Aerobic Hydrocarbon-Degrading Microbial Communities in Oilsands Tailings Ponds

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2016 Aerobic Hydrocarbon-degrading Microbial Communities in Oilsands Tailings Ponds

Rochman, Fauziah

Rochman, F. (2016). Aerobic Hydrocarbon-degrading Microbial Communities in Oilsands Tailings Ponds (Unpublished doctoral thesis). University of Calgary, Calgary, AB. doi:10.11575/PRISM/24733 http://hdl.handle.net/11023/3508 doctoral thesis

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Aerobic Hydrocarbon-degrading Microbial Communities in Oilsands Tailings Ponds

Fauziah Fakhrunnisa Rochman

A THESIS

SUBMITTED TO THE FACULTY OF GRADUATE STUDIES

IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE

DEGREE OF DOCTOR OF PHILOSOPHY

GRADUATE PROGRAM IN BIOLOGICAL SCIENCES

CALGARY, ALBERTA

DECEMBER, 2016

Abstract

Oilsands process-affected water (OSPW), produced by the surface-mining oilsands industry in Alberta, Canada, is alkaline and contains salts, various metals, and hydrocarbon compounds. In this thesis, aerobic communities involved in several key biogeochemical processes in OSPW were studied. Degradation of several key hydrocarbons was analyzed in depth.

Benzene and naphthalene were used as models for aromatic hydrocarbons, in which their oxidation rates, degrading communities, and degradation pathways in OSPW were researched.

The potential oxidation rates were 36.7 μmol L-1 day-1 for benzene and 85.4 μmol L-1 day-1 for naphthalene. Via stable isotope probing (SIP), and high-throughput sequencing of 16S rRNA gene amplicons, it was discovered that strains of the genera Methyloversatilis and Zavarzinia were the main benzene degraders, while Thiococcus and Pseudomonas were the main naphthalene degraders. Cultivated strains of Zavarzinia and Pseudomonas were shown to be growing on benzene and naphthalene.

Metagenomics analysis revealed genes encoding oxygenases active against aromatic compounds, as well as catechol oxidases. Although these belonged to many phylogenetically diverse bacteria, only few bacteria were predominant in the SIP experiments. A highly divergent pmoA-like gene was also detected in the metagenome data. Here, the possibility of this gene allowing growth on short alkanes (C1 to C3) was examined. This gene was investigated via SIP and quantitative PCR. Results showed that the monooxygenase encoded by the gene has high affinity toward ethane and mostly propane. For the study of lighter hydrocarbons, methane, ethane, and propane were chosen as model compounds. OSPW was capable of supporting methane oxidation with a rate of 108.2

−1 −1 −1 −1 μmol of CH4 L OSPW d , ethane oxidation with a rate of 83.2 μmol of C2H6 L OSPW d ,

−1 −1 and propane oxidation with a rate of 58.6 μmol of C3H8 L OSPW d . SIP analysis uncovered

Methyloparacoccus to be predominant in methane-incubated samples, whereas Methyloversatilis was predominant in ethane and propane-incubated samples.

SIP technique was also employed to study photosynthetic bacterial communities and indigenous aerobic bacterial communities that assimilate methanol, acetate, and protein extracts.

All OSPW photosynthetic ‘heavy-DNA’ samples were dominated by unidentified

Planctomycetes. Predominant groups in methanol, acetate, and protein extract-SIPs were

Betaproteobacteria, Alphaproteobacteria, and Bacteroidetes. Finally, via a modified cultivation technique, a novel Verrucomicrobia was isolated from OSPW. The aerobic bacterium was named Oleiharenicola alkalitolerans gen. nov., sp. nov., and it was studied in depth via phylogenetic, chemotaxonomic and whole-genome sequencing techniques.

iii Acknowledgements

One of the lessons that I learned while earning this degree is that none of my achievements could have been attained without the help of others. It is, therefore, a pleasure to thank the many people who made this dissertation possible. First and foremost, I would like to thank my supervisor, Dr. Peter Dunfield, for providing me the opportunity to explore the microbial world. I barely knew how to use a micropipette when I first came, and now I have learned so many scientific methods and laboratory techniques due to your encouragement and dedication. Thank you for your patience, thoughtful guidance, critical comments, and insights in my research. My sincerest thanks to my committee, Dr. Gerritt Voordouw, Dr. Lisa Gieg, and Dr. Patrick Hettiaratchi, for your insightful comments and valuable time. I’d also like to thank my examiners Dr. Thomas Oldenburg and Dr. Tariq Siddique for giving your time and insights to improve this dissertation. My dissertation would not be possible without the following funding providers: Institute for Sustainable Energy, Environment and Economy (ISEEE), Genome Alberta, Genome Canada, Hydrocarbon Metagenomics group, and Syncrude Canada, Ltd. Sincere thanks to Dr. Lisa Gieg for allowing me to complete some experiments in your lab. Also, to Dr. Carolina Berdugo and Courtney Toth from the Gieg lab who helped me operate their lab GC for aromatics detection. I really appreciate the work dedicated by the departmental staff: Karen Barron, Christine Goodwin, Sophia George, David Bininda, Rebecca Lee, Carol Sprague, Laureen Clement, Hayley Harris, Bill Huddleston, Cate McRae, and Paulette Harrison, who have all made my time as a graduate student and teaching assistant at the U of C a lot easier. I am indebted to Dr. Allyson Brady, who was very helpful and accommodating in my first two years in the lab. Thank you for patiently teaching me how to operate all lab equipment. Thank you to Dr. Joongjae Kim, my classic-microbiology mentor, for generously sharing your vast knowledge and thoughts in the field. I will be forever grateful! I am also very thankful to Dr. Ivica Tamas, for introducing me to the basics of bioinformatics in a way that made it seemed so easy, and to Dr. Christine Sharp for all your help with the GC, SIP, QIIME, and countless other things. To Gareth Jones, thank you for taking care of the gas tanks business, and for having your arms wide open to help with every little problem in and out of the lab that I had to deal with.

ii Thank you to Dr. Angela Smirnova, for helping me with so many orders, Roshan Khadka, for sharing the valuable pmoA trees and ARB databases, and Ilona Ruhl for the very helpful and useful Illumina guidance. Thanks to Alireza Saidi-Mehrabad, for the times we shared working together on the tailings, bugging Joongjae for help, and talking about microbes and politics. To Emily Wang, Evan Haupt, Gul Zeb, Emad Albakistani, and Andriy Sheremet: thanks for always being wonderful friends! And to all past and present Dunfield lab members: thank you for the great times and the valuable friendship. I wish you all the best in life! The journey for this degree was not always smooth, especially that it happened at the same time as I embarked the new adventure of motherhood. But I was lucky to have endless family support. This doctorate wouldn’t be possible if my father didn't let my mom travel halfway around the world and stay with me in Calgary for my first two years of motherhood. Thank you so much umi and buya for your prayers and sacrifices! I’d also like to thank my sisters, Rya and Ifa, for coming over to help keep me sane in numerous ways. Especially to my children, Ibrahim and Mariam, thank you for being the most wonderful and understanding kids! Finally, to my husband, Mochamad, thanks for the fantastic support – I couldn't have asked for more.

iii

To my family− whose encouragement, inspiration, prayers, support and love

sustained me throughout my years pursuing education.

iv Table of Contents

Abstract ...... ii Acknowledgements ...... ii Table of Contents ...... v List of Tables ...... viii List of Figures and Illustrations ...... xii List of Symbols, Abbreviations and Nomenclature ...... xviii

CHAPTER ONE: INTRODUCTION ...... 1 1.1 Research objectives...... 1 1.2 Dissertation structure ...... 4

CHAPTER TWO: LITERATURE REVIEW ...... 7 2.1 Oilsands in Alberta ...... 7 2.1.1 Background...... 7 2.1.2 Oilsands recovery ...... 7 2.2 Oilsands tailings ponds ...... 10 2.2.1 Oilsands process-affected water (OSPW) ...... 11 2.2.2 Layers of the tailings ponds ...... 12 2.3 Petroleum hydrocarbons ...... 13 2.3.1 Hydrocarbons in the tailings ponds ...... 14 2.3.2 Polycyclic aromatic hydrocarbons ...... 15 2.4 Microbial hydrocarbon degradation in OSPW ...... 16 2.4.1 Aerobic hydrocarbon degradation ...... 17 2.5 The oxygenases ...... 18 2.5.1 Oxygenases ...... 18 2.5.2 Aromatic hydrocarbon degradation via oxygenases ...... 19 2.5.3 Catechol dioxygenases ...... 21 2.5.4 Aerobic alkane oxidation ...... 21 2.6 Study of hydrocarbon-degrading bacteria in the environment ...... 23 2.6.1 Activity measurements to monitor microbial activity ...... 23 2.6.2 Enrichments: growing target organisms ...... 23 2.6.3 Culture-dependent methods: Isolation of microorganisms from oil-contaminated environments ...... 25 2.6.4 Culture-independent method: Next-generation sequencing to study oil- contaminated environments ...... 25 2.6.4.1 DNA based stable isotope probing ...... 26 2.6.4.2 16S rRNA gene sequencing ...... 28 2.6.4.3 Metagenomics ...... 30 2.7 Microbial communities in the tailings ...... 34 2.7.1 Aerobic microbial communities ...... 34 2.7.2 Anaerobic microbial communities ...... 35 2.8 Bioremediation...... 36 2.9 Conclusion ...... 37

v PREFACE ...... 39

CHAPTER THREE: BENZENE AND NAPHTHALENE DEGRADING BACTERIAL COMMUNITIES IN OILSANDS TAILINGS ...... 40 3.1 Abstract Art...... 41 3.2 Abstract ...... 42 3.3 Introduction ...... 43 3.4 Methods ...... 45 3.4.1 Sampling and water chemistry ...... 45 3.4.2 Potential hydrocarbon oxidation rates ...... 46 3.4.3 Stable isotope probing (SIP) ...... 47 3.4.4 Metagenomics...... 48 3.4.5 Isolation of hydrocarbon degrading bacteria ...... 49 3.4.6 Hydrocarbon degradation capabilities of bacterial isolates ...... 50 3.5 Results ...... 51 3.5.1 Hydrocarbon oxidation ...... 51 3.5.2 SIP and 16S rRNA gene sequencing ...... 53 3.5.3 WIP-OSPW metagenome ...... 56 3.5.4 Isolated bacterial strains and their degradation potentials ...... 61 3.6 Discussion ...... 61 3.7 Acknowledgments ...... 69

PREFACE ...... 70

CHAPTER FOUR: IDENTIFICATION OF METHANOL-, ACETATE-, PROTEIN- ASSIMILATING, AND PHOTOTROPHIC BACTERIAL COMMUNITIES IN THE SURFACE OILSANDS PROCESS-AFFECTED WATER BY STABLE-ISOTOPE PROBING ...... 71 4.1 Introduction ...... 71 4.2 Methods ...... 73 4.2.1 Sampling and water chemistry ...... 73 4.2.2 Stable isotope probing (SIP) and 16S rRNA gene sequencing ...... 73 4.3 Results ...... 75 4.4 Discussion ...... 80 4.5 Conclusions ...... 93

PREFACE ...... 94

CHAPTER FIVE: FUNCTIONAL ANALYSIS OF A NOVEL CU-MONOOXYGENASE DETECTED VIA METAGENOMIC ANALYSIS OF OSPW ...... 95 5.1 Introduction ...... 95 5.2 Methods ...... 97 5.2.1 Sampling and water chemistry ...... 97 5.2.2 qPCR primer design ...... 97 5.2.3 MLSB-OSPW samples enrichments ...... 98 5.2.4 Stable isotope probing preparation ...... 99 5.2.5 16S rRNA gene sequencing and analysis ...... 100

vi 5.3 Results and discussion ...... 101 5.4 Conclusions ...... 116

PREFACE ...... 117

CHAPTER SIX: OLEIHARENICOLA ALKALITOLERANS GEN. NOV., SP. NOV., A NEW MEMBER OF THE VERRUCOMICROBIA ISOLATED FROM AN OILSANDS TAILINGS POND ...... 118 6.1 Abstract ...... 119 6.2 Introduction ...... 120 6.3 Materials and Methods...... 122 6.3.1 Bacterial isolation ...... 122 6.3.2 Cell growing ...... 122 6.3.3 Physiological tests ...... 123 6.4 Results and discussion ...... 126 6.4.1 Morphology and physiology...... 126 6.4.2 Genome analysis ...... 133 6.4.3 Description of Oleiharenicola gen. nov...... 135 6.4.4 Description of Oleiharenicola alkalitolerans sp. nov...... 136 6.5 Acknowledgements ...... 137

CHAPTER SEVEN: GENERAL CONCLUSIONS AND FUTURE RESEARCH...... 138 7.1 Research summary ...... 138 7.2 Directions for future research ...... 141

REFERENCES ...... 144

APPENDIX A: SUPPLEMENTARY INFORMATION –BENZENE AND NAPHTHALENE DEGRADING BACTERIAL COMMUNITIES IN OILSANDS TAILINGS ...... 167

APPENDIX B: SUPPLEMENTARY INFORMATION – IDENTIFICATION OF METHANOL-, ACETATE-, PROTEIN-ASSIMILATING, AND PHOTOTROPHIC BACTERIAL COMMUNITIES IN THE SURFACE OILSANDS PROCESS- AFFECTED WATER BY STABLE ISOTOPE PROBING ...... 220

APPENDIX C: SUPPLEMENTARY INFORMATION – DETECTION AND FUNCTIONAL ANALYSIS OF A NOVEL CU-MONOOXYGENASE DETECTED VIA METAGENOMIC ANALYSIS OF WIP-OSPW ...... 228

APPENDIX D: COPYRIGHT PERMISSIONS ...... 237

vii List of Tables

Table 2-1. Enzyme classes related to alkane oxidation (Rojo, 2010). Copyright permission is available in Appendix D...... 22

Table 3-1. Identity of the major OTUs in each corresponding sample according to the closest cultured relative. Isotopically labelled benzene, naphthalene, and methanol, were amended into oilsands processed water sample (OSPW) samples. DNA from the samples were then fractionated by stable isotope probing (SIP). The table presents the heavy fractions of the SIP. OSPW-heavy: heavy fraction of OSPW receiving the same treatment as all of the amended samples. Only OTUs enriched at least 10-fold compared to the OSPW-heavy control are noted. UD indicates that no reads were detected in the original sample- the minimum x-fold enrichment in these cases is calculated assuming 1 read in the original sample...... 66

Table 4-1. Identity of the OTUs with the highest enrichments in each corresponding sample according to the closest cultured relative. Isotopically labelled methanol, acetate, protein extract, and CO2 were amended into oilsands processed water sample (OSPW) samples. DNA from the samples were then fractionated by stable isotope probing (SIP). The table presents the heavy fractions of the SIP. OSPW-heavy: heavy fraction of OSPW receiving the same treatment as all of the amended samples. UD: undetected...... 83

Table 6-1. Differential phenotypic characteristics of Oleiharenicola alkalitolerans NVTT and its closely related species...... 130

Table 6-2. Cellular fatty acid compositions of strain NVTT and its most closely related species. Values are the molar percentages of total cellular fatty acids...... 132

Table A1. List of bacterial isolates obtained from various OSPW enrichments and their closest relatives...... 175

Table A2 A-E. Top 25 OTUs of each sample site based on % of total reads in 16S rRNA gene sequencing analysis. OTUs were clustered based on their closest related species. ... 177

Table A2-A. Reads from DNA in OSPW sampled in August 2011...... 177

Table A2-B. Heavy SIP fraction (fraction 5) of OSPW sampled in August, 2011...... 178

Table A2-C. Heavy SIP Fractions (fractions 4 and 5) of OSPW sampled in August, 2011, amended with labelled benzene for 9 days...... 179

Table A2-D. Heavy SIP Fractions (fractions 5 and 6) of OSPW sampled in August, 2011, amended with labelled naphthalene for 7 days...... 180

Table A2-E. Heavy SIP Fractions (fractions 3 and 4) of OSPW sampled in August, 2011, amended with labelled methanol for 7 days...... 181

viii Table A3. Metagenome features of surface WIP-OSPW sampled in August 2011 of the combined Illumina and 454 assembly obtained from the IMG pipeline...... 182

Table A4 A-P. Genes encoding aerobic hydrocarbon (benzene and naphthalene) metabolism detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum aligned scaffold length was 2500 nucleotides (for 16S rRNA). All read depth = 1...... 183

Table A4-A. Genes encoding for dmpB/xylE (catechol 2,3-dioxygenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 183

Table A4-B. Genes encoding for praC/xylH (oxalocrotonate tautomerase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 186

Table A4-C. Genes encoding for dmpH/xylI/nahK (gene 2-oxo-3-hexenedioate decarboxylase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 190

Table A4-D. Genes encoding for todC1/bedC1 (benzene/toluene dioxygenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 194

Table A4-E. Genes encoding for mhpE (4-hydroxy 2-oxovalerate aldolase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 195

Table A4-F. Genes encoding for mhpF (acetaldehyde dehydrogenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 202

Table A4-G. Genes encoding for dmpC, xylG (aminomuconate-semialdehyde/2- hydroxymuconate-6-semialdehyde dehydrogenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 208

Table A4-H. Genes encoding for dmpD/xylF (hydroxymuconate-semialdehyde hydrolase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 211

Table A4-I. Genes encoding for nahAc/ndoB (naphthalene 1,2-dioxygenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 212

Table A4-J. Genes encoding for nahC (dihydroxynaphthalene dioxygenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 212

ix Table A4-K. Genes encoding for nahD (hydroxychromene-2-carboxylate isomerase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 213

Table A4 L Genes encoding for nahE (trans-o-hydroxybenzylidenepyruvate hydratase- aldolase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 215

Table A4-M. Genes encoding for nahF (salicylaldehyde dehydrogenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 216

Table A4-N. Genes encoding for sal-hyd (salicylate hydroxylase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 216

Table A4-P. Genes encoding for dmpK/poxA (phenol hydroxylase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides...... 219

Table B1 A-G. The top 10 OTUs of each sample site based on percent (%) of total reads in 16S rRNA gene sequencing analysis. OTUs were clustered based on their closest related species...... 221

Table B1-A. Reads from DNA in WIP-OSPW sampled in August 2011...... 221

Table B1-B. Reads from DNA of the heavy SIP fraction (fraction 5) of OSPW sampled in August, 2011...... 222

Table B1-C. Reads from DNA of the heavy SIP fraction (fractions 3 and 4) of OSPW sampled in August, 2011, amended with labelled methanol for 7 days...... 223

Table B1-D. Heavy SIP Fractions (fractions 4 and 5) of OSPW sampled in August, 2011, amended with labelled sodium acetate for 5 days...... 224

Table B1-E. Heavy SIP Fractions (fractions 4 and 5) of OSPW sampled in August, 2011, amended with labelled protein extract for 14 days...... 225

Table B1-F. Heavy SIP Fractions (fractions 4 and 5) of MLSB-OSPW sampled in August, 2011, amended with labelled CO2 for 12 weeks under light...... 226

Table B1-G. Heavy SIP Fractions (fractions 4 and 5) of WIP-OSPW sampled in August, 2011, amended with labelled CO2 for 12 weeks under light...... 227

Table C1. Complete contig sequence from which the xmoCAB primer set was designed from. Sequence was obtained from the metagenome of WIP-OSPW sampled in 2011. Sequence length: 392 bp...... 228

x Table C2. KAPA HiFi HotStart ReadyMix PCR reaction protocol for Illumina amplicon sequencing...... 228

Table C3. Calculated concentration copies formulae...... 229

Table C4. A-E The top 10 OTUs of each sample site based on percent (%) of total reads in 16S rRNA gene sequencing analysis. OTUs were clustered based on their closest related species...... 230

Table C4-A. Reads from DNA in MLSB-OSPW sampled in August 2015...... 230

Table C4-B. Heavy SIP fraction (fraction 5 and 6) of MLSB-OSPW sampled in August 2011.230

Table C4-C. Heavy SIP fractions (fractions 4 and 5) of MLSB-OSPW sampled in August 2015, amended with labelled methane for 10 days...... 231

Table C4-D. Heavy SIP fractions (fractions 4 and 5) of MLSB-OSPW sampled in August 2015, amended with labelled ethane for 10 days...... 231

Table C4-E. Heavy SIP fractions (fractions 3 and 4) of MLSB-OSPW sampled in August 2015, amended with labelled propane for 10 days...... 232

xi List of Figures and Illustrations

Figure 1-1. Satellite image of the Mildred Lake Mine with MLSB and WIP tailings pond, from which samples for the research were obtained, outlined in red. Image taken from Google Earth (Public Domain Image) and the exact lake positions were obtained from Dompierre & Barbour (2016)...... 3

Figure 2-1. Illustrative map showing three major oilsands reserves regions: Peace River (Bluesky-Gething deposit), Cold Lake (Clearwater deposit) and Athabasca (Wabiskaw- McMurray deposit) (Public domain image; Alberta Energy, 2016)...... 8

Figure 2-2. A) Surface mining to recover oilsands located close to the surface. This process could only recover 20% of oilsands (Sierra-Garcia & Oliveira, 2013). B) In-situ mining of bitumen in Alberta. This system recovers around 80% of bitumen beneath earth surface. In this system, steam is used to heat the bitumen so that it can flow and then be pumped up to the surface (Public domain image; Government of Alberta, 2009)...... 10

Figure 2-3. A cross-sectional diagram of an oilsands tailings pond. (1) Discharge of tailings waste, (2) coarse sands, (3) mature fine tailings which contain more solid and heavy materials than the fluid fine tailings, (4) fine materials, (5) clear water rising to the top layer, (6) pumped water for reuse, and (7) oilsands process-affected water that is also recyclable (Modified from a public domain image; Government of Alberta, 2009)...... 12

Figure 2-4. Some examples of PAHs, including their chemical structures, physical characteristics, and solubility. The figure shows that the more benzene rings a compound has, the more recalcitrant it is (Cerniglia, 1992). Copyright permission is available in Appendix D...... 14

Figure 2-5. Two aerobic biodegradation pathways of benzene to form catechol via benzene dioxygenase (Gibson et al., 1970) and via benzene monooxygenase through a benzene epoxide intermediate (Madigan et al., 2015). The above diagram is adapted from a previous study (Mancini et al., 2003). Copyright permission is available in Appendix D... 19

Figure 2-6. Aerobic naphthalene degradation catalyzed by dioxygenases and dehydrogenases (Bamforth & Singleton, 2005). Copyright permission is available in Appendix D...... 20

Figure 2-7. Principle of the DNA-based stable isotope probing technique...... 27

Figure 2-8. In conventional metagenomics, DNA is extracted directly from environmental samples. As for the scheme of a DNA-based stable isotope probing (SIP) experiment, the sample uses ‘heavy’ fraction of DNA incorporated with 13C-labelled substrates. The DNA or ‘heavy’ DNA from SIP experiments can be further utilized for downstream analyses, such as pyrosequencing or metagenomics (Chen & Murrell, 2010). Copyright permission is available in Appendix D...... 32

xii Figure 3-1. Community structures of oilsands processed water from the WIP tailings pond (OSPW-control) and from the heavy, 13C-labelled DNA fractions of samples amended with 13C benzene (Benzene), naphthalene (Naphthalene), and methanol (Methanol), incubated under aerobic conditions. Predominant taxa are identified to the class level based on the SILVA database. The OSPW-Heavy treatment is the community detected only in the heaviest DNA fraction of OSPW. OSPW-Control shows the entire unseparated DNA from OSPW. The communities demonstrated that the heavy fractions retrieved from the SIP experiments were primarily determined by 13C enrichment of DNA rather than variabilities in genomic G+C content...... 52

Figure 3-2. Bubble chart of all samples based on relative abundances (>1%). Classification percentage is shown for the rank of genus (% classified). Benzene: heavy-DNA fraction of benzene-amended OSPW incubated for 9 d; Naphthalene: heavy-DNA fraction of naphthalene- amended OSPW incubated for 7 d; Methanol: heavy DNA fraction of methanol-amended OSPW incubated for 7 d; OSPW-heavy: heavy fraction of OSPW receiving the same treatment as all of the amended samples; and OSPW-control: complete, unfractionated DNA from OSPW...... 55

Figure 3-3 Aerobic hydrocarbon degradation pathways of benzene, naphthalene, and the corresponding genes detected in the OSPW metagenome based on KEGG pathways. (a) Detected pathways (substrates) for aerobic hydrocarbon metabolism and associated enzymes. (b) Bar chart of various genes and putative genes counts involved in aerobic hydrocarbon metabolism in the metagenome with >2500 bp scaffold length. Pathways descriptions: 1, meta-cleavage of catechol; 2, dioxygenase and dehydrogenase reactions; 3, ring removal from polycyclic aromatic ring; and 4, two monooxygenase reactions. Enzyme abbreviations: dmpB, xylE, catechol 2,3-dioxygenase; dmpD, xylF, hydroxymuconate-semialdehyde hydrolase; dmpC, xylG, aminomuconate- semialdehyde/2-hydroxymuconate-6-semialdehyde dehydrogenase; praC, xylH, oxalocrotonate tautomerase; dmpH, xylI, nahK, 2-oxo-3-hexenedioate decarboxylase; mhpD, 2-keto-4-pentenoate hydratase; mhpE, 4-hydroxy 2-oxovalerate aldolase; mhpF, acetaldehyde dehydrogenase; todC1, bedC1, benzene/toluene dioxygenase; nahAc, ndoB, naphthalene 1,2-dioxygenase; nahC, dihydroxynaphthalene dioxygenase; nahD, hydroxychromene-2-carboxylate isomerase; nahE, trans-o-hydroxybenzylidenepyruvate hydratase-aldolase; nahF, salicylaldehyde dehydrogenase; sal-hyd, salicylate hydroxylase; dmpK, poxA, phenol hydroxylase...... 59

Figure 4-1. Relative DNA versus DNA density for SIP experiments of WIP-OSPW tailings water sampled in August, 2011, with and without the addition of labelled substrates and model hydrocarbons: sodium acetate (A), methanol (B), protein extract (C), MLSB 13 13 amended with CO2 (D), and WIP amended with CO2 (E). Dashed lines represent incubations of OSPW control (no substrate addition) and solid lines represent incubations with 13C substrates. The y-axis indicates the relative DNA concentrations recovered from each fraction, where the highest quantity detected in any fractions is set to 1.0. Red arrows indicate the representative “heavy” fractions that were combined and used for 16S rRNA gene pyrotag sequencing to identify the active communities, and black arrows indicate “heavy” fraction of OSPW (OSPW-heavy) used in this study...... 78

xiii Figure 4-2. Community structures of oilsands process-affected water from the WIP tailings pond (OSPW-control) and from the heavy fraction (OSPW-heavy), 13C-labelled DNA fractions of samples amended with 13C-methanol (Methanol), 13C-sodium acetate 13 13 (Acetate), C-protein extract (Protein extract), CO2-MLSB (light incubation), and 13 CO2-WIP (light incubation), incubated under aerobic conditions. Predominant taxa are identified to the class level based on the SILVA database. The OSPW-heavy treatment is the community detected only in the heaviest DNA fraction of OSPW. OSPW-control shows the entire unseparated DNA from OSPW...... 79

Figure 4-3. Bubble chart of all 13C-labelled DNA from SIP heavy fraction samples analysed in this study based on relative abundances (>1%). Classification percentage is shown for genus rank (% classified). Heavy fraction of OSPW (OSPW-heavy), and unfractionated DNA from fresh OSPW (OSPW) were used as controls...... 89

Figure 4-4. Phylogenetic tree based on 16S rRNA gene sequences of the 5 most abundant OTUs of OSPW-control (green), 13C-methanol SIP-heavy fraction (red), 13C-acetate SIP 13 13 13 (dark blue), C-protein extract SIP (orange), CO2 in MLSB-OSPW SIP (blue), CO2 in WIP-OSPW SIP (black). To create the most reliable phylogeny, reference sequences (>1400 bp; black) from NCBI database (04/05/2016) were obtained. Tree was constructed via Maximum Likelihood with 1,000 bootstraps using Seaview Phylogeny (Gouy et al., 2010). For comparison, total number of reads of each OTU is given in parentheses (entire dataset was 49,186 reads). Scale bar represents 0.1 change per nucleotide position. Bootstrap values >50% are indicated for key nodes...... 92

Figure 5-1. Phylogenetic tree of xmoA genes that includes sequences detected in WIP-OSPW metagenome sampled in 2011, and other bacteria known to possess pmoA/amoA/xmoA. The tree was constructed via Maximum Likelihood algorithm in Seaview (Gouy et al., 2010), using 1,000 bootstraps as measure of support and the scale bar represents 0.2 change per nucleotide position. The xmoA gene of interest is depicted in red...... 103

Figure 5-2 Number of xmoCAB gene copies in MLSB-OSPW (per ml) sampled in September 2014, with and without the addition of hydrocarbon substrates: DNA from fresh MLSB (MLSB), MLSB amended with benzene (Benzene), MLSB amended with toluene (Toluene), MLSB amended with xylene (Xylene), and MLSB amended with naphthalene (Naphthalene). The x-axis indicates the different samples and incubation time. The y-axis indicates the number of xmoCAB gene copies detected per ml of sample. Bars indicate ±SEM of true replicate samples...... 104

Figure 5-3 Number of xmoCAB gene copies in MLSB-OSPW (per ml) sampled in March 2015, with and without the addition of short chain alkanes as substrates: DNA from fresh MLSB (MLSB), MLSB incubated for 6 weeks (MLSB), MLSB amended with methane (Methane), MLSB amended with ethane (Ethane), and MLSB amended with propane (Propane). The x-axis indicates the different samples and incubation time. The y-axis indicates the number of xmoCAB gene copies detected per ml of sample. Bars indicate ±SEM...... 106

xiv Figure 5-4 Community structures (based on class) of oilsands process-affected water from the MLSB tailings pond (MLSB) and from the heavy fractions of MLSB-OSPW (Control), 13C-labelled DNA fractions of samples amended with 13C-methane (Methane), 13C-ethane (Ethane), and 13C-propane (Propane) incubated under aerobic conditions. Predominant taxa are identified to the class level based on the SILVA database. The OSPW-heavy treatment is the community detected only in the heaviest DNA fraction of OSPW. OSPW-control shows the entire unseparated DNA from OSPW...... 109

Figure 5-5 Bubble chart of all 13C-labelled DNA from SIP heavy fraction samples analysed in this study based on relative abundances (>1%). Community structures (based on genus) of oilsands process-affected water from the MLSB tailings pond (MLSB-OSPW) and from the heavy fraction of MLSB-OSPW (MLSB-heavy), 13C-labelled DNA fractions of samples amended with 13C-methane (Methane), 13C-ethane (Ethane), and 13C-propane (Propane) incubated under aerobic conditions. Predominant taxa are identified to the class level based on the SILVA database. The OSPW-heavy treatment is the community detected only in the heaviest DNA fraction of OSPW. OSPW-control shows the entire unseparated DNA from OSPW...... 111

Figure 5-6 Relative DNA versus DNA density for SIP experiments versus number of xmoCAB gene copies of MLSB-OSPW tailings water sampled in August, 2015, with and without the addition of labelled substrates and model alkanes: 12C-methane (A), 13C- methane (B), 12C-ethane (C), 13C-ethane (D), 12C-propane (E), 13C-propane (F), and DNA from fresh MLSB (G). The x-axis represents DNA density (g/ml) of each SIP fraction. The y-axis (solid line) indicates the relative DNA concentrations recovered from each fraction, where the highest quantity detected in any fractions is set to 1.0. The bars indicate the number of xmoCAB gene copies detected per SIP fraction. Error bars represent ±SEM...... 114

Figure 6-1. (A) Gram staining picture of strain NVTT cells under bright field microscopy showing cocci and diplococci cell morphologies. (B) Transmission electron micrograph T (TEM) of strain NVT cells fixed in OsO4 solution. Uranyl acetate was used to increase image contrast. (C) Inset: Note the presence of granular vesicle inside the cell indicated by an arrow (lower right inset; bar, 100 nm)...... 129

Figure 6-2. Maximum Likelihood phylogenetic tree based on 16S rRNA gene sequences, showing the position of strain NVTT within the Verrucomicrobia phylum. Numbers at branch nodes are bootstrap values based on 1000 replications. The accession numbers of the 16S rRNA gene sequences of the representative strains in the tree are indicated in brackets. Numbers inside wedges shows the number of species in the closed groups. Rubritalea, Roseibacillus and Haloferula are genus names within the order Verrucomicrobiales. Tree was constructed via SeaView and aligned via ARB. Bar, 0.05 substitutions per sit...... 134

Figure A1. Stable isotope probing figures of WIP-OSPW tailings water sampled in August, 2011, with and without the addition of labelled substrates and model hydrocarbons: sodium acetate (A), methanol (B), benzene, and (C) naphthalene. Twelve SIP fractions

xv are shown as filled circles (amended with substrates) and empty circles (DNA from OSPW tailings water or control). Fractions pointed by red arrows were taken for DNA pyrosequencing as 13C-labelled samples, fractions pointed by black arrows were the control fraction 5 (OSPW-heavy) used in this study...... 167

Figure A2-A. Benzene oxidation of OSPW sampled in August, 2011, with the results of linear regressions for benzene oxidation...... 168

Figure A2-B. Headspace CO2 production time courses of replicate samples from OSPW sampled in August, 2011...... 168

Figure A3 A-B. Representative naphthalene oxidation and headspace CO2 production time courses of replicate samples from OSPW sampled in August, 2011, with the results of linear regressions...... 169

Figure A3-A. Naphthalene oxidation of OSPW sampled in August, 2011, with the results of linear regressions...... 169

Figure A3-B. Representative CO2 production time courses of replicate samples from OSPW sampled in August, 2011...... 169

Figure A4 A-B. Representative CO2 production time courses of replicate samples used for labelled hydrocarbons SIP of WIP-OSPW sampled in August, 2011...... 170

Figure A4-B. Representative CO2 production time courses of replicate samples used for labelled substrates SIP of WIP-OSPW sampled in August, 2011...... 170

Figure A5. Bubble chart of all samples based on relative abundances (>1%). Classification percentage is shown for class rank (% classified)...... 171

Figure A6. Protein coding genes statistics of WIP-2011 tailings pond metagenome based on its KEGG Categories analyzed on IMG...... 172

Figure A6 A-B. Metagenomic community structure of OSPW sampled in August, 2011, with >30% number of hits on IMG/GenBank database. Entire dataset of all the genes was 172,777 reads...... 173

Figure A6-A. Metagenomic community structure based on domains...... 173

Figure A6-B. Metagenomic community structure of OSPW based on the orders of Proteobacteria using entire dataset of all the genes detected...... 173

xvi Figure A7. Benzene degradation by pure cultures of Xanthobacter sp. OSPW28 and Zavarzinia compransoris sp. OSPW6, isolated from OSPW. Experiments were done in 2 replicates...... 174

Figure B1. Representative CO2 production time courses of 2 replicate samples used for labelled substrates SIP of WIP-OSPW sampled in August, 2011. Bars represent SD...... 220

Figure C1. Simplified workflow of this study. All MLSB-OSPW samples fed with 13C labelled methane, ethane, and propane, was monitored for CO2 production. Samples were extracted and analyzed by qPCR, followed by 16S rRNA amplicon sequencing. Picture was modified from a previous SIP figure (Friedrich, 2006)...... 233

Figure C2. PCR reactions gel images of inserted E. coli vector via gradient PCR annealing temperatures (48–58°C), using primers xmoCAB designed in this study. Gel band in red box is the brightest, thus selected as optimal annealing temperature...... 233

Figure C3. Representative methane oxidation time courses of replicate samples from MLSB tailings pond sampled in August 2015. Bars represent ±SEM...... 234

Figure C4. Representative ethane oxidation time courses of replicate samples from MLSB tailings pond sampled in August 2015. Bars represent ±SEM...... 234

Figure C5. Representative propane oxidation time courses of replicate samples from MLSB tailings pond sampled in August 2015. Bars represent ±SEM...... 235

Figure C6. Relative DNA versus DNA density for SIP experiments of MLSB-OSPW tailings water sampled in August, 2015, with and without the addition of labelled (13C) and 12 unlabelled ( C) alkanes: methane (A), ethane (B), and propane (C). Dotted lines represent incubations of OSPW control, dashed lines represent incubations with 12C substrates, and solid lines represent incubations with 13C substrates. The y-axis indicates the relative DNA concentrations recovered from each fraction, where the highest quantity detected in any fractions is set to 1.0. Red arrows indicate the representative “heavy” fractions that were combined and used for 16S rRNA gene pyrotag sequencing to identify the active communities, and black arrows indicate “heavy” fraction of OSPW (OSPW-heavy) used in this study...... 236

xvii List of Symbols, Abbreviations and Nomenclature

Symbol Definition amoA α-subunit of gene encoding ammonia monooxygenase BLAST Basic Local Alignment Search Tool bp base pair BTEX benzene, toluene, ethylbenzene, and xylenes

C10H8 naphthalene

C2H6 ethane

C3H8 propane

C5H7O2N representative for bacterial tissue

CH2O formaldehyde

CH3OH methanol

CH4 methane CHOO− formate CO carbon monoxide

CO2 carbon dioxide CsCl cesium chloride CuMMO copper-containing membrane monooxygenase dmpB/xylE gene encoding catechol 2,3-dioxygenase dmpC/xylG/praB gene encoding 2-hydroxymuconic semialdehyde dehydrogenase dmpD/xylF gene encoding 2-hydroxymuconate semialdehyde hydrolase dmpH/xylI/nahK gene encoding 2-oxo-3-hexenedioate decarboxylase dmpK/poxA gene encoding phenol hydroxylase DNA deoxyribonucleic acid G+C content guanine-cytosine content

xviii KEGG Kyoto Encyclopedia of Genes and Genomes GC-MS gas chromatography–mass spectrometry

H2O water

HCO3 bicarbonate HGD homogentisate 1,2-dioxyge HMN 2,2,4,4,6,8,8-heptamethylnonane kDa kilodalton (kDa) is 1,000 daltons mhpE gene encoding 4-hydroxy-2-oxovalerate aldolase mhpF gene encoding acetaldehyde dehydrogenase MLSB Mildred Lake Settling Basin MMO methane monooxygenase NA naphthenic acid NGS Next-Generation Sequencing

NH2OH hydroxylamine

NH3 ammonia + NH4 ammonium − NO2 nitrate

O2 molecular oxygen OSPW oilsands process-affected water OTU operational taxonomic unit PAH polycyclic aromatic compounds PCR polymerase chain reaction PLFA phospholipid fatty acid pMMO particulate methane monooxygenase pmoA α-subunit of gene encoding particulate methane monooxygenase pracC/xylH 4-oxalocrotonate tautomerase QIIME Quantitative Insights Into Microbial Ecology qPCR quantitative polymerase chain reaction rRNA ribosomal ribonucleic acid SEM standard error of the mean

xix SIP stable isotope probing sMMO soluble methane monooxygenase TCA tricarboxylic acid (citric acid) cycle TCD thermal conductivity detector todC1/bedC1 gene encoding benzene/toluene/chlorobenzene dioxygenase WIP West-In-Pit tailings pond of Syncrude Canada, Ltd.

Chapter One: Introduction

1.1 Research objectives

The Athabasca oilsands reserves, located in northeastern Alberta, Canada, contribute to more than 50% of crude oil production in Canada (Budgell, 2015). Production has escalated in the last decade due to global energy demand, and it is estimated that bitumen production could reach 5 to 6 million barrels per day by 2050 if demand persists (Rahnama et al., 2013). Every m3 of mined oilsands consumes up to 2m3 of water and produces on average 4m3 of tailings, consisting of oilsands process-affected water (OSPW), sand, clays, residual bitumen and dissolved inorganic and organic compounds (Holowenko et al., 2002; Quagraine et al., 2005).

Tailings are deposited into open ponds to allow settling of sand and clay particles, releasing

OSPW for reuse in the extraction process, and long-term containment of tailings for on-site reclamation processes (Government of Alberta, 2009).

It is estimated that polycyclic aromatic hydrocarbons (PAHs) and heterocyclic aromatic compounds are present at 0.01–10 mg kg-1 in fine tails in Alberta tailings ponds (Wayland et al.,

2008). Environmental concerns have been raised in regards to the potential of hydrocarbon exposure to the surrounding environments (Wayland et al., 2008). Oilsands mining operators, who are responsible for remediating the tailings ponds once they are permanently closed, have proposed several reclamation strategies, such as the ‘wet-landscape’ approach (Allen, 2008;

Wayland et al., 2008). In this strategy, the function of the ponds is reclaimed into End-Pit lakes or other wetland ecosystems, in which a layer of process and river water is used to cap the ponds at a specific depth in order to increase the depth of the oxic surface water layer, provide seed microbes and algae, and dilute contaminants (Allen, 2008; Wayland et al., 2008; Penner, 2016).

It is expected that such ponds will detoxify naturally over time, until colonized by aquatic plants.

A small-scale study of this reclamation method also showed that aeration and nutrient additions could stimulate indigenous microbial populations that help increase detoxification rates (Allen,

2008).

Aerobic metabolisms of indigenous microbial populations are known to be pivotal in the reclamation of industrially affected waters (Oller et al., 2011; Tocchi et al., 2012). Aerobic hydrocarbon degradation involves the addition of oxygen to an organic compound via oxygenase enzymes (Atlas, 1981; Díaz et al., 2013). A previous study detected high proportions of aerobic hydrocarbon-degrading bacterial genes in the metagenome of surface OSPW of the tailings ponds, indicating potential for in situ degradation activities (An et al., 2013a). Although residual hydrocarbon removal in reclaimed OSPW and other ponds occur naturally via biodegradation, little is known about the aerobic hydrocarbon degradation activities in the OSPW.

OSPW samples used in this thesis research were obtained from two active tailings ponds:

West In Pit (WIP) and Mildred Lake Settling Basin (MLSB), both located at the Athabasca oilsands reserves in Alberta, Canada, and owned by Syncrude Canada, Ltd. (Figure 1). WIP, now called Base Mine Lake (BML), was closed in 2012 and is currently being reclaimed. At the time of writing, MLSB is still an active pond, harbouring the largest volume of tailings in the area.

Water samples used in this study were obtained from the surface aerobic zone of the tailings ponds (0-10 cm from the surface).

In this thesis, aerobic communities involved in several key biogeochemical processes in

OSPW were studied by combining metagenomics, DNA based stable isotope probing (DNA-

SIP), high-throughput sequencing of 16S rRNA gene amplicons, and quantitative PCR methods.

Degradation of several hydrocarbons was also analyzed in depth using benzene and naphthalene

as models of aromatic hydrocarbons, and methane, ethane, and propane for model compounds for light alkanes.

Figure 1-1. Satellite image of the Mildred Lake Mine with MLSB and WIP tailings pond,

from which samples for the research were obtained, outlined in red. Image taken from

Google Earth (Public Domain Image) and the exact lake positions were obtained from

Dompierre & Barbour (2016).

The three major objectives of this dissertation were to:

1. Identify indigenous microbial communities responsible for the aerobic degradation of several key hydrocarbons, as well as other aerobic biogeochemical processes in OSPW.

2. Explore the diversity of hydrocarbon degrading genes in tailings water using metagenomics.

3. Cultivate strains indigenous to OSPW by using improved cultivation techniques, and test their hydrocarbon degradation potentials.

1.2 Dissertation structure

Chapter 1 of this dissertation gives a brief introduction into the study and its main objectives. Chapter 2 provides a detailed overview of the concepts associated with the topic, including background information, previous related studies, and theories of the methodologies used. In-depth background reviews, including methods, results, and discussions are included in every chapter of this dissertation.

Chapter 3 describes the aerobic bacterial communities within the OSPW that degrade benzene and naphthalene. In this study, metagenomic sequencing was performed on fresh WIP-

OSPW sample, while the techniques of DNA-SIP coupled with 16S rRNA gene sequencing were used to target active bacterial communities using 13C-labelled benzene and naphthalene.

In Chapter 4, the SIP technique was employed to study the indigenous aerobic bacterial communities of WIP-OSPW. Acetate is a major product of anaerobic degradation, and methanol is a byproduct of methane oxidation. To study these communities, 13C-labelled methanol and sodium acetate were added as substrates to WIP-OSPW samples. 13C-labelled protein extract was used to study indigenous proteolytic bacterial communities. Photosynthetic bacterial

13 communities were studied by adding CO2 into WIP-OSPW samples incubated under photosynthetic lights.

Chapter 5 explores the metagenome results of WIP-OSPW sample from the previous chapter (Chapter 3). Here, a novel CuMMO-encoding operon was detected in the sample, with the possibility of short-chain alkanes (C2-C3) oxidation. This gene was investigated in depth, using SIP, quantitative PCR, and RNA expression study.16S rRNA gene sequencing coupled

13 13 13 13 with SIP of C-alkanes ( CH4, C2H6, and C3H8) were employed to study levels of CuMMO operon in response to added substrates and survey the microbial community responsible to 13C- alkanes degradation.

Chapter 6 presents a thorough characterization of a novel bacterium from the phylum

Verrucomicrobia, isolated from WIP-OSPW, which was studied in detail. The bacterium was named Oleiharenicola alkalitolerans, which means an alkali-tolerating microorganism that inhabits oilsand, referring to the oilsands process-affected water from where the type strain was isolated. Genomic information revealed catE (catechol 2,3-dioxygenase) and praC/xylH (4- oxalocrotonate tautomerase) in the strain, which both are involved in catechol degradation pathways. A brief genome description for this strain is also provided as part of the chapter.

Chapter 7 ties all the major findings and general conclusions of this dissertation, providing a broad overview of what has been accomplished, and also directions for future research.

Additional co-authored research papers where the dissertation author was part of the projects are listed below. In the papers below, the author was involved in part of the taxonomical data analysis of the 16S rRNA gene amplicons data output. These papers are not included in the dissertation:

An D, Caffrey SM, Soh J, Agrawal A, Brown D, Budwill K, Dong X, Dunfield PF, Foght J, Gieg LM, Hallam SJ, Hanson NW, He Z, Jack TR, Klassen J, Konwar KM, Kuatsjah E, Li C, Larter S, Leopatra V, Nesbø CL, Oldenburg T, Pagé AP, Ramos EP, Rochman FF, SaidiMehrabad A, Sensen CW, Sipahimalani P, Song YC, Wilson S, Wolbring G, Wong ML & Voordouw G. (2013). Metagenomics of hydrocarbon resource environments indicates aerobic taxa and genes to be unexpectedly common. Environmental Science & Technology. 47:10708- 10717.

Saidi-Mehrabad A, He Z, Tamas I, Sharp CE, Brady AL, Rochman FF, Bodrossy L, Abell GC, Dong X, Sensen C and Dunfield PF. (2013). Methanotrophic bacteria in oilsands tailings ponds of northern Alberta. The International Society for Microbial Ecology Journal. 7(5):908-21. doi:10.1038/ismej.2012.163.

Chapter Two: Literature Review

2.1 Oilsands in Alberta

2.1.1 Background

The Athabasca oilsands reserves, located in northeastern Alberta, Canada, contribute to more than 50% of crude oil production in Canada (Nedashkovskaya et al., 2015). Production has escalated in the last decade due to global energy demand. It is estimated that bitumen production could reach 5 to 6 million barrels per day by 2050 if demand persists (Rahnama et al., 2013), although the availability of shale oil and alternative fuels may depress this demand. Oilsands contain a highly viscous crude oil within an unconsolidated matrix of sand, clay, and water, referred to as bitumen (Government of Alberta, 2008; Cook, 1967; Cutler, 2004). They can be found in vast amounts worldwide, in countries such as Venezuela, Trinidad and Tobago, Canada, and Russia (Bradshaw et al., 2015). Covering over 140,000 km2, the Athabasca oilsands is the world’s third largest oilsands deposit, after Saudi Arabia's conventional oil and Venezuela's

Orinoco oilsands, containing an estimated 2.5 trillion barrels of total reserves (Government of

Alberta, 2008; Penner & Foght, 2010). This unconventional oil resource can be divided into the

Peace River (Bluesky-Gething deposit), Cold Lake (Clearwater deposit) and Athabasca

(Wabiskaw-McMurray deposit) regions (Figure 2-1) (Honarvar et al., 2011).

2.1.2 Oilsands recovery

Recovery of conventional oil is regularly done by drilling oil wells into natural reservoirs and letting the oil flow up to the surface through different techniques (Ancheyta et al., 2005), but unconventional oil sources need to be extracted and processed using a more sophisticated and

laborious technique. Since oilsands are ‘heavy’, their viscosity needs to be reduced so they can be refined further. Two techniques to extract bitumen in Alberta are: 1) surface mining, and 2) injecting steam or solvents into the sands, called in-situ mining (Figure 2-2) (Ancheyta et al.,

2005). The Athabasca oilsands reserves contain approximately 82% of the total distributed bitumen, and only 20% is recoverable through surface mining (Carrera-Hernández et al., 2012;

Peacock, 2010).

Figure 2-1. Illustrative map showing three major oilsands reserves regions: Peace River

(Bluesky-Gething deposit), Cold Lake (Clearwater deposit) and Athabasca (Wabiskaw-

McMurray deposit) (Public domain image; Alberta Energy, 2016).

Surface mining, a method only feasible for the shallow oilsands deposits (65 metres in depth or less), accounts for 52% of bitumen production in Alberta (Pembina Institute, 2008). In surface mining operations, bituminous oilsands are mined using trucks and shovels before being transported into a preparation plant, where the oilsands are crushed and combined with water.

Bitumen-water slurry is then transported to bitumen extraction plants, where water and sand are finally being separated from the bitumen (Figure 2-2) (Alberta Energy, 2016; Pembina Institute,

2008). The surface mining, or in situ method, require substantial energy and water, and are more expensive compared to surface mining (Dyer & Huot, 2010).

The Canadian Association of Petroleum Producers (CAPP) projects that in situ mining activities will surpass that of surface mining by 2017, as research develops more efficient oilsands recovery technologies (CAPP, 2011; Dyer & Huot, 2010). Currently, the Steam Assisted

Gravity Drainage (SAGD) is the most commonly used in situ recovery method (Figure 2-2). This method requires two horizontal wells, one above another, in which heated steam is injected into one well, causing the bitumen reservoir to melt and flow downward to the second well (Dyer &

Huot, 2010; Giacchetta et al., 2015; Irani & Ghannadi, 2013). Later, water is injected to maintain stability after the bitumen is removed.

A B

Figure 2-2. A) Surface mining to recover oilsands located close to the surface. This process

could only recover 20% of oilsands (Sierra-Garcia & Oliveira, 2013). B) In-situ mining of

bitumen in Alberta. This system recovers around 80% of bitumen beneath earth surface.

In this system, steam is used to heat the bitumen so that it can flow and then be pumped up

to the surface (Public domain image; Government of Alberta, 2009).

2.2 Oilsands tailings ponds

As a result of the extraction processes, bitumen is upgraded further, and tailings are deposited into tailings ponds. Oilsands-derived bitumen is upgraded into synthetic crude oil

(SCO) to reduce viscosity so they are suitable for production (Charpentier et al., 2009). Through this process, unwanted substances are eliminated. When tailings are deposited, sand particles quickly settle out to the bottom of the pond, releasing pore water, whereas clays form a fairly stable suspension (fine tailings) that will take decades or centuries to fully consolidate.

2.2.1 Oilsands process-affected water (OSPW)

Although both oilsands extraction operations consume substantial amounts of water, the water use in surface mining is twice as high as in situ (Pembina Institute, 2008). On average, in situ operations use 1.1 barrels of water for every barrel of bitumen produced, and surface mine uses 2.1 barrels of water for each barrel of bitumen produced (a barrel is equivalent to 159 litres)

(Pembina Institute, 2008). Produced tailings are deposited into open tailings ponds to allow settling of particles, reuse of surface water for extraction, and long-term pollutant containment for on-site reclamation processes (Government of Alberta, 2009). All mining operations draw water primarily from the nearby Athabasca River or recycle water from the surface layer of tailings ponds (Cowie et al., 2015).

In the Athabasca oilsands bitumen, long chain or cyclic aromatic carboxylic acids content is approximately 2% and about 90% primarily make up the naphthenic acid (NA) fraction

(Quagraine et al., 2005). NAs are soluble, therefore it partitions into the water and are deposited into the tailings ponds, causing toxicity and potential health risks to surrounding terrestrial ecosystems (Ancheyta et al., 2005; Penner & Foght, 2010; Quagraine et al., 2005; Rogers et al.,

2002b; Penner, 2016). Oilsands process-affected water (OSPW) in the ponds contains organic contaminants such as polycyclic aromatic hydrocarbons (PAHs) and a complex mixture of aliphatic and alicyclic carboxylic acids, mostly composed of NAs. The toxicity of the tailings ponds is attributed to NAs and other contaminants such as BTEX, ammonia, and polyphenols

(Allen 2008; Zhuo et al., 2012).

Figure 2-3. A cross-sectional diagram of an oilsands tailings pond. (1) Discharge of tailings

waste, (2) coarse sands, (3) mature fine tailings which contain more solid and heavy

materials than the fluid fine tailings, (4) fine materials, (5) clear water rising to the top

layer, (6) pumped water for reuse, and (7) oilsands process-affected water that is also

recyclable (Modified from a public domain image; Government of Alberta, 2009).

2.2.2 Layers of the tailings ponds

The tailings ponds consist of a thin oxic surface water layer and an extensive lower anoxic and methanogenic layer, releasing CO2 and CH4 (Holowenko et al., 2000). Methane gas production occurs at around 15 m depth and below, causing a serious concern to the environment. A study suggested that methane gas emission from a tailings pond, Mildred Lake

3 -1 Settling Basin (MLSB), is estimated to reach up to 43,000 m CH4 day (Holowenko et al.,

2000; Siddique et al., 2008).

Using the current water extraction methods to extract the bitumen, every barrel of oil produced needs 2.1 barrels of water to process, most of which is recycled. Thus, the size of tailings ponds keeps expanding, currently covering an area of approximately 88 km2 (Alberta

Energy, 2016). The slurry wastes from the extraction processes within the tailing ponds could take from decades to centuries to settle (Barton & Wallace, 1979; Quagraine et al., 2005). Finer solids remain in a floating layer forming the Fluid Fine Tailings (FFT) layer, while sand particles sink and settle to form the mature fine tailings (MFT) (Holowenko et al., 2000). Through time, densification allows the overlying layer of water to be reused in the oilsands extraction processes

(Government of Alberta, 2009). A cross-sectional diagram of the aforementioned layers is simplified in Figure 2-3.

2.3 Petroleum hydrocarbons

Petroleum is made up of a mixture of various molecular weights hydrocarbons and other organic compounds (Olah & Molnár, 2003; McCain, 1990). They are by far the most encountered pollutants in water and soil environments (Das & Chandran, 2011; Holliger et al.,

1997). Petroleum hydrocarbons can be classified as either saturated (e.g. alkanes) or unsaturated

(e.g. alkenes and alkynes) (Radke et al., 1980).

Figure 2-4. Some examples of PAHs, including their chemical structures, physical

characteristics, and solubility. The figure shows that the more benzene rings a compound

has, the more recalcitrant it is (Cerniglia, 1992). Copyright permission is available in

Appendix D.

2.3.1 Hydrocarbons in the tailings ponds

In general, OSPW is alkaline (pH 7.8 – 8), with high contents of salts (2.2 g L-1), various metals, sulfides, naphthenic acids (NAs), and polycyclic aromatic hydrocarbons (PAHs) (Kelly et al., 2010; Kelly et al., 2009; Quagraine et al., 2005; Saidi-Mehrabad et al., 2013). Other major organic contaminants in the tailings are derived from naphtha (a mixture of C3−C14 alkanes,

BTEX (benzene, toluene, ethylbenzene, and xylenes), and iso-parrafins) that is used as a diluent

in processing of bitumen from oilsands, and other low molecular weight iso-paraffins and naphthenes (Siddique et al., 2007). In particular, NAs and PAHs are compounds that contribute to the toxicity of the tailings ponds, and thus their presence and degradation have been extensively studied (Allen, 2008; Grewer et al., 2010; Han et al., 2009; Quagraine et al., 2005;

Rogers et al., 2002a; Scott et al., 2005; Wayland et al., 2008). It is estimated that PAHs and heterocyclic aromatic compounds account for 0.01–10 mg kg-1 of fine tails in Alberta oilsands tailings ponds (Wayland et al., 2008).

2.3.2 Polycyclic aromatic hydrocarbons

Out of the various groups of hydrocarbons, PAHs are one of the major concerns. PAHs refer to various chemical compounds having two or more fused benzene rings (Figure 2-4)

(Arulazhagan & Vasudevan, 2011; Eljarrat & Barceló, 2004). PAHs are listed as priority pollutants by the United States Environmental Protection Agency for their environmental persistence, and mutagenic and carcinogenic effects (Cerniglia, 1992). The binding of PAHs metabolic intermediates with DNA, RNA, and albumins could activate cellular carcinogenicity

(Pashin and Bakhitova, 1979). There is a positive link between cellular metabolic activity and the covalent bonding of the PAHs with DNA. Epoxide intermediates, formed via PAH degradation, can be incorporated into a DNA molecule and affect the insertions in DNA replication process and repair mechanism (Pashin and Bakhitova, 1979). Two major sources of

PAHs in the environment are from industrial and fuel combustion byproducts (Kasperski &

Mikula, 2011). Their persistence comes from the benzene rings; in general the more rings they have the more difficult they are to break down (Figure 2-4) (Peng et al., 2008).

Environmental concerns have been raised about the potential movement of these compounds into surrounding environments (Wayland et al., 2008). A recent study showed that the increase of oilsands production have contributed to the increase of PAHs to the surrounding environment (Kelly et al., 2009). The closest water bodies prone to contamination of the aforementioned compounds are the Athabasca river and its tributaries, with PAHs contamination reaching up to 4.8 μg/L of its dissolved concentration as ice melts in the spring (Kelly et al.,

2009). Previous studies on biodegradation of these compounds have emphasized degradation in anoxic conditions, which dominate in the tailings ponds, and relatively little is known about activities in the more oxic surface water cap (10 cm deep) of the ponds, from which recycle water is drawn. This surface layer is weakly oxygenated and supports some aerobic microbial activities such as methane oxidation (Saidi-Mehrabad et al., 2013).

2.4 Microbial hydrocarbon degradation in OSPW

Challenges to the oilsands industry include developing a more sustainable way of extracting bitumen, reducing the production of tailings, and promoting an effective waste treatment process. In environmental microbiology, one of the challenges posed is to find the microorganisms responsible for certain processes. In the context of the tailings, this means finding the bacteria responsible for biodegrading toxic components (Quagraine et al., 2005).

Bioremediation itself refers to the breaking down of contaminants into less-harmful substances

(Margesin & Schinner, 2001). The ability of microorganisms to utilize hydrocarbons has been known since the 1940s (Atlas, 1981). Although the main concern has usually been on microbial roles in degrading various marine petroleum spills, knowledge on the subject matter has expanded into freshwater, soil, and waste ecosystems (Atlas, 1981).

2.4.1 Aerobic hydrocarbon degradation

The presence of genes for anaerobic and aerobic degradation of aromatic hydrocarbons in several oilsands related samples from Alberta, Canada, has been previously described (An et al.,

2013b). High proportions of aerobic bacterial species were detected in the surface OSPW of various tailings ponds (>60% of all detected 16S rRNA genes). Genes encoding aerobic hydrocarbon-degradation enzymes such as dioxygenases, monooxygenases, and extradiol ring- cleavage enzymes were abundant in metagenomes of OSPW (An et al., 2013b; Saidi-Mehrabad et al., 2013), showing a diversity of hydrocarbon degradation mechanisms in the oxic layers of the ponds. Indigenous aerobic microbial communities in the tailings ponds have been shown to oxidise methane (Saidi-Mehrabad et al., 2013) and naphthenic acids (Herman et al., 1994), and may have potentials in biofilm technology applications for bioremediation (Golby et al., 2012).

Aerobic metabolisms of indigenous microbial populations are known to be pivotal in the reclamation of industrially affected waters (Oller et al., 2011; Tocchi et al., 2012). Previous studies have examined the significance of hydrocarbon degradation under aerobic conditions

(Atlas, 1991; Das & Chandran, 2011; DeLaune et al., 1980; Prince et al., 2010). Rapid and complete biodegradation in aerobic conditions has led to various waste management applications to help reduce the levels of` contaminants in different sites of interest such as in industrial wastewater.

Understanding aerobic degradation processes in OSPW is valuable because some of the proposed reclamation strategies for tailings ponds involve increasing the depth of the oxic water cap. One of the reclamation strategies, the wet-landscape approach, proposes the conversion of tailings ponds to End-Pit Lakes or other wetlands, in which natural detoxification would rely

mostly on aerobic processes (Allen, 2008; Wayland et al., 2008). In this approach, fine tailings would be transferred into an abandoned mined-pit, with a layer of fresh water placed on top, establishing a water-cap over the fine tailings base, thus allowing fine tailings to settle over time, and creating end pit lakes (Zhao et al., 2012; Penner, 2016).

2.5 The oxygenases

2.5.1 Oxygenases

The first step of aerobic hydrocarbon biodegradation is an initial attack by oxygenases or peroxidases, which is initiated by the incorporation of an oxygen molecule into the substrate

(Cerniglia, 1992). The first such enzyme was discovered in 1955 by two separate groups, led by

Osamu Hayaishi from Japan, and Howard Mason from USA. An in-depth study was carried out by Hayaishi (2005), describing the distinctive properties of this enzyme. From his studies, it was discovered that: 1) this enzyme does not contain heme, flavin, or other redox cofactors, 2) oxygen atoms incorporated into the substrate were derived from air, and 3) oxygenases can be found in diverse microorganisms, plants and animals.

Two types of oxygenases were then discovered: 1) monooxygenases, which was then termed ‘mixed function oxidase,’ transfer one oxygen atom from molecular O2 to the substrate, and consequently reduce the other oxygen atom to water, and 2) dioxygenases, or ‘oxygen transferases’, incorporate both atoms of molecular oxygen (O2) into the product(s) of the reaction

(Bugg, 2003).

Figure 2-5. Two aerobic biodegradation pathways of benzene to form catechol via benzene

dioxygenase (Gibson et al., 1970) and via benzene monooxygenase through a benzene epoxide intermediate (Madigan et al., 2015). The above diagram is adapted from a previous

study (Mancini et al., 2003). Copyright permission is available in Appendix D.

2.5.2 Aromatic hydrocarbon degradation via oxygenases

Simplified in Figure 2-5 and 2-6, hydrocarbon degradation involves breaking down organic contaminants into precursors of central metabolic pathways, i.e. the citric acid cycle

(TCA Cycle) (Das & Chandran, 2011). Monooxygenase enzymes could either attack methyl- or ethyl-substituents of aromatic rings, followed by a number of oxidations to form catechol, or alternatively, one oxygen atom may be incorporated into the aromatic ring while the second atom is used to oxidise either the aromatic ring or a methyl group (Jindrová et al., 2002). Figure 2-6 shows the biodegradation pathway of naphthalene, a type of PAH, by a number of enzymes.

Intermediate compounds shown in the pathway are gentistic acid and catechol, which would then

proceed to the TCA Cycle to produce CO2 (Bamforth & Singleton, 2005). Naphthalene is most

possibly mineralized via dioxygenase reactions, in which dioxygenase attacks aromatic rings to

form 1,2-dihydroxy-naphthalene compounds (Figure 2-6) (Jindrová et al., 2002). Through the

cycle, aerobic oxidation would produce CO2 and water (Figure 2-5 and 2-6) (Rich, 2003). 2‐rin ged PAH: Naphthalene

TCA cycle

Bamforth and Singleton, 2005, J. Chem. Technol. Biotechnol. 80: 723‐736 Figure 2-6. Aerobic naphthalene degradation catalyzed by dioxygenases and

dehydrogenases (Bamforth & Singleton, 2005). Copyright permission is available in

Appendix D.

2.5.3 Catechol dioxygenases

A key step in the degradation of aromatic compounds is the oxidative cleavage of catechol and substituted catechols. Two families of dioxygenase enzyme in charge of this activity are the intradiol and extradiol dioxygenases (Hayaishi, 2005; Bugg, 2003). The intradiol dioxygenases, represented by catechol 1,2-dioxygenase, break down carbon bonds between the catechol hydroxyl groups using Fe3+ as a cofactor to yield muconic acid. The extradiol dioxygenases, represented by catechol 2,3-dioxygenase, break the carbon bonds adjacent to the catechol oxygens, also with Fe2+ as a cofactor, yielding 2-hydroxymuconaldehyde as the product

(Hayaishi, 2005; Bugg, 2003; Lin et al., 2001). Catechol 2,3-dioxygenase is critical in the breakdown of toluene, benzoate, and their methyl derivatives via the meta-cleavage pathway

(Saunders et al., 1996). This enzyme breaks down catechol to 2-hydroxymuconic semialdehyde

(Saunders et al., 1996; Winstanley et al., 1991), and its gene is used as a marker to assess several microorganisms, such as Pseudomonas putida and Streptomyces lividans populations in the environment (Prosser, 1994; Winstanley et al., 1991).

2.5.4 Aerobic alkane oxidation

Alkanes are saturated hydrocarbons with the general chemical formula of CnH2n+2 (Smith,

2011). The most important commercial sources for alkanes are natural gas and crude oil, although many living organisms also produce them for protection and survival mechanisms

(Thom et al., 2007). Some microorganisms are known to be capable of metabolizing alkanes, utilizing them for both carbon and energy sources (Rojo, 2010).

Aerobic alkane degradation begins by the oxidation of a terminal methyl group into an alcohol, which is oxidized to aldehyde, converted into a fatty acid, and CoA, to be further

processed by β-oxidation to generate acetyl-CoA (Rojo, 2010; Van Agteren et al., 2013). A number of enzyme families in some bacterial strains can be related to the initial alkane hydroxylation (Table 2-1). Microorganisms degrading short-length alkanes (C2–C4) contain enzymes related to soluble di-iron methane monooxygenases, whereas other strains degrading medium-length alkanes (C5-C9) and long-chain-length alkanes (>C10) tend to contain the non- heme iron monooxygenases (Rojo, 2010).

Table 2-1. Enzyme classes related to alkane oxidation (Rojo, 2010). Copyright permission is available in Appendix D.

One enzyme family known to be present in alkane degrading bacteria is non-heme alkane hydroxylases, encoded by alkB. This gene has been used as a marker for alkane-degrading communities in the environment (Rojo, 2010). alkB can be associated with strains such as

Alcanivorax, Marinobacter, and Pseudomonas (Smith et al., 2013). Pseudomonas putida strain

GPo1 contain alkB that allows oxidation of propane, n-butane, as well as of C5–C13 alkanes

(Johnson and Hyman, 2006).

2.6 Study of hydrocarbon-degrading bacteria in the environment

2.6.1 Activity measurements to monitor microbial activity

Microbial activities of an environmental sample can be monitored by either the biodegradation rate of the pollutant or gas production rate (Boonchan et al., 2000; Healy &

Young, 1978). Indicators for hydrocarbon degrading activities include: the amount of compounds degraded, production of CO2 for aerobic samples, and production of H2S or CH4 from anaerobic samples. Various studies have quantified consumption of aromatic compounds and gas production periodically using gas chromatographs (GCs) equipped with a flame ionization detector, high-performance liquid chromatographs (HPLC), and other gas partitioning techniques (Boonchan et al., 2000; Healy & Young, 1978; Kästner & Mahro, 1996; Volkering et al., 1992). A study on aerobic degradation by Pseudomonas putida strain CSV86 used oxygen uptake rate as an indication of microbial activities. In addition to that, degradation rate of substrate compounds such as succinate and glucose could also measured to indicate metabolic capacity (Nigam et al., 2012).

2.6.2 Enrichments: growing target organisms

One common method to see if indigenous microorganisms can grow on target compounds in a specific environmental site is by setting up enrichment samples, by adding target compounds to a closed environmental microcosm (Leahy & Colwell, 1990a). This method has been

successful in growing both aerobic and anaerobic hydrocarbon-degrading microorganisms in laboratories (Berry et al., 1987; Fedorak & Westlake, 1984; Kuhn & Suflita, 2009; Meade et al.,

2002). Enrichment samples are set up by enriching a subset of an environmental sample in a microcosm, and improving its limiting nutrients and growth conditions in the laboratory (Li et al., 2000). This is to increase the number or hydrocarbon-degrading bacteria, and promote the enzymatic capacity of the resident microbial communities (Li et al., 2000). In this method, indigenous bacterial communities are grown in a laboratory system with hydrocarbon(s) of interest as their carbon source(s). Over time, the system will contain mostly ‘selected’ bacteria with the ability to breakdown the target hydrocarbon(s) (Atlas, 1991; Edward, 1993; Leahy and

Colwell, 1990; Robert, 1992).

Classic enrichment methods have been used to study both aerobic and anaerobic bacterial communities (Atlas, 1981). Different enrichment methods have been utilized to grow selected communities, such as the sequential enrichment method, to isolate organisms that could utilize progressively more complex compounds (Horowitz et al., 1975), and enrichment culture coupled with a bioreactor to investigate microbial communities originating from deep-sea hydrothermal vents (Postec et al., 2005). Enriched communities tend to respond differently to extrinsic factors compared to bacterial communities in situ, as bacterial communities adjust themselves to laboratory systems. Finally, enrichment method been one of the main methods utilized in the study of microbial hydrocarbon degradation over the past decades (Chang et al., 2000; Eriksson et al., 2003; Laban et al., 2015; Schink, 1985; Tan et al., 2015).

2.6.3 Culture-dependent methods: Isolation of microorganisms from oil-contaminated environments

The cultivation-based method in microbiology is a traditional way to understand microorganisms. The idea of this technique is to grow microorganisms in pure cultures in laboratories to study them (Sagaram et al., 2009). This could help determine the morphological, metabolic, and physiological characteristics of the species. Pure cultures are obtained from serial dilutions of sample enrichments (Cowan, 2011). However, this method is deemed difficult because cultivating a targeted organism can be hindered by slow growing time, ability to grow on selective laboratory media, competition with other organisms during incubation, and understanding of the conditions needed for growth that would mimic those in natural environments (Ellis et al., 2003; Prosser & Embley, 2002).

Biodegradation capabilities of various strains can also be further studied using combinations of pure and mixed cultures (Der Yang & Humphrey, 2004; Hanson et al., 1999;

Mo et al., 1997). Studies of the pure bacterial cultures have led to a further confirmation of their taxonomic groupings by using information from their genomic sequences and phenotypic characteristics (Joseph et al., 2003). This information became useful to further explain their distribution in natural environments and how they may respond to environmental changes

(Prosser & Embley, 2002).

2.6.4 Culture-independent method: Next-generation sequencing to study oil-contaminated environments

Growing microbial strains is not an easy task. Because of their specificity, current cultivation methods have only been successful to obtain around 0.1-1% of all microbial species

(Sagaram et al., 2009). Therefore, microbiological studies in this past decade have turned to cultivation-independent methods (Ellis et al., 2003).

16S ribosomal RNA (16S rRNA) gene sequences are the most common genetic marker to identify bacteria. The gene is highly conserved and present in all species of bacteria and archaea

(Janda & Abbott, 2007). 16S rRNA amplification of pure environmental DNA is one of the most definitive tools of microbial identification (Janda & Abbott, 2007). 16S rRNA gene sequencing has been used to identify bacteria as pure cultures, or as a whole community (Ward et al., 1990;

Will et al., 2010).

2.6.4.1 DNA based stable isotope probing

Stable isotope probing (SIP) is one of the cultivation-independent tools that have been widely used to understand microbial function. The purpose of this method is to link the identity of specific microorganisms with their function in an environment (Radajewski et al., 2000). In this method, it is possible to identify targeted microbial communities by adding a 13C-enriched carbon source in the enrichments. Microbial communities responsible in taking up the labelled substrates will incorporate 13C to their DNA (13C-DNA or ‘heavy’ DNA) (Radajewski et al.,

2000). After incubation, 13C and 12C-DNA are separated by cesium chloride (CsCl) density- gradient centrifugation into several fractions (Dumont & Murrell, 2005). The 13C ‘heavy’ fraction contains only DNA from species capable of assimilating the 13C as a carbon source, in which they are then incorporated into the DNA during DNA synthesis and replication (Neufeld et al., 2007) (Figure 2-7).

SIP as a tool in microbial ecology has been used to identify important microbial communities as early as 2000 (Radajewski et al., 2000). Since then, it has been frequently used

in identifying pollutant degraders in various environments. SIP analysis of salicylate-, naphthalene-, and phenanthrene-degrading microbial communities in a bioreactor treating contaminated soil detected primarily sequences belonging to the genera Pseudomonas, Ralstonia, and Acidovorax (Singleton et al., 2005). DNA based SIP also revealed groups of microorganisms not previously thought to contribute to methane, ethane, or propane oxidation, i.e. the family

Methylococcaceae, Methylophaga, and Methylophilaceae (Redmond et al., 2010). Other SIP studies also discovered novel uncultured microorganisms, or assigned new roles to groups of organisms both from aerobic (Gutierrez et al., 2011; Singleton et al., 2007; Xie et al., 2011) and anaerobic environmental samples (Jehmlich et al., 2008; Pilloni et al., 2011).

Figure 2-7. Principle of the DNA-based stable isotope probing technique.

SIP also enables the study of microbial communities that have specific functions, as it eliminates unnecessary data generated from sequencing of the whole community (Röling et al.,

2010). Some limitations of SIP are the dilution of labelled substrate before being incorporated, length of incubation time, and assimilation of intermediates by non-target organisms or cross- feeding (Radajewski et al., 2003; Radajewski et al., 2000). Dilution of labelled substrate, such as

13C, with the same unlabelled natural substrates, 12C, will make it difficult to target the main degraders. The length of incubation time relates to cross-feeding. Incubation time should be long enough to allow for the incorporation of a labelled substrate into DNA of target organisms, but not too long to cause the incorporation of labelled substrates into products and intermediates of microbial metabolism, which may be assimilated by non-target organisms (Radajewski et al.,

2003).

While SIP has its own benefits and limitations, this method can be modified based on needs. In addition to incubation combinations and the use of a long-term SIP which is known as

SIS (stable isotope switching), which is the simultaneous assessment of carbon uptake, turnover, and decay (Maxfield et al., 2012). Other studies have used multiple forms of SIP such as

13 18 combining C- mannitol with H2 O to see moisture effects on diversity (Saurey et al., 2012) and the integration of SIP with metagenomics and metaproteomics (Seifert et al., 2012; Uhlik et al., 2012).

2.6.4.2 16S rRNA gene sequencing

Although the cultivation approach is probably the best way to study microorganisms, it is limited by the difficulties of growing them. An estimate suggests that 99% of active natural bacteria are not cultivated using common cultivation-based methods and only the remaining is

cultivable (Aslam et al., 2010; Lee & Fuhrman, 1990; Pace, 1997). Comparison of cultivated microbes is also difficult for closely related species if it is based merely on the observable phenotypic characters (Hugenholtz & Pace, 1996). This problem has hindered the study of natural diversity of environmental samples as most microorganisms are still uncultured (Prosser

& Embley, 2002). Due to these limitations, scientists have turned to molecular methods as attempts to understand community structures.

Sequencing technology progressed from the 1980s when the first semi-automated DNA sequencing machines became available, to the 1990s when the concept of Next-Generation

Sequencing (NGS) methods were first introduced (Ansorge et al., 1986; Smith et al., 1998). But it was not until the year 2000 that NGS methods became commercially available (Mardis, 2008).

Pyrosequencing, a pyrophosphate sequencing concept which was first introduced by Hyman

(1988), was the first NGS technique to be available commercially (Hyman, 1988; Ronaghi, 2001;

Shendure & Ji, 2008). Commonly known as 454 pyrosequencing, based on the Roche/454 sequencer machine, the idea of this technique is to detect the release of pyrophosphate group after incorporation of a nucleotide base by DNA polymerase enzyme. The pyrophosphate can be detected as emitted light signals by luciferase enzyme, which indicates which nucleotide bases is incorporated throughout the DNA strand (Ansorge, 2009; Mardis, 2008).

In order to produce enough DNA copies for sequencing, such as via Roche/454 and

Illumina, DNA extracted from laboratory cultures or natural samples needs to be amplified to a certain number of DNA copies. With PCR amplification methods, a single or a few copies of DNA strands can be copied into several orders of magnitudes, generating thousands to millions of copies of the targeted DNA sequences (Wilson et al., 1990). Many studies have used

PCR primers which target 16S rRNA genes to analyze microbial diversity and functions in various environments.

The Illumina Genome Analyzer sequencing technique is based on sequencing by synthesis, in which four DNA bases, labelled with different fluorescent dyes, are present in every sequencing cycle, allowing base-by-base sequencing. To determine sequences, incorporation of

DNA and terminator nucleotide is identified via the fluorescent dyes (Ansorge, 2009).

Undoubtedly, while NGS methods produce high-throughput reads, their read lengths are limited of around 250 – 600 bp of the 1.5 kbp long 16S rRNA gene (Burke & Darling, 2014; Chaisson &

Pevzner, 2008).

16S rRNA gene analysis is used best to study the phylogeny of microbial communities, which is done by analyzing the total DNA from the environment. Analysis of microbial communities will provide information on all taxa that are present in the sample, which have helped in identifying uncultured microorganisms (Torsvik & Øvreås, 2002; Ward et al., 1990).

To classify groups of closely related individuals, groups of organisms are classified as operational taxonomic unit (OTU), based on DNA sequence similarity of a specific taxonomic marker gene (Blaxter et al., 2005).

2.6.4.3 Metagenomics

The rapid development of NGS makes it possible to further investigate microbial communities from the sequencing of 16S rRNA gene to fragments of the whole genome without the need for culturing (Cowan et al., 2005). The genomic analysis of microorganisms within an environmental community is termed as metagenomics (Cowan et al., 2005; Handelsman, 2004).

Different from 16S rRNA gene sequence, this approach targets the whole genome to perform a

deeper study of the organisms. Metagenomics unravels microbial genetic information, which can be further used to study the genome structure, gene function, population genetics, potentially novel enzymes, phylogenetic genomic linkages, lateral gene transfers, and evolutionary profiles

(DeLong, 2004). Information from metagenomic sequences may also facilitate a broader understanding of improved culturing strategies to further link genomic information to phenotypic characteristics of pure cultures (Handelsman, 2004; Thomas et al., 2012).

Figure 2-8. In conventional metagenomics, DNA is extracted directly from environmental

samples. As for the scheme of a DNA-based stable isotope probing (SIP) experiment, the

sample uses ‘heavy’ fraction of DNA incorporated with 13C-labelled substrates. The DNA

or ‘heavy’ DNA from SIP experiments can be further utilized for downstream analyses, such as pyrosequencing or metagenomics (Chen & Murrell, 2010). Copyright permission is

available in Appendix D.

Steps to metagenomics sequencing are: experimental design, sampling, DNA extraction,

DNA sequencing using NGS machines, DNA assembly to obtain longer genomes, binning to sort

DNA sequences into groups of closely related organisms, annotation of genes based on their functions and taxonomic groups, statistical analysis, and data storage (Thomas et al., 2012). This approach has offered comprehensive insights into understanding environmental biodegradation processes in situ with a new perspective (Trigo et al., 2008; Vilchez-Vargas et al., 2010).

A study of genes involved in microbial degradation of aromatic compounds was done by

Suenaga et al. (2007). In the study, they found common genes having a specific role in environmental aromatic degradation by an activated sludge sample. Based on a constructed metagenomic library, they were able to define an evolutionary lineage of the genes and identify potential adaptive mutations (Suenaga et al., 2007). Coupling of SIP and metagenomics helps us to simplify the complexity of metagenomics analyses. It is expected that the raw tailings metagenome will be very complex, but the SIP metagenomes will allow us to study only the genomes of a few species that consume particular substrates. Recent studies using the combination of SIP and metagenomics proved that it helps detect low-abundance species and enables the detection of novel enzymes in a targeted metabolically active community in environmental samples (Figure 2-8) (Chen & Murrell, 2010; Dumont et al., 2006; Neufeld et al.,

2008). 33

2.7 Microbial communities in the tailings

Alberta tailings ponds consist of a thin surface aerobic layer (0-10 cm) and an extensive lower anoxic layer. As the depth increases, it becomes anaerobic and methanogenic (Holowenko et al., 2000). Studies of the 16S rRNA gene sequences indicated that the aerobic surface layer had a very different community profile than the rest of the pond (An et al., 2013b).

2.7.1 Aerobic microbial communities

Aerobic microbial communities in the surface OSPW are known to include naphthenic acid degraders and methane oxidizers (methanotrophs) (Fedorak et al., 2002; Scott et al., 2005;

Whitby, 2010). A cultivation-based study by Foght et al. (1985) reported that the aerobic microbial community was dominated by either Alcaligenes sp. and Acinetobacter sp. (Foght et al., 1985; Herman et al., 1993). An initial microbial community study based on 16S rRNA genes was done in 2010 on the composition of the surface waters of the West-In Pit and Mildred Lake

Settling Basin (MLSB) by the Dunfield Laboratory (Saidi-Mehrabad et al., 2013). Two 16S- rRNA gene sequencing analyses conducted in August and October of 2010 for West-In Pit pond

(WIP) surface waters showed that predominant bacteria were members of the phyla

Proteobacteria, Bacteriodetes and Actinobacteria. Dominant genera discovered in the pond were

Rhodoferax (Betaproteobacteria), Hydrogenophaga (Betaproteobacteria), and Methylocaldum

(Gammaproteobacteria). Predominance of the above taxa suggests some conditions in the tailings ponds. For example, Methylocaldum as a methanotroph (Eshinimaev et al., 2004) provides evidence of aerobic respiration within the tailings ponds (Saidi-Mehrabad et al., 2013).

Bacterial communities of the surface tailings ponds studied in 2013 included presumed obligate aerobes (Methylocaldum, Xanthobacter, Flavobacterium), obligate anaerobes

(Methanoregula, Methanosarcina, or the order Clostridiales) and facultative anaerobes (family

Comamonadaceae) as predominant groups (Saidi-Mehrabad et al., 2013). Most predominant groups in OSPW belonged to the Betaproteobacteria (families Comamonadaceae and

Rhodocyclaceae). Known hydrocarbon-degrading genera, such as Acinetobacter, Geobacter,

Rhodoferax, Methylibium, Hydrogenophaga, Pseudomonas, Xanthobacter and Acidovorax, constituted 0.5 – 4% abundance in OSPW (Prince et al., 2010; Saidi-Mehrabad et al., 2013).

Methanotrophy in the surface OSPW can mostly be attributed to an aerobic methanotroph belonging to the Methylococcus/Methylocaldum cluster of Gammaproteobacteria

(1.5% abundance), whereas some other methanotrophs were detected at lower relative abundance

(Methylomonas, Crenothrix, Methylosoma and Methylobacter) (Saidi-Mehrabad et al., 2013).

2.7.2 Anaerobic microbial communities

Within the lower layer of the tailings ponds, hydrocarbon-degrading and methanogenic communities consist of a consortium of syntrophic bacteria, methanogenic archaea, sulfate- reducing bacteria (SRB) and nitrate-reducing bacteria (NRB) (Bordenave et al., 2010;

Holowenko et al., 2000). As syntrophic bacteria and methanogenic archaea break down residual hydrocarbons, these communities release methane into the atmosphere, reaching an amount of

12 g of emitted methane/m2/day (Holowenko et al., 2000).

Lower anaerobic layers of the tailings ponds were predominated by syntrophs, SRB

(Desulfocapsa and Desulfurivibrio spp.), methanogens, and other anaerobes with roles in hydrocarbon utilization or nutrient cycling (Ramos-Padrón et al., 2011). Most prominent methanogenic Archaea detected are in the genera Methanosaeta, Methanoregula, and

Methanolinea (An et al., 2013b). A functional gene analysis study of various OSPW and other

oilsands related samples confirmed the presence of genes responsible for both anaerobic and aerobic activities in situ, including genes for anaerobic aromatic hydrocarbon degradation and methanogenesis, as well as for aerobic aromatic hydrocarbon degradation and methane oxidation

(An et al., 2013b).

2.8 Bioremediation

Microbiological methods have been used to facilitate bioremediation of various industrial wastes, including wastes from both the hydrocarbon extraction and mining processes, and recovery (Allard & Neilson, 1997; Atlas, 1991). Microorganisms are the primary agents for the biodegradation of hydrocarbons and other pollutants in the environment (Balba et al., 1998;

Gutnick & Rosenberg, 1977). Widely known genera of hydrocarbon oxidizers in aquatic environments are Pseudomonas (Proteobacteria), Burkholderia (Proteobacteria), Arthrobacter

(Actinobacteria), Mycobacterium (Actinobacteria), Sphingomonas (Proteobacteria), and

Rhodococcus (Actinobacteria). Pseudomonas (Proteobacteria), Bacillus (Firmicutes),

Micrococcus (Actinobacteria), Nocardia (Actinobacteria), Vibrio (Proteobacteria),

Acinetobacter (Proteobacteria), Brevibacterium (Actinobacteria), Corynebacterium

(Actinobacteria), and Flavobacterium (Bacteriodetes) have been found in various oil-polluted environments (Atlas, 1991; Das & Chandran, 2011).

In a hydrocarbon polluted environment, known hydrocarbon degraders can constitute up to

10% of the microbial community, which is ten times more than their composition in a natural environment (Atlas, 1991). Atlas (1981) noted that these microorganisms could respond to pollutants within hours, with a biodegradation rate faster than the rate of evaporation. In an

aqueous ecosystem, the rate of the uptake and mineralization of the hydrocarbons is related to the aqueous solubility (Balba et al., 1998; Leahy & Colwell, 1990b).

Responses of microorganisms to oil contamination have been studied in depth. For instance, indigenous oil degrading microorganisms played a significant role in breaking down petroleum hydrocarbons and thus reducing overall environmental impact of two of the worst oil spills in U.S. history: the Exxon Valdez and the BP Deepwater Horizon oil spills (Atlas & Hazen,

2011). Studies of rRNA gene sequences and bacterial isolates implicated Gammaproteobacteria

(e.g. Alcanivorax, Marinobacter) and Alphaproteobacteria (Rhodobacteraceae) as key players in hydrocarbon degradation in the Gulf of Mexico beach sands impacted by the Deepwater Horizon oil spill (Kostka et al., 2011). Hydrocarbon degrading bacterial isolates closely related to the same genera were also successfully isolated from oil-contaminated sites in the Persian Gulf and the Caspian Sea, proving their prevalence in situ (Hassanshahian et al., 2012). This brings into light the importance of indigenous bacterial communities in response to oil disturbance and their bioremediation potentials.

2.9 Conclusion

Although hydrocarbon degradation has been extensively studied, NGS technology has opened up new possibilities. Uncultivated microorganisms and microorganisms not previously thought to contribute to hydrocarbon degradation, have been revealed recently via cultivation- independent techniques. Metagenomics have also shown that there are many undiscovered microorganisms and pathways, broadening our perspective of the microbial world. The aim of this dissertation is to contribute to the understanding of the aerobic microbial hydrocarbon degradation in Alberta tailings ponds. Some of the proposed reclamation strategies for tailings

ponds involve increasing the depth of the oxic water cap to allow aerobic biodegradation to naturally detoxify the tailings ponds to wetland ecosystems. It is therefore valuable to know the related mechanisms, as well as the microbial communities responsible in the bioremediation processes in situ.

Preface

Chapter 3 describes the aerobic bacterial communities within the OSPW that degrade benzene and naphthalene. In this study, metagenomic sequencing was performed on fresh WIP-

OSPW sample, while the techniques of DNA-SIP coupled with 16S rRNA gene sequencing were used to target active bacterial communities using 13C-labelled benzene and napthalene.

Dr. Ivica Tamas, Dr. Xiaoli Dong, and Dr. Christoph W Sensen, processed the metagenomic data used in this study, so it could be available online on JGI/IMG for public access. Alireza Saidi-Mehrabad extracted genomic DNA from WIP-OSPW. Dr. JoongJae Kim provided scientific suggestions on bacterial isolation strategies. Benzene and naphthalene measurements performed for this chapter were conducted according to Dr. Lisa M Gieg’s advice.

Dr. Peter Dunfield provided both scientific advice and financial means of this study.

Supplementary Information related to this chapter is provided in Appendix A.

Chapter Three: Benzene and naphthalene degrading bacterial communities in oilsands tailings

Fauziah F. Rochman1, Ivica Tamas1,2, Alireza Saidi-Mehrabad1,3, JoongJae Kim1, Xiaoli Dong4,

Christoph W Sensen1,5, Lisa M Gieg1, Peter F Dunfield1*

1Department of Biological Sciences, University of Calgary, 2500 University Drive NW, Calgary,

Alberta, Canada, T2N 1N4

2Departman Za Biologiju I Ekologiju, Prirodno-Matematicki Fakultet, Univerzitet u Novom

Sadu, Trg Dositeja Obradovića 2, 21000 Novi Sad, Serbia

3Department of Biological Sciences, University of Alberta, Edmonton, Alberta, P6G 2M7,

Canada

4Department of Geosciences, University of Calgary, 2500 University Drive NW, Calgary,

Alberta, Canada, T2N 1N4

5Institute of Molecular Biotechnology, Graz University of Technology, Petersgasse 14, A-8010

Graz, Austria

The Joint Genome Institute IMG/M submission ID number for the metagenome is 43209.

3.1 Abstract Art

3.2 Abstract

Oilsands process-affected water (OSPW), produced by the surface-mining oilsands industry in Alberta, Canada, is alkaline and contains high concentrations of salts, various metals, naphthenic acids, and polycyclic aromatic compounds. Residual hydrocarbon biodegradation occurs naturally in OSPW, but little is known about the aerobic hydrocarbon communities present. In this study, aerobic oxidation of benzene and naphthalene in the oxic surface layer of an oilsands tailings pond were measured. The potential hydocarbon oxidation rates were as high as 36.7 μmol L-1 day-1 for benzene and 85.4 μmol L-1 day-1 for naphthalene. To understand benzene and naphthalene degrading microbial communities in the OSPW, metagenomics was combined with stable isotope probing (SIP), high-throughput sequencing of 16S rRNA gene amplicons, and isolation of microbial strains. SIP using 13C-benzene and 13C-naphthalene detected Betaproteobacteria and Gammaproteobacteria as the most active biodegradation agents. Strains of Methyloversatilis and Zavarzinia were the main benzene degraders, while strains of Thiococcus and Pseudomonas were the main naphthalene degraders. SIP of 13C- methanol confirmed the methylotrophic capability of the predominant Methyloversatilis strain detected. Cultures closely related to some of these bacteria were successfully cultivated using

OSPW supplemented with benzene or naphthalene as a growth medium. Metagenomic analysis revealed a diversity of genes encoding oxygenases active against aromatic compounds, as well as catechol dioxygenases. Although these apparently belonged to many phylogenetically diverse bacteria, only few bacteria were predominant in the SIP experiments. This suggested that many members of the community are adapted to consuming other aromatic compounds, or are active only under specific conditions.

3.3 Introduction

For every cubic meter of bitumen extracted, 3 m3 of water is required, and 4 m3 of tailings consisting of OSPW, sand, clays, residual bitumen and dissolved inorganic and organic compounds are produced (Holowenko et al., 2002; Quagraine et al., 2005). Produced OSPW is deposited into open tailings ponds to allow settling of particles, reuse of surface water for extraction, and long-term pollutant containment for on-site reclamation processes (Government of Alberta, 2009).

In general, oilsands OSPW is alkaline (pH 7.8 – 8), with high contents of salt (2.2 g L-1), various metals, sulfides, naphthenic acids (NAs), and polycyclic aromatic compounds (PAHs)

(Kelly et al., 2010; Kelly et al., 2009; Quagraine et al., 2005; Saidi-Mehrabad et al., 2013).

Other major organic contaminants in OSPW are derived from naphtha (a mixture of C3−C14 alkanes, BTEX (benzene, toluene, ethylbenzene, and xylenes), and iso-parrafins) that is used as a diluent in processing of bitumen from oilsands, and other low molecular weight iso-paraffins and naphthenes (Siddique et al., 2007). In particular, NAs and PAHs are compounds of concern for ecosystems, and thus their presence in tailings ponds has been extensively studied (Allen, 2008;

Grewer et al., 2010; Han et al., 2009; Quagraine et al., 2005; Rogers et al., 2002a; Scott et al.,

2005; Wayland et al., 2008). It is estimated that PAHs and heterocyclic aromatic compounds account for 0.01–10 mg kg-1 of fine tails in Alberta tailings ponds (Wayland et al., 2008).

Environmental concerns have been raised about the potential movement of these compounds into surrounding environments (Wayland et al., 2008). Previous studies on biodegradation of these compounds have emphasized degradation in anoxic conditions, which dominate in most of the volume of the tailings ponds, and relatively little is known about activities in the more oxic surface water cap (c. 1-m deep) of the ponds, from which recycle water is drawn. This surface layer is weakly oxygenated and supports some aerobic microbial activities such as methane oxidation (Saidi-Mehrabad et al., 2013).

The presence of genes for anaerobic and aerobic degradation of aromatic hydrocarbons in several oilsands related samples from Alberta, Canada, has been previously described (An et al.,

Aerobic metabolisms of indigenous microbial populations are known to be pivotal in the reclamation of industrially affected waters (Oller et al., 2011; Tocchi et al., 2012).

Understanding aerobic degradation processes in oilsands process-affected water (OSPW) is valuable because some of the proposed reclamation strategies for tailings ponds involve increasing the depth of the oxic water cap. The ‘wet-landscape’ approach proposes the conversion of tailings ponds to End-Pit Lakes or other wetlands (Allen, 2008; Wayland et al.,

2008). In this strategy, the function of the ponds is reclaimed into End-Pit lakes or other wetland ecosystems, in which a layer of process and river water is used to cap the ponds at a specific depth in order to increase the depth of the oxic surface water layer, provide seed microbes and algae, and dilute contaminants (Allen, 2008; Wayland et al., 2008; Penner, 2016). It is expected that such ponds will detoxify naturally over time, until colonized by aquatic plants. A small-scale study of this reclamation method also showed that aeration and nutrient additions could stimulate indigenous microbial populations that help increase detoxification rates (Allen, 2008).

In this study, we combined metagenomics, stable isotope probing (SIP), high-throughput sequencing of 16S rRNA gene amplicons, and cultivation to examine bacterial communities responsible for the degradation of two model aromatic hydrocarbons, benzene and naphthalene, in the surface oxic layer of an oilsands tailings pond.

3.4 Methods

3.4.1 Sampling and water chemistry

The West-In-Pit (WIP) of Syncrude Canada, Ltd., constructed in 1995, was primarily used as a storage facility for fluid fine tailings and recycle water supply before its closure in

December 2012 (Syncrude Canada, 2010). The pond was closed for reclamation in 2012 to become the first model End-Pit Lake for the oilsands industry in Canada (Syncrude Canada,

2010). Before closure, the pond stored approximately 200 Mm3 of fluid fine tailings capped with a minimum 5 m (30 Mm3) of water. The OSPW used in this study was surface water (0-10 cm) sampled from WIP in August and December 2011. The chemical composition of the OSPW and the water sampling methods were described previously (Saidi-Mehrabad et al., 2013).

Throughout this article, OSPW sampled from WIP tailings pond is termed as WIP-OSPW.

3.4.2 Potential hydrocarbon oxidation rates

To concentrate bacteria, 150 ml of fresh OSPW was filtered through a 0.22-μm filter. The filter and 20 ml of OSPW were added to a 100-ml serum bottle (equivalent to 170 ml of water and 80 ml of headspace). Each sample was run in duplicate. Vials were sealed with butyl rubber stoppers (20 mm) and 40 uM of benzene (Sigma-Aldrich, St Louis, MO, USA) was added.

Naphthalene was prepared in 20 mL of the inert non-degradable carrier 2, 2, 4, 4, 6, 8, 8- heptamethylnonane (HMN; Sigma-Aldrich). The naphthalene stock solution was prepared at an initial concentration of 85 μM in HMN. Two-ml aliquots of HMN stock solution were added to duplicate incubations and abiotic leakage controls. Incubation length was 57 d for benzene and

14 d for naphthalene. At regular intervals one μl of headspace from the benzene incubations was injected into an Agilent 7890A model GC equipped with an Agilent 5975C mass spectrometer

(GC-MS) equipped with a DB-5ms column (30 m × 0.25 i.d. × 3 µm film; J & W Scientific,

Folsom, CA, USA). One μl of the naphthalene-containing HMN was injected by an auto-injector into the same GC-MS connected to a mass selective detector (5975C inert XL MSD series,

Agilent Technologies, Palo Alto, CA, USA), with He as the carrier gas. The GC-MS system was operated as described previously (Fowler et al., 2012; Okoro et al., 2012). Benzene and naphthalene oxidation rates were calculated by linear regression of benzene and naphthalene depletion over the course of 60 and 14 days respectively.

Headspace CO2 was monitored using a Varian 450-GC gas chromatograph equipped with a thermal conductivity detector (TCD) (150°C), after separation in a 2 mm × 0.5 m Hayesep N and a 2 mm × 1.2 m Molecular Sieve 16X column in series (70°C). CO2 production rates were

calculated by linear regression of CO2 production versus time in the vial headspaces. All bottles were incubated at 23°C on a rotary shaker at 180 rpm to ensure aeration.

3.4.3 Stable isotope probing (SIP)

To identify microbial communities actively assimilating the model aromatic compounds,

13 13 isotopically labelled benzene ( C6 99 atom%, Sigma-Aldrich) and naphthalene ( C10H8 99 atom%, Sigma-Aldrich) were used in DNA-SIP. An additional experiment also used isotopically

13 labelled methanol ( CH3OH 99 atom%, Sigma-Aldrich). Methanol, a byproduct of methane oxidation, was used because a large number of putative methylotrophs have been identified in the pond (Saidi-Mehrabad et al., 2013), which suggests that they might be cross-feeding on metabolic byproducts of methane oxidation.

Water samples were filtered as described above. The filter and 20 ml of OSPW were added to a 100-ml serum vial (equivalent to 170 ml of water sample and 80 ml of headspace) and supplemented with 0.05% (w/w) of labelled benzene, naphthalene, or methanol. Each sample was run in duplicate or triplicate. Vials containing 20 ml of filtered OSPW without added substrate were used as controls. Vials were sealed with butyl rubber stoppers and incubated at

23°C on a rotary shaker at 180 rpm in the dark. Headspace CO2 was monitored for all SIP- enrichment samples to determine incubation lengths. Incubations were terminated when CO2 concentration in the microcosms increased by around 0.0025 – 0.005 mol. After 7 days

(naphthalene and methanol), and 9 days (benzene), of incubation, cells were collected by centrifugation for 10 min at 10,000 ×g. DNA was extracted from the pellett using the FastDNA

Spin Kit for Soil (MP Biomedicals, Santa Ana, CA, USA) with additional purification steps using 5.5 M guanidine thiocyanate (Knief et al., 2003), and stored at −80°C. DNA was separated

by isopycnic centrifugation in CsCl and fractionated as described previously (Sharp et al., 2012).

DNA extracted from OSPW incubated for the same time without addition of substrates was used as a control to determine the expected position of unlabelled DNA in CsCl gradients. Heavy

DNA fractions of all the labelled samples were used for 16S rRNA gene amplification and sequencing (Supplementary Figure A1). Heavy fractions of OSPW receiving the same treatment as all of the amended samples (OSPW-heavy), and unfractionated DNA from fresh OSPW

(OSPW-control) were used as controls. Amplification of 16S rRNA genes was carried out as described previously (Sharp et al., 2014a). Purified PCR products (~150 ng total DNA) were sequenced using a 454 Life Sciences Genome Sequencer FLX (Roche, Branford, CT, USA) and analysed by QIIME as previously described (Caporaso et al., 2010; Sharp et al., 2014a). All sequence data have been submitted to the Sequence Read Archive (SRA) under accession number PRJNA338192.

3.4.4 Metagenomics

Genomic DNA was prepared and extracted as described previously (Saidi-Mehrabad et al., 2013). DNA concentration was determined with a Qubit Fluorometer using a Quant-iT dsDNA HS Assay Kit (Invitrogen, Carlsbad, CA, USA). Total genomic DNA (12 μg) was sequenced using a combination of Roche 454 GS FLX and paired-end Illumina HiSeq2000 platforms at the Genome Quebec and McGill University Innovation Center, Montreal, Quebec.

Quality control and assembly of metagenomic data was performed at the University of Calgary

Visual Genomics Center, Calgary, Canada according to previous reports (Aguilar et al., 2016;

Saidi-Mehrabad et al., 2013; Tan et al., 2013). Annotation was conducted by submission to the

IMG platform of the Joint Genome Institute (JGI) website (http://www.jgi.doe.gov/) (Markowitz et al., 2012).

3.4.5 Isolation of hydrocarbon degrading bacteria

Several media were prepared to cultivate bacterial strains, including complex media targeting versatile chemotrophs as well as media containing the model hydrocarbons as sole carbon and energy sources. OSPW was diluted 1 : 10 with milliQ water and 100-μl aliquots were spread on 20% strength R2A plates (20R2A) (Reasoner & Geldreich, 1985) adjusted to pH 8 with NaOH, as well as on a modification of the basal mineral salts medium M10. Medium M10 was prepared as described previously (Heyer et al., 2002), except that agar was substituted with

1.5% phytagel (Sigma-Aldrich), and milliQ water was substituted with 0.2-μm-filtered

(Pall Life Sciences, East Hills, NY, USA) OSPW to mimic its living environment. We named these modified media as 20R2A-T and M10-T. All plates were incubated aerobically at 20°C.

To specifically grow naphthalene-degrading microorganisms, OSPW samples were initially enriched with 0.05 % naphthalene (99.9%, Sigma-Aldrich) for 7 d, serially diluted, and plated onto solid media M10-T and 20R2A-T prepared in 125-ml wide-mouth jars (I-Chem,

VWR, Radnor, PA, USA) instead of Petri plates. These growth vessels were inverted with 3 g of naphthalene placed on the inverted lids to diffuse naphthalene vapour, as previously described

(Jeon et al., 2004), and incubated at 20°C for 3 months. For benzene and methanol-degrading microorganisms, OSPW samples enriched with both benzene and methanol (from the SIP experiments described above) were diluted serially, spread onto M10-T and 20R2A-T media in the wide-mouth jars, and incubated with the addition of 0.1 ml, 0.01 ml, and 0.001 ml of benzene and methanol per jar at 20°C for 3-6 months.

Colonies were picked and streaked to new plates of the same medium. This was repeated several times until purified strains were obtained. Colony PCR was performed to screen the isolates. Single colonies were picked with autoclaved toothpicks and frozen at -80°C overnight in 0.2-ml tubes to lyse cells. 16S rRNA genes were amplified using the universal primer set 9f and 1492b (Weisburg et al., 1991). PCR reaction conditions were: initial denaturation at 94°C for 10 min, followed by 35 cycles of 1 min at 94°C, 1 min at 56°C and 2 mins at 72°C, and a 10- min final elongation at 72°C. PCR products were visualized on a 1% agarose gel and purified with an EZ-10 Spin Column PCR Purification Kit (BioBasic Inc.). DNA concentration was determined via a Qubit Fluorometer using a Quant-iT™ dsDNA HS Assay Kit

(Invitrogen). Sanger sequencing performed at the University of Calgary Core DNA Services using Applied Biosystems 3730xl (96 capillary) genetic analyzer (Applied Biosystems, Foster

City, CA, USA). To identify the isolates, obtained 16S rRNA forward and reverse sequences of each isolate were combined in via EMBOSS 6.3.1: merger (http://mobyle.pasteur.fr/cgi- bin/portal.py?#forms::merger), using default settings (Néron et al., 2009). Nearly full length

(>1,300 bp) sequences were compared to the most related sequences via BLASTN (Altschul et al., 1997) against the database containing type strains with validly published prokaryotic names using the EzTaxon server (http://www.ezbiocloud.net/eztaxon; Kim et al., 2012).

3.4.6 Hydrocarbon degradation capabilities of bacterial isolates

Eight isolates were tested for hydrocarbon degradation potentials using benzene:

Xanthobacter strain OSPW1, Pseudomonas strain OSPW2, Thauera strain OSPW3,

Brevundimonas strain OSPW4, Microbacterium strain OSPW5, Zavarzinia strain OSPW6, and

Aquimonas strain OSPW7. Information regarding their isolation sources is provided in

Supplementary Table A1. Exponentially growing bacteria in 20R2A medium (pH 8.5) were transferred to sterile 20R2A liquid medium (20 ml in a 100-ml serum vials), and sealed with butyl rubber stoppers. Six mM of benzene (Sigma-Aldrich) was added to the cultures. All bottles were incubated at 23°C on a rotary shaker at 180 rpm to ensure aeration. Loss of benzene was measured regularly as described above. Growth was monitored by measuring OD600 on an

Ultrospec 10 Cell Density Meter (Amersham Biosciences, Piscataway, NJ). Each test was done in duplicate vials.

3.5 Results

3.5.1 Hydrocarbon oxidation

Direct measurement of headspace benzene and dissolved naphthalene loss in WIP samples incubated with these substrates demonstrated that oxidation occurred (Supplementary

Figures A2 and A3). The benzene oxidation rate was 36.7 μmol L-1 OSPW day-1 in the first 40 d. The naphthalene oxidation rate was 85.4 μmol L-1 OSPW day-1 in the 14 d incubation. In the

-1 -1 benzene enrichment samples, around 27.3 μmol of CO2 L OSPW day was produced, representing a 74.4% expected CO2 production based on the stoichiometric equation for direct benzene oxidation, which produce microbial cells, CO2, and water: C6H6 + 2.5O2 + NH3 

C5H7O2N + CO2 + H2O (Shuler and Kargi, 2002). In the naphthalene enriched samples, 232.3

-1 -1 μmol of CO2 L OSPW day was produced, showing 54.4% expected CO2 production, based on the following stoichiometric equation: C10H8 + 7O2 + NH3  C5H7O2N + 5CO2 + 2H2O, modified from Shuler and Kargi (2002).

51 1

2 Figure 3-1. Community structures of oilsands processed water from the WIP tailings pond (OSPW-control) and from the heavy, 13C-labelled

3 DNA fractions of samples amended with 13C benzene (Benzene), naphthalene (Naphthalene), and methanol (Methanol), incubated under

4 aerobic conditions. Predominant taxa are identified to the class level based on the SILVA database. The OSPW-Heavy treatment is the

5 community detected only in the heaviest DNA fraction of OSPW. OSPW-Control shows the entire unseparated DNA from OSPW. The

6 communities demonstrated that the heavy fractions retrieved from the SIP experiments were primarily determined by 13C enrichment of DNA

7 rather than variabilities in genomic G+C content.

3.5.2 SIP and 16S rRNA gene sequencing

13 OSPW samples fed with C-substrates showed higher CO2 production rates compared to control samples (i.e. WIP-OSPW without substrates added; Supplementary Figure A4). Density gradient separation of extracted DNA indicated that 13C-substrate incubations also showed a shift in the DNA distribution towards heavier values (Supplementary Figure A1). The positions of the

"heavy" DNA fractions representing the active 13C-assimilating communities were determined by comparing density gradients of the DNA extracted from the SIP incubations to that of the control samples. The fractions chosen for community analyses are indicated in Supplementary

Figure A1.

Community analysis of 16S rRNA genes amplified from the DNA extracted from OSPW prior to the incubations showed both methanogenic archaea and diverse bacteria. The combination of strict anaerobes, aerobes, and facultative anaerobes has been noted before (An et al., 2013b; Saidi-Mehrabad et al., 2013) and is probably the result of active water circulation between anoxic and slightly oxic water zones. Among the bacteria the dominant classes/phyla were Betaproteobacteria (9.6%), Alphaproteobacteria (7.3%) and Flavobacteria (4.9%; Figure

3-1; Supplementary Figure A-5). The predominant species were Hydrogenophaga taeniospiralis

(1.8% of 2,404 reads), Lutibacter maritimus (1.6% of 2,404 reads), and Enhydrobacter aerosaccus (1.4%, of 2,404 reads; Supplementary Table A2-A). The community detected in only the heaviest fractions of a DNA extract after separation in a CsCl gradient (Supplementary Table

A2-B, Figure 3-2) was much simpler, and dominated by the genera Devosia (23.3% of 6,098 reads), Methylomonas (12.6% of 6,098 reads), and Thauera (9.0% of 6,098 reads).

Figure 3-2. Bubble chart of all samples based on relative abundances (>1%). Classification percentage is shown for the rank of genus (% classified). Benzene: heavy-DNA fraction of

benzene-amended OSPW incubated for 9 d; Naphthalene: heavy-DNA fraction of

naphthalene- amended OSPW incubated for 7 d; Methanol: heavy DNA fraction of

methanol-amended OSPW incubated for 7 d; OSPW-heavy: heavy fraction of OSPW receiving the same treatment as all of the amended samples; and OSPW-control: complete,

unfractionated DNA from OSPW.

The OSPW-heavy community is therefore a simplified subset of the OSPW-control community, suggesting that a few species like Devosia sp. have high G+C contents and therefore high DNA densities. This is supported by sequenced genomes for these genera: genus Devosia has an average G+C content of 62.7%, genus Methylomonas has an average G+C content of

52.3%, and genus Thauera has an average G+C content of 66.8% (Nordberg et al., 2014).

Results of the SIP experiments were compared to both of these controls to verify that the species detected were truly enriched with 13C. Only OTUs enriched at least 10-fold in the SIP heavy fractions compared to both controls were considered to be clearly enriched in the added substrate.

The active communities detected in the heavy DNA fractions from SIP incubations were all dominated by Proteobacteria. 13C-benzene SIP was dominated by Betaproteobacteria

(88.0%), the 13C-naphthalene SIP by Gammaproteobacteria (59.2%), and the 13C-methanol SIP by Betaproteobacteria (57.1%; Figure 3-1). In the heavy DNA of the 13C-benzene SIP sample, a close relative of Methyloversatilis universalis accounted for 86.9% (11,071 out of 12,745 reads) in relative abundance and was enriched 20-fold compared to the OSPW-heavy control (Figure 3- 55

2, Table 3-1). Zavarzinia compransoris was the second most predominant OTU with 5.2% (660 out of 12,745 reads), an enrichment of 26-fold compared to OSPW-heavy controls. An OTU closely related to Geobacter sp. (Deltaproteobacteria) was also enriched (14.6-fold) in the 13C- benzene SIP (Table 3-1). All other OTUs accounted for <1.5% of reads (<80 reads or less).

The three most dominant OTUs in the 13C-naphthalene SIP heavy fraction were: a

Thiococcus sp. at 45.6% (4,045 out of 8,949 reads), a Thauera sp. at 20.5% (1,820 out of 8,949 reads), and a Pseudomonas sp. at 8.2% (727 out of 8,949 reads). The Thiococcus and

Pseudomonas OTUs were enriched >25 fold compared to the OSPW-heavy control (Table 3-1).

However, the Thauera was similarly enriched in the OSPW-heavy control (Figure 3-1) indicating that this organism has a naturally high G+C content, and we therefore cannot conclude that it assimilated the naphthalene. Other enriched OTUs in the 13C-naphthalene SIP were related to three different Pseudomonas sp. (increased around 37.7-62.0 times) and Gemmobacter sp.

(increased by 36.8 times; Table 3-1). The most dominant highly enriched OTUs (>10-fold compared to controls) in the 13C-methanol SIP were Methyloversatilis universalis (53.0%, 4945 out of 9,338 reads), and Hyphomicrobium sp. (2.1%, 199 out of 9,338 reads).

3.5.3 WIP-OSPW metagenome

An overview of the OSPW metagenome features is given in the Supplementary Table A3.

The metagenome data shows extensive evidence of the microbial community's capacity for aerobically degrading diverse petroleum hydrocarbons. This concurs with a previous study where the predominance of an aerobic bacterial community was evident in surface OSPW samples (An et al., 2013b). A predicted 39,257 out of 520,901 annotated genes belonged to the KEGG category ‘xenobiotics biodegradation and metabolism’ (Supplementary Figure A6). Genes

encoding p-cymene, benzene, toluene, ethylbenzene, trans-cinnamate, phenylpropionate, biphenyl, 4-chlorobiphenyl, carbazole, naphthalene, 1-methylnaphthalene, phenanthrene, benzoate, methane, 4-chlorophenoxyacetate, 2,4-dichlorophenoxyacetate, and cyclohexanol degradation through various dioxygenase and monooxygenase reactions are predicted.

In the OSPW sample, aromatic and polyaromatic compounds are predicted to be broken down through 3 different aerobic pathways: dioxygenase and dehydrogenase reactions, ring removal from polycyclic aromatic ring structures, and two-monooxygenase reactions. Each process forms catechol as a common intermediate. Catechol is broken down through the catechol meta-cleavage pathway into acetyl-CoA (Figure 3-3). Benzene is attacked in this scenario by either benzene/toluene dioxygenases, in which two O molecules are incorporated into benzene, forming catechol, or by phenol hydroxylases, whereby only one O atom is incorporated in two separate steps, forming phenol in the first stage, and catechol in the second. Naphthalene is attacked by dihydroxynaphthalene dioxygenases through the ring removal from polycyclic aromatic rings pathway, forming at least 3 different metabolites upstream of catechol (Figure 3-

3).

Detected genes encoding for dmpB/xylE (catechol 2,3-dioxygenase) showed best BLAST hits to diverse genera including the Gammaproteobacteria, the Alphaproteobacteria, and the

Betaproteobacteria, as well as others (Tables A4 A-P). Genes for todC1/bedC1 (benzene/toluene dioxygenase) can be linked to Gammaproteobacteria, Betaproteobacteria, and

Alphaproteobacteria), and others (Tables A4 A-P). Each of the genes encoding for nahAc/ndoB

(naphthalene 1,2-dioxygenase) and nahC (dihydroxynaphthalene dioxygenase) were most closely related to those of one organism: Liberibacter (Alphaproteobacteria) and Parvibaculum

(Alphaproteobacteria) respectively.

57 1

2 (A)

Figure 3-3 Aerobic hydrocarbon degradation pathways of benzene, naphthalene, and the

corresponding genes detected in the OSPW metagenome based on KEGG pathways. (a)

Detected pathways (substrates) for aerobic hydrocarbon metabolism and associated

enzymes. (b) Bar chart of various genes and putative genes counts involved in aerobic

hydrocarbon metabolism in the metagenome with >2500 bp scaffold length. Pathways descriptions: 1, meta-cleavage of catechol; 2, dioxygenase and dehydrogenase reactions; 3, ring removal from polycyclic aromatic ring; and 4, two monooxygenase reactions. Enzyme

abbreviations: dmpB, xylE, catechol 2,3-dioxygenase; dmpD, xylF, hydroxymuconate-

semialdehyde hydrolase; dmpC, xylG, aminomuconate-semialdehyde/2-hydroxymuconate-

6-semialdehyde dehydrogenase; praC, xylH, oxalocrotonate tautomerase; dmpH, xylI, nahK, 2-oxo-3-hexenedioate decarboxylase; mhpD, 2-keto-4-pentenoate hydratase; mhpE,

4-hydroxy 2-oxovalerate aldolase; mhpF, acetaldehyde dehydrogenase; todC1, bedC1,

benzene/toluene dioxygenase; nahAc, ndoB, naphthalene 1,2-dioxygenase; nahC,

dihydroxynaphthalene dioxygenase; nahD, hydroxychromene-2-carboxylate isomerase;

nahE, trans-o-hydroxybenzylidenepyruvate hydratase-aldolase; nahF, salicylaldehyde

dehydrogenase; sal-hyd, salicylate hydroxylase; dmpK, poxA, phenol hydroxylase.

Based on best BLAST hits, several genes in the metagenome appear to belong to the

Methyloversatilis strain identified in the SIP experiments as an important degrader, including dmpB/xylE (catechol 2,3-dioxygenase), praC/xylH (oxalocrotonate tautomerase), and dmpH/xylI/nahK (2-oxo-3-hexenedioate decarboxylase). Thiococcus sp. was predicted by SIP to be important, but reads mapping most closely to known Thiococcus genomes were not detected in the metagenome. However, another member of the Chromatiaceae family was detected as the closest BLAST hit to genes dmpB/xylE (catechol 2,3-dioxygenase), and todC1/bedC1

(benzene/toluene dioxygenase) (Tables A4 A-P). Pseudomonas spp. can be linked to dmpB/xylE, todC1/bedC1, dmpK/poxA (phenol hydroxylase), dmpD/xylF (hydroxymuconate-semialdehyde hydrolase), dmpC/xylG (2-hydroxymuconate-6-semialdehyde dehydrogenase), dmpH/xylI/nahK

(2-oxo-3-hexenedioate decarboxylase), praC/xylH (oxalocrotonate tautomerase), mhpE (4- hydroxy 2-oxovalerate aldolase), and mhpF (acetaldehyde dehydrogenase).

60 3.5.4 Isolated bacterial strains and their degradation potentials

A list of bacterial isolates obtained is presented in Appendix A: Table A1. Some bacteria representing major OTUs identified via the SIP experiments as active in degrading benzene, naphthalene or methanol were obtained in pure cultures. Pure cultures of Xanthobacter strain

OSPW1 and Zavarzinia strain OSPW2, isolated using 20R2A-T medium, were able to perform complete degradation of 8 mM benzene in 14 and 8 days, respectively. For both cultures, growth continued despite the complete degradation of benzene (Appendix A: Figures A7 A-B), presumably growing from nutrients dissolved in 20R2A medium. Pseudomonas stutzeri strain

OSPW8 was isolated on M10-T with naphthalene as a sole substrate. Two predominant species detected from both 13C-benzene and 13C-naphthalene SIP, Methyloversatilis universalis and

Thiococcus, sp. were not isolated. Two predominant species detected from both 13C-benzene and

13C-naphthalene SIP, Methyloversatilis and Thiococcus, sp. were not isolated.

3.6 Discussion

2011; Hazen et al., 2016). Studies of rRNA gene sequences and bacterial isolates implicated

Gammaproteobacteria (e.g. Alcanivorax, Marinobacter) and Alphaproteobacteria

(Rhodobacteraceae) as key players in hydrocarbon degradation in Gulf of Mexico beach sands impacted by the Deepwater Horizon oil spill (Kostka et al., 2011). Hydrocarbon degrading bacterial isolates closely related to the same genera were also successfully isolated from oil- contaminated sites in the Persian Gulf and the Caspian Sea, proving their prevalence in situ

(Hassanshahian et al., 2012). This brings into light the importance of indigenous bacterial communities in response to oil disturbance and their bioremediation potentials.

In oilsands tailings ponds, major organic contaminants in the fine tailings are derived from naphtha (C3−C14), BTEX, and other low molecular weight iso-paraffins and naphthenes, used as diluents in the bitumen extraction process, although small amounts of these compounds may also arise directly from the bitumen (Siddique et al., 2007). One plan for reclaiming oilsands tailings ponds and reducing their water toxicity is to convert them into End-

Pit Lakes or other wetlands (Allen, 2008; COSIA, 2016; Wayland et al., 2008). This method uses a layer of process and river water to cap the ponds in order to increase the depth of the oxic surface water layer, provide seed microbes and algae, and dilute contaminants (Allen, 2008;

Wayland et al., 2008; Penner, 2016). It is expected that such ponds will detoxify naturally over time, until colonized by aquatic plants. It was shown that aeration and nutrient additions could stimulate indigenous microbial populations that help increase decontamination rates (Allen,

2008), stressing the importance of understanding aerobic degradation processes of the oilsands process-affected water (OSPW) in tailings ponds.

The potential oxidation rates of the sampled OSPW were as high as 36.7 μmol L-1 day-1 for benzene and 85.4 μmol L-1 day-1 for naphthalene. Previous studies have shown complete mineralization of benzene in Salt Plains soil, amended with mineral salts medium, under aerobic conditions, with a rate of 600 and 640 μmol kg-1 day-1 at 37 and 45°C (Nicholson & Fathepure,

2005). A study also summarized aerobic benzene degradation rates in sandy aquifer samples and

Pseudomonas sp. cultures, showing benzene degradation rates more than 10 times higher than the rates in WIP-OSPW (Alvarez et al., 1991). Potential naphthalene degradation rates were therefore higher than those of benzene, which agrees with a previous study showing that

naphthalene had a higher biodegradation rate constant than other major hydrocarbons, including benzene in an enriched sludge culture (Li & Goel, 2011). When compared to other complex tricyclic or tetracyclic PAHs, rates of naphthalene degradation were generally faster (Chang et al., 2014). However, degradation rates in the tailings water were somewhat lower than previously measured rates in other oil-contaminated environments, probably because the naphthalene level in OSPW was low (101 ± 27 ng L−1) (Brownlee et al., 1999) compared to other organics such as naphthenic acids (40–120 mg L−1), leading to a lower naphthalene degradation activity (Demeter et al., 2014; Holowenko et al., 2002; Scott et al., 2005).

The use of metagenomics allowed us to identify complete genetic pathways encoding enzymes for aerobic degradation of diverse aromatic and polyaromatic hydrocarbons in the WIP-

OSPW community. Two degradation pathways (via dioxygenases and monooxygenases) were identified for aerobic benzene degradation (Jindrová et al., 2002), while a pathway for naphthalene degradation through a naphthalene 1,2-dioxygenase was also predicted (Figure 3-

3A) (Chang et al., 2014). Catechol, as an intermediate of all pathways, could be further mineralized via either ortho- and meta-cleavage pathways. Metagenomic analysis revealed abundant dmpB and xylE (catechol 2,3-dioxygenase) genes encoding enzymes responsible for converting catechol into formate (Figure 3-3A). However, metagenomes have limited value for identifying bacterial species involved in particular processes, in part because: i) species identifications of key genes can usually only be inferred from BLAST identities, which are not always accurate due to lateral gene transfer and other limitations, ii) some species may have the genetic systems required for a process but be only weakly active in situ, and iii) a large number of genes detected (in our case ring-cleaving oxygenases) are not closely homologous to studied enzymes and their exact substrates are therefore uncertain. For example, todC1/bedC1

(benzene/toluene dioxygenase) and dmpK/poxA (phenol hydroxylase) gene scaffolds in the metagenome were mapped to Pseudomonas sp. based on BLAST, but this bacterium was identified by SIP to be important in naphthalene but not benzene degradation. Benzene degradation genes from the metagenome were not mapped to Methyloversatilis sp., although this was identified by SIP as the most predominant organism in benzene degradation. There are clear limitations of the homology-based metagenome analysis. Therefore cultivation and SIP studies were also undertaken here.

13 OSPW samples fed with C-substrates showed higher CO2 production compared to control samples (fresh WIP-OSPW) and tails of heavy DNA peaks appeared on the SIP graphs, showing active uptake of 13C-substrates in the samples (Supplementary Figures A4). 16S rRNA gene sequencing of the 13C-benzene-labelled heavy DNA fractions revealed the prevalence of the methylotroph genus Methyloversatilis. In SIP of 13C-methanol Methyloversatilis was also dominant in the heavier methanol SIP fractions (fractions 3 – 4), verifying its methylotrophic capability (Supplementary Figure A4). The genome of the type strain FAM5T of

Methyloversatilis universalis has a membrane protein involved in catabolism of aromatic compounds via metacleavage pathway (Kittichotirat et al., 2011), and a recent bioinformatics approach discovered an uncultured bacterium related to M. universalis strain FAM5 (98% identity on BLASTN) that is a benzene degrader (Satija & Gore, 2014). Previous studies have also reported this bacterium in various petroleum related environments, but its relative abundance never reached more than 18% and its roles were not deemed crucial in hydrocarbon degradation (Golby et al., 2012; Lenchi et al., 2013; Noguchi et al., 2014). Over the course of this study, M. universalis was not isolated.

Another prevalent OTU identified in the heavy fractions of 13C-benzene SIP was a

Zavarzinia strain. This genus was originally isolated in 1974 from the sludge of the Moscwa

River and called Comamonas compransoris, but was renamed to Pseudomonas compransoris in

1980, and finally renamed to Zavarzinia compransoris in 1993 (Willems & De Vos, 2006). It has been detected in various hydrocarbon-contaminated sites (Al-Mailem et al., 2014a, 2015a; Al-

Mailem et al., 2015b; Al-Mailem et al., 2014b; Nies & Schlegel, 1982). A member of this OTU was isolated, named Zavarzinia strain OSPW6, and was shown to degrade 8 mM benzene in 8 days of incubation (Supplementary Figure A7). The growth of an unknown Geobacter was also stimulated by benzene (Table 3-1). Some Geobacter species are found in contaminated aquifers and are capable of anaerobic benzene degradation (Meckenstock et al., 2016; Zhang et al., 2012;

Zhang et al., 2013).

A different bacterial community was implicated in naphthalene degradation: predominantly the Gammaproteobacteria genera Thiococcus and Pseudomonas, and

Betaproteobacteria genus Thauera. Our previous study identified unknown Chromatiaceae related to Thiococcus in OSPW surface water sample (2.6-8.0% abundance) (Saidi-Mehrabad et al., 2013). The results obtained in this study study verified a role for an uncultured

Chromatiaceae (95% similarity to Thiococcus) in naphthalene degradation, and genes related to catechol 2,3-dioxygenase, benzene/toluene dioxygenase, and phenol hydroxylase in the metagenome were mapped by BLAST most closely to members of this family. The family

Chromatiaceae has previously been linked to homogentisate 1,2-dioxygenase (HGD), a unique aromatic ring-cleavage enzyme, which cleaves an aromatic ring between ortho carbon atoms substituted with carboxyl and hydroxyl groups (Pérez-Pantoja et al., 2010; Titus et al., 2000). In addition to the catabolism of aromatic rings, HGD genes are also involved in the breakdown of

amino acids tyrosine and phenylalanine (Titus et al., 2000). Although similarity is relatively low

(28.7% and 26.0%), two HGD genes are detected in the metagenome data are related to

Chromatiaceae, showing the possibility of the family Chromatiaceae in aerobic aromatic degradation in OSPW.

On the other hand, P. stutzeri strains have been studied biochemically and genetically for naphthalene degradation (Rosselló-Mora et al., 1994). A P. stutzeri strain OSPW 23 was successfully cultivated from our WIP-OSPW sample supplemented with naphthalene as a sole substrate (Jeon et al., 2004). In the WIP-OSPW metagenome dmpB/xylE (catechol 2,3- dioxygenase) genes with close homologs to Pseudomonas sp. (73% similarity) and

Methyloversatilis sp. (74% similarity) were detected in a single scaffold (Supplementary Table

A4-C).

OSPW-heavy control are noted. UD indicates that no reads were detected in the original sample- the minimum x-fold enrichment in these cases is calculated assuming 1 read in the original sample.

Relative % x-fold % Identity abundance enrichment SIP-Sample Number Relative % OTU ID Taxonomy (Phylum; Class) Closest relative (BLAST) to the closest in control compared to (reads) of reads Abundance relative heavy control heavy fraction fraction Benzene 5106 Proteobacteria; Betaproteobacteria Methyloversatilis universalis 100 11071 86.87 4.35 19.97

(12,745) 5634 Proteobacteria; Alphaproteobacteria Zavarzinia compransoris 99 660 5.18 0.20 25.89

873 Proteobacteria; Deltaproteobacteria Geobacter lovleyi 91 62 0.49 UD >14.55

Naphthalene 4006 Proteobacteria; Gammaproteobacteria Thiococcus pfennigii 95 4045 45.56 1.80 25.31

(8,879) 700 Proteobacteria; Gammaproteobacteria Pseudomonas stutzeri 99 727 8.19 0.23 35.60

5076 Proteobacteria; Gammaproteobacteria Pseudomonas anguilliseptica 96 184 2.69 UD >61.96

2995 Proteobacteria; Gammaproteobacteria Pseudomonas fulva 98 172 5.73 1.00 >191

7146 Proteobacteria; Gammaproteobacteria Pseudomonas knackmussii 97 100 1.13 UD >37.68

6096 Proteobacteria; Alphaproteobacteria Gemmobacter caeni 99 82 1.23 UD >36.78

Methanol 5106 Proteobacteria; Betaproteobacteria Methyloversatilis universalis 100 4945 52.96 4.35 12.17

(9,338) 197 Proteobacteria; Alphaproteobacteria Hyphomicrobium methylovorum 96 199 2.13 0.13 16.39

4723 Proteobacteria; Betaproteobacteria Delftia acidovorans 100 105 1.12 UD >37.33

816 Actinobacteria; Propionibacteriales Propionibacterium acnes 99 82 0.88 UD >29.33

337 Bacteroidetes; Sphingobacteriia Sediminibacterium goheungense 96 53 0.57 UD >19.00

4209 Armatimonadetes; Fimbriimonadia Fimbriimonas ginsengisoli 93 50 0.54 UD >18.00

6504 Planctomycetes; Planctomycetia Planctomicrobium piriforme 92 32 0.34 UD >11.33

The microbial community detected in the OSPW usually harbors many Burkholderiales

(Betaproteobacteria, Comamonadaceae) (An et al., 2013b; Saidi-Mehrabad et al., 2013). In this study, Burkholderiales was one of the most predominant groups in the WIP-OSPW accounting for close to 18.0% of all 16S rRNA genes of all the metagenome reads. This agrees to the analysis of 16S rRNA gene amplicons, which reveals Burkholderiales as predominant

(Supplementary Figure A4). Based on BLAST analysis of genes predicted in the metagenome, the family Comamonadaceae is associated with most of the genes associated to catechol meta- cleavage (Supplementary Tables A4). Recognizing the limitations of BLAST, this analysis nevertheless suggests some importance of this taxon in aromatic ring cleavage.

Members of the Rhizobiales (Alphaproteobacteria) also appear to be important in the metagenome, and genes encoding monooxygenasess for degradation of aromatic coupounds detected in the metagenome often matched most closely to genes of organisms in this group based on BLAST identities (Supplementary Table A3). Nevertheless, these two dominant groups did not appear in either 13C-benzene and 13C-naphthalene SIP heavy fractions. This finding indicates either that these genes cannot be mapped properly to any taxon with BLAST, or more likely that the detected genes are involved primarily in other degradation pathways, rather than degradation of the model aromatic compounds used in our experiments.

The coupling of metagenomic sequencing technology, SIP, and traditional microbiological methods allowed us to identify organisms with important roles within the

OSPW. Utilizing the metagenome alone was not very useful, since it predicted a huge number of potential hydrocarbon degraders (mostly Alphaproteobacteria) but in the actual SIP heavy fractions of both 13C-benzene and 13C-naphthalene, only a few of these potential bacteria were actually shown to be active. The others either concentrate on other substrates or are active under

other conditions. Predominant bacteria found to be important in our benzene and naphthalene amended environments, such as Methyloversatilis and Thiococcus, were not necessarily predicted by other related studies (Hassanshahian & Boroujeni, 2016; Jechalke et al., 2013;

Ortega-González et al., 2013; Patel et al., 2012; Pérez-Pantoja et al., 2010). The combination of both metagenome analyses and SIP enabled us to determine metabolic roles of some resident microbial communities.

3.7 Acknowledgments

The authors thank the mining companies for the support of this research. Particular thanks to Syncrude Canada, Ltd. for providing samples. This research was supported by Genome

Canada and Genome Alberta (Hydrocarbon Metagenomics), and the Natural Sciences and

Engineering Council of Canada (NSERC grant CRDPJ478071-14).

Preface

Alberta tailings ponds, 13C-labelled methanol and acetate were added as substrates to WIP-

OSPW samples. 13C-labelled protein extract was also used to study indigenous proteolytic

13 bacterial communities. Photosynthetic bacterial communities were studied by adding CO2 into

WIP and MLSB-OSPW samples incubated under constant light exposure.

Drs. Allyson Brady and Christine Sharp trained me to perform SIP technique, i.e. sample set up, and DNA fractionation by cesium chloride (CsCl) density-gradient centrifugation. QIIME analysis was done according to the pipeline outlined by Dr. Christine Sharp. Dr. Peter Dunfield provided scientific and financial support of this study.

Chapter Four: Identification of methanol-, acetate-, protein-assimilating, and phototrophic bacterial communities in the surface oilsands process-affected water by stable-isotope probing

4.1 Introduction

Oilsands tailings ponds are man-made dyke systems that contain a mixture of salts, suspended solids, residual hydrocarbons, and soluble organic compounds, with traces of solvents used for oilsands extraction process (Saidi-Mehrabad et al., 2013; Small et al., 2015). Large volumes of tailings are a byproduct of bitumen extraction, and incomplete bitumen and solvent recovery have caused organic compounds within tailings ponds to increase over time (Small et al., 2015). Currently covering an area of about 88 km2 (Alberta Energy, 2016), expanding storage of large volumes of tailings in the Alberta tailings ponds has become a major concern for

.(s oilsands industry (Schindler, 2013׳Canada

An active tailings pond consists of a thin layer of oxic surface water (1–2 m) overlying a deep (up to ~40 m in depth), viscous layer of unprocessed bitumen particles and mature fine tailings (MFT) (Nix & Martin, 1992; Saidi-Mehrabad et al., 2013). These fine tailings settle by gradual densification, forming mature fine tailings (MFT) in the first 3–4 years of deposition, before decades for accumulative settling (Penner & Foght, 2010b; Siddique et al., 2011). The presence of inorganic and organic contaminants in the lower anoxic layer create a condition conducive to methanogenesis (Quagraine et al., 2005; Saidi-Mehrabad et al., 2013). As a result, emission of biogenic greenhouse gases (CH4 and CO2) became one of the major concerns associated to the tailings ponds.

Methanogenesis is one of the anaerobic processes in the tailings ponds, causing the release of methane from the ponds (Holowenko et al., 2000). The largest tailings basin belongs

3 -1 to Syncrude Canada, Ltd., which effluxes an estimated 43,000 m CH4 d of methane to the atmosphere, equivalent to about 500,000 cattle (Holowenko et al., 2000). Acetate and

H2/CO2 are the most important immediate precursors for methanogenesis (Conrad, 1999).

Acetate levels are around 0.41 - 0.79 mM throughout the lower layer (>7 mbs) of the ponds, as residual hydrocarbons are degraded through anaerobic processes (Ramos-Padrón et al., 2011).

Our previous study of the surface layer bacterial communities detected predominant organisms known to use methanol (unpublished), such as Xanthobacter, Rhodoferax, Hyphomicrobium,

Methylophilum. We hypothesize that these organisms consume the byproducts of methane oxidation, particularly methanol or formaldehyde. In this study, SIP analysis was applied to clarify how growth of bacterial populations was stimulated by acetate, methanol, and protein extracts, as external carbon sources under oxic conditions. We also studied tailings phototrophic communities, by applying constant light exposure to the surface OSPW sample. In wastewater environments such as the tailings ponds, phototrophs are capable of assimilating nutrients from the wastewater, and in turn converting solar energy into products that are consumables for immediate heterotrophs (Koblížek, 2015;

Umamaheswari & Shanthakumar, 2016). This study will help to detect what communities are feeding on the byproducts of hydrocarbon degradation, as well as the phototrophic communities in the tailings ponds.

4.2 Methods

4.2.1 Sampling and water chemistry

The West-In-Pit (WIP) of Syncrude Canada, Ltd., constructed in 1995, was primarily used as a storage facility for fluid fine tailings and recycle-water supply before its closure in

December 2012 (Syncrude Canada, 2010). The pond was closed for reclamation in 2012 to become the first model End-Pit Lake for the oilsands industry in Canada (Syncrude Canada,

2010). Before closure, the pond stored approximately 200 Mm3 of fluid fine tailings capped with a minimum of 5 m layer (30 Mm3) of water.

After less than two decades of operation, the Mildred Lake Settling Basin (MLSB), is still the largest oilsands tailings ponds operated by Syncrude Canada, Ltd. (Syncrude Canada, 2010).

With a total fluid volume of 209.9 Mm3, MLSB currently provides storage for fluid fine tailings, coke solids, flotation tailings, froth treatment tailings, and recycled water (Syncrude Canada,

2010). The samples used for all experiments in this study came from WIP surface water (0-10 cm) sampled in August 2011 and MLSB surface water sampled in July 2015. The chemical composition of both the WIP and MLSB OSPW and the water sampling methods were described previously (Saidi-Mehrabad et al., 2013). Throughout this chapter, OSPW sampled from WIP and MLSB tailings ponds are termed as WIP-OSPW and MLSB-OSPW respectively.

4.2.2 Stable isotope probing (SIP) and 16S rRNA gene sequencing

To study the microbial communities growing in response to the addition of intermediate

13 compounds in the tailings ponds, isotopically labelled methanol ( CH3OH 99 atom%, Sigma-

13 Aldrich), sodium acetate ( C2H3NaO2 99 atom%, Sigma-Aldrich), and protein extracts (Algal

crude protein extract-13C and 15N, 98 atom % 13C, 98 atom % 15N, Sigma-Aldrich), and carbon

13 dioxide ( CO2, 99 atom%, Sigma-Aldrich), were used as substrates.

To concentrate bacteria, 150 ml of fresh OSPW was filtered through a 0.22-μm filter. The filter and 20 ml of OSPW were added to a 100-ml serum bottle (equivalent to 170 ml of water and 80 ml of headspace). The filter and 20 ml of OSPW were added to a 100-ml serum vial

(equivalent to 170 ml of water sample and 80 ml of headspace) and supplemented with 0.05%

(w/w) of labelled sodium acetate, methanol, protein extract, or 10% of labelled CO2. Vials containing 20 ml of filtered OSPW without added substrate were used as controls. Vials were sealed with butyl rubber stoppers and incubated in the dark. Vials containing 10% of labelled

CO2 were incubated under constant light exposure. Headspace CO2 and O2 were monitored for all SIP-enrichments samples to determine incubation lengths. Incubations were terminated when

CO2 levels in the microcosms increased by around 0.003 – 0.006 mol for methanol, acetate and protein extracts.

Headspace CO2 ans O2 were monitored using a Varian 450-GC gas chromatograph equipped with a thermal conductivity detector (TCD) (150°C), after separation in a 2 mm × 0.5 m Hayesep N and a 2 mm × 1.2 m Molecular Sieve 16X column in series (70°C). CO2 production rates were calculated by linear regression of CO2 production versus time in the vial headspaces. All bottles were incubated at 23°C on a rotary shaker at 180 rpm to ensure aeration.

After 5 days (13C-sodium acetate), 7 days (13C-methanol), 14 days (13C-protein extract),

13 and 3 months ( CO2) of incubation, cells were collected by centrifugation for 10 min at 10,000 × g. DNA was extracted from the pellett using the FastDNA Spin Kit for Soil (MP Biomedicals) with additional purification steps using 5.5 M guanidine thiocyanate (Knief et al., 2003), and stored at −80°C. DNA was separated by isopycnic centrifugation in CsCl and fractionated as

described previously (Sharp et al., 2012). DNA extracted from OSPW incubated for the same time without addition of substrates was used as a control to determine the expected position of unlabelled DNA in CsCl gradients (OSPW-heavy). Heavy DNA fractions of all the labelled samples were used for 16S rRNA gene amplification and sequencing (Supplementary Figure

B1). Heavy fractions of OSPW (OSPW-heavy), and unfractionated DNA from fresh OSPW

(OSPW-control) were used as controls. Amplification of 16S rRNA genes was carried out as described previously (Sharp et al., 2014a). Purified PCR products (~150 ng total DNA) were sequenced using a 454 Life Sciences Genome Sequencer FLX (Roche) and analysed by QIIME as previously described (Caporaso et al., 2010; Sharp et al., 2014a).

4.3 Results

OSPW samples fed with 13C-substrates (methanol, acetate, and protein extract) showed

-1 increased CO2 production rates of 28 mmol, 0.39 mmol, and 14.4 mmol of CO2 day

-1 respectively, compared to 0.53 mmol of CO2 day for control sample (OSPW with no added substrates; Supplementary Figure B1). Density gradient separation of extracted DNA indicated that 13C incubations also showed a shift in the DNA distribution towards heavier values (Figure

4-1). The "heavy" DNA fractions representing the active 13C-assimilating communities were determined by comparing density gradients of the DNA extracted from the SIP incubations to that of the control samples. The fractions chosen for community analyses are indicated in Figure

4-1.

16S rRNA gene sequencing of natural OSPW, which is used as control, gave around

2,404 - 12,731 reads (Table 4-1). The results, which are summarized at the phylum/class level, show the predominance of Betaproteobacteria (9.6%), Alphaproteobacteria (7.3%) and

Flavobacteria (4.9%; Figure 4-2). Heavy fraction of OSPW fractionated by SIP (OSPW-heavy) showed the dominance of Alphaproteobacteria (28.8%) and Gammaproteobacteria (21.7%).

Methanol-SIP is dominated by Betaproteobacteria (57.2%), sodium acetate-SIP is dominated by

Alphaproteobacteria (74.4%), protein extract-SIP is dominated by Alphaproteobacteria (24.4%)

13 and Bacteroidetes (22.8%), CO2-MLSB-SIP is dominated by Planctomycetacia (45.4%) and

13 Alphaproteobacteria (27.2%), while CO2-WIP-SIP is highly dominated by Sphingobacteria

(25%), Alphaproteobacteria (24.2%) and Planctomycetacia (23.3%; Figure 4-2).

DNA extracted from OSPW incubated for the same time without addition of substrates was used as a control to determine the expected position of unlabelled DNA in CsCl gradients

(OSPW-heavy). Both the OSPW-heavy and OSPW-control 16S rRNA gene sequences show disparate community structures, suggesting that some species present have higher G+C contents and therefore higher densities. Results from OSPW-heavy sequencing show the predominance of

Alphaproteobacteria (28.9%) and Gammaproteobacteria (21.7%; Figure 4-2). Major species are

Devosia sp. (23.3% of 6,098 reads), Methylomarinum sp. (12.6% of 6,098 reads), and Thauera phenylacetica (9.0% of 6,098 reads). .

The most dominant OTUs of 13C-labelled sodium acetate-SIP substrates belong to the classes Rhizobiales and Caulobacterales of Alphaproteobacteria: Devosia riboflavina (20.7% abundance), 15.3% abundance (61.1 times enriched) belonged to Xanthobacter tagetidis with a

99% identity, and 9.1% abundance belonged to Brevundimonas nasdae with a 100% identity

(Figure 4-3; Table 4-1). Stable isotope probing of 13C-labelled protein extract in WIP-OSPW sample shows the predominance of Alphaproteobacteria (24.4%), Bacteroidetes (22.8%) and

Betaproteobacteria (14.7%). Closely related major proteolytic species to the OTUs detected are:

Acetobacteroides (1,089 out of 5,999 reads), Fusibacter (636 out of 5,999 reads) (Ben Hania et al., 2012), and Azospirillum sp. (368 out of 5,999 reads).

Figure 4-1. Relative DNA versus DNA density for SIP experiments of WIP-OSPW tailings

water sampled in August, 2011, with and without the addition of labelled substrates and

model hydrocarbons: sodium acetate (A), methanol (B), protein extract (C), MLSB

13 13 amended with CO2 (D), and WIP amended with CO2 (E). Dashed lines represent incubations of OSPW control (no substrate addition) and solid lines represent incubations

with 13C substrates. The y-axis indicates the relative DNA concentrations recovered from each fraction, where the highest quantity detected in any fractions is set to 1.0. Red arrows indicate the representative “heavy” fractions that were combined and used for 16S rRNA

gene pyrotag sequencing to identify the active communities, and black arrows indicate

“heavy” fraction of OSPW (OSPW-heavy) used in this study.

13 Both CO2 incubation of MLSB-and WIP-OSPW under light over the duration of 12 weeks are dominated by unidentified Planctomycetes, a rarely cultured phylum. MLSB-OSPW

13 amended with CO2 was dominated by an unidentified Planctomycete related to Aquisphaera sp.

(87% similarity; 145.5 times enriched) and an OTU that is related to Oceanibaculum indicum

(99% similarity; 636.6 times enriched; Table 4-1; Figure 4-4).

Figure 4-2. Community structures of oilsands process-affected water from the WIP tailings

pond (OSPW-control) and from the heavy fraction (OSPW-heavy), 13C-labelled DNA fractions of samples amended with 13C-methanol (Methanol), 13C-sodium acetate (Acetate),

13 13 13 C-protein extract (Protein extract), CO2-MLSB (light incubation), and CO2-WIP (light

incubation), incubated under aerobic conditions. Predominant taxa are identified to the

class level based on the SILVA database. The OSPW-heavy treatment is the community

detected only in the heaviest DNA fraction of OSPW. OSPW-control shows the entire

unseparated DNA from OSPW. 79

4.4 Discussion

Community analysis of 16S rRNA genes from the complete DNA extract obtained from

OSPW (OSPW-control) shows a dominance of Betaproteobacteria (9.6%), Alphaproteobacteria

(7.3%) and Flavobacteria (4.9%; Figure 4-3). The predominant species are Hydrogenophaga taeniospiralis (4.7% of 2,404 reads), Lutibacter maritimus (3.9% of 2,404 reads), and

Enhydrobacter aerosaccus (3.4%, of 2,404 reads; Supplementary Table B1). H. taeniospiralis is a hydrogen-oxidizing bacterium (Willems et al., 1989), of which the type strain was isolated from sediment layers and feeds on organic acids such as pyruvate, acetate, and succinate (Staley et al., 1987; Willems et al., 1989). A member of the genus Hydrogenophaga is known capable to degrade 4-Aminobenzenesulfonate (sulfanilate), a recalcitrant and xenobiotic compound found in industrial wastewater. Genes involved in the biodegradation of other aromatic compounds such as terephthalate, phenol, phenoxybenzoate, protocatechuate, and phenylacetate, were also detected in its genome (Gan et al., 2012).

Abundance of D. riboflavina was multiplied by 80.2 times in OSPW-heavy versus

OSPW-control, M. vadi is undetected in OSPW-control, and T. phenylacetica was multiplied by

44.8-fold (Table 4-1). Overall, 16S rRNA genes from the OSPW-complete sample shows high community diversity with only 1.8% of the highest single OTU dominance, while OSPW-heavy shows the predominance of D. riboflavina (23.3% abundance; Figure 4-2). This result suggests that OTU 4969 (96% similarity to D. riboflavina) seem to have higher G+C contents compared to other species and therefore tends to fall in fractions with heavier densities (1.73-1.75 g ml-1).

SIP of 13C-methanol was carried out to observe active methylotrophs in WIP-OSPW, organisms that can use reduced one-carbon compounds as their carbon source, such as methanol, which is presumably abundant in the WIP tailings surface water as a result of methane oxidation.

Betaproteobacteria and Alphaproteobacteria were dominant in the heavier methanol SIP fractions (fractions 3 – 4), also with M. universalis as the most dominant OTU (53.0% abundance), verifying its methylotrophic capability (Figure 4-3). The most dominant OTUs in the 13C-methanol SIP are M. universalis (53.0% abundance, 25.6 times enriched),

Desulforhabdus sp. (3.5% abundance, 35.5 times enriched), and Hyphomicrobium sp. (2.1% abundance, 15.0 times enriched). Other than M. universalis, all other dominant OTUs of the methanol-SIP heavy fractions show <97% identity to any cultured species (Supplementary Table

B1-C).

On top of the major OTUs, some species such as Delftia acidovorans and

Propionibacterium acnes are highly enriched in the methanol enrichment sample (>50 times enriched; Table 4-1). Both members of the Delftia and Propionibacteriaceae were found in diverse habitats, from dairy products, sludge, soil, water, sewage treatment plants, to Antarctic sandstone (Stackebrandt, 2014; Stolze et al., 2012), but their roles in the methanol enriched sample is unknown.

Methyloversatilis universalis (OTU 5106), predominant in the 13C-methanol heavy fraction, is a methyl-oxidising bacterium (Kalyuzhnaya et al., 2006). Its dominance (79.0 times higher than OSPW) in the sample verifies its methylotrophic capability. Bacterial lineages of this species show exceptionally versatile metabolic capabilities, related to their ability to utilize single carbon compounds, such as methanol and methylated amines (Good et al., 2015;

Kalyuzhnaya et al., 2008). Genome analyses of M. universalis indicate that it possesses a heme- containing amine dehydrogenase (Qhp), which can be linked to aliphatic and aromatic amine oxidizing capabilities (Good et al., 2015), and a recent bioinformatics approach discovered an uncultured bacterium related to M. universalis strain FAM5 (98% identity on

BLASTN) that is a benzene degrader (Satija & Gore, 2014). Previous studies have also reported its presence in various petroleum related environments, but its relative abundance never reached more than 18% and its roles were not deemed crucial in hydrocarbon degradation (Golby et al.,

2012; Lenchi et al., 2013; Noguchi et al., 2014). This is probably why the same OTU (OTU

5106) also appeared in the 13C-benzene SIP with around 87% of abundance (Chapter 3).

Acetate is a common intermediate in the oxidation of aldehydes and aromatics, and is detected (0.41 - 0.79 mM) in the deeper sections (>7 m) of Alberta tailings ponds (Ramos-

Padrón et al., 2011). Of all the three major OTUs detected, X. tagetidis is probably the one responsible for acetate degradation in the tailings pond (61.1-fold increase compared to OSPW;

Table 4-1). Type strain of X. tagetidis was shown able to grow on various acids such as formate, acetate, and propionate (Oren, 2014). The other two major OTUs, D. riboflavina and B. nasdae, were present in OSPW heavy fractions with approximately the same percent coverage (less than

1.1% difference), indicating that they might have higher G+C contents than other species in the community. Although some Devosia isolates have shown promising biodetoxification abilities that could lead to their utilization as detoxifying agents (Hassan et al., 2014; Sato et al., 2012;

Zhou & He, 2010), the role of D. riboflavina in the tailings ponds is unknown.

(OSPW) samples. DNA from the samples were then fractionated by Stable isotope probing (SIP). The table presents the heavy fractions of the SIP. OSPW-heavy: heavy fraction of OSPW receiving the same treatment as all of the amended samples. UD: undetected.

x-fold Relative % % Identity Relative % enrichment abundance SIP-Sample OTU to the Number of Abundance in compared to Taxonomy (Phylum; Class) Closest relative (BLAST) in control (reads) ID closest reads DNA heavy control heavy relative fractions heavy fraction fraction Methanol 5106 Proteobacteria; Betaproteobacteria Methyloversatilis universalis 100 4945 52.96 2.07 25.58 9,434 197 Proteobacteria; Alphaproteobacteria Hyphomicrobium methylovorum 96 199 2.13 0.06 35.52 2961 Proteobacteria; Betaproteobacteria Azoarcus communis 99 138 1.48 0.10 15.08 4723 Proteobacteria; Betaproteobacteria Delftia acidovorans 100 105 1.12 UD >65.96 816 Actinobacteria; Propionibacteriales Propionibacterium acnes 99 82 0.88 UD >52.24

820 Actinobacteria; Acidimicrobiia Aciditerrimonas ferrireducens 91 78 0.84 0.07 12.73 337 Bacteroidetes; Sphingobacteriia Sediminibacterium goheungense 96 53 0.57 UD >19.00 4209 Armatimonadetes; Fimbriimonadia Fimbriimonas ginsengisoli 93 50 0.54 UD >18.00 6031 Planctomycetes; Planctomycetia Pirellula staleyi 96 38 0.41 0.03 12.72 6504 Planctomycetes; Planctomycetia Planctomicrobium piriforme 92 32 0.34 UD >11.33

10 Chlamydiae; Chlamydiae Estrella lausannensis 90 32 0.34 0.02 20.90

5098 Planctomycetes; Planctomycetia Planctomyces brasiliensis 98 30 0.32 0.02 19.59

2301 Proteobacteria; Alphaproteobacteria Methylobacterium fujisawaense 100 30 0.32 0.02 19.59

4341 Actinobacteria; Acidimicrobiia Ilumatobacter nonamiensis 98 28 0.30 0.02 18.28 Acetate 3316 Proteobacteria; Alphaproteobacteria Xanthobacter tagetidis 99 605 8.01 0.13 61.09

7,549 2492 Proteobacteria; Alphaproteobacteria Novosphingobium barchaimii 98 274 3.63 0.15 24.59

Proteobacteria; Betaproteobacteria Azoarcus communis 99 115 1.52 0.03 46.45 2961 Proteobacteria; Alphaproteobacteria Phenylobacterium falsum 100 104 1.38 0.05 28.00 1126 Planctomycetes; Planctomycetia Aquisphaera giovannonii 90 77 1.02 0.03 31.10 3517 Proteobacteria; Alphaproteobacteria Elstera litoralis 100 39 0.52 0.03 15.75 1158 Planctomycetes; Planctomycetia Planctomicrobium piriforme 91 33 0.44 UD >26.66 6751 Proteobacteria; Alphaproteobacteria Labrenzia alexandrii 99 32 0.42 UD >25.85 3083 Actinobacteria; Rubrobacteria Gaiella occulta 91 27 0.36 0.02 21.81 4743 Protein Firmicutes; Clostridia Fusibacter tunisiensis 94 1089 18.15 1106.97 extract 5576 0.02 5,999 5803 Proteobacteria; Alphaproteobacteria Azospirillum rugosum 93 636 10.60 UD >646.50 79 Proteobacteria; Betaproteobacteria Aquaspirillum serpens 99 231 3.85 UD >234.81

892 Proteobacteria; Alphaproteobacteria Defluviimonas alba 100 224 3.73 0.31 11.98

890 Proteobacteria; Betaproteobacteria Janthinobacterium lividum 100 213 3.55 UD >216.52

5880 Proteobacteria; Alphaproteobacteria Azospirillum palatum 92 209 3.48 UD >212.45

4082 Firmicutes; Clostridia Fusibacter tunisiensis 94 178 2.97 UD >180.94

4938 Spirochaetae; Spirochaetes Treponema bryantii 93 156 2.60 UD >158.57

3358 Proteobacteria; Betaproteobacteria Acidovorax caeni 99 108 1.80 0.02 109.78

1974 Bacteroidetes; Bacteroidia Acetobacteroides hydrogenigenes 96 87 1.45 UD >88.44 Proteobacteria; Citrobacter freundii 99 >74.20 117 Gammaproteobacteria 73 1.22 UD Proteobacteria; Aeromonas hydrophila 100 69.12 3382 Gammaproteobacteria 68 1.13 0.02

5257 Proteobacteria; Deltaproteobacteria Desulfatiferula berrensis 96 62 1.03 UD >63.02

4693 Bacteroidetes; Bacteroidia Anaerocella delicata 94 61 1.02 UD >62.01

2602 Proteobacteria; Deltaproteobacteria Desulfovibrio idahonensis 99 59 0.98 0.05 19.99

4131 Proteobacteria; Deltaproteobacteria Desulfovibrio psychrotolerans 98 58 0.97 UD >58.96

4328 Proteobacteria; Alphaproteobacteria Xanthobacter autotrophicus 98 55 0.92 UD >55.91

5412 Proteobacteria; Alphaproteobacteria Azospirillum doebereinerae 99 55 0.92 UD >55.91

817 Proteobacteria; Deltaproteobacteria Desulforhabdus amnigena 97 51 0.85 UD >51.84

4841 Firmicutes; Clostridia Fusibacter bizertensis 94 50 0.83 UD >50.83

4310 Firmicutes; Bacilli Paenibacillus turicensis 88 45 0.75 UD >45.74

4796 Bacteroidetes; Bacteroidia Anaerocella delicata 89 45 0.75 UD >45.74

200 Firmicutes; Clostridia Fusibacter tunisiensis 91 38 0.63 UD >38.63

1736 Proteobacteria; Deltaproteobacteria Desulfomicrobium hypogeium 100 35 0.58 UD >35.58

4656 Elusimicrobia; Elusimicrobia Elusimicrobium minutum 87 34 0.57 UD >34.56

640 Proteobacteria; Alphaproteobacteria Rhizobium selenitireducens 100 33 0.55 0.02 33.54

1212 Spirochaetae; Spirochaetes Sphaerochaeta globosa 98 31 0.52 UD >31.51

5935 Firmicutes; Clostridia Fusibacter tunisiensis 91 30 0.50 UD >30.50

1126 Proteobacteria; Alphaproteobacteria Phenylobacterium falsum 100 29 0.48 0.03 14.74

6395 Spirochaetae; Spirochaetes Sphaerochaeta globosa 98 29 0.48 UD >29.48

3066 Firmicutes; Bacilli Trichococcus pasteurii 100 28 0.47 UD >28.46

1938 Proteobacteria; Betaproteobacteria Comamonas testosteroni 99 24 0.40 UD >24.40

5807 Proteobacteria; Alphaproteobacteria Aquaspirillum polymorphum 97 22 0.37 UD >22.36

1634 Proteobacteria; Alphaproteobacteria Xanthobacter autotrophicus 98 21 0.35 UD >21.35

3124 Firmicutes; Clostridia Clostridium sticklandii 95 20 0.33 UD >20.33

3438 Proteobacteria; Alphaproteobacteria Azospirillum lipoferum 94 20 0.33 UD >20.33

360 Proteobacteria; Alphaproteobacteria Cloacibacillus porcorum 91 18 0.30 UD >18.30

2565 Firmicutes; Clostridia Clostridium sticklandii 100 17 0.28 UD >17.28

2416 Spirochaetes; Spirochaetales Treponema stenostreptum 93 17 0.28 UD >17.28

4459 Proteobacteria; Deltaproteobacteria Desulfobacula phenolica 94 17 0.28 UD >17.28

2521 Firmicutes; Clostridia Christensenella minuta 90 16 0.27 UD >16.26

6539 Planctomycetes; Planctomycetia Planctomyces limnophilus 95 15 0.25 0.02 15.25

MLSB-CO2 6596 Planctomycetes; Planctomycetacia Aquisphaera giovannonii 87 5299 45.11 0.31 145.50 11,748 6288 Proteobacteria; Alphaproteobacteria Oceanibaculum indicum 99 1227 10.44 UD >636.63 Bacteroidetes; Sphingobacteriia Haliscomenobacter hydrossis 92 >377.36 6340 727 6.19 UD Proteobacteria; Luteimonas aestuarii 97 >223.72 758 Gammaproteobacteria 431 3.67 UD Proteobacteria; Alphaproteobacteria Phenylobacterium zucineum 99 74.23 97 286 2.43 0.03 Chloroflexi; Anaerolineae Anaerolinea thermophila 90 >131.84 4533 254 2.16 UD Proteobacteria; Alphaproteobacteria Amphiplicatus metriothermophilus 97 27.90 2094 215 1.83 0.07 Bacteroidetes; Bacteroidetes Order II. >106.93 1338 Incertae sedis Salinibacter ruber 87 206 1.75 UD

3386 Acidobacteria; Solibacterales Bryobacter aggregatus 90 199 1.69 0.13 12.91

6690 Proteobacteria; Alphaproteobacteria Amphiplicatus metriothermophilus 90 145 1.23 0.05 25.09

5644 Proteobacteria; Alphaproteobacteria Rhodobacter megalophilus 99 138 1.17 0.10 11.94

2174 Proteobacteria; Alphaproteobacteria Bauldia consociata 99 113 0.96 0.07 14.66

5633 Proteobacteria; Alphaproteobacteria Azospirillum brasilense 92 81 0.69 UD >42.04

4360 Proteobacteria; Alphaproteobacteria Blastomonas natatoria 99 80 0.68 UD >41.53

6373 Chloroflexi; Anaerolineae Leptolinea tardivitalis 99 73 0.62 UD >37.89

6539 Planctomycetes; Planctomycetia Planctomyces limnophilus 95 70 0.60 0.02 36.33

16 Proteobacteria; Alphaproteobacteria Aureimonas altamirensis 96 64 0.54 UD >33.22

3976 Chloroflexi; Caldilineae Caldilinea aerophila 89 62 0.53 UD >32.18

6112 Proteobacteria; Alphaproteobacteria Thalassobaculum litoreum 96 61 0.52 UD >31.66

2517 Proteobacteria; Alphaproteobacteria Prosthecomicrobium pneumaticum 100 40 0.34 UD >20.76

4780 Proteobacteria; Alphaproteobacteria Roseomonas stagni 95 37 0.31 UD >19.21

233 Actinobacteria; Micrococcales Chryseoglobus frigidaquae 99 35 0.30 0.02 18.17

WIP-CO2 6340 Bacteroidetes; Sphingobacteriia Haliscomenobacter hydrossis 92 3050 23.96 0.02 1460.91 12,731 6915 Planctomycetes; Planctomycetia Thermogutta hypogea 91 1971 15.48 UD >944.09 3386 Acidobacteria; Solibacterales Bryobacter aggregatus 90 754 5.92 0.13 45.14

2174 Proteobacteria; Alphaproteobacteria Bauldia consociata 99 511 4.01 0.07 61.19

740 Planctomycetes; Planctomycetia Gemmata obscuriglobus 89 343 2.69 UD >164.29

3530 Acidobacteria; Solibacteres Paludibaculum fermentans 91 334 2.62 0.03 79.99

640 Proteobacteria; Alphaproteobacteria Rhizobium selenitireducens 100 332 2.61 0.02 159.02

4533 Chloroflexi; Anaerolineae Anaerolinea thermophila 90 320 2.51 UD >153.28

4674 Actinobacteria; Micrococcales Mesorhizobium ciceri 99 284 2.23 0.11 19.43

6373 Chloroflexi; Anaerolineae Leptolinea tardivitalis 84 258 2.03 UD >123.58

4716 Planctomycetes; Planctomycetia Thermogutta hypogea 89 243 1.91 0.13 14.55

1575 Planctomycetes; Planctomycetia Thermogutta hypogea 87 199 1.56 Ud >95.32

6421 Planctomycetes; Planctomycetia Thermogutta hypogea 88 181 1.42 0.02 86.70

6869 Acidobacteria; Acidobacteriales Granulicella aggregans 88 147 1.15 UD >70.41

2721 Proteobacteria; Alphaproteobacteria Brevundimonas viscosa 97 129 1.01 UD >61.79

1032 Chloroflexi; Caldilineae Litorilinea aerophila 90 95 0.75 UD 45.50

5949 Bacteroidetes; Sphingobacteriia Chitinophaga eiseniae 93 83 0.65 0.05 13.25

5633 Proteobacteria; Alphaproteobacteria Azospirillum brasilense 92 78 0.61 UD >37.36

3978 Actinobacteria; Micrococcales Micrococcus yunnanensis 100 77 0.60 0.03 18.44

1190 Cyanobacteria; Oscillatoriophycideae Leptolyngbya schmidlei 98 71 0.56 UD >34.01

2514 Planctomycetes; Planctomycetia Thermogutta hypogea 88 69 0.54 0.02 33.05 Proteobacteria; >32.09 5696 Gammaproteobacteria Thiofaba tepidiphila 97 67 0.53 UD

2947 Chloroflexi; Caldilineae Caldilinea aerophila 90 63 0.49 UD 30.18

1126 Proteobacteria; Alphaproteobacteria Phenylobacterium falsum 100 61 0.48 0.03 14.61

2824 Planctomycetes; Planctomycetia Thermogutta hypogea 92 59 0.46 UD >28.26

97 Proteobacteria; Alphaproteobacteria Phenylobacterium zucineum 99 56 0.44 0.03 13.41

1363 Planctomycetes; Planctomycetia Nostocoida limicola 85 43 0.34 UD >20.60

58 Planctomycetes; Planctomycetia Thermogutta hypogea 87 33 0.26 UD >15.81

DNA from fresh OSPW (OSPW) were used as controls.

Stable isotope probing of 13C-labelled protein extract in WIP-OSPW sample shows the predominance of Acetobacteroides hydrogenigenes, Fusibacter sp., and Azospirillum sp. Both

13 CO2 incubation of MLSB-and WIP-OSPW under light over the duration of 12 weeks are dominated by unidentified Planctomycetes, related to Aquisphaera sp, a chemoheterotrophic

Planctomycete known to feed on sugars and organic acids (Bondoso et al., 2011). Another dominant OTU identified under light incubated sample is related to Oceanibacilum (Table B-1).

The type strain of Oceanibaculum indicum is an aerobic, nitrate-reducing bacterium, isolated from a polycyclic aromatic hydrocarbon-degrading consortium, obtained from a deep-seawater sample of the Indian Ocean (Lai et al., 2009). Although this species has been detected mostly in

PAH-degrading environments (Lai et al., 2009; Yuan et al., 2015), it hasn't been linked to any hydrocarbon-degrading activities, therefore its roles remain unknown.

Other highly enriched OTUs (>100 times enriched; Table 4-1) are related to

Haliscomenobacter sp., Luteimonas sp., Anaerolinea sp., and Salinibacter sp. It is unclear as to why these species thrived under constant light exposure, since none of them can be related to photosynthetic activities. Haliscomenobacter sp., a filamentous bacteria, is commonly found in wastewater treatment environments and is known responsible for biomass bulking (Cao & Lou,

2016; Daligault et al., 2011), members of Luteimonas are mostly sedimentary organisms (Fan et al., 2014; Romanenko et al., 2013), Anaerolinea hydrocarbon contaminated wastewater (Yadav 89

et al., 2015), and Salinibacter, a halophilic bacterium (Oren, 2013). OSPW is known to have a high salt content (about 2.2 g l−1), with high contents of various metals, sulfides and hydrocarbons, which cause the above organisms to thrive with or without light exposure (Saidi-

Mehrabad et al., 2013).

WIP-OSPW heavy SIP fraction sample incubated under constant light (WIP-OSPW

3 amended with CO2) is dominated by Bacteroidetes and unidentified Planctomycetes. Two dominant OTUs are related to Haliscomenobacter sp. (92% similarity, 1460.9 times enriched), and Thermogutta sp. (91% similarity, >1,000 times enriched). Haliscomenobacter is a sheath- forming bacterium that grows on simple sugars and organics (Krul, 1977), and has mostly been detected in many wastewater environments (Austermann-Haun et al., 1998; Kragelund et al.,

2008; Morgan-Sagastume & Allen, 2003). Thermogutta is a facultatively anaerobic

Planctomycete (Slobodkina et al., 2015). The phylum Planctomycetes has been commonly associated with a wide range of algae and marine flowering plants, in which they grow as complex microbial biofilm and depend on polysaccharides secreted by algae (Bream; Kim et al.,

2016; Lage & Bondoso, 2015; Yoon et al., 2014). The phylum Planctomycetes has a distinctive morphology as its members appear to contain an unusual membrane-bound internal compartments, often referred to as the paryphoplasm, or a peripheral ribosome-free region, and an interior region (includes nucleoid regions and ribosome-like particles) (Lindsay et al., 2001).

During sample incubation, constant light exposure might have caused algae indigenous to OSPW to thrive. It has been reported before that micro-algae, such as Parachlorella kessleri, thrived on the surface of the tailings ponds, feeding on mainly nitrogen, phosphorus and heavy metals

(Mahdavi et al., 2012).

Other interesting OTUs that are >100 times enriched include two Chloroflexi species:

Anaerolinea sp. and Leptolinea sp. (Table 4-1). Members of the class Chloroflexi consist of phototrophic bacteria with filamentous morphology (Hanada, 2014), which is probably the reason why this group of organism thrive under light.

13 All of the OSPW photosynthetic samples amended with CO2 are dominated by unidentified Planctomycetes (>40% of all reads; Figure 4-4; Table 4-1), a rarely cultured

Phylum, suggesting a possible new method of cultivation for this phylum. This known relationship of Planctomycetes with photosynthetic organisms can lead to a hypothesis that

Planctomycetes indigenous to tailings ponds can be isolated from algal biofilm using cultivation techniques adapted for these organisms. Presence of algae in the surface OSPW incubated under lights may be related to high levels of algal secondary metabolites: organic compounds, amino acids, and sugars (McGenity et al., 2012) in OSPW that provide both the carbon and energy sources to the indigenous microbial community. These simple compounds in OSPW may have stimulated the growth of simple sugars and organic acid feeders in the community, which might as well perform hydrocarbon-degrading activities (McGenity et al., 2012).

Figure 4-4. Phylogenetic tree based on 16S rRNA gene sequences of the 5 most abundant

OTUs of OSPW-control (green), 13C-methanol SIP-heavy fraction (red), 13C-acetate SIP

13 13 13 (dark blue), C-protein extract SIP (orange), CO2 in MLSB-OSPW SIP (blue), CO2 in

WIP-OSPW SIP (black). To create the most reliable phylogeny, reference sequences (>1400

bp; black) from NCBI database (04/05/2016) were obtained. Tree was constructed via

Maximum Likelihood with 1,000 bootstraps using Seaview Phylogeny (Gouy et al., 2010).

For comparison, total number of reads of each OTU is given in parentheses (entire dataset

was 49,186 reads). Scale bar represents 0.1 change per nucleotide position. Bootstrap

values >50% are indicated for key nodes.

4.5 Conclusions

The surface layer of the oilsands process-affected water (OSPW) in Alberta contain diverse bacterial communities with various roles. Predominant OTU is related to

Hydrogenophaga, a genus that can be linked to degradation of xenobiotic compounds. The rest of the microbial community consist of methylotrophs, simple sugars and organic acid feeders, and others with unknown roles. Sequencing results based on DNA from untreated OSPW is predominated by Betaproteobacteria, OSPW heavy fraction by Alphaproteobacteria, methanol-

SIP by Betaproteobacteria, sodium acetate-SIP by Alphaproteobacteria, protein extract-SIP by

13 13 Bacteroidia, CO2-MLSB-SIP and CO2-WIP-SIP by Planctomycetia. All of the OSPW

13 photosynthetic samples amended with CO2 are dominated by unidentified Planctomycetes

(>40% of all reads), a rarely cultured phylum, suggesting a possible new method of cultivation for this phylum.

Preface

Chapter 5 explores the metagenome results of the WIP-OSPW sample from the previous chapter (Chapter 3). Here, a novel CuMMO-encoding operon was detected in the sample, grouping phylogenetically away from other previously known CuMMO operons. This operon was investigated in depth, using SIP, quantitative PCR, and a RNA expression study. 16S rRNA

13 13 13 13 gene sequencing coupled with SIP of C-alkanes ( CH4, C2H6, and C3H8) were employed to study levels of CuMMO operon in response to added substrates and survey the microbial community responsible to 13C-alkanes degradation.

In this Chapter, Drs. Peter Dunfield and Ivica Tamas screened for the novel CuMMO operon described in this Chapter from the WIP-OSPW metagenome data, and then contrasted it against previously known CuMMO operons. Roshan Khadka provided the latest pmoA- phylogeny database, which is presented as a pmoA gene based tree in this Chapter. A former colleague, Alireza Saidi-Mehrabad, performed the WIP-OSPW sample processing for metagenome sequencing. Dr. Peter Dunfield also provided scientific advice in exploring this operon.

Chapter Five: Functional analysis of a novel Cu-monooxygenase detected via metagenomic analysis of OSPW

5.1 Introduction

Alkanes are saturated hydrocarbons with a general chemical formula of CnH2n+2 (Smith,

2011). The most important commercial sources for alkanes are natural gas and crude oil, although many living organisms also produce them for protection and survival mechanisms

(Thom et al., 2007). Some microorganisms are known capable of metabolizing alkanes, utilizing them for both carbon and energy sources (Rojo, 2010).

Aerobic alkane degradation begins by the oxidation of a terminal methyl group into an alcohol, which is further oxidized to aldehyde, converted into a fatty acid, to be further processed by β-oxidation to generate acetyl-CoA (Rojo, 2010; Van Agteren et al., 2013). Microbial methane monooxygenase reactions involve the initial incorporation of oxygen into CH4, forming alcohols (Bürgmann, 2011; Prosser, 1989; Vajrala et al., 2013). A number of enzyme families in some bacterial strains can be related to the initial alkane hydroxylation (Table 2-1). Short-chain- length alkanes (C1-C4) can be linked to a set of enzymes related to sMMO (soluble methane monooxygenase), and long-chain-length alkenes (>C5) tend to contain the non-heme iron monooxygenases (Rojo, 2010).

The particulate methane monooxygenase (pMMO), the key enzyme in aerobic methane oxidation, is a member of the Cu-containing membrane-bound monooxygenases (CuMMO) enzyme family (Hainbuch, 2015; Murrell, 2010). pmoA, the gene encoding the α subunit of pMMO, is highly conserved, making this gene a suitable biomarker for detecting methanotrophs with pMMO activity (McDonald & Murrell, 1997). pmoA and amoA (encoding

ammonia monooxygenase) share a high degree of sequence identity (Holmes et al., 1995), and they have been utilized as environmental functional markers to detect natural methane and ammonia-oxidizing populations (McDonald & Murrell, 1997; Rotthauwe et al., 1997).

Only recently, it has been discovered that organisms containing the CuMMOs are capable of ethylene and C2–C4 alkane oxidation (Beman et al., 2008; Coleman et al., 2012; Tavormina et al., 2011, Suzuki et al., 2012). This enzyme group is deemed important as it contributes to the global cycling of methane, nitrogen, and CO2, as well as the degradation of hydrocarbon compounds (Stein et al., 2012; Coleman et al., 2012; Rotthauwe et al., 1997). A deeper understanding of this enzyme group enables us to see their roles outside methanotrophy and ammonia oxidation (Coleman et al., 2012; Hainbuch, 2015; Nielsen et al., 1997).

In this study, we discovered an operon possibly encoding a novel CuMMO via metagenomic analysis of WIP-OSPW. Represented by several full operon sequences, the operon was found on large contigs and were phylogenetically distinct from other previously known

CuMMOs. A further study was done to look into the enzyme capabilities via 1) a modified SIP

13 13 13 technique that was developed based on labeling with CH4, C2H6, and C3H8, which are suspected alkane substrates, followed by 16S rRNA gene sequencing, and via 2) qPCR to specifically detect the targeted gene in all of the alkane-SIP fractions, using an assay specific to the novel CuMMO-encoding operon. Summarized workflow of this Chapter is presented in

Supplementary Figure C1, modified from a previous SIP figure (Friedrich, 2006).

5.2 Methods

5.2.1 Sampling and water chemistry

OSPW sample used for metagenomic analysis was sampled from the West-In-Pit (WIP) pond of Syncrude Canada, Ltd. A description of the WIP pond can be found in Chapter One, and

MLSB pond was described in Chapter Four. Throughout this article, OSPW sampled from WIP tailings pond is termed as WIP-OSPW. OSPW sampled from MLSB tailings pond is termed as

MLSB-OSPW, and the novel CuMMO-encoding operon described in this Chapter is termed xmoCAB. MLSB surface water (0-10 cm) used for this experiment was sampled in September

2014, and March and Auguast 2015.

5.2.2 qPCR primer design

Two sequences grouping together were detected from WIP-OSPW metagenome analysis, separate from CuMMO operons. The longest contig sequence of the two was singled out for further analysis (Supplementary Table C1). A primer set xmoCAB-f

(5’AAATGACTTCGCTCGCTG 3’) and xmoCAB-r (5’CATGCTGCCACCTTCTTT 3’) based on the sequence was designed by using 'Probe Design' tool in ARB that searches for potential target sites (Ludwig et al., 2004a), with the end product of 271 bp. DNA extracted from fresh

WIP-OSPW from September 2014 was used as a positive control, from which the target gene was amplified via PCR. Cycling conditions were performed with a three-step reaction: an initial denaturation of 5 min at 94°C; 35 cycles of 94°C for 30 s, 55°C for 30 s and 72°C for 30 s; and a final elongation step of 72°C for 10 min. Control PCR product was then used for cloning of the gene into an E. coli vector by following the CloneJET PCR Cloning Kit (Thermo Fisher

Scientific, Waltham, MA, USA) protocols. PCR parameters were subsequently optimized on a

pure culture of transformed E. coli containing the vector and xmoCAB gene insert via gradient

PCR (48–58°C), optimal conditions determined based on the brightness of the PCR gel bands seen under blue-lights. Optimal condition of 56°C was selected (Supplementary Figure C2). PCR products were verified via 16S rRNA gene Sanger sequencing done at the University of

Calgary Genetic Analysis laboratory. Sequenced products displayed 100% similarity to the original target genes from the WIP-OSPW metagenome.

Quantitative PCR was performed for both the MLSB-OSPW hydrocarbon enrichments and SIP fractions, using primers xmoCAB-f and xmoCAB-r (this study). PCR assays for the primer pair was prepared as described previously (Sharp et al., 2012). Cycling conditions were performed with a three-step reaction: an initial denaturation of 5 min at 94°C; 35 cycles of 94°C for 60 s, 56°C for 45 s and 72°C for 45 s; and a final elongation step of 72°C for 10 min. The measured DNA amount could be converted to target molecules per ml of OSPW. PCR products of the inserted E. coli vector were used as xmoCAB standards. Standards for qPCR were prepared as per Sharp et al. (2014). Measured DNA amount was converted to target molecules per microliter, and dilutions were adjusted to 108, 106, 104 and 102 target molecules μl-1 (Sharp et al.,

2014b). Fluorescence data was obtained during the last step of each cycle. The specificity of each reaction was verified by melt curve analysis (data not shown). Efficiencies for standard curves used to calculate qPCR values ranged from 90–97%. Gene copy numbers were calculated as shown in Supplementary Table C3.

5.2.3 MLSB-OSPW samples enrichments

MLSB-OSPW samples were enriched by using the following hydrocarbon substrates: methane (Sigma), ethane (Sigma), propane (Sigma), benzene (Alfa Aesar), toluene (Sigma), p-

xylene (Sigma), and naphthalene (Sigma). Aromatics were tested to see the possibility of xmoCAB involvements in their degradation processes. To concentrate bacteria, 150 ml of fresh

OSPW was filtered through a 0.22-μm filter. The filter and 20 ml of OSPW were added to a 100- ml serum bottle (equivalent to 170 ml of OSPW and 80 ml of headspace). Each sample was run in duplicate. Vials containing 20 ml of milliQ water were used as abiotic leakage controls. Vials were sealed with butyl rubber stoppers (20 mm).

Water samples were filtered as described above. Twenty ml of filtered OSPW were added to a 100-ml serum vial and supplemented with 10% v/v methane, ethane, or propane, and 5 ml l-1 of benzene, toluene, p-xylene, or naphthalene, were added into 20 ml filtered samples. Each sample was run in duplicate or triplicate. 100-ml serum vials containing 20 ml of filtered OSPW were used as controls. Vials were sealed with butyl rubber stoppers. Bottles were incubated at

23°C on a rotary shaker at 180 rpm in the dark. After 1, 3, 6, and 8 weeks for methane, ethane, and propane, and 1, 2, and 4 weeks of other hydrocarbons, microcosms were opened, and samples were extracted and processed. All bottles were incubated at 23°C on a rotary shaker at

180 rpm to allow aeration. After incubation, cells were collected by centrifugation for 10 min at

10,000x g. DNA was extracted based on the FastDNA Spin Kit for Soil (MP Bio) and stored at

−80°C for qPCR and 16S rRNA gene sequencing.

5.2.4 Stable isotope probing preparation

To see if microbial communities capable of methane, ethane, and propane oxidation

13 relate to the levels of xmoCAB in OSPW, isotopically labelled methane ( CH4 99 atom%,

13 13 Sigma-Aldrich), ethane ( C2H6 99 atom%, Sigma-Aldrich), and propane ( C23H8 99 atom%,

Sigma-Aldrich), were used as substrates.

150 ml of OSPW (sampled in August 2015) were added to 1-l incubation bottles. 10% v/v of methane, ethane, and propane were added. Each sample was run in duplicate or triplicate.

Bottles containing 150 ml of filtered OSPW with no added substrates were used as controls.

Bottles were sealed with butyl rubber stoppers and incubated at 23°C on a rotary shaker at 180 rpm in the dark. After 10 days of incubation, bottles were opened, and samples were extracted and processed as described above.

Headspace CH4, C2H6, and C3H8 were monitored using a Varian 450-GC gas chromatograph equipped with a thermal conductivity detector (TCD) (150°C), after separation in a 2 mm × 0.5 m Hayesep N and a 2 mm × 1.2 m Molecular Sieve 16X column in series (70°C).

CO2 oxidation rates were calculated by linear regression of CO2 production versus time in the vial headspaces. All bottles were incubated at 23°C on a rotary shaker at 180 rpm to ensure aeration.

When approximately 5% of the CH4, C2H6, and C3H8 (10 days) was consumed, cells were collected by centrifugation for 10 min at 10,000 ×g. DNA was extracted as described above and stored at −80°C. DNA was separated by isopycnic centrifugation in CsCl and fractionated as described previously (Sharp et al., 2012). DNA extracted directly from the same OSPW incubated for the same time without addition of substrates was used as a control to determine the expected position of unlabelled DNA in CsCl gradients. Heavy DNA fractions of both the labelled and unlabelled samples were used for 16S rRNA gene amplification and sequencing.

5.2.5 16S rRNA gene sequencing and analysis

Around 4-5.0 ng/uL of input DNA (13C-labelled) was PCR amplified based on the KAPA

HiFi HotStart ReadyMix protocols (Kapa Biosystems), which is provided in Supplementary

100

Table C2. Primers used were 16S rRNA primer set: 9fw and 1492r (Eden et al., 1991). Cycling conditions were performed with a three-step reactions: an initial denaturation of 3 min at 95°C;

25 cycles of 95°C for 30 s, 55°C for 30 s and 72°C for 30 s; and a final elongation step of 72°C for 5 min. To confirm the presence of anticipated PCR reaction products, each reaction was run on an agarose gel. PCR products were purified using AMPure XP beads (Beckman Coulter).

Purified PCR products containing 16S rRNA genes were re-amplified from the labelled gradient fractions using Illumina barcoded amplicon primers containing 16S rRNA gene targeted primers

341fw and 785r at their 3ʹ′-ends. Second PCR reaction were rerun through agarose gel and purified via the methods described above. After Qubit quantification, DNA samples were normalized into 4 nM. High Sensitivity BioAnalyzer chip was used to analyze amplicon pools going on the same Illumina Miseq lane. Total genomic DNA was subjected to high-throughput sequencing using Illumina Miseq at the Harrison Laboratory, University of Calgary. Sequence output was analyzed by via Illumina Basespace 16S Metagenomics V1.0 software and QIIME

(Heruth et al., 2016; Kuczynski et al., 2012).

5.3 Results and discussion

Aerobic methane and ammonia oxidation are carried out by microorganisms possessing either methane monooxygenase (MMO) or ammonia monooxygenase (AMO) of the copper- containing membrane-bound monooxygenase superfamily (CuMMOs) (Stein et al., 2001).

CuMMOs are heterotrimers and are commonly genetically encoded in the subunit order CAB

(Hatzenpichler, 2012). Recently, it has been discovered that organisms containing the CuMMOs are capable of ethylene and C2–C4 alkanes oxidation (Beman et al., 2008; Coleman et al., 2012;

Tavormina et al., 2011, Suzuki et al., 2012). A deeper understanding of this enzyme group

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enables us to see their roles outside methanotrophy and ammonia oxidation (Coleman et al.,

2012; Hainbuch, 2015; Nielsen et al., 1997). Genes encoding all of the CuMMO family enzymes are known to have evolved from a single ancestral operon capable of oxidising either ammonia or methane (Stein et al., 2001). Here we generically refer to the related operons as xmoCAB.

Our initial interest in this possibly novel CuMMO operon detected in the WIP-OSPW began when we had its metagenome sequencing completed. A pmoA/amoA based gene tree generated from this metagenome showed two sequences grouping away from known pmoAs, suggesting a phylogenetically novel type of the CuMMO family, in which the term xmoCAB is used in this Chapter to refer to the operon (Figure 5-1).

Levels of xmoCAB were monitored in OSPW samples enriched with aromatic hydrocarbons. The level of xmoCAB gene decreased over time in all aromatic-hydrocarbons amended samples, after 4 weeks of incubation in benzene, toluene, xylene, and naphthalene enrichments (Figure 5-2). Reduced xmoCAB levels indicate that the gene doesn't take part in the degradation of the above hydrocarbon substrates. xmoCAB levels in MLSB-OSPW control sample (no added substrate) also decreased over the incubation time, which was probably affected by substrate loss. The aromatics test shows that organisms possessing the xmoCAB gene of interest are probably specialized to certain alkanes, and they do not grow on other substrates like aromatics. This experiment is also a useful control, since if the organisms with this gene are able to grow on everything, other experiments related to the gene would not be very informative.

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99 Methylacidiphilum fumariolicum SolV 100 Methylacidiphilum kamchatkense Kam1 100 Methylacidiphilum infernorum V4 Verrucomicrobia 61 Methylacidimicrobium tartarophylax 4AC LP2A isolate ARB 8C01A18 Methylosinus sp. LW4 98 96 Methylocystis echinoides IMET 10491 100 Methylocystis sp.SB12 81 Methylocystis sp.SC2 91 Methylocystis heyerii SAKB1 Methanocapsa acidophila B2 Alphaproteobacteria 100 Methylosinus sporium ATCC 35069 98 Methylosinus sporium SK13 95 Methylosinus trichosporium KS24b 100 Methylocystis sp. B2/7 Methylocystis parvus 81 62 100 Nitrosococcus oceani ATCC 19707 100 Nitrosococcus watsoni C-113 Nitrosococcus halophilus Nc4 99 87 Methylococcaceae bacterium AB302947 Methylomonas denitrificans FJG1 94 100 Methylobacter sp. BB5.1 Gammaproteobacteria 97 Methylobacter psychrophilus Z-0021 94 Methylovulum miyakonense HT12 Methylomicrobium alcaliphilum 20Z 78 92 Methylobacter sp. LW14 100 100 Haliea sp. ETY-NAG 90 Haliea sp. ETY-M xmoCAB 61 Hydrogenophaga sp. T4 Betaproteobacteria 100 Methylobacter marinus A45 99 79 Methylomicrobium album BG8 100 Methylocystis rosea SV97T Gammaproteobacteria 100 Methylomicrobium album BG8 Methylobacter luteus IMV-B-3098T Nitrosomonas eutropha C91 100 Nitrosomonas sp. ENI-11 89 Nitrosomonas europaea ATCC 19718 Nitrosomonas europaea ATCC 19718 99 Nitrosomonas sp. TK794 Betaproteobacteria 92 Nitrosovibrio tenuis NV-12 86 Nitrosospira multiformis ATCC 25196 Nitrosomonas sp. Is79A3 100 Crenothrix polyspora Crenothrix 100 Mycobacterium chubuense NBB4 Actinobacteria Mycobacterium rhodesiae NBB3 0.2 Figure 5-1. Phylogenetic tree of xmoA genes that includes sequences detected in WIP-

OSPW metagenome sampled in 2011, and other bacteria known to possess

pmoA/amoA/xmoA. The tree was constructed via Maximum Likelihood algorithm in

Seaview (Gouy et al., 2010), using 1,000 bootstraps as measure of support and the scale bar represents 0.2 change per nucleotide position. The xmoA gene of interest is depicted in red.

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Figure 5-2 Number of xmoCAB gene copies in MLSB-OSPW (per ml) sampled in

September 2014, with and without the addition of hydrocarbon substrates: DNA from

fresh MLSB (MLSB), MLSB amended with benzene (Benzene), MLSB amended with

toluene (Toluene), MLSB amended with xylene (Xylene), and MLSB amended with naphthalene (Naphthalene). The x-axis indicates the different samples and incubation time.

The y-axis indicates the number of xmoCAB gene copies detected per ml of sample. Bars

indicate ±SEM of true replicate samples.

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To study how organisms possessing the gene respond to the addition of short chain alkanes, we conducted an experiment using C1 (methane), C2 (ethane), and C3 (propane) as substrates. In the March 2015 MLSB control sample (with no added substrates), xmoCAB gene copy levels started very low, and remained approximately at the same levels over time.

Incubation of MLSB-OSPW sample with added ethane and propane showed an increase of xmoCAB gene copy levels after 6 incubation weeks, but a decrease was observed with added methane. MLSB amended with ethane was 1.7 times enriched, and MLSB amended with propane was 27.8 times enriched (Figure 5-3). The results show that xmoCAB operon has a possible role in oxidizing propane (Figure 5-3). The decrease of xmoCAB in the methane incubation confirms our hypothesis that this operon is dissimilar to pmoA gene responsible for methane oxidation in the tailings ponds. This brought up the questions: What organism can be linked to this gene? Does this gene take direct part in the oxidation of alkanes? Or does it only involve in processes not directly implicated to alkane oxidation?

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Figure 5-3 Number of xmoCAB gene copies in MLSB-OSPW (per ml) sampled in March

2015, with and without the addition of short chain alkanes as substrates: DNA from fresh

MLSB (MLSB), MLSB incubated for 6 weeks (MLSB), MLSB amended with methane

(Methane), MLSB amended with ethane (Ethane), and MLSB amended with propane

(Propane). The x-axis indicates the different samples and incubation time. The y-axis

indicates the number of xmoCAB gene copies detected per ml of sample. Bars indicate

±SEM.

To answer the first question of the organism that can be linked to this gene, a SIP experiment was conducted to: 1) see in which SIP fraction(s) the xmoCAB gene enriches, and 2) what possible organisms can be associated to the degradation of methane, ethane, and propane, by conducting a 16S rRNA gene sequencing of the heavy SIP fractions. MLSB-OSPW was

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amended with 12C- and 13C-methane, ethane, and propane, and was incubated for 10 days.

Results showed decreased levels of all three alkanes in 10 days, showing active oxidation of these alkanes by MLSB-OSPW (Supplementary Figures C3-C5). Methane oxidation by tailings water was expected and has been reported before, reaching an oxidation rate of up to 152 nmol

−1 −1 CH4 ml water d (Saidi-Mehrabad et al., 2013), but ethane and propane oxidation rates have never been reported before. In this study, it is discovered that OSPW sampled on August 2015 is

−1 −1 capable of methane oxidation with a rate up to 108.2 μmol of CH4 L OSPW d , almost

−1 −1 equivalent to the previous study that showed a rate of 152 μmol of CH4 L water d . Ethane

−1 −1 oxidation rate was 83.2 μmol of C2H6 L OSPW d , and propane oxidation rate was 58.6 μmol

−1 −1 of C3H8 L OSPW d (Supplementary Figures C3-C5).

Stable isotope probing (SIP) of MLSB-OSPW with and without addition of substrates was done after 10 days of incubation. Density gradient separation of extracted DNA indicated that 13C incubations also showed a shift in the DNA distribution towards heavier values (DNA density of 1.73 – 1.74 g ml-1, Supplementary Figure C6). This confirmed the incorporation of 13C into microbial DNA. The "heavy" DNA fractions representing the active 13C-assimilating communities were determined by comparing density gradients of the DNA extracted from the

SIP incubations to that of the control samples. The fractions chosen for community analyses are indicated in Figure C6.

Altogether, there were 65,002 reads from MLSB-OSPW 16S rRNA gene sequencing,

10,099 reads from MLSB-OSPW heavy SIP fractions, 17,180 reads from 13C-methane heavy SIP fractions, 29,718 reads from 13C-ethane heavy SIP fractions, and 9,080 reads from 13C-methane heavy SIP fractions. Gammaproteobacteria predominated MLSB-OSPW, its heavy SIP fraction, and the heavy SIP fraction of 13C-methane incubated sample: at 23.3%, 42.0%, and 57.2%

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relative abundance respectively (Figure 5-4). Betaproteobacteia predominated in 13C-ethane

(53.5%) and propane (78.1%) heavy SIP fractions (Figure 5-4). Both MLSB-OSPW and its heavy SIP fraction are diverse communities with the highest species predominance of only 9.6% and 9.8% coverage, as opposed to the other three enrichment samples (methane, ethane, and propane) which have around 33.5-55.0% predominance of one species (Supplementary Tables

C4 A-E).

An OTU closely related to Nitrosomonas oligotropha (98% similarity) dominates MLSB-

OSPW sampled in 2015, an organism known to have high affinity for ammonium (Figure 5-5)

(Ma et al., 2015; Saidi-Mehrabad et al., 2013). Ammonium concentration in OSPW can reach up to 17 mg l−1 (Saidi-Mehrabad et al., 2013), which likely represents the level of ammonium in the

MLSB-OSPW at the time it was sampled. The genus Nitrosomonas is known as an ammonia oxidizing group of bacteria adapted to low ammonium concentration (Bollmann et al., 2002).

This is probably why this genus appeared in both MLSB-OSPW and its SIP heavy fraction.

Dominant in the heavy fraction of MLSB-OSPW are organisms presumably contain high G+C levels: Porticoccus sp. (9.8% abundance), a hydrocarbon-degrading bacterium (Gutierrez et al.,

2012); Arenimonas sp. (8.3% abundance), a freshwater bacterium (Yuan et al., 2014), and

Nitrosomonas sp. (7.9% abundance), an ammonia oxidizing bacterium (Figure 5-5;

Supplementary Table C4) (Bollmann et al., 2002).

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Figure 5-4 Community structures (based on class) of oilsands process-affected water from the MLSB tailings pond (MLSB) and from the heavy fractions of MLSB-OSPW (Control),

13C-labelled DNA fractions of samples amended with 13C-methane (Methane), 13C-ethane

(Ethane), and 13C-propane (Propane) incubated under aerobic conditions. Predominant

taxa are identified to the class level based on the SILVA database. The OSPW-heavy treatment is the community detected only in the heaviest DNA fraction of OSPW. OSPW-

control shows the entire unseparated DNA from OSPW.

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110 Figure 5-5 Bubble chart of all 13C-labelled DNA from SIP heavy fraction samples analysed

in this study based on relative abundances (>1%). Community structures (based on genus)

of oilsands process-affected water from the MLSB tailings pond (MLSB-OSPW) and from

the heavy fraction of MLSB-OSPW (MLSB-heavy), 13C-labelled DNA fractions of samples

amended with 13C-methane (Methane), 13C-ethane (Ethane), and 13C-propane (Propane) incubated under aerobic conditions. Predominant taxa are identified to the class level based

on the SILVA database. The OSPW-heavy treatment is the community detected only in the

heaviest DNA fraction of OSPW. OSPW-control shows the entire unseparated DNA from

OSPW.

The strain most responsible for methane oxidation in MLSB-OSPW is closely related to

Methyloparacoccus murrellii (97% similarity), in congruence to Saidi-Mehrabad et al. (2013). It is predominant in the 13C-methane SIP heavy fraction, with 56.2% coverage (Figure 5-5;

Supplementary Table C4). An OTU closely related to Methyloversatilis thermotolerans (99% similarity) is responsible to both ethane and propane degradation in MLSB-OSPW at around

40.3% and 33.5% abundance in the heavy DNA, respectively (Figure 5-5; Supplementary Table

C4). Surprisingly, this Methyloversatilis genus is also predominant in both methanol and benzene

SIP heavy fractions (Chapter 3). This organism, which is a methylotroph (Doronina et al., 2014), is therefore capable of short chain alkanes (C2, C3, possibly C4–C5) and benzene (C6) degradation in MLSB-OSPW. Methyloversatilis thermotolerans is known capable of utilizing methanol, methylamine and a variety of multicarbon compounds as carbon and energy sources (Doronina et al., 2014; Kalyuzhnaya et al., 2006).

At the time of writing, the authors hypothesized the possibility of Hydrogenophaga sp. being the organism that possessed the novel xmoCAB gene of interest (Figure 5-1), because of its 111

presence in the heavy propane SIP fraction and fresh OSPW sample (7.4% predominance, Table

C4-D). The genome of Hydrogenophaga can be accessed on JGI but it is not published.

Therefore, nothing is as yet known about the strain. Accordingly, no definite conclusion can be made in regards of the organism related to xmoCAB gene, as a pure isolate of this organism was not cultured in this study. Nevertheless, the target organism did appear in either the ethane- or propane-SIP heavy fraction community.

To see if the xmoCAB gene of interest is found in the heavy SIP fractions of methane, ethane, and propane, combined SIP and qPCR methods was employed for MLSB-OSPW, as well as the 12C- and 13C-alkane substrates. The purpose of this is to detect the presence of xmoCAB in the heavy DNA fraction. SIP of MLSB-OSPW showed the presence of xmoCAB gene in 11 out of

12 SIP fractions. The highest number of xmoCAB copies was detected in the light fraction (DNA density 1.69 g ml-1) at 8,834.4 ± 2,589.6 number of gene copies ml-1 (Figure 5-6). This shows that in natural OSPW environment, this gene doesn't belong to organisms having high G+C contents and it is therefore detected in the lighter fractions (DNA density 1.69 – 1.7 g ml-1). Similar trend is seen in 12C- and 13C- methane incubations, in which the highest number of xmoCAB copies was detected in the light DNA fraction (DNA density 1.70 and 1.68 g ml-1) at 676,365 ± 188,685 and

10,905 ± 165 number of gene copies ml-1 (Figure 5-6). Based on the results, it can be inferred that xmoCAB does not respond to methane addition, and instead, pmoA genes can be linked to methane oxidation in OSPW (Saidi-Mehrabad et al., 2013).

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SIP of 12C- and 13C-ethane and propane incubations show distinct results. At the 12C- substrate incubation, highest number of xmoCAB copies was detected in the light fractions: at

DNA density of 1.70 g ml-1 for ethane and 1.69 g ml-1 for propane (Figure 5-6). xmoCAB copy number was 531.9 times enriched in 12C-propane compared to 12C-ethane SIP fractions. From this, it can be hypothesized that xmoCAB gene of interest tends to have an affinity to ethane, and especially propane, and that it is most likely involved in these alkanes oxidation. 114

Highest number of xmoCAB copies was detected in their 13C- ethane and propane heavy fractions: at DNA density of 1.71 – 1.72 g ml-1 for ethane and 1.73 g ml-1 for propane (Figure 5-

6). xmoCAB copy number was 58.4 times enriched in 13C-propane compared to 13C-ethane heavy

SIP fractions, indicating that the organism(s) assimilated both ethane and propane, but grew more on propane. Propane can be oxidized in 3 different pathways: 1) by monooxygenase-mediated terminal oxidation via 1-propanol or 2) subterminal oxidation via 2-propanol producing methyl acetate, or 3) hydroxyacetone (Kotani et al., 2003; Stephens & Dalton, 1986). It is interesting to note that xmoCAB copy number was the highest in the heavy fraction, which harbours heavy 13C-

DNA from organisms that assimilated the added substrates, i.e. xmoCAB was found in organisms responsible for ethane and propane oxidation (Figure 5-6). Although this is not proof, it does suggest that the enzyme encoded by this operon may be involved directly in ethane and propane oxidation.

It was previously reported that some methanotrophs such as Methylococcaceae (Hou et al., 1980), can be linked to intermediate ethane and propane oxidation. pMMOs in these organisms are possibly capable of co-oxidation functions, enabling them to oxidize a range of intermediate alkanes and alkenes with up to five carbons (Redmond et al., 2010; Cuny et al.,

2011). This shows that the CuMMO enzymes can act on other hydrocarbons, and perhaps can be evolved to specialize on these compounds. The study of gaseous alkanes oxidation has received little attention compared to the microbial oxidation of methane. Nevertheless, a number of microorganisms possessing CuMMO enzymes with the abilities to oxidise gaseous n-alkanes or alkenes as energy source have been discovered. Some Mycobacterium strains have been shown to oxidize C2–C4 alkanes with a CuMMO (Coleman et al., 2012), and two Haliea strains ETY-M and ETY-NAG are known to grow on ethylene using a similar enzyme (Suzuki et al., 2012). Our results show that 13C-ethane and propane were consumed by the genus Methyloversatilis, a 115

betaproteobacterial species not previously linked to either ethane or propane oxidation (Redmond et al., 2010).

Although its functions remain to be determined, the discovery of the divergent xmoCAB operon in WIP-OSPW expands our understanding of the evolutionary relationships among members of the CuMMO superfamily. It is possible that the main substrate of xmoCAB is a compound other than methane or ammonia, but the complexity of the CuMMO enzyme family and their substrate ranges make it difficult to verify this without having its pure isolate.

5.4 Conclusions

A highly divergent pmoA-like gene detected in the metagenome data of the oilsands process-affected water (OSPW) sample from West-In-Pit tailings pond may encode a novel alkane monooxygenase. This Cu-MMO enzyme group is known to catalyze the oxidation of methane, ammonia and potentially C2–C4 alkanes and alkenes. In this study, it is discovered that

−1 MLSB-OSPW is capable of methane oxidation with a rate up to 108.2 μmol of CH4 L OSPW

−1 −1 −1 d , ethane oxidation with a rate up to 83.2 μmol of C2H6 L OSPW d , and propane oxidation

−1 −1 with a rate up 58.6 μmol of C3H8 L OSPW d . It is suspected that some organisms in the

OSPW possess divergent pmoA-like gene with alternate oxidation functions, enabling them to oxidize a range of short-chain alkanes such as ethane and propane A qPCR system designed to detect this novel CuMMO combined with SIP suggested that this gene is present in both propane and ethane degrading organisms. Future direction in this study is to look at the gene expression for this particular operon in C2–C4 alkane enriched environments.

116 Preface

Chapter 6 presents a thorough characterization of a novel bacterium from the phylum

Genomic information revealed the genes catE (catechol 2,3-dioxygenase) and praC/xylH (4- oxalocrotonate tautomerase) in the organism, in which both are involved in catechol degradation pathways.

This paper is formatted for the International Journal of Systematic and Evolutionary

Microbiology where it will be submitted. Phospholipid-derived fatty acids (PLFA) analysis was done by Dr. Jaap Sinninghe-Damsté. Dr. Joongjae Kim guided me on most of the characterization experiments. Quinone analysis was done by Dr. Irene C Rijpstra and Dr. Peter Schumann. Dr.

Peter F Dunfield isolated the bacterium, provided funding, and supervised me throughout the study.

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Chapter Six: Oleiharenicola alkalitolerans gen. nov., sp. nov., a new member of the Verrucomicrobia isolated from an oilsands tailings pond

Fauziah F. Rochman1, Joong-Jae Kim1, Irene C. Rijpstra2, Jaap S. Sinninghe-Damsté2, Peter

Schumann3, and Peter F Dunfield1*

1 Department of Biological Sciences, University of Calgary, Calgary, Canada; 2 Department of

Marine Organic Biogeochemistry, NIOZ Royal Netherlands Institute for Sea Research, P.O. Box

59, 1790 AB, Den Burg, Texel, The Netherlands; 3 Leibniz Institute DSMZ-German Collection of Microorganisms and Cell Cultures, Braunschweig, Germany.

The Genbank accession number for the 16S rRNA gene sequence of Oleiharenicola alkalitolerans NVTT is KJ721192. The Joint Genome Institute IMG/M submission ID number for the draft genome is 43214.

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6.1 Abstract

A novel member of the phylum Verrucomicrobia was isolated from an oilsands tailings pond in Alberta, Canada. Cells of isolate NVTT are Gram-negative, strictly aerobic, non- pigmented, non-motile cocci to diplococci 0.5-1.0 μm in diameter. The bacterium is neutrophilic

(optimum pH 6.0–7.0) but alkalitolerant, capable of growth between pH 5.5 and 11.0. The temperature range for growth is 15-40°C (optimum 25–37°C). Energy sources include sugars and organic acids. Nitrogen sources include nitrate, urea, L-glycine, L-alanine, L-proline, and L- serine. Does not fix atmospheric nitrogen, and nitrogenase-encoding genes are absent from a draft genome. Does not require NaCl and is inhibited at NaCl concentrations above 3.0% (w/v). The

DNA G+C content of strain NVTT, based on a draft genome sequence, is 66.13 mol%. MK-6 and

MK-7 are the major respiratory quinones. Major cellular fatty acids and polar lipids are isoC15:0 and anteisoC15:0. Phylogenetic analysis of 16S rRNA gene sequences revealed that the strain belongs to the family Opitutaceae of the phylum Verrucomicrobia. The most closely related validated species is Opitutus terrae PB90-1T (93.7% 16S rRNA gene sequence identity). Based on genotypic, phenotypic and chemotaxonomic characteristics, it was concluded that this strain represents a novel genus and species, for which the name Oleiharenicola alkalitolerans gen. nov., sp. nov. is proposed. The type strain is NVTT (=ATCC BAA-2697).

The Genbank accession number for the 16S rRNA gene sequence of Oleiharenicola alkalitolerans NVTT is KJ721192. The Joint Genome Institute IMG/M submission ID number for the draft genome is 43214.

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6.2 Introduction

The phylum Verrucomicrobia (Hedlund et al., 1997) contains a diverse range of microorganisms utilizing various carbon and energy sources. Cultivation-independent approaches have revealed a broad distribution of this phylum in various niches. Members of this phylum have been detected in open oceans and sediments (Freitas et al., 2012), geothermal fields

(Dunfield et al., 2007; Sharp et al., 2014b), hot springs (Lau et al., 2009), soils (Chin et al.,

2001), fresh waters (Lindström et al., 2004; Lindström et al., 2005), and termite guts ecosystems

(Hongoh et al., 2003), with wide ranges of temperature, nitrate and oxygen levels, depth, salinity, and nutrient availabilities. However, since only a handful of representatives have been successfully cultivated, little is understood about the phylum Verrucomicrobia.

At the time of writing, three classes of Verrucomicrobia have been formally defined:

Verrucomicrobiae, Spartobacteria, and Opitutae (Hedlund, 2010). The class Opitutae, whose members have mostly been cultivated from marine environments, is one of the most commonly detected lineages of the phylum (Hedlund, 2010; Hugenholtz et al., 1998). Molecular studies have detected the class Opitutae in diverse habitats including Arctic sea ice and seawater, shrimp and tilapia culture ponds, and surface soils in Antarctica, Europe, and the Americas (Bergmann et al., 2011; Bowman et al., 2012; Fan et al., 2016; Wu et al., 2014). Aside from the not validated

“Diplosphaera colitermitum TAV2” which was isolated from a termite gut (Wertz et al., 2012;

Wu et al., 2014), no novel members from the family Opitutaceae, Order Opitutales, Class

Opitutae, have been cultivated and validly reported in the last decade. Altogether, there are only

10 validated species in the class Opitutae.

The two recognized members of the family Opitutaceae are Alterococcus agarolyticus

ADT3T, which is the type strain for the genus Alterococcus (Shieh & Jean, 1998), and

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Opitutus terrae PB90-1T, which is the type strain for the genus Opitutus (Chin et al., 2001), as well as the order Opitutales and the family Opitutaceae (Choo et al. 2007).

Alterococcus agarolyticus ADT3T is a halophilic thermophilic bacterium isolated from hot springs in Taiwan (Shieh & Jean, 1998). Five strains belonging to the genus Alterococcus isolated in the study were all able to produce extracellular agarase on agar medium, and were capable of both aerobic growth and anaerobic growth by fermenting glucose to various organic acids (Shieh & Jean, 1998). Opitutus terrae PB90-1T is an obligately anaerobic bacterium commonly found in rice paddy soils (Chin et al., 2001; van Passel et al., 2011). The strain is a chemoheterotroph that grows on mono-, di- and polysaccharides, with propionate and acetate as the major fermentation products (Chin et al., 2001).

In this study, a novel mesophilic Verrucomicrobia belonging to the family Opitutaceae was isolated from an oilsands tailings pond located near Ft. McMurray, Alberta, Canada. The tailings water sample was collected in June 2012 from 0-10 cm below the surface and about 15 m from the shore. Tailings ponds contain process-affected waters and fluid fine tailings from the industrial oilsands extraction processes (Quagraine et al., 2005; Saidi-Mehrabad et al., 2013).

The sample used in this study was alkaline (pH 8), 20°C, contained 2770 ± 12.4 mg L-1 of total major ions, 9.9 ± 0.1 mg N L-1 of total nitrogen, and 40 ± 1.0 mg C L-1 of dissolved organic carbon (Saidi-Mehrabad et al., 2013; Syncrude Canada, 2010). In 2012, a sequencing analysis of

16S rRNA amplicons from the water sample found that an OTU from the phylum

Verrucomicrobia, class Opitutae, order Opitutales, and family Opitutaceae was quite abundant, accounting for 0.66% of all reads in the tailings surface water (24 out of 3636) (Saidi-Mehrabad et al., 2013).

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6.3 Materials and Methods

6.3.1 Bacterial isolation

To isolate the bacterium, sample water was diluted 1:10 with milliQ water and 100-μl aliquots were spread on plates of the mineral salts medium 10. Plates were incubated at 25-35°C under air. Medium 10 was prepared as described previously (Heyer et al., 2002), except that agar was substituted with 1.5% phytagel (Sigma Aldrich, St Louis, MO, USA), and milliQ water was substituted with filtered sample water to mimic the native environment and provide potential energy sources. The sample water was filtered through a 0.2-μm filter (Pall Life Sciences, East

Hills, NY, USA). We named this modified medium as M10-T. Colonies growing on the M10-T were picked and streaked to new M10-T plates. This was repeated several times until purified strain NVTT was obtained.

6.3.2 Cell growing

Strain NVTT was initially found to be growing well on Medium 10 plates together with an

Ancylobacter sp. (identified via colony PCR of 16S rRNA genes as having 100% sequence similarity to Ancylobacter polymorphus DSM 2457T). Members of the genus Ancylobacter are abundant in nature, especially in aquatic environments (Xin et al., 2006). Some strains are methylotrophs (Van den Wijngaard et al., 1992), and they have been found in various water reservoirs, such as lowland marshes, shallow-waters and aerobic wastewater treatment ponds

(Doronina et al., 2001; Fulthorpe et al., 1993; Raj, 1989; Zaichikova et al., 2010). When strain

NVTT was obtained into pure culture, it grew much more slowly than in mixed culture. It was hypothesized that the companion bacterium provided growth factors not present in the mineral medium. Therefore growth was tested in media supplemented with cell extract of the companion

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bacterium, as well as on several complex media. The optimal complex medium was a diluted

20% strength R2A medium, a medium recommended for growing heterotrophic bacteria

(Reasoner & Geldreich, 1985) adjusted to pH 8 with NaOH, and supplemented with an

Ancylobacter sp. cell-free extract prepared based on a previously described method (Kim et al.,

2008). To prepare the extract Ancylobacter sp. culture was harvested in its exponential growth phase. The culture was centrifuged (Beckman Coulter, Miami, FL, USA) to separate the cell pellet (C1) and supernatant (S1). C1 was sonicated, suspended in 500 ml of 20% strength R2A medium, and centrifuged (Beckman Coulter) at 4°C to separate C1 from its second supernatant

(S2). Supernatant (S1) was filtered with a 0.2-μm filter (Pall Life Sciences). S1, S2, and 1.5% phytagel (Sigma Aldrich) were combined to make plates of R2A-Ancylo medium and C1 was discarded.

This method increased the cell growth of strain NVTT in terms of masses of cells and growth rate, so that visible growth occurred in 2-3 d instead of 5-7 d of incubation in liquid medium. However, after several transfers, cells grown on the Ancylobacter sp. extract medium could then be grown subsequently on 20% R2A Medium with the similar enhanced growth rate.

Therefore, subsequently strain NVTT was cultured routinely at 25°C under aerobic conditions on

20% strength R2A (pH 8.5) plates without the laborious addition of cell extract. Except where noted, this medium was used for all subsequent physiological tests.

6.3.3 Physiological tests

Gram-staining was performed according to the protocol provided by the Gram Stain Kit

No. 212539 (Becton Dickinson, Franklin Lakes, NJ, USA). Cell morphology was observed via light microscopy (BX51; Olympus Life Science Solutions, Waltham, MA, USA) using cells from

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exponentially growing culture in 20% R2A broth (pH 8.5, 25°C). Bacterial flagella staining was performed as described previously (Moyes et al., 2009). A motility test was done as described previously (Shields & Cathcart, 2011) except that R2A was used as a medium and agar was substituted with 0.4% phytagel. Escherichia coli K-12 strain W3110T was used as a positive control.

To determine the temperature (°C) range for growth, isolate NVTT was grown at 10-50°C

(at intervals of 5°C) and was observed for 2 weeks in 20% R2A broth (pH 8.5). Growth at pH

4.0-12.0 (at intervals of 0.5 pH units) was evaluated for 7 d in 20% R2A broth adjusted with 10%

NaOH (Sigma Aldrich) and 10 M H2SO4 (Sigma Aldrich). pH was measured with an AB15 pH

Meter (Fisher Scientific, Waltham, MA, USA) before and after medium autoclaving with no observed pH change. Growth in the presence of 0-4% (w/v) NaCl (at intervals of 0.5% NaCl) was investigated for 2 weeks. Anaerobic growth was tested under N2 and N2/CO2 (80 : 20, v/v) atmospheres, using resazurin sodium salt (1 mg per litre; Sigma Aldrich) as an indicator. Growth was observed for 2 weeks. All tests were performed in duplicate 100-ml serum bottles (VWR,

Radnor, PA, USA) capped with butyl rubber stoppers (VWR) and filled with 20 ml of the broth.

T Growth of strain NVT was determined by monitoring the optical density at 600 nm (OD600) using an Ultrospec 10 Cell Density Meter (Amersham Biosciences, Little Chalfont, UK) daily.

The ability of NVTT cells to grow aerobically on a variety of nitrogen sources was investigated by growing them in a nitrogen-free medium (Medium 10 without NH4Cl) with 0.1%

Glucose (Sigma Aldrich) as a carbon source. Cells were grown in 20 ml of nitrogen-free Medium

10 in 100-ml serum bottles. 10 mM of sodium nitrite (Sigma Aldrich), sodium nitrate (Sigma

Aldrich), urea (Sigma Aldrich), aspartic acid (Sigma Aldrich), L-valine (Sigma Aldrich), L- glycine (Sigma Aldrich), L-alanine (Sigma Aldrich), L-proline (Sigma Aldrich), L-serine (Sigma

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Aldrich), or 10% of N2 gas were added as nitrogen sources. Nitrite and nitrate reduction were tested by growing the cells anaerobically in nitrogen-free Medium R2A amended with 10 mM of sodium nitrite and 10 mM of sodium nitrate. Tests were done using 100-ml serum bottles.

Catalase and oxidase activities were tested using 3% (v/v) H2O2 and an oxidase reagent dropper (Becton Dickinson, Franklin Lakes, NJ, USA), respectively. Other enzymatic tests, and the substrate range for growth were determined using API 20NE, API 50CH and API ZYM kits

(bioMérieux, Marcy-l'Étoile, France) according to the manufacturer's instructions. Growth on catechol was tested by growing strain NVTT in 0.25% (w/v) of dissolved catechol (99%, Sigma

Aldrich) in Medium 10. For cellular fatty acids and respiratory quinones analyses cells were harvested during the late exponential growth phase (3 d). Cell culture was centrifuged and the pellet was freeze-dried overnight. Isoprenoid quinones were extracted from lyophilized cells according to the method of Collins et al. (1977) and the profile was analysed by HPLC

(Shimadzu LC 20A; Shimadzu Corporation, Kyoto, Japan) (Groth et al., 1997).

Cellular fatty acids analysis was carried out as described previously (Klatte et al., 1994), and the fatty acid methyl esters were then separated using the Microbial Identification System (MIDI,

Inc., Sherlock version 6.1) (Sasser, 1990).

A draft genome of strain NVTT was generated to support genomic characterization. The

G+C content, the 16S rRNA sequence, and information about the enzyme pathways present were determined from the draft genome. Genomic DNA was extracted using the FastDNA SPIN kit

(MP Biomedical, Santa Ana, CA, USA). A paired end library was sequenced in 1/10 of a lane of a HiSeq2000 Illumina platform at McGill University and Genome Québec Innovation Center following the center guidelines (http://gqinnovationcenter.com/index.aspx). Raw reads were passed through an in-house quality control program as described previously (Saidi-Mehrabad et

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al., 2013). The draft genome was assembled via Velvet Software Version (Zerbino & Birney,

2008). A total of thirteen assemblies using different k-mer size values (51, 55, 59, 63, 67, 71, 75,

79, 83, 87, 91, 95, and 99) were performed; the 95-k-mer assembly was selected as the highest quality for the further characterization. The total assembly contained 5.72 Mbp in 322 contigs

(max length: 225,843 bp, min length: 201 bp, median length: 1381 bp, mean length: 17,772 bp,

N50: 81,720 bp). Annotation was performed by submission to the IMG platform of the Joint

Genome Institute (JGI) website (http://www.jgi.doe.gov/) (Markowitz et al., 2006; Mavromatis et al., 2009).

A Neighbour-Joining phylogenetic tree was generated to determine the phylogenetic position of strain NVTT compared to other closely related Verrucomicrobia species. All other

16S rRNA gene sequences of reference strains were obtained from the ARB SSURef119 SILVA

Database updated on July 14 2014 (Ludwig et al., 2004b). NVTT was aligned to the database using the SINA aligner (Pruesse et al., 2012).

6.4 Results and discussion

6.4.1 Morphology and physiology

A novel mesophilic Verrucomicrobia belonging to the family Opitutaceae was isolated from an oilsands tailings pond located near Ft. McMurray, Alberta, Canada. The organism apparently belongs to a new genus with a proposed name: Oleiharenicola, under the family

Opitutaceae, order Verrucomicrobiales, class Verrucomicrobiae, and division Verrucomicrobia.

The type strain of the genus is Oleiharenicola alkalitolerans sp. nov. strain NVTT.

Colonies grown on R2A plates after 2-3 d were pale white, smooth, moist, raised, opaque, and circular with smooth margins, measuring 0.5-4 mm in diameter. Cells of strain NVTT were

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Gram-stain negative, coccoid in shape, and ranged from 0.5 to 1.0 μm in diameter. Flagella were not observed. Cells were not motile after 5 d of incubation at 37°C. Cells were able to grow at

15–40°C (optimum 25-37°C). No growth was observed at 10°C or 45°C. Growth was observed from pH 5.5–11 (optimum pH 6.0–7.0). No growth was observed at pH 5.0 or pH 11.5. Cells could tolerate up to 3% of NaCl (w/v) but did not require supplemental NaCl to grow. The doubling time on 20% R2A liquid medium at pH 7 was 0.6 h based on OD600 measurements of its growth in 25°C. No growth was observed under anaerobic conditions. No growth was observed

T under a 100% N2 atmosphere with 10 mM added sodium nitrite, indicating that strain NVT was unable to grow by anaerobic respiration with nitrite.

Growth was positive for sodium nitrate, urea, L-glycine, L-alanine, L-proline, and L- serine, but no growth was seen for L-valine and N2 gas after 6 days of incubation. Strain

NVTT was unable to grow by anaerobic respiration with nitrite or nitrate (10 mM). The catalase test was positive while the oxidase test was negative. The results of the substrate test with the

50CH kit are listed in the species description. The predominant respiratory quinones detected in strain NVT were menaquinone MK-7 and MK-6 (80:12). This result was in congruence to the characteristics found in the type strains of the order Puniceicoccales – one of the closest relatives to the order Opitutales – whereby MK-7 was also found to be the major respiratory quinone

(Chung & King, 2001; Yoon et al., 2007). There is no quinone data available for the closest relative of strain NVTT, i.e. Opitutus terrae PB90-1T.

For Transmission Electron Microscopy (TEM), 3-d old cells from R2A medium were centrifuged and preparation was done based on a previously described general protocol (Bozzola

& Russell, 1999). Cells were washed with 0.01 N NaOH (Sigma-Aldrich) and fixed with 3% glutaraldehyde (Alfa Aesar) in cacodylate buffer (Electron Microscopy Sciences) for 2 h at 25°C.

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Washing was done with buffer 5 times every 10 m at 25°C. Cells were fixed with 1% OsO4

(Sigma-Aldrich) in buffer for 2 h at 4°C and washed. Staining was done with 2% uranyl acetate

(Electron Microscopy Sciences) for 20 m in the dark. In the dehydration step, 30, 50, 75, 95, and

100% alcohol was used to wash the cells 3 times in every 15 m at 25°C. Two changes of 100% propylene oxide solvent (Sigma-Aldrich) every 20 m were done as a transition. In the resin infiltration step, cells were incubated with 2:1, 1:1, 1:2, and 0:1 mixtures of propylene oxide solvent and epoxy resin (Sigma-Aldrich) for 1 h each and overnight for the last mixture. Before embedding, cells were incubated in fresh resin for 1 h. Resin was prepared and polymerized according to the protocol provided by the Epoxy Embedding Medium Kit No. 45359 (Fluka

Analytical). Embedded cells sample was sectioned with thickness of 50-70 nm. Cells morphology was imaged under Tecnai F20 Transmission Electron Microscope (FEI) at the Microscopy and

Imaging Facility at the University of Calgary.

A thorough examination of the cells morphology under TEM (Figure 6-1) shows distinct rounded structures close to eukaryotic nuclear envelopes within cells’ inner membrane in addition to aggregations of granular particles that are evenly distributed within the cells. It is suggested that the rounded structure resembles a vesicle membrane formed inside the cells, which is uniquely found in the Planctomycetes, Verrucomicrobia, and Chlamydiae superphylum

(Santarella-Mellwig et al., 2010). The previous study also argued that such protein architectures play important roles in eukaryogenesis, a hypothesis that the cell nucleus of eukaryotic life forms evolved from the endosymbiosis of a large DNA virus (Santarella-Mellwig et al., 2010).

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Figure 6-1. (A) Gram staining picture of strain NVTT cells under bright field microscopy

showing cocci and diplococci cell morphologies. (B) Transmission electron micrograph

T (TEM) of strain NVT cells fixed in OsO4 solution. Uranyl acetate was used to increase image contrast. (C) Inset: Note the presence of granular vesicle inside the cell indicated by

an arrow (lower right inset; bar, 100 nm).

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Table 6-1. Differential phenotypic characteristics of Oleiharenicola alkalitolerans NVTT and its closely related species.

Characteristic 1 2 3 4 Isolation sources industrial rice paddy termite gut hot springs

16S rRNA similarity processNA - soil93.78 93.05 88.65 Aerobic growth + - + + Anaerobic(%) growth affected- water + - + Motility - + -(microaerophile) + Colony colour colorless unpigmented colorless white Growth temperature 15-40 (25-37) 10-37 15–35 (30) 38-58 (48) Growth pH 5.5-11 (6-8) 5.5-9 (7.5-8) 5.5–7.5 (7) 7-8.5 NaCl(°C) tolerance (%, 3 3 1.5 1-3.5 (2) G+C contents (mol%) 66.13 74 60.5 65.8 Nitratew/v) reduction - + ND ND Catalase + - - + Oxidasecapacity - - - + Assimilation test Arabinose w + - - Mannose - + - w Mannitol + + - - Maltose - + + ND Sucrose - + - + Lactose - + ND + Xylose + - - + Melibiose - + ND - Fructose + + - ND Cellobiose - + + + Quinone MK-6 (%) 80 ND ND ND MK-7 (%) 12 ND ND ND Taxa: 1, Oleiharenicola alkalitolerans NVTT; 2, Opitutus terrae PB90-1T (Chin et al., 2001); 3, Diplosphaera colitermitum strain TAV2 (Wertz et al., 2012); 4, Alterococcus agarolyticus ADT3T (Shieh & Jean, 1998). Abbreviations: +, Positive; W, weakly positive; −, negative; ND, no data available; NA, not applicable. Data for strain NVTT was obtained from present research and the rest was gathered from previously published sources referred above. For growth temperature, pH, and NaCl levels, values in brackets are the known optimum conditions. All four strains are gram negative, with cocci and some diplococci cells morphology ranging from 0.5 to 1 um in length.

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Aerobic respiration is a characteristic of strain NVTT that distinguishes it from its closest neighbour Opitutus terrae PB90-1T. Opitutus terrae is an obligate anaerobe, capable of fermentation, and reducing nitrate to nitrite (Chin et al., 2001). Of the other related strains,

“Diplosphaera colitermitum strain TAV2” prefers microaerophilic conditions to grow (Wertz et al., 2012), whereas Alterococcus agarolyticus ADT3T is a facultative anaerobe (Table 6-1) (Choo et al., 2007). G+C contents for all the above species are approximately the same as strain NVTT

(60.5 and 66 mol% respectively), except for O. terrae PB90-1T, which has a higher G+C content of 74 mol% (Chin et al., 2001). O. terrae PB90-1T can grow to as low as 10°C, and A. agarolyticus ADT3T is capable of growth at 58°C, while strain NVTT grows between 15-40°C

(Table 6-2). No information is available on quinones or nitrogen sources for all of the above strains except for strain “D. colitermitum strain TAV2”, that does perform nitrogen fixation

(Wertz et al., 2012). Several other characteristics that differentiate strain NVTT from four other closest strains of phylum ‘Verrucomicrobia’ are provided in Table 6-2.

Based on the results of the phylogenetic, biochemical, and physiological analyses, strain

NVTT represents a novel genus within the family Opitutaceae. Strain NVTT is proposed as the type strain of the new genus and species, Oleiharenicola alkalitolerans gen. nov., sp. nov.

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Table 6-2. Cellular fatty acid compositions of strain NVTT and its most closely related species. Values are the molar percentages of total cellular fatty acids.

Total fatty acids (Molar %) Fatty acids 1 2 3 4 5 Straight chain C14:0 1.2 11 3.8 42 - C16:0 8.5 15 8.9 - 7.9 C16:1 8.3 - - 17.7 - 2-OH-C16:0 - - 6.1 - - C17:0 - - 4.7 - 7 C18:0 - - 4.1 - 24.7 C18:1 - - 1.7 - - C18:2 - - 1.9 - - C19:0 - - 2.5 - - C20:0 - - - - -

Branched chain Iso-C13:0 6 - - - - Anteiso -C13:0 3 Iso-C 14:0 7.5 - - 5.7 - Iso-C15:0 20.3 21 2.1 - - Anteiso-C15:0 27.7 33 51.5 9.7 30.9 I so-C16:0 4.6 - 8.1 - - A nteiso-C17:0 0.5 - 1.1 - -

Total 87.6 80 96.5 75.1 70.5 Taxa: 1, Oleiharenicola alkalitolerans NVTT; 2, Diplosphaera colitermitum strain TAV2 (Wertz et al., 2012); 3, Alterococcus agarolyticus ADT3T (Shieh & Jean, 1998); 4, Prosthecobacter fluviatilis HAQ-1T (Takeda et al., 2008), 5, Puniceicoccus vermicola IMCC1545T (Choo et al., 2007). Opitutus terrae PB90-1T had no reported cellular fatty acid compositions (Chin et al., 2001). Values are molar percentages of total fatty acids; –, not detected/low detection.

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6.4.2 Genome analysis

The 1440 bp 16S rRNA gene sequence of strain NVTT obtained from the genome was phylogenetically compared to the most related sequences via BLASTN (Altschul et al., 1997) against a database containing type strains with validly published prokaryotic names and representatives of uncultured phylotypes (Kim et al., 2012). Strain NVTT was phylogenetically related to the members of the family Opitutaceae, with its closest relatives being Opitutus terrae

PB90-1T (93.7 %), "Diplosphaera colitermitum strain TAV2” (not validated; 93.1 %),

Alterococcus agarolyticus ADT3T (88.4 %), and less than 86 % similarity with other members of the phylum Verrucomicrobia.

A phylogenetic tree was constructed with a Jukes–Cantor correction and 1000 bootstrap replicates to ensure confidence (Figure 6-2). The phylogenetic tree confirmed that the closest relatives of strain NVTT are Opitutus terrae PB90-1T and "Diplosphaera colitermitum strain

TAV2” (Figure 6-2). An in silico DNA–DNA hybridization (DDH) experiment was also performed between the genome of NVTT and the genome of Opitutus terrae PB90-1T

(http://ggdc.dsmz.de) (Meier-Kolthoff et al., 2014). The DDH value was 19.10 ± 2.28%.

Some major pathways based on the KEGG categories of strain NVTT genome sequence were retrieved: amino acids metabolism (13%), carbohydrate metabolism (12%), energy metabolism (10%), metabolism of cofactors and vitamins (7%), and signal transduction (6%).

Around 3% of the pathways are for xenobiotics biodegradation and metabolism genes. Complete genes were retrieved for general prokaryotic metabolisms, such as glycolysis, TCA cycle, and carbon and fatty acids metabolism. For nitrogen metabolism, strain NVTT harbours nrfA (nitrite reductase), nirK (nitrite reductase), and a putative gene encoding nosZ (nitrous-oxide reductase).

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numbers of the 16S rRNA gene sequences of the representative strains in the tree are indicated in brackets. Numbers inside wedges shows the number of species in the closed groups. Rubritalea, Roseibacillus and Haloferula are genus names within the

order Verrucomicrobiales. Tree was constructed via SeaView and aligned via ARB. Bar, 0.05 substitutions per sit

134 KEGG pathway based on genomic annotation of this strain detected catE (catechol 2,3- dioxygenase) and praC, xylH (4-oxalocrotonate tautomerase), both are involved in catechol degradation pathways, and aldH NADP-dependent aldehyde dehydrogenase, from ring cleavage via Baeyer-Villiger oxidation pathway. These pathways relate to the aerobic degradation of aromatic compounds, possibly revealing its ecological role in the tailings. Although this strain is likely incapable of performing aromatic compounds degradation by itself, it may have been feeding on intermediate products from aromatic compounds degradation in the tailings basin. It can also be postulated that plain R2A medium might not provide conditions favourable to genes involved in aromatic compounds degradation.

6.4.3 Description of Oleiharenicola gen. nov.

Oleiharenicola (O.le.i.ha.re.ni'co.la. L. n. oleum, oil; L. n. harena, sand; L. suffix -cola, inhabitant, dweller; N.L. masc. n. Oleiharenicola an inhabitant of oilsands, referring to the oilsands process-affected water, from where the type strain was isolated).

Non-spore forming, non-motile, mesophilic and strictly aerobic chemoorganoheterotrophic bacteria. Gram-negative cocci or diplococci (0.5-1.5 μm) that occur mainly as single cells or pairs. No flagella are observed. Belongs to the family Opitutae, closest neighbor is the genera Opitutus. Menaquinone MK-7 and MK-6 are major respiratory quinones.

Major cellular fatty acids are isoC15:0 and anteisoC15:0. The type strain of the genus is

Oleiharenicola alkalitolerans sp. nov. strain NVTT.

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6.4.4 Description of Oleiharenicola alkalitolerans sp. nov.

Oleiharenicola alkalitolerans (al.ka.li.to'le.rans. N.L. n. alkali (from Arab. al qali) alkaline, L. part. pres. tolerans, tolerating. N.L. part, adj. alkalitolerans, referring to an alkali- tolerating micro-organism).

Colonies grown on 20% R2A plates are pale white, smooth, moist, raised, opaque, and circular with smooth margins, measuring around 0.5-4 mm in diameter. Cells are able to grow at

15–40°C (optimum 25-37°C) and pH 5.5–11 (optimum pH 6.0–7.0). Cells can tolerate up to 3 % of NaCl (w/v) but doesn't require NaCl to grow. Doubling time under optimum conditions is 0.6 h. Catalase positive and oxidase negative. Cells are negative for indole production, glucose fermentation, arginine dihydrolase, gelatin hydrolysis, and assimilation of mannose, n-acetylglucosamine, D-maltose, potassium gluconate, capric acid, adipic acid, trisodium citrate, and phenylacetic acid. Cells are positive for urease, β-glucosidase hydrolysis (esculin), protease hydrolysis (gelatin), β-galactosidase, and glucose, arabinose, mannitol, and malate assimilation.

The strain is able to metabolize the following carbon sources: of D-galactose, D-glucose,

D-fructose, D-mannitol, D-sorbitol, esculin ferric acetate, xylitol, D-lyxose, D-fucose, and D- arabitol. The cells fail to metabolize the following compounds: or D-mannose, L-sorbose, L- rhamnose, dulcitol, inositol, methyl-α-D-mannopyranoside, methyl-α-D-glucopyranoside, N- acetylglucosamine, amygdalin, arbutin, salicin, D-cellobiose, D-maltose, D-lactose, D-melibiose,

D-saccharose, D-trehalose, inulin, D-melezitose, D-raffinose, amidon, glycogen, gentiobiose, D- turanose, D-tagatose, L-fucose, L-arabitol, potassium gluconate, potassium 2-ketogluconate, and potassium 5-ketogluconate. For enzymatic activities, positive reactions are observed for alkaline phosphatase, esterase lipase (C8), trypsin, α-chymotrypsin, acid phosphatase, and naphtol-AS-BI- phosphohydrolase, and negative for esterase (C4), lipase (C14), leucine arylamidase, valine

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arylamidase, cystine arylamidase, α-galactosidase, β-galactosidase, β-glucuronidase, α- glucosidase, β-glucosidase, N-acetyl-β-glucosaminidase, α-mannosidase, and α-fucosidase. Strain

NVTT was able to use sodium nitrate, urea, L-glycine, L-alanine, L-proline, and L-serine, but not

T L-valine and N2 gas, as nitrogen sources. The type strain, NVT (=ATCC BAA-2697 =DSMZ), was isolated from an oilsands process-affected water in Alberta, Canada. The DNA G+C content of the type strain is 66.13 mol%.

6.5 Acknowledgements

This work was made possible through financial assistance from Genome Canada, Genome

Alberta, Genome BC and the Government of Alberta (GC Grant 1203). PD acknowledges the support of a visiting scholar grant from the German Academic Exchange Service (DAAD). We thank Bernhard Schink for nomenclature advice, Anika Wasner for technical assistance with quinone analysis, and Xiaoli Dong and Christoph Sensen for assistance with the genome assembly.

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Chapter Seven: General conclusions and future research

This dissertation applied 16S rRNA gene sequencing via both 454 pyrotag and Illumina

MiSeq sequencing along with stable isotope probing (SIP) approaches to investigate microbial communities responsible for hydrocarbon degradation in oilsands process-affected water (OSPW). Hydrocarbons investigated in this study were methane, ethane, propane, benzene, and naphthalene. Other substrates studied via stable isotope probing approach were methanol, sodium acetate, protein extract and CO2.

The objectives directed in this thesis were to: 1) identify indigenous microbial communities of WIP-OSPW pond surface responsible for benzene and naphthalene degradation, as well as other major aerobic activities, 2) use metagenomics to explore the diversity of hydrocarbon degrading genes in the WIP-OSPW, 3) cultivate strains indigenous to OSPW by using improved cultivation techniques, and test their hydrocarbon degradation potentials.

7.1 Research summary

In recent years, our understanding of microbial systems has expanded, in which many undiscovered microorganisms and pathways were discovered due to the advances of sequencing technologies. In this dissertation, Next-Generation Sequencing technology is mainly harnessed in conjunction with SIP to complete the classic microbial hydrocarbon degradation methodologies, such as sample enrichment and bacterial cultivation.

The study in Chapter 3 discusses the organisms and pathways that are responsible for aerobic hydrocarbon degradation of the surface OSPW layer. In the study, metagenomic sequencing of the WIP-OSPW sample was performed, in which a number of pathways were outlined and linked to microorganisms responsible for the processes. Nevertheless, we noted that

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metagenomics itself cannot stand alone, therefore cultivation and SIP studies were also undertaken. SIP of model aromatics (benzene and naphthalene) and 16S rRNA gene sequencing were performed. To strengthen the hypothesis, traditional cultivation of hydrocarbon degrading organisms from OSPW was also done. The coupling of metagenomics sequencing technology,

SIP, and traditional microbiological methods allowed us to identify organisms with important roles within the OSPW. Utilizing the metagenome alone was not very useful, since it predicted a huge number of potential hydrocarbon degraders (mostly Alphaproteobacteria) but in the actual

SIP heavy fractions of both 13C-benzene and 13C-naphthalene, only a few of these potential bacteria were actually shown to be active. The others either concentrate on other substrates or are active under other conditions.

Predominant bacteria found to be important in our benzene and naphthalene amended environments, such as Methyloversatilis and unknown Chromatiaceae (related to Thiococcus), were not predicted by other related studies, or that their previous roles in hydrocarbon degradation were of minor importance. Chances are that these organisms may have been overlooked in the past, or that there are certain environmental conditions in OSPW that caused them to prosper. The combination of both metagenome analyses and SIP enabled us to determine metabolic roles of some resident microbial communities.

In Chapter 4, the SIP technique was employed to study the indigenous aerobic bacterial communities of WIP-OSPW. Methanol is a byproduct of methane oxidation, and acetate is a major product of anaerobic degradation. To study these communities in the Alberta tailings ponds, labelled methanol and acetate were added as substrates to WIP-OSPW samples. Labelled protein extract was also used to study indigenous proteolytic bacterial communities. Although methanol, a byproduct of methane oxidation, is probably abundant in the tailings pond, methylotrophs are not particularly predominant in natural OSPW. Instead, detected organisms are 139

mostly simple sugars and organic acid feeders, such as Devosia riboflavina from the phylum

Alphaproteobacteria. Sequencing results based on DNA from untreated OSPW is predominated by Betaproteobacteria, OSPW heavy fraction by Alphaproteobacteria, methanol-SIP by

Betaproteobacteria, sodium acetate-SIP by Alphaproteobacteria, protein extract-SIP by

Alphaproteobacteria, Photosynthetic bacterial communities were studied incubating WIP and

MLSB-OSPW samples under constant light exposure. All of the OSPW photosynthetic samples are predominated by unidentified Planctomycetes (>40% of all reads), a rarely cultured phylum, suggesting a possible new method of cultivation for this phylum.

A novel operon encoding an enzyme belonging to the copper-containing membrane- bound monooxygenases (CuMMOs) enzymes group was detected in the metagenome data of the

WIP-OSPW (described in Chapter 3). This enzyme group is known capable to catalyze the oxidation of methane, ammonia and potentially C2–C4 alkanes and alkenes. In Chapter 5, it is noted that MLSB-OSPW is capable of methane oxidation with a rate up to 108.2 μmol of CH4

−1 −1 −1 −1 L OSPW d , ethane oxidation with a rate up to 83.2 μmol of C2H6 L OSPW d , and propane

−1 −1 oxidation with a rate up 58.6 μmol of C3H8 L OSPW d . Organisms in the OSPW are likely to possess divergent CuMMOs with alternate oxidation functions, enabling them to oxidize a range of short-chain alkanes such as ethane and propane. A qPCR system designed to detect the genes encoding this novel CuMMO combined with SIP identified the presence of the genes in both propane and ethane amended systems. Future direction in this study is to look at the gene expression for this particular operon in C2–C4 alkane enriched environments.

Chapter 6 describes the characterization of a novel member of the Verrucomicrobia, a rarely cultured phylum, isolated from the hydrocarbon-rich WIP-OSPW. The type strain of this isolate is NVTT. Genomic information revealed genes involved in catechol degradation pathways in the strain. Although NVTT did not grow in catechol under laboratory conditions, it can be 140

postulated that plain R2A medium might not provide conditions favourable to genes involved in aromatic compounds degradation. It is also probable that this strain is incapable of performing aromatic compounds degradation by itself, and that it may have been feeding on intermediate products from aromatic compounds degradation in the tailings pond. Phylogenetic analysis of

16S rRNA gene sequences revealed that the strain belongs to the family Opitutaceae of the phylum Verrucomicrobia. Based on genotypic, phenotypic and chemotaxonomic characteristics, it was concluded that this strain represents a novel genus and species, for which the name

Oleiharenicola alkalitolerans gen. nov., sp. nov. is proposed.

7.2 Directions for future research

Although the basic concepts of hydrocarbon degradation in polluted environments have been extensively studied, advances in sequencing technologies have opened up many possibilities for further research.

 Detection of hydrocarbon-assimilating bacteria in other environments. The SIP

technique (Chapter 3, 4, and 5) designed to detect various hydrocarbon-assimilating

bacteria that are substrate-specific in OSPW samples can also be applied to other

polluted environments to detect what organisms play the major roles in

bioremediation. In Chapter 3, we discovered that organisms such as Methyloversatilis

and an unknown Chromatiaceae (related to Thiococcus) played major degradation

roles of benzene and naphthalene in OSPW. These organisms have never been related

to hydrocarbon degradation. They may have been overlooked in the past, or that there

are certain environmental conditions in OSPW that caused them to thrive.

Identification of a more ‘unusual’ hydrocarbon-degrading organism might reveal the

possibility of studying how bacteria previously unrelated to hydrocarbon degradation

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'switch on' their metabolic machinery in the presence of abundant hydrocarbon

sources.

 Development of better primers to detect genes encoding aromatic hydrocarbon-

degrading enzymes. Unlike methanotrophy activities in the environment that can be

detected via a universal particulate methane monooxygenase (pmoA) PCR primer sets,

there is no universal set of hydrocarbon-degrading specific primers. Some

hydrocarbon-degradation related primer sets can only be linked to their corresponding

organisms, such as nah-like genes for Pseudomonas species and some

Sphingomonads. Development of targeted primer sets that improve the detection of a

certain group of taxa would enable for rapid detection of hydrocarbon-degrading

organisms in the environment for bioremediation monitoring.

13  Cultivation of novel Planctomycetes from OSPW. In Chapter 4, both CO2

incubation of MLSB-and WIP-OSPW under light over the duration of 12 weeks are

dominated by unidentified Planctomycetes, a rarely cultured phylum. Both samples

13 amended with CO2 was dominated by an unidentified Planctomycete related to

Aquisphaera sp. The phylum Planctomycetes, a rarely cultured phylum, has been

commonly associated with a wide range of algae in which they grow as complex

microbial biofilm and depend on polysaccharides secreted by algae. During sample

incubation, constant light exposure might have caused algae indigenous to OSPW to

thrive. It has been reported before that micro-algae, such as Parachlorella kessleri,

thrived on the surface of the tailings ponds. This experiment suggests a possible new

method of cultivation for this phylum by incubating OSPW samples under constant

light exposure for >4 weeks of incubation. Isolation of members from this phylum

might reveal so many interesting aspects about Planctomycetes that are previously 142

unknown, such as its roles in bioremediation, and especially that this phylum appears

to have a distinctive membrane morphology.

 Gene expression studies of the novel xmoCAB operon. In Chapter 5, it is explained

that a unique xmoCAB operon was detected in WIP-OSPW sample. Studies conducted

on this operon were merely on its detection levels in different hydrocarbon enrichment

samples. Future direction as a follow up of this xmoCAB operon study is to perform a

gene expression experiment in C2–C4 alkane enriched environments via qPCR and

RNA sequencing.

 Metagenomics of samples enriched with xmoCAB operons. In addition to gene

expression study, a metagenomic sequencing of SIP fractions that are high in the

unusual xmoCAB operon levels can also be done to detect organisms possessing the

operon.

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166

APPENDIX A: SUPPLEMENTARY INFORMATION –BENZENE AND NAPHTHALENE DEGRADING BACTERIAL COMMUNITIES IN OILSANDS TAILINGS

Figure A1. Stable Isotope Probing figures of WIP-OSPW tailings water sampled in August, 2011, with and without the addition of labelled substrates and model hydrocarbons: sodium acetate (A), methanol (B), benzene, and (C) naphthalene. Twelve SIP fractions are shown as filled circles (amended with substrates) and empty circles (DNA from OSPW tailings water or control). Fractions pointed by red arrows were taken for DNA pyrosequencing as 13C-labelled samples, fractions pointed by black arrows were the control fraction 5 (OSPW-heavy) used in this study.

167

Figure A2 A–B. Representative headspace benzene oxidation and CO2 production time courses of replicate samples from OSPW sampled in August, 2011, with the results of linear regressions for benzene oxidation. Figure A2-A. Benzene oxidation of OSPW sampled in August, 2011, with the results of linear regressions for benzene oxidation.

)

M m

( 5

t a

r 4

c n

o 3

n e

z 2

e B 1

0 0 10 20 30 40 50 60 Time (d) OSPW + Benzene y = -0.0367x + 4.5563 (R² = 0.71742) OSPW Control y = -0.0072x + 5.8197 (R² = 0.17438)

Figure A2-B. Headspace CO2 production time courses of replicate samples from OSPW sampled in August, 2011. 1.8

1.6

1.4

)

M 1.2

(

O 1

c 0.8

p s

d 0.6

e H 0.4

0.2

0 0 10 20 30 40 50 60 Time (d) OSPW + BENZENE y = 0.0273x - 0.0215 (R² = 0.95944) OSPW Control y = 0.0021x + 0.0719 (R² = 0.92847)

168

Figure A3 A-B. Representative naphthalene oxidation and headspace CO2 production time courses of replicate samples from OSPW sampled in August, 2011, with the results of linear regressions. Figure A3-A. Naphthalene oxidation of OSPW sampled in August, 2011, with the results of linear regressions. 5

4.5

) 3.5

m (

n e

l 2.5

h t

h 2

a N 1.5

0.5

0 0 2 4 6 8 10 12 14 Time (d)

OSPW + Naphthalene y = -0.0854x + 3.823 (R² = 0.90852) OSPW Control y = 0.0048x + 4.2556 (R² = 0.13138)

Figure A3-B. Representative CO2 production time courses of replicate samples from OSPW sampled in August, 2011. 5

4.5

)

M 3.5

(

2 3

e 2.5

a p

s 2

d a

e 1.5 H

0.5

0 0 2 4 6 8 10 12 14 Time (d)

OSPW + Naphthalene y = 0.2323x + 0.9559 (R² = 0.97646) OSPW Control y = 0.0024x + 0.0786 (R² = 0.75967)

169

Figure A4 A-B. Representative CO2 production time courses of replicate samples used for labelled hydrocarbons SIP of WIP-OSPW sampled in August, 2011.

Figure A4-B. Representative CO2 production time courses of replicate samples used for labelled substrates SIP of WIP-OSPW sampled in August, 2011.

170

Figure A5. Bubble chart of all samples based on relative abundances (>1%). Classification percentage is shown for class rank (% classified).

171

Figure A6. Protein coding genes statistics of WIP-2011 tailings pond metagenome based on its KEGG Categories analyzed on IMG.

Amino acid metabolism Carbohydrate metabolism Energy metabolism Metabolism of cofactors and vitamins Membrane transport Nucleotide metabolism Signal transduction Lipid metabolism Xenobiotics biodegradation and metabolism Metabolism of other amino acids Translation Replication and repair Metabolism of terpenoids and polyketides Folding, sorting and degradation Glycan biosynthesis and metabolism Cell motility Cell growth and death Biosynthesis of other secondary metabolites Infectious diseases: Bacterial Endocrine system Transport and catabolism Transcription Environmental adaptation Neurodegenerative diseases Nervous system Endocrine and metabolic diseases Circulatory system Infectious diseases: Parasitic Excretory system Digestive system Immune system Immune diseases Infectious diseases: Viral Cardiovascular diseases Chemical structure transformation maps Substance dependence Cell communication Signaling molecules and interaction Development Sensory system

172

Figure A6 A-B. Metagenomic community structure of OSPW sampled in August, 2011, with >30% number of hits on IMG/GenBank database. Entire dataset of all the genes was 172,777 reads. Figure A6-A. Metagenomic community structure based on domains.

Figure A6-B. Metagenomic community structure of OSPW based on the orders of Proteobacteria using entire dataset of all the genes detected.

0 Orders in Proteobacteria Burkholderiales Rhizobiales Sphingomonadales Rhodobacterales Alteromonadales Rhodocyclales Rhodospirillales Methylophilales Oceanospirillales Thiotrichales Myxococcales Pseudomonadales Vibrionales Campylobacterales Rickettsiales Unclassiﬁed Proteobacteria Caulobacterales Chromatiales Methylococcales Enterobacteriales Xanthomonadales Aeromonadales 173

Figure A7. Benzene degradation by pure cultures of Xanthobacter sp. OSPW28 and Zavarzinia compransoris sp. OSPW6, isolated from OSPW. Experiments were done in 2 replicates.

8 Blank Xanthobacter 7

Zavarzinia

)

M 6

(

n 5

n e

b 4

c a

p 3

d a

e 2 H

0 0 2 4 6 8 10 12 14 Time (d)

174 Table A1. List of bacterial isolates obtained from various OSPW enrichments and their closest relatives.

Percen Source/ Sample(s) t Benze Query GenBank Medium where numbe % Strain ne Phylum Class Order Family Genus length taxonomy of OTUs r of Identity name degra (bp) isolation were found reads dation (%) Proteobacteria Alphaproteobacteri Rhizobiales Xanthobacteraceae Xanthobacter Xanthobacter 20R2A-T 1205 97 OSPW - a tagetidis 1 Proteobacteria Gammaproteobacte Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas 20R2A-T 1329 99 OSPW - ria xanthomarina 2 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera Thauera 20R2A-T SA, M, N, 6, <1, 1387 99 OSPW - phenylacetica with C, OSPW 21, 9, 2 3 naphthale ne vapour Proteobacteria Alphaproteobacteri Caulobacterales Caulobacteraceae Brevundimona Brevundimonas 20R2A-T SA, C, 9, 9, 6 1241 99 OSPW - a s nasdae OSPW 4 Proteobacteria Actinobacteria Micrococcales Microbacteriaceae Microbacteriu Microbacterium 20R2A-T SA 1 1322 98 OSPW - m paraoxydans 5 Proteobacteria Alphaproteobacteri Rhodospirillales Acetobacteraceae Zavarzinia Zavarzinia M1O-T B 5 1355 99 OSPW + a compransoris 6 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Variovorax Variovorax 20R2A-T ALL <1 1361 100 OSPW - paradoxus 7 Proteobacteria Alphaproteobacteri Sphingomonadales Erythrobacteraceae Porphyrobacte Pseudomonas 20R2A-T SA, M, N, 5, 2, 8, 1374 99 OSPW - a r stutzeri with OSPW 1 8 naphthale ne vapour Proteobacteria Gammaproteobacte Xanthomonadales Xanthomonadaceae Aquimonas Aquimonas voraii M1O-T C, SA, N <1 1303 99 OSPW ria 9 Bacteroidetes Cytophagia Cytophagales Cyclobacteriaceae Algoriphagus Algoriphagus 20R2A-T OSPW 3 1383 99 OSPW - alkaliphilus 10

Proteobacteria Alphaproteobacteri Rhizobiales Xanthobacteraceae Ancylobacter Ancylobacter M1O-T SA 1 1296 99 OSPW - a polymorphus 11 Proteobacteria Betaproteobacteria Burkholderiales Methylibium Aquabacterium M1O-T C, OSPW <1 1069 98 OSPW - fontiphilum 12

Actinobacteria Actinobacteria Micrococcales Micrococcaceae Arthrobacter Arthrobacter M1O-T C <1 1353 99 OSPW - agilis 13 Proteobacteria Alphaproteobacteri Caulobacterales Caulobacteraceae Caulobacter Caulobacter 20R2A-T SA, M, N 12, <1, 1327 98 OSPW - a fusiformis <1 14 Proteobacteria Alphaproteobacteri Rhizobiales Hyphomicrobiaceae Devosia Devosia 20R2A-T SA, C, 21, 23, 1303 97 OSPW - a submarina OSPW 8 15

175

Bacteroidetes Sphingobacteria Sphingobacteriales Cyclobacteriaceae Indibacter Indibacter M1O-T 1385 93 OSPW - alkaliphilus with 16 methanol Proteobacteria Alphaproteobacteri Rhizobiales Phyllobacteriaceae Mesorhizobiu Mesorhizobium 20R2A-T M <1 1336 99 OSPW - a m alhagi 17 Actinobacteria Actinobacteria Micrococcales Microbacteriaceae Chryseoglobus Chryseoglobus 20R2A-T C, OSPW <1 1354 99 OSPW - frigidaquae 18

Proteobacteria Alphaproteobacteri Rhodospirillales Rhodospirillaceae Oceanibaculu Oceanibaculum 20R2A-T C, OSPW <1 1277 99 OSPW a m indicum 19 Proteobacteria Alphaproteobacteri Rhodospirillales Acetobacteraceae Oleomonas Oleomonas 20R2A-T B, OSPW <1 1314 99 OSPW a sagaranensis 20 Verrucomicrobi Opitutae Opitutales Opitutaceae Opitutus Opitutus terrae M1O-T C, OSPW <1 1368 94 OSPW - a 21 Proteobacteria Alphaproteobacteri Sphingomonadales Erythrobacteraceae Porphyrobacte Porphyrobacter 20R2A-T N, C, <1, 2, 4 1247 99 OSPW - a r sanguineus OSPW 22 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax Acidovorax 20R2A-T N 2 1364 93 OSPW facilis 23 Proteobacteria Gammaproteobacte Xanthomonadales Xanthomonadaceae Pseudoxantho Pseudoxanthomo 20R2A-T 1405 99 OSPW ria monas nas mexicana 24

Proteobacteria Betaproteobacteria Burkholderiales Ralstoniaceae Ralstonia Ralstonia 20R2A-T N, C, <1 1387 99 OSPW - pickettii OSPW 25 Proteobacteria Alphaproteobacteri Rhizobiales Rhizobiaceae Rhizobium Rhizobium 20R2A-T NA NA 1310 99 OSPW a naphthalenivoran 26 s Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Rhodoferax Rhodoferax M10-T C 1.5 1393 99 OSPW antarticus with 27 propane

Proteobacteria Alphaproteobacteri Rhizobiales Xanthobacteraceae Xanthobacter Xanthobacter 20R2A-T OSPW, M, 3, <1, 1287 98 OSPW + a autotrophicus SA 15 28

176

Table A2 A-E. Top 25 OTUs of each sample site based on % of total reads in 16S rRNA gene sequencing analysis. OTUs were clustered based on their closest related species. Table A2-A. Reads from DNA in OSPW sampled in August 2011.

Rank % Reads Phylum Class Order Family Genus % Identity 1 4.66 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Hydrogenophaga 99 2 3.88 Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Lutibacter 98 3 3.38 Proteobacteria Gammaproteobacteria Pseudomonadales Moraxellaceae Enhydrobacter 99 4 2.99 Proteobacteria Betaproteobacteria Burkholderiales Burkholderiaceae Limnobacter 99 5 2.99 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylogaea 94 6 2.89 Bacteroidetes Saprospiraceae Sphingobacteriales Sphingobacteriia Haliscomenobacter 93 7 2.59 Bacteroidetes Bacteroidia Bacteroidales Porphyromonadaceae Barnesiella 90 8 2.59 Bacteroidetes Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola 96 9 2.39 Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Flavobacterium 98 10 2.29 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Xanthobacter 99 11 2.19 Proteobacteria Betaproteobacteria Burkholderiales Burkholderiaceae Sphaerotilus 99 12 1.89 Bacteroidetes Sphingobacteriia Sphingobacteriales Chitinophagaceae Balneola 89 13 1.79 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Rhodocyclus 96 14 1.59 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Roseimaritima 93 15 1.39 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Variovorax 99 16 1.39 Proteobacteria Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Variibacter 94 17 1.29 Planctomycetes Planctomycetia Planctomycetales Isosphaeraceae Aquisphaera 90 18 1.19 Proteobacteria Betaproteobacteria Burkholderiales Alcaligenaceae Achromobacter 96 19 1.19 Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Flavobacterium 98 20 1.09 Tenericutes Mollicutes Acholeplasmatales Acholeplasmataceae Acholeplasma 95 21 1.09 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Zoogloea 95 22 1.09 Proteobacteria Betaproteobacteria NA NA Imtechium 99 23 1.00 Proteobacteria Alphaproteobacteria Rhodobacterales Hyphomonadaceae Hyphomonas 92 24 0.90 Proteobacteria Betaproteobacteria Methylophilales Methylophilaceae Methylotenera 98 25 0.90 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Gimesia 90

177

Table A2-B. Heavy SIP fraction (fraction 5) of OSPW sampled in August, 2011.

Rank % Reads Phylum Class Order Family Genus % Identity

1 23.31 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Devosia 96 2 12.64 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylomonas 96 3 9.03 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera 99 4 8.53 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas 100 5 8.29 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylogaea 94 6 6.79 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 87 7 4.95 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylomicrobium 99 8 4.52 Proteobacteria Alphaproteobacteria Magnetococcales Magnetococcaceae Magnetococcus 90 9 4.25 Proteobacteria Alphaproteobacteria Rhodospirillales Acetobacteraceae Acidiphilium 99 10 4.21 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 100 11 3.28 Euryarchaeota Methanomicrobia Methanomicrobiales Methanoregulaceae Methanoregula 99 12 2.98 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 86 13 2.88 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Hydrogenophaga 99 14 2.47 Proteobacteria Gammaproteobacteria Xanthomonadales Algiphilaceae Algiphilus 94 15 2.34 Proteobacteria Alphaproteobacteria Sphingomonadales Erythrobacteraceae Altererythrobacter 99 16 2.21 Euryarchaeota Methanomicrobia Magnetococcales 99 17 2.17 Euryarchaeota Methanomicrobia Methanosarcinales Methanosaetaceae Methanosaeta 100 18 1.97 Proteobacteria Alphaproteobacteria Sphingomonadales Erythrobacteraceae Porphyrobacter 100 19 1.81 Proteobacteria Gammaproteobacteria Chromatiales Chromatiaceae Thiococcus 95 20 1.81 Proteobacteria Alphaproteobacteria Rhodocyclales Rhodocyclaceae Azospirillum 93 21 1.81 Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Lutibacter 98 22 1.74 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 89 23 1.57 Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Flavobacterium 99 24 1.51 Proteobacteria Gammaproteobacteria Magnetococcales Methylococcaceae Methylomicrobium 97 25 1.40 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Rhodoferax 98

178

Table A2-C. Heavy SIP Fractions (fractions 4 and 5) of OSPW sampled in August, 2011, amended with labelled benzene for 9 days.

Rank % Reads Phylum Class Order Family Genus % Identity

1 86.87 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 100 2 5.18 Proteobacteria Alphaproteobacteria Rhodospirillales Acetobacteraceae Zavarzinia 99 3 1.39 Proteobacteria Gammaproteobacteria Xanthomonadales Algiphilaceae Algiphilus 94 4 0.62 Proteobacteria Deltaproteobacteria Syntrophobacterales Syntrophobacteraceae Desulforhabdus 90 5 0.51 Proteobacteria Deltaproteobacteria Desulfuromonadales Geobacteraceae Geobacter 91 6 0.42 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Rhodocyclus 96 7 0.24 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Dechloromonas 94 8 0.23 Proteobacteria Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium 95 9 0.19 Chloroflexi Anaerolineae Anaerolineales Anaerolineaceae Bellilinea 92 10 0.17 Proteobacteria Gammaproteobacteria Chromatiales Chromatiaceae Thiococcus 95 11 0.17 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Gemmata 91 12 0.16 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 86 13 0.13 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Inquilinus 94 14 0.11 Tenericutes Mollicutes Entomoplasmatales Spiroplasmataceae Spiroplasma 75 15 0.09 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Sulfuritalea 99 16 0.08 Proteobacteria Deltaproteobacteria Desulfuromonadales Desulfuromonadaceae Desulfuromusa 90 17 0.07 Acidobacteria Solibacteres Solibacterales Solibacterales Bryobacter 93 18 0.07 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Caenimonas 98 19 0.07 Proteobacteria Alphaproteobacteria Sphingomonadales Erythrobacteraceae Altererythrobacter 99 20 0.06 Proteobacteria Deltaproteobacteria Desulfobacterales Desulfobacteraceae Desulfotignum 85 21 0.06 Chloroflexi Caldilineae Caldilineales Caldilineaceae Litorilinea 88 22 0.05 Proteobacteria Gammaproteobacteria Alteromonadales Alteromonadaceae Microbulbifer 95 23 0.05 Proteobacteria Alphaproteobacteria Rhodospirillales Acetobacteraceae Oleomonas 93 24 0.05 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 94 25 0.05 Proteobacteria Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum 100

179

Table A2-D. Heavy SIP Fractions (fractions 5 and 6) of OSPW sampled in August, 2011, amended with labelled naphthalene for 7 days.

Rank % Reads Phylum Class Order Family Genus % Identity

1 45.56 Proteobacteria Gammaproteobacteria Chromatiales Chromatiaceae Thiococcus 95 2 20.50 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera 99 3 8.19 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 100 4 2.69 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 96 5 5.73 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 98 6 1.61 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax 99 7 1.24 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 97 8 1.23 Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Gemmobacter 99 9 0.73 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 98 10 0.57 Chloroflexi Caldilineae Caldilineales Caldilineaceae Caldilinea 83 11 0.39 Proteobacteria Betaproteobacteria Rhodocyclales Rhodobacteraceae Azoarcus 99 12 0.39 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Azotobacter 100 13 0.39 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 98 14 0.35 Proteobacteria Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium 98 15 0.34 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 98 16 0.30 Proteobacteria Alphaproteobacteria Sphingomonadales Erythrobacteraceae Porphyrobacter 100 17 0.28 Proteobacteria Gammaproteobacteria Legionellales Coxiellaceae Aquicella 95 18 0.24 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 99 19 0.23 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Phenylobacterium 97 20 0.19 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 98 21 0.17 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Rubinisphaera 91 22 0.15 Proteobacteria Gammaproteobacteria Xanthomonadales Algiphilaceae Algiphilus 94 23 0.15 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 100 24 0.12 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 97 25 0.11 Chloroflexi Caldilineae Caldilineales Caldilineaceae Litorilinea 88

180

Table A2-E. Heavy SIP Fractions (fractions 3 and 4) of OSPW sampled in August, 2011, amended with labelled methanol for 7 days.

Rank % Reads Phylum Class Order Family Genus % Identity

1 52.96 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 100 2 3.54 Proteobacteria Deltaproteobacteria Syntrophobacterales Syntrophobacteraceae Desulforhabdus 90 3 2.13 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Hyphomicrobium 96 4 1.49 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus 99 5 1.19 Proteobacteria Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium 98 6 1.12 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Delftia 100 7 1.04 Chloroflexi Caldilineae Caldilineales Caldilineaceae Litorilinea 86 8 1.03 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 96 9 1.01 Proteobacteria Gammaproteobacteria Chromatiales Chromatiaceae Thiococcus 95 10 1.00 Actinobacteria Acidimicrobiia Acidimicrobiales Acidimicrobiaceae Aciditerrimonas 91 11 0.88 Actinobacteria Actinobacteria Propionibacteriales Propionibacteriaceae Propionibacterium 99 12 0.85 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 99 13 0.80 Proteobacteria Alphaproteobacteria Rhizobiales Phyllobacteriaceae Mesorhizobium 99 14 0.58 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera 99 15 0.57 Bacteroidetes Sphingobacteriia Sphingobacteriales Chitinophagaceae Sediminibacterium 96 16 0.54 Armatimonadetes Fimbriimonadia Fimbriimonadales Fimbriimonadaceae Fimbriimonas 93 17 0.46 Proteobacteria Betaproteobacteria Methylophilales Methylophilaceae Methylotenera 98 18 0.43 Chloroflexi Caldilineae Caldilineales Caldilineaceae Litorilinea 88 19 0.43 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Xanthobacter 99 20 0.41 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Caulobacter 99 21 0.41 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Pirellula 96 22 0.35 Chloroflexi Chloroflexia Chloroflexales Chloroflexaceae Chloroflexus 78 23 0.35 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Ancylobacter 99 24 0.35 Proteobacteria Betaproteobacteria Burkholderiales Alcaligenaceae Achromobacter 100 25 0.35 Chlamydiae Chlamydiae Chlamydiales Criblamydiaceae Criblamydia 90

181 Table A3. Metagenome features of surface WIP-OSPW sampled in August 2011 of the combined Illumina and 454 assembly obtained from the IMG pipeline.

Assembled

Number % of Assembled

Number of sequences 1,520,282 100.00% Number of bases 728,742,540 100.00% GC count 426,300,665 58.50%

Genes RNA genes 11,467 0.63% rRNA genes 2,282 0.13% 5S rRNA 396 0.02% 16S rRNA 632 0.03% 18S rRNA 38 0.00% 23S rRNA 1,170 0.06% 28S rRNA 46 0.00% tRNA genes 9,185 0.50% Protein coding genes 1,811,780 99.37% with Product Name 852,731 46.77% with COG 1,088,850 59.72% with Pfam 961,630 52.74% with KO 873,050 47.88% with Enzyme 511,566 28.06% with MetaCyc 360,503 19.77% with KEGG 520,901 28.57% COG Clusters 4,559 98.45% Pfam Clusters 6,650 40.81% Biosynthetic Clusters 554 Genes in Biosynthetic Clusters 2,711

182 Table A4 A-P. Genes encoding aerobic hydrocarbon (benzene and naphthalene) metabolism detected in the OSPW metagenome, and their top

BLAST hits (to a cultured organism). The minimum aligned scaffold length was 2500 nucleotides (for 16S rRNA). All read depth = 1.

Hits Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species (%)

93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.4 Pseudomonas stutzeri A1501 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 89.5 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 88.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 87.9 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 87.9 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 87.9 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 87.9 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 87.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.3 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 83.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 82.1 Chelativorans sp. BNC1 Alphaproteobacteria Rhizobiales Phyllobacteriaceae Chelativorans Chelativorans sp. BNC1 81.9 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 81.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 80.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 80 Streptomyces scabrisporus DSM 41855 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces scabrisporus 79.8 Pseudomonas sp. CT14 plasmid pCT14 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas

183

79.6 Cupriavidus metallidurans CH34 Betaproteobacteria Burkholderiales Burkholderiaceae Cupriavidus Cupriavidus metallidurans 78.7 Thiocapsa marina 5811, DSM 5653 Gammaproteobacteria Chromatiales Chromatiaceae Thiocapsa Thiocapsa marina 76.1 Bradyrhizobium sp. Th.b2 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. th.b2 76 Amycolicicoccus subflavus DQS3-9A1 Actinobacteria Actinomycetales Mycobacteriaceae Amycolicicoccus Amycolicicoccus subflavus 75.4 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 74.3 Methyloversatilis sp. RZ18-153 Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis Methyloversatilis sp. RZ18-153 73.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.9 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 72.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 Saccharomonospora glauca K62, DSM 72.4 43769 Actinobacteria Actinomycetales Pseudonocardiaceae Saccharomonospora Saccharomonospora glauca 71.8 Humibacter albus DSM 18994 Actinobacteria Actinomycetales Microbacteriaceae Humibacter Humibacter albus Promicromonospora sukumoe 71.2 327MFSha3.1 Actinobacteria Actinomycetales Promicromonosporaceae Promicromonospora Promicromonospora sukumoe 69.5 Meganema perideroedes DSM 15528 Alphaproteobacteria Rhizobiales Methylobacteriaceae Meganema Meganema perideroedes 68.7 Azospira suillum PS Betaproteobacteria Rhodocyclales Rhodocyclaceae Azospira Azospira suillum 68.4 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 67.8 Azospira suillum PS Betaproteobacteria Rhodocyclales Rhodocyclaceae Azospira Azospira suillum 67 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 66.3 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 65.5 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri Polynucleobacter necessarius 65 asymbioticus QLW-P1DMWA-1 Betaproteobacteria Burkholderiales Burkholderiaceae Polynucleobacter Polynucleobacter necessarius Microbacterium paraoxydans 64.2 77MFTsu3.2 Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium paraoxydans 62.3 Aromatoleum aromaticum EbN1 Betaproteobacteria Rhodocyclales Rhodocyclaceae Aromatoleum Aromatoleum aromaticum 62 Methylobacter tundripaludum SV96 Gammaproteobacteria Methylococcales Methylococcaceae Methylobacter Methylobacter tundripaludum Uliginosibacterium gangwonense DSM Uliginosibacterium 60.9 18521 Betaproteobacteria Rhodocyclales Rhodocyclaceae Uliginosibacterium gangwonense marine actinobacterium 60.8 marine actinobacterium PHSC20C1 Actinobacteria unclassified unclassified unclassified PHSC20C1 60.6 Azonexus hydrophilus DSM 23864 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azonexus Azonexus hydrophilus Nakamurella multipartita Y-104, DSM 60.4 44233 Actinobacteria Actinomycetales Nakamurellaceae Nakamurella Nakamurella multipartita 59.3 Herbaspirillum sp. GW103 Betaproteobacteria Burkholderiales Oxalobacteraceae Herbaspirillum Herbaspirillum sp. GW103 57.4 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus

184

57.1 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 56.6 Frankia sp. EAN1pec Actinobacteria Actinomycetales Frankiaceae Frankia Frankia sp. EAN1pec 56 Nocardia brasiliensis ATCC 700358 Actinobacteria Actinomycetales Nocardiaceae Nocardia Nocardia brasiliensis Polynucleobacter necessarius 55.9 necessarius STIR1 (Endosymbiont) Betaproteobacteria Burkholderiales Burkholderiaceae Polynucleobacter Polynucleobacter necessarius 55.3 Tepidiphilus margaritifer DSM 15129 Betaproteobacteria Hydrogenophilales Hydrogenophilaceae Tepidiphilus Tepidiphilus margaritifer Nakamurella multipartita Y-104, DSM 54.7 44233 Actinobacteria Actinomycetales Nakamurellaceae Nakamurella Nakamurella multipartita 52.9 Roseobacter sp. GAI101 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Sulfitobacter Roseobacter sp. GAI101 52.4 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 52.4 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 51.8 Microbacterium sp. 11MF Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium sp. 11MF 50.9 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 47.9 Desulfobacula toluolica Tol2 Deltaproteobacteria Desulfobacterales Desulfobacteraceae Desulfobacula Desulfobacula toluolica 47.6 Mesorhizobium loti R88b Alphaproteobacteria Rhizobiales Phyllobacteriaceae Mesorhizobium Mesorhizobium loti 47.1 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 44.4 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 43.9 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 43.8 Rhodococcus opacus B4 Actinobacteria Actinomycetales Nocardiaceae Rhodococcus Rhodococcus opacus 42.4 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 41.9 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 41.9 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 35.4 Dickeya zeae Ech1591 Gammaproteobacteria Enterobacteriales Enterobacteriaceae Dickeya Dickeya zeae 34.3 Thioalkalivibrio sp. ALE11 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Thioalkalivibrio Thioalkalivibrio sp. ALE11 30.5 Teredinibacter turnerae T8513 Gammaproteobacteria Alteromonadales unclassified Teredinibacter Teredinibacter turnerae

185

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

96.7 Thiobacillus denitrificans ATCC 25259 Betaproteobacteria Hydrogenophilales Hydrogenophilaceae Thiobacillus Thiobacillus denitrificans 96.3 Thiobacillus denitrificans DSM 12475 Betaproteobacteria Hydrogenophilales Hydrogenophilaceae Thiobacillus Thiobacillus denitrificans 93.7 Verminephrobacter eiseniae EF01-2 Betaproteobacteria Burkholderiales Comamonadaceae Verminephrobacter Verminephrobacter eiseniae 93.6 Thiobacillus denitrificans DSM 12475 Betaproteobacteria Hydrogenophilales Hydrogenophilaceae Thiobacillus Thiobacillus denitrificans 93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 92.9 Alicycliphilus denitrificans BC Betaproteobacteria Burkholderiales Comamonadaceae Alicycliphilus Alicycliphilus denitrificans 92.5 Thiobacillus thioparus DSM 505 Betaproteobacteria Hydrogenophilales Hydrogenophilaceae Thiobacillus Thiobacillus thioparus 92.1 Thiobacillus thioparus DSM 505 Betaproteobacteria Hydrogenophilales Hydrogenophilaceae Thiobacillus Thiobacillus thioparus 91.7 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.4 Pseudomonas stutzeri A1501 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90 Azoarcus sp. BH72 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus sp. BH72 89.5 Dechloromonas aromatica RCB Betaproteobacteria Rhodocyclales Rhodocyclaceae Dechloromonas Dechloromonas aromatica 89.5 Dechloromonas aromatica RCB Betaproteobacteria Rhodocyclales Rhodocyclaceae Dechloromonas Dechloromonas aromatica 89.5 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 89.1 Dechloromonas aromatica RCB Betaproteobacteria Rhodocyclales Rhodocyclaceae Dechloromonas Dechloromonas aromatica 88.9 Alicycliphilus denitrificans BC Betaproteobacteria Burkholderiales Comamonadaceae Alicycliphilus Alicycliphilus denitrificans 88.9 Alicycliphilus denitrificans BC Betaproteobacteria Burkholderiales Comamonadaceae Alicycliphilus Alicycliphilus denitrificans 88.4 Thiobacillus denitrificans DSM 12475 Betaproteobacteria Hydrogenophilales Hydrogenophilaceae Thiobacillus Thiobacillus denitrificans 88.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 87.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 87.1 Bradyrhizobium elkanii WSM2783 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium elkanii 86.4 Methyloversatilis universalis EHg5 Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis Methyloversatilis universalis 86 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri

186

85.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.3 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 83.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 81.9 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 81.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 81.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 80.9 Burkholderia vietnamiensis G4 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia vietnamiensis 80.9 Burkholderia vietnamiensis G4 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia vietnamiensis 80.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 80.7 Burkholderia mimosarum STM3621 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia mimosarum 80.7 Burkholderia mimosarum STM3621 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia mimosarum 79.4 Silicibacter sp. TrichCH4B Alphaproteobacteria Rhodobacterales Rhodobacteraceae Ruegeria Silicibacter sp. TrichCH4B 79.2 Pseudomonas sp. GM78 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM78 78.4 Nitrosococcus halophilus Nc4 Gammaproteobacteria Chromatiales Chromatiaceae Nitrosococcus Nitrosococcus halophilus 78.3 Rhodobacter capsulatus SB1003 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Rhodobacter Rhodobacter capsulatus 76.4 Achromobacter piechaudii HLE Betaproteobacteria Burkholderiales Alcaligenaceae Achromobacter Achromobacter piechaudii 75.8 Bradyrhizobium sp. S23321 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. S23321 75.4 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 75.3 Starkeya novella DSM 506 Alphaproteobacteria Rhizobiales Xanthobacteraceae Starkeya Starkeya novella 74.1 Roseovarius sp. TM1035 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Roseovarius Roseovarius sp. TM1035 73.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.9 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 72.6 Bordetella petrii Se-1111R, DSM 12804 Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella petrii 72.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.3 Bordetella petrii Se-1111R, DSM 12804 Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella petrii 71.8 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 70.8 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 70.8 Nonomuraea coxensis DSM 45129 Actinobacteria Actinomycetales Streptosporangiaceae Nonomuraea Nonomuraea coxensis 70.1 Bradyrhizobium elkanii USDA 3254 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium elkanii

187

70 Pseudomonas sp. GM78 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM78 69.7 Bordetella petrii Se-1111R, DSM 12804 Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella petrii 68.7 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 68.5 Marinobacter adhaerens HP15 Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Marinobacter adhaerens 68.4 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 67.9 Melitea salexigens DSM 19753 Gammaproteobacteria Alteromonadales Alteromonadaceae Melitea Melitea salexigens 67 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 66.9 Bordetella petrii Se-1111R, DSM 12804 Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella petrii 66.7 Bordetella petrii Se-1111R, DSM 12804 Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella petrii 66.7 Methyloversatilis universalis EHg5 Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis Methyloversatilis universalis 66.3 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 66 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 66 Rhodococcus opacus B4 Actinobacteria Actinomycetales Nocardiaceae Rhodococcus Rhodococcus opacus 65.5 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri Polaromonas 64.9 Polaromonas naphthalenivorans CJ2 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas naphthalenivorans 64.2 Burkholderia pseudomallei 1026b Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia pseudomallei 63.7 Alcaligenes faecalis phenolicus DSM 16503 Betaproteobacteria Burkholderiales Alcaligenaceae Alcaligenes Alcaligenes faecalis 63.4 Burkholderia pseudomallei 1026b Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia pseudomallei 63.3 Oxalobacter formigenes HOxBLS Betaproteobacteria Burkholderiales Oxalobacteraceae Oxalobacter Oxalobacter formigenes 62.2 Anabaena variabilis ATCC 29413 unclassified Nostocales Nostocaceae Anabaena Anabaena variabilis 62.2 Loktanella hongkongensis DSM 17492 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Loktanella Loktanella hongkongensis 61.1 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 60.8 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 60.3 Pseudomonas sp. GM55 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM55 59.6 Methyloversatilis universalis EHg5 Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis Methyloversatilis universalis 58.8 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 58.5 Acidovorax sp. CF316 Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax Acidovorax sp. CF316 56.5 Pseudomonas protegens Pf-5 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas protegens 56.5 Rhodobacter sphaeroides 2.4.1 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Rhodobacter Rhodobacter sphaeroides 56.4 Haliea salexigens DSM 19537 Gammaproteobacteria Alteromonadales Alteromonadaceae Haliea Haliea salexigens

188

55.6 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 54.2 Gordonia rhizosphera NBRC 16068 Actinobacteria Actinomycetales Gordoniaceae Gordonia Gordonia rhizosphera 53.6 Rhizobium rhizogenes bv. II K84 Alphaproteobacteria Rhizobiales Rhizobiaceae Agrobacterium Agrobacterium radiobacter 52.8 Salinarimonas rosea DSM 21201 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Salinarimonas Salinarimonas rosea Bradyrhizobium sp. 49.3 Bradyrhizobium sp. WSM2254 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium WSM2254 48.1 Nocardia sp. 348MFTsu5.1 Actinobacteria Actinomycetales Nocardiaceae Nocardia Nocardia sp. 348MFTsu5.1 46.2 Chelativorans sp. BNC1 Alphaproteobacteria Rhizobiales Phyllobacteriaceae Chelativorans Chelativorans sp. BNC1 41.7 Chelativorans sp. BNC1 Alphaproteobacteria Rhizobiales Phyllobacteriaceae Chelativorans Chelativorans sp. BNC1 Novosphingobium 37.9 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium nitrogenifigens

34.3 Thioalkalivibrio sp. ALE11 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Thioalkalivibrio Thioalkalivibrio sp. ALE11

189

Table A4-C. Genes encoding for dmpH/xylI/nahK (gene 2-oxo-3-hexenedioate decarboxylase) detected in the OSPW metagenome, and their top

BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

96.3 Burkholderia cepacia 2a plasmid pIJB1 Betaproteobacteria Betaproteobacteria Burkholderiales Burkholderiaceae cepacia 94.9 Burkholderia cepacia 2a plasmid pIJB1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia cepacia 93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 91.7 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.6 Thauera aminoaromatica MZ1T Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera Thauera aminoaromatica 90.4 Pseudomonas stutzeri A1501 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 89.5 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus Pseudomonas sp. Chol1 88.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 88.3 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 87.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Burkholderia mimosarum 86.2 Burkholderia mimosarum LMG 23256 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Azoarcus sp. BH72 86.1 Azoarcus sp. BH72 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 85.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus 85.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.3 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus Azoarcus toluclasticus 84.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Pseudomonas sp. Chol1 83.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Flavobacteria bacterium MS220-5C 82.3 (unscreened) Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium Flavobacterium sp. MS220-5C 81.9 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 81.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 Flavobacteria bacterium MS390-1F Flavobacteria bacterium 81.2 (unscreened) Flavobacteriia unclassified unclassified unclassified MS390-1F

190

80.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 79.4 Silicibacter sp. TrichCH4B Alphaproteobacteria Rhodobacterales Rhodobacteraceae Ruegeria Silicibacter sp. TrichCH4B 78.4 Nitrosococcus halophilus Nc4 Gammaproteobacteria Chromatiales Chromatiaceae Nitrosococcus Nitrosococcus halophilus 78.3 Rhodobacter capsulatus SB1003 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Rhodobacter Rhodobacter capsulatus Polaromonas 77 Polaromonas naphthalenivorans CJ2 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas naphthalenivorans 75.4 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 75.2 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 74.1 Roseovarius sp. TM1035 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Roseovarius Roseovarius sp. TM1035 73.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.9 Thiothrix nivea JP2, DSM 5205 Gammaproteobacteria Thiotrichales Thiotrichaceae Thiothrix Thiothrix nivea 72.9 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 72.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.1 Polaromonas sp. JS666 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas Polaromonas sp. JS666 Mucilaginibacter paludis TPT56, DSM 71.9 18603 Sphingobacteriia Sphingobacteriales Sphingobacteriaceae Mucilaginibacter Mucilaginibacter paludis 71.4 Renibacterium salmoninarum ATCC 33209 Actinobacteria Actinomycetales Micrococcaceae Renibacterium Renibacterium salmoninarum 70.9 Burkholderia sp. JPY347 Betaproteobacteria Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia sp. JPY347 70.8 Nonomuraea coxensis DSM 45129 Actinobacteria Actinomycetales Streptosporangiaceae Nonomuraea Nonomuraea coxensis Intrasporangium calvum 7KIP, DSM 70.8 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 70.2 Ensifer sp. BR816 Alphaproteobacteria Rhizobiales Rhizobiaceae Ensifer Ensifer sp. BR816 68.6 Methylibium petroleiphilum PM1 Betaproteobacteria Burkholderiales unclassified Methylibium Methylibium petroleiphilum 68.5 Marinobacter adhaerens HP15 Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Marinobacter adhaerens 68.4 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 67.9 Melitea salexigens DSM 19753 Gammaproteobacteria Alteromonadales Alteromonadaceae Melitea Melitea salexigens 67.8 Bradyrhizobium elkanii WSM2783 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium elkanii 67 Ramlibacter tataouinensis TTB310 Betaproteobacteria Burkholderiales Comamonadaceae Ramlibacter Ramlibacter tataouinensis 67 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 66.3 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 66 Rhodococcus opacus B4 Actinobacteria Actinomycetales Nocardiaceae Rhodococcus Rhodococcus opacus 65.8 Methylocystis rosea SV97T Alphaproteobacteria Rhizobiales Methylocystaceae Methylocystis Methylocystis rosea 65.5 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 191

65.2 Azospirillum sp. B510 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum Azospirillum sp. B510 62.2 Loktanella hongkongensis DSM 17492 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Loktanella Loktanella hongkongensis Intrasporangium calvum 7KIP, DSM 60.8 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 60.1 Alkalilimnicola ehrlichii MLHE-1 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Alkalilimnicola Alkalilimnicola ehrlichii Flavobacteria bacterium MS390-1F Flavobacteria bacterium 58.8 (unscreened) Flavobacteriia unclassified unclassified unclassified MS390-1F 58 Microbacterium sp. 292MF Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium sp. 292MF 58 Acidovorax delafieldii 2AN Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax Acidovorax delafieldii 57.1 Mycobacterium sp. JLS Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium sp. JLS 56.6 Meiothermus rufus DSM 22234 Deinococci Thermales Thermaceae Meiothermus Meiothermus rufus 56.5 Rhodobacter sphaeroides 2.4.1 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Rhodobacter Rhodobacter sphaeroides 56.4 Haliea salexigens DSM 19537 Gammaproteobacteria Alteromonadales Alteromonadaceae Haliea Haliea salexigens 55.6 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 54.2 Gordonia rhizosphera NBRC 16068 Actinobacteria Actinomycetales Gordoniaceae Gordonia Gordonia rhizosphera 53.6 Rhizobium rhizogenes bv. II K84 Alphaproteobacteria Rhizobiales Rhizobiaceae Agrobacterium Agrobacterium radiobacter 52.6 Cellulomonas flavigena 134, DSM 20109 Actinobacteria Micrococcales Cellulomonadaceae Cellulomonas Cellulomonas flavigena 52.3 Microbacterium testaceum StLB037 Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium testaceum 52.1 Microbacterium testaceum StLB037 Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium testaceum 48.7 Meiothermus ruber 21, DSM 1279 Deinococci Thermales Thermaceae Meiothermus Meiothermus ruber 48.1 Nocardia sp. 348MFTsu5.1 Actinobacteria Actinomycetales Nocardiaceae Nocardia Nocardia sp. 348MFTsu5.1 47.5 Leptothrix cholodnii SP-6 Betaproteobacteria Burkholderiales unclassified Leptothrix Leptothrix cholodnii 47.5 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 47.1 Renibacterium salmoninarum ATCC 33209 Actinobacteria Actinomycetales Micrococcaceae Renibacterium Renibacterium salmoninarum 46.4 Marinobacter manganoxydans MnI7-9 Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Marinobacter manganoxydans 46.2 Chelativorans sp. BNC1 Alphaproteobacteria Rhizobiales Phyllobacteriaceae Chelativorans Chelativorans sp. BNC1 Anaerophaga thermohalophila Fru22, 45.5 DSM 12881 Bacteroidia Bacteroidales Marinilabiliaceae Anaerophaga Anaerophaga thermohalophila 43.5 Starkeya novella DSM 506 Alphaproteobacteria Rhizobiales Xanthobacteraceae Starkeya Starkeya novella 41.7 Chelativorans sp. BNC1 Alphaproteobacteria Rhizobiales Phyllobacteriaceae Chelativorans Chelativorans sp. BNC1 39.3 Polaromonas sp. JS666 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas Polaromonas sp. JS666 38.7 Mycobacterium vanbaalenii PYR-1 Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium vanbaalenii 37.9 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium 192

nitrogenifigens

34.3 Thioalkalivibrio sp. ALE11 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Thioalkalivibrio Thioalkalivibrio sp. ALE11 32.3 Ulvibacter sp. SCB49 unclassified unclassified unclassified Ulvibacter Ulvibacter sp. SCB49 30.6 Methyloversatilis universalis EHg5 Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis Methyloversatilis universalis

193

Table A4-D. Genes encoding for todC1/bedC1 (benzene/toluene dioxygenase) detected in the OSPW metagenome, and their top BLAST hits

(to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species Thiocapsa marina 5811, DSM 91.9 5653 Gammaproteobacteria Chromatiales Chromatiaceae Thiocapsa Thiocapsa marina Thiocapsa marina 5811, DSM 79.8 5653 Gammaproteobacteria Chromatiales Chromatiaceae Thiocapsa Thiocapsa marina 71.1 Thauera aminoaromatica MZ1T Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera Thauera aminoaromatica 66.3 Nitrococcus mobilis Nb-231 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Nitrococcus Nitrococcus mobilis 61.1 Burkholderia sp. WSM4176 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. WSM4176 56.1 Cycloclasticus sp. P1 Gammaproteobacteria Thiotrichales Piscirickettsiaceae Cycloclasticus Cycloclasticus sp. P1 55.8 Pseudonocardia sp. P1 Actinobacteria Actinomycetales Pseudonocardiaceae Pseudonocardia Pseudonocardia sp. P1 Marinobacter manganoxydans 51.2 MnI7-9 Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Marinobacter manganoxydans Methylobacterium extorquens 50.2 DM4 Alphaproteobacteria Rhizobiales Methylobacteriaceae Methylobacterium Methylobacterium extorquens Cupriavidus metallidurans 47.7 CH34 Betaproteobacteria Burkholderiales Burkholderiaceae Cupriavidus Cupriavidus metallidurans Marinobacter manganoxydans 45.7 MnI7-9 Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Marinobacter manganoxydans Rudaea cellulosilytica DSM 45.1 22992 Gammaproteobacteria Lysobacterales Lysobacteraceae Rudaea Rudaea cellulosilytica Thermocrispum municipale DSM 43.5 44069 Actinobacteria Actinomycetales Pseudonocardiaceae Thermocrispum Thermocrispum municipale Parvibaculum lavamentivorans 42.5 DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans Granulicoccus phenolivorans 41.3 DSM 17626 Actinobacteria Actinomycetales Propionibacteriaceae Granulicoccus Granulicoccus phenolivorans 39.2 Achromobacter piechaudii HLE Betaproteobacteria Burkholderiales Alcaligenaceae Achromobacter Achromobacter piechaudii 35.2 Pseudomonas mendocina NK-01 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas mendocina Acinetobacter nosocomialis 35 Ab22222 Gammaproteobacteria Pseudomonadales Moraxellaceae Acinetobacter Acinetobacter nosocomialis Bradyrhizobium japonicum 33.6 USDA 122 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium japonicum

194

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

96.3 Burkholderia cepacia 2a plasmid pIJB1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia cepacia 94.9 Burkholderia cepacia 2a plasmid pIJB1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia cepacia 93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 91.7 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.8 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 90.6 Thauera aminoaromatica MZ1T Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera Thauera aminoaromatica 90.4 Pseudomonas stutzeri A1501 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 89.8 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 89.7 Sphingobium chlorophenolicum L-1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingobium Sphingobium chlorophenolicum 89.5 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 89.5 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 88.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 88 Meganema perideroedes DSM 15528 Alphaproteobacteria Rhizobiales Methylobacteriaceae Meganema Meganema perideroedes 87.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 86.2 Burkholderia mimosarum LMG 23256 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia mimosarum Novosphingobium aromaticivorans DSM 85.6 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 85.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.3 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.9 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 84.8 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y Novosphingobium aromaticivorans DSM 84.7 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 84.6 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii

195

84.5 Bradyrhizobium sp. EC3.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Ec3.3 84.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.3 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 83.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 83.7 Sphingobium chlorophenolicum L-1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingobium Sphingobium chlorophenolicum 82.6 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y Flavobacteria bacterium MS220-5C 82.3 (unscreened) Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium Flavobacterium sp. MS220-5C 81.9 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 81.5 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 81.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 81.4 Rhizobium etli CFN 42, DSM 11541 Alphaproteobacteria Rhizobiales Rhizobiaceae Rhizobium Rhizobium etli 81.4 Meganema perideroedes DSM 15528 Alphaproteobacteria Rhizobiales Methylobacteriaceae Meganema Meganema perideroedes 81.3 Bradyrhizobium sp. Cp5.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Cp5.3 Flavobacteria bacterium MS390-1F Flavobacteria bacterium MS390- 81.2 (unscreened) Flavobacteriia unclassified unclassified unclassified 1F 80.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 80.3 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 80 Streptomyces scabrisporus DSM 41855 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces scabrisporus 79.2 Tistrella mobilis KA081020-065 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Tistrella Tistrella mobilis Novosphingobium aromaticivorans DSM 79 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans Novosphingobium aromaticivorans DSM 79 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 77.9 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium nitrogenifigens Novosphingobium aromaticivorans DSM 77.3 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 76.9 Streptomyces sp. AA4 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces sp. AA4 76.7 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium nitrogenifigens 76.6 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 76.5 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 76 Amycolicicoccus subflavus DQS3-9A1 Actinobacteria Actinomycetales Mycobacteriaceae Amycolicicoccus Amycolicicoccus subflavus 75.8 Rhizobium sp. AP16 Alphaproteobacteria Rhizobiales Rhizobiaceae Rhizobium Rhizobium sp. AP16 Novosphingobium aromaticivorans DSM 75.7 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans

196

75.4 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 75.2 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 75.1 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 74.6 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium nitrogenifigens 74.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 73.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 73.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 73.2 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y Novosphingobium aromaticivorans DSM 73.2 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 72.9 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 72.9 Thiothrix nivea JP2, DSM 5205 Gammaproteobacteria Thiotrichales Thiotrichaceae Thiothrix Thiothrix nivea 72.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.5 Polymorphum gilvum SL003B-26A1 Alphaproteobacteria unclassified unclassified Polymorphum Polymorphum gilvum Saccharomonospora glauca K62, DSM 72.4 43769 Actinobacteria Actinomycetales Pseudonocardiaceae Saccharomonospora Saccharomonospora glauca Mucilaginibacter paludis TPT56, DSM 71.9 18603 Sphingobacteriia Sphingobacteriales Sphingobacteriaceae Mucilaginibacter Mucilaginibacter paludis 71.8 Humibacter albus DSM 18994 Actinobacteria Actinomycetales Microbacteriaceae Humibacter Humibacter albus 71.2 Methylobacterium nodulans ORS 2060 Alphaproteobacteria Rhizobiales Methylobacteriaceae Methylobacterium Methylobacterium nodulans 71.2 Promicromonospora sukumoe 327MFSha3.1 Actinobacteria Actinomycetales Promicromonosporaceae Promicromonospora Promicromonospora sukumoe 70.9 Burkholderia sp. JPY347 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. JPY347 70.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y Novosphingobium aromaticivorans DSM 70.3 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans Novosphingobium aromaticivorans DSM 70 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans Novosphingobium aromaticivorans DSM 69.5 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 69.5 Meganema perideroedes DSM 15528 Alphaproteobacteria Rhizobiales Methylobacteriaceae Meganema Meganema perideroedes 69.3 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y Novosphingobium aromaticivorans DSM 68.7 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 68.7 Acetobacter aceti ATCC 23746 Alphaproteobacteria Rhodospirillales Acetobacteraceae Acetobacter Acetobacter aceti 68.6 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 68.4 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis

197

68.1 Bradyrhizobium sp. WSM471 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. WSM471 Novosphingobium aromaticivorans DSM 67.5 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 67 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 66.4 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 66.3 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 66.2 Octadecabacter antarcticus 307 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Octadecabacter Octadecabacter antarcticus 66 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 65.5 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri Novosphingobium aromaticivorans DSM 65.2 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 65 Caulobacter crescentus OR37 Alphaproteobacteria Caulobacterales Caulobacteraceae Caulobacter Caulobacter vibrioides Novosphingobium aromaticivorans DSM 64.9 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 64.7 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 64.2 Microbacterium paraoxydans 77MFTsu3.2 Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium paraoxydans Novosphingobium aromaticivorans DSM 63.9 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 63.9 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 63.8 Pseudomonas fluorescens HK44 (Draft 1) Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas fluorescens 63.7 Bordetella avium 197N Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella avium Marine gamma proteobacterium sp. marine gamma proteobacterium 63.2 HTCC2080 Gammaproteobacteria unclassified unclassified unclassified HTCC2080 63.2 Polymorphum gilvum SL003B-26A1 Alphaproteobacteria unclassified unclassified Polymorphum Polymorphum gilvum 63 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 62.9 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 62.3 Aequorivita capsosiphonis DSM 23843 Flavobacteriia Flavobacteriales Flavobacteriaceae Aequorivita Aequorivita capsosiphonis 61.4 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 61.2 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 61.2 Bordetella avium 197N Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella avium 61 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 61 Pseudomonas sp. GM84 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM84 marine actinobacterium 60.8 marine actinobacterium PHSC20C1 Actinobacteria unclassified unclassified unclassified PHSC20C1 60.7 Actinomadura rifamycini DSM 43936 Actinobacteria Actinomycetales Thermomonosporaceae Actinomadura Actinomadura rifamycini

198

60.7 Pedobacter sp. BAL39 Sphingobacteriia Sphingobacteriales Sphingobacteriaceae Pedobacter Pedobacter sp. BAL39 60.4 Burkholderia nodosa DSM 21604 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia nodosa Nakamurella multipartita Y-104, DSM 60.4 44233 Actinobacteria Actinomycetales Nakamurellaceae Nakamurella Nakamurella multipartita 60.1 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y Flavobacteria bacterium MS390-1F Flavobacteria bacterium MS390- 58.8 (unscreened) Flavobacteriia unclassified unclassified unclassified 1F 58.4 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y Novosphingobium aromaticivorans DSM 58.2 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 58 Acidovorax delafieldii 2AN Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax Acidovorax delafieldii 57.8 Tistrella mobilis KA081020-065 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Tistrella Tistrella mobilis 57.7 Burkholderia nodosa DSM 21604 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia nodosa 57.5 Bradyrhizobium sp. Cp5.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Cp5.3 57.5 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 57.4 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 57.4 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 57.1 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 56.6 Frankia sp. EAN1pec Actinobacteria Actinomycetales Frankiaceae Frankia Frankia sp. EAN1pec 56.6 Meiothermus rufus DSM 22234 Deinococci Thermales Thermaceae Meiothermus Meiothermus rufus 56 Nocardia brasiliensis ATCC 700358 Actinobacteria Actinomycetales Nocardiaceae Nocardia Nocardia brasiliensis 55.8 Solimonas variicoloris DSM 15731 Gammaproteobacteria Xanthomonadales Sinobacteraceae Solimonas Solimonas variicoloris 55.8 Mycobacterium sp. JLS Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium sp. JLS 55.7 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 55.7 Caulobacter sp. AP07 Alphaproteobacteria Caulobacterales Caulobacteraceae Caulobacter Caulobacter sp. AP07 55.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 54.8 Hoeflea sp. 108 Alphaproteobacteria Rhizobiales Phyllobacteriaceae Hoeflea Hoeflea sp. 108 Nakamurella multipartita Y-104, DSM 54.7 44233 Actinobacteria Actinomycetales Nakamurellaceae Nakamurella Nakamurella multipartita 54.4 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 54.4 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 54.3 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 53.9 Amycolatopsis taiwanensis DSM 45107 Actinobacteria Actinomycetales Pseudonocardiaceae Amycolatopsis Amycolatopsis taiwanensis

199

53.8 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 52.6 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii Novosphingobium aromaticivorans DSM 52.2 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 51.9 Pseudomonas fluorescens HK44 (Draft 1) Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas fluorescens 51.8 Microbacterium sp. 11MF Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium sp. 11MF 50.5 Aequorivita capsosiphonis DSM 23843 Flavobacteriia Flavobacteriales Flavobacteriaceae Aequorivita Aequorivita capsosiphonis Novosphingobium aromaticivorans DSM 50.3 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans Novosphingobium aromaticivorans DSM 50 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 49.1 Dechloromonas aromatica RCB Betaproteobacteria Rhodocyclales Rhodocyclaceae Dechloromonas Dechloromonas aromatica Novosphingobium aromaticivorans DSM 49 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 48.8 Burkholderia nodosa DSM 21604 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia nodosa 48.7 Meiothermus ruber 21, DSM 1279 Deinococci Thermales Thermaceae Meiothermus Meiothermus ruber Novosphingobium aromaticivorans DSM 48.4 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans Novosphingobium aromaticivorans DSM 47.6 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 47.5 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 47.1 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 46.8 Cupriavidus necator N-1, ATCC 43291 Betaproteobacteria Burkholderiales Burkholderiaceae Cupriavidus Cupriavidus necator 46.7 Sphingobium chlorophenolicum L-1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingobium Sphingobium chlorophenolicum 46.4 Marinobacter manganoxydans MnI7-9 Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Marinobacter manganoxydans 45.7 Citreicella sp. SE45 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Citreicella Citreicella sp. SE45 Anaerophaga thermohalophila Fru22, DSM 45.5 12881 Bacteroidia Bacteroidales Marinilabiliaceae Anaerophaga Anaerophaga thermohalophila 45.2 Pseudomonas sp. GM84 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM84 44.6 Bradyrhizobium sp. EC3.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Ec3.3 44.5 Burkholderia ambifaria IOP40-10 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia ambifaria 43.8 Rhodococcus opacus B4 Actinobacteria Actinomycetales Nocardiaceae Rhodococcus Rhodococcus opacus 43.6 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 42.4 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 42.1 Pseudomonas sp. GM78 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM78 41.6 Sphingomonas sp. LH128 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas sp. LH128

200

40.8 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 40.8 Pseudomonas sp. GM49 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM49 40.2 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 39.3 Polaromonas sp. JS666 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas Polaromonas sp. JS666 Amycolatopsis thermoflava N1165, DSM 38.3 44574 Actinobacteria Actinomycetales Pseudonocardiaceae Amycolatopsis Amycolatopsis thermoflava 37.4 Pseudomonas fluorescens HK44 (Draft 1) Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas fluorescens 34.3 Thioalkalivibrio sp. ALE11 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Thioalkalivibrio Thioalkalivibrio sp. ALE11 32.3 Ulvibacter sp. SCB49 unclassified unclassified unclassified Ulvibacter Ulvibacter sp. SCB49 32 Ensifer meliloti GVPV12 Alphaproteobacteria Rhizobiales Rhizobiaceae Ensifer Ensifer meliloti

201

Table A4-F. Genes encoding for mhpF (acetaldehyde dehydrogenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

96.3 Burkholderia cepacia 2a plasmid pIJB1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia cepacia 94.9 Burkholderia cepacia 2a plasmid pIJB1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia cepacia 93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 91.7 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.8 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 90.6 Thauera aminoaromatica MZ1T Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera Thauera aminoaromatica 90.4 Pseudomonas stutzeri A1501 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 89.8 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 89.7 Sphingobium chlorophenolicum L-1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingobium Sphingobium chlorophenolicum 89.5 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 89.5 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 88.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 88 Meganema perideroedes DSM 15528 Alphaproteobacteria Rhizobiales Methylobacteriaceae Meganema Meganema perideroedes 87.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 86.2 Burkholderia mimosarum LMG 23256 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia mimosarum 85.6 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 85.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.3 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.9 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 84.8 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 84.7 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 84.6 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 84.5 Bradyrhizobium sp. EC3.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Ec3.3 84.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.3 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 202

83.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 83.7 Sphingobium chlorophenolicum L-1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingobium Sphingobium chlorophenolicum 82.6 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 82.3 Flavobacteria bacterium MS220-5C (unscreened) Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium Flavobacterium sp. MS220-5C 81.9 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 81.5 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 81.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 81.4 Rhizobium etli CFN 42, DSM 11541 Alphaproteobacteria Rhizobiales Rhizobiaceae Rhizobium Rhizobium etli 81.4 Meganema perideroedes DSM 15528 Alphaproteobacteria Rhizobiales Methylobacteriaceae Meganema Meganema perideroedes 81.3 Bradyrhizobium sp. Cp5.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Cp5.3 81.2 Flavobacteria bacterium MS390-1F (unscreened) Flavobacteriia unclassified unclassified unclassified Flavobacteria bacterium MS390-1F 80.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 80.3 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 80 Streptomyces scabrisporus DSM 41855 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces scabrisporus 79.2 Tistrella mobilis KA081020-065 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Tistrella Tistrella mobilis 79 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 79 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 77.9 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium nitrogenifigens 77.3 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 76.9 Streptomyces sp. AA4 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces sp. AA4 76.7 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium nitrogenifigens 76.6 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 76.5 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 76 Amycolicicoccus subflavus DQS3-9A1 Actinobacteria Actinomycetales Mycobacteriaceae Amycolicicoccus Amycolicicoccus subflavus 75.8 Rhizobium sp. AP16 Alphaproteobacteria Rhizobiales Rhizobiaceae Rhizobium Rhizobium sp. AP16 75.7 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 75.4 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 75.2 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 75.1 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 74.6 Novosphingobium nitrogenifigens Y88 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium nitrogenifigens

203

74.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 73.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 73.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 73.2 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 73.2 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 72.9 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 72.9 Thiothrix nivea JP2, DSM 5205 Gammaproteobacteria Thiotrichales Thiotrichaceae Thiothrix Thiothrix nivea 72.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.5 Polymorphum gilvum SL003B-26A1 Alphaproteobacteria unclassified unclassified Polymorphum Polymorphum gilvum 72.4 Saccharomonospora glauca K62, DSM 43769 Actinobacteria Actinomycetales Pseudonocardiaceae Saccharomonospora Saccharomonospora glauca 71.9 Mucilaginibacter paludis TPT56, DSM 18603 Sphingobacteriia Sphingobacteriales Sphingobacteriaceae Mucilaginibacter Mucilaginibacter paludis 71.8 Humibacter albus DSM 18994 Actinobacteria Actinomycetales Microbacteriaceae Humibacter Humibacter albus 71.2 Methylobacterium nodulans ORS 2060 Alphaproteobacteria Rhizobiales Methylobacteriaceae Methylobacterium Methylobacterium nodulans 71.2 Promicromonospora sukumoe 327MFSha3.1 Actinobacteria Actinomycetales Promicromonosporaceae Promicromonospora Promicromonospora sukumoe 70.9 Burkholderia sp. JPY347 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. JPY347 70.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 70.3 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 70 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 69.5 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 69.5 Meganema perideroedes DSM 15528 Alphaproteobacteria Rhizobiales Methylobacteriaceae Meganema Meganema perideroedes 69.3 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 68.7 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 68.7 Acetobacter aceti ATCC 23746 Alphaproteobacteria Rhodospirillales Acetobacteraceae Acetobacter Acetobacter aceti 68.6 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 68.4 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 68.1 Bradyrhizobium sp. WSM471 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. WSM471 67.5 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 67 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 66.4 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 66.3 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis

204

66.2 Octadecabacter antarcticus 307 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Octadecabacter Octadecabacter antarcticus 66 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 65.5 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 65.2 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 65 Caulobacter crescentus OR37 Alphaproteobacteria Caulobacterales Caulobacteraceae Caulobacter Caulobacter vibrioides 64.9 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 64.7 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 64.2 Microbacterium paraoxydans 77MFTsu3.2 Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium paraoxydans 63.9 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 63.9 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 63.8 Pseudomonas fluorescens HK44 (Draft 1) Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas fluorescens 63.7 Bordetella avium 197N Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella avium 63.2 Marine gamma proteobacterium sp. HTCC2080 Gammaproteobacteria unclassified unclassified unclassified marine gamma proteobacterium HTCC2080 63.2 Polymorphum gilvum SL003B-26A1 Alphaproteobacteria unclassified unclassified Polymorphum Polymorphum gilvum 63 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 62.9 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 62.3 Aequorivita capsosiphonis DSM 23843 Flavobacteriia Flavobacteriales Flavobacteriaceae Aequorivita Aequorivita capsosiphonis 61.4 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 61.2 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 61.2 Bordetella avium 197N Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella avium 61 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 61 Pseudomonas sp. GM84 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM84 60.8 marine actinobacterium PHSC20C1 Actinobacteria unclassified unclassified unclassified marine actinobacterium PHSC20C1 60.7 Actinomadura rifamycini DSM 43936 Actinobacteria Actinomycetales Thermomonosporaceae Actinomadura Actinomadura rifamycini 60.4 Burkholderia nodosa DSM 21604 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia nodosa 60.4 Nakamurella multipartita Y-104, DSM 44233 Actinobacteria Actinomycetales Nakamurellaceae Nakamurella Nakamurella multipartita 60.1 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 58.8 Flavobacteria bacterium MS390-1F (unscreened) Flavobacteriia unclassified unclassified unclassified Flavobacteria bacterium MS390-1F 58.4 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 58.2 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans

205

58 Acidovorax delafieldii 2AN Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax Acidovorax delafieldii 57.8 Tistrella mobilis KA081020-065 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Tistrella Tistrella mobilis 57.7 Burkholderia nodosa DSM 21604 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia nodosa 57.5 Bradyrhizobium sp. Cp5.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Cp5.3 57.5 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 57.4 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 57.4 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 57.1 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 56.6 Frankia sp. EAN1pec Actinobacteria Actinomycetales Frankiaceae Frankia Frankia sp. EAN1pec 56.6 Meiothermus rufus DSM 22234 Deinococci Thermales Thermaceae Meiothermus Meiothermus rufus 56 Nocardia brasiliensis ATCC 700358 Actinobacteria Actinomycetales Nocardiaceae Nocardia Nocardia brasiliensis 55.8 Solimonas variicoloris DSM 15731 Gammaproteobacteria Xanthomonadales Sinobacteraceae Solimonas Solimonas variicoloris 55.8 Mycobacterium sp. JLS Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium sp. JLS 55.7 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 55.7 Caulobacter sp. AP07 Alphaproteobacteria Caulobacterales Caulobacteraceae Caulobacter Caulobacter sp. AP07 55.5 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 54.8 Hoeflea sp. 108 Alphaproteobacteria Rhizobiales Phyllobacteriaceae Hoeflea Hoeflea sp. 108 54.7 Nakamurella multipartita Y-104, DSM 44233 Actinobacteria Actinomycetales Nakamurellaceae Nakamurella Nakamurella multipartita 54.4 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 54.4 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 54.3 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 53.9 Amycolatopsis taiwanensis DSM 45107 Actinobacteria Actinomycetales Pseudonocardiaceae Amycolatopsis Amycolatopsis taiwanensis 53.8 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 52.6 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 52.2 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 51.9 Pseudomonas fluorescens HK44 (Draft 1) Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas fluorescens 51.8 Microbacterium sp. 11MF Actinobacteria Actinomycetales Microbacteriaceae Microbacterium Microbacterium sp. 11MF 50.3 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 50 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 49.1 Dechloromonas aromatica RCB Betaproteobacteria Rhodocyclales Rhodocyclaceae Dechloromonas Dechloromonas aromatica

206

49 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 48.8 Burkholderia nodosa DSM 21604 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia nodosa 48.7 Meiothermus ruber 21, DSM 1279 Deinococci Thermales Thermaceae Meiothermus Meiothermus ruber 48.4 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 47.6 Novosphingobium aromaticivorans DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium aromaticivorans 47.5 Kyrpidia tusciae T2, DSM 2912 Bacilli Bacillales Alicyclobacillaceae Kyrpidia Kyrpidia tusciae 47.1 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 46.8 Cupriavidus necator N-1, ATCC 43291 Betaproteobacteria Burkholderiales Burkholderiaceae Cupriavidus Cupriavidus necator 46.7 Sphingobium chlorophenolicum L-1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingobium Sphingobium chlorophenolicum 46.4 Marinobacter manganoxydans MnI7-9 Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter Marinobacter manganoxydans 45.7 Citreicella sp. SE45 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Citreicella Citreicella sp. SE45 45.5 Anaerophaga thermohalophila Fru22, DSM 12881 Bacteroidia Bacteroidales Marinilabiliaceae Anaerophaga Anaerophaga thermohalophila 45.2 Pseudomonas sp. GM84 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM84 44.6 Bradyrhizobium sp. EC3.3 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. Ec3.3 44.5 Burkholderia ambifaria IOP40-10 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia ambifaria 43.8 Rhodococcus opacus B4 Actinobacteria Actinomycetales Nocardiaceae Rhodococcus Rhodococcus opacus 43.6 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 42.4 Actinoplanes globisporus DSM 43857 Actinobacteria Actinomycetales Micromonosporaceae Actinoplanes Actinoplanes globisporus 42.1 Pseudomonas sp. GM78 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM78 41.6 Sphingomonas sp. LH128 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas sp. LH128 40.8 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 40.8 Pseudomonas sp. GM49 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. GM49 40.2 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 39.3 Polaromonas sp. JS666 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas Polaromonas sp. JS666 38.3 Amycolatopsis thermoflava N1165, DSM 44574 Actinobacteria Actinomycetales Pseudonocardiaceae Amycolatopsis Amycolatopsis thermoflava 37.4 Pseudomonas fluorescens HK44 (Draft 1) Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas fluorescens 34.3 Thioalkalivibrio sp. ALE11 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Thioalkalivibrio Thioalkalivibrio sp. ALE11 32.3 Ulvibacter sp. SCB49 unclassified unclassified unclassified Ulvibacter Ulvibacter sp. SCB49 32 Ensifer meliloti GVPV12 Alphaproteobacteria Rhizobiales Rhizobiaceae Ensifer Ensifer meliloti

207

Table A4-G. Genes encoding for dmpC, xylG (aminomuconate-semialdehyde/2-hydroxymuconate-6-semialdehyde dehydrogenase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species (%)

98 Pseudomonas sp. CT14 plasmid pCT14 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas sp. 96.9 Pseudomonas sp. CT14 plasmid pCT14 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas sp. 95.5 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 90.8 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 90 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 88.1 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis Mesoflavibacter 87.7 Mesoflavibacter zeaxanthinifaciens DSM 18436 Flavobacteriia Flavobacteriales Flavobacteriaceae Mesoflavibacter zeaxanthinifaciens 86.1 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 81.5 Flavobacterium tegetincola DSM 22377 Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium Flavobacterium tegetincola 81.4 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 80.8 Thauera aminoaromatica MZ1T Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera Thauera aminoaromatica 80.6 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 79.6 Leadbetterella byssophila 4M15, DSM 17132 Cytophagia Cytophagales Cytophagaceae Leadbetterella Leadbetterella byssophila 78.8 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 75.8 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 75.4 Acidovorax sp. JS42 Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax Acidovorax sp. JS42 75.3 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 75 Rhodanobacter sp. 2APBS3 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Rhodanobacter Rhodanobacter sp. 2APBS3 74.1 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 73.8 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 71.7 Flavobacteria bacterium BAL38 Flavobacteriia Flavobacteriales unclassified unclassified Flavobacteria bacterium BAL38 71.2 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 70.6 Marivirga tractuosa H-43, DSM 4126 Cytophagia Cytophagales Flammeovirgaceae Marivirga Marivirga tractuosa 70.1 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis

208

70.1 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 69.9 Thiothrix nivea JP2, DSM 5205 Gammaproteobacteria Thiotrichales Thiotrichaceae Thiothrix Thiothrix nivea Pseudoxanthomon 69.7 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae as Pseudoxanthomonas spadix Flavobacteriaceae bacterium 69.7 Flavobacteriaceae bacterium S85 Flavobacteriia Flavobacteriales Flavobacteriaceae unclassified S85 69.5 Luteimonas mephitis DSM 12574 Gammaproteobacteria Lysobacterales Lysobacteraceae Luteimonas Luteimonas mephitis 68.8 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 68.2 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 68.1 Luteimonas mephitis DSM 12574 Gammaproteobacteria Lysobacterales Lysobacteraceae Luteimonas Luteimonas mephitis Oscillatoriales cyanobacterium 68 Oscillatoriales sp. JSC-12 unclassified Oscillatoriales unclassified unclassified JSC-12 66.9 Rudaea cellulosilytica DSM 22992 Gammaproteobacteria Lysobacterales Lysobacteraceae Rudaea Rudaea cellulosilytica 66.7 Microscilla marina ATCC 23134 Cytophagia Cytophagales Cytophagaceae Microscilla Microscilla marina 66.3 Ochrobactrum anthropi ATCC 49188 Alphaproteobacteria Rhizobiales Brucellaceae Ochrobactrum Ochrobactrum anthropi 65.4 Bizionia argentinensis JUB59, DSM 19628 Flavobacteriia Flavobacteriales Flavobacteriaceae Bizionia Bizionia argentinensis 61.2 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 60.4 Aequorivita sublithincola QSSC9-3, DSM 14238 Flavobacteriia Flavobacteriales Flavobacteriaceae Aequorivita Aequorivita sublithincola 59.5 Niveispirillum irakense DSM 11586 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Niveispirillum Niveispirillum irakense 58.9 Massilia alkalitolerans DSM 17462 Betaproteobacteria Burkholderiales Oxalobacteraceae Massilia Massilia alkalitolerans 58.5 Syntrophus aciditrophicus SB Deltaproteobacteria Syntrophobacterales Syntrophaceae Syntrophus Syntrophus aciditrophicus 58.5 Flavobacterium frigidarium DSM 17623 Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium Flavobacterium frigidarium 58.4 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 57.8 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 56.8 Marivirga tractuosa H-43, DSM 4126 Cytophagia Cytophagales Flammeovirgaceae Marivirga Marivirga tractuosa Emticicia oligotrophica GPTSA100-15, DSM 56.2 17448 Cytophagia Cytophagales Cytophagaceae Emticicia Emticicia oligotrophica 55.7 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 55.7 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 55.4 Aquimarina latercula DSM 2041 Flavobacteriia Flavobacteriales Flavobacteriaceae Aquimarina Aquimarina latercula 53.7 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 53.2 Owenweeksia hongkongensis DSM 17368 Flavobacteriia Flavobacteriales Cryomorphaceae Owenweeksia Owenweeksia hongkongensis 53.1 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis

209

53 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 52.9 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 51.8 Xanthomonas vesicatoria Maraite, ATCC 35937 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Xanthomonas Xanthomonas vesicatoria 51.8 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 48.6 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 46.9 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 44.7 Niveispirillum irakense DSM 11586 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Niveispirillum Niveispirillum irakense 43.5 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 41.7 Reinekea blandensis MED297 Gammaproteobacteria unclassified unclassified Reinekea Reinekea blandensis 41.5 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 39.6 Methylobacterium radiotolerans JCM 2831 Alphaproteobacteria Rhizobiales Methylobacteriaceae Methylobacterium Methylobacterium radiotolerans 37.9 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis Mesoflavibacter 36.8 Mesoflavibacter zeaxanthinifaciens DSM 18436 Flavobacteriia Flavobacteriales Flavobacteriaceae Mesoflavibacter zeaxanthinifaciens 34.5 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis 30.8 Fluviicola taffensis RW262, DSM 16823 Flavobacteriia Flavobacteriales Cryomorphaceae Fluviicola Fluviicola taffensis Flavobacteriales bacterium 30.1 Flavobacteriales sp. ALC-1 Flavobacteriia Flavobacteriales unclassified unclassified ALC-1

210

Table A4-H. Genes encoding for dmpD/xylF (hydroxymuconate-semialdehyde hydrolase) detected in the OSPW metagenome, and their top

BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species (50%)

95 Polaromonas naphthalenivorans CJ2 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas Polaromonas naphthalenivorans 92.1 Pseudomonas sp. CT14 plasmid pCT14 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas sp. 92.1 Pseudomonas sp. CT14 plasmid pCT14 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas sp. 85.4 Cupriavidus metallidurans CH34 Betaproteobacteria Burkholderiales Burkholderiaceae Cupriavidus Cupriavidus metallidurans 83.5 Pseudoxanthomonas spadix BD-a59 Gammaproteobacteria Xanthomonadales Xanthomonadaceae Pseudoxanthomonas Pseudoxanthomonas spadix 80.4 Pseudomonas sp. CT14 plasmid pCT14 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas sp. 77.7 Comamonas testosteroni S44 Betaproteobacteria Burkholderiales Comamonadaceae Comamonas Comamonas testosteroni 76.6 Comamonas testosteroni S44 Betaproteobacteria Burkholderiales Comamonadaceae Comamonas Comamonas testosteroni 75.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus Pseudomonas putida MT53 plasmid 74.6 pWW53 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas putida 73.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 71.2 Thiothrix nivea JP2, DSM 5205 Gammaproteobacteria Thiotrichales Thiotrichaceae Thiothrix Thiothrix nivea 69.9 Burkholderia vietnamiensis G4 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia vietnamiensis 60.4 Herbaspirillum sp. GW103 Betaproteobacteria Burkholderiales Oxalobacteraceae Herbaspirillum Herbaspirillum sp. GW103 59.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 Intrasporangium calvum 7KIP, DSM 58.4 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum Thermomonosporace 52.2 Actinomadura rifamycini DSM 43936 Actinobacteria Actinomycetales ae Actinomadura Actinomadura rifamycini Intrasporangium calvum 7KIP, DSM 50 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum

211

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

40.4 Liberibacter crescens BT-1 Alphaproteobacteria Rhizobiales Rhizobiaceae Liberibacter Liberibacter crescens

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

95.2 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 98.6 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 97.7 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 95.2 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 88 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 75.4 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 74.1 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 85.8 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 70.3 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 71.6 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 72.6 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 73.6 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 80.3 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 88.5 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans

212

87.5 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 81.8 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 74.5 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 82.6 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 85.5 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 87.3 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 84.3 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 64.3 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 69.4 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 71.5 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 40.6 Parvibaculum lavamentivorans DS-1 Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum Parvibaculum lavamentivorans 46.6 Burkholderia sp. WSM2230 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. WSM2230

Table A4-K. Genes encoding for nahD (hydroxychromene-2-carboxylate isomerase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

86.2 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.6 Porphyrobacter cryptus DSM 12079 Alphaproteobacteria Sphingomonadales Erythrobacteraceae Porphyrobacter Porphyrobacter cryptus 80.6 Novosphingobium sp. PP1Y Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium sp. PP1Y 78.2 Sphingopyxis alaskensis RB2256 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingopyxis Sphingopyxis alaskensis 77.9 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 74.7 Azoarcus sp. BH72 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus sp. BH72 70.5 Beijerinckia indica indica ATCC 9039 Alphaproteobacteria Rhizobiales Beijerinckiaceae Beijerinckia Beijerinckia indica 70.1 Sphingopyxis alaskensis RB2256 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingopyxis Sphingopyxis alaskensis 69.4 Sphingopyxis alaskensis RB2256 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingopyxis Sphingopyxis alaskensis

213

68.6 Thiomonas arsenitoxydans 3As Betaproteobacteria Burkholderiales unclassified Thiomonas Thiomonas arsenitoxydans Novosphingobium acidiphilum DSM 68.5 19966 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium Novosphingobium acidiphilum 67 Sphingopyxis alaskensis RB2256 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingopyxis Sphingopyxis alaskensis 66.7 Porphyrobacter cryptus DSM 12079 Alphaproteobacteria Sphingomonadales Erythrobacteraceae Porphyrobacter Porphyrobacter cryptus Novosphingobium aromaticivorans Novosphingobium 66.1 DSM 12444 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium aromaticivorans 64.8 Sphingomonas wittichii RW1 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingomonas Sphingomonas wittichii 63.4 Sphingopyxis alaskensis RB2256 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingopyxis Sphingopyxis alaskensis 61.9 Alcanivorax dieselolei B5 Gammaproteobacteria Oceanospirillales Alcanivoracaceae Alcanivorax Alcanivorax dieselolei Uliginosibacterium gangwonense DSM Uliginosibacterium 58.9 18521 Betaproteobacteria Rhodocyclales Rhodocyclaceae Uliginosibacterium gangwonense 58.6 Sphingopyxis alaskensis RB2256 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingopyxis Sphingopyxis alaskensis 57.3 Dethiobacter alkaliphilus AHT1 Clostridia Clostridiales Syntrophomonadaceae Dethiobacter Dethiobacter alkaliphilus 56.3 Sphingopyxis alaskensis RB2256 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingopyxis Sphingopyxis alaskensis 51.9 Cohnella thermotolerans DSM 17683 Bacilli Bacillales Paenibacillaceae Cohnella Cohnella thermotolerans 51.3 Nevskia ramosa DSM 11499 Gammaproteobacteria Xanthomonadales Nevskiaceae Nevskia Nevskia ramosa 47.9 Sphingobium sp. AP49 Alphaproteobacteria Sphingomonadales Sphingomonadaceae Sphingobium Sphingobium sp. AP49 44.8 Chitiniphilus shinanonensis DSM 23277 Betaproteobacteria Neisseriales Neisseriaceae Chitiniphilus Chitiniphilus shinanonensis

214

Table A4 L Genes encoding for nahE (trans-o-hydroxybenzylidenepyruvate hydratase-aldolase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

74.7 Azoarcus sp. BH72 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus sp. BH72 70.5 Beijerinckia indica indica ATCC 9039 Alphaproteobacteria Rhizobiales Beijerinckiaceae Beijerinckia Beijerinckia indica 77.9 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1 68.6 Thiomonas arsenitoxydans 3As Betaproteobacteria Burkholderiales unclassified Thiomonas Thiomonas arsenitoxydans 86.2 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus Uliginosibacterium gangwonense DSM 58.9 18521 Betaproteobacteria Rhodocyclales Rhodocyclaceae Uliginosibacterium Uliginosibacterium gangwonense 44.8 Chitiniphilus shinanonensis DSM 23277 Betaproteobacteria Neisseriales Neisseriaceae Chitiniphilus Chitiniphilus shinanonensis 51.3 Nevskia ramosa DSM 11499 Gammaproteobacteria Xanthomonadales Nevskiaceae Nevskia Nevskia ramosa 57.3 Dethiobacter alkaliphilus AHT1 Clostridia Clostridiales Syntrophomonadaceae Dethiobacter Dethiobacter alkaliphilus 72.1 Ralstonia solanacearum Po82 Betaproteobacteria Burkholderiales Burkholderiaceae Ralstonia Ralstonia solanacearum 64.2 Cupriavidus basilensis OR16 Betaproteobacteria Burkholderiales Burkholderiaceae Cupriavidus Cupriavidus basilensis 52.5 Burkholderia sp. Ch1-1 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. Ch1-1

95.2 Delftia sp. CS3-4 Betaproteobacteria Burkholderiales Comamonadaceae Delftia Delftia sp. CS3-4 97.3 Delftia sp. CS3-4 Betaproteobacteria Burkholderiales Comamonadaceae Delftia Delftia sp. CS3-4 43.1 Pseudogulbenkiania sp. NH8B Betaproteobacteria Neisseriales Neisseriaceae Pseudogulbenkiania Pseudogulbenkiania sp. NH8B

215

Homolog Homolog Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Species Family Genus

83.2 Leptothrix cholodnii SP-6 Betaproteobacteria Burkholderiales Unclassified Leptothrix Leptothrix cholodnii 82.7 Leptothrix cholodnii SP-6 Betaproteobacteria Burkholderiales Unclassified Leptothrix Leptothrix cholodnii

Table A4-N. Genes encoding for sal-hyd (salicylate hydroxylase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

92.9 Phaeobacter gallaeciensis 2.10 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Phaeobacter Phaeobacter gallaeciensis 90.7 Polaromonas sp. JS666 Betaproteobacteria Burkholderiales Comamonadaceae Polaromonas Polaromonas sp. JS666 87.9 Leptothrix cholodnii SP-6 Betaproteobacteria Burkholderiales unclassified Leptothrix Leptothrix cholodnii 85.1 Rhodoferax ferrireducens T118 Betaproteobacteria Burkholderiales Comamonadaceae Rhodoferax Rhodoferax ferrireducens 84.3 Roseovarius sp. 217 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Roseovarius Roseovarius sp. 217 83.9 Actinopolyspora halophila DSM 43834 Actinobacteria (class) Actinomycetales Actinopolysporaceae Actinopolyspora Actinopolyspora halophila 83.1 Roseovarius sp. 217 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Roseovarius Roseovarius sp. 217 81 Streptomyces griseoflavus Tu4000 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces griseoflavus 80.2 Saccharomonospora azurea SZMC 14600 Actinobacteria Actinomycetales Pseudonocardiaceae Saccharomonospora Saccharomonospora azurea 79.6 Nonomuraea coxensis DSM 45129 Actinobacteria Actinomycetales Streptosporangiaceae Nonomuraea Nonomuraea coxensis Novispirillum itersonii itersonii ATCC 78.1 12639 Alphaproteobacteria Rhodospirillales Rhodospirillaceae Novispirillum Novispirillum itersonii 77.6 Roseovarius sp. 217 Alphaproteobacteria Rhodobacterales Rhodobacteraceae Roseovarius Roseovarius sp. 217 76.9 Mycobacterium smegmatis JS623 Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium smegmatis

216

Nitratireductor aquibiodomus RA22 75 (Draft1) Alphaproteobacteria Rhizobiales Phyllobacteriaceae Nitratireductor Nitratireductor aquibiodomus 74.8 Rhodoferax ferrireducens T118 Betaproteobacteria Burkholderiales Comamonadaceae Rhodoferax Rhodoferax ferrireducens 74.6 Bradyrhizobium sp. Th.b2 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium sp. th.b2 74.5 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 74.4 Amycolatopsis balhimycina FH 1894 Actinobacteria Pseudonocardiales Pseudonocardiaceae Amycolatopsis Amycolatopsis balhimycina Arthrobacter sp. 74.1 Arthrobacter sp. 131MFCol6.1 Actinobacteria Actinomycetales Micrococcaceae Arthrobacter 131MFCol6.1 73.8 Saccharomonospora cyanea NA-134 Actinobacteria Actinomycetales Pseudonocardiaceae Saccharomonospora Saccharomonospora cyanea 73 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum 72.3 Mycobacterium sp. JLS Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium sp. JLS Amycolatopsis halophila YIM 93223, DSM 72.2 45216 Actinobacteria Actinomycetales Pseudonocardiaceae Amycolatopsis Amycolatopsis halophila 70.9 Saccharomonospora azurea SZMC 14600 Actinobacteria Actinomycetales Pseudonocardiaceae Saccharomonospora Saccharomonospora azurea 70.6 Bordetella bronchiseptica RB50 Betaproteobacteria Burkholderiales Alcaligenaceae Bordetella Bordetella bronchiseptica 70.6 Bradyrhizobium elkanii WSM1741 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium Bradyrhizobium elkanii Comamonadaceae bacterium 69.2 Comamonadaceae bacterium URHA0028 Betaproteobacteria Burkholderiales Comamonadaceae unclassified URHA0028 69.2 Curvibacter lanceolatus ATCC 14669 Betaproteobacteria Burkholderiales Comamonadaceae Curvibacter Curvibacter lanceolatus Bradyrhizobium sp. CCGE- 69 Bradyrhizobium sp. CCGE-LA001 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium LA001 Pseudonocardia asaccharolytica DSM Pseudonocardia 65.9 44247 Actinobacteria Actinomycetales Pseudonocardiaceae Pseudonocardia asaccharolytica 65 Streptomyces ghanaensis ATCC 14672 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces ghanaensis 64.5 Rhodopseudomonas palustris BisB5 Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Rhodopseudomonas Rhodopseudomonas palustris 61.6 Gordonia sihwensis NBRC 108236 Actinobacteria Actinomycetales Gordoniaceae Gordonia Gordonia sihwensis 60.4 Mycobacterium sp. JLS Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium sp. JLS 60 Burkholderia sp. CCGE1001 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia sp. CCGE1001 Pseudonocardia asaccharolytica DSM Pseudonocardia 59.6 44247 Actinobacteria Actinomycetales Pseudonocardiaceae Pseudonocardia asaccharolytica 58.9 Granulicella mallensis MP5ACTX8 Acidobacteriia Acidobacteriales Acidobacteriaceae Granulicella Granulicella mallensis Pseudonocardia asaccharolytica DSM Pseudonocardia 58.3 44247 Actinobacteria Actinomycetales Pseudonocardiaceae Pseudonocardia asaccharolytica 57.9 Saccharomonospora azurea SZMC 14600 Actinobacteria Actinomycetales Pseudonocardiaceae Saccharomonospora Saccharomonospora azurea 57.7 Rhizobium tropici CIAT899 Alphaproteobacteria Rhizobiales Rhizobiaceae Rhizobium Rhizobium tropici 57.7 Intrasporangium calvum 7KIP, DSM 43043 Actinobacteria Actinomycetales Intrasporangiaceae Intrasporangium Intrasporangium calvum

217

57.2 Streptomyces ghanaensis ATCC 14672 Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces ghanaensis 55.8 Streptomyces sp. SA3_actG Actinobacteria Actinomycetales Streptomycetaceae Streptomyces Streptomyces sp. SA3_actG 54.4 Mycobacterium sp. MCS Actinobacteria Actinomycetales Mycobacteriaceae Mycobacterium Mycobacterium sp. MCS 54.1 Roseovarius nubinhibens ISM Alphaproteobacteria Rhodobacterales Rhodobacteraceae Roseovarius Roseovarius nubinhibens 53.2 Burkholderia oklahomensis C6786 Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia Burkholderia oklahomensis 50.9 Alcaligenes faecalis phenolicus DSM 16503 Betaproteobacteria Burkholderiales Alcaligenaceae Alcaligenes Alcaligenes faecalis 49.8 Frankia sp. EUN1f Actinobacteria Actinomycetales Frankiaceae Frankia Frankia sp. EUN1f 48.1 Gordonia sihwensis NBRC 108236 Actinobacteria Actinomycetales Gordoniaceae Gordonia Gordonia sihwensis 47.3 Gordonia sihwensis NBRC 108236 Actinobacteria Actinomycetales Gordoniaceae Gordonia Gordonia sihwensis 45.6 Methylobacterium extorquens DSM 13060 Alphaproteobacteria Rhizobiales Methylobacteriaceae Methylobacterium Methylobacterium extorquens 45.2 Blastococcus saxobsidens DD2 Actinobacteria Actinomycetales Geodermatophilaceae Blastococcus Blastococcus saxobsidens 44.6 Methylopila sp. M107 Alphaproteobacteria Rhizobiales Methylocystaceae Methylopila Methylopila sp. M107 39.5 Gordonia sihwensis NBRC 108236 Actinobacteria Actinomycetales Gordoniaceae Gordonia Gordonia sihwensis Amycolatopsis thermoflava N1165, DSM 36.5 44574 Actinobacteria Actinomycetales Pseudonocardiaceae Amycolatopsis Amycolatopsis thermoflava Bradyrhizobiaceae bacterium 36.1 Bradyrhizobiaceae bacterium SG-6C Alphaproteobacteria Rhizobiales Bradyrhizobiaceae unclassified SG-6C

218

Table A4-P. Genes encoding for dmpK/poxA (phenol hydroxylase) detected in the OSPW metagenome, and their top BLAST hits (to a cultured organism). The minimum scaffold length was 2500 nucleotides.

Hits (%) Homolog Genome Homolog Class Homolog Order Homolog Family Homolog Genus Homolog Species

93.1 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 91.7 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 90.3 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 89.5 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 88.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 87.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.4 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 85.3 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 84.4 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 83.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1

81.9 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 81.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 80.9 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 75.4 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 73.6 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 72.9 Azoarcus toluclasticus ATCC 700605 Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus Azoarcus toluclasticus 72.5 Pseudomonas sp. Chol1 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas sp. Chol1 68.4 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 67 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 66.3 Halomonas zhanjiangensis DSM 21076 Gammaproteobacteria Oceanospirillales Halomonadaceae Halomonas Halomonas zhanjiangensis 65.5 Pseudomonas stutzeri KOS6 Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas Pseudomonas stutzeri 34.3 Thioalkalivibrio sp. ALE11 Gammaproteobacteria Chromatiales Ectothiorhodospiraceae Thioalkalivibrio Thioalkalivibrio sp. ALE11

219

Figure B1. Representative CO2 production time courses of 2 replicate samples used for labelled substrates SIP of WIP-OSPW sampled in August, 2011. Bars represent SD.

220

Table B1 A-G. The top 10 OTUs of each sample site based on percent (%) of total reads in 16S rRNA gene sequencing analysis. OTUs were clustered based on their closest related species. Table B1-A. Reads from DNA in WIP-OSPW sampled in August 2011.

221

Table B1-B. Reads from DNA of the heavy SIP fraction (fraction 5) of OSPW sampled in August, 2011.

Rank % Reads Phylum Class Order Family Genus % Identity

1 22.27 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Devosia 96 2 12.64 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylomonas 96 3 9.03 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera 99 4 8.53 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas 100 5 8.29 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylogaea 94 6 6.79 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 87 7 4.95 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylomicrobium 99 8 4.52 Proteobacteria Alphaproteobacteria Magnetococcales Magnetococcaceae Magnetococcus 90 9 4.25 Proteobacteria Alphaproteobacteria Rhodospirillales Acetobacteraceae Acidiphilium 99 10 4.21 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 100 11 3.28 Euryarchaeota Methanomicrobia Methanomicrobiales Methanoregulaceae Methanoregula 99 12 2.98 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 86 13 2.88 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Hydrogenophaga 99 14 2.47 Proteobacteria Gammaproteobacteria Xanthomonadales Algiphilaceae Algiphilus 94 15 2.34 Proteobacteria Alphaproteobacteria Sphingomonadales Erythrobacteraceae Altererythrobacter 99 16 2.21 Euryarchaeota Methanomicrobia Magnetococcales 99 17 2.17 Euryarchaeota Methanomicrobia Methanosarcinales Methanosaetaceae Methanosaeta 100 18 1.97 Proteobacteria Alphaproteobacteria Sphingomonadales Erythrobacteraceae Porphyrobacter 100 19 1.81 Proteobacteria Gammaproteobacteria Chromatiales Chromatiaceae Thiococcus 95 20 1.81 Proteobacteria Alphaproteobacteria Rhodocyclales Rhodocyclaceae Azospirillum 93 21 1.81 Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Lutibacter 98 22 1.74 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 89 23 1.57 Bacteroidetes Flavobacteria Flavobacteriales Flavobacteriaceae Flavobacterium 99 24 1.51 Proteobacteria Gammaproteobacteria Magnetococcales Methylococcaceae Methylomicrobium 97 25 1.40 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Rhodoferax 98

222

Table B1-C. Reads from DNA of the heavy SIP fraction (fractions 3 and 4) of OSPW sampled in August, 2011, amended with labelled methanol for 7 days.

Rank % Reads Phylum Class Order Family Genus % Identity

223

Table B1-D. Heavy SIP Fractions (fractions 4 and 5) of OSPW sampled in August, 2011, amended with labelled sodium acetate for 5 days.

Rank % Reads Phylum Class Order Family Genus % Identity

1 20.73 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Devosia 96 2 15.27 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Xanthobacter 99 3 9.11 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas 100 4 7.40 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylomarinum 96 5 7.17 Proteobacteria Alphaproteobacteria Sphingomonadales Sphingomonadaceae Novosphingobium 98 6 6.31 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylogaea 94 7 5.71 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera 99 8 4.72 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 93 9 2.98 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Azoarcus 100 3.00 Thaumarchaeota Nitrosopumilales Nitrosopumilaceae Candidatus 97 10 Nitrosoarchaeum 11 2.73 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Phenylobacterium 98 12 2.42 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylomicrobium 99 13 2.50 Proteobacteria Gammaproteobacteria Pseudomonadales Moraxellaceae Enhydrobacter 99 14 2.50 Proteobacteria Alphaproteobacteria Rhodospirillales Acetobacteraceae Acidiphilum 99 15 2.30 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Hydrogenophaga 99 16 2.17 Proteobacteria Gammaproteobacteria Pseudomonadales Pseudomonadaceae Pseudomonas 96 17 1.94 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Aquisphaera 90 18 1.64 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 100 19 1.44 Proteobacteria Alphaproteobacteria Sphingomonadales Erythrobacteraceae Altererythrobacter 99 20 1.24 Actinobacteria Micrococcales Microbacteriaceae Microbacterium 100 21 1.21 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Ancylobacter 99 22 1.21 Euryarchaeota Methanomicrobia Methanomicrobiales Methanoregulaceae Methanoregula 99 1.19 Proteobacteria Alphaproteobacteria Sphingomonadales Unclassified 100 23 Sphingmonadales 24 1.01 Proteobacteria Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium 95 25 0.98 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Elstera 100

224

Table B1-E. Heavy SIP Fractions (fractions 4 and 5) of OSPW sampled in August, 2011, amended with labelled protein extract for 14 days.

% Rank % Reads Phylum Class Order Family Genus Identity 1 18.15 Bacteroidetes Bacteroidia Bacteroidales Rikenellaceae Acetobacteroides 98 2 10.60 Firmicutes Clostridia Clostridiales Clostridiales Fusibacter 94 3 6.13 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 93 4 3.85 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 99 5 3.73 Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Defluviimonas 100 6 3.55 Proteobacteria Betaproteobacteria Burkholderiales Oxalobacteraceae Janthinobacterium 99 7 3.48 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 92 8 2.97 Firmicutes Clostridia Clostridiales Clostridiales Fusibacter 94 9 2.60 Spirochaetes Spirochaetes Spirochaetales Spirochaetaceae Treponema 92 10 2.12 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Thauera 100 11 1.80 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax 99 12 1.45 Bacteroidetes Bacteroidia Bacteroidales Rikenellaceae Acetobacteroides 96 13 1.22 Proteobacteria Gammaproteobacteria Enterobacteriales Enterobacteriaceae Citrobacter 100 14 1.13 Proteobacteria Gammaproteobacteria Aeromonadales Aeromonadaceae Aeromonas 100 15 1.03 Proteobacteria Deltaproteobacteria Desulfobacterales Desulfobacteraceae Desulfatiferula 96 16 1.02 Bacteroidetes Bacteroidia Bacteroidales Rikenellaceae Anaerocella 94 17 0.98 Proteobacteria Deltaproteobacteria Desulfovibrionales Desulfovibrionaceae Desulfovibrio 99 18 0.97 Proteobacteria Deltaproteobacteria Desulfovibrionales Desulfovibrionaceae Desulfovibrio 96 19 0.92 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Xanthobacter 98 20 0.92 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 98 21 0.85 Proteobacteria Deltaproteobacteria Syntrophobacterales Syntrophobacteraceae Syntrophobacter 97 22 0.83 Firmicutes Clostridia Clostridiales Clostridiales Fusibacter 94 23 0.75 Bacteroidetes Bacteroidia Bacteroidales Rikenellaceae Anaerocella 89 24 0.75 Proteobacteria Firmicutes Bacilli Bacillales Bacillaceae 88 25 0.73 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Xanthobacter 98

225

Table B1-F. Heavy SIP Fractions (fractions 4 and 5) of MLSB-OSPW sampled in August, 2011, amended with labelled CO2 for 12 weeks under light.

% Rank % Reads Phylum Class Order Family Genus Identity 1 45.11 Planctomycetes Planctomycetacia Planctomycetales Planctomycetaceae Aquisphaera 87 2 10.44 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Oceanibaculum 99 3 6.19 Bacteroidetes Sphingobacteriia Sphingobacteriales Saprospiraceae Haliscomenobacter 92 4 6.15 Proteobacteria Alphaproteobacteria Magnetococcales Magnetococcaceae Magnetococcus 90 5 3.67 Proteobacteria Gammaproteobacteria Xanthomonadales Xanthomonadaceae Luteimonas 97 6 2.43 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Phenylobacterium 99 7 2.16 Chloroflexi Anaerolineae Anaerolineles Anaerolineaceae Anaerolinea 90 8 1.83 Proteobacteria Alphaproteobacteria Parvularculales Parvularculaceae Amphiplicatus 97 9 1.75 Bacteroidetes Bacteroidetes Bacteroidetes Order II. Incertae sedis Rhodothermaceae Salinibacter 87 10 1.69 Acidobacteria Solibacterales Solibacterales Solibacterales Bryobacter 90 11 1.23 Proteobacteria Alphaproteobacteria Parvularculales Parvularculaceae Amphiplicatus 90 12 1.17 Proteobacteria Alphaproteobacteria Rhodobacterales Rhodobacteraceae Rhodobacter 99 13 0.96 Proteobacteria Alphaproteobacteria Rhizobiales Rhizobiaceae Bauldia 99 14 0.69 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 92 15 0.68 Proteobacteria Alphaproteobacteria Rhodospirillales Acetobacteraceae Roseomonas 99 16 0.66 Bacteroidetes Bacteroidia Bacteroidales Porphyromonadaceae Porphyrobacter 100 17 0.63 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 94 18 0.62 Chloroflexi Anaerolineae Anaerolineales Anaerolineaceae Leptolinea 84 19 0.60 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Planctopirus 95 20 0.54 Proteobacteria Alphaproteobacteria Rhizobiales Aurantimonadaceae Aureimonas 96 21 0.53 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 87 22 0.53 Chloroflexi Caldilineae Caldilineales Caldilineaceae Caldilinea 89 23 0.52 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Thalassobaculum 96 24 0.44 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Prosthecomicrobium 97 25 0.43 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Pedomicrobium 99

226

Table B1-G. Heavy SIP Fractions (fractions 4 and 5) of WIP-OSPW sampled in August, 2011, amended with labelled CO2 for 12 weeks under light.

% % Rank Phylum Class Order Family Genus Reads Identity 1 23.96 Bacteroidetes Sphingobacteriia Sphingobacteriales Saprospiraceae Haliscomenobacter 92 2 22.93 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 91 3 5.95 Acidobacteria Solibacterales Solibacterales Solibacterales Bryobacter 90 4 4.01 Proteobacteria Alphaproteobacteria Rhizobiales Rhizobiaceae Bauldia 99 5 3.72 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas 100 6 3.28 Proteobacteria Alphaproteobacteria Rhizobiales Bradyrhizobiaceae Bradyrhizobium 95 7 2.95 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Gemmata 89 8 2.62 Acidobacteria Solibacteres Solibacterales unclassified Solibacterales Paludibaculum 91 9 2.61 Proteobacteria Alphaproteobacteria Rhizobiales Rhizobiaceae Rhizobium 100 10 2.51 Chloroflexi Anaerolineae Anaerolineales Anaerolineaceae Anaerolinea 90 11 2.23 Proteobacteria Alphaproteobacteria Rhizobiales Phyllobacteriaceae Mesorhizobium 99 12 2.05 Proteobacteria Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum 89 13 2.03 Chloroflexi Anaerolineae Anaerolineales Anaerolineaceae Leptolinea 84 14 1.91 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 89 15 1.56 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 87 16 1.42 Planctomycetes Planctomycetia Planctomycetales Planctomycetaceae Thermogutta 88 17 1.41 Planctomycetes Planctomycetacia Planctomycetales Planctomycetaceae Aquisphaera 87 18 1.15 Acidobacteria Acidobacteria Acidobacteriales Acidobacteriaceae Granulicella 88 19 1.01 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas 97 20 0.75 Chloroflexi Caldilineae Caldilineales Caldilineaceae Litorilinea 90 21 0.68 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 94 22 0.65 Bacteroidetes Sphingobacteriia Sphingobacteriales Chitinophagaceae Chitinophaga 93 23 0.61 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Azospirillum 92 24 0.60 Actinobacteria Actinobacteria Micrococcales Micrococcaceae Micrococcus 100 25 0.59 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Pedomicrobium 95

227

APPENDIX C: SUPPLEMENTARY INFORMATION – DETECTION AND FUNCTIONAL ANALYSIS OF A NOVEL CU-MONOOXYGENASE DETECTED VIA METAGENOMIC ANALYSIS OF WIP-OSPW

Table C1. Complete contig sequence from which the xmoCAB primer set was designed from. Sequence was obtained from the metagenome of WIP-OSPW sampled in 2011.

Sequence length: 392 bp.

CGA CTC ACT ATA GGG AGA GCG GCC GCC AGA TCT TCC GGA TGG CTC GAG TTT

TTC AGC AAG AT AAA TGA CTT CGC TCG CTG ATT GGC TTG GCT ACA CAT TCC

CTC GAG CAG GCA CGC CTG AAT ACA TCC GGA TCA TTG AAC GCG GAA CCC

TGC GCA CGT TTC AAG ACT CTG CGC AGT GGG TGT CGG CAT TCT TCT CTG GGT

TTA TCT GCG TTT TCA TGT ACT ACA CCT TCT GGT GGA TTG GCA ACG CGG CCT

GTT CCA TAA AGT ATG TTC CTA TCG GAA TCA AAA TGA AGG AAG CGT TTG GAG

CCA ATG ACG CAG CAA AGA AGG TGG CAG CAT G A TCT TTC TAG AAG ATC TCC

TAC AAT ATT CTC AGC TGC CAT GGA AAA TCG ATG TTC TTC T

Table C2. KAPA HiFi HotStart ReadyMix PCR reaction protocol for Illumina amplicon sequencing.

Reagent Volume Microbial DNA 2.5 μL Forward Primer 1 μM 5.0 μL Reverse Primer 1 μM 5.0 μL 2x KAPA HiFi HotStart ReadyMix 12.5 μL

Total 25 μL

228

Table C3. Calculated concentration copies formulae.

A. Calculated Concentration for enrichment samples

CC x 100 μl* = Copies ml-1 of OSPW sample

18 ml of OSPW**

CC is Calculated Concentration (copies) per μl-1 of sample (values automatically generated based on qPCR standard curve).

* Each DNA extracted sample was diluted to 100 μl.

** The amount of each OSPW enrichment sample was 20 ml, 2 ml being preserved and 18 ml DNA-extracted.

B. Calculated Concentration for SIP samples

CC x 30 μl* = Copies per SIP fraction

* Each SIP fraction contain 30 μl of DNA sample.

C. Standard Error (SE)

s is the sample standard deviation (i.e., the sample-based estimate of the standard deviation of the population), and n is the size (number of observations) of the sample.

229

Table C4. A-E The top 10 OTUs of each sample site based on percent (%) of total reads in 16S rRNA gene sequencing analysis. OTUs were clustered based on their closest related species. Table C4-A. Reads from DNA in MLSB-OSPW sampled in August 2015.

Rank % Reads Phylum Class Order Family Genus % Identity 1 9.62 Proteobacteria Betaproteobacteria Nitrosomonadales Nitrosomonadaceae Nitrosomonas 98 2 9.21 Bacteroidetes Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium 99 3 6.80 Proteobacteria Gammaproteobacteria Cellvibrionales Porticoccaceae Porticoccus 93 4 6.66 Acidobacteria Blastocatellia Blastocatellales Blastocatellaceae Stenotrophobacter 94 5 4.86 Proteobacteria Gammaproteobacteria Xanthomonadales Xanthomonadaceae Arenimonas 97 6 2.72 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Hyphomicrobium 96 7 2.67 Proteobacteria Gammaproteobacteria Chromatiales Granulosicoccaceae Granulosicoccus 92 8 2.14 Proteobacteria Gammaproteobacteria Chromatiales Chromatiaceae Rheinheimera 94 9 2.13 Planctomycetes Phycisphaerae Phycisphaerales Phycisphaeraceae Algisphaera 89 10 2.00 Proteobacteria Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter 94

Table C4-B. Heavy SIP fraction (fraction 5 and 6) of MLSB-OSPW sampled in August 2011.

Rank % Reads Phylum Class Order Family Genus % Identity 1 9.81 Proteobacteria Gammaproteobacteria Cellvibrionales Porticoccaceae Porticoccus 93 2 8.33 Proteobacteria Gammaproteobacteria Xanthomonadales Xanthomonadaceae Arenimonas 97 3 7.85 Proteobacteria Betaproteobacteria Nitrosomonadales Nitrosomonadaceae Nitrosomonas 98 4 6.54 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Hyphomicrobium 96 5 6.23 Proteobacteria Gammaproteobacteria Chromatiales Granulosicoccaceae Granulosicoccus 92 6 5.04 Proteobacteria Gammaproteobacteria Alteromonadales Alteromonadaceae Marinobacter 93 7 3.51 Acidobacteria Blastocatellia Blastocatellales Blastocatellaceae Stenotrophobacter 94 8 2.88 Proteobacteria Gammaproteobacteria Ectothiorhodospiraceae Methylohalomonas 91 9 2.27 Proteobacteria Alphaproteobacteria Rhizobiales Rhodobiaceae Parvibaculum 98 10 1.86 Proteobacteria Gammaproteobacteria Chromatiales Chromatiaceae Thiocapsa 99

230

Table C4-C. Heavy SIP fractions (fractions 4 and 5) of MLSB-OSPW sampled in August 2015, amended with labelled methane for 10 days. % % Rank Phylum Class Order Family Genus Reads Identity 1 56.21 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methyloparacoccus 97 2 11.85 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas 99 3 9.16 Proteobacteria Betaproteobacteria Methylophilales Methylophilaceae Methylotenera 98 4 3.42 Proteobacteria Gammaproteobacteria Methylococcales Methylococcaceae Methylomonas 98 5 3.00 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Caulobacter 99 6 2.99 Actinobacteria Actinobacteria Actinomycetales Microbacteriaceae Microbacterium 99 7 1.77 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 99 8 1.40 Bacteroidetes Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium 98 9 1.19 Acidobacteria Blastocatellia Blastocatellales Blastocatellaceae Stenotrophobacter 94 10 0.70 Planctomycetes Planctomycetia Candidatus Brocadiales Candidatus Brocadiaceae Candidatus Kuenenia 84

Table C4-D. Heavy SIP fractions (fractions 4 and 5) of MLSB-OSPW sampled in August 2015, amended with labelled ethane for 10 days.

Rank % Reads Phylum Class Order Family Genus % Identity 1 41.26 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 99 2 21.70 Proteobacteria Alphaproteobacteria Rhizobiales Xanthobacteraceae Xanthobacter 99 3 3.40 Proteobacteria Gammaproteobacteria Xanthomonadales Xanthomonadaceae Lysobacter 98 4 2.43 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Hyphomicrobium 99 5 1.76 Proteobacteria Alphaproteobacteria Rhodospirillales Rhodospirillaceae Lacibacterium 93 6 1.70 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Blastochloris 94 7 1.61 Proteobacteria Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas 99 8 1.17 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Sulfuritalea 97 9 0.98 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Ramlibacter 97 10 0.94 Planctomycetes Planctomycetia Candidatus Brocadiales Candidatus Brocadiaceae Candidatus Kuenenia 84

231

Table C4-E. Heavy SIP fractions (fractions 3 and 4) of MLSB-OSPW sampled in August 2015, amended with labelled propane for 10 days. % % Rank Phylum Class Order Family Genus Reads Identity 1 33.46 Proteobacteria Betaproteobacteria Rhodocyclales Rhodocyclaceae Methyloversatilis 99 2 11.41 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Acidovorax 99 3 7.40 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Hydrogenophaga 99 4 5.56 Proteobacteria Alphaproteobacteria Rhizobiales Hyphomicrobiaceae Hyphomicrobium 97 5 4.75 Proteobacteria Betaproteobacteria Burkholderiales Comamonadaceae Rhodoferax 99 6 3.47 Proteobacteria Gammaproteobacteria Xanthomonadales Xanthomonadaceae Arenimonas 97 7 2.17 Acidobacteria Blastocatellia Blastocatellales Blastocatellaceae Stenotrophobacter 94 8 1.96 Proteobacteria Betaproteobacteria Burkholderiales Oxalobacteraceae Oxalicibacterium 97 9 1.18 Proteobacteria Betaproteobacteria Burkholderiales Burkholderiaceae Burkholderia 95 Candidatus Candidatus 10 0.81 Planctomycetes Planctomycetia Brocadiales Brocadiaceae Candidatus Kuenenia 84

232

233

Figure C3. Representative methane oxidation time courses of replicate samples from MLSB tailings pond sampled in August 2015. Bars represent ±SEM.

Figure C4. Representative ethane oxidation time courses of replicate samples from MLSB tailings pond sampled in August 2015. Bars represent ±SEM.

234

Figure C5. Representative propane oxidation time courses of replicate samples from MLSB tailings pond sampled in August 2015. Bars represent ±SEM.

235

236

APPENDIX D: COPYRIGHT PERMISSIONS

One Table used in Chapter 2, taken from: Rojo, F. (2010). Enzymes for Aerobic Degradation of Alkanes. In Handbook of Hydrocarbon and Lipid Microbiology, pp. 781-797. Edited by K. N. Timmis. Berlin, Heidelberg: Springer Berlin Heidelberg.

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One Figure used in Chapter 2, taken from: Cerniglia, C. E. (1992). Biodegradation of polycyclic aromatic hydrocarbons. Biodegradation 3, 351-368.

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One Figure used in Chapter 2, taken from: Bamforth, S. M. & Singleton, I. (2005). Bioremediation of polycyclic aromatic hydrocarbons: current knowledge and future directions. Journal of Chemical Technology & Biotechnology 80, 723-736.

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Title: Bioremediation of polycyclic Logged in as: aromatic hydrocarbons: current Fauziah Rochman knowledge and future directions Author: Selina M Bamforth,Ian Singleton Publication: Journal of Chemical Technology & Biotechnology Publisher: John Wiley and Sons Date: Apr 22, 2005 Copyright © 2005 Society of Chemical Industry

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License Number 3924360101247 License date Aug 08, 2016 Licensed Content John Wiley and Sons Publisher Licensed Content Journal of Chemical Technology & Biotechnology Publication Licensed Content Title Bioremediation of polycyclic aromatic hydrocarbons: current knowledge and future directions Licensed Content Selina M Bamforth,Ian Singleton Author Licensed Content Date Apr 22, 2005 Licensed Content Pages 14 Type of use Dissertation/Thesis Requestor type University/Academic Format Print and electronic Portion Figure/table Number of 1 figures/tables Original Wiley Figure 3 figure/table number(s) Will you be translating? No Title of your thesis / Aerobic hydrocarbondegrading microbial communities in oilsands tailings ponds dissertation Expected completion Sep 2016 date Expected size (number 280 of pages) Requestor Location Fauziah Rochman 118 Kincora Glen Mews NW

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One Figure used in Chapter 2, taken from: Chen, Y. & Murrell, J. C. (2010). When metagenomics meets stable-isotope probing: progress and perspectives. Trends in Microbiology 18, 157-163.

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Title: When metagenomics meets Logged in as: stableisotope probing: progress Fauziah Rochman and perspectives Author: Yin Chen,J. Colin Murrell Publication: Trends in Microbiology Publisher: Elsevier Date: April 2010 Crown copyright © 2010 Published by Elsevier Ltd. All rights reserved.

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License Number 3924360362798 License date Aug 08, 2016 Licensed Content Elsevier Publisher Licensed Content Trends in Microbiology Publication Licensed Content Title When metagenomics meets stableisotope probing: progress and perspectives Licensed Content Author Yin Chen,J. Colin Murrell Licensed Content Date April 2010 Licensed Content Volume 18 Licensed Content Issue 4 Licensed Content Pages 7 Type of Use reuse in a thesis/dissertation Portion figures/tables/illustrations Number of 1 figures/tables/illustrations Format both print and electronic Are you the author of this No Elsevier article? Will you be translating? No Order reference number Original figure numbers Figure 1 Title of your Aerobic hydrocarbondegrading microbial communities in oilsands tailings ponds thesis/dissertation Expected completion date Sep 2016 Estimated size (number 280 of pages) Elsevier VAT number GB 494 6272 12 Requestor Location Fauziah Rochman 118 Kincora Glen Mews NW

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