likely cluster together due to this reduced diversity, not due to similarities in abundances of OTUs present, as the OTUs present in the two cultures are different according to Figure 4.3. Cultures that originated from Swamp material cluster within the Cartwright-origin samples, showing that culture origin has an effect even after 18 years enrichment. The 2013 Cart T3g and 2013 Cart T3h, which were set up with material from Cart T3 in 2012, cluster together and with Cart T3 2013, but show some

2014 Cart T3

2014 Cart T3f

2012 Swamp T3

2012 Swamp T2a

2012 Cart T3

2013 Cart T3

2013 Cart T3g

2013 Cart T3h

2012 Cart T2d

2012 Cart T3c

Figure 4.5: Bootstrap tree using UPGMA (Unweighted Pair Group Method with A rithmetic mean) values to show clustering of samples. Red nodes = 75 – 100% support, yellow nodes = 50 – 75% support.

differences from each other and their parent culture even after less than a year of cultivation. Cart T3c and Cart T2d, both cultures grown from thousand-fold or greater dilutions, cluster together. Samples cluster by the year they were taken and sequenced. This could be due to DNA extraction effects, methodology changes, or reflect the point in the degradation cycle that samples were taken, which varied from year to year. Samples could be taken over the course of a single degradation cycle to see if this has an effect.

4.2.4 Phylogenetic grouping of OTUs

The classifications assigned by QIIME were checked through BLAST searching reference sequences for a selection of OTUs against the NCBI nr/nt nucleotide database and comparing the sequence hits to the phylogenies assigned. QIIME classifications matched the identities of the nearest BLAST hits. The phylogenetic trees shown in this section further confirm the accuracy of QIIME’s classification algorithm. OTUs fall within the same clade on the tree as QIIME would assign them.

Figures 4.6 and 4.7 show phylogenetic trees based on pyrotag sequencing results. All OTUs present in any culture at greater than one percent abundance are represented, along with relevant reference sequences. Betaproteobacteria and Gammaproteobacteria OTUs are shown in Figure 4.6. OTUs outside the Betaproteobacteria and Gammaproteobacteria groups are shown in Figure 4.7.

These trees show that groups present in the pyrotag sequencing data are highly similar to sequences found in the cultures previously. Closest BLAST hits to pyrotag sequences were often from other benzene-degrading enrichments or aromatics-contaminated environments. Branch lengths are short in Figure 4.6 due to the short genetic distance between members of the Beta- and Gammaproteobacteria clades. Pyrotag sequences of 200 – 300 base pairs were used to produce the trees, which contributes to the low bootstrap values.

While many of the OTUs found in the pyrotag sequencing data are similar to known sequences, others group into less well-known clades. The Chlorobea/Ignavibacterium clade is

not well understood, and Ignavibacterium class was only recognized as a separate division to the Chlorobea class in 2010 (Iino et al., 2010). Pyrotag OTUs 1537, 911, 1530 and 1698 are not closely related to any isolated species. Closest BLAST hits for these OTUs to the 16S ribosomal RNA sequence database are a mixture of Firmicutes and Chloroflexi.

Rhodocyclaceae

Azoarcus

Burkholderiales

Figure 4.6: Phylogenetic tree of Beta- and Gammaproteobacteria showing pyrotag sequences in blue, earlier sequences from these cultures in red, and root sequence in purple. OTUs shown are greater than one percent abundance in sequenced cultures. Boxes indicate OTUs belonging to groups of interest.

Chlorobi/Ignavibacterium

Peptococcaceae/Clostridiales

Figure 4.7: Phylogenetic tree of all groups outside Beta- and Gammaproteobacteria showing pyrotag sequences in blue, earlier sequences from these cultures in red, and root sequence in purple. OTUs shown are greater than one percent abundance in sequenced cultures. Boxes indicate OTUs belonging to groups of interest. 33

4.2.5 Comparison to other benzene-degrading enrichment cultures

Table 4.2 compares groups present in benzene degrading consortia with different electron donors and the presence or absence of those groups in pyrotag data from the cultures studied in this project.

Table 4.2: Groups present during benzene degradation with different electron acceptors, compared to the presence (indicated by X) or absence (indicated by -) at greater than one percent abundance in pyrotag sequencing data Group associated with Present at >1% abundance in benzene degradation pyrotag data Reference Electron Acceptor: Nitrate Dechloromonas X (Coates et al., 2001) Burkholderia X Azoarcus X Rhodocyclaceae X Chlorobi/Ignavi X Acidobacteria/Holophaga X Decholoromonas X (Ulrich & Edwards, 2003) Peptococcaceae X Azoarcus X Decholoromonas X Aquaspirillum - Comamonas X Zoogloea - Hylemonella - Novosphignobium - Xanthobacter - (Kasai et al., 2006) Peptococcaceae X Rhodocyclaceae X Burkholderiaceae X Gemmatimonadetes X Chlorobia X (van der Zaan et al., 2012) Electron Acceptor: Sulfate Thiomicrospira - Epsilonproteobacteria - Cytophagales - (Phelps et al., 1998) Desulfobacteraceae -

Electron Acceptor: Iron (III) Geobacter - (Rooney-Varga et al., 1999) Variovorax - Methanogenic Desulfosporosinus - Desulfobacterium - Dehalospirillum - Chloroflexi X Methanosaeta - Methanomicrobiales - Methanosarcinales - (Ulrich & Edwards, 2003) Methanobacteriales -

From this table, it is apparent that the phylogenetic groups present in nitrate-reducing benzene-degrading cultures share considerable similarities, while different phylogenetic groups are present in cultures with alternate electron acceptors. In nitrate-reducing cultures, Dechloromonas, Peptococcaceae and Rhodocyclaceae are common groups. Methanogens were also present in all the cultures sequenced at very low abundances (shown in Table 4.3).

Table 4.3: Percent abundances of methanogenic OTUs in consortia according to pyrotag data

2012 2012 2012 2012 2014 2013 2013 #OTU 2012 2013 2014 Swamp Swamp Cart Cart Cart Cart Cart Order Cart T3 Cart T3 Cart T3 ID T3 T2a T2d T3c T3f T3g T3h 613 Methanobacteriales 0.027 0.014 0.059 0.024 0.048 0.068

1639 Methanomicrobiales 0.054 0.014 0.065 0.012 0.210 0.126

1095 Methanomicrobiales 0.027 0.029 0.106 0.032 0.014 0.037

795 Methanosarcinales 0.082

4.2.6 Comparison of OTUs to culture parameters

The variables “benzene rate”, “sediment presence”, “generation” and “sequencing run” were tested against an NMDS (non-metric multidimensional scaling) model of pyrotag sequence abundances in R (R Core Team, 2014). Values used in this analysis are shown in Table 4.4.

Benzene degradation rate is in mg/L/day. “Solids” is based on a visual estimate of how much sediment is present in the enrichment bottles, “0” = little/none, “1” = some, “2” = maximum level of solids observed. “Regularity” is based on culture history, “0” = culture has stalled multiple times in the past, “1” = the culture has stalled at some point but not frequently, and “2” indicates a culture that has not stalled. The values for “Generation” indicate: “1” = no dilution in the past, “2” = culture is a transfer of a previous culture, “3” = culture was generated by a deep dilution of an older culture.

Table 4.4: Metadata used in analysis of Pyrotag sequencing data

Benzene Pyrotag Degradation Sequencing Culture Rate Solids Regularity Generation Run 2012Swamp_T3 0.3 2 2 1 2012 2012Swamp_T2a 0.4 1 2 2 2012 2012Cart_T2d 0.2 0 0 3 2012 2012Cart_T3c 0.2 0 0 3 2012 2014Cart_T3f 0.7 0 2 2 2014 2012Cart_T3 0.25 1 1 2 2012 2013Cart_T3 0.25 1 1 2 2013 2014Cart_T3 0.25 1 1 2 2014

This analysis indicated which variables might correlate with OTU abundance. The variables “sediment presence”, “generation” and “sequencing run” were non-significant. “Benzene rate” had a p-value of 0.002. Figure 4.8 shows the NMDS with the correlation with benzene rate indicated. OTU 561, which correlates with benzene degradation rate, is a Chlorobi, present at less than 0.5% in two cultures only. OTU 1150, which also correlates with benzene degradation rate is an Anaerolineae present at 0.6% in one culture (Cart T3c). Other groups that cluster with these are also very low abundance OTUs, found only in some of the cultures. Their potential impact on benzene degradation rate is difficult to estimate.

0.8

Benzene.rate 0.6

OTU.561 OTU.197OTU.1597OTU.1764OTU.55 OTU.1150 OTU.912 OTU.810OTU.1807 OTU.1516OTU.676 OTU.235OTU.816 OTU.126OTU.861OTU.712OTU.995OTU.1347 OTU.1698 0.4 OTU.286 OTU.1451OTU.99OTU.1247OTU.153 OTU.689OTU.1102OTU.172OTU.957OTU.814OTU.402 OTU.1107OTU.773OTU.327OTU.1822OTU.855OTU.1175 OTU.1134OTU.959OTU.525OTU.958OTU.459 OTU.1359OTU.1603OTU.1589 OTU.364OTU.266OTU.507 OTU.376OTU.40OTU.1532OTU.1795 OTU.1743 OTU.1731OTU.1219 OTU.637 OTU.193OTU.57 OTU.801OTU.1434 OTU.1642 OTU.456OTU.1550 OTU.1527 OTU.1063 OTU.1157OTU.626OTU.1309OTU.1650 OTU.871 OTU.1234OTU.373OTU.537 OTU.78 0.2 OTU.1034 OTU.1340OTU.650OTU.1397 OTU.718 OTU.360 OTU.288 OTU.115OTU.862OTU.1040OTU.15OTU.348 OTU.323 OTU.1183OTU.1442OTU.753OTU.838 OTU.1295OTU.258OTU.1742OTU.439 OTU.910 OTU.1140 OTU.1078 OTU.1407 OTU.1765OTU.540OTU.476OTU.265OTU.1638OTU.573OTU.1626OTU.1044OTU.1423OTU.1215 OTU.620 OTU.1508 OTU.965OTU.1544OTU.1238OTU.1754 OTU.691OTU.1551OTU.284 OTU.1687OTU.1076 OTU.762OTU.1465OTU.927OTU.474OTU.516 OTU.1332OTU.59OTU.1598 OTU.1552OTU.652 OTU.215 NMDS2 OTU.1540OTU.251OTU.728OTU.1439OTU.890OTU.203OTU.1567OTU.1399OTU.1419 OTU.864OTU.1647OTU.1752 OTU.1031OTU.692OTU.1808OTU.1046OTU.1120OTU.780OTU.37OTU.1093OTU.1641OTU.1553OTU.440OTU.1164OTU.1101OTU.536OTU.831 OTU.80 OTU.1591 OTU.477OTU.993OTU.271OTU.580OTU.280OTU.952OTU.656OTU.813OTU.1021OTU.1194OTU.1304OTU.817OTU.352OTU.1438OTU.1643OTU.577OTU.721OTU.975OTU.11 OTU.1192OTU.1390 OTU.176OTU.241OTU.1803OTU.774OTU.782OTU.1543OTU.502 OTU.1320 OTU.763OTU.710OTU.1734OTU.489OTU.1534OTU.797OTU.1697OTU.918OTU.212OTU.214OTU.1370OTU.253OTU.788OTU.468OTU.4 OTU.442OTU.443OTU.601 OTU.1590OTU.287 OTU.1424OTU.195 OTU.366 OTU.920OTU.735OTU.30OTU.42OTU.843OTU.532OTU.872OTU.1536OTU.1672 OTU.1648OTU.1565OTU.426OTU.1696OTU.14 OTU.1579OTU.388 OTU.1593OTU.1623OTU.1128OTU.453OTU.1018 OTU.310 OTU.853 OTU.145OTU.1297 OTU.586OTU.1061OTU.829 OTU.719 OTU.885OTU.1602OTU.950OTU.1006OTU.209OTU.804OTU.752OTU.492OTU.178OTU.962OTU.570OTU.906 OTU.24OTU.1254 OTU.1187OTU.1281 0.0 OTU.556OTU.1651OTU.551OTU.951OTU.669OTU.73OTU.938OTU.899OTU.166OTU.1727OTU.1409OTU.32OTU.1487OTU.1690OTU.65OTU.1144 OTU.1322OTU.1330OTU.648OTU.25OTU.568 OTU.33OTU.863OTU.1279OTU.1156 OTU.1028OTU.407OTU.1123OTU.1179OTU.415OTU.610OTU.1067OTU.1195OTU.1813 OTU.1095OTU.647OTU.1639OTU.289OTU.1530OTU.898OTU.1051OTU.1717 OTU.1301OTU.164 OTU.1578OTU.942OTU.1342OTU.220OTU.945OTU.89OTU.883OTU.1576OTU.1174OTU.1180OTU.699OTU.1305OTU.1169OTU.1791OTU.724OTU.1338OTU.1127OTU.1366 OTU.347OTU.560OTU.546 OTU.423OTU.9OTU.1396OTU.1715OTU.1750OTU.302OTU.466OTU.1710OTU.1317OTU.1345OTU.157OTU.23OTU.246OTU.1426OTU.1408OTU.1558 OTU.50 OTU.273OTU.398OTU.252OTU.834OTU.255OTU.1677OTU.1560 OTU.1363OTU.835OTU.1276OTU.617OTU.748OTU.554OTU.1148OTU.1382OTU.1712OTU.304OTU.795OTU.1472OTU.1615OTU.68OTU.74OTU.1449OTU.1475OTU.895OTU.686OTU.1373OTU.1431OTU.1738OTU.591OTU.826OTU.1493OTU.923OTU.632OTU.263OTU.1757OTU.707OTU.218 OTU.385OTU.128OTU.264OTU.1466OTU.846OTU.547OTU.1585OTU.1447OTU.444OTU.1401OTU.1500OTU.606OTU.605OTU.1702OTU.868OTU.450OTU.750OTU.949OTU.992 OTU.201OTU.578OTU.943OTU.341OTU.723OTU.997OTU.293OTU.6OTU.1769OTU.1375OTU.1226OTU.977OTU.46 OTU.1537OTU.1260OTU.1355OTU.931 OTU.1695 OTU.595OTU.387OTU.187OTU.1085OTU.624OTU.581OTU.1811OTU.839OTU.82OTU.1110OTU.1685OTU.1549OTU.1344OTU.1722OTU.1245OTU.837OTU.52OTU.515OTU.380OTU.185OTU.517OTU.194OTU.1385 OTU.1542OTU.1503 OTU.604OTU.428 OTU.736 OTU.911 OTU.528OTU.1242OTU.1762OTU.1313OTU.22OTU.1600OTU.974OTU.77OTU.940OTU.1429OTU.1291OTU.1573OTU.588OTU.44OTU.845OTU.755OTU.437OTU.314OTU.732OTU.257OTU.1129OTU.1307OTU.487OTU.150OTU.1001OTU.1283OTU.1725OTU.524OTU.332 OTU.1227OTU.1789OTU.39 OTU.206OTU.1654OTU.1607OTU.1196OTU.1416OTU.877OTU.1190OTU.10OTU.1218OTU.1083OTU.1629OTU.521OTU.514OTU.1017OTU.511OTU.160OTU.623 OTU.242 OTU.613OTU.668OTU.349OTU.1737OTU.498 OTU.716OTU.1748OTU.368OTU.1668OTU.821OTU.1202OTU.53OTU.1631OTU.1011OTU.1611OTU.1272OTU.1331OTU.1022OTU.134OTU.379OTU.318OTU.531OTU.937OTU.1352OTU.870OTU.1441OTU.1232OTU.1619OTU.256OTU.12 OTU.1282OTU.1020 OTU.535 OTU.1249OTU.1730OTU.397 OTU.711 OTU.1112OTU.848OTU.809OTU.191OTU.1414OTU.1267OTU.1362OTU.354OTU.1073OTU.1776OTU.1239 OTU.1459OTU.1739OTU.97OTU.248OTU.63OTU.811OTU.1502OTU.1172OTU.827OTU.125 OTU.1403 OTU.694OTU.1115OTU.1693OTU.1029OTU.1694OTU.932OTU.488OTU.1181OTU.96OTU.140OTU.1358OTU.100OTU.1243OTU.1088OTU.1074OTU.772OTU.103 OTU.1299 OTU.1662 OTU.1367OTU.1772OTU.48OTU.1113OTU.331OTU.1666OTU.500OTU.1221 OTU.1026OTU.29OTU.683OTU.463 OTU.226 OTU.173 OTU.494OTU.311OTU.56 OTU.1333OTU.471 OTU.884OTU.1793 OTU.1251OTU.196 OTU.844OTU.467OTU.1720 OTU.1388OTU.1185 OTU.1767 OTU.858 OTU.1119 OTU.1818OTU.1065 OTU.1729 -0.2 OTU.1786OTU.1784OTU.1806 OTU.372OTU.1284OTU.590OTU.575 -0.4

-0.6 -0.4 -0.2 0.0 0.2 0.4 0.6 0.8

NMDS1 Figure 4.8: NMDS showing clustering of all pyrotag OTUs and the correlation of benzene degradation rate with OTUs.

A second NMDS was generated, using only OTUs present at one percent abundance or greater in at least one culture. The same variables were tested against this model. “Generation” correlated with this model with a p-value <0.01 (Figure 4.9). OTUs that correlate with “generation” are mostly Chlorobi/Ignavibacteriales. As the cultures are transferred, these OTUs are increasing in abundance. Finally, an NMDS was generated using only OTUs belonging to the groups tracked in a previous study on this culture (Gitiafroz, Devine, Raskin, & Edwards, n.d.). None of the variables tested against this model showed a significant correlation with the OTUs present.

OTU_57_Bacteria.Acidobacteria.Sva0725.Sva0725

OTU_1698_Bacteria.Chloroflexi.Anaerolineae.SBR1031.A4bGeneration

OTU_366_Bacteria.Proteobacteria.Gammaproteobacteria.Xanthomonadales.Xanthomonadaceae.Luteimonas OTU_197_Bacteria.Chlorobi.Ignavibacteria.Ignavibacteriales.Ignavibacteriaceae 0.5 OTU_810_Bacteria.Chlorobi.Ignavibacteria.Ignavibacteriales.Ignavibacteriaceae OTU_55_Bacteria.Proteobacteria.Betaproteobacteria.Rhodocyclales.Rhodocyclaceae.Dechloromonas OTU_1407_Bacteria.Proteobacteria.Betaproteobacteria.Rhodocyclales.Rhodocyclaceae OTU_1516_Bacteria.Chlorobi.Ignavibacteria.Ignavibacteriales.IgnavibacteriaceaeOTU_1552_Bacteria.Chlorobi.SJA.28

OTU_586_Bacteria.Acidobacteria.Sva0725.Sva0725

OTU_145_Bacteria.Proteobacteria.Betaproteobacteria.Burkholderiales.Comamonadaceae OTU_287_Bacteria.Proteobacteria.Betaproteobacteria.Rhodocyclales.Rhodocyclaceae.Dok59OTU_736_Bacteria.Proteobacteria.Betaproteobacteria.Hydrogenophilales.Hydrogenophilaceae.Thiobacillus OTU_12_Bacteria.Chlorobi.OPB56 OTU_1530_Bacteria.Chloroflexi.Anaerolineae.SBR1031.SHA.31OTU_604_Bacteria.Chlorobi.Ignavibacteria.Ignavibacteriales.IgnavibacteriaceaeOTU_668_Bacteria.Bacteroidetes.Bacteroidia.BacteroidalesOTU_1537_Bacteria.WPS.2 OTU_1254_Bacteria.Thermi.Deinococci.Deinococcales.Deinococcaceae.DeinococcusOTU_226_Bacteria.Planctomycetes..Brocadiae..Brocadiales.Brocadiaceae OTU_1164_Bacteria.Gemmatimonadetes.GemmatimonadetesSum_low_abundance_organisms OTU_1757_Bacteria.Acidobacteria.iii1.8.SJA.36OTU_1299_Bacteria.Proteobacteria.Betaproteobacteria.Hydrogenophilales.Hydrogenophilaceae.Thiobacillus 0.0 OTU_853_Bacteria.Proteobacteria.Betaproteobacteria.Burkholderiales.BurkholderiaceaeOTU_1322_Bacteria.Firmicutes.Clostridia.Clostridiales.Peptococcaceae OTU_1465_Bacteria.Bacteroidetes.Sphingobacteriia.Sphingobacteriales.Chitinophagaceae.FlavisolibacterOTU_1282_Bacteria.Proteobacteria.Alphaproteobacteria.Rhizobiales.Hyphomicrobiaceae.RhodoplanesOTU_263_Bacteria.Proteobacteria.Alphaproteobacteria.Rhodospirillales.Rhodospirillaceae OTU_788_Bacteria.Firmicutes.Clostridia.Clostridiales.PeptococcaceaeOTU_1793_Bacteria.Proteobacteria.Deltaproteobacteria.Desulfovibrionales.Desulfovibrionaceae.Desulfovibrio.mexicanus

NMDS2 OTU_535_Bacteria.Firmicutes.Clostridia.Clostridiales.Veillonellaceae.Desulfosporomusa OTU_1031_Bacteria.Proteobacteria.BetaproteobacteriaOTU_1409_Bacteria.Firmicutes.Clostridia.Clostridiales.Peptococcaceae OTU_692_Bacteria.Firmicutes.Clostridia.Clostridiales.Peptococcaceae OTU_372_Bacteria.Chlorobi.SJA.28 OTU_1067_Bacteria.Proteobacteria.Betaproteobacteria.Rhodocyclales.Rhodocyclaceae.AzoarcusOTU_920_Bacteria.Proteobacteria.Betaproteobacteria OTU_827_Bacteria.Proteobacteria.Betaproteobacteria.Rhodocyclales.Rhodocyclaceae.AzoarcusOTU_1767_Bacteria.Chlorobi.Ignavibacteria.Ignavibacteriales OTU_575_Bacteria.Proteobacteria.Gammaproteobacteria.Pseudomonadales.Pseudomonadaceae.Pseudomonas OTU_437_Bacteria.Proteobacteria.Betaproteobacteria.Burkholderiales.Comamonadaceae.RamlibacterOTU_293_Bacteria.Firmicutes.Clostridia.Clostridiales.Peptococcaceae OTU_895_Bacteria.Bacteroidetes.Bacteroidia.Bacteroidales.Porphyromonadaceae

OTU_1185_Bacteria.Proteobacteria.Betaproteobacteria OTU_1818_Bacteria.Proteobacteria.Deltaproteobacteria.DTB120

OTU_1115_Bacteria.Proteobacteria.Gammaproteobacteria OTU_196_Bacteria.Chlorobi.Ignavibacteria.Ignavibacteriales.Ignavibacteriaceae

-0.5 OTU_911_Bacteria

OTU_1607_Bacteria.Bacteroidetes.Bacteroidia.Bacteroidales.Porphyromonadaceae

OTU_694_Bacteria.Chlorobi.OPB56

-0.5 0.0 0.5

NMDS1 Figure 4.9: NMDS showing clustering of pyrotag OTUs present at greater than one percent abundance in any of the enrichments, and the correlation with generation.

4.2.7 Multiple OTUs of a single phylogeny

Many phylogenetic groups are represented by more than one OTU in this analysis. A high similarity threshold (99% sequence identity) was used to group sequences into OTUs for analysis. This has caused sequences that differ very slightly, likely due to sequencing error, to be grouped in to separate OTUs. Multiple Peptococcaceae OTUs are present, according to QIIME

classification. This could be due to sequencing error, where a random sequencing error results in reads being classified as separate OTUs. It could also reflect the presence of multiple Peptococcaceae OTUs in the enrichments. To ascertain the more likely scenario, all pyrotag OTUs classified as “Peptococcaceae” by QIIME were aligned in Geneious (version 6.1.7) (Kearse et al., 2012). This alignment is shown in Figure 4.10. Sequences fall into two groups. Sequences 1 - 18 form one group, and sequences 19 – 22 form another group. Two strains (at least) of Peptococcaceae are present in the enrichments. Within-group sequence variations were likely due to sequencing error. A similar scenario is envisioned for other phylogenetic groups observed in this study. Multiple OTUs with the same phylogenetic classification were considered to comprise a single population in this analysis.

Figure 4.10: Geneious alignment of Peptococcaceae OTUs showing two distinct groupings.

4.3 Key points

• Abundances of different OTUs varies widely between cultures.

• 18 OTUs are common to all cultures, and these OTUs are likely to be directly involved in benzene degradation or cycling of metabolites and inorganics produced during benzene degradation with nitrate reduction.

• There is redundancy of some functional groups in the cultures, which provides resilience to changing environmental pressures.

• OTUs present are highly similar to the groups found in other nitrate-reducing benzene- degrading enrichment cultures.

• No relationship was found between the abundances of the groups proposed to play key roles in the degradation process and benzene degradation rates.

• Two strains of Peptococcaceae are present in the cultures.

Chapter 5 Metagenomic analysis of a nitrate-reducing benzene-degrading culture 5 5.1 Introduction

Potential roles of some microbial groups in the nitrate-reducing, benzene-degrading enrichments used in this study have been elucidated by former students using qPCR and metatranscriptomic studies (Gitiafroz, 2012, Luo et al., 2014). Other nitrate-reducing, benzene-degrading enrichments also provide evidence of the roles of some key organisms. Peptococcaceae are thought to be responsible for the initial step of benzene degradation (Abu Laban et al., 2010, van der Zaan et al., 2012). Nitrate-reducing Azoarcus and Burkholderiales members are thought to be responsible for downstream metabolite degradation (Ulrich & Edwards, 2003). Azoarcus- specific genes increase in abundance when cultures are fed benzoate (Luo et al., 2014). Anammox bacteria may have a role in nitrogen cycling in this culture (Nandi, 2006). These noted groups were the dominant phylotypes detected in previous clone libraries, along with members of the Chlorobi/Ignavibacter clade (Ulrich & Edwards, 2003). As pyrotag sequencing shows, however, many other OTUS are present at similar abundances as these “key players”. The roles of other OTUs can be better predicted through analysis of their genomes. Furthermore, while complete genomes of members of the same phylogenetic clades as the “key players” have been released, genomes of strains actively carrying out benzene degradation under nitrate- reducing conditions have not been produced. Differences between the genomes of organisms enriched solely on benzene as an electron donor for 19 years and their close relatives that have not been exposed to benzene may provide insight in to adaptations that allow successful benzene degradation.

Advances in sequencing technologies and the development of software to deal with huge datasets allow more extensive analyses than could be obtained previously (Solomon et al., 2014). Assembly of genomes of organisms at low abundances in highly diverse enrichments with strain

variation present is still challenging though (Tang et al., 2012). Multiple assemblies are often required to optimize the assembly for different genomes, as genomes at different read depths can assemble better under different parameters (Tang et al., 2012, Devine, 2013). Automated assembly programs are often unable to carry on the assembly of a contig due to potential branching options, where repetitive regions or strain variation exists. Paired end and mate pair data can be used together to bridge gaps in some of these cases. Paired end data consists of 150 base pair (bp) reads, with an insert size of 1,500bp. Mate pair data consists of 100bp reads, with an insert size ~10,000bp. Mate pair data can be used to find potential bridges between contigs, where one read lies on one contig, and its pair lies on another contig. This technique can produce high-quality long scaffolds suitable for some analyses and manual assembly efforts.

The three goals of this metagenome assembly work were to 1) Produce nearly-complete genomes from the most abundant organisms, along with a set of unjoined contigs for downstream analysis, 2) Provide further evidence of the benzene degradation pathway in these cultures, and 3) Reveal novel insights into the potential roles of supplementary organisms present in the nitrate-reducing benzene-degrading cultures.

5.2 Results and discussion

Metagenomic sequencing was carried out for one actively degrading anaerobic benzene- degrading nitrate-reducing consortium. The consortium contains material consolidated from past experiments that were inoculated with soil taken from a contaminated retired gas station in Ontario (Cartwright Gas Station), and is referred to as "Cart T3" throughout this thesis. Degradation rate was 0.36 mg benzene/L/day at the time of sampling in 2013. Mate pair sequence data and paired end data were both produced by Genome Quebec. The locations of raw data files are given in Appendix 5.

Four sequencing runs of paired end (PE) libraries were carried out. Paired end libraries produced ~35M reads each, ~60% of which survived trimming. One sequencing run of a mate pair (MP) library was carried out, producing ~200M reads, ~75% of which survived trimming.

Trimming was carried out using the default parameters in Trimmomatic (Bolger et al., 2014). Only reads for which both the forward and reverse read of the pair passed the trimming step were used for subsequent assembly (121M PE reads, 153M MP reads).

5.2.1 ABySS Assembly

ABySS was used to generate contigs (Simpson et al., 2009). Different parameters were varied to optimize the assembly of different OTUs. The value of the parameter “K” (kmer) was kept at 96, the maximum kmer length possible in ABySS. The term kmer refers to all possible substrings within the string of length “k”. A lower kmer length can increase the chances of a contig growing, as more substrings are likely to overlap. This is a problem where there are ambiguities, as it further increases the number of options for contig extension. As ambiguities were considered likely, the longest kmer length available was chosen. The parameter “c” refers to the cut-off threshold. Contigs with mean kmer coverage below c are removed from the assembly before it continues. This can be useful to remove high-abundance or repetitive regions from the assembly that may be confounding the assembler. c values between 10 – 100 were used. The amount of data fed into ABySS was also optimized. Assembly using all PE reads that passed quality trimming were used in the first assembly (k96, c20). This assembly produced contigs up to 500,000 bp. Further assemblies used a single PE library, and different values of c. Results of selected assemblies are presented in Figure 5.1. Data for these figures were pulled out of ABySS assembly files using a PERL script written by Cheryl Devine. The plots show contig length versus Normalized Kmer Depth (NKD). NKD is a proxy for read depth in contigs, calculated by dividing the number of kmers in a contig by the contig length minus the kmer length plus one. Comparing Figure 5.1a and 5.1b, the former used all four PE libraries whereas 5.1b used just one PE library (a quarter of the amount of sequence data). Both used k96, c20. The use of four times the amount of data in the first assembly did not result in a much better assembly (not significantly longer contigs; and not better separation of different OTUs by NKD); therefore, a single library was used for further assemblies. Lower and higher c values were used (Figures 5.1c and5.1d). These showed that a high c value (c=100) did not produce relatively long contigs.

The assemblies with one PE library, k96, c10 and c20 were very alike. These assemblies were used in further steps.

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Figure 5.1:a) Contigs from an ABySS assembly using four paired end libraries (k96, c20).

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Figure 5.1:c) Contigs from an ABySS assembly using one paired end library (k96, c10).

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Figure 5.1:d) Contigs from an ABySS assembly using one paired end library (k96, c100).

5.2.2 ALLPATHS-LS Assembly

ALLPATHS-LG was used to produce long scaffolds. This program makes use of both PE reads, and MP reads (Gnerre et al., 2011). Different amounts of sequence data were also sent through this program. Better results were generated when more data was used (~4M PE reads and ~6M MP reads). This is likely due to the low abundance of all OTUs of interest in the enrichment. Using this amount of data and ALLPATHS-LG default parameters, only three scaffolds were produced that were longer than 2M bp. When less data was used, these scaffolds broke up.

Reads were digitally normalized by DigiNorm (Brown, Howe, Zhang, Pyrkosz, & Brom, 2012). Digital normalization can facilitate the assembly of low abundance organisms that might otherwise be lost amongst the overwhelming volume of sequence data. It does however cause the loss of read depth data during assembly. Read depth of scaffolds can be approximated after assembly by mapping reads back to assembled scaffolds and calculation mean read depth from this. Digitally normalized reads (~4.5 million paired end reads and ~6.5 million mate pair reads) were assembled using ALLPATHS-LG default parameters. From this assembly, 20 scaffolds were generated longer than three million base pairs. Scaffolds from this assembly were used for further analysis.

Table 5.1 shows the long scaffolds produced by ALLPATHS-LG that could be identified by searching for 16S rRNA genes in the scaffolds. Shown is the scaffold number assigned by ALLPATHS-LG, the length of the scaffold, the percent of the scaffold that is gaps, the percent pairwise identity of the 16S rRNA gene to the nearest match in the NCBI Whole Genome Database, and the genus and order of that match. Long scaffolds are present for a number of groups of interest. Scaffold 5 (4.7M bp) is a Burkholderiales. This scaffold is 20% gaps. Another scaffold of interest is Scaffold 2, a 4.5M bp scaffold that is most similar to Melioribacter, with 3.7% gaps. Scaffold 10 is another Iganvibacteriales. Scaffold 12 is an Aromatoleum. Scaffold 29 and 36 are Peptococcaceae, which both have around 13% gaps, and are 2.1M and 1.6M bp respectively.

Table 5.1: Scaffolds that contain a 16S rRNA sequence identified from ALLPATHS-LG assembly using digitally normalized data.

Scaffold % pairwise number Most similar 16SrRNA Scaffold identity to Order ( - where genus (diginorm % gaps sequence in whole length (bp) closest 16S in already covered) ALLPATHS genome database WG database run)

Scaffold 1 4,786,733 11.6 95.1 Opitutus Verrucomicrobiales Scaffold 5 4,780,120 20.1 97.3 Curvibacter Burkholderiales Scaffold 5 4,780,120 20.1 99.7 Comamonas Burkholderiales Scaffold 3 4,763,066 9.2 91.7 Chitinophaga Sphignobacteriales Scaffold 0 4,599,941 5.1 93.6 Oceanibaculum Rhodospirillales Scaffold 0 4,599,941 5.1 88.5 Phenylobacterium Caulobacterales Scaffold 0 4,599,941 5.1 89.2 Nisaea Rhodospirillales Scaffold 2 4,558,958 3.7 84.5 Melioribacter Ignavibacteriales Scaffold 4 4,433,340 1.2 92.8 Bradyrhizobium Rhizobiales Scaffold 6 4,339,521 1.5 84.5 Anaerolinea Anaerolineales Scaffold 6 4,339,521 1.5 84.5 Anaerolinea - Scaffold 6 4,339,521 1.5 84.5 Anaerolinea - Scaffold 10 4,157,143 4.5 93.4 Ignavibacterium Ignavibacteriales Scaffold 7 4,146,149 4.2 91.2 Gemmatimonas Gemmatimonadales Scaffold 7 4,146,149 4.2 91.2 Gemmatimonas - Scaffold 8 4,143,667 2.3 98.9 Pseudomonas Pseudomonadales Scaffold 8 4,143,667 2.3 99.2 Pseudomonas - Scaffold 8 4,143,667 2.3 99.0 Pseudomonas - Scaffold 11 4,115,703 9 98.8 Thiobacillus Hydrogenophilales Scaffold 12 3,738,300 2.4 98.2 Nitrosomonas Nitrosomonadales Scaffold 12 3,738,300 2.4 98.2 Aromatoleum Rhodocyclales Scaffold 14 3,443,583 5.4 97.8 Thiobacillus - Scaffold 15 3,162,071 2 95.9 Terrimonas Sphignobacteriales Scaffold 16 3,106,881 0.8 94.4 Prosthecochloris Chlorobiales Scaffold 17 3,007,566 0.5 86.2 Desulfovibrio Desulfovibrionales Scaffold 20 2,981,289 5.5 94.5 Melioribacter - Scaffold 19 2,849,200 1.6 90.9 Truepera Deinococcales Scaffold 21 2,670,708 6.8 95.8 Terrimonas - Scaffold 22 2,575,370 0 92.2 Desulfovibrio - Scaffold 24 2,394,295 14.5 85.5 Ignavibacterium - Scaffold 24 2,394,295 14.5 87.3 Ignavibacterium -

Scaffold 25 Candidatus 2,351,371 5.6 81.2 omnitrophus Unclassified Scaffold 27 2,249,054 8.6 93.7 Ignavibacterium - Scaffold 29 2,120,880 12.5 90 Pelotomaculum Clostridales Scaffold 28 2,049,086 6.8 100 Methyloversatilis Rhodocyclales Scaffold 36 1,646,985 13.7 89.9 Pelotomaculum - Scaffold 36 1,646,985 13.7 92.3 Thermincola Clostridales Scaffold 37 1,455,376 1.9 84.9 Holophaga Acidobacteriales Scaffold 42 981,250 22.5 98.7 Gallionella Nitrosomonadales Scaffold 51 827,218 32.4 90.8 Aciditerrimonas Acidimicrobiales Scaffold 51 827,218 32.4 91.6 Acidimicrobium Acidimicrobiales Scaffold 45 813,270 7.4 90.6 Nisaea - Scaffold 48 769,958 0 76.4 Muricauda Flavobacteriales Scaffold 75 Prolixibacter Unclassified 725,663 47.8 90.5 bacteriodetes Scaffold 70 680,155 25.9 99.5 Thiobacillus - Scaffold 77 658,858 30.6 99.1 Thiobacillus - Scaffold 94 619,385 39.4 100 Thiobacillus - Scaffold 138 534,803 70.7 88.8 Anaeromyxobacter Myxococcales Scaffold 113 473,638 34.7 100 Thiobacillus - Scaffold 260 356,475 87.6 100 Pseudomonas - Scaffold 282 356,475 82.5 89.9 Byssovorax Myxococcales Scaffold 161 319,134 47.7 95.6 Parvibaculum Rhizobiales Scaffold 152 267,307 10.1 97 Dechlorosoma Rhodocyclales Scaffold 152 267,307 10.1 92.4 Sideroxydans Gallionellales Scaffold 206 216,852 27.8 92.5 Sideroxydans - Scaffold 200 186,020 12.3 100 Nitrosomonas - Scaffold 302 85,966 10.1 100 Nitrosomonas - Scaffold 694 83,993 88.6 96.3 Thermanaerothrix Anaerolineales Scaffold 358 66,727 42.9 94.3 Lautropia Burkholderiales Scaffold 511 51,843 78 87.3 Ignavibacterium - Scaffold 1323 27,196 89.7 93.6 Geobacter Desulfuromonadales Scaffold 1456 No hits for putative 16S gene in whole 25,396 91.7 genome database Scaffold 1187 No hits for putative 16S gene in whole 10,473 45.6 genome database

Potential roles of organisms in the enrichment can be hypothesized from phylogeny of the organisms. Table 5.2 shows functions ascribed to representatives of the genera mentioned in Table 5.1. The organisms in the enrichment may be carrying out these functions. There is considerable variation in the capabilities of different members of genera, so identifying specific functional genes in the genomes of these organisms would provide stronger evidence of their capabilities.

Table 5.2: Functions ascribed to genera present in assembly.

Genus Functions ascribed to type strain of genus Reference Opitutus Obligate anaerobe, fermentation of organic (Chin, Liesack, & Janssen, compounds, nitrate reducer 2001) Curvibacter Chemoheterotroph (Ding & Yokota, 2004) Comamonas Aerobic or anoxic denitrifier (Gumaelius et al., 2001) Chitinophaga (Sangkhobol & Skerman, Chemoheterotroph 1981) Oceanibaculum Polycyclic aromatic hydrocarbon degrader, facultatively anaerobic (Lai & Shao, 2012) Phenylobacterium Aerobic xenobiotic degrader (Lingens et al., 1985) Nisaea (Urios, Michotey, Intertaglia, Lesongeur, & Anaerobic denitrifier Lebaron, 2008) Melioribacter (Podosokorskaya et al., Facultatively anaerobic chemoorganotroph 2013) Bradyrhizobium Nitrogen fixation (Kaneko et al., 2002) Anaerolinea Anaerobic chemoorganotroph (Sekiguchi, 2003) Ignavibacterium Anaerobic chemoorganotroph (Iino et al., 2010) Gemmatimonas Aerobic chemoorganotroph, phosphate accumulator (H. Zhang, 2003) Pseudomonas Aerobic, great metabolic diversity (Spencer et al., 2003) Thiobacillus Chemolithoautotroph, thiosulfate oxidation/denitrification, CO2 fixation, can also carry out aerobic respiration (Beller et al., 2006) Nitrosomonas Chemolithoautotroph, ammonia oxidiser, CO2 fixation (Chain et al., 2003) Aromatoleum Anaerobic hydrocarbon degrader, denitrifier (Wöhlbrand et al., 2007) Terrimonas Aerobic chemoorganotroph, nitrate reducer (Xie & Yokota, 2006) Prosthecochloris Strictly anaerobic photolithotroph (Gorlenko, 1970)

Desulfovibrio Anaerobic sulfate reducer (Devereux et al., 1990) Truepera Aerobic chemoorganotroph, radiation resistant (Albuquerque et al., 2005) Pelotomaculum (Imachi et al., 2002) Anaerobic syntrophic propionate oxidiser Methyloversatilis Facultative methylotroph (Kalyuzhnaya et al., 2006) Thermincola Anaerobic, carboxydotrophic, hydrogenogenic (Sokolova et al., 2005) Holophaga Obligately anaerobic chemoorganotroph, (Liesack, Bak, Kreft, & aromatics degrader, acetogen Stackebrandt, 1994) Gallionella Iron oxidising chemolithotroph (Emerson et al., 2013) Aciditerrimonas (Itoh, Yamanoi, Kudo, Capable of anaerobic or aerobic respiration, Ohkuma, & Takashina, iron reducer 2011) Acidimicrobium Aerobic iron oxidizer (Clark & Norris, 1996) Muricauda (Bruns, Rohde, & Berthe- Faculative aerobe, chemoheterotroph Corti, 2001) Prolixibacter (Holmes, Nevin, Woodard, Sugar- fermenting anaerobe Peacock, & Lovley, 2007) Anaeromyxobacter Facultatively anaerobic chemoorganotroph, (Sanford, Cole, & Tiedje, chlororespiring 2002) Byssovorax (Reichenbach, Lang, Schumann, & Spröer, Aerobic chemoorganotroph 2006) Parvibaculum (Schleheck, Tindall, Rosselló-Mora, & Cook, Aerobic chemoorganotroph 2004) Dechlorosoma (Achenbach, Michaelidou, Bruce, Fryman, & Coates, Anaerobic perchlorate reduction 2001) Sideroxydans Obligate lithotrophic iron oxidizers (Weiss et al., 2007) Thermanaerothrix Strictly anaerobic chemoorganotroph (Grégoire et al., 2011) Lautropia Aerobic chemoorganotroph, nitrite and nitrate reducer (Kei et al., 1994) Geobacter Anaerobic organotroph, iron reducer (Lovley et al., 1993)

5.2.3 Assessment of assemblies

Selected scaffolds from digitally normalized ALLPATHS-LG assembly were analysed by myRAST (Aziz et al., 2008). Theoretical protein encoding genes (PEGs) were called, and these PEGs were then identified. Where no identity to known proteins was found, PEGs were labeled “hypothetical proteins”. Proportions of identified proteins are listed in Table 5.3. The genomes analysed vary between 33 – 71% unidentified proteins, showing a lot of novel genes are potentially present in the genomes. Table 5.3: Proportions of potential genes in each scaffold putatively identified by myRAST Number Protein Encoding Genes Number of % unidentified Scaffold identity (PEGs) hypothetical proteins by myRAST Thiobacillus 3977 1486 37.4 Aromatoleum 3506 1161 33.1 Melioribacter 4143 2939 70.9 Pelotomaculum 2157 1319 61.1 Thermincola 1597 951 59.5

A set of genes found in all bacterial genomes in single copy was identified by Cheryl Devine (Devine, 2013). These genes can be used to assess genome completeness. The proportion of these genes found in an assembled genome or scaffold belonging to a single OTU reflects the proportion of the genome represented in the sequence scaffold. Selected scaffolds were assessed using this test. All genes used for this assessment are shown in Appendix 1, along with hit/miss results for the chosen scaffolds. Overall genome completeness estimates are shown in Table 5.4. The later shows that some assemblies, such as Scaffold 2, are more complete than others, such as Scaffolds 29 and 36. This is evident from Table 5.1 also, where the lengths of scaffolds and the percent gaps give a measure of how much of a genome is actually present. More work can be

Table 5.4: Proportions of genes in each scaffold from single-copy gene set putatively identified by myRAST

Thiobacillus Aromatoleum Melioribacter Peptococcaceae Peptococcaceae Scaffold 11 Scaffold 12 Scaffold 2 Scaffold 29 Scaffold 36 % Complete 73.8 60.7 83.6 18.0 16.4 (Observed/Expected*100%)

done on these assemblies to further assess how much of the genomes are present, and methods such as that described in (Tang et al., 2012) could be used to reduce the number of gaps in the scaffolds, and to find the bridges between scaffolds. Near-complete scaffolds can be produced from this metagenome assembly.

5.2.4 Multiple Peptococcaceae present in the enrichment?

A key question of the Peptococcaceae scaffolds (Scaffold 29 and Scaffold 36) is whether they represent a single organism that has not assembled together, or whether more than one Peptococcaceae is present in the cultures. These scaffolds do not map to each other, and when mapped separately to a completed Thermincola potens genome (Byrne-Bailey et al., 2010) using Mauve (Darling, Mau, Blattner, & Perna, 2004), they show very different results (Figure 5.2). This supports the evidence presented in Chapter 4, where Peptococcaceae pyrotag 16S rRNA sequences clustered into two different groups when aligned together. Two Peptococcaceae OTUs are present in the cultures. The similar genomes may have interfered with each other during assembly. Similar but slightly different genomes present a challenge to assemblers because the choices of which of multiple possibilities for extending contigs to follow or for joining together different contigs become more difficult.

Figure 5.2a) Scaffold 29 shows 69% pairwise identity to Thermincola potens genome

Figure 5.2b) Scaffold 36 shows 26% pairwise identity to Thermincola potens genome

5.2.5 Identification of putative anaerobic benzene carboxylase operon

The putative anaerobic benzene carboxylase operon abcD → abcA → bzlA → ubiX is thought to be responsible for the first step of benzene degradation by carboxylation of benzene. This operon was first proposed in (Abu Laban et al., 2010), and the complete operon was found in the transcriptome of the nitrate-reducing, benzene-degrading enrichment studied in this project by Fei Luo (Luo et al., 2014). This operon was present (100% sequence identity) on Scaffold 559 from the digitally normalized ALLPATHS-LG assembly. Scaffold 559 is 34,305bp, with no gaps. This provides more context of the gene neighbourhood around the abc operon. Scaffold 559 does not contain a 16S rRNA gene to identify which organism it belongs to. (Luo et al., 2014) found that this operon was up-regulated when other Peptococcaceae -associated genes were up-regulated. The abc operon is greater than 80% identical at the nucleotide level to the sequence from (Abu Laban et al., 2010). This study also suggested that the operon belongs to Peptococcaceae, as their enrichment was highly dominated by Peptococcaceae. In addition to this, when raw reads are mapped back to assembled scaffolds, Scaffold 559 has a kmer depth of 13, similar to Scaffold 29. This evidence all supports the abc operon belonging to Peptococcaceae.

Scaffold 559 was annotated using myRAST. Putative annotations are presented in Table 5.5, along with the percent similarity, identity and accession number of the closest BLAST hits in the nr database on NCBI. Shaded PEGs are those included in the abc operon (prot_05 = ubiX, prot_06 = bzlA, prot_07 = abcA, prot_08 = abcD). Three additional PEGs similar to abcA are present (prot_20, prot_24 and prot_25). These may have arisen through duplication of sections of the DNA, and suggest that the gene neighbourhood is under intense selection pressure. Many PEGs were highly similar to those found in Abu Laban et al., (2010), and were found in the same order on Scaffold 559 as on Contig BF_11418 from Abu Laban et al., (2010.

53 (

( ( ( ( ( ( ( ( ( ( ( ( ( ( Accession( number(of( closest(hit GU357996. 1 CP000612. 1 GU357995. 1 GU357994. 1 GU357993. 1 GU357992. 1 GU357991. 1 GU357990. 1 GU357988. 1 GU357987. 1 GU357986. 1 GU357985. 1 GU357984. 1 GU357983. 1 GU357982. ( ( ( ( ( ( ( ( ( ( ( ( ( BF( BF( BF( BF( BF( BF( BF( BF( BF( BF( BF( BF( BF( BF( ( ( ( ( ( ( ( ( ( ( ( ( * Contig& & ( operon&are& Source(of(closest( hit Clostridia* enrichment Desulfotomaculum* reducens Clostridia* enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( abc$ Contig&BF_11418& 82 84 83 97 96 98 97 97 98 98 97 93 96 88 94 & on ( %( identity( to(closest( hit ( ( ( ( ( ( r&is&found&in&Scaffold&559&as ( ( ( ( binding(protein( J CoA(ligase(BzlA( J orde & family(transcriptional( ( ( ( like(carboxylase(gene protein(gene J ( ( ORF MarR(family(transcriptional( multidrug(resistance(protein( ( ( ( similar&to&those&previously&found& & ) putative(benzoate hypothetical(protein(gene hypothetical(protein(gene hypothetical hypothetical(protein(gene The&same& Closest(BLAST(hit(to(nr/nt(database putative(MarR regulator(gene transposase,(IS4(family putative( regulator(gene putative(UbiX ( gene putative(anaerobic(benzene( carboxylase(AbcA(gene putative(anaerobic(benzene( carboxylase(AbcD(gene putative gene putative(ATPase(gene ( ( ( putative(FmdB(family(regulatory( protein(gene ( putative(nucleotide ( ( ORFs ( ) like J J 2010).& & ., ( ( ( 4S]( ( ) J J & hydroxybenzoate( hydroxybenzoate( J J ( ( ( ( 4 4 ( S(cluster(assembling( S(cluster(assembling( J J reading&frames&( J J & ( CoA(ligase((EC(6.2.1.25) J lyase(UbiX((EC(4.1.1. lyase((EC(4.1.1. J J Spn1,(transposase Abu&Laban&et&al J ( ( ( prot_14. open polyprenyl polyprenyl haloalkanoic(acid(dehalogenase( J J J Annotation(from(myRAST Transcriptional(regulator,(MarR( family IS1380 Transcriptional(regulator,(MarR( family 3 carboxy Benzoate 3 carboxy No(annotation Cytosolic(Fe factor(NBP35 Scaffold(protein(for([4Fe cluster(assembly(ApbC,(MRP No(annotation 2 (EC(3.8.1.2) No(annotation Type(I(antifreeze(protein No(annotation Cytosolic(Fe ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( blzA ubiX abcA abcD enrichment& & ( ” abc$ operon subunit s ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( BF & ( 1523 2540 4350 5485 6134 7821 9379 10008 11015 12101 12559 13308 13777 14025 14469 Stop between&prot_04&and& Putative&annotation&of&protein&encoding&genes&on&Scaffold&559.&The&four&subunits&of&the& & & ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( Clostridia “ & 2119 3778 5381 6078 7759 9347 9759 11015 12064 12496 13236 13745 13980 14393 15392 Start ! ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( Table&5.5: present,&along&with&many&other& from BF_11418

Putative( protien( number prot_01 prot_03 prot_04 prot_05 prot_06 prot_07 prot_08 prot_09 prot_10 prot_11 prot_12 prot_13 prot_14 prot_15 prot_16

( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( 1 GU357981. 1 GU357980. 1 GU357979. 1 GU357978. 1 GU357981. 1 GU357977. 1 GU357975. 1 GU357974. 1 GU357995. 1 GU357981. 1 CP001336. 1 CP000724. 1 CP000612. 1 CP000724. 1 GQ214703. 1 AP009050. 1| * * * ( ( ( ( ( ( ( ( ( ( ( ( BF( BF( BF( BF( BF( BF( BF( BF( BF( BF( * ( ( ( ( ( ( ( ( ( ( * enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Clostridia enrichment Desulfitobacterium* hafniense Alkaliphilus* metalliredigens Desulfotomaculum* reducens Alkaliphilus* metalliredigens Bacillus(phage( Clostridium*kluyveri* ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( 99 99 99 59 99 96 32 43 55 52 56 60 48 69 100 100 ( ( ( ( ( ( ( lyase(gene alanine( ( ( J J ( ( L J J ( 4 J pCKL1(DNA ( carboxylase(gene J polyprenyl J transcriptase) phenylphosphate(carboxylase( ( acetylmuramoyl J directed(DNA(polymerase( ( ( ( ( J gene putative(multidrug(resistance(protein( gene hypothetical(protein(gene hypothetical(protein(gene putative(3 hydroxybenzoate(carboxy putative(multidrug(resistance(protein( gene putative alpha(subunit(PpcA(gene putative(phenylphosphate(carboxylase( alpha(subunit(PpcA(gene putative(UbiD putative(MarR(family(transcriptional( regulator(gene putative(multidrug(resistance(protein( gene transcriptional(regulator,(Fis(family conserved(hypothetical(protein conserved(hypothetical(protein RNA (Reverse( Frp2(N amidase(family(2(protein(gene NBRC(12016(plasmid( ( ( ( ( ( ( ( ) ) ) ) ) ( J J J J J ( directed(DNA( J hydroxybenzoate( hydroxybenzoate( hydroxybenzoate( hydroxybenzoate( hydroxybenzoate( J J J J J ( ( ( 4 4 4 4 4 ( ( ( ( S(cluster(assembling( S(cluster(assembling( S(cluster(assembling( J J J J J J J J ( lyase((EC(4.1.1. lyase((EC(4.1.1. lyase((EC(4.1.1. lyase((EC(4.1.1. lyase((EC(4.1.1. J J J J J type(RNA J ( annotation polyprenyl polyprenyl polyprenyl polyprenyl polyprenyl J J J J J factor(NBP35 Cytosolic(Fe factor(NBP35 3 carboxy No( 3 carboxy Cytosolic(Fe factor(NBP35 3 carboxy 3 carboxy 3 carboxy Transcriptional(regulator,(MarR family Cytosolic(Fe factor(NBP35 two(component,(sigma54( specific,(transcriptional(regulator,( Fis(family No(annotation No(annotation Retron polymerase((EC(2.7.7.49) Autolysin,(amidase DNA(adenine(methylase ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( Similar( Similar( Similar( to(abcA to(abcA to(abcA ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( 15436 16459 17981 18344 21092 22021 23626 25229 26578 27125 28762 29738 30247 31677 32623 33467 ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( ( 16365 17955 18277 19888 22024 23568 25050 26560 27138 28156 28622 28815 29924 30580 31784 32712

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prot_17 prot_18 prot_19 prot_20 prot_22 prot_23 prot_24 prot_25 prot_26 prot_27 prot_28 prot_29 prot_30 prot_31 prot_32 prot_33

5.2.6 Identification of other enzymes in the anaerobic benzene degradation pathway

Selected scaffolds were checked for genes relating to anaerobic benzene degradation pathways. PEG annotations from myRAST (Aziz et al., 2008) were used as the basis for this search. Scaffolds that were used were chosen as they represent the organisms most likely to be involved in the benzene degradation pathway at some stage. The results are shown in Table 5.6. Most scaffolds checked have a benzoyl-CoA reductase, and half have a benzoyl-CoA ligase. None of the long scaffolds checked have genes that encode the proposed enzymes for the first steps of benzene degradation. This is the anticipated result, as the abc operon was only found in a short scaffold, and no other proposed benzene degradation mechanism is expected in the metagenome. Other enzymes of interest, such as nitrate reduction enzymes can be located and their host organisms identified using this method.

Table 5.6: Presence of genes encoding enzymes responsible for different stages of anaerobic benzene degradation in selected scaffolds (X indicates present, - indicates absent)

Pathway Key Gene Scaffold 2 Scaffold 5 Scaffold 11 Scaffold 12 Scaffold 29 Scaffold 36 Melioribacter Burkholderia- Thiobacillus Aromatoleum Peptococca- Peptococca- les ceae ceae Toluene Benzylsucc------inate synthase Phenol Phenylpho------sphate synthase Phenol Phenylpho------sphate carboxylase Benzoate Anaerobic ------benzene carboxylase Benzoate Benzoate- - X - X - X central CoA ligase metabolism Benzoate Benzoyl- X X - X X X central CoA metabolism reductase

5.3 Key points

• Metagenomic assembly produced a number of large scaffolds and a mixture of contigs. These scaffolds and contigs vary in completeness and quality.

• There are two strains of Peptococcaceae present in the enrichment.

• The putative anaerobic benzene carboxylase operon abcD → abcA → bzlA → ubiX is present in the assembly. It did not assemble to the longer Peptococcaceae scaffolds.

• Genes encoding enzymes from later steps of the anaerobic benzene degradation pathway are present in many of the long scaffolds. Genes encoding enzymes for alternative benzene degradation pathways are absent in all scaffolds checked.

• Comparisons of the scaffolds and contigs assembled here to other related organisms that do not live on benzene could yield interesting results.

• Further assembly efforts on this dataset are worthwhile.

• More investigation of the functional capabilities of the organisms represented by the assembled metagenome, or of the potential activities of the microbiome treated as a whole, is also justified.

Chapter 6 Methanogenic capability of nitrate-reducing benzene-degrading culture 6 6.1 Introduction

The organism thought responsible for the initial step in benzene degradation under nitrate reducing conditions is a Peptococcaceae. This is supported by evidence from independent cultures (van der Zaan et al., 2012, Luo et al, 2014). In contrast, the proposed benzene degrader under methanogenic conditions is a Syntrophobacterales OTU known as ORM2 (Devine, 2013). There is evidence of syntrophic processes in these cultures. No isolation efforts have been successful, and some terminal electron acceptor variability has been demonstrated (Ulrich and Edwards, 2003, van der Zaan et al., 2012). Terminal electron acceptor variability is evidence of syntrophy because intermediates produced by the initiator of benzene degradation can be consumed by a variety of other organisms during syntrophy, each of which may utilize different electron acceptors. If one organism carried out the entire process, wide terminal electron acceptor variability would not be expected.

Peptococcaceae may hand off benzene degradation products to syntrophic partners at different stages of the degradation pathway compared to ORM2. This may influence which potential syntrophic partners are most competitive under each condition. Methanogens present in nitrate-reducing cultures may or may not be preferred syntrophic partners of ORM2, and vice versa, ORM2-preferred methanogens may not be the most competitive syntrophic partners for Peptococcaceae. Other intermediary organisms capable of steps between benzoate and fermentation end-products could also be more competitive under nitrate reducing or methanogenic conditions.

Benzene degrading cultures maintained during this project were supplied nitrate as the dominant electron acceptor for 17 years before the experiment described in this chapter was

initiated. Despite this, a small amount of methane production was occasionally observed, when nitrate was depleted before benzene was fully removed. Methane production was observed through a small peak on GC, and by pressure build-up under butyl rubber stoppers on maintenance bottles. Pyrotag sequencing showed that nitrate-reducing benzene-degrading cultures have methanogens present at low abundances (Table 4.3) An attempt was made to switch consortia from degrading benzene with nitrate as the terminal electron acceptor to degradation with methanogenesis by allowing nitrate to become depleted so that carbon dioxide was the only terminal electron acceptor available.

Undiluted nitrate-reducing benzene-degrading culture was used to set up experimental bottles as the organisms involved have slow growth rates and the use of undiluted culture should allow the activity of the cultures to be observed sooner. Experimental bottles were also set up with nitrate-reducing benzene-degrading culture blended with methanogenic toluene-degrading culture. The aim of these bottles was to provide a high-abundance methanogenic partner for the benzene degraders, presuming the Peptococcaceae could hand off metabolites to the methanogens present in the toluene-degrading culture. Experimental bottles with toluene- degrading methanogenic culture were set up to test whether the toluene-degrading culture could switch to benzene degradation.

ORM2 is not present in the nitrate-reducing benzene-degrading cultures. Several Syntrophobacterales OTUs are present in the nitrate reducing cultures at low abundance, but none of the Syntrophobacterales present are greater than 87% similar at the 16S level to ORM2. Any benzene degradation that takes place in this experiment, therefore, is likely to be carried out by Peptococcaceae.

More energy is potentially available from benzene degradation with nitrate reduction than via benzene degradation with methanogenesis (ΔG° –1672 kJ/mol benzene with nitrate reduction versus ΔG° -108 kJ/mol benzene with methanogenesis (values for calculating standard free change energies from (Thauer et al., 1977), according to the methods in (Rittmann & McCarty, 2001)). Nitrate reduction should, therefore, be favoured when nitrate is available, but

methanogenesis presents the advantage that no external electron acceptor is required, and potentially inhibiting products (such as nitrite) are not formed.

6.2 Results

Experimental bottles were initially set up in serum bottles with butyl rubber stoppers. Benzene sorption to the rubber was a problem, and cultures were moved to mininert-cap sealed bottles on Day 133. One replicate (“Toluene culture only Replicate C”, Figure 6.9) was maintained in a butyl-rubber stoppered bottle, as it had shown a decrease in benzene concentration and increase in methane concentration by that point. After replicates were moved to mininert-stoppered bottles, less variation in measurements was produced. Gas chromatograph results for benzene, methane, and toluene (where relevant) are shown from Day 133 (Figure 6.1 – 6.12). Arrows indicate feeding points, and are labeled with the substrate added. Amounts fed per feeding point - for the respective substrate: benzene: 11umol/bottle; toluene: 11umol/bottle; NO3 : 80umol/bottle.

18" Benzene Benzene 16" Methane" 14" (uM"total)"

12" Benzene" (uM"total)" 10"

uM# 8"

6" umol/bottle

0" 0" 100" 200" 300" 400" 500" 600" 700" Days#

Figure 6.1: Nitrate-reducing benzene degrading culture only (without nitrate). Replicate A.

30" Benzene Benzene

25"

20"

15" uM# Methane" (uM"total)" umol/bottle 10" Benzene" (uM"total)"

0" 0" 100" 200" 300" 400" 500" 600" 700" Days#

Figure 6.2: Nitrate-reducing benzene degrading culture only (without nitrate). Replicate B.

30" Benzene Benzene Benzene Benzene

25"

20" Methane"(uM"total)"

15" uM#

umol/bottle 10"

5" Benzene"(uM"total)"

0" 0" 100" 200" 300" 400" 500" 600" 700" Days#

Figure 6.3: Nitrate-reducing benzene degrading culture only (without nitrate). Replicate C.

14" - Benzene NO3 Benzene

12"

Benzene"(uM"total)" 10"

uM# 6" Methane"(uM"total)" umol/bottle 4"

0" 0" 100" 200" 300" 400" 500" 600" 700" Days#

Figure 6.4: Nitrate-reducing benzene culture blended with methanogenic toluene- degrading culture. Amended with nitrate on Day 325. Replicate A.

14" Benzene - Benzene NO3

12"

Methane"(uM" 10" total)"

Benzene"(uM"

uM# total)" 6" umol/bottle 4"

0" 0" 100" 200" 300" 400" 500" 600" 700" Days# Figure 6.5: Nitrate-reducing benzene culture blended with methanogenic toluene- degrading culture. Amended with nitrate on Day 325. Replicate B.

16"

- Benzene NO3 Benzene 14"

12" Methane"(uM" total)" 10" Benzene"(uM" total)"

8" uM#

6" umol/bottle

0" 0" 100" 200" 300" 400" 500" 600" 700" Days# Figure 6.6: Nitrate-reducing benzene culture blended with methanogenic toluene- degrading culture. Amended with nitrate on Day 325. Replicate C.

14" Benzene Toluene

Methane"(uM"total)" 12"

10" Benzene"(uM"total)" 8" uM# 6"

umol/bottle Toluene"(uM"total)"

0" 0" 100" 200" 300" 400" 500" 600" 700" Days#

Figure 6.7: Methanogenic toluene-degrading culture only. Amended with toluene on Day 242. Replicate A.

Benzene Toluene 30"

25" Methane"(uM" total)"

20"

15" uM# Benzene"(uM"total)"

umol/bottle 10"

Toluene"(uM"total)" 0" 0" 100" 200" 300" 400" 500" 600" 700" Days# Figure 6.8: Methanogenic toluene-degrading culture only. Amended with toluene on Day 242. Replicate B.

16"

Toluene 14" Methane"(uM" total)" 12"

10"

8" uM# Benzene"(uM"

umol/bottle 6" total)"

Toluene"(uM" 0" total)" 0" 100" 200" 300" 400" 500" 600" 700" Days# Figure 6.9: Methanogenic toluene-degrading culture only. Amended with toluene on Day 242. Replicate C.

64 Benzene 12"

10" Methane"(uM" total)"

Benzene"(uM" 6" total)" uM# umol/bottle 4"

0" 0" 100" 200" 300" 400" 500" 600" 700" Days#

Figure 6.10: Autoclaved control. Replicate A. 20" Benzene

18"

16"

14"

12"

10" uM#

umol/bottle 8"

0" 0" 100" 200" 300" 400" 500" 600" 700" Days#

Figure 6.11: Autoclaved control. Replicate B.

Benzene 14"

12"

10"

8" uM# 6" umol/bottle

0" 0" 100" 200" 300" 400" 500" 600" 700" Days# Figure 6.12: Autoclaved control. Replicate C.

“Benzene culture-only” treatments (without nitrate, Figures 6.1 - 6.3) each showed hints at possible benzene degradation, but only Replicate C eventually revealed reproducible benzene degradation and associated methane production. Replicates B and C were both re-fed at day ~250 as a faulty GC reading showed no benzene present in the bottles. Replicate A (Figure 6.1) showed some evidence of potential benzene degradation with methanogenesis as benzene concentration decreased while methane concentration increased. At the end of the monitoring period, the concentration of methane and benzene both decreased to trace amounts. This was most likely due to a gas leak from the bottle, although no damage was visible to the mininert cap or the bottle. The cap was tightened and bottle re-fed benzene to check for remaining benzene degradation potential. Benzene culture-only replicate B (Figure 6.2) also showed evidence of potential benzene degradation. Benzene culture-only replicate C (Figure 6.3) degraded three feedings of benzene completely with methanogenesis, and is continuing to degrade benzene at a rate of 0.14 uM/Day. The ratio of methane produced: benzene consumed over three degradation cycles was 2.1.

No evidence of methanogenic benzene degradation was present in any of the “nitrate- reducing benzene culture blended with toluene culture” replicates (Figures 6.4 – 6.6). To verify the viability and benzene degradation capacity of the nitrate-reducing benzene-degrading consortium in the blend, nitrate was added on Day 325. Benzene degradation and nitrate reduction followed with no lag. When nitrate was fully depleted, benzene degradation ceased. This indicated that the initial benzene degrader (likely Peptococcaceae) was still viable in these cultures, and the lack of methanogenic benzene degradation must be attributed to other causes.

“Toluene culture only” replicates did not degrade benzene (Figures 6.7 – 6.9). Toluene was added on Day 242 to test the viability of the methanogenic partners in these replicates. Toluene was degraded rapidly with methanogenesis, indicating that methanogens present were viable.

The autoclaved controls (Figures 6.10 – 6.12) have not degraded benzene, but (A) and (B) replicates began to produce methane, indicating that they were contaminated at some stage. Methane was generated from the degradation of dead biomass in the culture. Contamination is possible from the gas-tight needles used to remove headspace samples for GC. The long incubation period that this experiment was conducted over increases the chance of contamination occurring during repeated sampling, and allows a very small amount of contamination time to grow to a detectable abundance.

For all bottles, cell pellets were taken at Day 303 and Day 393 and stored frozen at -20C. These time points were chosen to capture the composition of the cultures before nitrate was added and after benzene degradation under nitrate reducing conditions concluded. Cultures are active and still being maintained. DNA from different treatments at specific time points has been submitted for pyrotag sequencing. Points chosen were: 1) Toluene culture only Replicate B Day 303, 2) Benzene culture blended with toluene culture replicate B Day 393, and 3) Benzene culture only Replicate C Day 671. Parent Toluene culture DNA was also submitted. Results of this sequencing will be returned in January 2015. These samples will enable comparisons of the different treatments. The impact of the treatments on community diversity can be determined.

6.3 Discussion

Nitrate-reducing, benzene-degrading consortia can be switched from nitrate as electron acceptor to degradation with methanogenesis, after a lag period of approximately 450 days. This is further evidence of syntrophy in benzene-degrading enrichment cultures. A single organism is extremely unlikely to be capable of both nitrate reduction and methanogenesis, showing that a different organism must have taken over the role of consuming fermentation products from benzene degradation during the course of this experiment. This is not possible if the entire benzene degradation pathway were carried out in a single organism.

This experiment also provided evidence against the hypothesis that intracellular disproportionation of nitrate to nitrogen and oxygen is carried out in nitrate-reducing benzene- degrading cultures and the oxygen generated is subsequently used to activate the benzene ring. Benzene degradation continued in the absence of nitrate, showing that nitrate, and therefore oxygen, is not required for the initial attack on benzene. A different mechanism must be used (such as carboxylation).

The long lag phase experienced in this experiment may be due to the slow growth rate of the methanogens present in the consortium. Benzene degradation with nitrate reduction was taking place at the beginning of the cultivation period, showing that initial benzene degrader (likely Peptococcaceae) abundance was sufficient for benzene degradation. Methanogen abundance was very low (<0.5%) however, and the period required for methanogen growth to a sufficient level to consume the products of benzene degradation may have caused the long lag period before the benzene degradation rate became detectable.

According to the stoichiometric equation for methanogenic benzene degradation from (Ulrich & Edwards, 2003) (equation (1)), theoretically, 3.6 moles of methane are produced for every mole of benzene consumed. The ratio of methane produced: benzene consumed over three degradation cycles was 2.1. This is relatively close to the theoretical value. Methane consumption by other organisms may be taking place, or slight leakage when bottles become over pressurized.

+ - + (1) C6H6 + 6.55 H2O + 0.05 NH4 → 2.15 HCO3 + 3.6 CH4 + 0.05 C5H7O2N + 2.4H

Changes of the electron acceptor for benzene degradation from nitrate to alternate electron acceptors have been reported previously (nitrate to chlorate, oxygen, sulfate and ferrous iron). Previous work on cultures related to the Cart T3 culture used in this experiment (nitrate- reducing benzene degrading enrichment cultures from Cartwright Gas Station microcosms) did not show electron acceptor variability (Ulrich and Edwards, 2003). This may be due to more highly enriched cultures being used in the previous study, compared to Cart T3. The parent culture studied in that experiment was an 8th generation transfer, and was sediment – free (Ulrich and Edwards, 2003). The more highly enriched cultures may have lost the methanogens needed for methanogenesis from benzene, or the long lag phase before methanogenesis occurs may have prevented its detection.

The lack of methanogenic benzene degradation in the “benzene plus toluene culture” replicates is curious. The organism responsible for the initial attack on benzene is viable in these replicates, as is shown by benzene degradation in the “benzene plus toluene culture” replicates after the addition of nitrate. It is not inhibited by anything introduced with the toluene culture. The lack of methanogenic benzene degradation could therefore be due to either a lack of activity of the methanogens in these replicates, or an incompatibility between the benzene-attacking organism and the methanogens. An intermediary organism may also be missing in this culture.

The methanogens in the “toluene culture only” replicates are viable, as shown by the toluene-degrading activity of these replicates. Methanogens are not inhibited by benzene, as they produced methane from toluene in the “toluene culture only” replicates while benzene remained present in the bottles. Methanogens in the “benzene culture only” replicates are also active. Together, these points suggest that the methanogens in the “benzene plus toluene culture” replicates ought to be viable. There could be something about the methanogens from the toluene- degrading methanogenic culture that makes them unsuited to partner benzene degradation by the initial benzene degrader in these cultures (likely Peptococcaceae). They could be affected by a metabolite present in the culture, or prefer a different energy source than the metabolites produced by benzene degradation in these cultures. Methanogens introduced with the nitrate-

reducing benzene-degrading consortia may still be present and undergoing growth to a sufficient abundance for methanogenesis to be detectable in the “benzene plus toluene culture” replicates. The longer lag time in these replicates can be accounted for, if that is the case, by the 50% dilution of nitrate-reducing benzene-degrading consortia into toluene-degrading methanogenic consortia in these replicates, compared to the 100% nitrate-reducing benzene-degrading consortia used in the “benzene culture only” replicates.

6.4 Key points

• Benzene degrading cultures can switch terminal electron acceptor from nitrate to carbon dioxide. This is further evidence that benzene degradation is carried out through a syntrophic process in these enrichment cultures.

• Evidence against intracellular disproportionation of nitrate to nitrogen and oxygen and the use of the oxygen generated to initiate the degradation of benzene was established.

• An extended lag period of 450 days was observed before methanogenesis and benzene degradation occur. This was likely due to the time required for slow-growing methanogens to increase in abundance.

• It is possible that the initial benzene degrader (likely Peptococcaceae) preferentially carries out benzene degradation in syntrophy with indigenous methanogens, not with methanogens from the toluene-degrading methanogenic culture.

• Future work includes pyrotag sequencing and analysis to compare the community composition between the replicates that switched electron donor successfully, parent cultures, replicates that did not show electron acceptor variability, and a “toluene- degrading culture only” replicate.

Chapter 7 Conclusions and Engineering significance 7

This work provided a more detailed examination of the community involved in anaerobic benzene-degrading nitrate-reducing enrichment cultures. It provided more evidence for carboxylation as the activation mechanism for benzene degradation. An enrichment was established from the nitrate-reducing culture which repeatedly degraded benzene with CO2 as electron acceptor, rather than nitrate, confirming the presence of a syntrophic association between fermenting and nitrate-reducing or methanogenic organisms. The metagenome assembly presented here can be further used to examine the metabolic potential of the microbiome and attribute functions to community members. 7.1 A highly diverse community with functional redundancy is present

There are many different phylotypes present in the various enrichments cultures. Organisms common to all nitrate-reducing benzene-degrading enrichments include Peptococcaceae, Azoarcus, Burkholderiaceae, and Ignavibacteriaceae. Peptococcaceae is the organism thought to be responsible for the initial attack on the benzene ring. Azoarcus and Burkholderiaceae are both nitrate reducers that may degrade intermediates of anaerobic benzene degradation. Many other groups were present at similar abundances to these key organisms. These supplementary groups may take part in nutrient cycling, or scavenging dead biomass. Methanogens are present in the nitrate-reducing enrichments at very low abundances, and may account for the terminal fate of benzene-derived electrons when nitrate is depleted in the enrichments. Both pyrotag sequencing data and the assembled metagenome support this picture of enrichments with high diversity and multiple organisms being present that can carry out similar functions. Two strains of Peptococcaceae are present in the enrichments. The abundances of OTUs common to all cultures varied between cultures. Abundances of groups proposed to take part in the benzene degradation

pathway did not correlate significantly with benzene degradation rate in this study. Presence, absence, and abundances of specific OTUs do not have a significant effect on benzene degradation rate. Overall community composition and richness is more important to benzene degradation than any individual organism.

7.2 The putative anaerobic benzene carboxylase “Abc” is responsible for the initiation of benzene degradation

The only gene found encoding an enzyme in any of the proposed pathways for anaerobic conversion of benzene → benzoate was the abc operon. This supports evidence from a prior transcriptomic study on this enrichment (Luo et al., 2014). The enzymes responsible for other key functions in the community can be found within the assembled metagenome, and can be ascribed to host organisms.

7.3 Nitrate-reducing benzene degrading enrichments can utilize alternative electron acceptors

No nitrate-reducing, benzene-degrading enrichments have previously been successfully switched to methanogenesis. This is likely due to the extended lag period before methanogenesis occurs (~450 days). The long lag period can be attributed to the requirement for extremely low abundance methanogens to increase in abundance. The lack of methanogenic benzene degradation to date in nitrate-reducing, benzene-degrading enrichments supplemented with methanogen-rich, toluene-degrading culture suggests that there may be something distinctive about the methanogens present in the nitrate-reducing culture that enables them to operate as syntrophic partners of the initial benzene degrader (likely Peptococcaceae), whereas methanogens from the toluene-degrading culture cannot. The assumption is made that Peptococcaceae are still the dominant benzene degrading organisms in these cultures as the organism proposed to be responsible for methanogenic benzene degradation, ORM2, was not

present in the nitrate-reducing, benzene-degrading enrichment used as inoculum. Pyrotag sequencing or qPCR can be used to test this assumption. The community present in this enrichment is of interest because methanogenic and nitrate-reducing benzene degrading enrichments typically have very dissimilar populations. The community may now be more similar to methanogenic enrichment communities, or it may have retained similarity to its parent nitrate-reducing enrichment. This experiment provided more evidence for syntrophy in benzene- degrading enrichments, and evidence against nitrate disproportionation to nitrogen and oxygen as a step in benzene degradation in nitrate-reducing enrichment cultures.

7.4 Future directions suggested

The metagenomic assembly described here can be enhanced by further work. Methods presented in (Tang et al., 2012) can be used to close some of the gaps in scaffolds, and link different scaffolds together. The assembly can also be further mined to locate all the genes involved in the benzene degradation pathway, and investigate the different organisms that may be carrying out different stages of the pathway. The metagenome can be compared to the metatranscriptome generated in (Luo et al., 2014). Individual genomes can also be compared to genomes of closely related organisms adapted to alternative substrates to see what adaptations are present only after growth on benzene. Once the full genomic potential of Peptococcaceae is understood, this information could be used to attempt to create culture conditions where Peptococcaceae could be isolated. An isolate of benzene-degrading Peptococcaceae from a nitrate-reducing culture would be valuable for detailed biochemical characterization of its unique metabolism. This characterization would help to figure out the handoff point between the benzene fermenter and other microbes in the community. Roles of other organisms in the microbiome can also be elucidated though isolation or inference.

7.5 Engineering significance and contributions of this research

This research has several applications. An understanding of the biological anaerobic degradation pathway of benzene in enrichment cultures can be used to generate hypotheses about the fate of benzene in the environment. This is an important issue, as benzene is a common environmental contaminant and current knowledge states that benzene persists in anaerobic environments for a long period of time. If the biodegradation of benzene in anaerobic environments can be proven to commonly take place, this can be used to better focus remediation efforts on scenarios where no natural attenuation is occurring. Biodegradation of benzene could be estimated by detecting biomarkers of benzene degradation. Biomarkers are genes or transcripts that are directly linked to the process of interest. One potential biomarker for anaerobic benzene would be the abc operon. This marker is specific to degradation through carboxylation, so a suite of biomarkers that are specific to different degradation mechanisms may be necessary. This research also opens the path towards generating inocula that could be used to bioaugment a site to enhance the rate of anaerobic benzene degradation in contaminated environments, much like is already done for chlorinated solvents with Dehalococcoides-containing enrichment cultures.

Another product of this research is a metagenome assembly that can be mined for functional pathways present in the enrichment. Novel enzymes may be found in the metagenome, which could have industrial applications as they may be capable of catalyzing industrially relevant reactions. Benzene is a molecule of interest to industry, and the reactions involved in its degradation are unusual, so this is likely. This research also generated a methanogenic benzene degrading enrichment from a nitrate-reducing enrichment that can be used in future research.

References

Abu Laban, N., Selesi, D., Jobelius, C., & Meckenstock, R. U. (2009). Anaerobic benzene degradation by Gram-positive sulfate-reducing bacteria. FEMS Microbiology Ecology, 68(3), 300–11. doi:10.1111/j.1574-6941.2009.00672.x

Abu Laban, N., Selesi, D., Rattei, T., Tischler, P., & Meckenstock, R. U. (2010). Identification of enzymes involved in anaerobic benzene degradation by a strictly anaerobic iron-reducing enrichment culture. Environmental Microbiology, 12(10), 2783–96. doi:10.1111/j.1462- 2920.2010.02248.x

Achenbach, L. A., Michaelidou, U., Bruce, R. A., Fryman, J., & Coates, J. D. (2001). Dechlorosoma suillum gen. nov., sp. nov., two novel environmentally dominant (per)chlorate- reducing bacteria and their phylogenetic position. International Journal of Systematic and Evolutionary Microbiology, 51, 527–533.

Albuquerque, L., Simões, C., Nobre, M. F., Pino, N. M., Battista, J. R., Silva, M. T., … da Costa, M. S. (2005). Truepera radiovictrix gen. nov., sp. nov., a new radiation resistant species and the proposal of Trueperaceae fam. nov. FEMS Microbiology Letters, 247(2), 161–9. doi:10.1016/j.femsle.2005.05.002

Amann, R. I., Ludwig, W., & Schleifer, K. H. (1995). Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiological Reviews, 59(1), 143–69.

Aziz, R. K., Bartels, D., Best, A. A, DeJongh, M., Disz, T., Edwards, R. A, … Zagnitko, O. (2008). The RAST Server: rapid annotations using subsystems technology. BMC Genomics, 9, 75. doi:10.1186/1471-2164-9-75

Beller, H. R., Letain, T. E., Chakicherla, A., Kane, S. R., Legler, T. C., & Coleman, M. A. (2006). Whole-genome transcriptional analysis of chemolithoautotrophic thiosulfate

oxidation by Thiobacillus denitrificans under aerobic versus denitrifying conditions. Journal of Bacteriology, 188(19), 7005–15. doi:10.1128/JB.00568-06

Bento, F. M., Camargo, F. O., Okeke, B. C., & Frankenberger, W. T. (2005). Comparative bioremediation of soils contaminated with diesel oil by natural attenuation, biostimulation and bioaugmentation. Bioresource Technology, 96(9), 1049–55. doi:10.1016/j.biortech.2004.09.008

Bolger, A. M., Lohse, M., & Usadel, B. (2014). Trimmomatic : A flexible trimmer for Illumina Sequence Data. Bioinformatics, 1–7.

Breinig, S., Schiltz, E., & Fuchs, G. (2000). Genes Involved in Anaerobic Metabolism of Phenol in the Bacterium Thauera aromatica. Journal of Bacteriology, 182(20), 5849–5863. doi:10.1128/JB.182.20.5849-5863.2000

Brown, C., Howe, A., Zhang, Q., Pyrkosz, A., & Brom, T. (2012). A Reference-Free Algorithm for Computational Normalization of Shotgun Sequencing Data. arXiv, 1203.408. Retrieved from http://arxiv.org/abs/1203.4802

Bruns, A., Rohde, M., & Berthe-Corti, L. (2001). Muricauda ruestringensis gen. nov., sp. nov., a facultatively anaerobic, appendaged bacterium from German North Sea intertidal sediment. International Journal of Systematic and Evolutionary Microbiology, 51(6), 1997–2006. doi:10.1099/00207713-51-6-199A

Burland, S. M., & Edwards, E. A. (1999). Anaerobic benzene biodegradation linked to nitrate reduction. Applied and Environmental Microbiology, 65(2), 529–533.

Byrne-Bailey, K. G., Wrighton, K. C., Melnyk, R. A, Agbo, P., Hazen, T. C., & Coates, J. D. (2010). Complete genome sequence of the electricity-producing “Thermincola potens” strain JR. Journal of Bacteriology, 192(15), 4078–9. doi:10.1128/JB.00044-10

Caldwell, M. E., & Suflita, J. M. (2000). Detection of Phenol and Benzoate as Intermediates of Anaerobic Benzene Biodegradation under Different Terminal Electron-Accepting

Conditions. Environmental Science & Technology, 34(7), 1216–1220. doi:10.1021/es990849j

Caporaso, J. G., Kuczynski, J., Stombaugh, J., Bittinger, K., Bushman, F. D., Costello, E. K., … Knight, R. (2010). QIIME allows analysis of high- throughput community sequencing data Intensity normalization improves color calling in SOLiD sequencing. Nature Publishing Group, 7(5), 335–336. doi:10.1038/nmeth0510-335

Carmona, M., Zamarro, M. T., Blázquez, B., Durante-Rodríguez, G., Juárez, J. F., Valderrama, J. A., … Díaz, E. (2009). Anaerobic catabolism of aromatic compounds: a genetic and genomic view. Microbiology and Molecular Biology Reviews : MMBR, 73(1), 71–133. doi:10.1128/MMBR.00021-08

Chain, P., Lamerdin, J., Larimer, F., Regala, W., Lao, V., Land, M., … Arp, D. (2003). Complete Genome Sequence of the Ammonia-Oxidizing Bacterium and Obligate Chemolithoautotroph Nitrosomonas europaea. Journal of Bacteriology, 185(9), 2759–2773. doi:10.1128/JB.185.9.2759-2773.2003

Chakraborty, R., & Coates, J. D. (2005). Hydroxylation and Carboxylation — Two Crucial Steps of Anaerobic Benzene Degradation by Dechloromonas Strain RCB. Applied and Environmental Microbiology, 71(9), 5427–5432. doi:10.1128/AEM.71.9.5427

Chin, K. J., Liesack, W., & Janssen, P. H. (2001). Opitutus terrae gen. nov., sp. nov., to accommodate novel strains of the division “Verrucomicrobia” isolated from rice paddy soil. International Journal of Systematic and Evolutionary Microbiology, 51(6), 1965–1968. doi:10.1099/00207713-51-6-1965

Clark, D. A., & Norris, P. R. (1996). Acidimicrobium ferrooxidans gen. nov., sp. nov.: mixed- culture ferrous iron oxidation with Sulfobacillus species. Microbiology, 142, 785–790.

Clinton, M. (1948). Benzene. API Toxicological Review, 1–6. American Petroleum Institute. Department of Safety, New York.

Coates, J. D., Chakraborty, R., Lack, J. G., Connor, S. M. O., Cole, K. A., Bender, K. S., & Achenbach, L. A. (2001). Anaerobic benzene oxidation coupled to nitrate reduction in pure culture by two strains of Dechloromonas. Letters to Nature, 411(June), 1039–1043.

Darling, A. C. E., Mau, B., Blattner, F. R., & Perna, N. T. (2004). Mauve : Multiple Alignment of Conserved Genomic Sequence With Rearrangements. Genome Research, 14, 1394–1403. doi:10.1101/gr.2289704.tion

Devereux, R., He, S., Doyle, C. L., Orkland, S., Stahl, D. A., LeGall, J., & Whitman, W. B. (1990). Diversity and Origin of Desulfovibrio Species : Phylogenetic Definition of a Family. Journal of Bacteriology, 172(7), 3609 – 3619.

Devine, C. E. (2013). Identification of Key Organisms, Genes and Pathways in Benzene- Degrading Methanogenic Cultures. University of Toronto, Toronto.

Ding, L., & Yokota, A. (2004). Proposals of Curvibacter gracilis gen. nov., sp. nov. and Herbaspirillum putei sp. nov. for bacterial strains isolated from well water and reclassification of [Pseudomonas] huttiensis, [Pseudomonas] lanceolata, [Aquaspirillum] delicatum and [Aquaspirillum. International Journal of Systematic and Evolutionary Microbiology, 54(Pt 6), 2223–30. doi:10.1099/ijs.0.02975-0

Eckardt, R. E. (1973) Recent development in industrial carcinogens. Journal of Occupational Medicine, 15, 904–907. Retrieved from http://www.scopus.com/inward/record.url?eid=2- s2.0-0015698199&partnerID=tZOtx3y1

Edgar, R. C. (2004). MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research, 32(5), 1792–7. doi:10.1093/nar/gkh340

Emerson, D., Field, E. K., Chertkov, O., Davenport, K. W., Goodwin, L., Munk, C., … Woyke, T. (2013). Comparative genomics of freshwater Fe-oxidizing bacteria: implications for physiology, ecology, and systematics. Frontiers in Microbiology, 4(September), 254. doi:10.3389/fmicb.2013.00254

Gibson, J., & Harwood, C. S. (2002). Metabolic diversity in aromatic compound utilization by anaerobic microbes. Annual Review of Microbiology, 56, 345–69. doi:10.1146/annurev.micro.56.012302.160749

Gitiafroz, R. (2012). Benzene biodegradation under nitrate-reducing conditions. University of Toronto, Toronto.

Gitiafroz, R., Devine, C. E., Raskin, L., & Edwards, E. A. (n.d.). Peptococcaceae intiates benzene mineralization in a nitrate-reducing enrichment culture.

Gnerre, S., Maccallum, I., Przybylski, D., Ribeiro, F. J., Burton, J. N., Walker, B. J., … Jaffe, D. B. (2011). High-quality draft assemblies of mammalian genomes from massively parallel sequence data. Proceedings of the National Academy of Sciences of the United States of America, 108(4), 1513–8. doi:10.1073/pnas.1017351108

Gorlenko, V. M. (1970). A new phototrophic green sulfur bacterium - Prosthecochloris aestuarii nov. gen. nov. spec. Zeitschrift Fur Allg. Mikrobiologie, 10(2), 147 – 149.

Government of Canada. (1997). Benzene in Gasoline Regulations Règlement sur le benzène dans l ’ essence. Retrieved from http://laws-lois.justice.gc.ca/PDF/SOR-97-493.pdf

Government of Canada. (2013). Groundwater. Retrieved from https://www.ec.gc.ca/eau- water/default.asp?lang=En&n=300688DC-1#sub5

Grégoire, P., Fardeau, M.-L., Joseph, M., Guasco, S., Hamaide, F., Biasutti, S., … Ollivier, B. (2011). Isolation and characterization of Thermanaerothrix daxensis gen. nov., sp. nov., a thermophilic anaerobic bacterium pertaining to the phylum “Chloroflexi”, isolated from a deep hot aquifer in the Aquitaine Basin. Systematic and Applied Microbiology, 34(7), 494– 7. doi:10.1016/j.syapm.2011.02.004

Guindon, S., Dufayard, J.-F., Lefort, V., Anisimova, M., Hordijk, W., & Gascuel, O. (2010). New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Systematic Biology, 59(3), 307–21. doi:10.1093/sysbio/syq010

Gumaelius, L., Magnusson, G., Pettersson, B., & Dalhammar, G. (2001). Comamonas denitrificans sp. nov., an efficient denitrifying bacterium isolated from activated sludge. International Journal of Systematic and Evolutionary Microbiology, 51(3), 999–1006. doi:10.1099/00207713-51-3-999

Health Canada. (2009). Guidelines for Canadian Drinking Guideline Technical Document. Retrieved from http://www.hc-sc.gc.ca/ewh-semt/alt_formats/hecs-sesc/pdf/pubs/water- eau/benzene/benzene-eng.pdf

Heider, J., Spormann, A. M., Beller, H. R., & Widdel, F. (1999). Anaerobic bacterial metabolism of hydrocarbons. FEMS Microbiology Reveiws, 22(22), 459 – 473.

Holmes, D. E., Nevin, K. P., Woodard, T. L., Peacock, A. D., & Lovley, D. R. (2007). Prolixibacter bellariivorans gen. nov., sp. nov., a sugar-fermenting, psychrotolerant anaerobe of the phylum Bacteroidetes, isolated from a marine-sediment fuel cell. International Journal of Systematic and Evolutionary Microbiology, 57(Pt 4), 701–7. doi:10.1099/ijs.0.64296-0

Holmes, D. E., Risso, C., Smith, J. a, & Lovley, D. R. (2011). Anaerobic oxidation of benzene by the hyperthermophilic archaeon Ferroglobus placidus. Applied and Environmental Microbiology, 77(17), 5926–33. doi:10.1128/AEM.05452-11

Hug, L. A., Maphosa, F., Leys, D., Lo, F. E., Smidt, H., Edwards, E. A., & Adrian, L. (2013). Overview of organohalide-respiring bacteria and a proposal for a classification system for reductive dehalogenases. Philosophical Transactions of the Royal Society B, 368(20120322).

Iino, T., Mori, K., Uchino, Y., Nakagawa, T., Harayama, S., & Suzuki, K.-I. (2010). Ignavibacterium album gen. nov., sp. nov., a moderately thermophilic anaerobic bacterium isolated from microbial mats at a terrestrial hot spring and proposal of Ignavibacteria classis nov., for a novel lineage at the periphery of green sulfur bacteria. International Journal of Systematic and Evolutionary Microbiology, 60(Pt 6), 1376–82. doi:10.1099/ijs.0.012484-0

Imachi, H., Sekiguchi, Y., Kamagata, Y., Hanada, S., Ohashi, A., & Harada, H. (2002). Pelotomaculum thermopropionicum gen. nov., sp. nov., an anaerobic, thermophilic, syntrophic propionate-oxidizing bacterium. International Journal of Systematic and Evolutionary Microbiology, 52(Pt 5), 1729–35. doi:10.1099/ijs.0.02212-0.Abbreviations

Itoh, T., Yamanoi, K., Kudo, T., Ohkuma, M., & Takashina, T. (2011). Aciditerrimonas ferrireducens gen. nov., sp. nov., an iron-reducing thermoacidophilic actinobacterium isolated from a solfataric field. International Journal of Systematic and Evolutionary Microbiology, 61(Pt 6), 1281–5. doi:10.1099/ijs.0.023044-0

Jindrova, E., Chocova, M., Demnerova, K., & Brenner, V. (2002). Bacterial Aerobic Degradation of Benzene, Toluene, Ethylbenzene and Xylene. Folia Microbiologica, 47(2), 83 – 93.

Kadnikov, V. V, Mardanov, A. V, Podosokorskaya, O. a, Gavrilov, S. N., Kublanov, I. V, Beletsky, A. V, … Ravin, N. V. (2013). Genomic analysis of Melioribacter roseus, facultatively anaerobic organotrophic bacterium representing a novel deep lineage within Bacteriodetes/Chlorobi group. PloS One, 8(1), e53047. doi:10.1371/journal.pone.0053047

Kalyuzhnaya, M. G., De Marco, P., Bowerman, S., Pacheco, C. C., Lara, J. C., Lidstrom, M. E., & Chistoserdova, L. (2006). Methyloversatilis universalis gen. nov., sp. nov., a novel taxon within the Betaproteobacteria represented by three methylotrophic isolates. International Journal of Systematic and Evolutionary Microbiology, 56(Pt 11), 2517–22. doi:10.1099/ijs.0.64422-0

Kaneko, T., Nakamura, Y., Sato, S., Minamisawa, K., Uchiumi, T., Sasamoto, S., … Wada, T. (2002). Complete Genomic Sequence of Nitrogen-fixing Symbiotic Bacterium Bradyrhizobium japonicum USDA110, 197, 189–197.

Kasai, Y., Takahata, Y., Manefield, M., & Watanabe, K. (2006). RNA-based stable isotope probing and isolation of anaerobic benzene-degrading bacteria from gasoline-contaminated

groundwater. Applied and Environmental Microbiology, 72(5), 3586–92. doi:10.1128/AEM.72.5.3586-3592.2006

Kazumi, J., Caldwell, M. E., Suflita, J. M., Lovley, D. R., & Young, L. Y. (1997). Anaerobic Degradation of Benzene in Diverse Anoxic Environments. Environmental Science & Technology, 31(3), 813–818.

Kearse, M., Moir, R., Wilson, A., Stones-Havas, S., Cheung, M., Sturrock, S., … Drummond, A. (2012). Geneious Basic: an integrated and extendable desktop software platform for the organization and analysis of sequence data. Bioinformatics, 28(12), 1647–1649.

Kei, H., Ursing, J., Blom, J., Christen, A. C., Christensen, J., Frederiksen, W., … Ying, Y. T. (1994). Lautropia mirabilis gen. nov., sp. nov., a Gram-negative motile coccus with unusual morphology isolated from the human mouth. Microbiology, 140(1787 - 1797).

Key, K. C., Sublette, K. L., Johannes, T. W., Ogles, D., Baldwin, B., & Biernacki, A. (2014). Assessing BTEX Biodegradation Potential at a Refinery Using Molecular Biological Tools. Groundwater Monitoring and Remediation, 1(1), 35–48. doi:10.1111/gwmr.12037

Kosaka, T., Kato, S., Shimoyama, T., Ishii, S., Abe, T., & Watanabe, K. (2008). The genome of Pelotomaculum thermopropionicum reveals niche-associated evolution in anaerobic microbiota. Genome Research, 18(3), 442–8. doi:10.1101/gr.7136508

Kunapuli, U., Griebler, C., Beller, H. R., & Meckenstock, R. U. (2008). Identification of intermediates formed during anaerobic benzene degradation by an iron-reducing enrichment culture. Environmental Microbiology, 10(7), 1703–12. doi:10.1111/j.1462- 2920.2008.01588.x

Lai, Q., & Shao, Z. (2012). Genome sequence of Oceanibaculum indicum type strain P24. Journal of Bacteriology, 194(24), 6942. doi:10.1128/JB.01859-12

Landon, M. K., & Belitz, K. (2012). Geogenic sources of benzene in aquifers used for public supply, California. Environmental Science & Technology, 46(16), 8689–97. doi:10.1021/es302024c

Liang, X., Devine, C. E., Nelson, J., Lollar, B. S., Zinder, S., & Edwards, E. A. (2013). Anaerobic Conversion of Chlorobenzene and Benzene to CH 4 and CO 2 in Bioaugmented Microcosms.

Liesack, W., Bak, F., Kreft, J.-U., & Stackebrandt, E. (1994). Holophaga foetida gen. nov., sp. nov., a new , homoacetogenic bacterium degrading methoxylated aromatic compounds. Arch Microbiol, 162(December 1992), 85–90.

Lingens, F., Blecher, R., Blecher, H., Blobel, F., Frohner, C., Gorisch, H., … Layh, G. (1985). Bacterium That Degrades the Herbicide Chloridazon. International Journal of Systematic and Evolutionary Microbiology, 35(1), 26–39.

Lovley, D. R., Coates, J. D., Woodward, J. C., & Phillips, E. (1995). Benzene oxidation coupled to sulfate reduction. Applied and Environmental Microbiology, 61(3), 953–958.

Lovley, D. R., Giovannoni, S. J., White, D. C., Champine, J. E., Phillips, E. J. P., Gorby, Y. A., & Goodwin, S. (1993). Geobacter metallireducens gen. nov. sp. nov., a microorganism capable of coupling the complete oxidation of organic compounds to the reduction of iron and other metals D. Archives of Microbiology, 159, 336–344.

Lovley, D. R., Woodward, J. C., & Chapelle, F. H. (1994). Stimulated anoxic biodegradation of aromatic hydrocarbons using Fe(III) ligands. Letters to Nature, 370, 128–131.

Luo, F., Gitiafroz, R., Devine, C. E., Gong, Y., Hug, L. A, Raskin, L., & Edwards, E. A. (2014). Metatranscriptome of an anaerobic benzene-degrading, nitrate-reducing enrichment culture reveals involvement of carboxylation in benzene ring activation. Applied and Environmental Microbiology, 80(14), 4095–107. doi:10.1128/AEM.00717-14

MacLeod, M., & Mackay, D. (1999). An assessment of the environmental fate and exposure of benzene and the chlorobenzenes in Canada. Chemosphere, 38(8), 1777–1796. Retrieved from http://www.scopus.com/inward/record.url?eid=2-s2.0- 0033120867&partnerID=40&md5=7fc09c4463a07db08357e782e6e00c2a

Mancini, S. A, Devine, C. E., Elsner, M., Nandi, M. E., Ulrich, A. C., Edwards, E. A, & Lollar, B. S. (2008). Isotopic evidence suggests different initial reaction mechanisms for anaerobic benzene biodegradation. Environmental Science & Technology, 42(22), 8290–6. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/19068808

Moran, N. A., Hansen, A. K., Powell, J. E., and Sabree, Z. L.(2012) Distinctive Gut Microbiota of Honey Bees Assessed Using Deep Sampling from Individual Worker. PLoS One, 7(4): e36393

Mouttaki, H., Johannes, J., & Meckenstock, R. U. (2012). Identification of naphthalene carboxylase as a prototype for the anaerobic activation of non-substituted aromatic hydrocarbons. Environmental Microbiology, 14(10), 2770–4. doi:10.1111/j.1462- 2920.2012.02768.x

Nales, M., Butler, B. J., & Edwards, E. A. (1998). Anaerobic Benzene Biodegradation: A Microcosm Survey. Bioremediation Journal, 2(2), 125–144. doi:10.1080/10889869891214268

Nandi, M. E. (2006). Biochemical and Molecular Characterization of Anaerobic Benzene- Degrading Nitrate-Reducing Cultures. University of Toronto, Toronto.

Patel, S. V, & Tolley, W. K. (2014). Developments toward a low-cost approach for long-term, unattended vapor intrusion monitoring. Analyst, 139(15), 3770–80. doi:10.1039/c4an00736k

Phelps, C. D., Kerkhof, L. J., & Young, L. Y. (1998). Molecular characterization of a sulfate- reducing consortium which mineralizes benzene. FEMS Microbiology Ecology, 27(3), 269– 279. doi:10.1111/j.1574-6941.1998.tb00543.x

Phelps, C. D., Zhang, X., & Young, L. Y. (2001). Brief report: Use of stable isotopes to identify benzoate as a metabolite of benzene degradation in a sulphidogenic consortium, Environ Microbiol, 3, 600–603.

Podosokorskaya, O. a, Kadnikov, V. V, Gavrilov, S. N., Mardanov, A. V, Merkel, A. Y., Karnachuk, O. V, … Kublanov, I. V. (2013). Characterization of Melioribacter roseus gen. nov., sp. nov., a novel facultatively anaerobic thermophilic cellulolytic bacterium from the class Ignavibacteria, and a proposal of a novel bacterial phylum Ignavibacteriae. Environmental Microbiology, 15(6), 1759–71. doi:10.1111/1462-2920.12067

Ponsin, V., Coulomb, B., Guelorget, Y., Maier, J., & Höhener, P. (2014). In situ biostimulation of petroleum hydrocarbon degradation by nitrate and phosphate injection using a dipole well configuration. Journal of Contaminant Hydrology, 171, 22–31. doi:10.1016/j.jconhyd.2014.10.003

R Core Team. (2014). R: A Language and Environment for Statistical Computing. Vienna, Austria: R Foundation for Statistical Computing.

Rabus, R., Kube, M., Heider, J., Beck, A., Heitmann, K., Widdel, F., & Reinhardt, R. (2005). The genome sequence of an anaerobic aromatic-degrading denitrifying bacterium, strain EbN1. Archives of Microbiology, 183(1), 27–36. doi:10.1007/s00203-004-0742-9

Ramos, D. T., da Silva, M. L. B., Chiaranda, H. S., Alvarez, P. J. J., & Corseuil, H. X. (2013). Biostimulation of anaerobic BTEX biodegradation under fermentative methanogenic conditions at source-zone groundwater contaminated with a biodiesel blend (B20). Biodegradation, 24(3), 333–41. doi:10.1007/s10532-012-9589-y

Reichenbach, H., Lang, E., Schumann, P., & Spröer, C. (2006). Byssovorax cruenta gen. nov., sp. nov., nom. rev., a cellulose-degrading myxobacterium: rediscovery of “Myxococcus cruentus” Thaxter 1897. International Journal of Systematic and Evolutionary Microbiology, 56(Pt 10), 2357–63. doi:10.1099/ijs.0.63628-0

Rooney-Varga, J. N., Anderson, R. T., Fraga, J. L., & Lovley, D. R. (1999). Microbial Communities Associated with Anaerobic Benzene Degradation in a Petroleum- Contaminated Aquifer. Applied and Environmental Microbiology, 65(7), 3056–3063.

Salanitro, J. P., Wisniewski, H. L., Byers, D. L., Neaville, C. C., & Schroder, R. A. (1997). Use of Aerobic and Anaerobic Microcosms to Assess BTEX Biodegradation in Aquifers. Groundwater Monitoring and Remediation, 210–221.

Salinero, K. K., Keller, K., Feil, W. S., Feil, H., Trong, S., Di Bartolo, G., & Lapidus, A. (2009). Metabolic analysis of the soil microbe Dechloromonas aromatica str. RCB: indications of a surprisingly complex life-style and cryptic anaerobic pathways for aromatic degradation. BMC Genomics, 10, 351. doi:10.1186/1471-2164-10-351

Sanford, R. A., Cole, J. R., & Tiedje, J. M. (2002). Characterization and Description of Anaeromyxobacter dehalogenans gen. nov., Anaerobic Myxobacterium Characterization and Description of Anaeromyxobacter dehalogenans gen. nov., sp . nov., an Aryl- Halorespiring Facultative Anaerobic Myxobacterium. Applied and Environmental Microbiology, 68(2), 893 – 900. doi:10.1128/AEM.68.2.893

Sangkhobol, V., & Skerman, V. B. D. (1981). Chitinophaga , a New Genus of Chitinolytic Myxobacteria. International Journal of Systematic and Evolutionary Microbiology, 31(3), 285–293.

Schleheck, D., Tindall, B. J., Rosselló-Mora, R., & Cook, A. M. (2004). Parvibaculum lavamentivorans gen. nov., sp. nov., a novel heterotroph that initiates catabolism of linear alkylbenzenesulfonate. International Journal of Systematic and Evolutionary Microbiology, 54(Pt 5), 1489–97. doi:10.1099/ijs.0.03020-0

Schwarzenbach, R. P., Escher, B. I., Fenner, K., Hofstetter, T. B., Johnson, C. A., von Gunten, U., & Wehrli, B. (2006). The challenge of micropollutants in aquatic systems. Science (New York, N.Y.), 313(5790), 1072–7. doi:10.1126/science.1127291

Sekiguchi, Y. (2003). Anaerolinea thermophila gen. nov., sp. nov. and Caldilinea aerophila gen. nov., sp. nov., novel filamentous thermophiles that represent a previously uncultured lineage of the domain Bacteria at the subphylum level. International Journal of Systematic and Evolutionary Microbiology, 53(6), 1843–1851. doi:10.1099/ijs.0.02699-0

Sieber, J. R., McInerney, M. J., & Gunsalus, R. P. (2012). Genomic insights into syntrophy: the paradigm for anaerobic metabolic cooperation. Annual Review of Microbiology, 66, 429–52. doi:10.1146/annurev-micro-090110-102844

Simpson, J. T., Wong, K., Jackman, S. D., Schein, J. E., Jones, S. J. M., & Birol, I. (2009). ABySS: a parallel assembler for short read sequence data. Genome Research, 19(6), 1117– 23. doi:10.1101/gr.089532.108

Smith, R. L., Repert, D. A., Barber, L. B., & LeBlanc, D. R. (2013). Long-term groundwater contamination after source removal—The role of sorbed carbon and nitrogen on the rate of reoxygenation of a treated-wastewater plume on Cape Cod, MA, USA. Chemical Geology, 337-338, 38–47. doi:10.1016/j.chemgeo.2012.11.007

Sogin, M. L., Morrison, H. G., Huber, J. A, Mark Welch, D., Huse, S. M., Neal, P. R., … Herndl, G. J. (2006). Microbial diversity in the deep sea and the underexplored “rare biosphere”. Proceedings of the National Academy of Sciences of the United States of America, 103(32), 12115–20. doi:10.1073/pnas.0605127103

Sokolova, T. G., Kostrikina, N. A, Chernyh, N. A, Kolganova, T. V, Tourova, T. P., & Bonch- Osmolovskaya, E. a. (2005). Thermincola carboxydiphila gen. nov., sp. nov., a novel anaerobic, carboxydotrophic, hydrogenogenic bacterium from a hot spring of the Lake Baikal area. International Journal of Systematic and Evolutionary Microbiology, 55(Pt 5), 2069–73. doi:10.1099/ijs.0.63299-0

Solomon, K. V, Haitjema, C. H., Thompson, D. A, & O’Malley, M. A. (2014). Extracting data from the muck: deriving biological insight from complex microbial communities and non-

model organisms with next generation sequencing. Current Opinion in Biotechnology, 28, 103–10. doi:10.1016/j.copbio.2014.01.007

Spencer, D. H., Kas, A., Smith, E. E., Christopher, K., Sims, E. H., Hastings, M., … Burns, J. L. (2003). Whole-Genome Sequence Variation among Multiple Isolates of Pseudomonas aeruginos. J. Bacteriol, 185(4) 1316 - 1325. doi:10.1128/JB.185.4.1316

Strous, M., Fuerst, J., Kramer, E. H. M., Logemann, S., Gerard, M., Pas-schoonen, K. Van De, … Jetten, M. S. M. (1999). Missing lithotroph identifed as new planctomycete. Nature, 400(July), 446 – 449.

Strous, M., Van Gerven, E., Zheng, P., Kuenen, J. G., & Jetten, M. S. M. (1997). Ammonium removal from concentrated waste streams with the anaerobic ammonium oxidation (Anammox) process in different reactor configurations. Water Research, 31(8), 1955–1962. doi:10.1016/S0043-1354(97)00055-9

Tang, S., Gong, Y., & Edwards, E. a. (2012). Semi-automatic in silico gap closure enabled de novo assembly of two Dehalobacter genomes from metagenomic data. PloS One, 7(12), e52038. doi:10.1371/journal.pone.0052038

U.S. Environmental Protection Agency. (1993). Technical Factsheet on : BENZENE, 1–5. Retrieved from http://www.epa.gov/airtoxics/hlthef/benzene.html

U.S. Environmental Protection Agency. (1995). OPPT Chemical Fact Sheets: Chlorobenzene Fact Sheet. Pollution Prevention and Toxics, 7407. Retrieved from http://www.epa.gov/chemfact/chlor-sd.pdf

U.S. Environmental Protection Agency. (2007). Control of Hazardous Air Pollutants from Mobile Sources: Final Rule to Reduce Mobile Source Air Toxics, (February). Retrieved from http://www.epa.gov/oms/regs/toxics/420f07017.pdf

U.S. Environmental Protection Agency. (2008). EPA’s 2008 Report on the Environment. Retrieved from http://cfpub.epa.gov/ncea/cfm/recordisplay.cfm?deid=190806#Download

U.S. Environmental Protection Agency. (2009). National Primary Drinking Water Regulations. Retrieved from http://www.epa.gov/ogwdw/consumer/pdf/mcl.pdf

U.S. Environmental Protection Agency. (2011). Common Chemicals Found at Superfund Sites. Retrieved from http://www.epa.gov/superfund/health/contaminants/radiation/chemicals.htm

Ulrich, A. C., Beller, H. R., & Edwards, E. A. (2005). Metabolites detected during biodegradation of 13C6-benzene in nitrate-reducing and methanogenic enrichment cultures. Environmental Science & Technology, 39(17), 6681–91. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/16190227

Ulrich, A. C., & Edwards, E. A. (2003). Physiological and molecular characterization of anaerobic benzene-degrading mixed cultures. Environmental Microbiology, 5(2), 92–102. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/12558592

Urios, L., Michotey, V., Intertaglia, L., Lesongeur, F., & Lebaron, P. (2008). Nisaea denitrificans gen. nov., sp. nov. and Nisaea nitritireducens sp. nov., two novel members of the class Alphaproteobacteria from the Mediterranean Sea. International Journal of Systematic and Evolutionary Microbiology, 58(Pt 10), 2336–41. doi:10.1099/ijs.0.64592-0

Van der Zaan, B. M., Saia, F. T., Stams, A. J. M., Plugge, C. M., de Vos, W. M., Smidt, H., … Gerritse, J. (2012). Anaerobic benzene degradation under denitrifying conditions: Peptococcaceae as dominant benzene degraders and evidence for a syntrophic process. Environmental Microbiology, 14(5), 1171–81. doi:10.1111/j.1462-2920.2012.02697.x

Varga, J. J., Losada, L., Zelazny, A. M., Brinkac, L., Harkins, D., Radune, D., … Goldberg, J. B. (2012). Draft genome sequence determination for cystic fibrosis and chronic granulomatous disease Burkholderia multivorans isolates. Journal of Bacteriology, 194(22), 6356–7. doi:10.1128/JB.01306-12

Vogel, T. M., & Grbić-Galić, D. (1986). Incorporation of Oxygen from Water into Toluene and Benzene during Anaerobic Fermentative Transformation. Applied and Environmental Microbiology, 52(1), 200–202.

Weiss, J. V., Rentz, J. A., Plaia, T., Neubauer, S. C., Merrill-Floyd, M., Lilburn, T., … Emerson, D. (2007). Characterization of Neutrophilic Fe(II)-Oxidizing Bacteria Isolated from the Rhizosphere of Wetland Plants and Description of Ferritrophicum radicicola gen. nov. sp. nov., and Sideroxydans paludicola sp. nov. Geomicrobiology Journal, 24(7-8), 559–570. doi:10.1080/01490450701670152

Wöhlbrand, L., Kallerhoff, B., Lange, D., Hufnagel, P., Thiermann, J., Reinhardt, R., & Rabus, R. (2007). Functional proteomic view of metabolic regulation in “Aromatoleum aromaticum” strain EbN1. Proteomics, 7(13), 2222–39. doi:10.1002/pmic.200600987

Xie, C.-H., & Yokota, A. (2006). Reclassification of [Flavobacterium] ferrugineum as Terrimonas ferruginea gen. nov., comb. nov., and description of Terrimonas lutea sp. nov., isolated from soil. International Journal of Systematic and Evolutionary Microbiology, 56(Pt 5), 1117–21. doi:10.1099/ijs.0.64115-0

Yumoto, I., Yamazaki, K., Hishinuma, M., Nodasaka, Y., Suemori, a, Nakajima, K., … Kawasaki, K. (2001). Pseudomonas alcaliphila sp. nov., a novel facultatively psychrophilic alkaliphile isolated from seawater. International Journal of Systematic and Evolutionary Microbiology, 51(Pt 2), 349–55. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/11321079

Zhang, H. (2003). Gemmatimonas aurantiaca gen. nov., sp. nov., a Gram-negative, aerobic, polyphosphate-accumulating micro-organism, the first cultured representative of the new bacterial phylum Gemmatimonadetes phyl. nov. International Journal of Systematic and Evolutionary Microbiology, 53(4), 1155–1163. doi:10.1099/ijs.0.02520-0

Zhang, T., Bain, T. S., Nevin, K. P., Barlett, M. a, & Lovley, D. R. (2012). Anaerobic benzene oxidation by Geobacter species. Applied and Environmental Microbiology, 78(23), 8304– 10. doi:10.1128/AEM.02469-12

Appendix 1: Assessment of genome completeness

Single copy genes found in all bacterial and archaeal genomes, and their presence in selected scaffolds, identified by “x” if present or “-“ if absent. Single copy Thiobacillus Aromatoleum Melioribacter Peptococcaceae Peptococcaceae gene (or gene Scaffold 11 Scaffold 12 Scaffold 2 Scaffold 29 Scaffold 36 cluster) RIBOSOMAL PROTEINS – ALL ORGANISMS LSU ribosomal - - x - - protein L1p (L10Ae) LSU ribosomal x x x - - protein L2p (L8e), L3p (L3e), L4p (L1e), L5p (L11e), L6p (L9e), L14p (L23e), L15p (L27Ae),L18p (L5e), L22p (L17e), L23p (L23Ae), L24p (L26e), L29p (L35e)* LSU ribosomal - - - - - protein L10p (P0) LSU ribosomal - - x - - protein L11p (L12e) LSU ribosomal - - x - - protein L7/L12p LSU ribosomal x x x - - protein L13p (L13Ae) LSU ribosomal x x x - - protein L30p (L7e) SSU ribosomal - - x - - protein S2p

(SAe) SSU ribosomal x x x - - protein S3p (S3e), S8p (S15Ae), S17p (S11e), S19p (S15e) SSU ribosomal x x x - - protein S4p (S9e), S11p (S14e), S13p (S18e) SSU ribosomal x x x - - protein S5p (S2e) SSU ribosomal - - x - - protein S7p (S5e) SSU ribosomal x x x - - protein S9p (S16e) SSU ribosomal x x x - - protein S10p (S20e) SSU ribosomal - - x - - protein S12p (S23e) SSU ribosomal x x x - - protein S14p (S29e) SSU ribosomal x x x - - protein S15p (S23e) LSU ribosomal - - - - - L7Ae Total (out of 10 10 16 0 0 18)

RIBOSOMAL PROTEINS – BACTERIA

ONLY LSU ribosomal x x x - x protein L9p LSU ribosomal x x x - - protein L16p LSU ribosomal x x x - - protein L17p LSU ribosomal x - x x - protein L19p LSU ribosomal x x x x - protein L20p LSU ribosomal x x x - - protein L21p LSU ribosomal x x x - - protein L27p LSU ribosomal - - - - - protein L28p Ribosomal ? ? - - ? protein L31 Ribosomal - - - ? - protein L32 Ribosomal ? ? ? - - protein L33 Ribosomal ? ? - - ? protein L34 Ribosomal ? ? ? ? - protein L35 Ribosomal - - - - - protein L36 SSU ribosomal x x x - x protein S6p SSU ribosomal x - x x - protein S16p SSU ribosomal x x x - - protein S18p SSU ribosomal x x x - - protein S20p Total (out of 11 9 11 3 2 18) tRNA

SYNTHETASES AND OTHER SINGLE COPY GENES Alanyl-tRNA x x x x - synthetase (EC 6.1.1.7) Arginyl-tRNA x x x - x synthetase (EC 6.1.1.19) aspartyl-tRNA x x x x - synthetase asparaginyl------trna synthetase Cysteinyl-tRNA x - x - - synthetase (EC 6.1.1.16) Glutaminyl- x x x - x tRNA synthetase (EC 6.1.1.18) Glutamyl-tRNA x - x - - synthetase (EC 6.1.1.17) Glycyl-tRNA x - x - - synthetase (EC 6.1.1.14) Histidyl-tRNA x x x x - synthetase (EC 6.1.1.21) Isoleucyl-tRNA x x x - - synthetase (EC 6.1.1.5) Leucyl-tRNA x x x - - synthetase (EC 6.1.1.4) Lysyl-tRNA x - x - x synthetase (class II)(EC 6.1.1.6) Methionyl- x x x - x

tRNA synthetase (EC 6.1.1.10) Phenylalanyl- x x x x - tRNA synthetase alpha chain (EC 6.1.1.20) Prolyl-tRNA x x x - - synthetase (EC 6.1.1.15) Seryl-tRNA x - x - x synthetase (EC 6.1.1.11) Threonyl-tRNA x x x x - synthetase (EC 6.1.1.3) Tryptophanyl- x x - x - tRNA synthetase (EC 6.1.1.2) Tyrosyl-tRNA x x x x - synthetase (EC 6.1.1.1) Valyl-tRNA x - x - - synthetase (EC 6.1.1.9) preprotein - - - - x translocase secG (Archaea: SEC61 complex) Ribonuclease P x x x - - protein RecA protein x x x - - (Archaea: RadA) DNA gyrase x x x - x subunit A (EC 5.99.1.3) DNA gyrase x x x - x subunit B (EC 5.99.1.3)

DNA-directed - - x - - RNA polymerase beta subunit (EC 2.7.7.6) Translation x x x x - elongation factor P (Archaea: translation elongation factor 1 alpha subunit) TOTAL (out of 24 18 24 8 8 27)

TOTAL – ALL 45 37 51 11 10 GENES (out of 61) % Complete 73.8 60.7 83.6 18.0 16.4 (Observed/Exp ected*100%)

Appendix 2: Degradation of monochlorobenzene

A2.1 Introduction

One side project I was involved in concerned monochlorobenzene (MCB) degrading consortia. These consortia were created by a previous Post Doctoral Fellow in Edwards’ Laboratory, Xiaoming Liang (Liang et al., 2013), by blending together a monochlorobenzene → benzene enrichment with benzene → CO2 + CH4. Blended enrichments were developed that degraded repeated doses of MCB to HCl, CO2 and CH4. The combination these two cultures is especially exciting because chlorobenzene degradation generally does not proceed further than benzene. Attempts to remediate monochlorobenzene can result in a more problematic contaminant being produced, as benzene is more toxic than monochlorobenzene (MCL 0.005mg/L versus MCL 0.1mg/L for MCB (U.S. Environmental Protection Agency, 1995)). Linking benzene degradation to monochlorobenzene dehalogenation removes this problem. Benzene degradation is an oxidation reaction, and an electron acceptor is required. Monochlorobenzene → benzene is a reduction reaction, and an electron donor is required. The two reactions are coupled in the blendedBenzene)degrada

Benzene&oxida)on&coupled& MCB&dechlorina)on& to&methanogenesis&

Cl&

- e +&&&& & 2eK&&&&+&&&2H+&&

CO2

CH4

Figure A2.1: Benzene oxidation coupled to MCB dechlorination 6&

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acceptor (see Figure A2.1).

The key organisms involved in MCB → benzene and benzene → CO2 + CH4 in these cultures are proposed to be a Dehalobacter (MCB → benzene) and the Syntrophobacterales ORM2

(benzene → CO2 + CH4).

A2 Results and discussion

A2.1 Microscopy reveals a very diverse enrichment

While morphology can not be used to conclusively determine phylogeny, an estimate of enrichment diversity can be made from microscopy. If a wide range of morphologies are present, the culture is likely more diverse than if a single morphology is present. Figure A2.2 shows MCB and benzene degrading consortium scale-up 2 (MCBBSU2) stained with Gram’s Stain. A large protist is present, along with a mixture of microbial cell types (cocci and rods). Sediment from the culture is also visible. Figure A2.3 shows a similar image, with a possible diatom present.

Figure A2.2: MCBBSU2 with Gram’s Stain at 600x magnification.

Figure A2.3: MCBBSU2 with Gram’s Stain at 600x magnification.

Figure A2.4: MCBBSU2 with DAPI staining at 100x magnification.

DAPI (4',6-diamidino-2-phenylindole) is a fluorescent general nucleic acid stain, used to enhance visibility of cells for microscopy. Figure A2.4 shows a DAPI-stained MCBBSU2 sample. The stain has bound to sediment in the culture, making it difficult to distinguish cells from sediment. Figure A2.5a) shows another DAPI-stained sample from MCBBSU2. In this field of view, a protist had bound sufficient DAPI that it fluoresces brightly and drowns out background fluorescence. Figure A2.5b) shows the bright-field image of the same field of view for comparison.

A B

Figure A2.5: a) shows a DAPI stained protist from the MCBBSU2 enrichment (1000x magnification). B) shows the bright field image of the same field of view.

Microscopy of this MCB and benzene degrading enrichment blend shows that the enrichment is very diverse. While a mixture of Archaea and Bacteria were anticipated, the presence of protists was unanticipated. Protists have been introduced to the enrichment with the sediment used to initiate the microcosms they descend from. Figure A2.5b) raises the possibility that protist predation of microorganisms may be an issue. It appears to show a protist that has engulfed two rod-shaped microbes. Protist predation can further hinder the growth of already- slow-growing organisms. This protist may be a Tetrahymena, a pear-shaped ciliate, based on morphology.

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A2.2 Pyrotag sequencing

Pyrotag sequencing was carried out to examine the community present in two different MCB →

CO2 + CH4 cultures. Cultures are designated MCBBSU3 (MCB and benzene degrading consortium scale-up 3) and MCBBSU4 (MCB and benzene degrading consortium scale-up 4). Organisms present in the scale-up cultures at greater than 0.5% abundance are shown in Table A2.1.

Table A2.1: Taxonomy of groups present at abundances greater than 0.5% in MCBBSU3 and MCBBSU4 MCB Scale- MCB Scale- Domain Phylum Class Order up 3 (2012) up 4 (2013) Archaea Other Other Other 0.7% <0.5% Archaea Crenarchaeota C2 pGrf C26 0.5% <0.5% Archaea Euryarchaeota Methanomicrobia Methanomicrobiales 1.3% 1.2% Archaea Euryarchaeota Methanomicrobia Methanosarcinales 0.9% <0.5% Archaea Euryarchaeota Thermoplasmata Other 0.5% <0.5% Bacteria Other Other Other 21.2% 10.7% Bacteria AC1 SHA-114 Other 0.7% <0.5% Bacteria AC1 TA06 Other <0.5% 15.7% Bacteria Bacteriodetes Bacteroidia Bacteriodales 2.9% 1.9% Bacteria Bacteriodetes Sphingobacteria Sphignobacteriales 0.9% <0.5% Bacteria Caldiserica Caldisericia Caldisericales 1.1% 2.1% Bacteria Chlorobi BSV19 Other 0.7% <0.5% Bacteria Chlorobi Ignavibacteria Ignavibacteriales 0.9% 0.7% Bacteria Chloroflexi Anaerolineae Anaerolinales <0.5% 0.8% Bacteria Chloroflexi Anaerolineae envOPS12 1.0% <0.5% Bacteria Chloroflexi Anaerolineae GCA004 <0.5% 0.7% Bacteria Chloroflexi Anaerolineae SHA-20 <0.5% 1.0% Bacteria Chloroflexi Anaerolineae Other 1.5% <0.5% Bacteria Chloroflexi Dehalococcoidetes Dehalococcoidales 0.5% 3.8% Bacteria Chloroflexi Dehalococcoidetes NT-B4 0.6% <0.5% Bacteria Chloroflexi Other Other 0.7% <0.5% Bacteria Elusimicrobia 4-29 Other <0.5% 0.7% Bacteria Firmicutes Clostridia Clostridiales 11.6% 7.1% Bacteria Firmicutes Clostridia SHA-98 <0.5% 0.7% Bacteria Firmicutes Clostridia Other 0.7% <0.5% Bacteria Firmicutes Other Other 0.7% <0.5%

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Bacteria Nitrospirae Nitrospira Nitrospirales 4.5% 0.9% Bacteria OD1 ABY1 Other <0.5% 2.6% Bacteria OP11 Other Other <0.5% 3.5% Bacteria OP3 koll11 GIF10 0.5% 0.7% Bacteria OP8 OP8_1 SHA-124 0.5% <0.5% Bacteria OP8 OP8_1 Other 0.6% 0.5% Bacteria Planctomycetes Phycisphaerae Other 1.7% <0.5% Bacteria Planctomycetes Other Other <0.5% 0.8% Bacteria Proteobacteria Betaproteobacteria Burkholderiales <0.5% 0.6% Bacteria Proteobacteria Betaproteobacteria Hydrogenophilales 0.8% 0.5% Bacteria Proteobacteria Betaproteobacteria Rhodocyclales <0.5% 0.8% Bacteria Proteobacteria Betaproteobacteria Other 0.7% <0.5% Bacteria Proteobacteria Deltaproteobacteria Desulfobacterales 0.5% <0.5% Bacteria Proteobacteria Deltaproteobacteria Syntrophobacterales 17.0% 18.7% Bacteria Proteobacteria Deltaproteobacteria Other 2.2% <0.5% Bacteria Proteobacteria Gammaproteobacteria Chromatiales 1.2% 0.7% Bacteria Proteobacteria Gammaproteobacteria Oceanospirillales 0.5% <0.5% Bacteria SAR406 AB16 noFP_H7 <0.5% 1.5% Bacteria Spirochaetes Brevinematae Brevinematales <0.5% 3.2% Bacteria Spirochaetes Spirochaetes Spirochaetales 1.1% 2.2% Bacteria Synergistetes Synergistia Synergistales 0.6% 2.2% Bacteria Tenericutes Mollicutes Other <0.5% 3.5% Bacteria Tenericutes Other Other 0.7% <0.5% Bacteria WS3 PRR-12 SSS58A 0.7% <0.5%

A large number of OTUs were unclassifiable beyond "Bacteria" in MCBBSU3 (21.2%). This may be due to the fact that the MCB scale-up contains large numbers of environmental bacteria that do not have cultured, sequenced representatives. Another high-abundance group is Syntrophobacterales (17% in MCBBSU3, 18.7% in MCBBSU4). This order includes the proposed benzene-degrading organism in these cultures, an uncultured Deltaproteobacteria, ORM2. The proposed monochlorobenzene dechlorinating organism in the microcosms is a Dehalobacter, which is classified within the Clostridiales order. Clostridiales were present at 11.6% abundance in MCBBSU3. In MCBBSU4, its abundance was 7.1%. Another dechlorinating clade, Dehalococcoidales, was present at 0.5% abundance in MCBBSU3, and 3.8% abundance in MCBBSU4. This may be carrying out MCB dechlorination also.

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To identify the OTUs of interest, reference sequences for some OTUs were BLASTed. The high-abundance Syntrophobacterales OTU returned hits for the same "uncultured bacterium clones" as ORM2 (top 5 hits are clones SHRa-h11, Hasda-A, D15-36, SHRa-65 and F5OHPNU07ILFVC, 4/5 of the top 5 BLAST hits for ORM2). The high-abundance Clostridiales OTU was also BLASTed. Hits for this OTU were all Dehalobacter. The high-abundance group classified as "bacteria/other" returns hits to uncultured bacterium clones. The top hit came from an oils sands tailings clone, another extreme, hydrocarbon rich environment.

A2.2 Metagenome assembly

DNA was extracted from MCBBSU4 and sequenced using Illumina HiSeq. A single library of paired end reads was sequenced. 27M read pairs were generated. Reads were trimmed for quality using Trimmomatic (Bolger et al., 2014). 21M paired reads passed quality testing and were used for assembly. Assembly was carried out using ABySS (Simpson et al., 2009). Different k values and c values were tested. The best assembly was generated using k 64, and c automatic (c value gets set to square root of median normalized kmer depth). A graph of contig length versus normalized kmer depth is shown in Figure A2.6. The 16S rRNA gene from ORM2 was identified on a contig with NKD = 17. Other contigs belonging to the ORM2 genome are likely to also be NKD =17. Contigs were mapped to ORM2 contigs, and Dehalobacter and Dehalococcoides genomes to check that these genomes were represented in the assembly. This showed that ORM2 was present, although not all ORM2 contigs were fully covered by MCBBSU4 contigs. Comparable numbers of reads mapped to Dehalobacter and Dehalococcoides genomes. The genomes were not covered fully either.

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Normalized kmer depth

Methanosaeta (nkd 17)

Contig length (bp)Contig length (bp)

Figure A2.6: NKD versus contig length for ABySS assembly (k64, c automatic)

A2.3 Identification of putative reductive dehalogenase genes

One aim for this project was to identify potential rdh genes, the genes responsible for reductive dehalogenation reactions. A large number of rdh genes have been identified in previous studies. Organisms usually have multiple rdh genes which are each specific to a particular substrate (Hug et al., 2013). This unique system may contain novel rdh genes.

Olivia Molenda identified potential rdh genes in the contigs from MCBBSU4. Potential rdh sequences were identified in the contigs by BLAST. These hits were annotated using RAST. Results are shown in Table A2.2. Highlighted genes are those that contain TAT export & 4Fe-4S regions essential to rdh. These sequences had BLAST hits to Dehalobacter FTH1 (91-100% identity). Futher analysis of the MCBBSU4 metagenome could include binning contigs by read

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depth or tetranucleotide frequency to create “bins” containing contigs from a single organism (Devine, 2013). More annotation of the contigs is also required to identify unique features of the organisms in this valuable enrichment.

Table A2.2: Putative rdh genes in MCBBSU4 contigs. Those containing TAT export and 4Fe-4S regions necessary for rdh activity are shaded grey.

Contig GC Contig Contig Normalized content ID length kmer depth (%) start stop strand Annotation Tetrachloroethene reductive dehalogenase 1030332 847 3 44 5 517 + TceA 1030332 847 3 44 837 688 - hypothetical protein Tetrachloroethene reductive dehalogenase 1372608 1375 6 47 103 570 + TceA Tetrachloroethene reductive dehalogenase 1467415 4690 5 38 58 483 + TceA 1467415 4690 5 38 678 541 - hypothetical protein 1467415 4690 5 38 772 1536 + Transcriptional regulator, Crp/Fnr family 1467415 4690 5 38 2485 1541 - Cell division trigger factor (EC 5.2.1.8) 1467415 4690 5 38 2683 3396 + Transcriptional regulator, Crp/Fnr family 1467415 4690 5 38 4526 3549 - Cell division trigger factor (EC 5.2.1.8) Tetrachloroethene reductive dehalogenase 1730258 2829 5 42 1955 561 - PceA (EC 1.97.1.8) 1730258 2829 5 42 2339 1956 - hypothetical protein 1730258 2829 5 46 2690 2571 - Transcriptional regulator, Crp/Fnr family Tetrachloroethene reductive dehalogenase 1747532 2146 5 46 5 229 + TceA Tetrachloroethene reductive dehalogenase 1747532 2146 5 42 403 1935 + PceA (EC 1.97.1.8) Tetrachloroethene reductive dehalogenase 257284 2945 5 42 17 1396 + PceA (EC 1.97.1.8) Phosphatidylserine/phosphatidylglycerophos phate/cardiolipi n synthases and related 257284 2945 5 42 1409 1714 + enzymes 681041 2557 5 41 132 19 - hypothetical protein 681041 2557 5 41 799 86 - Transcriptional regulator, Crp/Fnr family Tetrachloroethene reductive dehalogenase 681041 2557 5 41 2555 1224 - TceA Tetrachloroethene reductive dehalogenase 746870 1388 4 40 1235 684 - TceA

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Appendix 3: Attempted Anammox enrichment

A3.1 Introduction

Nitrogen cycling is a key issue in the nitrate-reducing cultures, affecting benzene degradation rates and community structure. If nitrite accumulates as a result of partial nitrate reduction, it can inhibit benzene degradation, when concentrations build up to around 5mM. One group of bacteria that may assist benzene degradation under nitrate reducing conditions are anaerobic ammonium oxidizing bacteria (anammox) (Strous et al., 1999). These bacteria oxidize ammonium with nitrite derived from the incomplete reduction of nitrate (Strous, Van Gerven, Zheng, Kuenen, & Jetten, 1997). Anammox are present in some of the cultures maintained in the lab (Gitiafroz, 2012). Isolation of anammox bacteria would enable us to characterize their activity, and recombine them with nitrate-reducing benzene-degrading consortia with a low abundance of anammox bacteria or no anammox bacteria present to examine their effect.

A3.2 Methods

Anammox enrichment was attempted using batch cultures inoculated with benzene-degrading culture and supplied with 12 mM ammonium and 2mM nitrite as the sole electron donor and acceptor. Enrichments were set up from both Cart T3 and Swamp T1. Controls were enrichments with no nitrite supplied, killed controls, and sterile medium. Ammonium concentration was measured by Hach High Ammonium kit, and nitrite and nitrate concentrations measured by IC. Presence and quantity of anammox bacteria were detected by qPCR. DNA extraction for qPCR was the same as for PCR. Quantitative-PCR (qPCR) was carried out using the Pla46f forward primer (5′-GAC TTG CAT GCC TAA TCC-3′) and Amx368r reverse primer (5′-CCT TTC GGG CAT TGC GAA-3′). The AMX8 plasmid was used for standard curve generation. Reaction components were 10uL SsoFast Evagreen mastermix (Bio-Rad Laboratories Inc., Ca), 0.5uL 10uM forward primer, 0.5uL 10uM reverse primer, 7uL ultrapure water and 2uL sample or standard DNA. PCR conditions were as follows: initial denaturation at 98°C for 2 minutes, followed by 40 cycles of: denaturation at 98°C for 5 seconds, annealing at 59°C for 5 seconds,

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and chain extension at 65°C for 10 seconds on a CFX96 Real-Time System C1000 Thermocycler (Bio-Rad Laboratories Inc., Ca).

A3.3 Results and discussion

Table A3.1 shows the abundances of anammox bacteria in enrichments. Anammox bacteria were present in samples, but at a lower proportion of the total bacteria than in one of the parent cultures, Cart T3. The exception was in Replicate C of the enrichment from the Swamp culture. This value is likely to be inaccurate as the copies of genbac per mL of culture was significantly lower compared to the other cultures so is indicative of poor amplification.

Table A3.1: Copies of amx (anammox-specific) 16SrRNA gene and copies of genbac (universal) 16SrRNA gene in anammox enrichments compared to parent culture (July 2013, after 10 months incubation)

Anammox Copies of Copies of as percent Sample Replicate amx per genbac per mL of total mL culture: culture: bacteria Anammox enrichment of cart A 4x10^6 1x10^9 0.32 cons Anammox enrichment of cart B 3x10^6 2x10^9 0.16 cons Anammox enrichment of cart C 6x10^5 5x10^8 0.13 cons Anammox enrichment of swamp A 2x10^5 8x10^8 0.03 cons Anammox enrichment of swamp B 2x10^5 1x10^9 0.02 cons Anammox enrichment of swamp C 3x10^5 9x10^6 2.93 cons Cart NO3- consolidated 3x10^6 6x10^8 0.42

Potential anammox activity was observed, as the culture removed nitrite without accumulating nitrate. After three cycles of nitrite removal, anammox activity ceased, and the cultures were

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transferred to fresh medium to attempt to re-activate the anammox activity. Figure A3.1 shows the nitrite concentrations in the cultures, and Figure A3.2 shows the nitrate concentrations, averaged between three repeats for each condition. There are no differences between autoclaved controls and the test cultures. Nitrate is being produced, but at very low levels near the detection threshold.

4.5"

3.5" Cart"cons"+NO2)" 3" Cart"cons,"no"NO2)" 2.5" Cart"cons"Killed" Swamp"cons"+NO2)" 2" Swamp"cons,"no"NO2)" 1.5" Swamp"cons"Killed" Sterile"

NO2$%concentra-on%(mM)% 1"

0.5"

0" 2013)01)26" 2013)03)17" 2013)05)06" 2013)06)25" 2013)08)14" Date%

Figure A3.1: Nitrite concentrations in enrichment bottles (error bars show standard deviation)

3.5"

2.5" Cart"cons"+NO2(" Cart"cons,"no"NO2("

2" Cart"cons"Killed" Swamp"cons"+NO2(" 1.5" Swamp"cons,"no"NO2(" NO3$%concentra-on%(mM)% Swamp"cons"Killed" 1" Sterile"

0.5"

0" 2013(01(26" 2013(03(17" 2013(05(06" 2013(06(25" 2013(08(14" Date% Figure A3.2: Nitrate concentrations in enrichment bottles (error bars show standard deviation)

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Further monitoring showed no activity. Results of pyrotag sequencing (See Chapter 4) did not show any Anammox bacteria. The closest relative to known Anammox organisms was a Brocadiaceae belonging to the Planctomycetes phylum. Anammox have been lost from the culture. Nitrite accumulation is not observed, which could be due to another organism capable of nitrite reduction taking over this role.

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Appendix 4: Microscopic techniques to examine microbial communities

A4.1 Introduction

Syntrophic associations may be promoted by close physical proximity, as short distances between cells exchanging metabolites may overcome diffusion constraints (van der Zaan et al., 2012). The benefits associated with close physical proximity may outweigh the metabolic cost associated with production of extra-cellular matrix substances to stay attached. Cells may also use iron sulfide or other surfaces in the cultures for electron transfer (Sieber et al., 2012). Physical relationships can be observed via microscopy. Cell morphologies may also provide insight into culture diversity. Morphological diversity may represent phylogenetic diversity, but a lack of morphological diversity does not necessarily reflect phylogenetic homogeneity.

A4.2 Methods

Different types of microscopy were used to examine cell morphology and adhesion to solid particles. Solid particles caused several difficulties during microscopy. Fluorescence microscopy using DAPI as a general nucleic acid stain was used to look for different cell morphologies and flocculation. Fluorescence in situ hybridization (FISH) was used to find out the possibility of identifying different phylogenetic groups, and scanning electron microscopy (SEM) was used to investigate the interactions of bacteria with solid particles in the culture. To carry out FISH, an oligonucleotide complementary to a target 16SrRNA sequence is tagged with a fluorescent molecule (the combination is known as a "probe") and hybridized with a fixed sample to enable base-pairing. Unbound probe is then washed off the sample, and the sample is examined using the appropriate filter for its excitation/emission spectra (Amann, Ludwig, & Schleifer, 1995). Cultures investigated included previously described benzene-degrading nitrate-reducing cultures, other cultures grown by lab members were also investigated to provide points of comparison. These included a methanogenic toluene-degrading culture and a methanogenic

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monochlorobenzene-degrading. Wild-type E. coli overnight cultures grown in LB were also used for probe validation.

A4.2.1 Fluorescence microscopy

Different sample preparation protocols were trialed to determine which protocols gave the clearest images for our cultures. Samples were withdrawn using sterile, disposable anaerobic syringes from various cultures. Samples were stained using 25 ug/mL 4',6-diamidino-2- phenylindole (DAPI). A variety of modifications were made to sample preparation protocols. Some samples were air-dried and flame-fixed on glass slides. In this procedure, 10-100uL sample dried onto the slide, with 100uL DAPI added after flame-fixation. Slides were then incubated with DAPI for 3-15 minutes and rinsed in sterile filtered MilliQ water. Alternately, samples were diluted with 1x phosphate buffered saline (PBS) to a total volume of 10 mL and vacuum filtered onto 0.2 micron polycarbonate membranes (EMD Millipore, MA), using 0.45 micron nitrocellulose membranes (EMD Millipore, MA) as support membranes. Polycarbonate membranes were then rinsed with 5mL sterile filtered MilliQ water before they were air-dried. Once membranes were dried, 100uL DAPI was added and membranes were incubated in the dark for 3-15 minutes. Membranes were rinsed by placing them in a petri dish filled with sterile filtered MilliQ water for several seconds before air-drying them again. All samples were examined using either an Olympus BX51 fluorescence microscope fitted with an Olympus DP72 camera and Cellsense software, or an Olympus IX83 inverted microscope fitted with a Photometrics Coolsnap HQ2 CCD camera and Metamorph Premier software, both equipped with filter cubes with DAPI-specific excitation/emission spectra supplied by Olympus.

A4.2.2 Fluorescence in situ Hybridization

Fluorescence in situ hybridization (FISH) is an established technique for visualization of different phylogenetic groups simultaneously, but its application requires optimization for culture conditions and organisms of interest. Established general archaea and bacteria probes were used to optimize protocols. These probes are included in Table A4.1 Different fluorescent

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dyes were attached to probes so that multiple probes could be hybridized to the same sample and detected simultaneously. Dyes used were Cy3, Cy5 and 6-FAM, which do not have overlapping excitation/emission spectra.

Table A4.1: Probes used to optimize FISH protocols

Probe name Specificity 5' modification Sequence (5'-3')

Eub338(I) hits 90% of domain bacteria Cy3 GCT GCC TCC CGT AGG AGT

Eub338(II) hits 69% of planctomycetales Cy3 GCA GCC ACC CGT AGG TGT

Eub338(III) hits 93% order Verrucomicrobiales Cy3 GCT GCC ACC CGT AGG TGT

ARCH915 Euryarchaea 6-FAM GTG CTC CCC CGC CAA TTC CT nonEUB none Cy5 ACTCCTACGGGAGGCAGC

The protocols steps are 1) Sample 2) Fixation in paraformaldehyde (PFA) 3) Re- suspension in 1:1 1x PBS: ethanol 4) Attachment onto gelatin-coated glass slide or 0.2 micron polycarbonate filter 5) Dehydration 6) Hybridization mix preparation 7) Hybridization 8) Washing 9) Counterstaining 10) Examination. Variations of these steps were used to optimize hybridization of our samples. The optimized protocol is as follows:

One mL samples were preserved in 10mL 1% paraformaldehyde (PFA) in 1x PBS overnight. The 10mL fixation mixture was filtered onto a 0.2-micron polycarbonate filter (EMD Millipore, MA), using 0.45-micron nitrocellulose membranes (EMD Millipore, MA) as support membranes under vacuum. Polycarbonate membranes were then rinsed with 5mL sterile filtered MilliQ water before they were air-dried. Membranes were then placed onto parafilm-wrapped glass slides and 100uL hybridization mix was added. Excess hybridization mix was poured on a

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Kimwipe and placed in a 50mL falcon tube with the slides. Slides were incubated at 46°C for 1.5 hours. Membranes were then removed from the slides and placed in 50mL wash buffer in polyethylene tubes at 48°C for 15 minutes. Membranes were then rinsed in a petri dish of sterile milliQ water and air dried in the dark. To counter stain the membranes, 1 drop DAPI solution (Life Technologies ready-to-use formulation DAPI) was added to each membrane and incubated for 5 minutes in the dark. Membranes were then rinsed in sterile milliQ water and air dried in the dark before being covered with citifluor and a coverslip. Coverslips were sealed with nail polish. Slides were stored in the dark until examination using a BX51 Olympus fluorescence microscope under with filters supplied by Olympus for the different dyes.

A4.2.3 Scanning electron microscopy

Scanning electron microscopy (SEM) was carried out to examine finer details of cell morphology and physical interactions with solids. 5mL Cart NO3- consolidated culture was centrifuged at 3,000g for 20 minutes. Supernatant was removed, and the sample was resuspended in 4% gluteraldehyde in Sorenson’s phosphate buffer (0.1M, pH 7.2) and fixed for 2 hours at room temperature. The sample was then centrifuged again and resuspended in 0.1M phosphate buffer 3x before being re-suspended in 1% osmium tetroxide in 0.1M phosphate buffer and incubated for 1 hour at room temperature. The sample was then centrifuged again and resuspended in 0.1M phosphate buffer 3x. 200uL of fixed sample was then dropped onto poly-l- lysine coated coverslips and settled for 30 minutes. The coverslips were then dehydrated in an ethanol series (30%, 50%, 70%, 80%, 90% and 100%) for 5 minutes each before they were dehydrated in an ethanol: hexamethyldisilazane (HMDS) series (3:1, 1:1, 1:3, 100% HMDS) for 5 minutes each. The sample was then left to dry overnight. Aluminium stubs were covered with a square of carbon tape, and cover slips placed on these, sample-side up. Silver paint was used to trail a conductive strip from the top surface of the coverslip to the aluminium stub. Samples were then sputter-coated with a gold-palladium alloy under an argon atmosphere. A Hitachi S-2500 Scanning Electron Microscope equipped with Quartz PC1 software was used to examine coverslips.

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A4.3 Results

A4.3.1 DAPI-stained microscopy results

DAPI staining was a useful tool for examining cultures. Cultures with large amounts of sediment were not suited to this technique as sediment bound the DAPI and caused large amounts of background fluorescence (see A2.4). It was used to look at the differences between the abundance of microbes in Cart T3 supernatant (Figure A4.1), versus the abundance of microbes seen after the culture is shaken ((Figure A4.2). It was also useful to determine the effects of PFA fixation on cells. Figure A4.3 shows cells stained with DAPI without fixation, compared to Figure A4.4, where cells were fixed in 4% PFA. Fixed cells are less bright than unfixed cells. This may be due to the fixation cross-linking cells walls to such a degree that DAPI is not able to penetrate as easily,

Figure A4.1: Cart T3 (shaken) flame fixed smear (400x magnification)

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Figure A4.2: Cart T3 (supernatant) flame fixed smear (400x magnification)

Figure A4.3: Unfixed DAPI stained E. coli cells (1000x magnification)

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Figure A4.4: E. coli cells fixed in 4% PFA and DAPI stained (1000x magnification)

A4.3.2 FISH results

Two FISH images are presented which illustrate the two main issues encountered during FISH. Figure A4.5 shows an E. coli cells which have been hybridized with the probe Eub338(I) according to the protocol in A4.2.1. There are unhybridized cells present. Unhybridized cells are blue as they have been stained with DAPI, whereas hybridized cells are red plus blue (magenta), as the Cy3 dye attached to the Eub338 probe was false-coloured red in this image. Figure A4.6 shows the opposite problem, where probes have hybridized non-specifically. The sample is from a toluene-degrading methanogenic enrichment. This sample was hybridized directly on a well on glass slide, at 50°C. Hybridization temperature was increased from 46°C to attempt to increase the stringency of binding. Again, blue is DAPI (all cells), and red is Cy3 (bacterial cells). Green here is representing 6-FAM, the dye on the Arch915 probe. Cells show dual hybridization with both the archaeal and bacterial probe, where red and green combine to yellow.

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Figure A4.5: E. coli cells fixed in 1% PFA and hybridized with Eub338(I) (red) and stained with DAPI (blue) (1000x magnification)

Figure A4.6: Toluene-degrading methanogenic culture fixed in 4% PFA and hybridized with Eub338(I) (red)and Arch915 (green) and stained with DAPI (blue) (1000x magnification)

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A4.3.3 SEM results

Cultured cells have been visualized by SEM, using the methods described previously, to check for any physical associations with the iron sulfide particles, which is included in the medium as a reducing agent. SEM imaging of Cart T3 culture shows a clump of iron sulfide with rod-shaped bacteria attached (Figure A4.7). Figure A4.7 shows an overview of a whole iron sulfide clump, with bacteria visible on the surface. Figure A4.8 shows a close-up of a region of the iron sulfide, with the 1-2µm by 0.4µm rod shapes more visible. Besides cells attached to iron sulfide clumps, chains of bacteria were also observed. Figure A4.9 shows a chain of rod-shaped 1-2µm by 0.4µm bacteria. Chains of bacteria or archaea may promote attachment to surfaces and biofilm formation. These images provide evidence that of physical associations between cells and iron sulfide particles. Surface attachment may assist syntrophic processes.

Figure A4.7: SEM image of Cart T3 Figure A4.8: SEM image of Cart T3 showing an iron sulfide particle with showing a closer look at the attachment of microbial rods attached microbial rods to iron sulfide

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Figure A4.9: SEM image of Cart T3 showing a chain of rod-shaped microorganisms

A4.4 Key points

• Microscopy can be a valuable technique to directly examine communities and reveal features such as morphological diversity and physical associations. • Physical associations exist between microbial cells and iron sulfide particles. These associations may enable tighter co-operation during syntrophic processes. • Optimization of protocols is crucial to obtain good quality images.

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Appendix 5: Location of sequence data files

A5.1 Pyrotag Sequence Data

Raw reads and quality files for all pyrotag sequencing runs are located on Syntrophy, within the folder /EdLab_FETA/Pyrotag/Pyrotag_sequencing_files_EdLab/…

Sequence data from separate rounds of sequencing are in different sub-folders. Listed are the names of the folders containing data used in this thesis, followed by the files the relevant data is included in. Bullet points list which samples’ sequence data are found in those files (simplified names and historical names are both included as simplified names are used in this thesis, but historical names were used during data collection and analysis).

Raw_Data_First_Round_2012 (files 1.722.TCA.454Reads.fna & 1.722.TCA.454Reads.qual)

• Swamp T1 (Swamp NO3 cons) • Swamp T2a (Swamp NO3 1ab) • Cart T2d (Cart NO3 PW1 10-1 third) • Cart T3c (Cart NO3 cons 10-2 third) • Cart T3 (Cart NO3 cons) • MCBBSU3 (M3B Scale up)

Raw_Data_Third_Round_July2013 (IEF2H5Y04Olivia.fna & IEF2H5Y04Olivia.qual)

• Cart T3 (Cart NO3-cons) • MCBBSU4 (MCBBscaleup4) • Cart T3g (FL Cart NO3 cons) • Cart T3h (FL Cart NO3 cons2)

Raw_Data_Fifth_Round_July2014 (2.864.TCA.454Reads.fna & 2.864.TCA.454Reads.qual)

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• Cart T3 (CartNO3-cons) • Cart T3f (Cartrevival)

QIIME input, scripts and analyzed data are in the folder Syntrophy/People/Students/Current/Sarah McRae 2012/Pyrotag

A5.2 Cart T3 Metagenome Data

Scripts and processed data are located on Sunfire in the folder /data4/sarah/Cart_NO3_metagenome/…

Raw read files (which include quality scores) are located on Sunfire. File paths (paired end data):

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1247.001.N701---N502.cartNO3- cons_pairedend_R1.fastq

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1247.001.N701---N502.cartNO3- cons_pairedend_R2.fastq

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1247.002.N701---N502.cartNO3- cons_pairedend_R1.fastq data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1247.002.N701---N502.cartNO3- cons_pairedend_R2.fastq

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1291.001.N701---N502.cartNO3- cons_pairedend_R1.fastq'

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1291.001.N701---N502.cartNO3- cons_pairedend_R2.fastq

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/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1291.002.N701---N502.cartNO3- cons_pairedend_R1.fastq

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1291.002.N701---N502.cartNO3- cons_pairedend_R2.fastq

File paths (mate pair data):

/data4/GQ-data/UofT_EdLab_MP_august2013_unzip/HI.1216.002.Index_12.cartNO3- cons_matepair_R1.fastq'

/data4/GQ-data/UofT_EdLab_MP_august2013_unzip/HI.1216.002.Index_12.cartNO3- cons_matepair_R2.fastq

A5.3 MCBBSU4 Metagenome Data

Scripts and processed data are located on Sunfire in the folder /data4/sarah/MCB_metagenome/…

Raw read files (which include quality scores) are located on Sunfire. File paths:

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1284.002.N704--- N504.MCBBSU4_R1.fastq

/data4/GQ-data/UofT_EdLab_PE_july2013_unzip/HI.1284.002.N704--- N504.MCBBSU4_R2.fastq