Biodegradation of Aromatic Hydrocarbons by Methanogenic Consortia and Groundwater-Associated Microbial Communities

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2021-01-08 Biodegradation of Aromatic Hydrocarbons by Methanogenic Consortia and Groundwater-Associated Microbial Communities

Taylor, Nicole

Taylor, N. (2021). Biodegradation of Aromatic Hydrocarbons by Methanogenic Consortia and Groundwater-Associated Microbial Communities (Unpublished master's thesis). University of Calgary, Calgary, AB. http://hdl.handle.net/1880/112987 master thesis

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Biodegradation of Aromatic Hydrocarbons by Methanogenic Consortia and Groundwater-

Associated Microbial Communities

Nicole Taylor

A THESIS

SUBMITTED TO THE FACULTY OF GRADUATE STUDIES

IN PARTIAL FULFILMENT OF THE REQUIREMENTS FOR THE

DEGREE OF MASTER OF SCIENCE

GRADUATE PROGRAM IN BIOLOGICAL SCIENCES

CALGARY, ALBERTA

JANUARY, 2021

The biodegradation of hydrocarbons is an important environmental process responsible for in situ remediation of crude oil and gas components. Microorganisms of many lineages and redox conditions have been characterized to degrade numerous types of petroleum hydrocarbons, including those with aromatic structures. Alkyl-substituted mono- and polycyclic aromatic hydrocarbons are more chemically reactive than their unsubstituted counterparts, and as such their anaerobic degradation pathways have been studied to varying degrees. Aromatic hydrocarbons require enzymatic functionalization before biodegradation can occur; these activation enzymes and products are often unique to anaerobic reactions, therefore identifying the metabolites produced or the enzymes carrying out these reactions lends evidence to identifying in situ bioremediation of aromatic hydrocarbon contamination. Hydrocarbon biodegradation in the deep subsurface is often associated with methanogenesis.

Anaerobic toluene degradation has been extensively studied and has been shown in multiple studies to involve an activation process known as fumarate addition, however methanogenic biodegradation of other alkylbenzenes and polycyclic aromatic hydrocarbons is comparatively poorly understood. In this work, the biodegradation of ethylbenzene and p-xylene was examined in the presence of toluene; p-toluic acid was found as a metabolite of p-xylene biotransformation, but no evidence of fumarate addition to either p-xylene or ethylbenzene were observed. A second methanogenic biodegradation study of naphthalene, 2-methylnaphthalene, and phenanthrene revealed 2-naphthoic acid as the primary metabolite produced by microbial cultures. A third study involved evaluating the use of a trapping device for passively sampling microorganisms from groundwater contaminated with aromatic hydrocarbons; this study showed that the chosen sorptive material did not influence the biodiversity of microbial communities, did ii

not influence the rate of hydrocarbon biodegradation, and the presence of hydrocarbons was correlated to higher biomass recovery.

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Acknowledgements

My time in this program has been some of the best years of my life. The support of my family, friends and colleagues have been instrumental to my success and I am incredibly grateful.

In particular I would like to thank my sister, parents, and grandparents, who despite the physical distance have always been there for me when I needed them; and to all the members of the

University of Calgary community who have made my time here a success. To my supervisor Dr.

Lisa Gieg for her encouragement, guidance, advice, and faith in me; and to my supervisory committee Drs. Hubert and Dunfield for their valuable insight and feedback. To Gabrielle,

Gurpreet, Danika, Julie, Ciara, Natalie, Yin, Rita, Gloria, Nuno, Mohita, and all past or present members of the Gieg lab family for being friends more than colleagues. Your constant advice, suggestions, problem solving, and sometimes just venting about frustrations made a molehill out of what would otherwise have been a mountain. Lunchtime laughs, coffee walks, and ice cream trips were indispensable mental health breaks. Special thanks to my mentor Dr. Courtney Toth for teaching me the essentials for working with hydrocarbons and TOLDC, to Gurpreet Kharey for teaching me the fundamentals of molecular biology, and to Dr. Victoria Collins for collaborating on experiments described in Chapter Six. Funding for this project was provided by NSERC

Discovery, Western Canadian Innovation Offices, and the Genome Canada LSARP grants all awarded to Dr. Lisa Gieg. Isaac Newton once wrote, “if I have seen further it is by standing on the shoulders of giants.” By lifting me up, you are all giants to me and for that I will always be grateful.

Dedication

To Bradley: my partner, my love.

I owe this all to you.

Table of Contents

Abstract ...... ii Acknowledgements ...... iv Dedication ...... v Table of Contents ...... vi List of Tables ...... x List of Figures and Images ...... xii List of Symbols, Abbreviations and Nomenclature ...... xviii

Chapter One: Introduction ...... 1 1.1 Research objectives ...... 1 1.2 Thesis structure and summary ...... 2

Chapter Two: Literature Review ...... 4 2.1 Hydrocarbons in the environment ...... 4 2.2 Origins of crude oil ...... 5 2.3 Chemical and physical properties of petroleum hydrocarbons ...... 5 2.3.1 Features of aromatic hydrocarbons ...... 6 2.4 Hydrocarbon biodegradation ...... 8 2.5 Aerobic biodegradation of aromatic hydrocarbons ...... 11 2.6 Anaerobic biodegradation of toluene ...... 14 2.6.1 Hydrocarbon addition to fumarate (fumarate addition) ...... 16 2.6.2 Benzylsuccinate synthase ...... 17 2.7 Anaerobic activation of other alkylbenzenes ...... 19 2.7.1 Ethylbenzene ...... 20 2.7.2 Xylenes ...... 22 2.7.3 Alkylated PAHs ...... 23 2.8 Activation of unsubstituted aromatic hydrocarbons ...... 24 2.9 Substrate range of hydrocarbon-biodegrading microorganisms ...... 26 2.10 Remediation technologies ...... 27 2.11 Research needs ...... 30

Chapter Three: General Materials and Methods ...... 32 3.1 Microbial cultures and strains ...... 32 3.2 General cultivation of anaerobic microorganisms ...... 32 3.2.1 Media recipes and procedures ...... 32 3.2.2 Addition of BTEX substrates ...... 33 3.2.3 Addition of PAH substrates ...... 33 3.2.4 Adsorptive materials (Amberlite and Tenax) ...... 33 3.3 Gas chromatography-flame ionization detection (GC-FID) ...... 34 3.3.1 Hydrocarbon analysis...... 34 3.3.2 Methane analysis ...... 35 3.4 Gas chromatography-thermal conductivity detection (GC-TCD) ...... 35 3.5 High-performance liquid chromatography (HPLC)...... 35

3.5.1 Quantification of organic acids ...... 35 3.5.2 Detection of anions ...... 36 3.6 Spectrophotometric analysis of iron ...... 36 3.7 ATP assay ...... 36 3.8 Chemical analysis via gas chromatography-mass spectrometry (GC-MS) ...... 36 3.8.1 PAH quantification ...... 36 3.8.2 Metabolite identification ...... 37 3.9 DNA analysis ...... 38 3.9.1 DNA extraction ...... 38 3.9.2 16S rRNA gene sequencing with Illumina MiSeq ...... 38 3.9.3 Bioinformatics processing ...... 39 3.9.4 Data interpretation and ecological comparisons using R ...... 39 3.10 RNA extraction, purification, and reverse transcription ...... 40 3.11 qPCR assays ...... 40 3.12 Scanning electron microscopy ...... 41 3.13 Statistical analyses ...... 41

Chapter Four: Elucidating activation metabolites of alkylbenzenes in a toluene-degrading methanogenic enrichment culture ...... 43 4.1 Introduction ...... 43 4.2 Methods...... 46 4.2.1 Toluene-degrading methanogenic enrichment culture (TOLDC)...... 46 4.2.2 Syntrophus aciditrophicus bioaugmentation ...... 47 4.2.3 Phase 1 enrichments ...... 48 4.2.4 Phase 2 enrichments ...... 48 4.2.5 Phase 3 enrichments ...... 49 4.3 Results ...... 50 4.3.1 S. aciditrophicus bioaugmentation ...... 50 4.3.2 Phase 2 experiments ...... 50 4.3.3 Phase 3 enrichments ...... 59 4.4 Discussion ...... 62 4.4.1 Ethylbenzene ...... 62 4.4.2 Hydrocarbon transformation ...... 62 4.4.3 Metabolite analyses ...... 64 4.4.3.1 p-Toluic acid ...... 64 4.4.3.2 Benzylsuccinic acid ...... 67 4.4.4 qPCR assay optimization ...... 68 4.4.5 Reverse-transcription bssA gene expression analysis...... 70 4.4.6 Microbial community analysis ...... 71 4.5 Conclusions ...... 78

Chapter Five: Characterizing the biodegradation of polycyclic aromatic hydrocarbons by methanogenic microbial communities ...... 80 5.1 Introduction ...... 80 5.2 Methods...... 84

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5.3 Results ...... 86 5.3.1 Methane production and degradation of substrates ...... 86 5.3.2 Metabolite analysis ...... 90 5.3.3 Microbial community analysis ...... 94 5.4 Discussion ...... 96 5.4.1 PAH transformation and methane production ...... 96 5.4.2 Metabolite analysis ...... 98 5.4.3 Microbial community analysis ...... 102 5.5 Conclusions ...... 105

Chapter Six: The effect of a sorbent matrix on recovery of microorganisms from contaminated groundwater ...... 106 6.1 Introduction ...... 106 6.2 Materials and methods ...... 109 6.2.1 Survey of different adsorbent materials ...... 109 6.2.2 Experimental microcosms ...... 110 6.2.3 Analytical procedures ...... 112 6.3 Results ...... 113 6.3.1 Field testing of microbial trapping matrices ...... 113 6.3.2 Hydrocarbon degradation and mineralization of electron acceptors ...... 115 6.3.3 Microbial community analysis ...... 119 6.3.4 Visualization of Tenax-TA ...... 122 6.4 Discussion ...... 124 6.4.1 Matrix effects ...... 124 6.4.2 Biodegradation of hydrocarbons ...... 125 6.4.3 Mineralization analysis ...... 126 6.4.4 Microbial community analysis ...... 128 6.5 Conclusions ...... 131

Chapter Seven: Conclusions ...... 133 7.1 Research objectives and hypotheses ...... 133 7.2 Conclusions and future directions ...... 134 7.2.1 Chapter Four ...... 134 7.2.2 Chapter Five ...... 135 7.2.3 Chapter Six...... 137

Literature Cited ...... 139

Appendix A: Supplementary Material from Chapter Four ...... 160

Appendix B: Supplementary Material from Chapter Six ...... 174

Appendix C: Medium Recipes ...... 182

Appendix D: R Scripts for Diversity Analyses ...... 184

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Appendix E: Bioelectrochemical remediation of phenanthrene in a microbial fuel cell using an anaerobic consortium enriched from a hydrocarbon-contaminated site ...... 186

Appendix F: Comparative evaluation of coated and non-coated carbon electrodes in microbial fuel cells for treatment of municipal sludge ...... 197

List of Tables

Table 3-1: Primer-pair sequences used for Illumina MiSeq. Primers 926Fi5 and 1392Ri7 targeted the V6 to V8 hypervariable region of the 16S rRNA gene and were used for microbial community analysis. Nextera adaptor sequences are in bold...... 39

Table 3-2: Primer-pair sequences targeting bssA used in qPCR assays...... 40

Table 4-1: Predicted metabolites of fumarate addition or metabolites reported in literature sought in co-metabolic experiments with toluene, ethylbenzene, and p-xylene. Fragment ions are reported by Gieg and Toth (2017)...... 54

Table 5-1: Calculated stoichiometric yields for the complete degradation of naphthalene, 2- methylnaphthalene, and phenanthrene under methanogenic conditions (Symons and Buswell, 1933; Meckenstock et al., 2016)...... 89

Table 5-2: Predicted metabolites of naphthalene, 2-methylnaphthalene, or phenanthrene biodegradation surveyed. Compounds were searched for by their diagnostic ions when authentic standards were not available. A “-” indicates that the putative metabolite was not detected, while a “+” indicates that the compound was detected in the methanogenic enrichments...... 90

Table 6-1: Conditions and various controls of experimental microcosms. Each combination was conducted under either aerobic, nitrate-reducing, iron(III)-reducing, or sulfate- reducing conditions. A fifth condition included no additional electron acceptor to examine the effect of endogenous electron accepting processes...... 111

Table 6-2: Stoichiometric equations for the complete metabolism of benzene and toluene - under the experimental electron-accepting conditions surveyed. Depletion of NO3 and 2- SO4 (from nitrate and sulfate-reducing treatments, respectively) and accumulation of 2+ CO2 (from aerobic treatments), Fe (from iron-reducing treatments), and CH4 (from no EA added treatments) were monitored in this microcosm study and results are reported in Table 6-3...... 118

Table 6-3: Predicted and actual hydrocarbon transformation yields in microcosms under various electron-accepting conditions. Predicted yields were calculated using stoichiometric equations shown in Table 6-2. Actual yields were measured as described in Chapter Three and averaged from triplicate microcosms; error bars indicate standard error of the mean. An unpaired, two-tailed t-test was calculated to determine statistically significant differences between the product yields from Tenax-TA and no Tenax-TA treated microcosms...... 119

Table A-1: Relative abundance of the complete 16S rRNA gene sequencing lineages from TOLDC incubated with toluene, ethylbenzene, or p-xylene in phase 2 enrichments. Lineages not fully characterized to the species level are denoted by NA. Taxa believed to be important in toluene metabolism are shown in bold...... 163 x

Table A-2: Detection of bssA by qPCR from cDNA for expression analysis. Technical replicates were analyzed in triplicate...... 173

Table A-3: Diversity metrics from TOLDC microbial communities incubated with toluene, ethylbenzene, and p-xylene and sequenced 100 days into the experiment and again after 600 days of further incubation...... 173

Table B-1: DNA recoveries from the initial survey of trapping materials efficacy in water and soil to loosely approximate a sampling well with exposed soil (5 g soil in 30 ml sterile DI water). Fertilizer (Miracle-Gro Garden Feeder, 28-8-16) was added at 0.33 g/L to one treatment to enhance growth. Matrix materials included zeolite (molecular sieve, 8-12 mesh, 3Å, 208582; Sigma Aldrich, Oakville, Canada), activated carbon (CAS 7440-44-0, L16334; Alfa Aesar, Haverhill, USA), Mat540 (porous 30 μm silica microspheres; Materium Innovations, Ithaca, USA), diatomaceous earth (Red Lake Earth, Kamloops, Canada), and ZMM® T-carbon (biochar from 2 mm woody feedstock; Canada Minerals Corp., Peachland, Canada). DNA was extracted from two replicates of each material following 6 days of incubation at room temperature in the dark and removal of excess soil by gentle rinsing with sterile DI water. No DNA was recovered from activated carbon or Mat540, while only 0.3-0.6 ng/µL was recovered from zeolite. Average DNA recoveries from duplicate extractions are shown...... 174

Table B-2: DNA extraction concentrations from field trials with diatomaceous earth (DE), T- carbon (TC), and Tenax-TA (TA). Trap samplers were deployed into a hydrocarbon- contaminated aquifer and recovered in one-month intervals for a total of three months. Recovered and extracted DNA concentrations are provided for each replicate. Some samples had DNA concentrations higher than the detection limit (> 60 ng/μL) of the instrument and are reported as “too high” or TH. Values for Shannon and Simpson diversity indices were computed in R using vegan...... 175

Table B-3: Normalized DNA recoveries from experimental Tenax-TA incubations, comparing incubations with or without hydrocarbons (HCs), with or without Tenax-TA, and the fraction the sample was collected from (planktonic or sessile). Raw extractant DNA concentrations were normalized based on the amount of starting material (0.13 g for sessile samples, 5 mL for planktonic samples). DNA recoveries that were too low to quantify (< 0.05 ng/μL) are denoted as NA...... 176

List of Figures and Images

Figure 2-1: Structures of common monoaromatic hydrocarbons including (from left to right) benzene, toluene, ethylbenzene, o-xylene, m-xylene, and p-xylene (BTEX)...... 7

Figure 2-2: Generalized depiction of the redox zonation effect to describe how electron acceptors are distributed and sequentially mineralized in a hydrocarbon plume in groundwater. Aerobic microorganisms are generally the fastest degraders and first to be recruited, followed by nitrate reduction as oxygen is depleted, then iron(III) reduction, sulfate reduction, and finally methanogenesis when other electron acceptors are depleted (adapted from Meckenstock et al., 2015)...... 10

Figure 2-3: Examples of activation pathways of toluene common to many aerobic bacteria mediated by mono- or dioxygenase activity. Modified from Parales et al. (2008) and Martínez-Lavanchy et al. (2015)...... 13

Figure 2-4: Generalized reaction of toluene (a) with fumarate (b) mediated by benzylsuccinate synthase (BSS) to form benzylsuccinate (c). Benzylsuccinyl-CoA (d) is formed through the activity of succinyl-CoA:(R)-benzylsuccinate CoA-transferase. A series of hydratase and dehydrogenase reactions results in formation of benzoylsuccinyl-CoA (e). Benzoylsuccinyl-CoA thiolase cleaves succinyl-CoA (to reform fumarate in the cycle) from benzoyl-CoA (f). Benzoyl-CoA is then used by fermentative microorganisms such as Desulfovibrio or Syntrophus via the β-oxidation pathway. Summarized from Biegert et al. (1996), Boll et al. (2014), and Michas et al. (2017)...... 17

Figure 4-1: Hydrocarbon degradation in phase 2 enrichments over 150 days after pooling and scaling up. Separate incubations of TOLDC were established with different carbon sources including toluene only, ethylbenzene only, p-xylene only, toluene + ethylbenzene, toluene + p-xylene, and an unamended treatment. After toluene (A) was depleted by approximately day 60, enrichments were reamended with 100 μmoles of toluene. Ethylbenzene (B) and p-xylene (C) were not reamended as they never reached zero. n=1...... 51

Figure 4-2: Methane production from TOLDC enriched on various carbon sources from phase 2 enrichments, then monitored for a further 150 days during the RNA and metabolite extraction experiment. Carbon sources in this experiment included: toluene only, ethylbenzene only, p-xylene only, toluene + ethylbenzene, toluene + p-xylene, and an unamended control treatment. n 1...... 52

Figure 4-3: Microbial community composition of TOLDC enriched on different carbon substrates at day 100 of the initial phase 1 incubation and again at day 600. The top 10 taxa from each sample with their most detailed taxonomic classification are displayed here, with all others being grouped as “Other”. Total taxonomic lineages of all samples can be found in Appendix Table A-1...... 53

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Figure 4-4: Bray-Curtis dendrograms to highlight the diversity changes in the microbial communities of TOLDC enriched on various substrates when sequenced at day 100 (A) and day 600 (B). Analyses were completed in R (scripts described in Appendix D)...... 54

Figure 4-5: Benzylsuccinic acid detected in phase 2 enrichments over 60 days of monitoring. Toluene only treatments lost almost all benzylsuccinic acid by day 60, while it persisted at higher amounts in co-amended treatments (toluene with ethylbenzene and toluene with p-xylene). n=2, error bars ± SEM...... 56

Figure 4-6: (A) Chromatograph overlay of TOLDC sample extracts from cultures amended with toluene, p-xylene, or toluene + p-xylene, an unamended control, and a p-toluic acid standard (all TMS-derivatized). The peak with a retention time of 31.8 minutes (denoted by the black arrow; blue trace) was observed only in the TOLDC with toluene + p-xylene culture extracts, matching with a p-toluic acid standard (black trace). This peak was not observed in toluene only, p-xylene only, or unamended treatments. (B) Fragment ion profile of authentic TMS-derivatized p-toluic acid, with diagnostic fragment ions at m/z 193 and 208 (denoted by black arrows). (C) Fragment ion profile of the metabolite detected in TOLDC amended with toluene + p-xylene, with diagnostic ions at m/z 193 and 208...... 57

Figure 4-7: The amounts of p-toluic acid detected in the toluene + p-xylene amended incubations. This metabolite was not detected in toluene only or unamended treatments; p-xylene only treatments contained roughly 1.5 nmoles of p-toluic acid throughout the experimental monitoring period while levels in toluene with p-xylene treatments ranged from 40 to 85 nmoles. Error bars represent the standard error of the mean of 2 replicates...... 58

Figure 4-8: Fold changes in TOLDC bssA gene expression relative to the unamended treatment, quantified via qPCR from cDNA over time when incubated with toluene or ethylbenzene (A), toluene + ethylbenzene (B), p-xylene (C), and toluene + p-xylene (D). Error bars represent the standard error of the mean of 3 technical replicates...... 59

Figure 4-9: Average methane production from phase 3 experiments of TOLDC amended with toluene and/or p-xylene. Error bars represent the standard error of the mean of 3 replicates...... 60

Figure 4-10: Hydrocarbon loss from phase 3 TOLDC with toluene (A) and p-xylene (B). Unpaired two-tailed t-tests of the change in toluene concentration relative to the sterile control from day 1 to day 50 (A) determined p-values for the toluene only treatment (blue) of 0.0005% and 0.0011% for toluene from the co-amended treatment (green). Non- significant p-values were calculated for the changes in all p-xylene treatments. Error bars ± SEM, n=3...... 61

Figure 4-11: Benzylsuccinic acid (A) and p-toluic acid (B) in phase 3 enrichments with toluene, p-xylene, toluene + p-xylene or unamended treatments. Error bars ± SEM, n=3. . 61

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Figure 5-1: Methane production from the initial soil enrichment cultures (T0) over 600 days of monitoring. Amended enrichments were established with a cocktail of PAHs containing naphthalene, 2-methylnaphthalene, and phenanthrene in HMN as the carbon sources. Unamended incubations received HMN with no PAHs. Breaks in the lines denote when enrichment bottles were opened in an anoxic glove bag to filter and transfer to subsequent incubations. n=1...... 86

Figure 5-2: Methane production from the first round of transfers (T1) from filtered and diluted enrichment stock cultures. Consistently, the unamended incubation produced more methane than that amended with PAHs. n=1...... 87

Figure 5-3: Methane production from the second round of transfers (T2) from the stock enrichment cultures. Methane produced by these treatments parallels all subsequent transfers wherein approximately 1 μmole of methane was produced. n=3, error bars ± SEM...... 87

Figure 5-4: PAHs degraded in a time-course incubation (T4), displayed as the percent of hydrocarbons remaining relative to the sterile controls (100%) (not shown). By day 240, naphthalene was depleted by 81%, 2-methylnaphthalene by 78%, and phenanthrene by 61%. Statistically significant loss of hydrocarbons was observed by day 240 (** p-value ≤ 0.01, *** p-value ≤ 0.001) relative to the sterile controls as determined by t-tests. Error bars ± SEM of three replicates...... 89

Figure 5-5: TMS-derivatized mass spectral profiles of an authentic 2-naphthoic acid standard (A) and 2-naphthoic acid detected in live PAH-amended incubations (B). Diagnostic fragment ions of m/z 229 and 244 were detected in both samples and are denoted by black arrows...... 91

Figure 5-6: 2-Naphthoic acid quantified from PAH-amended live and sterile incubations. n=3, error bars ± SEM...... 92

Figure 5-7: Total ion chromatogram and mass spectral profiles of a metabolite formed in the live PAH-amended cultures. A peak with a retention time of 48.8 minutes was observed in all live incubations at after 90, 120 and 240 days of incubation but was absent at day 30 (A). TMS-derivatized diagnostic fragment ions of m/z 367 and 382 were identified in all samples with this peak (B)...... 93

Figure 5-8: Peak area integration of a putative metabolite detected only in live PAH-amended incubations (Figure 5-7) showing its accumulation and subsequent decrease in abundance during the incubation. Error bars represent the standard error of the mean of three replicates...... 94

Figure 5-9: Microbial community analysis of the soil inoculum, enrichment cultures (T0) after 150 days of incubation, T1 and T4 transfers after 650 day of total incubation. Taxa representing the 10 most abundant of each sample are displayed, the rest are grouped as Other...... 95 xiv

Figure 5-10: Possible structures of a metabolite detected in GC-MS analysis arising naphthalene or 2-methylnaphthalene transformation (A and B) or phenanthrene transformation (C) from Figure 5-7. (A) IUPAC: 2-[(2E,4E)-5-carboxypenta-2,4-dien-1- yl]cyclohexane-1-carboxylic acid. Chemical formula C13H18O4 and molecular weight 238.28 g/mol. (B) IUPAC: 2-(carboxymethyl)-1,2,3,4,5,6,7,8-octahydronaphthalene-1- carboxylic acid. Chemical formula C13H18O4 and molecular weight 238.28 g/mol; the exact location of the carboxylic acid, carboxymethyl group, and double bond are unknown. (C) IUPAC: 2-hydroxyphenanthrene-1-carboxylic acid. Chemical formula C15H10O3 and molecular weight 238.24 g/mol; the exact location of the carboxylic acid and alcohol groups are unknown...... 101

Figure 6-1: Average DNA recoveries from various trap matrices (blue bars) initially tested in groundwater samples, and the associated average Shannon diversity indices (black points) of the microbial communities analyzed through 16S rRNA gene sequencing and R (vegan). Error bars represent the standard error of the mean of 18 replicates per treatment...... 114

Figure 6-2: Microbial community composition of field samples collected in 2017. The percent relative abundances of the top ten taxa recovered from each material from all sampling dates (August, September, and October) and depths (3, 4, and 5 m) surveyed were averaged and are displayed. All other taxa (406 ASVs with less than 0.2% relative abundance) are grouped as “Other”...... 115

Figure 6-3: Hydrocarbon biodegradation profiles from aerobic microcosms with (A) and without Tenax-TA (B) as well as no electron acceptor (EA) added control microcosms with (C) and without Tenax-TA (D). Toluene and benzene were both degraded in aerobic microcosms within 14 days; no EA added microcosms required an initial lag of 40 days for toluene degradation to occur after which it was degraded within 14-20 days. Benzene was not degraded during the monitoring period. Gaps in the plots represent depletion and re-amendment of hydrocarbons. Live and heat-killed controls (HK) were established. Error bars depict the standard error of the mean of 3 replicates...... 118

Figure 6-4: Top five most abundant microorganisms from each hydrocarbon-amended, Tenax- TA pouch-containing microcosms by treatment as determined using 16S rRNA gene amplicon sequencing. Total reads of three replicates were averaged and are displayed as percent relative abundance. Taxa that did not make the top five are grouped as “Other”. Sessile samples from the no electron acceptor added treatment could not be amplified through PCR and thus are not included...... 121

Figure 6-5: NMDS analysis of 88 microbial communities (microcosms) as analyzed by 16S rRNA gene sequencing. Hydrocarbon-amended treatments are denoted by closed circles (●) while unamended treatments are indicated by closed triangles (▲). Analyses were completed in R (scripts in Appendix D)...... 122

Figure 6-6: Microbially-colonized Tenax-TA. Panels A-C: false coloured scanning electron micrographs of microbe-colonized Tenax-TA beads retrieved from aerobic incubations. xv

(A) The sterile control, (B) the live, hydrocarbon-free control, and (C) the live, hydrocarbon-amended treatment. Tenax-TA beads in panel C has visible microorganisms adhering to its surface while beads in panels A and B show little to no microbial colonization. Panels D and E: Tenax-TA pouches recovered from aerobic microcosms after 80 days of incubation. (D) A Tenax-TA pouch from a live, hydrocarbon-free microcosm and (E) a Tenax-TA pouch from a hydrocarbon-amended treatment. The Tenax-TA pouch from the hydrocarbon treatment is visibly darkened, presumably with biomass, compared to the unamended treatment...... 123

Figure A-1: TOLDC broth culture following 2 years of consistent, semi-monthly feeding with toluene and supplementation with Balch vitamins. When undisturbed for at least three days, cells were found to settle and concentrate into large distinct clumps. Vigorous shaking would break apart these clumps, but after a few days they were observed to form again...... 160

Figure A-2: Benzoic acid detected in phase 2 incubations, error bars ± SEM, n=3...... 160

Figure A-3: (A) Electrophoresis gel of bssA amplified in randomly selected qPCR reactions, with a positive control bssA amplicon denoted by a white arrow. (B) Melt peaks from qPCR standards (blue traces) compared to “unknown” samples assayed (green traces), all with melt peak temperatures of 86-87˚C. The no template control (red trace) did not amplify...... 161

Figure A-4: Benzoic acid production from phase 3 incubations, error bars ± SEM, n=3...... 162

Figure B-1: Hydrocarbons measured in abiotic sorption tests one day after addition. Toluene sorption to Tenax-TA represented 77% of available hydrocarbons (without Tenax-TA), while benzene sorption represented 53%. Asterisks represent statistically significant differences as calculated by t-tests (** p-value ≤ 0.01, *** p-value ≤ 0.001)...... 177

Figure B-2: Design of experimental microcosms. Glass serum bottles were sealed with the Tenax-TA filled pouch suspended into the aqueous phase. Groundwater was added; associated sand settled to the bottom over time. Aerobic treatments received air as the headspace while anoxic treatments were flushed with a headspace of N2 gas...... 177

Figure B-3: Toluene and benzene degradation profiles in aerobic (panels A and B), nitrate- reducing (C and D), iron(III)-reducing (E and F), sulfate-reducing (G and H), and no electron acceptor-added (I and J) microcosms, respectively, over 80 days of incubation in the presence and absence of Tenax-TA. Both live incubations and heat-killed controls (HK) were established. Error bars represent the standard error of the mean of 3 replicates. Breaks in plot lines represent re-amendment of hydrocarbons after a time point where all previously added hydrocarbons had been consumed...... 179

was below detection limits in iron(III)-reducing microcosms regardless of the presence of hydrocarbons. Error bars represent the standard error of the mean of 3 replicates. Asterisks represent statistically significant differences as calculated by a t-test (** p-value ≤ 0.01)...... 179

Figure B-5: Microbial community composition of aerobic microcosms including the groundwater used as the inoculum as well as all treatments and controls. Samples were analyzed through 16S rRNA gene sequencing in triplicate and the averages of that analysis are shown here. Hydrocarbon-amended samples with Tenax-TA (TA) have already been discussed in detail in Figure 4. In the planktonic hydrocarbon-unamended treatments (no HCs), Candidatus Nitrocosmicus was found to be the most abundant...... 180

Figure B-6: Microbial community composition of nitrate-reducing microcosms including the groundwater used as the inoculum as well as all treatments and controls. Samples were analyzed through 16S rRNA gene sequencing in triplicate and the averages of that analysis are shown here. Hydrocarbon-amended samples with Tenax-TA were dominated by Azoarcus and have already been discussed in detail in Figure 4. Hydrocarbon- unamended planktonic communities were dominated by Rhodoferax and Candidatus Roizmanbacteria, while sessile communities were incredibly diverse, with 66% the total relative abundance comprising a variety of taxa each making up less than 0.2% of the overall community...... 181

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List of Symbols, Abbreviations and Nomenclature

Symbol/Abbreviations Definition

16S rRNA 16S subunit of ribosomal RNA AE Activating enzyme ASV Amplicon sequence variant BCR Benzoyl-CoA reductase BLASTn Basic local alignment search tool – nucleotide database bp Base pairs BSS Benzylsuccinate synthase bssA Benzylsuccinate synthase gene, α-subunit BSTFA N, O-bis(trimethylsilyl)trifluoroacetamide BTEX Benzene, toluene, ethylbenzene, and xylenes cDNA Complementary DNA CoA Coenzyme A DCM Dichloromethane DSMZ German Collection of Microorganisms and Cell Cultures GmbH EBD Ethylbenzene dehydrogenase GC-FID Gas chromatography with flame ionization detection GC-MS Gas chromatography-mass spectrometry GC-TCD Gas chromatography with thermal conductivity detection GRE Glycyl radical enzyme GRE-AE Glycyl radical enzyme-activating enzyme HMN 2,2,4,4,6,8,8-heptamethylnonane HPLC High-performance liquid chromatography m/z Mass to charge ratio NCBI National Center for Biotechnology Information NMDS Non-metric multidimensional scaling NTC No template control PAH Polycyclic aromatic hydrocarbon PCR Polymerase chain reaction qPCR Quantitative PCR RT Reverse transcriptase SEM Standard error of the mean Tenax-TA Tenax®-trapping agent TMS Trimethylsilyl TOLDC Toluene-degrading methanogenic enrichment culture XSS X-succinate synthase

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Chapter One: Introduction

For this thesis, I conducted research to further knowledge in the field of aromatic hydrocarbon biodegradation by methanogenic and groundwater-associated microbial communities. The results of these experiments advance the understanding of microbial metabolism and hydrocarbon degradation under some of the most extremely energy-limited conditions known.

1.1 Research objectives

I. Characterize the initial activation metabolites associated with methanogenic aromatic

hydrocarbon biodegradation (toluene, ethylbenzene, xylenes, naphthalene, 2-

methylnaphthalene, and phenanthrene).

Hypothesis: Alkyl-substituted aromatic hydrocarbons will be activated through fumarate addition mechanisms in a model methanogenic culture.

II. Quantify changes in real-time expression of key hydrocarbon-degradation genes when

different substrates are metabolized.

Hypothesis: Transcription of hydrocarbon biodegradation genes (such as bssA) will be upregulated when the methanogenic culture is growing on its preferred substate and not transcribed or downregulated on non-target substrates.

III. Identify key microorganisms and link their activity to hydrocarbon degradation observed

in experiments.

Hypothesis: Microorganisms carrying out biodegradation will be found in higher relative abundance in hydrocarbon-amended treatments compared to unamended treatments.

IV. Examine the features and feasibility of implementing a passive sorbent trap for monitoring

in situ bioremediation. 1

Hypothesis: The sorbent trap material will not influence the rate of intrinsic bioremediation and will not significantly alter the microbial community structure of associated microorganisms.

1.2 Thesis structure and summary

This thesis is separated into three main sections: introductory content, research chapters, and conclusions. The introductory chapters (1-3) give an overview of the content herein, the current knowledge and literature associated with microbial hydrocarbon biodegradation, and a general summary of the common materials and methods used throughout this work. In the research chapters (4-6) several aspects of methanogenic mono- and polycyclic aromatic hydrocarbon biodegradation that pertain to objectives I through III were explored, while in Chapter Six research outcomes addressing objectives III and IV are presented as a version of a manuscript recently published in the peer-reviewed journal Microorganisms. Supplemental data relating to Chapters

Four and Six are included in Appendices A and B.

Chapter Two is a review of current literature and describes existing knowledge of the field of hydrocarbon biodegradation, specifically focused on the enzymatic functionalization necessary for degradation of aromatic hydrocarbons. Techniques that were employed to examine a spectrum of features of hydrocarbon biodegradation are described in Chapter Three, including consumption of hydrocarbon substrates, production of metabolites and intermediates, mineralization of electron acceptors, production of metabolic end-products, as well as analyzing changes in microbial community structure through both 16S rRNA gene sequencing and diversity metrics. Experiments conducted and presented in Chapter Four were designed to explore the activation processes and pathways of benzene, toluene, ethylbenzene, and xylenes under methanogenic conditions. In

Chapter Five, I investigated the biodegradation pathways and intermediates of different polycyclic aromatic hydrocarbons (PAHs) by a methanogenic consortium. The research described in Chapter 2

Six involved the characterization and implementation of a sorbent sampling device to monitor in situ bioremediation of aromatic hydrocarbons in contaminated groundwater. Chapter Seven provides a summary of the key findings and conclusions gleaned from the experimental chapters; it also outlines future directions and investigations that would be useful to the field of microbial hydrocarbon degradation.

The research conducted throughout this thesis is my own work product, apart from some experiments described in Chapter Six which integrated previous work initiated by Dr. Courtney

Toth and was done in collaboration with Dr. Victoria Collins and Dr. Paolo Mussone of the

Applied Bio/Nanotechnology Industrial Research Group at the Northern Alberta Institute of

Technology. Not included in this thesis but also part of my work product were my contributions as a co-author on two publications which examined the use of microbial fuel cell technology in the remediation of phenanthrene and the optimization of anode material design to improve bioelectrical energy recovery (Appendices E and F).

Chapter Two: Literature Review

2.1 Hydrocarbons in the environment

Hydrocarbon contamination of pristine areas is one of the leading environmental concerns of the 21st century. Anticipated effects associated with failures in oil production equipment such as pipeline leaks, tanker disasters, and contamination of water sources have largely influenced legislation and public policies surrounding the expansion of oil-related industries over the past several decades. With increased use of petroleum products—not just as fuels but also as components of plastics, asphalt, motor oil, cosmetics, paraffin wax, fertilizers, and other products—hydrocarbons have inexorably become a part of daily modern life.

Oil industry operations take place in a variety of environments, including drilling in oceans, hydraulic fracturing in prairies, surface mining in forests, and underground storage in urban settings (Foght et al., 2017; Ossai et al., 2020). Due to the global nature of fossil fuel production and use, any of these diverse environments can be threatened by the potential of hydrocarbon release. Accidental spillage can occur at any point during extraction, refinement, transportation, or storage. The effects of these spills are often underestimated in their extent, as diffusion, sorption, and evaporation can account for rapid loss of the free source contaminant (Lueders, 2017;

Speight and El-Gendy, 2018).

Petrochemicals are harmful to the environment not only through their accidental release but also through their combustion, a process that releases billions of tons of carbon dioxide into the atmosphere every year (Abas et al., 2015). Carbon dioxide and methane—an important fossil fuel- related hydrocarbon—are the two most important greenhouse gases that contribute to anthropogenic climate change through the absorption of solar energy in the atmosphere and increasing radiative forcing effects (Saunois et al., 2016). Efficiently curbing emissions from 4

hydrocarbon combustion is a major modern scientific obstacle which is being investigated by scientists of many disciplines and perspectives around the world.

The impact of hydrocarbons on the environment is complex and of wide-reaching concern.

Technological advances in alternative energies are occurring at a rapid rate, however, they have not yet overcome our reliance on fossil fuels and related products. At current global crude oil production rates of roughly 30 billion barrels per year, there remains a threat of petroleum release related to hydrocarbon extraction and use (EIA, 2019; OPEC, 2019).

2.2 Origins of crude oil

Formed over geological time, fossil fuels are primarily found in the subsurface, originating from the decomposition and chemical transformation of buried organic matter. Through the processes of diagenesis, catagenesis, and metagenesis, complex macromolecules from organic matter decompose to varying degrees (Gray et al., 2010; Speight and El-Gendy, 2018).

Condensation of these breakdown products, called kerogen, and association with mineral components results in the formation of oil shale (Widdel and Rabus, 2001; Speight and El-Gendy,

2018). Following millions of years at high pressure and temperature, kerogen matures and breaks down further to form petroleum hydrocarbons, notably crude oil and natural gas. Depending on the source material, nitrogen, sulfur, and oxygen-containing compounds may also be found in these deposits. Migration of hydrocarbons through porous geological features of the subsurface results in hydrocarbon concentration in areas where impermeable rock blocks further flow, forming oil reservoirs (Head et al., 2003).

2.3 Chemical and physical properties of petroleum hydrocarbons

Hydrocarbons are, by definition, compounds consisting of hydrogen and carbon. The simplest hydrocarbon is methane, with a single carbon atom surrounded by four hydrogen atoms 5

in a tetrahedral arrangement. Structures include alkanes (single bonds connecting adjacent carbons), alkenes (double bonds), alkynes (triple bonds), as well as cyclic compounds such as cycloalkanes (single bonds only), aromatics (delocalized electrons resulting in resonance bonds), resins, and asphaltenes (large, polar, aromatic structures with limited solubility) (Jones and

Fleming, 2010; Ossai et al., 2020). Hydrocarbons are hydrophobic, non-polar compounds which results in them having poor water solubility. Light fraction alkanes (C1 to C4) are volatile at standard pressure and temperature, as they have high vapor pressures and naturally occur in a gaseous state. Alkanes with 5 to 8 carbons are considered moderately volatile, in that they mainly exist as a liquid, have comparatively low vapor pressures, but can evaporate and condense within a closed system. The boiling point (or range) of various hydrocarbons can be exploited during the refining process of crude oil, whereby hydrocarbons can be distilled or separated into fractions for downstream use (Speight and El-Gendy, 2018). Vapor pressure, which is the defining characteristic for predicting volatility and boiling points, tends to decrease as molecular weight increases (Eastcott et al., 1988).

2.3.1 Features of aromatic hydrocarbons

Aromatic hydrocarbons display a broad range of chemical and physical properties.

Monoaromatic hydrocarbons such as benzene, toluene, ethylbenzene, and xylenes (BTEX; Figure

2-1) are liquid under ambient temperatures and pressures but will rapidly volatilize in an open system (Brown et al., 2017). Volatilization is directly correlated with vapor pressure and dissolution rates (governed by the water solubility of a given chemical), and thus can be described and calculated by Henry’s Law (Zwolinski and Wilhoit, 1971; Eastcott et al., 1988). Benzene is the most water soluble of the BTEX chemicals, which is one reason it is so toxic in aquatic systems

(Headley et al., 2001; Johnson et al., 2003). 6

Figure 2-1: Structures of common monoaromatic hydrocarbons including (from left to right) benzene, toluene, ethylbenzene, o-xylene, m-xylene, and p-xylene (BTEX).

In contrast to BTEX hydrocarbons, polycyclic aromatic hydrocarbons (PAHs) are generally solid at room temperature and largely insoluble in water (less than 1 mg/L). The prevalence of aromatics in petroleum deposits is directly related to the level of degradation of the oil—heavy oil has a much larger proportion of PAHs than light oil. Heavy oil, in part due to the abundance of these large aromatics, is much more recalcitrant due to its low biodegradability (Brown et al.,

2017).

Aromatic hydrocarbons are generally considered chemically unreactive due to their resonance stability (Rabus et al., 2016a). Delocalization of electrons between overlapping orbitals evenly spreads the electron density both above and below the planar ring, resulting in the high energy of activation of benzene and similar aromatic molecules (Jones and Fleming, 2010).

Substituent alkyl groups marginally increase the reactivity of the TEX compounds because they are electron donating, and as such are considered “moderately” biodegradable compared to benzene. While there are thousands of aromatic compounds that may be found in petroleum, this work will focus on the BTEX hydrocarbons and specific PAHs including naphthalene, 2- methylnaphthalene, and phenanthrene.

2.4 Hydrocarbon biodegradation

The activities of bacteria and archaea can have numerous effects on crude oil biodegradation

(an economically detrimental process) and the bioremediation of oil (environmentally beneficial).

These impacts even now are not fully understood, as various external factors including temperature, oxygen availability, and geochemistry play a role microbial growth and metabolism.

In petroleum deposits, microbes preferentially degrade small, light hydrocarbons before more complex substrates like aromatics; this results in heavy oil formation which is less valuable, more viscous, has higher sulfur content, and is harder to obtain and process for market (Head, 2017;

Speight and El-Gendy, 2018). Conversely, this process is desirable when fuels are accidentally released during midstream and downstream processing in order to facilitate proper remediation and reclaim the affected environment. Microbial biodegradation is the only self-sustaining mechanism of natural attenuation used in hydrocarbon remediation efforts (Lueders, 2017).

Microorganisms play a central role in carbon cycling over geological time (Fuchs et al.,

2011). The Earth’s subsurface is rich with petroleum deposits and is home to microbes with unique metabolic capabilities. Prokaryotes can subsist on chemicals that would be toxic or fatal to larger multicellular organisms (Ward et al., 2009), and their metabolism is orders of magnitude more diverse than that of eukaryotes. Subsurface-associated microorganisms have several important adaptations and metabolic capabilities that allow them to succeed. As subsurface environments are usually depleted in oxygen, microorganisms have the ability to use alternative electron acceptors such as nitrate, manganese(IV), iron(III), sulfate, and even carbon dioxide for their metabolism (Boll et al., 2014; Meckenstock et al., 2015). In addition, the oil reservoir environment can include generally unfavorable conditions for growth including high temperature, salinity, pressure, low pH, and low water availability (Pannekens et al., 2019). Conditions in a 8

contaminated aquifer produce a different set of challenges; while there is high water availability, temperatures are often low, carbon sources are generally scarce, and electron acceptors are often depleted. Microorganisms living in these environments must therefore be adapted to such conditions, as well as be metabolically flexible as conditions change over time.

Water availability is a critical factor in facilitating hydrocarbon biodegradation in a reservoir environment. Most microbial activity is believed to occur at the oil-water transition zone, where microbes are in contact with both the hydrocarbon substrate and the aqueous environment

(Korenblum et al., 2012; Head, 2017; Speight and El-Gendy, 2018; Pannekens et al., 2019). In low water reservoirs (1-5% water content), biodegradation is generally slow due to the lack of microbial activity. In a comparison study, Korenblum et al. (2012) found that high water content positively correlated with increased microbial diversity, with communities from high water content reservoirs (40-60%) displaying up to 2.6 times more species richness than those with low water content. Meckenstock et al. (2014) found evidence that microscopic water droplets (1-3 μL) emulsified in oil within low-water reservoirs were home to microorganisms actively degrading hydrocarbons away from the oil-water interface. Thin films of water on the surface or within the pore spaces of rock are microhabitats where microorganisms may live and utilize hydrocarbons.

Additionally, some microbes can adhere to hydrophobic surfaces to increase the bioavailability of otherwise water-insoluble hydrocarbons through the process of interfacial accession (Bouchez-

Naïtali et al., 1999).

Studies of contamination plumes in groundwater describe a predictable and sequential degradation process known as the redox zonation effect (Figure 2-2). This involves rapid recruitment and competition of aerobic and nitrate-reducing microbes on the fringes or leading edge of a new contamination plume, followed by iron(III)-reducing, sulfate-reducing, and finally 9

methanogenic biodegradation as electron acceptors of higher reducing strengths and small, easily biodegraded hydrocarbons are depleted (Meckenstock et al., 2015). In cases of point source spills such as these, the rapid influx of available carbon substrates (with 20% or more aromatic hydrocarbon composition) causes a massive population boom among the indigenous microorganisms followed by a slower dying-off event as essential nutrients are exhausted until only the hardiest and metabolically flexible organisms remain. This gradient effect may be horizontal or vertical through the system.

While it has been well established that many types of microbial metabolisms are active in hydrocarbon-contaminated environments, they differ greatly in terms of the microorganisms 10

involved, the mechanisms employed, and the energetics of degradation. The electron accepting process is the largest predictor of the hydrocarbon degradation mechanism of a given microbe or community of microorganisms. Many studies have examined how different groups of microorganisms biodegrade hydrocarbons on a molecular level, however there is still some debate as to the exact mechanisms employed in degradation of complex hydrocarbons (like aromatics) in extremely low energy or methanogenic environments.

2.5 Aerobic biodegradation of aromatic hydrocarbons

Due to the inherent stability of aromatic hydrocarbons, all microbial degradation processes are known to require an initial activation step to increase the reactivity of the hydrocarbon before further degradation can occur. Aerobic microorganisms are known to use molecular oxygen and oxygenases to directly attack the aromatic ring of both mono- and polycyclic aromatic hydrocarbons (Speight and El-Gendy, 2018). In the case of toluene, a model aromatic hydrocarbon, oxygenases add hydroxyl groups to either of the ortho, meta, or para positions (one atom at a time by monooxygenases or two adjacent carbon atoms at the same time for dioxygenases; Figure 2-3) of the aromatic ring to form a diol intermediate, followed by dehydrogenation to form a methylcatechol compound (Parales et al., 2008; Philipp and Schink,

2012; Boll et al., 2014; Dobslaw and Engesser, 2015). Cleavage of the catechol molecule between the two hydroxyl groups then yields a muconate intermediate before further metabolism through the tricarboxylic acid (TCA) cycle (Parales et al., 2008; Vogt et al., 2011). Dioxygenase activity has been identified in aerobic microorganisms such as Pseudomonas putida F1 and Burkholderia fungorum FLU100, but not in anaerobic microorganisms (Parales et al., 2008; Dobslaw and

Engesser, 2015).

Aerobic toluene degradation genes in some microorganisms are encoded on the TOL plasmid, as is the case of P. putida mt-2 (Assinder and Williams, 1990; Parales et al., 2008). As these plasmid-encoded genes have high homology to chromosomally encoded toluene degradation genes in other microorganisms, this may represent a case of horizontal gene transfer and subsequent adaptation of previously non-hydrocarbon degrading microorganisms to this carbon source. In contrast to other types of oxygenases, TOL plasmid oxygenases appear to selectively hydroxylate the methyl carbon of toluene instead of the aromatic ring to form benzyl alcohol

(Figure 2-3 scheme 1; Parales et al., 2008). This unusual strategy is not well understood, but may indicate an adaptation to a broader range of substrates, as P. putida mt-2 has been shown to also degrade 1,2,4-trimethylbenzene, 3-ethyltoluene, m- and p-xylene (Kunz and Chapman, 1981). It has also been reported that TOL-encoded and chromosome-encoded oxygenases can be active synergistically in the same organism (Kunz and Chapman, 1981).

Additional aromatic rings present further challenges to biological degradation. PAHs are especially toxic to mammals and fish (Canadian Environmental Protection Act: Polycyclic aromatic hydrocarbons, 1994; Abdel-Shafy and Mansour, 2016). The mechanism of toxicity in eukaryotic cells involves the activity of aerobic cytochromes (such as cytochrome P450) which have monooxygenase activity on PAHs once they enter the cell (as PAHs are generally fat soluble, they often accumulate in adipose tissue), causing epoxidation, hydration, and hydrogenation of a double bond of the PAH to form a diol-like intermediate (Xue and Warshawsky, 2005; Jones and

Fleming, 2010). This PAH-diol compound can then hydrogen bond with guanine, leading this bulky molecule to impair DNA replication and transcription, resulting in mutations and potential carcinogenicity (Jones and Fleming, 2010; Abdel-Shafy and Mansour, 2016).

A major hindrance to the biodegradation of PAHs is the limited bioavailability of these hydrocarbons. As these compounds are largely insoluble in water and tend to adsorb to soil particles (particularly PAHs with four or more rings), they are physically separated from bacterial enzymes which exist in an aqueous environment (Speight and El-Gendy, 2018). PAHs also vary widely in their structure and this influences their relative biodegradability. PAHs in a straight conformation (such as anthracene) are more thermodynamically stable than angular arrangements

(like phenanthrene), as the open areas or “bay regions” increase the hydrocarbon’s susceptibility to enzymatic attack (Abdel-Shafy and Mansour, 2016). Temperature is also believed to be an important factor regulating PAH biodegradation. In a study of a mixture of PAHs, Eriksson et al.

(2003) found that degradation of 3-ringed PAH or larger was mostly inhibited at temperatures of

7˚C or less. Despite the abundance of psychrophilic or psychrotolerant microbes in low 13

temperature contaminated environments, the minimal energetics of the system and low solubility of PAHs at these temperatures becomes increasingly difficult to overcome.

2.6 Anaerobic biodegradation of toluene

Anaerobic hydrocarbon biodegradation is a slow process compared to aerobic biodegradation, however it likely accounts for more hydrocarbon attenuation than aerobic metabolism in contaminated environments and the subsurface (Gray et al., 2010; Speight and El-

Gendy, 2018). Oxygen is generally depleted within the first few metres of soil depth due to microbial activity, however this layer can be even more shallow in water-saturated soils (Lueders,

2017). Most microorganisms capable of anaerobic degradation are found in the subsurface, associating with oil deposits, groundwater aquifers, and soil particles (Pannekens et al., 2019). In the deep biosphere, oxygen and nitrate are believed to be completely absent unless anthropogenically added (Youssef et al., 2009). In addition, solid iron is not believed to be a major electron acceptor as it has been reduced over millions of years of microbial activity and is difficult to replenish (Pannekens et al., 2019). The dominant processes in subsurface environments therefore are those involving marginal energy transformations such as fermentation, sulfate reduction, and methanogenesis (Head, 2017; Pannekens et al., 2019). Despite methanogenic hydrocarbon biodegradation being a slower process than those previously described, it may have a competitive advantage in these depleted environments compared to other electron accepting processes because it does not require input or replenishment of an external electron acceptor (Gray et al., 2010; Jiménez et al., 2016).

Two main groups are involved in the anaerobic biodegradation of hydrocarbons: heterotrophic bacteria, usually of the Firmicutes or Bacteroidetes phyla that have a broad substrate range; and archaea, which often display lithotrophic or organotrophic metabolisms (Leng et al., 14

2018; Dong et al., 2019). In a hydrocarbon-containing environment, bacteria are responsible for the hydrolysis of polymers and large hydrocarbons into smaller compounds which then may be consumed wholly by a single bacterium such as Pseudomonas thivervalensis MAH1 or Thauera aromatica, or passed to acetogenic bacteria (such as members of the Firmicutes, Bacteroidetes,

Proteobacteria, Chloroflexi, or Actinobacteria phyla) in a consortium (Biegert et al., 1996; Qu et al., 2015; Leng et al., 2018). Shared electron donation and accepting relationships in these systems sustain many bacterial taxa that would otherwise not survive. Archaea such as methanogens in these environments consume by-products of the acetogenic bacteria, usually acetate, formate, or

H2 (Leng et al., 2018; Dong et al., 2019); these compounds are then used in conjunction with CO2 as a terminal electron acceptor to produce methane. Methanogens thrive in this environment due to their syntrophic relationship with bacteria.

Several studies have indicated that the dominant lifestyle in subsurface anoxic environments is that of a biofilm (Smith et al., 2018; Flemming and Wuertz, 2019; Pannekens et al., 2019). A sessile lifestyle conveys several advantages to the biofilm-differentiated microorganisms such as increased resistance to external stressors (e.g., changing geochemical conditions), retention and absorption of water, and evolutionary advantages such as shared genetic elements (Smith et al.,

2018). There is also an aspect of sharing metabolites and intermediates to the benefit of the entire community, a process known as syntrophy. Due to the low energy yields in these systems, recycling and using every by-product is essential to survival. Living in a biofilm allows these individual microbes the close contact necessary to share these essential metabolites and support the entire community.

2.6.1 Hydrocarbon addition to fumarate (fumarate addition)

The mechanism by which anaerobic microorganisms degrade the monoaromatic hydrocarbon toluene has been studied extensively in the past 25 years (Biegert et al., 1996; Beller and Edwards, 2000; Kühner et al., 2005; Winderl et al., 2007; Fowler et al., 2012; Funk et al.,

2015). In 1996, Biegert and colleagues discovered benzylsuccinate synthase (BSS) activity in cell- free extracts of Thauera aromatica, a nitrate-reducing, toluene-degrading bacterium. Since then

BSS, or evidence of its activity in the form of its product benzylsuccinate, have been found in many other pure cultures of hydrocarbon-degrading microorganisms (Beller and Spormann, 1997;

Leuthner et al., 1998; Kane et al., 2002) as well as mixed consortia (Gieg et al., 1999; Beller and

Edwards, 2000; Washer and Edwards, 2007) under all anaerobic electron accepting conditions.

Toluene degradation by anaerobic bacteria (both by pure culture or by a consortium) involves first functionalizing the hydrocarbon by the addition of fumarate across the double bond to the alkyl side chain of toluene by benzylsuccinate synthase, as shown in Figure 2-4. The resulting activated product, benzylsuccinate, is unique to this process and is therefore considered a signature metabolite. Benzylsuccinate then undergoes a CoA-transferase reaction, dehydrogenation, hydration, and finally a thiolase reaction to form the central intermediate benzoyl-CoA (Michas et al., 2017). Benzoyl-CoA is metabolized through β-oxidation to form acetyl-CoA, which is later used to generate ATP. Additionally, some bacteria such as Syntrophus aciditrophicus make use of a reversible benzoate pathway where ATP is used to functionalize benzoate to benzoyl-CoA

(McInerney et al., 2007). This intermediate is then oxidized to crotonyl-CoA via a catabolic mechanism, which can also function biosynthetically to form benzoate from crotonate.

– O SCoA SCoA

O O O O O O O O SCoA

– – – BSS O O O

a c d e f O – O – O b O

Figure 2-4: Generalized reaction of toluene (a) with fumarate (b) mediated by benzylsuccinate synthase (BSS) to form benzylsuccinate (c). Benzylsuccinyl-CoA (d) is formed through the activity of succinyl-CoA:(R)-benzylsuccinate CoA-transferase. A series of hydratase and dehydrogenase reactions results in formation of benzoylsuccinyl-CoA (e). Benzoylsuccinyl-CoA thiolase cleaves succinyl-CoA (to reform fumarate in the cycle) from benzoyl-CoA (f). Benzoyl- CoA is then used by fermentative microorganisms such as Desulfovibrio or Syntrophus via the β- oxidation pathway. Summarized from Biegert et al. (1996), Boll et al. (2014), and Michas et al. (2017).

2.6.2 Benzylsuccinate synthase

Benzylsuccinate synthase is a glycyl radical enzyme (GRE), a family of enzymes that are unique to strictly anaerobic bacteria (Li et al., 2009). Six subunits make up the quaternary protein structure: 2 units of each α, β, and γ in a heterohexamer formation (Funk et al., 2014). The α- subunit encodes the enzyme’s active site, which is broadly believed to be the most conserved region of the enzyme across different bacterial groups. The gene encoding this subunit, bssA, is the primary target of molecular-based assays to identify putative toluene degraders (Winderl et al.,

2007; von Netzer et al., 2013; Fowler et al., 2014; Kharey et al., 2020). Knockout mutants of bssA are unable to grow on toluene or xylene but can metabolize benzoate (Achong et al., 2001).

Priming of the active site is mediated by a GRE-activating enzyme (AE) encoded by bssD (Achong

et al., 2001; Widdel and Rabus, 2001; Hermuth et al., 2002). The β-subunit, encoded by bssB, contains several iron-sulfur clusters and is believed to act both as a binding domain for the AE and also to cause conformational change to the α-subunit to expose the active site when the AE is bound (Funk et al., 2014). The γ-subunit (encoded by bssC) is less well understood, but it is believed to stabilize the conformational state and folding of the α-subunit due to hydrophobic and

Fe-S interactions (Widdel and Rabus, 2001; Li et al., 2009). The final component of the operon, bssE, functions as a chaperone for assembly and activation of the BSS superstructure (Hermuth et al., 2002).

The mechanism of hydrocarbon activation by BSS appears to be conserved across microorganisms with varying metabolic capabilities. GREs broadly require a GRE-AE to generate the glycine radical via removal of a hydrogen atom in the active site of BSSα (Funk et al., 2014).

This glycine radical is then brought in close contact with a cysteine residue (Cys493) to abstract another hydrogen atom, thereby generating the catalytic cysteine-thiyl radical. The cysteine radical is subsequently sequestered in a folded barrel-shaped core to prevent its reaction with non- target substrates (Funk et al., 2014). Fumarate hydrogen bonds to carboxyl residues on BSSαγ above the cysteine radical, followed by recruitment of toluene in an adjacent hydrophobic binding pocket (Funk et al., 2015). This pocket is tightly arranged which allows for specificity of toluene binding; BSS also discriminates against dicarboxylic acids other than fumarate through steric hindrance (favoring planar molecules) in the active site. The conformation of fumarate binding exposes the C2 carbon to the alkyl group of toluene and the catalytic cysteine radical. The space separating these three constituents is only 3.4–3.9 Å, indicating the high specificity for toluene and fumarate in the active site necessitates very little movement once they are bound (Funk et al.,

2015). 18

Despite the generally accepted notion that a gene encoding a catalytic site would largely be conserved between microbial taxa carrying out the same reaction, various assays have shown this not to be the case for BSS. There is diversity in bssA sequences across different bacterial phyla; in fact, Kharey et al. (2020) found distinct sequences of bssA from BTEX-contaminated environmental samples with homology to established sequences from Firmicutes and

Proteobacteria. In a study examining bssA in a toluene-degrading methanogenic enrichment culture, Washer and Edwards (2007) had limited success with PCR using primer sets originally designed for bssA from toluene-degrading nitrate-reducers (Beller et al., 2002). In a later study of a methanogenic toluene-degrading culture (of separate origin), Fowler et al. (2012) attempted to use primers redesigned by Washer and Edwards (2007), again with limited success. This illustrates the diversity that exists in bssA sequences even across metabolically similar microorganisms and consortia. Difficulty in amplifying or detecting bssA in assays of environmental samples, which would in theory be a quick indicator of the potential for monoaromatic hydrocarbon metabolism, has led many researchers to also look for benzylsuccinate and other metabolites (a slower, more labor intensive laboratory process) as definitive evidence of active in situ hydrocarbon degradation

(Beller and Edwards, 2000; Gieg and Suflita, 2002).

2.7 Anaerobic activation of other alkylbenzenes

While the biodegradation of toluene alone has been well studied, degradation pathways of other aromatic hydrocarbons have not been as extensively characterized. Funk et al. (2014) identified a subgroup of GREs called the X-succinate synthase (XSS) group which carries out hydrocarbon addition to fumarate (herein simply referred to as fumarate addition) reactions on various substrates including 2-methylnaphthalene, p-cresol, and some alkanes such as n-hexane.

Therefore, there may be specific XSSs for the other alkyl-substituted aromatics. Despite the 19

findings of Funk et al. (2015) that the binding pocket for toluene in BSS is stereospecific, several studies have shown activity of purified BSS extracts (from toluene-grown Azoarcus spp.) on xenobiotics including xylenes, cresol isomers, fluorinated toluene derivatives, and certain alkyl- substituted cyclohexenes (Beller and Spormann, 1999; Verfürth et al., 2004). Alternatively, these hydrocarbons may be activated in a fumarate addition-independent manner. Hydroxylation, methylation, and carboxylation have been proposed as alternative activation reactions both for alkyl-substituted and unsubstituted aromatics (Foght, 2008; Meckenstock et al., 2016).

2.7.1 Ethylbenzene

Ethylbenzene is structurally similar to toluene (Figure 2-1); the addition of the methylene carbon provides steric hindrance to BSS, but may be acted on by an as to now uncharacterized ethylbenzene-specific XSS. Kniemeyer et al. (2003) and Elshahed et al. (2001) found metabolites in ethylbenzene-degrading, sulfate-reducing enrichments indicating that fumarate was added to ethylbenzene at the subterminal carbon position, forming (1-phenylethyl)succinate. This activity on the subterminal carbon is analogous to fumarate addition processes in alkanes which also target this more reactive carbon, as opposed to the terminal carbon in the case of toluene and BSS

(Speight and El-Gendy, 2018). To date, an ethylbenzene-specific XSS has not been isolated and characterized.

In addition to the known fumarate addition mechanism, a separate pathway for ethylbenzene activation also exits. Ball et al. (1996) found evidence that ethylbenzene is activated through an oxygen-independent hydroxylation reaction carried out by ethylbenzene dehydrogenase (EBD) in the denitrifying bacterium EB1. The product of this hydroxylation, 1-phenylethanol, is then sequentially transformed to acetophenone, benzoyl acetate, benzoyl acetyl-CoA, then hydrolyzed to form acetyl-CoA and benzoyl-CoA (Ball et al., 1996; Rabus et al., 2002). Benzoyl-CoA then 20

feeds into the β-oxidation pathway as previously discussed. Carboxylation and methylation are not believed to be mechanisms of activation for ethylbenzene. Supporting this were later studies by Champion et al. (1999) and Dorer et al. (2014) who found similar evidence for nitrate or iron(III) reduction linked to ethylbenzene hydroxylation. Weelink et al. (2009) suggested that pure strains of nitrate-reducing bacteria such as Azoarcus, Thauera, and Dechloromonas could hydroxylate a broad range of substrates due to their facultatively aerobic metabolism. This would imply that ethylbenzene dehydrogenases may be evolutionarily related to aerobic oxygenases in terms of function, if not origin.

Ethylbenzene dehydrogenase is believed to be a membrane-bound protein, and analysis of upstream gene clusters reveled a predicted transporter protein (Orf90) capable of aromatic and organic solvent transport (Rabus et al., 2002; Kühner et al., 2005; Weelink et al., 2010).

Ethylbenzene dehydrogenase genes (ebdABCD) have been detected in metagenomic surveys of different hydrocarbon-degrading consortia (albeit these cultures were not enriched on ethylbenzene as the primary substrate), however the presence of the gene did not always correlate with translation of EBD (Tan et al., 2015). Using a toluene-degrading methanogenic enrichment culture (examined further in Chapter Four of this work), Fowler (2014) previously surveyed for the presence of ebd, but could not find sequences with high homology to known ebd genes.

Instead, it was speculated that some type of XSS was functional (due to previous reports of alkylsuccinate metabolites under sulfate-reducing conditions) and proposed a possible requirement for co-metabolism of ethylbenzene, where it can only be degraded or biotransformed when toluene is present in at least trace amounts (Fowler, 2014).

2.7.2 Xylenes

Xylenes, which are structural analogues of ethylbenzene, would also be expected to be degraded through a fumarate addition mechanism. The stereochemistry of the secondary alkyl groups at the ortho, meta, and para positions can have large bearing on the overall biodegradability of the compound, due to the conformations recognized by enzyme active sites and the resonance stability afforded by the methyl groups (Widdel and Rabus, 2001; Foght, 2008; Weelink et al.,

2010). In the case of o-xylene, studies have shown complete transformation of this substrate by denitrifiers (Beller and Spormann, 1997), iron-reducers (Jahn et al., 2005), sulfate-reducers

(Harms et al., 1999; Morasch et al., 2004b), and methanogenic consortia (Edwards and Grbić-

Galić, 1994) to succinate derivatives indicative of fumarate addition reactions. Similarly, m- xylene has been shown to be used under the same reducing conditions (Harms et al., 1999;

Morasch et al., 2004b). It is generally accepted that meta substituted xylenes are more easily degraded than ortho substitutions, while p-xylene is the most recalcitrant (Weelink et al., 2010).

Several studies have failed to demonstrate p-xylene biodegradation despite the same cultures successfully degrading other xylene isomers, toluene, ethylbenzene, alkylbenzoates or alkylbenzenes (Harms et al., 1999; Morasch et al., 2004b). In the few studies that reported p- xylene loss, metabolites such as (4-methylbenzyl)succinate (indicative of fumarate addition) have been detected (Biegert and Fuchs, 1995; Rabus and Widdel, 1995b; Rotaru et al., 2010), while others have suggested a hydroxylation reaction on the benzylic methyl group (from evidence observed in denitrifying p-cymene degrading strains pCyN1 and pCyN2) that may occur concurrently with fumarate addition pathways (Rabus et al., 2016a). Despite this initial transformation, many of these studies proposed that a dead-end metabolite forms due to the lack of corresponding nitrate or sulfate reduction, carbon dioxide production, or steric hindrance of the 22

intermediate (Biegert and Fuchs, 1995; Harms et al., 1999; Lahme et al., 2012). However, select studies have shown stoichiometric conversion of electron acceptors indicative of complete degradation (Higashioka et al., 2012). This bottleneck arises from transformation of the intermediate 4-methylbenzoyl-CoA to 4-methylbenzoate (also known as p-toluate), due to the para position alkyl group interfering with electron transfers by benzoyl-CoA reductases (BCRs) (Rabus et al., 2016a). While BCRs work on many alkylbenzenes, it appears that para substitutions are incompatible with normal BCR functioning. Instead, microorganisms that successfully degrade p-xylene must use a unique 4-methylbenzoyl-CoA reductase similar to that employed by strains pCyN1 and pCyN2, that is as of yet uncharacterized (Rabus et al., 2016a).

2.7.3 Alkylated PAHs

Methylated PAHs add a layer of complexity to the previously described fumarate addition and hydroxylation activation processes carried out for the biodegradation of TEX hydrocarbons.

The additional rings lower the water solubility (hence potential bioavailability) and add further steric hindrance. Therefore, aromaticity must be reduced before ring opening can occur. Studies of naphthalene degradation by sulfate reducers and methanogenic consortia revealed dearomatized intermediates, where each ring was sequentially hydrogenated before ring opening occurred

(Annweiler et al., 2002; Aitken et al., 2004; Toth et al., 2018). Similar to other anaerobic processes discussed, before ring reduction can occur there must be an activation reaction. In the case of alkyl-substituted naphthalenes, fumarate addition is believed to be the initial activation process across most anaerobic conditions (Annweiler et al., 2002; Meckenstock et al., 2004; Marozava et al., 2018). XSSs such as naphthyl-2-methylsuccinate synthase have been detected in 2- methylnaphthalene-degrading cultures to produce transient activation intermediates such as naphthyl-2-methylsuccinate (Annweiler et al., 2000). While other studies imply a naphthyl- 23

methylsuccinate synthase was active, the lack of succinate derivatives detected contradicts this assumption. Instead naphthoic acids were identified and assumed to be central intermediates

(analogous to benzoate as an intermediate in toluene degradation), rather than the result of direct carboxylation reactions (Berdugo-Clavijo et al., 2012; Marozava et al., 2018).

2.8 Activation of unsubstituted aromatic hydrocarbons

Using benzene as a model unsubstituted hydrocarbon, the proposed products include phenol

(hydroxylation; Zhang et al., 2013), toluene (methylation; Ulrich et al., 2005), and benzoate

(carboxylation; Luo et al., 2014). One issue with evaluating the veracity of these proposed activation mechanisms is that these products are not uniquely anaerobic metabolites like benzylsuccinate, and can therefore be easily introduced by secondary means, aerobic biodegradation, or through natural cellular processes unrelated to hydrocarbon degradation. For example, phenol can form through abiotic exposure to oxygen during sampling (Kunapuli et al.,

2008), toluene can be introduced as a contaminant of pure benzene (Ulrich et al., 2005), and benzoate is known to be a central intermediate in the degradation of all aromatic hydrocarbons through the β-oxidation pathway. Thus, identifying the biotic origin of these intermediates is not a trivial process. In addition, there is commonality in these pathways and mechanisms; one line of evidence suggests an initial methylation reaction followed by subsequent XSS activity and formation of benzylsuccinate derivatives (Safinowski and Meckenstock, 2006). To refute this,

Musat et al. (2009) found succinate derivatives in extracts from naphthalene and 2- methylnaphthalene-grown enrichments but no evidence of naphthalene methylation as a precursor reaction. Due to the low concentration of naphthyl-2-methylsuccinate in naphthalene-amended enrichments, the authors postulated that carboxylation of naphthalene could have occurred initially, followed by a condensation reaction with succinyl-CoA to form this product. 24

Additionally, multiple pathways or activation processes may be operational at the same time.

Ulrich et al. (2005) found isotopic evidence of toluene, benzoate, and phenol as intermediate metabolites of benzene degradation in a methanogenic culture, but only toluene and benzoate in a similar nitrate-reducing experiment. From this they proposed a methylation pathway for the nitrate-reducing culture and two parallel pathways in the methanogenic culture: one involving hydroxylation, and a second involving methylation. Mancini et al. (2008) postulated that benzene may be activated through different mechanisms under nitrate-reducing conditions than methanogenic conditions due differential abundance of 13C/12C and 2H/1H enrichment factors. In a similar study, Keller et al. (2018) also found evidence of separate benzene degradation mechanisms in nitrate-reducing and sulfate-reducing experiments. In a study using a pure denitrifying bacterium, Chakraborty and Coates (2005) found that benzene was activated by an initial hydroxylation reaction to phenol followed by carboxylation to form benzoate. The conflicting findings from a range of studies leads many researchers to conclude multiple pathways may be responsible for benzene activation depending on the electron-accepting condition and the origin of the enrichment culture (Ulrich et al., 2005; Mancini et al., 2008; Keller et al., 2018).

Select studies have found non-substituted PAHs such as naphthalene may undergo a methylation reaction before further transformation through a fumarate addition process as previously discussed in the context of benzene (Safinowski and Meckenstock, 2006). Other studies have shown carboxylated intermediates without detection of corresponding alkylated or succinate derivatives, implying that non-substituted PAHs like naphthalene, anthracene, and phenanthrene may be activated through carboxylation without fumarate addition (Zhang and

Young, 1997; Annweiler et al., 2002; Berdugo-Clavijo et al., 2012; Marozava et al., 2018; Toth and Gieg, 2018; Toth et al., 2018). Due to the nanomolar concentrations of these metabolites, 25

their transient nature, slow microbial generation times, and the lack of unique signature metabolites produced, it is difficult to be certain these processes are not occurring in each of these experiments; intermediates may be produced and consumed so quickly that the monitoring methods employed simply cannot detect them.

2.9 Substrate range of hydrocarbon-biodegrading microorganisms

Many studies that have demonstrated the successful degradation of single aromatic hydrocarbons also report a somewhat broad substrate range for similarly structured aromatic substrates, including the option or requirement for co-metabolism (Chakraborty et al., 2005).

There is a distinction between microorganisms or consortia that carry out concomitant

(simultaneous) degradation of two or more substrates, versus those which use substrates sequentially (i.e., a preference for toluene but can use ethylbenzene when toluene is depleted).

Additionally, transformation of a secondary substrate may depend on the degradation of a primary substrate; this secondary substrate may be completely mineralized or partially transformed into a dead-end product (Speight and El-Gendy, 2018). Lueders (2017) suggests that co-metabolism is an important feature in environments that are depleted in substrates and electron acceptors.

Flexible metabolic capabilities are the hallmark of environmental microorganisms who must be able to adapt to changing situations in order to succeed on an evolutionary timeframe.

There are several lines of evidence that indicate co-metabolism is necessary for the degradation of hydrocarbons in certain mixtures, particularly non-target xenobiotics. Select studies have shown that in cultures enriched with toluene as the sole carbon source, xylenes and ethylbenzene can also be degraded but this only occurs when toluene is present (Rabus et al.,

2016b). Similarly, Dorer et al. (2014) found that P. putida could only degrade ethylbenzene when naphthalene was also present. Champion et al. (1999) found a denitrifying bacterium grown on 26

toluene and ethylbenzene used two independent pathways – toluene was activated by BSS while ethylbenzene used EBD. Even when both substrates were present, these pathways operated independently and produced unique metabolites. To contradict this, Evans et al. (1992) found that denitrifying bacterium T1 could completely degrade toluene, however, when o-xylene was also present, both were transformed into dead-end benzylsuccinate derivatives. While alkylbenzenes have similar physical structures, subtle differences in resonance stability, location of substituent groups, and enzymatic susceptibility lead to a range of possible reaction mechanisms that depend largely on the enrichment factors and the microorganisms present in the consortium. The effect of multiple substrate availability may therefore be synergistic or antagonistic depending on the substrates in question, the composition of the microbial community, and the metabolic capacity of that community.

2.10 Remediation technologies

The abundance of hydrocarbons in the environment and their ubiquitous use by humans leads the field of remediation to be of interest by many stakeholders, particularly in light of the demonstrated toxic effects of hydrocarbon pollution to humans, wildlife, plants, and aquatic systems (Ossai et al., 2020). Many so-called “conventional” remediation techniques have been used industrially for many years, including physical and chemical methods. Mechanical recovery involves physically removing the contaminated material either through digging (in the case of a spill on land), skimming (in the case of a spill on water), pumping, or vacuuming (Speight and El-

Gendy, 2018). Chemical methods may be combined with physical removal, as in the case of sorbents used to soak up oil for short term transport or storage to a landfill, surfactants or solvents to emulsify oil mixtures for easier skimming, and dispersants to break up floating oil slicks for dispersal into a body of water (Youssef et al., 2009; Speight and El-Gendy, 2018). These 27

techniques range greatly in their efficacy, from as little as 5% recovery in skimming efforts up to

90% removal with the use of dispersants or in situ controlled burning (Speight and El-Gendy,

2018). These practices seldom result in complete remediation of the contaminant, rather they often move contaminated material from one location to another (mechanical recovery) or create more contaminated material than was originally was present (adsorbents). Additionally, these techniques can be expensive, with estimates of up to $35,000 USD to physically remediate one ton of contaminated material (Prendergast and Gschwend, 2014).

Abiotic attenuation of hydrocarbons can involve evaporation and photolysis, both of which disproportionately affect BTEX and short-chain hydrocarbons (Logeshwaran et al., 2018).

However, both processes necessitate exposure to oxygen or light in an open system which greatly limits their scope of use as passive remediation techniques. Natural attenuation by exploiting the normal metabolic processes of indigenous microorganisms is a sustainable, relatively cheap, and non-invasive option (Johnson et al., 2003). This approach generally involves intrinsic bioremediation, wherein natural microbial processes occur while the site is periodically assessed for decreasing contaminant or electron acceptor concentrations, increasing microbial byproducts like CO2, and for common biomarkers indicative of microbial degradation such as benzylsuccinate

(Weelink et al., 2010). Bioaugmentation may also be used, which involves the addition of nutrients that are often limited (such as nitrogen and phosphorus) to supplement the native microbial community and facilitate hydrocarbon degradation. Additionally, these microorganisms can contribute to remediation not just through degradation of the hydrocarbons themselves, but also in the production of various intermediates and byproducts such as biosurfactants and biopolymers that change the physical state or neutralize the contaminant (Ossai et al., 2020).

Biosparging is sometimes employed, where oxygen or air is injected to stimulate aerobic 28

biodegradation. This technique can be problematic, particularly when used to remediate contaminated groundwater as oxygen is minimally soluble (at 25˚C, O2 saturation in freshwater is approximately 8 mg/L; Miura et al., 2015) and oxygen is rapidly depleted through microbial activity (Logeshwaran et al., 2018). Despite these challenges, some successful studies have achieved up to 70% BTEX removal from contaminated groundwater after 10 months of treatment

(Kao et al., 2008).

With increased interest and use of bioremediation methods, it is more vital than ever to develop robust monitoring technologies that accurately assess not just the overall contaminant load, but that also provide evidence of specific microbial processes and active remediation. In dynamic environments such as subsurface aquifers, hydrocarbon and electron acceptor concentrations fluctuate as groundwater flows making it difficult to quantify changes over time

(Logeshwaran et al., 2018). In the case of alkane and toluene contamination, surveying environmental samples for anaerobic hydrocarbon degradation genes such as assA and bssA is an attractive and relatively quick strategy to determine if in situ biodegradation is occurring

(Bombach et al., 2010; von Netzer et al., 2013). Furthermore, signature metabolites such as alkylsuccinates and benzylsuccinates which only form transiently during anaerobic biodegradation can be detected in contaminated samples and can be conclusively linked to microbial activity

(Elshahed et al., 2001; Bombach et al., 2010; Gieg and Toth, 2017). The fumarate addition genes and metabolites associated with anaerobic alkane and toluene degradation have been thoroughly studied, detected in contaminated environments, and are generally well characterized; however, no such assays that detect degradation genes or metabolites of other hydrocarbons (BEX, PAHs, heterocycles) are used regularly. Conflicting evidence of specific degradation pathways, the

absence of extensively characterized enzymes, and the lack of signature metabolite formation are some reasons why the in situ biodegradation of these hydrocarbons is difficult to assess.

Recovering microorganisms to analyze the community structure or the presence of functional hydrocarbon degradation genes is an important aspect of monitoring natural attenuation.

This usually involves collecting groundwater from monitoring wells, however this method primarily recovers planktonic cells and therefore underrepresents the sessile microbial communities that are associated with subsurface structures (McMahon and Parnell, 2014;

Flemming and Wuertz, 2019). Obtaining core samples from the subsurface is a more ideal method of recovering sessile microorganisms, however this is expensive and logistically challenging.

Sorbent sampling devices emplaced in contaminated groundwater have been successful in capturing sessile or biofilm-associated hydrocarbon degrading microorganisms with considerably less cost than core sampling (Peacock et al., 2004; Sublette et al., 2006; Biggerstaff et al., 2007;

Busch-Harris et al., 2008). Multiple approaches are necessary to successfully demonstrate in situ natural attenuation is occurring in contaminated environments. Optimizing recovery, processing, and analysis of microbial samples will help further the use and viability of in situ bioremediation techniques.

2.11 Research needs

A clear gap in these previously discussed studies is the lack of experimentation under methanogenic conditions. While this is the least energetically favorable and slowest occurring biodegradation process, carbon cycling over geological time is often dependent on the activity of methanogenic consortia. Characterizing precisely how methanogenic microorganisms or consortia degrade recalcitrant aromatic hydrocarbons on a molecular level has been attempted by several researchers, however these experiments often require long incubations, precious enrichment 30

cultures, and have extremely slow generation times. Despite the best efforts of researchers, these methanogenic experiments often fail to achieve meaningful biodegradation or enzymatic activity within the time frame allotted for grants and thesis programs. Long-term experiments involving a variety of possible hydrocarbon substrates could more accurately reflect the conditions encountered by microorganisms in the environment and may elucidate synergistic or antagonistic effects that would not be observed in experiments when using only one carbon source.

Additionally, determining the mode of action of microorganisms and enzymes that target poorly soluble hydrocarbons (like PAHs) has larger implications not just in remediation and upgrading of heavy oil, but also potentially in the biodegradation of chemically complex petroleum-based plastic compounds such as polystyrene, phenolic resins, and polycarbonates. Finally, isolating and engineering microorganisms capable of targeted, efficient, and rapid aromatic hydrocarbon degradation, then optimizing them to operate in a range of conditions, would benefit environmentally conscious remediation efforts of petroleum spills. Uncovering the specific mechanisms and enzymes natively used by microorganisms could give rise to new biotechnologies that lessen the impact of petroleum hydrocarbons on the environment. The study of hydrocarbon biodegradation is an ever-expanding discipline with frequent new discoveries and promising implications to the broader scientific field.

Chapter Three: General Materials and Methods

This chapter describes the commonly used materials and methods used throughout this thesis. Certain chapters involve unique techniques that will be described in more detail in the appropriate section.

3.1 Microbial cultures and strains

Pure strains of Syntrophus aciditrophicus (DSM 26646), Thauera aromatica (DSM 6984), and Desulfobacula toluolica (DSMZ 7467) were used in the experiments described in Chapter

Four and were obtained from the Leibniz-Institute DSMZ (Braunschweig, Germany). For many experiments undertaken in this thesis work, a highly enriched toluene-degrading methanogenic culture (TOLDC) was used (Gieg et al., 1999; Fowler, 2014; Toth, 2017) and will be described further in Chapter Four. For experiments in Chapter Five, a new anoxic enrichment culture was established from soil contaminated with hydrocarbons, solvents, and sewage. For experiments in

Chapter Six, groundwater from a hydrocarbon-contaminated aquifer associated with a decommissioned gas station near Stony Plain, Alberta, Canada was used as the inoculum.

3.2 General cultivation of anaerobic microorganisms

3.2.1 Media recipes and procedures

Anoxic minimal freshwater medium (Pfennig medium) was prepared according to laboratory standard operating procedures, modified from McInerney et al. (1979) (recipe in Appendix C).

All anaerobic cultures were prepared in glass serum bottles, flushed with 10% CO2 in N2 gas, closed with butyl rubber stoppers (2048-11800; Bellco Glass, Vineland, USA), and sealed with aluminum crimps. All cultures were incubated without shaking at room temperature (20-22°C) in the dark otherwise noted. Culture transfers were performed with glassware, syringes and equipment flushed with 10% CO2 in N2 gas or in an anoxic glove bag containing the same gas 32

mixture. For most experiments, a 50% v/v transfer was performed unless otherwise noted.

Pelotomaculum medium for S. aciditrophicus was prepared using the above procedures, with crotonate substituted for sodium pyruvate as indicated (Appendix C).

3.2.2 Addition of BTEX substrates

Benzene, toluene, ethylbenzene, and xylenes (o-, m-, and p- isomers) (BTEX) were added from pure undiluted stocks (Sigma Aldrich, Oakville, Canada) into closed, stoppered serum bottles via a 10 µL glass syringe. Great care had to be taken when handling these hydrocarbons, as rapid volatilization and abiotic loss from closed serum bottles was common. Newly purchased butyl rubber stoppers were used in experiments to minimize abiotic hydrocarbon loss. Sterile controls were established for all experiments that contained hydrocarbons to account for any abiotic losses

(such as sorption to stoppers).

3.2.3 Addition of PAH substrates

Naphthalene, 2-methylnaphthalene, and phenanthrene (Sigma Aldrich) were used as carbon substrates for the experiments described in Chapter Five. As these PAHs are insoluble in water, they were first solubilized in an inert solvent [2,2,4,4,6,8,8-heptamethylnonane (HMN) or dichloromethane (DCM); Sigma Aldrich] for consistent and reproducible addition of milligram quantities of substrates. Solubilization of PAHs in HMN had mixed success in stimulating hydrocarbon degradation, so absorptive materials were subsequently used.

3.2.4 Adsorptive materials (Amberlite and Tenax)

Amberlite® XAD-7 polymeric beads, 20-60 mesh (Sigma Aldrich) were also used in PAH adsorption experiments (Chapter Five) as detailed by Berdugo-Clavijo et al. (2012). Amberlite was washed and prepared according to the methods outlined by Morasch et al. (2001) by first washing the beads with anhydrous ethanol five times, then with sterile distilled water five times, 33

drying under vacuum overnight, and finally baking at 90˚C for three days. The cleaned beads were stored in a tightly sealed sterile container until they were used. Adsorption of Amberlite involved distributing 0.3 g into sterile serum bottles to which a 100 μM stock of PAHs dissolved in DCM was added. DCM was evaporated before Pfennig medium was added, bottles were flushed with

10% CO2 in N2 gas, and sealed with butyl rubber stoppers and aluminum crimp seals. After autoclaving, 50% v/v culture transfers were performed. Experimental set-up is described in more detail in Chapter Five.

Tenax®-trapping agent (Tenax-TA), 60-80 mesh (Restek, Bellefonte, USA) was used as an adsorbent to trap aromatic hydrocarbons and microorganisms for the work described in Chapter

Six. Tenax-TA is a porous polymer of 2,6-diphenyl-p-phenylene oxide commonly used in air purification systems and some purge and trap analytical chemistry applications due to its ability to trap volatile organic compounds (VOCs) (Zhao and Pignatello, 2004). Tenax-TA was pre- weighed (125 ± 5 mg) and filled into nylon pouches of 60 μm pore mesh. These pouches were suspended from rubber stoppers into sealed serum bottles and autoclaved before inoculum was added (see Appendix Figure B-2 for images of the pouches in the serum bottles).

3.3 Gas chromatography-flame ionization detection (GC-FID)

3.3.1 Hydrocarbon analysis

BTEX concentrations were monitored by sampling 50 µL of headspace with a glass gas- tight syringe flushed with 10% CO2 in N2 and injecting on an Agilent gas chromatograph model

7890A with flame ionization detection (GC-FID). The analytical column used was HP-5 (30 m x

320 μm). The detector temperature was held at 250˚C, the inlet at 250˚C, and the oven at 100˚C.

Helium was used as the carrier gas. To calculate the headspace concentration of BTEX

compounds, calibration standards and constants of Henry’s Law were used (Eastcott et al., 1988;

Heath et al., 1993).

3.3.2 Methane analysis

Methane production was monitored by sampling 200 µL aliquots of headspace with a sterile plastic syringe flushed with 10% CO2 in N2 gas. Samples were immediately injected on a Hewlett

Packard GC-FID model 5890 equipped with an 80/100 Porapak R column (13156-U; Sigma

Aldrich). Oven temperature was held isothermally at 100˚C with the injector inlet at 150˚C and detector at 200˚C. Helium and nitrogen were used as the carrier and make-up gases, respectively.

3.4 Gas chromatography-thermal conductivity detection (GC-TCD)

Carbon dioxide and oxygen were monitored by sampling 100 µL of headspace with a plastic syringe and injecting on an Agilent 7890A gas chromatograph equipped with thermal conductivity detection (GC-TCD). An HP-PLOT-Q (30 m x 530 μm) column was used with a detector temperature of 200˚C, inlet of 250˚C and oven at 80˚C. Helium was used as the carrier gas.

3.5 High-performance liquid chromatography (HPLC)

3.5.1 Quantification of organic acids

Analysis of organic acids by high-perfomance liquid chromatography (HPLC) was performed on a Waters 1515 HPLC apparatus equipped with a Prevail™ Organic Acid Column

88645 (5 μm particle size, 250 x 4.6 mm). A Waters 2489 UV/Vis detector was operated at 210 nm for detection of organic acids, according to the methods described by Mouttaki et al. (2007).

Benzoate and crotonate were detected using an isocratic mobile phase of 60% 25 mM KH2PO4

(pH 2.5) and 40% acetonitrile. Acetate, propionate, and butyrate were detected using 25 mM

KH2PO4 (pH 2.5) buffer only. Samples were acidified with 1 M H3PO4 before injection. All flow rates were set to 1 mL/min. 35

3.5.2 Detection of anions

Nitrate, nitrite, and sulfate concentrations were measured using a Waters 1515 high- performance liquid chromatography ion exchange protocol with UV/Vis and conductivity detection. A Waters IC-Pak anion HC column (4.6 x 150 mm; WAT026770) was used for separations, while an acetonitrile-butanol-borate/gluconate (12%, 2% and 2% respectively) solution with a flow rate of 2 mL/min was used as the mobile phase.

3.6 Spectrophotometric analysis of iron

The ferrozine assay was used to monitor ferrous and total iron according to the method outlined by Lovley and Phillips (1987) and detected spectrophotometrically at 562 nm.

3.7 ATP assay

LifeCheck ATP test kits were supplied by OSP Microcheck (Calgary, Canada) and used according to the manufacturer’s instructions. Due to slow generation times and almost negligible doubling times for the cultures used in this thesis work (e.g., previous analyses showed TOLDC increased its total bacterial count by a factor of only 0.6 over 20 days; Fowler, 2014), ATP assays were used to monitor actual activity in cultures periodically, providing a log value of active microbes per milliliter of culture fluid.

3.8 Chemical analysis via gas chromatography-mass spectrometry (GC-MS)

3.8.1 PAH quantification

Amberlite-adsorbed PAHs including naphthalene, 2-methylnaphthalene, and phenanthrene were analyzed by acidifying samples to pH < 2, and subjecting these samples to organic extraction with dichloromethane prior to analysis according to the methods outlined by Gieg and Suflita

(2002) and Berdugo-Clavijo and Gieg (2014). These compounds were measured using an Agilent

7890A GC system coupled to a 5975C mass selective detector (GC-MS). This instrument was 36

equipped with Agilent HP-1 capillary column (1901Z-115E, 50 m x 0.32 mm x 0.52 μm), and the inlet was operated in split mode (50:1, 270˚C). The oven temperature was held at 100˚C for 5 minutes, then increased by 8˚C per minute to a final temperature of 250˚C where it was held for a further 5 minutes. Identification of PAHs of interest was achieved by comparing the retention times with authentic PAH standards of known concentration.

3.8.2 Metabolite identification

For extraction of metabolites from cultures, samples were acidified to pH 2 or less before extracting thrice with ethyl acetate and drying over anhydrous sodium sulfate (VWR, Radnor,

USA). Solvent extracts were concentrated by rotary evaporation to 50 μL and derivatized with 50

μL N, O-bis(trimethylsilyl)trifluoroacetamide (BSTFA) at 55˚C for 20 minutes before injection on the GC-MS apparatus described previously. For BTEX metabolites, the inlet was operated in split mode (20:1, 270˚C) with the oven holding at 45˚C for 5 minutes, then increasing by 4˚C per minute to 270˚C and holding for a further 5 minutes. For PAH metabolites, the protocol was the same as above with a final oven hold stage at 270˚C for 15 minutes instead of 5 minutes. Predicted metabolites, including benzylsuccinic acid (from toluene) and p-toluic acid (from p-xylene), were obtained from Sigma Aldrich and used as positive standards with known fragment ion profiles to compare and quantify metabolites from culture extracts in Chapter Four. Also sought were 1- phenylethanol (VWR) and (1-phenylethyl)succinic acid (both putative ethylbenzene metabolites), as well as 2-(4-methylbenzyl)succinic acid (from p-xylene). The putative fumarate addition metabolites [(1-phenylethyl)succinic acid and 2-(4-methylbenzyl)succinic acid] were unavailable from chemical suppliers and were therefore synthesized in-house by Dr. Courtney Toth according to the method described by Bickford et al. (1948).

3.9 DNA analysis

3.9.1 DNA extraction

Isolation of DNA from cultures and soil samples was performed using the FastDNA Spin

Kit for Soil™ (MP Biomedicals, Santa Ana, USA). DNA extractions for the experiments described in Chapter Six included an additional proteinase K digestion step after bead beating (20 mg/mL; Thermo Fisher Scientific, Waltham, USA) and 20% sodium dodecyl sulfate solution (SDS

20%; Thermo Fisher Scientific) to disrupt lipid membranes. Extracted DNA was quantified by

Qubit fluorometry (Invitrogen, Carlsbad, USA) and stored at -20˚C before use in downstream 16S rRNA gene sequencing.

3.9.2 16S rRNA gene sequencing with Illumina MiSeq

To prepare for microbial community analysis, the extracted DNA was subject to a two-step

PCR protocol to target the V6-V8 hypervariable region of the 16S rRNA gene (Toth and Gieg,

2018). The first round of PCR used the KAPA Hifi HotStart Ready Mix 2x DNA polymerase

(Roche, Basel, Switzerland) for high-fidelity binding and the Nextera-adapted primer combination

926Fi5 and 1392Ri7 (Table 3-1). Amplified products were verified for purity and size using gel electrophoresis and cleaned using the QIAquick® PCR Purification kit (Qiagen, Hilden, Germany),

Axygen® AxyPrep Magnetic Bead PCR Purification kit (Corning Inc., Union City, USA), or the

Select-A-Size DNA Clean and Concentrator™ kit (Zymo Research, Irvine, USA) depending on the quality of amplicons. Amplified products were then subject to a second round of PCR to attach the barcoded primers P5-S50X-OHAF and P7-N7XX-OHAF necessary for Illumina MiSeq®. This was done using the Thermo Scientific 2x PCR Master Mix with Taq DNA polymerase (Thermo

Fisher Scientific). Amplified products were again cleaned and checked for purity before

quantification by fluorometry. Prepared samples were sequenced using the Illumina MiSeq platform at the Alberta Children’s Hospital Research Institute (Calgary, Canada).

Primer Length Sequence Source name (bp) TCGTCGGCAGCGTCAGATGTGTATAAGAGACAG 926Fi5 52 AAACTYAAKGAATTGACGG Toth GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAG (2017) 1392Ri7 49 ACGGGCGGTGTGTRC

3.9.3 Bioinformatics processing

Amplicon data obtained from Illumina MiSeq were processed through standard bioinformatics pipelines for post-processing clean-up, quality check and annotation. Primers were removed using cutadapt (Martin, 2011), QIIME 2.0 was used to verify amplicon sequence quality

(Bolyen et al., 2018), and DADA2 was used for denoising (Callahan et al., 2016). Reads with forward reads shorter than 280 bp, and reverse reads shorter than 260 bp were removed with these programs before clustering, classifying with Naïve Bayes algorithms, and annotating as amplicon sequence variants (ASVs) against the SILVA 132 database (Quast et al., 2012).

3.9.4 Data interpretation and ecological comparisons using R

Annotated sequencing data were analyzed in R (version 3.6.2) to identify ecological relationships. Non-metric multidimensional scaling (NMDS) plots compared distance relationships between microbial communities of discrete samples using Bray-Curtis dissimilarity.

Scripts are described in Appendix D. Packages used included vegan (Oksanen et al., 2019), ape

(Paradis and Schliep, 2018), picante (Kembel et al., 2010), ggplot2 (Wickham, 2016), and phyloseq (McMurdie and Holmes, 2013).

3.10 RNA extraction, purification, and reverse transcription

Total RNA was extracted from microbial cells after bead beating using phenol-chloroform- isopropanol precipitation with PureZOL (Bio-Rad, Hercules, USA), a guanidine thiocyanate reagent. Glycogen (20 mg/mL; Thermo Fisher Scientific) was added following phase separation to help precipitate and stabilize RNA. RNA was quantified by fluorometry then purified with the

TURBO DNA-free kit (Invitrogen) and precipitated in 6 M lithium chloride to remove contaminating DNA. Purified RNA was subject to reverse transcription as soon as possible after extraction using the iScript cDNA Synthesis kit (Bio-Rad) with 2 μL RNA template and the following reaction conditions: 25˚C for 5 minutes, 46˚C for 20 minutes, and finally 95˚C for 1 minute to inactivate the reverse transcriptase. cDNA was quantified by fluorometry and stored at

-20˚C for downstream applications with time-course qPCR assays. RNA was frozen at -80˚C for long-term storage.

3.11 qPCR assays

Bioline 2x SensiFAST™ SYBR® No-ROX kit was used for qPCR analysis, with 0.8 μM of each forward and reverse primer per reaction. Multiple primer combinations and thermocycling conditions were tested for their use in targeting bssA in TOLDC. Following many trials, the following optimized thermocycling protocol for bssA was determined: 95˚C for 3 minutes, 40 cycles of 95˚C for 15 seconds and 58˚C for 25 seconds, 2 minutes of 58˚C followed by a melt curve starting at 58˚C and increasing by 0.5˚C every 5 seconds until 95˚C. Primers and their various targets and annealing temperatures are listed in Table 3-2. All qPCR assays were conducted on a Bio-Rad CFX96™ C1000 Real-Time PCR Detection System connected to CFX

Manager™ (version 3.1) software.

Table 3-2: Primer-pair sequences targeting bssA used in qPCR assays. 40

Primer Amplicon Sequence Tm (˚C) Source name size (bp) WinF CAATCCGTGGCTTCAGGTTCAT 57.7 Kharey et ~140 HitR TCCTCGTAGCCTTCCCAGTT 57.9 al. (2020)

Due to the complex, mixed nature of TOLDC, identifying a pure bssA gene amplicon was an involved process. Following many trials and attempts at using pure cultures including Thauera aromatica (DSMZ 6984) and Desulfobacula toluolica (DSMZ 7467), the primer pair WinF and

HitR were used to amplify bssA from TOLDC itself, separated by gel electrophoresis, then further gel purified with the QIAquick Gel Extraction kit (Qiagen) for its use as a standard in this qPCR assay. The purified amplicon was diluted across a range from 10-1 to 10-9 with three technical replicates for each dilution.

3.12 Scanning electron microscopy

As described in Chapter Six, aerobically incubated Tenax-TA pouches were preserved for visualization by scanning electron microscopy. Samples were first preserved with a 2.5% glutaraldehyde (Alfa Aesar, Haverhill, USA) in phosphate buffer solution. The solution was decanted and replaced with phosphate buffer three times. The buffer was again decanted, and the samples were dehydrated with increasing concentrations of ethanol. Finally, the samples were dried in hexamethyldisilazane (Sigma Aldrich) overnight. Preserved samples were loaded on carbon black, sputtered with gold and visualized with a TESCAN VEGA3 scanning electron microscope (Tescan Analytics, Fuveau, France).

3.13 Statistical analyses

Where appropriate, two-tailed unpaired t-tests with null hypothesis testing were calculated to determine statistically significant variances between means. Probability values (p-values) less

than 0.05 (5%) were considered statistically significant and are denoted by asterisks in graphs or tables.

Chapter Four: Elucidating activation metabolites of alkylbenzenes in a toluene-degrading methanogenic enrichment culture

4.1 Introduction

Toluene degradation by a range of anaerobic microorganisms has been well characterized and previously discussed in detail (section 2.6). Under anaerobic (including methanogenic) conditions, benzylsuccinate synthase has been shown to activate the alkyl carbon of toluene by adding a fumarate molecule, forming the signature metabolite benzylsuccinate (Figure 2-4).

Homologous X-succinate synthases may add a fumarate molecule to similarly substituted alkylbenzenes such as 2-methylnaphthalene and other aromatic compounds such as p-cresol (Funk et al., 2014). Hydrocarbon addition to fumarate (referred to as “fumarate addition” throughout this chapter) reactions have been characterized to occur for other alkylbenzenes, including ethylbenzene and xylenes (Beller and Spormann, 1999; Elshahed et al., 2001; Verfürth et al.,

2004). In addition, several studies have purported co-metabolism may be necessary to facilitate the biodegradation of non-target aromatic compounds (i.e. a primary substrate must be present for a secondary substrate to be metabolized or biotransformed into a dead-end product; discussed in section 2.9) (Evans et al., 1992; Champion et al., 1999; Chakraborty et al., 2005; Rabus et al.,

2016a). There are also reports that multiple hydrocarbon functionalization mechanisms can operate simultaneously in experiments with two or more substrates (Evans et al., 1992; Champion et al., 1999; Dorer et al., 2014; Rabus et al., 2016b). In nutrient or electron acceptor-depleted environments, these strategies would convey a competitive advantage to microbes capable of co- metabolism compared to those with limited substrate range (Lueders, 2017).

There are a few reported studies demonstrating methanogenic benzene, ethylbenzene or xylene degradation (Weiner and Lovley, 1998; Ulrich and Edwards, 2003; Da Silva and Alvarez,

2004; Reinhard et al., 2005). In an experiment comparing sulfate-reducing versus methanogenic

TEX biodegradation, Elshahed et al. (2001) found that toluene could only be degraded under methanogenic conditions while ethylbenzene and the xylene isomers required sulfate as the reductant. They also detected benzylsuccinate derivatives of all TEX hydrocarbons either from contaminated groundwater extracts or after enrichment on these substrates in laboratory microcosms. Other studies conducted under sulfate-reducing conditions indicated fumarate addition is indeed the mechanism for ethylbenzene activation (Elshahed et al., 2001; Kniemeyer et al., 2003; Heider, 2007) while under nitrate-reducing conditions, hydroxylation appears to be the dominant pathway (Champion et al., 1999; Johnson and Spormann, 1999; Knack et al., 2012;

Dorer et al., 2016). Carboxylation and methylation have not been reported as mechanisms of ethylbenzene activation. Ethylbenzene is most chemically reactive at the α-carbon position

(closest to the aromatic ring) due to the effect of resonance stabilization of the resulting benzylic radical after hydrogen has been abstracted (Jones and Fleming, 2010). As such, 1-phenylethanol

(from hydroxylation) and (1-phenylethyl)succinate (from fumarate addition) are hypothesized as the most likely metabolites arising from these reactions (Table 4-1). Most broad surveys of activation process across different reducing conditions assume that degradation patterns and pathways utilized by methanogenic microbial communities are largely shared by sulfate-reducing microorganisms, thus we predict the most likely product from methanogenic ethylbenzene activation is (1-phenylethyl)succinate (the product of fumarate addition).

The biodegradability of xylenes is influenced by the location of the second alkyl group, due to the conformations recognized by enzyme active sites and the resonance stability of methyl groups at certain positions (Widdel and Rabus, 2001; Foght, 2008; Weelink et al., 2010). Across numerous studies and reviews, fumarate addition is largely the accepted mechanism activating all 44

three xylene isomers regardless of the reducing condition (Foght, 2008; Weelink et al., 2010;

Callaghan, 2013b; Gieg and Toth, 2017). Carboxylation is not believed to be a major activation pathway, as this would involve addition of CO2 directly to the aromatic ring (creating a thrice- substituted product) or replacement of a methyl group. Despite this, methylbenzoates as intermediates in xylene degradation have been reported (Krieger et al., 1999; Elshahed et al., 2001;

Morasch et al., 2004b). Also known as toluates, these metabolites form downstream of the initial fumarate addition product. Certain methylbenzoate conformations (in particular para substitutions) are suspected not to be further metabolized due to the stereochemistry of their interactions with downstream enzymes (Rabus et al., 2016a; Sperfeld et al., 2019).

Benzene lacks alkyl substitutions, making it considerably more stable and less reactive than the alkylbenzenes. A range of conflicting studies have suggested that methylation, hydroxylation, or carboxylation reactions occur before further metabolism (Chakraborty and Coates, 2005; Ulrich et al., 2005; Mancini et al., 2008; Atashgahi et al., 2018). Universally, however, it is accepted that the difficulty in anaerobic benzene biodegradation lies in overcoming the aromatic resonance stabilizing effects while also withstanding the cytotoxic properties of this relatively highly water soluble chemical (Headley et al., 2001; Vogt et al., 2011).

Due to the long generation times required for methanogenic hydrocarbon-degrading microorganisms (sometimes with 150-600 day lag periods, Fowler et al., 2016; and others with upwards of 150 days to observe complete degradation, Elshahed et al., 2001), most studies explore hydrocarbon-activating mechanisms by sulfate-reducers or those undergoing higher energy electron accepting processes. The lack of successful methanogenic BEX degradation experiments presents a gap in knowledge in establishing the precise mechanisms by which these hydrocarbons are degraded under the most marginal energy yielding conditions. Further, co-metabolic effects 45

during hydrocarbon biodegradation under methanogenic conditions are not clear. With these gaps in mind, we sought to explore the activation processes of benzene, toluene, ethylbenzene, and the xylenes (BTEX) under methanogenic conditions, both as sole substrates and in conjunction with toluene using a well-established methanogenic culture enriched on toluene as the sole carbon source.

4.2 Methods

4.2.1 Toluene-degrading methanogenic enrichment culture (TOLDC)

A toluene-degrading methanogenic enrichment culture, colloquially known as “TOLDC”, has been studied for 20 years. It was initially established as a sulfate-reducing enrichment derived from sediments in a gas condensate-contaminated aquifer near Denver, Colorado (Gieg et al.,

1999). Through multiple transfers and continued amendment with toluene, sediments were removed, and sulfate was depleted; thus, a methanogenic enrichment was developed that utilized toluene as the sole carbon source. Later work with this enrichment culture by Fowler (2014) and

Toth (2017) demonstrated that complete toluene degradation occurred within 30 days (up to 500

μM) with concomitant production of methane. Attempts to isolate individual microorganisms from this consortium through serial dilution and plating have thus far proved unsuccessful, which has been observed by other studies attempting similar isolation techniques (Ficker et al., 1999).

Metagenomic analysis of TOLDC by Fowler (2014) and Tan et al. (2015) revealed that

TOLDC was primarily composed of Firmicutes such as Desulfosporosinus and Clostridium,

Deltaproteobacteria such as Desulfovibrio or Syntrophus, Chloroflexi including the family

Anaerolineaceae, Spirochaetes, and methanogens such as Methanosaeta, Methanoculleus,

Methanoregula, and Methanolinea. Based on stable isotope probing experiments conducted with

TOLDC, Fowler et al. (2014) proposed that Desulfosporosinus activates toluene to 46

benzylsuccinate before downstream metabolism by Desulfovibrio, Syntrophus and possibly some role of Clostridium. Methanogens notably only made up 1.34% of the TOLDC metagenome (Tan et al., 2015), which when considering the demonstrated efficient methane production of this culture were less abundant than expected. The use of whole shotgun sequencing metagenomic techniques it believed to reduce amplification bias associated with PCR-based 16S rRNA gene sequencing, so these proportions of taxa are likely more accurate than 16S rRNA gene sequencing methods (Rausch et al., 2019; Brumfield et al., 2020).

4.2.2 Syntrophus aciditrophicus bioaugmentation

Syntrophus sp. has been hypothesized to play an important role in toluene biodegradation in TOLDC based on a stable isotope probing study by Fowler et al. (2014) and experiments by

Toth (2017), which indicated that the proportion of Syntrophus sp. artificially added to TOLDC positively correlated to the amount of methane produced. From these findings, several bioaugmentation experiments were undertaken to explore if increasing the relative abundance of this organism could stimulate faster toluene consumption or conversion. The first trial involved inoculating a pure culture of S. aciditrophicus (DSMZ 26646) grown in Pelotomaculum medium

(DSMZ 960; Appendix C) into TOLDC in varying proportions (0-40% v/v) and amending with toluene. In the second and third trials, S. aciditrophicus was grown until it reached log phase (as measured by the ATP test; section 3.7) and crotonate from the medium was depleted (10 mM; measured according to methods in section 3.5.1). The culture was then centrifuged at 6,000 x g for 10 minutes, the cell pellet was washed with fresh crotonate-free medium, then the biomass was added across logarithmic dilutions (104-106 active cells/mL as quantified by the ATP assay) into

TOLDC and amended with toluene. Methane production and toluene loss were monitored throughout (measured according to methods in section 3.3). 47

4.2.3 Phase 1 enrichments

With TOLDC as a model, well-studied methanogenic culture, we sought to examine its degradation capacity for other aromatic hydrocarbons including benzene, ethylbenzene, and xylenes with either a single BTEX hydrocarbon alone or in combinations with toluene. To establish these incubations, TOLDC was grown until no toluene could be detected by GC-FID then anoxically transferred (50% v/v) into new Pfennig medium in 120 mL serum bottles as previously described (section 3.2). BTEX hydrocarbons were added to create co-amended incubations (a

BEX hydrocarbon, 300 μM; with toluene, 500 μM) or incubation with single BTEX hydrocarbons.

All incubations were set up in triplicate. Approximately 300 days of incubation were required before any notable methane production was observed in these incubations (data not shown).

Following a further 100 days of monitoring, the most promising combinations (in terms of literature-reported metabolites, methane production, and hydrocarbon loss) were selected for continued monitoring, metabolite analysis, and real-time gene expression assays in phase 2. These included incubations of toluene + ethylbenzene, toluene + p-xylene, toluene alone, ethylbenzene alone, and p-xylene alone. No significant methane was produced (or hydrocarbon loss was observed) from incubations amended with benzene, o-xylene, or m-xylene, either alone or in the presence of toluene thus additional studies with these cultures were not pursued. Incubations were periodically assessed for changes in the microbial community composition by 16S rRNA gene sequencing (methods according to section 3.9).

4.2.4 Phase 2 enrichments

Triplicate incubations from phase 1 (totaling about 180 mL culture volume) were pooled and transferred (50% v/v) into new medium in 1 L bottles. Over the next 150 days, these cultures were regularly reamended with hydrocarbons, additional medium (approximately 100 mL every month), 48

as well as with a 10X concentrated Balch vitamin solution (Appendix E; 100 μL/100 mL culture) to stimulate growth. After a prolonged period of regular toluene loss and increasing amounts of methane, incubations were sampled at distinct time intervals for quantitative PCR analysis

(methods described in section 3.10 and 3.11) to determine real-time expression of the bssA gene

(encoding the most conserved subunit of benzylsuccinate synthase). Additionally, samples were collected to screen for predicted metabolites such as benzylsuccinates and other literature-reported metabolites (Table 4-1) via gas chromatography-mass spectrometry (GC-MS; section 3.8.2). Due to previous reports of low extracellular benzylsuccinate yields and therefore operating under the assumption that most benzylsuccinate would be intracellular (Beller and Edwards, 2000), the decision was made to collect two samples per time point for metabolite analysis. For the extracellular sample, 20 mL of culture volume was withdrawn and passed through a 0.2 μm filter to collect the cells for RNA extraction. The filtrate was passed immediately into sterilized serum bottles containing 12 N HCl to acidify metabolites to a pH of < 2. A second, unfiltered sample was passed directly onto the acid to lyse cells and acidify metabolites for whole cell extract analysis. Following examination of these two samples, metabolite concentrations were not found to correlate with the fraction analyzed therefore only whole cell extracts were analyzed in phase 3 experiments. Due to inconsequential findings in ethylbenzene-containing treatments during phase

2, phase 3 experiments involved only toluene and p-xylene.

4.2.5 Phase 3 enrichments

Following analysis of phase 2 treatments, a third round of incubations were established in triplicate to validate and confirm metabolites identified previously. New 20 mL incubations were established without diluting into new medium from the 1 L phase 2 stock cultures (to eliminate lag time) and reamended with toluene only, p-xylene only, or toluene + p-xylene. Hydrocarbon 49

unamended TOLDC and sterile hydrocarbon control treatments were also included. These incubations were monitored for methane production, hydrocarbon loss, and were routinely sacrificed for metabolite analysis.

4.3 Results

4.3.1 S. aciditrophicus bioaugmentation

Three bioaugmentation experiments with S. aciditrophicus and TOLDC were conducted to evaluate the role of Syntrophus sp. in toluene degradation in TOLDC. The first experiment involved transferring pure S. aciditrophicus into TOLDC in proportions ranging 0-40% (v/v) inocula. Toluene loss was not affected by the percent of S. aciditrophicus added, while increasing methane production correlated directly with the proportion of S. aciditrophicus added. In a second trial, S. aciditrophicus was grown until log phase before transferring, but this resulted in the same trends in methane and toluene concentrations as were previously observed. Thirdly, S. aciditrophicus were grown until cells reached log phase and crotonate was depleted before washing the cells with fresh crotonate-free medium, then transferring in logarithmic proportions

(104-106 cells/mL) into static amounts of TOLDC. Again, methane production correlated with the inocula added and toluene loss rates was unaffected. Following this third experiment, further attempts at bioaugmentation in this manner were abandoned.

4.3.2 Phase 2 experiments

Following an initial survey of BTEX biodegradation in phase 1 (section 4.2.3), toluene, ethylbenzene, and p-xylene-amended cultures were selected (based on their activity) to proceed to phase 2 experiments for further in-depth study. Methane production and hydrocarbon loss in phase

2 enrichments were monitored for approximately 150 days while samples were periodically removed for RNA analysis (Figure 4-8 and Appendix Table A-2) and metabolite extractions 50

(Figures 4-5, 4-6 and 4-7). Toluene (approximately 100 μmoles) was completely depleted in all treatments twice during the experiment within roughly 50 days (Figure 4-1A) while statistically significant loss of ethylbenzene and p-xylene was not observed (Figures 4-1B and C).

Ethylbenzene decreased in the toluene + ethylbenzene treatments by 2.8% relative to the ethylbenzene only treatment, while p-xylene in the toluene + p-xylene treatments decreased by

16.7% relative to the p-xylene only treatment. Stoichiometric conversion of toluene to methane

(1 mole toluene to 4.5 moles methane; Symons and Buswell, 1933), amounted to yields of 52% to

82% for each toluene amendment (Figure 4-2). Trace amounts of methane (less than 20 μmoles) were produced in the ethylbenzene only, p-xylene only, and unamended treatments (Figure 4-2).

A 100 B 150 C 200

)

e l

e 150 l

75 m

o 

100 m

(



(

e e

e 50 100

y l

b 50

- y

T 25 50

t E 0 0 0 0 50 100 150 0 50 100 150 0 50 100 150 Time (days) Time (days) Time (days) Toluene Ethylbenzene p-Xylene Unamended Toluene + Ethylbenzene Toluene + p-Xylene

Figure 4-1: Hydrocarbon degradation in phase 2 enrichments over 150 days after pooling and scaling up. Separate incubations of TOLDC were established with different carbon sources including toluene only, ethylbenzene only, p-xylene only, toluene + ethylbenzene, toluene + p- xylene, and an unamended treatment. After toluene (A) was depleted by approximately day 60, enrichments were reamended with 100 μmoles of toluene. Ethylbenzene (B) and p-xylene (C) were not reamended as they never reached zero. n=1.

800

Toluene

) s

e Ethylbenzene

l 600 o

m p-Xylene

 ( 400

e Toluene + Ethylbenzene

n a

h Toluene + p-Xylene t e 200

M Unamended

0 0 50 100 150 Time (days)

Figure 4-2: Methane production from TOLDC enriched on various carbon sources from phase 2 enrichments, then monitored for a further 150 days during the RNA and metabolite extraction experiment. Carbon sources in this experiment included: toluene only, ethylbenzene only, p- xylene only, toluene + ethylbenzene, toluene + p-xylene, and an unamended control treatment. n 1.

Enrichments were sampled at day 100 and day 600 for 16S rRNA gene sequencing to determine the microbial community composition of TOLDC. At day 100, methanogens belonging to the genera Methanosaeta, Methanoculleus, and Methanolinea made up over 90% of the relative abundance of all incubations (Figure 4-3). By day 600, communities in incubations containing toluene shifted to a 50-60% relative abundance of Desulfosporosinus, while those without toluene

(including the unamended) instead shifted to a 50-60% relative abundance of Desulfovibrio.

Toluene-containing incubations also contained Desulfovibrio, albeit at a lower relative proportion

(~22%). The relative abundance of methanogens decreased greatly in all treatments by day 600, with less than 1% relative abundance across the five taxa of methanogens detected. The diversity of TOLDC shifted between the communities analyzed on day 100 compared to those at day 600.

At day 100, very low diversity was observed across all samples, as shown by the low Bray-Curtis dissimilarity depicted in Figure 4-4A (and the Shannon diversity scores in Appendix Table A-3).

By day 600, diversity increased and enrichments with toluene grouped distinctly from those without toluene (Figure 4-4B).

Organic extractions were performed on all phase 2 TOLDC treatments, which were then surveyed for the presence of literature predicted metabolites (Table 4-1). By searching for diagnostic fragment ions, benzylsuccinic acid was detected in all incubations containing toluene

(Figure 4-5) and in trace amounts (less than 0.5 nmoles) in unamended and toluene-free treatments.

No other succinate derivatives such as (1-phenylethyl)succinic acid from ethylbenzene or 2-(4- methylbenzyl)succinic acid from p-xylene were detected. Also sought but not detected was 1- phenylethanol (from ethylbenzene).

TMS- Molar derivatized Parent Predicted activation Compound mass diagnostic compound metabolites (protonated) structure (g/mol) fragment ions (m/z) COOH

COOH Toluene α-benzylsuccinic acid 208.21 337 & 352

COOH

(1-phenylethyl)succinic COOH 222.24 351 & 366 acid

Ethylbenzene

1-phenylethanol 122.17 180 & 195

COOH

COOH 2-(4-methylbenzyl)succinic 222.24 351 & 366 acid

p-Xylene

COOH

p-toluic acid 136.15 193 & 208 (or 4-methylbenzoic acid)

) s

e 125

o m

n 100 Toluene

(

i Ethylbenzene c

a 75

p-Xylene

i n i Toluene + Ethylbenzene

c 50

c u

s Toluene + p-Xylene l

y 25

z Unamended

n e

b 0 0 20 40 60 Time (days)

In the toluene + p-xylene treatments, a peak with a retention time of 31.8 minutes was observed (Figure 4-6A); this was not similarly present in toluene only, p-xylene only, or unamended treatments. Investigation of the diagnostic fragments associated with this chromatographic peak revealed two ions with mass to charge ratios (m/z) of 193 and 208 (Figure

4-6C), which matched the trimethylsilyl (TMS) derivatized p-toluic acid standard (Figure 4-6B).

This p-toluic acid standard also had a matching chromatogram peak at 31.8 minutes. p-Toluic acid was detected in all treatments amended with toluene and p-xylene (Figure 4-7) with amounts ranging from 40 to 85 nmoles; it was also found in some time-points in p-xylene only amended culture extracts (less than 1.5 nmoles).

100

)

e l

o 80 Toluene m

n Ethylbenzene (

60 d

i p-Xylene

c 40 Toluene + Ethylbenzene

u l

o Toluene + p-Xylene t

- 20

p Unamended 0 0 20 40 60 Time (days)

During the metabolite sampling period, additional samples were withdrawn for RNA extraction to quantify changes in expression of the bssA gene over time when incubated on the various substrates. Toluene only treatments increased in bssA gene expression between days 1 through 21 (3 times more abundant than the unamended) relative to the unamended control before decreasing (Figure 4-8). The unamended treatment did not similarly increase in bssA expression at day 21. In the ethylbenzene only treatments (Figure 4-8A) bssA expression also appeared to increase by more than 3-fold at day 11, however there was large standard error associated with these values. The fold change of bssA was not observed to increase in treatments with toluene and either ethylbenzene (B) or p-xylene (C), nor in treatments with p-xylene alone in a manner that was comparable to that of toluene alone. In all treatments, the highest increase in bssA gene expression was in the toluene only amended incubations.

A 6 B 4

s b

b 3

f f

o 4

g n

n 2

d l

l 1

F F 0 0 0 10 20 30 40 0 10 20 30 40 Time (days) Time (days)

C 4 D15

s b

3 b

f o

o 10

g n

2 n

d l

1 l

F F 0 0 0 10 20 30 40 0 10 20 30 40 Time (days) Time (days)

Toluene Ethylbenzene p-Xylene Unamended Toluene + Ethylbenzene Toluene + p-Xylene

4.3.3 Phase 3 enrichments

Conversion of toluene to methane (Figure 4-9) in the toluene only treatments corresponded to 103% of the predicted stoichiometric yield and 94% in treatments with toluene + p-xylene

(assuming only toluene was consumed). A t-test calculated a p-value of 0.024 for the means of methane produced from the toluene only treatment compared to the unamended, a statistically significant difference. Toluene was completely consumed in both toluene alone and in the toluene

+ p-xylene treatments within 50 days relative to the sterile control (Figure 4-10A). In contrast, p-

xylene loss was not significant (Figure 4-10B), as incubations with toluene + p-xylene demonstrated an average 6.4% reduction relative to the p-xylene only treatment. Abiotic loss of p-xylene from sterile controls amounted to approximately 0.063 ± 0.005 μmoles, while p-xylene only treatments lost 0.074 ± 0.123 μmoles and toluene + p-xylene lost 0.110 ± 0.212 μmoles of p- xylene.

)

s e

l 15

o Toluene

 (

10 p-Xylene

e n

a Toluene + p-Xylene

h t

e 5 Unamended M

0 0 10 20 30 40 50 Time (days)

Figure 4-9: Average methane production from phase 3 experiments of TOLDC amended with toluene and/or p-xylene. Error bars represent the standard error of the mean of 3 replicates.

A 4 B 4

)

e l

l 3 3



(

2 e 2

X o

1 - 1

T p

0 0 0 10 20 30 40 50 0 10 20 30 40 50 Time (days) Time (days) Toluene p-Xylene Toluene + p-Xylene Sterile

Figure 4-10: Hydrocarbon loss from phase 3 TOLDC with toluene (A) and p-xylene (B). Unpaired two-tailed t-tests of the change in toluene concentration relative to the sterile control from day 1 to day 50 (A) determined p-values for the toluene only treatment (blue) of 0.0005% and 0.0011% for toluene from the co-amended treatment (green). Non-significant p-values were calculated for the changes in all p-xylene treatments. Error bars ± SEM, n=3.

The amount of benzylsuccinic acid (Figure 4-11A) in the toluene + p-xylene incubations was low at day 0 (36.9 ± 8.5 nmoles) but more than doubled immediately upon addition of hydrocarbons (day 1; 85.3 ± 6.2 nmoles) and peaked by day 5 before leveling out at approximately

30 to 50 nmoles for the remainder of the monitoring period. The amount of benzylsuccinic acid in the toluene + p-xylene treatments did not disappear after 50 days, contrary to what was observed in toluene only treatments (Figure 4-5). In the same treatments, p-toluic acid levels were highest at day 0 (96.1 ± 11.3 nmoles; Figure 4-11B) before decreasing to ~50 nmoles by day 12 and remaining relatively constant throughout the rest of the experiment. The difference in p-xylene loss between the sterile control (accounting for abiotic loss) and the toluene + p-xylene treatment was approximately 47 nmoles, which may be attributed to initial p-toluic acid production and transient transformation.

A 125 B 125

)

i l

c 100 100

)

(

i 75 75

n l

50 c 50

(

e t

25 - 25

b p 0 0 0 10 20 30 40 50 0 10 20 30 40 50 Time (days) Time (days) Toluene p-Xylene Toluene + p-Xylene Unamended

Figure 4-11: Benzylsuccinic acid (A) and p-toluic acid (B) in phase 3 enrichments with toluene, p-xylene, toluene + p-xylene or unamended treatments. Error bars ± SEM, n=3.

4.4 Discussion

These experiments were undertaken with the goal of elucidating activation metabolites of monoaromatic hydrocarbons by a long-established toluene degrading methanogenic enrichment.

Experiments focused on ethylbenzene and xylene activation metabolites under these conditions.

Additionally, we sought to explore the effects of co-metabolism on the biotransformation of these alkylbenzenes of interest. Changes in both the microbial community composition and the expression of bssA were assayed over time in cultures amended with the various substrates, and experiments were conducted to evaluate the role of Syntrophus sp. in toluene biodegradation.

4.4.1 Ethylbenzene

No metabolites indicative of ethylbenzene degradation [(1-phenylethyl)succinic acid or 1- phenylethanol; Table 4-1] were detected in these experiments. Ethylbenzene loss was negligible in both ethylbenzene-amended and toluene + ethylbenzene-amended treatments (Figure 4-1B) and methane production did not rise above that of the incubations amended with toluene alone. As such, these incubations will not be further discussed in detail.

4.4.2 Hydrocarbon transformation

Methane production in the phase 1 enrichments was very slow, involving a lag phase of approximately 250 days before production from hydrocarbon-amended incubations exceeded that of the unamended (data not shown). After toluene was completely depleted in all co-amended incubations (roughly 300 days), experimental replicates were pooled and scaled up as described in section 4.2.4. Phase 2 enrichments demonstrated repeated, complete toluene loss in 50-60 days

(Figure 4-1A), in contrast to ethylbenzene and p-xylene which did not undergo any notable loss over 150 days (Figures 4-1B and C). Consumption of toluene yielded 52-82% of the predicted stoichiometric methane yield (Figure 4-2), which is in line with previous studies of methanogenic 62

toluene conversion (Fowler et al., 2012), suggesting the remaining carbon was likely incorporated to biomass.

Phase 3 enrichments with toluene or toluene + p-xylene produced 103% and 94% respectively of the predicted methane yield (Figure 4-9), assuming negligible degradation of p- xylene occurred. Similar to observations in phase 2 experiments, no significant p-xylene loss was observed (Figure 4-10B). p-Xylene is generally considered a poorly-degradable hydrocarbon

(Foght, 2008; Weelink et al., 2010; Rabus et al., 2016a), although select studies have demonstrated its degradation coupled to nitrate reduction (Häner et al., 1995; Rotaru et al., 2010) and sulfate reduction (Morasch and Meckenstock, 2005; Nakagawa et al., 2008). Additionally, there are reports of p-xylene loss that only occurred co-metabolically with toluene degradation (Kasai et al.,

2007). In the present experiment, only toluene degradation was observed without significant p- xylene loss relative to the sterile control. Despite this, the toluene + p-xylene enrichments lost

76% more p-xylene than the sterile control (approximately 48 nmoles) and 49% more than the p- xylene only treatment (36 nmoles). It is possible this missing p-xylene in the co-amended treatments was transformed into p-toluic acid (Figure 4-11B), as the variance in p-toluic acid from day 0 to day 50 was approximately 50 nmoles (discussed further in section 4.4.3.1).

The recalcitrance of p-xylene is not believed to be due to the stereochemistry of p-xylene itself, but rather the difficulty in transforming the central intermediate benzoyl-CoA analogue (4- methylbenzoyl-CoA) for entry into the β-oxidation pathway. Steric hindrance of the para substituted methyl group is believed to prevent electron transfers carried out by benzoyl-CoA reductases (BCRs) and destabilizes the radical-anion intermediate (Rabus et al., 2016a; Sperfeld et al., 2019). Callaghan (2013a) suggested that another issue with further degradation of methyl- substituted succinates derived from xylenes is the presence of a second chiral carbon, as opposed 63

to benzylsuccinate which has only one. Normal BCR functioning (in the case of toluene degradation pathways) involves several reactions to reduce the aromaticity of benzoyl-CoA in order to break open the ring, form a muconate intermediate, and enter the TCA cycle (Abu Laban et al., 2010; Porter and Young, 2014). A 4-methylbenzoyl-CoA reductase was identified in a p- toluate (i.e. 4-methylbenzoate) degrading pure culture of Magnetospirillum sp. MbN1, however the authors noted p-xylene itself could not be degraded (Lahme et al., 2012). No taxon annotated as Magnetospirillum sp. or in fact any higher-order lineage of the Rhodospirillales was detected in the p-xylene amended incubations (Appendix Table A-1).

4.4.3 Metabolite analyses

4.4.3.1 p-Toluic acid

In incubations containing p-xylene, succinate and carboxylated derivatives were predicted to be potential but thus far uncharacterized intermediates of p-xylene degradation by TOLDC

(Table 4-1). In all live toluene + p-xylene containing incubations (and select timepoints of p- xylene only analysis), a peak was observed with a retention time of 31.8 minutes that was not present in the toluene alone or unamended samples (Figures 4-6A and C). This compound was positively identified as it matched the diagnostic fragment ion pattern (m/z 193 and 208) of an authentic p-toluic acid (i.e. 4-methylbenzoic acid) standard that was also analyzed (Figure 4-6B). p-Toluic acid was detected in toluene + p-xylene-amended cultures at concentrations up to 50-fold higher than those containing p-xylene alone (Figures 4-7 and 4-11B). The predicted fumarate addition product, 2-(4-methylbenzyl)succinic acid, was sought but not detected in any samples. p-

Toluic acid has previously been identified as a dead-end product produced by Desulfobacula toluolica in sulfate-reducing incubations containing toluene and p-xylene as substrates (Rabus and

Widdel, 1995b). Aerobically, p-xylene has been characterized to form p-toluic acid before further 64

oxidation and decarboxylation to form 4-methylcatechol (Jindrová et al., 2002). The aerobic experiment showed complete mineralization of p-xylene to CO2 via the p-toluic acid intermediate, which to date no anaerobic studies have shown. In their study, Rabus and Widdel (1995b) noted the requirement for toluene to be present to facilitate p-xylene transformation, which was similarly observed in other studies (Kasai et al., 2007) including the present experiment. All currently available evidence suggests anaerobic p-xylene transformation to p-toluic acid is invariably a dead-end process (Biegert and Fuchs, 1995; Rabus and Widdel, 1995b; Kasai et al., 2007), while all reports of 2-(4-methylbenzyl)succinic acid as an intermediate were associated with complete or near complete mineralization (Elshahed et al., 2001; Rotaru et al., 2010). No studies were found attesting to methanogenic p-xylene degradation or characterizing the associated metabolites when provided as a sole substrate.

p-Toluic acid in this experiment is not likely to have arisen from direct carboxylation of p- xylene, as aromatic carboxylation enzymes typically act on the ring itself rather than an existing methyl group (Zhang and Young, 1997; Foght, 2008). Direct carboxylation would likely have resulted in a thrice-substituted compound such as 2,5-dimethylbenzoate, which was also not detected. This intermediate may have first been transformed into 2-(4-methylbenzyl)succinic acid

(perhaps at levels below the instrument detection limits) then followed a pathway with intermediates analogous to that of toluene (Figure 2-4) to form 4-methylbenzoyl-CoA. While thioester bonds (connecting the sulfur atom of coenzyme A to the benzylic carbon of benzoyl-CoA or 4-methylbenzoyl-CoA) are generally considered stable at neutral pH, acid-catalyzed hydrolysis is a thermodynamically favorable non-enzymatic reaction which can irreversibly convert CoA- substituted intermediates to their carboxylated forms (Horton et al., 2006; Bracher et al., 2011).

The acidification step necessary for organic extraction which reduced the pH to 2 or less (and in 65

some cases pH 0) likely altered the metabolites from their CoA-substituted form, therefore it is impossible with the analytical techniques employed to determine if the p-toluic acid detected was formed directly or if it was metabolically transformed all the way to the purported dead-end 4- methylbenzoyl-CoA.

While p-toluic acid is not exclusively a metabolite formed through anaerobic microbial metabolism, its absence in toluene only or unamended enrichments strongly indicates it arose through the biotic transformation of p-xylene. The fact that p-toluic acid was formed in larger amounts in the toluene + p-xylene treatment relative to the p-xylene only treatment supports the argument that the presence of toluene was in some way necessary to initiate p-xylene transformation. Toluene binds to the promoter of the bss operon to initiate transcription (Hermuth et al., 2002), however antagonistic effects with p-xylene may have been responsible for the low transcription of bssA that was observed in this study when both substrates were provided together

(discussed further in section 4.4.5). Despite reports that BSS is very specific for toluene (Funk et al., 2014, 2015), evidence exists from other studies that found BSS could act upon other aromatic compounds including xylenes (Beller and Spormann, 1999; Verfürth et al., 2004). In the latter two studies, both reported BSS purified from Azoarcus sp. strain T displayed activity on p-xylene and that 2-(4-methylbenzyl)succinate was formed; however, it is worth pointing out that the use of a purified enzyme from a cell-free extract would negate the need for toluene to regulate transcription so it is difficult to determine from these studies if the same activity would be observed in a whole-cell bacterial culture. Although no 2-(4-methylbenzyl)succinic acid was detected in the present study, we cannot rule out that fumarate addition to p-xylene did not initially occur prior to p-toluate formation.

4.4.3.2 Benzylsuccinic acid

Benzylsuccinic acid was detected in all treatments containing toluene. Notably, in co- amended experiments where toluene was present with either ethylbenzene or p-xylene, benzylsuccinic acid was shown to persist without obvious reduction in concentration (Figure 4-5) in contrast to toluene only-amended enrichments, in which a decrease in benzylsuccinic acid was observed by day 60. Well-known to be a transient metabolite, benzylsuccinic acid was expected to be produced and consumed, correlating with increasing methane production (Figures 4-9 and 4-

11A, day 12). Curiously, in phase 2 experiments, benzylsuccinic acid was easily detected in the toluene only treatments (Figure 4-5), however in phase 3 experiments only trace amounts were detected (Figure 4-11A). As benzylsuccinate is typically found at concentrations 1000-fold less than the parent compound (Elshahed et al., 2001; Fowler et al., 2012), this would represent only 3 nmoles, which may fall below the detection limits of our instrument. Interestingly though, the toluene plus either ethylbenzene or p-xylene-amended incubations produced approximately double the amount of benzylsuccinic acid compared to those containing toluene alone (Figures 4-5 and 4-

11A); in phase 3 experiments, there was a 231% increase in the amount of benzylsuccinic acid formed in the toluene + p-xylene treatment within the first 24 hours after toluene addition.

Contrary to expectations, co-amended incubations in both phase 2 and phase 3 demonstrated persistent benzylsuccinic acid amounts rather than what should be a rapidly formed and consumed metabolite. In phase 3, the fact that the toluene + p-xylene treatments had initial high benzylsuccinic acid levels before decreasing by day 12 then remaining relatively constant around

40 nmoles for the rest of the experiment raises several questions: was further benzylsuccinic acid transformation being inhibited, and was this occurring due to accumulation of metabolic intermediates or inhibition of an enzyme? Also, why was some benzylsuccinic acid produced and 67

consumed quickly, yet the remainder was not consumed? Benzoic acid, the central intermediate leading to the β-oxidation pathway was found not to accumulate in any treatments in either phase

2 or phase 3 experiments (Appendix Figures A-2 and A-4), indicating that any benzylsuccinic acid formed was indeed converted at least to benzoyl-CoA if not further metabolized. As benzoic acid levels were relatively consistent across all treatments (2-6 nmoles), it does not appear likely that p-toluic was demethylated or contributed to the overall amount of benzoic acid detected.

4.4.4 qPCR assay optimization

Attempts were made to quantify changes in bssA gene expression over time when TOLDC was amended with different aromatic substrates. Initially, several trials were conducted with established primer sets from Winderl et al. (2007) to amplify bssA from pure cultures of known toluene-degrading microorganisms (T. aromatica and D. toluolica) in a qPCR assay. While this was relatively successful, the same primer sets failed to reproducibly amplify bssA in the mixed consortium TOLDC. Eventually, primer sets were obtained from a colleague (Table 3-2; Kharey et al., 2020) that were designed to target a broad range of bssA sequences in environmental samples and were shown to successfully do so with good coverage of Clostridia sequences (the lineage to which Desulfosporosinus belongs) following Illumina sequencing of qPCR products. Early tests with these primers were successful at producing a single small molecular weight amplicon from

TOLDC DNA and thus were considered qPCR compatible; however, testing with T. aromatica resulted in two amplicons with two melt peaks (not qPCR compatible) and tests with D. toluolica failed to produce an amplicon (the authors noted limited coverage of the Deltaproteobacteria bssA sequences with this primer pair). These results necessitated using a purified bssA PCR product from TOLDC as a standard in the qPCR assay, which was successful when implemented.

The purified 140 bp fragment obtained from amplifying TOLDC DNA with WinF/HitR primers was subsequently Sanger sequenced. Despite several attempts, the sequences returned did not match established bssA sequences when analyzed through NCBI BLASTn; in fact, no hits of high homology were returned. Most bases from the sequencing run were called with high confidence, while select regions of the amplicon had base calls with medium or low confidence.

As a result of issues with timing and access to sequencing facilities due to COVID-19, this bssA amplicon was unable to be sequenced for positive identification with next-generation technologies such as Illumina MiSeq. Despite this, these primers had previously been shown (with sequence confirmation) to amplify bssA from diverse environmental samples including those belonging to

Clostridia (Kharey et al., 2020) and the assay did not produce an amplicon from benzoate-grown

T. aromatica or E. coli used as gene-negative controls. Primers were analyzed in silico through the BLASTn database and returned benzylsuccinate synthase and Desulfosporosinus-specific hits.

Amplicons resulting from the qPCR assay were checked for size and purity by gel electrophoresis and found to match the positive control bssA DNA amplicon (Appendix Figure A-3A), and all amplified products had melting temperatures from 86-87˚C (Appendix Figure A-3B) which were in line with standards also assayed. Metagenomic analysis by Tan et al. (2015) revealed bssA sequences in TOLDC diverged considerably from bssA sequences in two other metagenomes enriched from environmental sources that were also analyzed. The authors postulated that divergent evolution occurred in TOLDC bssA genotypes due to the long-term enrichment (at that time, 10+ years) compared to the other two cultures (2-3 years) and possibly was also influenced by the carbon sources (TOLDC enriched solely on toluene for over a decade while the other two received hydrocarbon cocktails of short-chain alkanes or naphtha components). While the exact identity of this amplicon is uncertain because Sanger sequencing did not conclusively confirm it 69

was a bssA sequence, all other lines of evidence supported that the correct gene fragment was amplified.

4.4.5 Reverse-transcription bssA gene expression analysis

Following RNA extraction, purification, and reverse transcription, cDNA was subsequently used in qPCR assays to evaluate changes in bssA expression over time when TOLDC was incubated on various substrates. As shown in Figure 4-8, toluene grown TOLDC was shown to increase its fold expression of bssA (relative to the unamended) between days 11 and 21

(maximally 3 times higher) before decreasing in expression by day 36. This was in contrast to a similar assay conducted previously for bssA quantification in TOLDC (Fowler et al., 2014), which used DNA instead of cDNA. That study reported a consistent level of 1.5 to 2-fold higher quantity of bssA gene during toluene degradation relative to the unamended throughout 20 days of monitoring. The use of RNA isolation in the present study may shed light on changes in expression level rather than basal presence of bssA in the genome, which appeared unaffected by the presence of toluene in the Fowler et al. (2014) study.

Ethylbenzene only amended incubations (Figure 4-8A) had higher fold change in bssA than toluene at day 11 (3.5 times) relative to the unamended control, however this data had high error associated (raw gene copies detected ranged from 9 to 141 copies across three technical replicates) so it can only be tentatively discussed. Contrary to expectations, treatments with toluene + ethylbenzene had lower fold change in expression than the unamended treatment (Figure 4-8B), despite the addition and consumption of toluene as the primary substrate. In fact, several samples from the toluene + ethylbenzene incubations had no increased expression in any replicates (days

1, 11, and 21). A similar trend was observed in the toluene + p-xylene samples (Figure 4-8D), where the fold change in bssA expression was lower than with p-xylene alone (Figure 4-8C). 70

Patterns in bssA expression in the p-xylene alone treatment most closely mirrored that of the unamended with fold change ranging from 1.37 on day 1 to 0.54 at day 36. An unusual result from these assays was the lack of higher bssA expression in treatments co-amended with toluene.

Evidence indicates that toluene must bind to the promoter of the bss operon to initiate transcription

(Hermuth et al., 2002), therefore the presence of toluene should catalyze bssA production. The slight repression observed in both co-amended incubations suggests that an antagonistic effect was exerted by the xenobiotic compounds on bss regulation. It is also possible that rather than transcribing more BSS, a basal level was maintained within the population from when the enrichment was transferred from toluene grown TOLDC (over 600 days prior).

4.4.6 Microbial community analysis

The contamination of different environments with petroleum hydrocarbons exhibits a substantial selective pressure on diversity of microbial communities. In a study examining the microbial communities in diesel-contaminated soils and comparing the communities from adjacent clean soils, Sutton et al. (2013) found that the presence of petroleum hydrocarbons positively correlated with an increase in the relative abundance of Choroflexi, Firmicutes, and Euryarcheota compared to the uncontaminated site. Multiple other studies have found these taxa associated with anaerobic degradation of complex carbon sources such as hydrocarbons or wastewater. The success of these taxa may be attributed to one of several factors: their ability to metabolize hydrocarbons compared to other native microbes which cannot, their ability to withstand the localized cytotoxicity exhibited by the hydrocarbons compared to other microorganisms that are harmed, depletion of nitrogenous and phosphorous compounds while the ratio of carbon compounds remains high, and their ability to ferment or use reduced electron acceptors in mature contaminated systems (Sutton et al., 2013). TOLDC has been previously characterized to possess 71

many of these lineages, with Firmicutes such as Desulfosporosinus identified as the most likely initiators of fumarate addition to toluene via RNA-SIP experiments (Fowler et al., 2014). In the same experiment, Deltaproteobacteria such as Desulfovibrio and Syntrophus were found to incorporate labelled 13C from toluene at later sampling points than Desulfosporosinus, leading to the inference that these organisms are involved in the downstream metabolism of toluene or its byproducts. Chloroflexi, including several taxa of the Anaerolineaceae family, made up 1-4% of

TOLDC’s relative abundance (Figure 4-3 and Appendix Table A-1), which have been associated with alkane degradation in long term incubations. Methanogens belonging to the Euryarcheota phylum round out the core microorganisms believed to be involved in toluene degradation in

TOLDC.

After just 100 days of enrichment on the various hydrocarbon substrates (and negligible methane production; phase 1 experiments), Desulfosporosinus was minimally present (0-4% relative abundance; Figure 4-3) while methanogens such as Methanosaeta (37-75%),

Methanoculleus (9-34%), and Methanolinea (0-12%) dominated the microbial communities.

When the same incubations were sequenced after 500 additional days of continued monitoring and enrichment, a dramatic shift was observed from a dominance of methanogens (collectively less than 1%) to that of communities dominated by Desulfosporosinus (51-60% relative abundance in toluene-containing samples) and Desulfovibrio (22-60% relative abundance in all treatments).

This shift could be due to the state of TOLDC when the experiment began; the culture had not been amended with toluene in over a year and was likely in a starvation state prior to its use for the experiments described in this chapter. Additionally, the day 100 sequencing itself was not presented with a great deal of confidence, as < 1000 reads were obtained from this analysis which indicated poor sequencing depth (usually > 10,000 reads). In contrast, all day 600 samples had 72

over 20,000 reads which conveys greater confidence and sequencing depth. Deltaproteobacteria such as Desulfovibrio and Syntrophus are often underrepresented when conducting Illumina MiSeq sequencing using V6-V8 primers and methanogens are generally overrepresented (Tremblay et al.,

2015). Additionally, multiple copies of 16S rRNA genes (i.e. certain Clostridium sp. have been shown to have 11 copies) may contribute to overrepresentation of certain taxa (Tan et al., 2015), thus their actual abundances in these incubations may be slightly distorted.

Desulfosporosinus sp. is believed to be primary initiator of toluene degradation in TOLDC

(Fowler et al., 2014); this taxon was found in high abundance in both the toluene only treatment

(51% relative abundance; Figure 4-3) and in the treatments amended with toluene and a second hydrocarbon (50-60% relative abundance). Accordingly, Desulfosporosinus was found at comparatively low relative abundance (approximately 1%) in all incubations that did not contain toluene. Illumina-sequenced reads from TOLDC that annotated as Desulfosporosinus were additionally analyzed through BLASTn, which returned 9 reads with 95.3-99.8% sequence homology to a Desulfosporosinus sp. designated “Tol M” from a methanogenic oil sands tailings enrichment characterized by Abu Laban et al. (2015). This taxon was identified through

13 incorporation of C6-labelled toluene and subsequent DNA stable isotope probing (SIP) fractionation of the heavy DNA. Metagenomic analysis revealed bssABC genes for fumarate addition, bbsBDEF for the enzymes transforming benzylsuccinate to benzoyl-CoA, and bamA-I genes which encode the enzymes involved in ring opening (Kube et al., 2004; Porter and Young,

2014; Abu Laban et al., 2015). Complete sequences of these genes were also detected in a metagenome analysis of TOLDC (Tan et al., 2015).

Unlike other Desulfosporosinus species, this taxon described by Abu Laban et al. (2015) possessed no dsrAB genes for sulfate reduction, which was consistent with data collected by Tan 73

et al. (2015) in their analysis of TOLDC. It is unknown if benzylsuccinate synthase possessed by

Desulfosporosinus sp. Tol M is capable of non-specific hydrocarbon activation (i.e., p-xylene) or if another bacterium may be carrying out this reaction (e.g., Clostridium, Desulfovibrio, and

Desulfobacteraceae increased in relative abundance in Figure 4-3 in p-xylene-containing treatments). Two additional reads clustering as a second Desulfosporosinus sp. was identified in the same 16S rRNA gene analysis; BLASTn analysis of these reads returned 95.4-95.8% sequence identity with Desulfosporosinus clone F5OHPNU07H9HB2, which was reported by Ramos-

Padrón et al. (2011) in oil sands tailings ponds samples. This Desulfosporosinus lineage was found between 0-0.11% relative abundance (Figure 4-3 grouping as Other and Appendix Table A-1); it is unknown if this second Desulfosporosinus also possesses toluene degradation or sulfate reduction genes. As Desulfosporosinus sp. Tol M was found at very low abundances in toluene- free treatments, this indicates it was not enriched or that its growth was not encouraged on other alkylbenzene substrates. Both organisms identified by Rabus and Widdel (1995b) and Kasai et al.

(2007) to putatively form p-toluic acid from p-xylene (Desulfobacula toluolica and Azoarcus sp.

DN11, respectively) were not detected in TOLDC, however unclassified members of the

Desulfobacteraceae family were detected (0.4 to 1.6% relative abundance; Figure 4-3) which could include the Desulfobacula genus. No organisms previously proposed to degrade p-xylene

(including genera such as Aromatoleum, Azoarcus, Thauera, Sulfuritalea, Georgfuchsia,

Magnetospirillum, and Desulfosarcina; Nakagawa et al., 2008; Sperfeld et al., 2019) were identified in the community analyses of TOLDC in our experiments.

There is little consensus as to the fate of BSS activation products in vivo, particularly in methanogenic cultures. Beller and Edwards (2000) suggested that there may be transfer of benzylsuccinate between members of a toluene-degrading methanogenic consortium in syntrophic 74

association before further degradation occurs. A Desulfovibrio sp. (designated “SRL8083” through BLASTn analysis) was also identified in all treatments (Figure 4-3), however its variable abundance in treatments with toluene raises several questions about its role in aromatic hydrocarbon metabolism. In incubations without toluene (ethylbenzene only, p-xylene only, and unamended), Desulfovibrio made up 50-60% relative abundance of the total community composition. This was in contrast to toluene-amended treatments which, while dominated by

Desulfosporosinus, also had on average 22% relative abundance of Desulfovibrio, suggesting that

Desulfovibrio plays an important role both in toluene metabolism and in the fermentation or necromass cycling occurring in the toluene-free treatments. Fowler et al. (2014) suggested that

Desulfovibrio was not involved in the initial activation of toluene in TOLDC but rather participated in a downstream step (as in a SIP experiment it became labelled with 13C a few days after

Desulfosporosinus). Several strains of Desulfovibrio have been characterized to degrade a range of aromatic compounds including di- and trinitro-substituted toluene, hydroxy and methoxy- substituted benzaldehydes and benzoates, and trihydroxybenzene (Zellner et al., 1990; Boopathy and Kulpa, 1993; Boopathy et al., 1993; Reichenbecher et al., 2000), therefore there is a possibility this lineage of Desulfovibrio could be similarly involved in toluene degradation or transformation of aromatic intermediates. As Desulfosporosinus sp. Tol M possesses bam genes encoding ring opening enzymes (Abu Laban et al., 2015; Tan et al., 2015), it is likely the dicarboxylic or fatty acids generated from these reactions are then passed to Desulfovibrio sp. SRL8083 or

Syntrophaceae (Figure 4-3) through a syntrophic mechanism. While Desulfovibrio species are known sulfate-reducers, they have been observed to switch to a fermentative or syntrophic metabolism once sulfate is depleted and can even become obligate syntrophs in these conditions

(Meyer et al., 2013; Gieg et al., 2014). Species of Desulfovibrio have been found to syntrophically 75

support methanogens, such as D. vulgaris which grew in association with Methanosarcina barkeri by producing acetate, CO2, and H2 which were then taken up by the methanogen partner (Stams and Plugge, 2009). We propose that a similar association likely exists between Desulfovibrio and the methanogens of TOLDC, and additionally that Desulfovibrio may play a role in the biotransformation of benzoate or other downstream metabolites produced during toluene metabolism.

Taxa of the Syntrophaceae lineage (Syntrophus and Smithella) were also identified by

Fowler et al. (2014) to be involved in toluene degradation in TOLDC, as they became labelled in

13C between days 8-20 in a SIP experiment conducted with this methanogenic consortium. In the present experiment, members of the Syntrophaceae family were detected at 1 to 9% relative abundance (with some grouping as “Other” in Figure 4-3) with slight enrichment in toluene-free or unamended treatments by day 600 (2.6-2.7%) compared to toluene amended treatments (1.0-

1.3%). Well-characterized facultative syntrophs such as Syntrophus aciditrophicus can metabolize benzoate or fatty acids to acetate via a reversible pathway involving crotonate, and can generate

ATP through a novel substrate-level phosphorylation mechanism (McInerney et al., 2007; Stams and Plugge, 2009; James et al., 2016, 2019). These enzymes operate near-equilibrium, meaning this pathway can easily become inhibited, cease, or reverse depending on accumulation of byproducts and the rate by which they are consumed; in particular, levels of cyclohexane carboxylate, formate, acetate, and H2 must be kept low to drive the reaction forward (James et al., 2019).

Additionally, benzoate may serve as an electron acceptor when cyclohexane carboxylate accumulates (Mouttaki et al., 2008). The degradation of benzoate to acetyl-CoA by Syntrophus sp. was hypothesized to be the rate-limiting process in TOLDC (Toth, 2017), thus this bacterium requires further investigation in the context of its metabolic role in TOLDC. 76

S. aciditrophicus can act either as a primary or secondary syntrophic fermenter due to its reported incorporation of substrates; it is not able to metabolize toluene or alkyl-substituted aromatics directly (McInerney et al., 2009). The strategy of cycling end products and substrates depending on the external conditions gives S. aciditrophicus a competitive advantage in carbon and electron acceptor depleted environments because it does not have to rely on input of new substrates. Due to the predicted role of Syntrophus sp. in TOLDC, several bioaugmentation experiments were undertaken to explore if increasing the relative abundance of this organism could stimulate faster toluene consumption or conversion. Toth (2017) found that the proportion of a

Syntrophus sp. artificially added to TOLDC positively correlated to the amount of methane produced. As a result, several attempts to evaluate this phenomenon more thoroughly were undertaken in the present study. However, given that augmenting TOLDC with S. aciditrophicus did not yield improved rates of toluene biodegradation relative to treatments without additional S. aciditrophicus (section 4.3.1), it appears more likely that the crotonate present in the S. aciditrophicus medium (Appendix C) was metabolized or that additional biomass was simply consumed by known necromass cyclers (such as the Spirochaetaceae; Dong et al., 2018) that were also detected in TOLDC (Figure 4-3). While these experiments did not support the hypothesis that simply bioaugmenting TOLDC with S. aciditrophicus would improve toluene degradation, it did reveal that additional input of carbon sources (crotonate and foreign biomass) was insufficient to stimulate faster toluene degradation and may have actually inhibited degradation of the more complex substrate (toluene) as has been previously reported (Edwards et al., 1992).

The final component to this consortium is the methanogens. Due to the longstanding relationship of the bacteria and archaea in this culture, intricate metabolic relationships developed over time. Reducing equivalents are shuttled between bacteria and methanogens in the form of 77

acetate, formate, and H2 through the process of interspecies electron transfer (Stams and Plugge,

2009; Morris et al., 2013; Gieg et al., 2014). While there was relatively low abundance of methanogens, particularly at the end-point analysis (0.17-0.87%; Appendix Table A-1), this was in line with analysis of the metagenome by Fowler (2014) who found similarly low abundances of methanogens (approximately 1%). Nevertheless, the methanogens within TOLDC were still able to convert acetate or H2 (following breakdown of toluene) completely to methane and CO2 in near- stoichiometric amounts (94-103% yield) in approximately 50 days (Figures 4-9 and 4-10A).

4.5 Conclusions

The results of these experiments lend new evidence to the field of TEX biodegradation by methanogenic consortia, both in the context of p-xylene biotransformation and the apparent requirement for co-metabolism with toluene to facilitate this process. Little success was achieved in bioaugmentation experiments with S. aciditrophicus to improve toluene degradation or in co- metabolism experiments with benzene, o-xylene, m-xylene, and ethylbenzene. While it is generally assumed that fumarate addition reactions are solely responsible for anaerobic aromatic hydrocarbon biodegradation under marginal energy conditions, the lack of corresponding metabolites in this experiment such as 2-(4-methylbenzyl)succinate as found in other studies (e.g.

Elshahed et al., 2001; Morasch and Meckenstock, 2005; Rotaru et al., 2010), casts doubt on this assumption and indicates that far more study is required to fully understand these dynamic and diverse hydrocarbon activation/biodegradation strategies. Here we presented evidence of an intermediate forming from a recalcitrant BTEX hydrocarbon, p-xylene, without corresponding detection of 2-(4-methylbenzyl)succinate in a co-metabolic mechanism with toluene. Degradation of toluene was not affected by the lack of p-xylene loss. p-Toluic acid (4-methylbenzoyl-CoA) was identified as a product of the p-xylene co-metabolic biotransformation. It remains unknown 78

which microorganism(s) carried out this reaction, however the increased abundance of

Desulfovibrio relative to toluene-containing treatments indicates that this microbe may be involved. We also found evidence that bssA expression may be repressed by non-target hydrocarbons such as p-xylene or ethylbenzene, despite toluene also being present. Co-metabolic effects of methanogenic BTEX biodegradation are varied and complex, and the present study only scratches the surface of the dynamics involved.

Chapter Five: Characterizing the biodegradation of polycyclic aromatic hydrocarbons by methanogenic microbial communities

5.1 Introduction

Polycyclic aromatic hydrocarbons (PAHs) are important targets for bioremediation due to their recalcitrance in the environment and extremely low chemical reactivity (Ghosal et al., 2016;

Meckenstock et al., 2016; Logeshwaran et al., 2018). These chemicals form either through biotic

(certain plant and bacterial biosynthetic reactions) or abiotic processes (charring of food, incomplete combustion of organic material, geothermal reactions) (Ghosal et al., 2016).

Anthropogenic activities such as the burning of fossil fuels and industrial activities also contribute to the release of PAHs into the environment (Wilson and Jones, 1993). In anoxic environments, such as water-logged soils and deep subsurface environments, PAHs adsorb to particulate matter and can remain unchanged in this state for hundreds or even thousands of years (Wilson and Jones,

1993; Mouttaki et al., 2012). These large, multi-ringed molecules are known to be lethal to aquatic invertebrates at very low concentrations and carcinogenic to humans, making their presence in the environment a concern (Canadian Environmental Protection Act: Polycyclic aromatic hydrocarbons, 1994; Meckenstock et al., 2016). Biodegradation by microorganisms is one of the few processes that can naturally attenuate these types of contaminants. In the deep subsurface, the biodegradation of hydrocarbons is usually associated with methane production (Head et al., 2003;

Jiménez et al., 2016; Toth and Gieg, 2018).

The biodegradation of 2- or 3-ringed PAHs has been examined under denitrifying, iron(III)- reducing, and sulfate-reducing conditions (Coates et al., 1997; Annweiler et al., 2002; Safinowski and Meckenstock, 2006; Kleemann and Meckenstock, 2011). PAHs comprised of four or more rings are believed to not be metabolized by anaerobic microorganisms, or potentially only under

co-metabolic conditions as a side reaction (Meckenstock et al., 2016). Comparatively few studies have reported methanogenic PAH biodegradation, and some have reported dubious evidence when compared to other available literature (failure to add electron acceptors such as bicarbonate or

CO2, unreasonably fast degradation rates, missing citations, numerous spelling errors, etc.; e.g.,

Wan et al., 2012; Zhang et al., 2012; Ye et al., 2018). That said, studies presenting high quality evidence of anaerobic PAH biodegradation do exist (Zhang and Young, 1997; Annweiler et al.,

2000; Chang et al., 2006; Safinowski and Meckenstock, 2006; Musat et al., 2009; Berdugo-Clavijo et al., 2012; Mouttaki et al., 2012; Eberlein et al., 2013; Toth et al., 2018; Shin et al., 2019;

Weyrauch et al., 2020), however the slow generation times of PAH-degrading cultures and conflicting results warrant much additional study. There is still debate as to the mechanisms involved in functionalizing PAHs under anaerobic conditions, and these mechanisms are likely not shared between substituted and unsubstituted hydrocarbons.

There are few studies examining the biodegradation of unsubstituted PAHs (such as naphthalene or phenanthrene) under anaerobic conditions, likely due to the very long generation times involved in culturing these microorganisms (Annweiler et al., 2002). In contrast, PAHs are relatively easily degraded by aerobic microorganisms. As previously discussed in section 2.5, highly energetic oxygenases can easily hydroxylate and break open aromatic rings. Aerobically, monoaromatics such as BTEX go through up to two rounds of oxidative attack, producing catechols and dicarboxylic acid intermediates; PAHs on the other hand require multiple rounds of oxidative attack to cleave aromatic rings, reducing the number of rings one at a time (Hidalgo et al., 2020). A similar strategy is believed to be employed under anaerobic conditions, wherein evidence suggests that anaerobic PAH biodegradation involves an initial functionalization of the hydrocarbon, then reduction of aromaticity before ring opening (Eberlein et al., 2013; Estelmann 81

et al., 2015). Under sulfate-reducing conditions, biochemical evidence has been presented that supports a methylation reaction for unsubstituted PAHs (Safinowski and Meckenstock, 2006) while other studies purport a carboxylation mechanism involving a recently characterized naphthalene carboxylase belonging to the UbiD family of carboxylases (Bergmann et al., 2011;

Mouttaki et al., 2012; Meckenstock et al., 2016). Carboxylation of unsubstituted PAHs involves high activation energy, so it may be limited to higher-energy reducing conditions such as nitrate reduction (Christensen et al., 2004). Methylation is generally believed to be a precursor reaction to fumarate addition which has been characterized in some studies (Safinowski and Meckenstock,

2006; Musat et al., 2009), before proceeding through succinate intermediates similar to those observed in toluene degradation (Meckenstock et al., 2016; Gieg and Toth, 2017). Zhang et al.

(2020) recently characterized a nitrate-reducing isolate closely related to Achromobacter denitrificans that produced both 2-phenanthroic acid and 2-methylphenanthrene (and postulated subsequent transformation to phenanthryl-2-methylsuccinate) from phenanthrene. Regardless of the activation process, these reactions appear to occur at the C2 position of unsubstituted PAHs, possibly due to electrophilic substitution reactions, radical intermediate formation (as the result of

C-H bond dissociation), or enzyme specificity for the 2-substituted structural isomer (Mouttaki et al., 2012).

In the case of methyl-substituted PAHs (such as 2-methylnaphthalene), studies have found evidence of a fumarate addition pathway involving naphthyl-methylsuccinate synthases (encoded by nmsABC), either through detection of naphthyl-methylsuccinate or assaying for the presence of nmsA (Annweiler et al., 2000; Marozava et al., 2018). Annweiler et al. (2002) found that under sulfate-reducing conditions, 2-methylnaphthalene was degraded through fumarate addition while in the same culture, only 2-naphthoic acid was produced from naphthalene. Several studies have 82

shown carboxylated PAHs from alkyl-substituted substrates without detection of the corresponding succinate derivatives indicative of fumarate addition (Annweiler et al., 2002;

Berdugo-Clavijo et al., 2012; Marozava et al., 2018; Toth and Gieg, 2018), indicating these succinate intermediates may occur at concentrations too low for the method of detection or that the long length of experimental incubations means certain sampling points simply miss the metabolite. All previously described activation processes inevitably converge to form the central intermediate naphthoyl-CoA or phenanthroyl-CoA (analogous to benzoyl-CoA in toluene degradation), which have been detected in their carboxylic acid forms in several studies and would represent the precursor to dearomatization (Zhang and Young, 1997; Safinowski and

Meckenstock, 2006; Musat et al., 2009; Mouttaki et al., 2012; Shin et al., 2019).

Metabolites of dearomatization are believed to be exclusively anaerobic and thus can serve as signature metabolites for evidence of anaerobic PAH biodegradation (Gieg and Toth, 2017).

Annweiler et al. (2002) found that naphthalene degraded by sulfate-reducers formed a 5,6,7,8- tetrahydro-2-naphthoic acid intermediate, where the ring furthest from the carboxyl group was hydrogenated first. In a study of methanogenic naphthalene degradation, Toth et al. (2018) detected a putative isomer of decahydro-2-naphthoic acid, which indicated that a carboxylated intermediate and dearomatization reactions were likely involved. Following dearomatization, ring opening occurs through hydrolytic cleavage to form (in the case of naphthalene) a proposed carboxylic acid-substituted cyclohexane with an adjacent hydroxybutyryl-CoA group (Weyrauch et al., 2020). Multiple rounds of dehydrogenase and thiolase reactions produce units of acetyl-

CoA and shorten the fatty acid chain before the remaining ring is also opened via a second hydrolytic reaction followed again by dehydrogenase and thiolase reactions.

While much has been learned about PAH biodegradation in the past 10-20 years, there remains a gap in providing supporting evidence of methanogenic PAH degradation activation mechanisms, the microorganisms involved, and the enzymes responsible for hydrocarbon transformation. In this study, we established a new enrichment culture of methanogenic PAH degraders. Using hydrocarbon-contaminated soil as the inoculum, culture enrichments were established with a cocktail of PAHs as carbon sources: naphthalene, 2-methylnaphthalene, and phenanthrene. By enriching a microbial community from soil contaminated with hydrocarbons, we aimed to further study the community structure and pathways involved in methanogenic PAH biodegradation.

5.2 Methods

The soil used as the inoculum in this experiment was obtained from a rural waste site contaminated with hydrocarbons, solvents, and sewage near Galahad, Alberta. The hydrocarbon load at this site and its general chemical features were not characterized. To prepare a methanogenic enrichment, 175 g of this soil were added to a 1 L glass bottle in an anoxic glove bag (headspace of 10% CO2 in N2 gas) containing 400 mL Pfennig medium (Appendix C), stoppered, and incubated at 30˚C in the dark for 400 days. A single incubation amended with PAHs was initially established to determine whether methanogenic activity would be observed from the environmental sample; similarly, a single incubation without added PAHs was also established.

In subsequent transfers, the slurry was pipetted from this initial enrichment inside of a glove bag and passed through 10 μm pore size filter paper to remove most of the sediment but still allow microorganisms to pass through.

Previous work with PAHs made use of HMN as an inert carrier to solubilize the PAHs and introduce them into the liquid phase, with the aim of increasing their bioavailability to the 84

microorganisms (Toth et al., 2018). Naphthalene, 2-methylnaphthalene, and phenanthrene were solubilized in 100 mL of degassed (anoxic) HMN, of which 20 mL were transferred to the enrichment cultures for a final experimental concentration of 1 mM of each hydrocarbon.

Unamended treatments received HMN with no dissolved PAHs. Following 400 days of enrichment with little methane production from the PAH-amended incubations compared to the

PAH-free control, an alternative substrate introduction method was used – the addition of a sorbent, Amberlite® XAD-7. Amberlite forms small, inert resin beads to which PAHs can be adsorbed (Morasch et al., 2001; Berdugo-Clavijo et al., 2012; Zhang et al., 2021). The benefit of this method was that rather than applying PAHs in an oil phase (HMN layer on top of the liquid),

Amberlite would sink to the bottom of the liquid phase which is where microbial biomass commonly accumulates; additionally, Zhang et al. (2021) observed improved rates of phenanthrene degradation when Amberlite was used as the PAH carrier compared to HMN.

To prepare the Amberlite beads, naphthalene, 2-methylnaphthalene, and phenanthrene were added to a 100 mL volumetric flask with dichloromethane (DCM) to create a working stock of 1 mM of each chemical. One hundred fifty milligrams of Amberlite were weighed into 60 mL serum bottles, to which 2.5 mL of the PAH stock solution were added; the DCM was allowed to evaporate in a fume hood for several hours to achieve full PAH adsorption. Bottles were supplemented with

Pfennig medium, then flushed with 10% CO2 in N2 gas before sealing and autoclaving. The enrichment culture was transferred (50% v/v) into these bottles for a final volume of 25 mL for further incubation, with a final aqueous hydrocarbon concentration of approximately 100 μM each.

Incubations were periodically sampled to assess changes to the microbial community (methods described in section 3.9), quantify methane production (section 3.3.2), and analyze for

hydrocarbon loss or metabolite production (section 3.8). Statistical tests conducted on some of the data are described in section 3.13.

5.3 Results

5.3.1 Methane production and degradation of substrates

Methane production in the initial soil enrichment cultures (Figure 5-1) was considerable, however at no point did the incubation with PAHs produce more methane than the substrate- unamended control. PAH-amended incubations from the first transfer (Figure 5-2) also did not produce more methane than the unamended control; in fact, the PAH-containing treatment produced 6-times less methane than the unamended incubation. Additionally, these transfers resulted in 120-fold reduction in methane production compared to the initial enrichments. Further transfers (second, third, and fourth; Figure 5-3) also produced considerably less methane which never exceeded 1 μmole total throughout the remainder of the experimental monitoring period.

6000

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Hydrocarbon loss was quantified during the fourth round of transfers from triplicate incubations with PAHs adsorbed to Amberlite. When sampled after 30, 90 and 240 days of incubation

(analysis between 90 and 240 days was not possible because of lab closure due to COVID-19), 10 to 80% hydrocarbon loss was observed across all three substrates relative to the sterile controls

(Figure 5-4). Phenanthrene demonstrated the least hydrocarbon loss relative to the sterile control, with only 61% loss by day 240 which corresponded to 0.94 ± 0.07 μmoles. The percent loss of 2- methylnaphthalene was greater than that for phenanthrene; by day 240, 78% of 2- methylnaphthalene was depleted relative to the sterile control (0.53 ± 0.02 μmoles). Naphthalene was the most extensively diminished relative to the sterile control, with 0.24 ± 0.01 μmoles lost by day 240, representing 81% depletion (Figure 5-4). Additionally, 0.26-0.40 μmoles were lost from sterile controls in all treatments between day 30 and day 240 however this is accounted for in the previous calculations. Hydrocarbon loss in these treatments did not result in a stoichiometric production of methane. The mineralization of 0.24 moles naphthalene, 0.53 μmoles of 2- methylnaphthalene, and 0.94 μmoles of phenanthrene should have resulted in 13.0 μmoles of CH4 based on the stoichiometric reactions for the methanogenic degradation of these PAHs (Table 5-

1), while only 0.8 μmoles CH4 were measured (Figure 5-3). Thus, additional avenues were explored as to the fate of carbon in the PAHs. Methodologies for metabolite analysis are described in section 3.8.2 and the results are outlined in section 5.3.2. Additional intermediates that were also sought via HPLC but not detected (approximately 100 μM detection limit; methods described in section 3.5.1) included acetic acid, propionic acid, butyric acid, fumaric acid, succinic acid, pyruvic acid, benzoic acid, pimelic acid, crotonic acid, and cyclohexane carboxylic acid. Live treatments were also surveyed for nitrate, sulfate, and iron(II) (methods described in section 3.5.2

and 3.6) which could possibly have been carried over from the initial enrichment, however none of these were detected.

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0 Naphthalene 2-Methylnaphthalene Phenanthrene

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Figure 5-4: PAHs degraded in a time-course incubation (T4), displayed as the percent of hydrocarbons remaining relative to the sterile controls (100%) (not shown). By day 240, naphthalene was depleted by 81%, 2-methylnaphthalene by 78%, and phenanthrene by 61%. Statistically significant loss of hydrocarbons was observed by day 240 (** p-value ≤ 0.01, *** p- value ≤ 0.001) relative to the sterile controls as determined by t-tests. Error bars ± SEM of three replicates.

Substrate Stoichiometry

naphthalene C10H8 + 8 H2O → 6 CH4 + 4 CO2

2-methylnaphthalene C11H10 + 8.5 H2O + → 6.75 CH4 + 4.25 CO2

phenanthrene C14H10 + 11.5 H2O + → 8.5 CH4 + 5.75 CO2

5.3.2 Metabolite analysis

Following PAH quantification, samples were concentrated, TMS-derivatized and surveyed for the presence of predicted metabolites including n-naphthoic acid, n-naphthol, n- methylnaphthalene, dimethylnaphthalene, 1,2,3,4-tetrahydro-2-naphthoic acid, 5,6,7,8-tetrahydro-

2-naphthoic acid, and n-phenanthrenecarboxylic acid (Table 5-2). Succinate derivatives from fumarate addition that may have formed from the PAHs were also sought based on predicted fragment ions as no authentic standards were commercially available. A peak with a retention time of 44.2 minutes matching the fragment ion profile of naphthoic acid was detected (Figure 5-

5; having fragment ions at m/z 229 and 244), and subsequent analysis of authentic standards distinguished it from 1-naphthoic acid (43.4 min). Instead, this metabolite was positively identified as 2-naphthoic acid (44.2 min); this peak was not observed in any sterile controls.

Quantification of this metabolite revealed a relatively constant amount throughout the experiment of 0.12-0.20 μmoles (Figure 5-6). This amount represented approximately a 23% conversion of the total two-ringed substrates lost by the end of the experiment. No other metabolites sought were detected (Table 5-2).

TMS- Molecular Metabolite derivatized Substrate Metabolite weight or isomer diagnostic (g/mol) detected? ions (m/z) 1-naphthoic acid 172.18 229 & 244 - Naphthalene 1-naphthol 144.17 201 & 216 - 1-methynaphthalene 142.20 142 - 2-naphthoic acid 172.18 229 & 244 + 2-naphthol 144.17 201 & 216 - 2-methylnaphthalene 142.20 142 +* 2,6-dimethylnaphthalene 152.20 152 - Naphthalene or 2- Naphthyl-2-methyl methylnaphthalene 258.27 387 & 402 - benzylsuccinic acid

1,2,3,4-tetrahydro-2-naphthoic 176.21 233 & 248 - acid 5,6,7,8-tetrahydro-2-naphthoic 176.21 233 & 248 - acid 2-phenanthrenecarboxylic acid 222.24 279 & 294 - Phenanthrene 2-(phenanthren-2- 308.33 437 & 452 - ylmethyl)succinic acid * already present as a substrate

) 0.30

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Figure 5-6: 2-Naphthoic acid quantified from PAH-amended live and sterile incubations. n=3, error bars ± SEM.

A second peak was found in the organic extracts from the live PAH-amended cultures that was not present in any sterile extracts (Figure 5-7A). This peak was found to be minimal at day

30 (a small, non-integrable peak at 48.8 minutes in live replicate 2), but by day 90 this peak was considerable (present in all 3 live replicates) before decreasing in relative abundance at day 120 and day 240 (Figure 5-8). Analysis of this peak revealed fragment ions at m/z 367 and 382

(denoted by black arrows; Figure 5-7B). Acting under the assumption these fragment ions represented the TMS-derivatized compound mass (M) and the corresponding M-15, the mass of this metabolite was calculated to be 310 (if derivatized once) or 238 g/mol (derivatized twice). As this peak did not match any of the standards surveyed, it could not be conclusively identified. It also did not match the calculated masses of any TMS-derivatized Pfennig medium components such as vitamins (Appendix C), ruling out the possibility that this compound originated from the culture medium.

Figure 5-7: Total ion chromatogram and mass spectral profiles of a metabolite formed in the live PAH-amended cultures. A peak with a retention time of 48.8 minutes was observed in all live incubations at after 90, 120 and 240 days of incubation but was absent at day 30 (A). TMS- derivatized diagnostic fragment ions of m/z 367 and 382 were identified in all samples with this peak (B).

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5.3.3 Microbial community analysis

PAH-amended methanogenic enrichments were analyzed for their microbial community composition based on 16S rRNA gene sequencing several times over two years of monitoring.

There were 861 distinct taxa identified in the analysis of these six samples. The microbial community composition in the soil inoculum shifted largely over this time (Figure 5-9). The original sample (inoculum) was dominated by aerobes such as Bacillus (31%) and facultative anaerobes like Porphyromonadaceae (22%), but by the fourth transfer, the PAH-containing enrichment was largely made up of Pelobacter (28% relative abundance), Desulfovibrio (11%) and Clostridium (5%). Mid-point sampling of unamended and PAH enrichments and transfers

(day 150 and 650) revealed almost identical microbial communities with similar taxa and distribution of microorganisms; no taxon represented more than 12% relative abundance of the total community. Methanogens such as Methanobacterium and Methanosarcina were detected at 94

day 150 in the initial enrichments (2-5%) and were not detected in similar relative abundance in later analyses, aligning with the decreased detection of methane upon repeated culture transfers

(section 5.3.1).

5.4 Discussion

5.4.1 PAH transformation and methane production

In the case of unsubstituted PAHs such as naphthalene, anthracene, and phenanthrene, there have been studies successfully demonstrating their biodegradation even under low energy-yielding conditions such as methanogenesis (Berdugo-Clavijo et al., 2012; Wan et al., 2012; Zhang et al.,

2012; Jiménez et al., 2016; Toth and Gieg, 2018; Toth et al., 2018). Zhang et al. (2021) compared the use of Amberlite, HMN, and no carrier to introduce phenanthrene to an iron(III)-reducing culture; they found that the Amberlite approach resulted in rapid phenanthrene removal (less than

10 days) while HMN facilitated only 80% loss and with no carrier only 70% was removed over 30 days of monitoring. The use of Amberlite resins to adsorb PAHs facilitates the slow introduction of insoluble and recalcitrant compounds to the enrichment while avoiding the cytotoxic effects of direct substrate addition (Foght, 2008). Data from this present study indeed indicated that hydrocarbon loss occurred (Figure 5-4) when PAHs were supplied in Amberlite beads, however stoichiometric conversion to methane was not observed (Figure 5-3). Similar non-stoichiometric loss of aromatic hydrocarbons in conjunction with minimal methane production has been reported in other biodegradation studies (Reinhard et al., 2005; Ye et al., 2018); both studies also surveyed for the presence of alternative reduced electron acceptors and found their concentrations could not explain the amount of hydrocarbon loss observed. Sulfate is believed to be inhibitory to methanogenesis at concentrations as low as 30 μM (Lovley and Klug, 1986), therefore incubations were also surveyed for the presence of other electron acceptors such as nitrate, iron(II), and sulfate, however these were not detected. Putative β-oxidation fatty acids or possible dead-end intermediates based on the degradation pathways outlined by Meckenstock et al. (2016) were also sought via HPLC (listed in section 5.3.1), however none were found. Chang et al. (2006) reported 96

that even when 2-bromoethanesulfonate (BES) was added (an inhibitor of methanogenesis), phenanthrene loss was still observed despite methane production ceasing. Reports of near stoichiometric conversion of methane from 2-methylnaphthalene or 2,6-dimethylnaphthalene in as little as 100 days do exist (Berdugo-Clavijo et al., 2012), likely due to the increased reactivity of methyl-substituted PAHs compared to those that are non-substituted. In the present study, we did observe the formation of a predicted intermediate in the form of 2-naphthoic acid as well as a putative novel metabolite (discussed further in section 5.4.2), indicating that some hydrocarbon bioconversion did occur.

Chang et al. (2006) established methanogenic cultures from river sediments that degraded naphthalene and phenanthrene but reported unusual evidence of PAH degradation. When methanogenesis was inhibited through the addition of BES, some PAH loss was still observed, albeit less than in the uninhibited incubations. They noted that some bacteria (likely syntrophs) were also eliminated and postulated that the sediment itself could be a source of electron acceptors for microbes carrying out higher energy electron accepting processes, such as iron(III)- or sulfate- reducers. While other studies have also achieved PAH degradation with sediment-containing enrichments (Zhang and Young, 1997; Chang et al., 2002; Morris et al., 2014), the findings of

Chang et al. (2006) underscores the necessity to achieve sediment-free incubations in assessing the metabolites of degradation while also removing confounding factors such as secondary organics or substrates, unknown abiotic factors, and minerals. Continually enriching and transferring these cultures removes sediment and ensures dilution of non-hydrocarbon degrading microorganisms. However, this approach is a double-edged sword, as numerous studies have attested to the need for sediments or co-culture conditions to support a diverse microbial population; Stewart (2012) highlighted the difficulties associated with cultivating “unculturable” 97

bacteria in the laboratory. From coculture dependency to substrate sorption kinetics, there may have been several factors that influenced poor methane production in the later transfers (Figure 5-

3) observed in the present study as sediments were removed and the culture was diluted through successive transfers.

5.4.2 Metabolite analysis

A metabolite positively identified as 2-naphthoic acid was found in live incubations but was not detected in sterile controls (Figure 5-5). The concentration of this metabolite increased only marginally over the course of the experiment, from 0.12 ± 0.03 μmoles at day 30 to 0.18 ± 0.00

μmoles by day 240. The detection of 2-naphthoic acid in the PAH-degrading enrichments is in line with other studies which have identified naphthoic acids but not fumarate addition products such as naphthyl-2-methylsuccinic acid or 2-(phenanthren-2-ylmethyl)succinic acid (Annweiler et al., 2002; Bergmann et al., 2011; Berdugo-Clavijo et al., 2012; Marozava et al., 2018) in PAH- amended cultures. The 2-naphthoic acid metabolite detected in this experiment could have arisen from either naphthalene via direct carboxylation (Zhang and Young, 1997; Mouttaki et al., 2012;

Eberlein et al., 2013) or from 2-methylnaphthalene degradation (via fumarate addition to naphthyl-

2-methylsuccinic acid that is then converted to 2-naphthoic acid; Annweiler et al., 2000, 2002,

Marozava et al., 2018). Naphthyl-2-methylsuccinic acid was sought but not detected at any sampling point. As losses of 81% naphthalene and 78% 2-methylnaphthalene were observed by the end of the experiment without corresponding production of methane, either or both compounds could have served as the parent substrate. Previous work seeking metabolites in anaerobic hydrocarbon-degrading cultures indicated up to a 1000-fold decrease in concentration of a metabolite relative to the parent compound (Elshahed et al., 2001; Fowler et al., 2012). In the present experiment, however, the 2-naphthoic acid formed represents a relatively high rate of 98

bioconversion (approximately 23% of the available 2-ringed substrates) which may indicate accumulation. Methane was negligibly produced in this experiment and CO2 production could not be measured (due to CO2 already being present in the headspace); therefore, it is possible that these

2-ringed substrates were converted into a dead-end product (other than the fatty acids surveyed in section 5.3.1) such as 2-naphthoic acid or another downstream intermediate. The findings of this experiment are in line with results observed by Chang et al. (2006), where PAH loss compared to sterile controls was observed despite little to no methane production. The authors postulated that the initial mechanism of PAH activation still occurred, however due to a metabolic backlog could not be fully converted to methane. In the present study, the processes of dilution and removing sediments may have eliminated crucial organism(s) necessary for the complete biotransformation via the methanogenic pathway.

A second metabolite was detected in live incubations only and increased in abundance over the course of the experiment (maximally at day 90 before decreasing; Figure 5-8). This compound was not detected in any sterile controls and did not match any of the predicted metabolites surveyed

(Table 5-2). From the fragment ion pattern (Figure 5-7B), diagnostic ions at fragment ions m/z

367 and 382 were observed which are consistent with a parent chemical mass of 310 g/mol (if the compound was TMS-derivatized once, i.e. one hydroxyl or carboxylic acid group) or 238 g/mol

(if the compound was TMS-derivatized twice, i.e. two hydroxyl or carboxylic acid groups). Large dearomatized intermediates such as 1,2,3,4-tetrahydro-2-naphthoic acid or 5,6,7,8-tetrahydro-2- naphthoic acid were sought but not detected, nor were carboxylation or fumarate addition products of phenanthrene. No obvious or appropriate structures matching these masses were found after a survey of the available literature. It also did not match the approximate mass (TMS-derivatized or underivatized) of any medium components (Appendix C) or reasonably predictable products of 99

Amberlite XAD-7 (an acrylic ester resin) breakdown (Domínguez et al., 2011). Amberlite is not expected to be friable in a manner that would make it susceptible to microbial degradation, and it has been used in similar PAH biodegradation studies previously with success (Morasch et al.,

2001; Berdugo-Clavijo et al., 2012; Zhang et al., 2021).

Of the possible substrates available for transformation (naphthalene, 2-methylnaphthalene, and phenanthrene), the most likely conclusion was a structure that had been TMS-derivatized twice thereby having a molar mass of approximately 238 g/mol. Structures hypothesized to meet these requirements, and which could have conceivably arisen from the substrates in question, are shown in Figure 5-10. Regardless of whether naphthalene underwent methylation followed by fumarate addition (Safinowski and Meckenstock, 2006; Musat et al., 2009) or direct carboxylation (Zhang and Young, 1997; Bergmann et al., 2011; Mouttaki et al., 2012), these initial reactions are universally expected to involve naphthoyl-CoA or phenanthroyl-CoA as intermediates, dearomatized structures, or hydrogenated rings with associated diacids as they proceed through modified β-oxidation intermediates (Gieg and Toth, 2017; Weyrauch et al., 2020). As discussed in Chapter Four, acidification of samples as part of the organic extraction process would hydrolyze any coenzyme A thioester linkages to generate carboxylic acids in their place, so it is difficult to be certain how far along metabolically these intermediates proceed with the analytical techniques used.

100

Hypothesized metabolite A Hypothesized metabolite B O O OH

OH O OH

O OH

Hypothesized metabolite C O OH

Figure 5-10: Possible structures of a metabolite detected in GC-MS analysis arising naphthalene or 2-methylnaphthalene transformation (A and B) or phenanthrene transformation (C) from Figure 5-7. (A) IUPAC: 2-[(2E,4E)-5-carboxypenta-2,4-dien-1-yl]cyclohexane-1-carboxylic acid. Chemical formula C13H18O4 and molecular weight 238.28 g/mol. (B) IUPAC: 2-(carboxymethyl)- 1,2,3,4,5,6,7,8-octahydronaphthalene-1-carboxylic acid. Chemical formula C13H18O4 and molecular weight 238.28 g/mol; the exact location of the carboxylic acid, carboxymethyl group, and double bond are unknown. (C) IUPAC: 2-hydroxyphenanthrene-1-carboxylic acid. Chemical formula C15H10O3 and molecular weight 238.24 g/mol; the exact location of the carboxylic acid and alcohol groups are unknown.

Metabolite A is a dicarboxylic acid which could conceivably form as a result of ring opening, and it is similar to a coenzyme A thioester-associated structure proposed by Weyrauch et al. (2020) who examined downstream degradation intermediates after naphthalene underwent carboxylation, dearomatization, and ring opening. These authors note this was a predicted structure they hypothesized based on an unexplained peak observed in LC-MS analysis. Metabolite B could possibly be an intermediate of dearomatization similar to a 1-hydroxy-octahydro-2-naphthoic acid described by Meckenstock et al. (2016) and Gieg and Toth (2017), while metabolite C could theoretically arise from phenanthrene activation.

101

While this metabolite did not match molecular weights of the few metabolites of anaerobic

PAH biodegradation reported in the available literature, it did increase in concentration from zero at day 30 maximally to day 90 before decreasing through the rest of the experiment (Figure 5-8).

This potentially novel intermediate likely does not represent a dead-end intermediate such as decahydro-2-naphthoic acid which has been reported in other studies (Meckenstock et al., 2016;

Toth et al., 2018), since this intermediate varied in abundance despite 2-naphthoic acid concentration remaining relatively constant.

5.4.3 Microbial community analysis

Of the 17000 to 35000 reads annotated in the six samples in Figure 5-9, the relative abundance of methanogens varied widely. In the soil inoculum, 0.23% of total reads belonged to methanogens; by day 150, methanogens totaling 8.56% relative abundance were found in the unamended enrichments and 12.71% relative abundance in the PAH-amended treatment. This again changed by day 650 in the subsequent transfers (T4), with 0.58% detected in the first unamended transfer and 1.25-1.54% in the PAH-amended transfers with and without the sorbent material, respectively (Figure 5-9). While this seems like a very small abundance of methanogens, this was in line with our previously characterized TOLDC studies (Chapter 4) where stoichiometric hydrocarbon degradation and methane production occurred despite very small relative abundances of methanogens. However, this contrasts with the results observed in those experiments, where decreasing abundance of methanogens resulted in greater methane production and hydrocarbon degradation. Here, increasing hydrocarbon degradation correlated with decreased methane production and decreased abundance of methanogens.

The enrichment of Betaproteobacteria has been reported in multiple anaerobic PAH degradation studies (Berdugo-Clavijo, 2015; Folwell et al., 2016; Ghosal et al., 2016; Zhang et 102

al., 2020). Some Betaproteobacteria were observed in the initial inoculum (5%), fewer were found in the enrichments after 150 days (1.6-1.8% relative abundance), and none were observed in subsequent analyses at day 650. It appears that in this study, members of the Betaproteobacteria did not contribute to PAH loss.

Pelobacter increased by the fourth transfer more than any other taxon (27.9% relative abundance) followed by Desulfovibrio (10.5%), and Clostridium sensu stricto 13 (4.5%).

Pelobacter was detected in sulfate-reducing enrichments biodegrading hexadecane and phenanthrene (Shin et al., 2019) and certain species are capable of anaerobic fermentation of acetylene (Schink, 1985). This organism has been characterized to have diverse metabolic capabilities including indirect reduction of solid iron in biofilm associations with Desulfovibrio and methanogens through a sulfide/S0 cycling pathway (Vigneron et al., 2016), nitrogen fixation during anaerobic hydrocarbon degradation (Akob et al., 2017; Davidova et al., 2018), 2,3- butanediol fermentation (Aklujkar et al., 2012), and acetogenesis from H2 (Rabus et al., 2016b).

Pelobacter acetylenicus is the only known bacterium capable of anaerobic activation of triple- bonded hydrocarbons, carrying out an oxygen-independent hydrogenation of acetylene to acetaldehyde then further to acetate and ethanol (Schink, 1985; Abbasian et al., 2015; Davidova et al., 2018). While triple-bonded hydrocarbons are not common in the environment (Spormann and Widdel, 2000), it is unknown if this enzyme could hydrogenate double bonded hydrocarbons or aromatic rings. Pelobacter acidigallici is unable to metabolize acetylene; instead it grows on trihydroxy-substituted benzoates such as gallic acid and produces acetate and CO2 (Schink, 2006), which would be amenable to the syntrophic associations necessary for methanogenesis to occur.

Members of the Pelobacter genus are frequently considered as obligate anaerobic fermenters, as most species characterized are deficient in most or all respiratory enzyme encoding genes (similar 103

to Smithella; Davidova et al., 2018). Pelobacter, or its family lineage Desulfuromonadaceae, has been identified in numerous fermentative or methanogenic hydrocarbon degrading communities

(Ramos et al., 2013; Berdugo-Clavijo, 2015; Rabus et al., 2016b; Toth and Gieg, 2018) thus we can speculate that this phylotype plays a role in PAH biodegradation in the present study. However, its mechanistic role remains uncharacterized and warrants further study.

The role of Desulfovibrio in aromatic hydrocarbon degradation was discussed at length in

Chapter Four. Desulfovibrio became isotopically labelled in a study examining naphthalene degradation under methanogenic conditions (Toth et al., 2018) and likely uses fatty acids or acetate as electron donors in a syntrophic oxidation mechanism for the downstream processes of fermentative metabolism (Fowler et al., 2014; Rabus et al., 2016b; Vigneron et al., 2016). Meyer et al. (2013) found that Desulfovibrio genes were differentially expressed when growing in a syntrophic co-culture with a methanogen partner when compared to growth alone in pure culture, which supports the position of Christensen et al. (2004) who found naphthalene oxidation under such conditions was only thermodynamically favorable when products are continuously removed by methanogens.

In addition to Desulfovibrio, Clostridium sensu stricto 13 was found to be enriched by day

650 in all transfers (4.5-12.0% relative abundance; Figure 5-9). Clostridiaceae or Clostridium has been found in several studies of naphthalene and 2-methylnaphthalene degradation, either through

16S rRNA gene sequencing or analysis of isotopically labelled fractions, and has frequently been associated with Desulfovibrio (Berdugo-Clavijo et al., 2012; Jiménez et al., 2016; Martirani-Von

Abercron et al., 2016; Toth et al., 2018). Similar findings were observed by Fowler et al. (2014), who identified Clostridium as a key microbe involved in toluene degradation. While these studies indicated that Clostridium is not likely the hydrocarbon-activating taxon, it has been strongly 104

implicated in the downstream metabolism of intermediates of hydrocarbon metabolism. This organism may be playing a similar role in our PAH-amended enrichments, but more investigation is needed to verify this and the roles of all the taxa identified in the enrichments.

5.5 Conclusions

The results of this study indicated that PAH biotransformation occurred under methanogenic conditions (e.g., in the absence of an added electron acceptor), however I was unable to conclusively link this process to the expected corresponding methane production in later transfers.

2-Naphthoic acid was detected in all live treatments which supports carboxylation as the activation mechanism of naphthalene or 2-methylnaphthalene having formed naphthyl-2- methylbenzylsuccinic acid before further transformation to a dead-end naphthoyl-CoA (similar to what was observed in Chapter Four); however, succinate derivatives indicative of fumarate addition to 2-methylnaphthalene were not detected in the present study. A putative novel metabolite was detected in live treatments, but it did not match the established masses or fragment patterns of metabolites available in the literature or predicted to occur as intermediates of PAH biodegradation. Pelobacter, Desulfovibrio, and Clostridium were identified as likely being involved in PAH biotransformation due to their increasing abundance (Figure 5-8). Ultimately, evidence has been presented supporting biotransformation of naphthalene, 2-methylnaphthalene, and phenanthrene in a methanogenic enrichment culture but more investigation is needed.

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Chapter Six: The effect of a sorbent matrix on recovery of microorganisms from contaminated groundwater

A version of this paper has been submitted for peer-review to the journal Microorganisms, with me as the lead author. Some of the initial work described in this chapter was performed by researchers that preceded me, but I conducted all the microcosm experiments including associated sequencing analyses, hydrocarbon sampling, and chemical analyses. In addition, I processed and interpreted most of the data and wrote the manuscript, which subsequently was reviewed, edited, and accepted by all coauthors. To be specific about the other contributors to this work, Dr.

Courtney Toth performed 16S rRNA gene sequencing and analysis of the field samples in sections

6.2.1 and 6.3.1. Drs. Victoria Collins and Paolo Mussone provided groundwater samples and pouches filled with the various sorbent materials; additionally, Dr. Collins and Arantxa Pino

Persico provided access to and equipment for preparing Tenax-TA samples for scanning electron microscopy imaging. Dr. Lisa Gieg provided funding, supervision, and manuscript editing.

6.1 Introduction

Monoaromatic hydrocarbons found in polluted environments, such as benzene and toluene, are chemically unreactive due to their non-polar nature and aromatic character (Widdel and Musat,

2010). Many microorganisms in hydrocarbon-contaminated environments can metabolize aromatic hydrocarbons as their carbon and energy sources, thus contributing to site remediation processes. The microbial degradation of aromatic hydrocarbons is well characterized under aerobic conditions (Jindrová et al., 2002; Fahy et al., 2006; Widdel and Musat, 2010). Anaerobic hydrocarbon biodegradation pathways have also been characterized to varying extents in recent years and involve different activation reactions in the absence of oxygen (Foght, 2008). While many polycyclic aromatic hydrocarbons are recalcitrant under anoxic conditions (with a few

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notable exceptions e.g. Meckenstock et al., 2016 and Toth et al., 2018), several studies have shown that monoaromatic hydrocarbons such as toluene can be metabolized when coupled with available anaerobic electron acceptors or under methanogenic conditions (Lovley and Lonergan, 1990;

Beller et al., 1992; Biegert et al., 1996; Fowler et al., 2012, 2014). Benzene biodegradation has also been reported under a variety of anoxic conditions (Lovley et al., 1995; Burland and Edwards,

1999; Ulrich and Edwards, 2003; Abu Laban et al., 2010; van der Zaan et al., 2012; Luo et al.,

2016; Keller et al., 2018), though the mechanism(s) catalyzing its degradation are still not fully understood.

The heterogeneity and diversity of subsurface groundwater environments constitute a major challenge in assessing the in situ role of microorganisms for hydrocarbon remediation. Most degradation studies focus heavily on the activity of planktonic cells, however sessile or biofilm- associated cells are likely more important in subsurface environments. Microorganisms in groundwater ecosystems can be attached to solid matrices or exist planktonically; several reports have estimated that only a small fraction (0.058-0.22%) remains unattached (McMahon and

Parnell, 2014), aligning with the recent estimate that up to 80% of all microbial life across major global habitats is surface-attached (Flemming and Wuertz, 2019). In nature, communities of diverse microorganisms typically work together in biofilms or aggregates to carry out metabolic processes (including hydrocarbon degradation) and nutrient cycling either synergistically or syntrophically (Morris et al., 2013; Gieg et al., 2014), thus capturing these aggregate communities from different environments can allow for a deeper understanding of their ecosystem functioning.

Collecting groundwater samples from monitoring wells is the simplest and most common way to identify microorganisms and their potential activities in fuel-contaminated sites (Reinhard et al., 2005; Sublette et al., 2006; Ahad et al., 2018). This method of sample collection primarily 107

targets planktonic cells, and therefore may greatly underestimate the microbial communities inhabiting groundwater systems given the presumed predominance of surface-attached organisms.

Obtaining core samples from the subsurface is a more ideal method to capture surface-attached cells, however this can be logistically challenging and costly. An alternative to these methods is to use samplers containing adsorptive solid support material that can be emplaced into groundwater monitoring wells to collect microorganisms that are prone to form biofilms. This approach has been explored by several researchers who have shown the efficacy of using solid support materials to sample microorganisms in subsurface environments that may be contributing to hydrocarbon metabolism (Bombach et al., 2010). Materials such as granular activated carbon (Ahad et al.,

2018) and Bio-Sep® beads (comprised of activated charcoal in a polymer matrix; Williams et al.,

2013) have been successfully used to sample microorganisms from hydrocarbon-contaminated groundwater environments for microbial community member identification (Peacock et al., 2004;

Sublette et al., 2006; Biggerstaff et al., 2007; Busch-Harris et al., 2008). Materials such as the

Bio-Sep® beads have been incorporated into commercially available Bio-Trap® samplers

(Microbial Insights, Knoxville, USA). Increased appreciation of the role of surface-attached microorganisms highlights the need to assess the impact of these solid supports in groundwater ecosystems.

In this study, we sought to test a variety of other high surface area materials for their ability to trap and enrich putative fuel-degrading microorganisms from hydrocarbon-contaminated groundwater. This work is part of a larger study that is developing an “all-in-one” prototype groundwater sampling device equipped with multiple components for measuring different parameters of contaminated groundwater. As such, the sampler includes a component to collect and identify surface-attached microorganisms. Following a preliminary sorption test with soil 108

(Appendix Table B-1) and a subsequent field-based experiment (Figures 6-1 and 6-2, Appendix

Table B-2), one commercially available material, Tenax®-trapping agent (Tenax-TA), was found to capture the most microbial diversity. This sorptive material was then further evaluated through a series of laboratory microcosm experiments to assess whether the trapping agent enriched microorganisms capable of degrading common groundwater contaminants (benzene and toluene) under various electron-accepting conditions. We also sought to compare how microorganisms grown in this sessile manner differed from the corresponding planktonic communities. Our findings can be applied to future studies to better characterize passively trapped hydrocarbon- degrading microorganisms in field experiments and subsurface environments.

6.2 Materials and methods

6.2.1 Survey of different adsorbent materials

In preliminary microcosm tests, five adsorbent materials [zeolite, activated carbon, Mat540, diatomaceous earth (DE), and T-carbon (TC)] were surveyed for their use as microbial traps in soil and groundwater environments. Material specifications for each material are included as supplementary materials (Appendix Table B-1). Nylon pouches (60 μm mesh, approximately 1.5 x 2.0 cm) containing the porous trapping agents (125 ± 5 mg) were added to anoxic microcosms containing 5 g soil and 30 mL sterile distilled water to simulate an environmental sampling well.

After six days of incubation at room temperature, DNA was extracted from biomass adsorbed to the trapping agents as described below and recoveries were quantified by fluorometry. Based on greater DNA recoveries (Appendix Table B-1), DE and TC were selected for field testing in a hydrocarbon-contaminated aquifer located near Saskatoon, Saskatchewan, Canada. A third material, Tenax-TA (60-80 mesh, 25550; Restek, Bellefonte, USA), was also included in this field trial given its known ability to adsorb hydrocarbons and its moderate sorption surface area (35 to 109

40 m2/g), which is a desirable feature to facilitate colonization by microorganisms (Dettmer and

Engewald, 2002; Zhao and Pignatello, 2004). The hydrocarbon sorption efficiency of Tenax-TA was verified in sterile experimental trials prepared with distilled water and known amounts of two hydrocarbons of interest: benzene and toluene (Appendix Figure B-1).

To deploy the trapping agents at the Saskatchewan field site, the sorbents DE, TC, and

Tenax-TA were prepared as described above. Eighteen pouches were prepared for each sorbent, for a total of 54 samplers. These adsorbent traps were then attached with nylon thread onto the prototype sampling device so that they could be suspended into a test groundwater monitoring well. Duplicate samplers were deployed directly into the groundwater aquifer at depths of 3, 4, and 5 meters below ground surface. Once a month, for a total of three months, two replicates of each sorbent sampler were recovered at each experimental depth for molecular analysis. Samplers were stored immediately at -20℃ and transported on ice to the University of Calgary for DNA extraction and microbial community analysis, according to the methods detailed below. DNA recoveries for each pouch are reported in Appendix Table B-2. As described in section 6.3.1, the samplers containing Tenax-TA captured greater species richness, more high-quality reads, and greater microbial diversity than DE or TC. As such, Tenax-TA was chosen for further study in controlled laboratory incubations.

6.2.2 Experimental microcosms

In a third series of experiments, we sought to further assess the performance of Tenax-TA samplers in capturing both hydrocarbons and microorganisms. The purpose of this test was to differentiate surface-attached versus planktonic microorganisms, if any, and to determine whether the use of such sorptive traps can help to stimulate hydrocarbon biodegradation under different electron-accepting conditions. Groundwater slurries (containing some sand particles) from a 110

hydrocarbon-contaminated aquifer located near Stony Plain, Alberta, Canada were collected in

June 2018 and served as the inoculum for new microcosm experiments. Chemical analyses revealed the groundwater to be neutral in pH (7.72 ± 0.03) and contain 1.84 ± 0.07 mM total iron, a sulfate concentration of 0.74 ± 0.04 mM, and freshwater salinity (0.03%). No nitrate, nitrite, organic acids (such as acetate) or aromatic hydrocarbons were detected in these samples, however, previous groundwater analyses (in 2016) indicated benzene levels in this site reached 8.39 mg/L and toluene 0.377 mg/L (Kharey et al., 2020), both exceeding Canadian maximum allowable concentrations (0.005 mg/L and 0.024 mg/L, respectively; AEP, 2019). The lack of hydrocarbons detected in the present groundwater samples was likely due to volatilization that occurred during transportation and storage.

Table 6-1: Conditions and various controls of experimental microcosms. Each combination was conducted under either aerobic, nitrate-reducing, iron(III)-reducing, or sulfate-reducing conditions. A fifth condition included no additional electron acceptor to examine the effect of endogenous electron accepting processes.

Electron Matrix Amendment Treatment acceptor 5.6 μmoles benzene + live One of: 4.7 μmoles toluene heat-killed Tenax-TA 313 mM O2, live - unamended 10 mM NO3 , heat-killed 30 mM Fe(OH)3, 5.6 μmoles benzene + live 10 mM SO 2-, or 4.7 μmoles toluene heat-killed 4 No Tenax-TA none added live unamended heat-killed

Experimental microcosms were established in triplicate using 30 mL of groundwater-sand slurry transferred anoxically under N2 gas into sterile 60 mL glass serum bottles, closed with butyl rubber stoppers and sealed with aluminum crimps, creating a total of 120 microcosms (Table 6-1).

Half of the microcosms received a nylon pouch containing Tenax-TA beads (prepared as described

111

above) which were suspended from the rubber stopper using nylon thread (as shown in Appendix

Figure B-2). The remaining half were not provided a pouch to determine any selective pressures that might be exerted by Tenax-TA. Microcosms were amended with 0.5 μL each of benzene and toluene, representing 5.6 μmoles and 4.7 μmoles, respectively. Select bottles also received exogenous electron acceptors as per Table 6-1. Also established were a set of heat-killed controls

(prepared by autoclaving groundwater three times with 24-hour intervals to ensure the sterilization of spore-forming organisms) and hydrocarbon-free (unamended) controls to ensure all variations of possible experimental outcomes were accounted for. All microcosms were incubated at room temperature (21-22°C) in the dark for 80 days.

6.2.3 Analytical procedures

Benzene and toluene were measured weekly over 80 days of incubation by gas chromatography-flame ionization detection (GC-FID) and quantified based on calibration curves prepared from known concentrations of standards (Eastcott et al., 1988; Heath et al., 1993).

Operating parameters for hydrocarbon, methane, CO2, O2, nitrate, sulfate, and iron(II) detection are described in Chapter Three.

After 80 days of incubation, live microcosms were sacrificed for DNA extraction. DNA was extracted from the both the planktonic and sessile (Tenax-TA) fractions separately. Planktonic microorganisms were collected by centrifuging 5 mL of the liquid fraction at 15,000 x g for 10 minutes and discarding the supernatant. Tenax-TA pouches were aseptically transferred to sterile tubes, cut in half and vortexed for 1 minute with sodium phosphate buffer to detach microbial cells from the Tenax-TA matrix. The DNA extraction protocol and Illumina MiSeq® preparation methods are described in section 3.9. R was used for computing statistical analysis of microbial communities (vegan, ape, and picante) and associated graphics (ggplot2). Due to highly diverse 112

samples each containing hundreds of unique taxa, only the top ten or top five most abundant taxa from each sample are reported in sections 6.3.1 and 6.3.3; however, all taxa were included in diversity analyses.

Some aerobically incubated Tenax-TA pouches were preserved for biofilm visualization by scanning electron microscopy, according to the fixation and dehydration method outlined by

Lewandowski and Beyenal (2013) and described in section 3.12. Preserved samples were loaded on carbon tape, sputtered with gold, and visualized with a TESCAN VEGA3 scanning electron microscope at the NAIT Nanotechnology lab (Edmonton, Canada).

6.3 Results

6.3.1 Field testing of microbial trapping matrices

Sampling pouches containing DE, TC, or Tenax-TA materials were deployed at three different depths into a contaminated aquifer near Saskatoon, Canada in July 2017. Pouches were recovered from each depth in one-month intervals for a total of three months, and two pouches per time point were extracted for genomic DNA and their microbial community profiles analyzed by

16S rRNA gene amplicon sequencing. Similar microbial communities were captured across all sampling dates, depths, and materials, albeit in varying relative abundances. Tenax-TA had lower average DNA recoveries than DE or TC (Figure 6-1), however this material trapped more diverse taxa (measured by the Shannon index; Figure 6-1 and Appendix Table B-2) and had a greater proportion of unique or low abundance taxa (less than 0.2%) than DE or TC (Figure 6-2).

Rhodoferax dominated the microbial communities in DE and TC sampling pouches (mean relative abundances of 22.9 and 25.2%, respectively; Figure 6-2), while this taxon was significantly less abundant in Tenax-TA (7.7%, p-values ≤ 0.01 and ≤ 0.001, respectively). Putative hydrocarbon degraders were captured by all materials, including taxa such as Azoarcus, Desulfosporosinus, 113

Geobacter, Pseudomonas, and Thermincola. Due to superior taxonomic diversity, capture of a variety of hydrocarbon-degraders, and low bias for Rhodoferax (Figure 6-2), Tenax-TA was chosen for further analysis in the microcosm experiments.

10 3.5

)

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/ g

n 8

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t t

6 i

4 n

n N

2.5 n

g a

r 2

v A 0 2.0 Diatomaceous earth Tenax-TA T-carbon

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6.3.2 Hydrocarbon degradation and mineralization of electron acceptors

Here, Tenax-TA was screened for its ability to sorb hydrocarbons (and thus associate with potential hydrocarbon-utilizing microorganisms), differentiate surface-attached versus planktonic microorganisms, and whether its use in sampler traps would stimulate hydrocarbon biodegradation under different electron-accepting conditions. We began by assessing the ability of Tenax-TA to sorb aromatic hydrocarbons, compared to treatments without the traps in an abiotic experiment.

115

Toluene (77%) and benzene (53%) were sorbed significantly within one day of hydrocarbon addition (Appendix Figure B-1) compared to those without Tenax-TA (representing 100% of available hydrocarbons). This result was consistent with other studies of benzene and toluene adsorption (Baimatova et al., 2016).

Microorganisms in aerobically prepared microcosms (O2) completely oxidized both toluene and benzene after approximately 14 days of incubation, which was reproducible after refeeding bottles additional hydrocarbon substrate (Figure 6-3). They also demonstrated the fastest overall rate of hydrocarbon degradation (Table 6-3). Carbon dioxide production in aerobic microcosms with Tenax-TA was 80% higher than those without Tenax-TA (97.9 ± 0.7 compared to 55.0 ± 2.0

μmoles, respectively), a statistically significant difference (p-value ≤ 0.01) representing 49% and

28% of the predicted stoichiometric yield. The difference in this yield may be explained by conversion to biomass. In contrast, all microcosms incubated anoxically were capable of toluene degradation, but benzene metabolism was never observed during the 80-day monitoring period

(Appendix Figure B-3). Nitrate-reducing microcosms had the fastest rate of toluene degradation

(0.2 μmoles/day; Table 6-3) and displayed only a slightly significant difference in the extent of nitrate reduction between Tenax-TA and Tenax-TA free treatments (91.8 ± 5.5 and 113.2 ± 4.6

μmoles; Table 6-3). No nitrite was detected, suggesting that denitrification to N2 or dissimilatory

+ nitrate reduction to NH4 occurred in both Tenax-TA and Tenax-TA-free treatments (however

Azoarcus—the dominant taxon detected in nitrate-reducing treatments—is not known to carry out the latter process). Iron(III)-reducing microcosms natively contained 32.5 ± 1.3 μmoles of Fe(II) at the outset of the experiment (day 0). Of the predicted stoichiometric yield (Table 6-3), only 7% of this amount was realized in incubations with Tenax-TA by the end of the experiment. Tenax-

TA free treatments yielded 53% of the predicted stoichiometric yield, indicating that Tenax-TA 116

could in some way be inhibitory to iron(III) reduction or the growth of iron-reducing microorganisms in this experiment. The exact cause of these results is unknown. Interestingly, toluene degradation was unaffected (Appendix Figure B-3E) by the lack of iron reduction, suggesting an alternate anaerobic respiration pathway was coupled to the mineralization of this substrate. Sulfate-reducing treatments did not display a significant difference in sulfate reduction between Tenax-TA-containing and Tenax-TA-free treatments (32.4 ± 7.2 and 38.3 ± 0.6 μmoles respectively) and had the slowest rate of toluene degradation (0.1 μmoles/day). This corresponded to 77% stoichiometric yield in these microcosms (Table 6-3). No methane was produced in any of anoxic microcosms, even in no electron acceptor added treatments which could have facilitated methanogenic activity.

A 30 +Tenax-TA B 30 -Tenax-TA

)

l o

m 20 20



(

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a 10 10

y H 0 0 0 20 40 60 80 0 20 40 60 80

C 30 D 30

)

l o

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

(

r a

c 10 10

y H 0 0 0 20 40 60 80 0 20 40 60 80 Time (days) Time (days)

Toluene HK Toluene Benzene HK Benzene

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Figure 6-3: Hydrocarbon biodegradation profiles from aerobic microcosms with (A) and without Tenax-TA (B) as well as no electron acceptor (EA) added control microcosms with (C) and without Tenax-TA (D). Toluene and benzene were both degraded in aerobic microcosms within 14 days; no EA added microcosms required an initial lag of 40 days for toluene degradation to occur after which it was degraded within 14-20 days. Benzene was not degraded during the monitoring period. Gaps in the plots represent depletion and re-amendment of hydrocarbons. Live and heat- killed controls (HK) were established. Error bars depict the standard error of the mean of 3 replicates.

Table 6-2: Stoichiometric equations for the complete metabolism of benzene and toluene under - 2- the experimental electron-accepting conditions surveyed. Depletion of NO3 and SO4 (from nitrate and sulfate-reducing treatments, respectively) and accumulation of CO2 (from aerobic 2+ treatments), Fe (from iron-reducing treatments), and CH4 (from no EA added treatments) were monitored in this microcosm study and results are reported in Table 6-3.

Electron acceptor Stoichiometric equation Source (oxidized/reduced) Weelink et al., C H + 7.5O → 6CO + 3H O 6 6 2 2 2 2010 O /CO 2 2 Su and C H + 9O → 7CO + 4H O 7 8 2 2 2 Kafkewitz, 1994 Burland and C H + 6NO - → 6HCO - + 3N 6 6 3 3 2 Edwards, 1999 NO -/N 3 2 Su and C H + 7.2NO - + 7.2H+ → 7CO + 7.6H O + 3.6N 7 8 3 2 2 2 Kafkewitz, 1994 Burland and C H + 18H O + 30Fe3+ → 6HCO - + 30Fe2+ + 36H+ 6 6 2 3 Edwards, 1999 Fe3+/Fe2+ Lovley and C H + 21H O + 36Fe3+ → 7HCO - + 36Fe2+ + 43H+ 6 6 2 3 Lonergan, 1990 Burland and C H + 3.75SO 2- + 3H O → 3.75HS- + 6HCO - + 2.25H+ 6 6 4 2 3 Edwards, 1999 SO 2-/H S 4 2 Beller et al., C H + 4.5SO 2- + 3H O → 2.25H S + 2.25HS- + 7HCO - + 0.25H+ 7 8 4 2 2 3 1992 Symons and C H + 4.5H O → 3.75CH + 2.25CO 6 6 2 4 2 Buswell, 1933 CO /CH 2 4 Symons and C H + 5H O → 4.5CH + 2.5CO 7 8 2 4 2 Buswell, 1933

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Total Degradation Predicted Actual yield/consumption Redox Chemical hydrocarbons rate yield/ (μmoles) p-value condition monitored consumed (μmoles/ consumption (μmoles) day) (μmoles) +Tenax-TA -Tenax-TA

Aerobic CO2 31.0 0.4 199.9 97.9 ± 0.7 55.0 ± 2.0 **

Nitrate- NO - 18.8 0.2 135.4 91.8 ± 5.5 113.2 ± 4.6 * reducing 3

Iron(III)- Fe2+ 14.1 0.2 507.6 35.9 ± 1.7 267.0 ± 68.4 ns reducing

Sulfate- SO 2- 9.4 0.1 42.3 32.4 ± 7.2 38.3 ± 0.6 ns reducing 4

No EA CH 14.1 0.2 63.5 0.0 0.0 ns added 4 ns = not significant, * p-value ≤ 0.05, ** p-value ≤ 0.01

6.3.3 Microbial community analysis

DNA recoveries from sessile (Tenax-TA pouch-associated) samples were greater on average

than those recovered from planktonic samples (Appendix Table B-3 and Figure B-4). Sessile

fractions on average yielded 1.40 ± 1.00 ng/μL (n=30) with a maximum of 12.3 ng/μL, however

several sessile samples yielded DNA concentrations too low to quantify. All planktonic samples

had quantifiable DNA recoveries (maximum 0.48 ng/μL, on average 0.15 ± 0.02 ng/µL; n=63);

this represented a statistically significant difference from sessile DNA recoveries (p-value ≤ 0.05).

Trends in DNA recoveries followed that of the energetics of the electron-accepting processes,

where aerobic treatments yielded the most DNA and sulfate-reducing/no electron acceptor-added

treatments yielded the least amount of DNA. Concentrations were normalized to the amount of

material extracted (liquid or solid; Appendix Table B-3). 119

The composition of microbial communities from Tenax-TA containing, hydrocarbon- amended treatments diverged from the initial inoculum over the course of the experiment (Figure

6-4). The groundwater inoculum was primarily composed of Sediminibacterium and Rhodoferax.

The aerobic microcosms became enriched in taxa such as Rhodoferax, Saprospiraceae, and

Sulfuritalea. Nitrate-reducing microcosms were dominated to a large extent by Azoarcus with a smaller proportion of Candidatus Roizmanbacteria. Communities from iron(III)-reducing incubations varied considerably, consisting of taxa identified as Desulfoprunum,

Sedminibacterium, Azoarcus, Rhodoferax, and Candidatus Roizmanbacteria. Sulfate-reducing and the no electron acceptor added communities were similar to each other, with Desulfoprunum in highest relative abundance (36-53%) and Rhodoferax in second highest relative abundance (9-

10%; Figure 6-4).

The primary selective pressure driving community divergence in this study appeared to be the electron-accepting condition, as different redox potentials resulted in distinctive clustering compared to hydrocarbon treatment for most conditions (Figure 6-5). There was slight grouping within aerobic and nitrate-reducing treatments based on the presence or absence of hydrocarbons, while iron(III)-reducing, sulfate-reducing, and no electron acceptor added treatments all clustered similarly regardless of the presence of hydrocarbons.

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Figure 6-4: Top five most abundant microorganisms from each hydrocarbon-amended, Tenax-TA pouch-containing microcosms by treatment as determined using 16S rRNA gene amplicon sequencing. Total reads of three replicates were averaged and are displayed as percent relative abundance. Taxa that did not make the top five are grouped as “Other”. Sessile samples from the no electron acceptor added treatment could not be amplified through PCR and thus are not included.

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6.3.4 Visualization of Tenax-TA

Scanning electron microscopy was used to visualize microorganisms colonized on the

Tenax-TA beads following 80 days of incubation. A single Tenax-TA bead was zoomed in on to see the overall arrangement of microorganisms. In the sterilized control (Figure 6-6A), no microorganisms are visible on the surface of the beads. In the no hydrocarbon, live control (Figure

6-6B), few microorganisms are observed on the surface of the bead. In contrast, on the live,

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hydrocarbon-amended treatment (Figure 6-6C), numerous flaky structures that are believed to be biofilms are visible (structures larger than single bacterial cells), which were not observed on the sterilized or no hydrocarbon treatments. The presence of hydrocarbons appeared to be necessary for microbial colonization (Figures 6-6D and E). This notion is supported by the hydrocarbon consumption observed (Appendix Figure B-3), notable differences in the composition of microbial communities between sessile and planktonic fractions (Figure 6-4), and distinct grouping of samples from hydrocarbon amended and unamended treatments (Figure 6-5).

A B C

50 μm

D E

Figure 6-6: Microbially-colonized Tenax-TA. Panels A-C: false coloured scanning electron micrographs of microbe-colonized Tenax-TA beads retrieved from aerobic incubations. (A) The sterile control, (B) the live, hydrocarbon-free control, and (C) the live, hydrocarbon-amended treatment. Tenax-TA beads in panel C has visible microorganisms adhering to its surface while beads in panels A and B show little to no microbial colonization. Panels D and E: Tenax-TA pouches recovered from aerobic microcosms after 80 days of incubation. (D) A Tenax-TA pouch from a live, hydrocarbon-free microcosm and (E) a Tenax-TA pouch from a hydrocarbon-amended

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treatment. The Tenax-TA pouch from the hydrocarbon treatment is visibly darkened, presumably with biomass, compared to the unamended treatment.

6.4 Discussion

6.4.1 Matrix effects

Tenax-TA is an inert resinous polymer that forms porous beads. It is used in many industrial and analytical applications to purge and trap sorbent chemicals, particularly those with four or more carbons (Dettmer and Engewald, 2002). Its low affinity for water makes it an ideal matrix to reversibly trap airborne volatile organic compounds (VOCs) and aqueous environmental pollutants (Headley et al., 2001; Kuntasal et al., 2005). To the authors’ knowledge, Tenax-TA has not been used as a growth surface for microorganisms previously.

Biochars such as T-carbon and diatomaceous earth have similar hydrocarbon adsorption properties to Tenax-TA. Highly pyrolyzed biochars (like graphene) and those that were acid treated demonstrated strong but reversible gas adsorption kinetics for both benzene (75-91%;

Bansode et al., 2003) and toluene (93-98%; Lim et al., 2019). Sheshdeh et al. (2013, 2014) found nickel oxide-modified diatomaceous earth adsorbed up to 98% of benzene and 97% of toluene

(compared to 74% and 71% from unmodified diatomaceous earth, respectively). Raw diatomaceous earth is capable of adsorbing all BTEX hydrocarbons; studies have found in a BTEX mixture, xylenes were more readily sorbed to diatomaceous earth while benzene sorbed the least

(Aivalioti et al., 2010). Previous findings have shown that Tenax-TA is capable of adsorbing approximately 78% of benzene and 99% of toluene (Baimatova et al., 2016), which is comparable albeit slightly higher than our abiotic tests (53-77% adsorption; Appendix Figure B-1).

Biochars generally have high surface areas (up to 340 m2/g compared to Tenax-TA’s 35-40 m2/g) but are noted to have basic pH in water (9-12) which may have discouraged acidophilic or

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neutrophilic microorganisms from associating with this material in our field study (Demeyer et al.,

2001; Manariotis et al., 2015). Diatomaceous earth varies widely in its particle size due to its irregular shape and biotic origins, and ranges in surface area from 4 m2/g (untreated) up to 50 m2/g when decarbonated through acid treatment (Tsai et al., 2006; Benkacem et al., 2016). In a study of diatomaceous earth as a bacterial delivery vehicle, the authors found hollow inner structures of diatomaceous earth that sheltered bacteria, as well as some bacteria that sorbed to the surface of the material (Wang et al., 2012). This may be one reason why diatomaceous earth materials yielded high DNA recoveries in our field experiment (Figure 6-1). We found diversity capture was inversely correlated with DNA recovery for all materials tested, but that high surface area positively correlated with DNA recovery. Overall, Tenax-TA captured the highest diversity of microbial taxa, indicating its usefulness to trap microorganisms in field environments. Sorption of aromatic hydrocarbons was a secondary benefit that was shared between the three materials tested and helped facilitate capture of putative hydrocarbon degraders.

6.4.2 Biodegradation of hydrocarbons

Microorganisms within aerobic treatments were able to degrade benzene, and they did so within 14 days of incubation (Figure 6-3). In contrast, microbial communities incubated under anoxic conditions failed to oxidize any benzene over 80 days of monitoring (Appendix Figure B-

3). Given the levels of benzene detected at the Stony Plain site in 2016 (8.39 mg/L; Kharey et al.,

2020), we hypothesized that anaerobic benzene degradation would occur due to its historical exposure to the indigenous microbial population, however this was not realized within the 80-day incubation period across the anaerobic conditions tested. It has been observed in the few studies that successfully demonstrated anaerobic benzene degradation that long lag times (upwards of 300 days) can be necessary, while many other microcosm studies failed to achieve benzene degradation 125

even after long incubation times (Wilson et al., 1986; Edwards and Grbić-Galić, 1992; Nales et al., 1998). In the present work, additional incubation time may have been required to observe benzene biodegradation under anoxic conditions.

In contrast to benzene, toluene biodegradation occurred under all electron-accepting conditions (Table 6-3), with initial lag times generally corresponding to the strength of the associated redox potential (Appendix Figure B-3). It has been suggested that toluene can act as an inhibitor of benzene biodegradation in anaerobic co-culture studies, which may be another reason that benzene persisted in the microcosms (Nales et al., 1998). Across all the conditions tested, the presence of Tenax-TA did not appear to stimulate or inhibit toluene degradation. Microorganisms within live microcosms with and without Tenax-TA consumed toluene at approximately the same rate (Table 6-3). We hypothesize that hydrocarbon-degrading microorganisms were recruited to the Tenax-TA pouches (Figure 6-6), however they were also present planktonically in the liquid phase and were able to readily degrade solubilized aromatic substrates in both conditions.

6.4.3 Mineralization analysis

In the aerobic incubations, we observed a significant increase in CO2 produced from hydrocarbon-degrading microorganisms associated with Tenax-TA compared to incubations without Tenax-TA (Table 6-3). This amounted to 1.1 μmoles/day of CO2 produced from hydrocarbon-amended Tenax-TA microcosms compared to 0.6 μmoles/day in those without

Tenax-TA. Differentiated sessile microorganisms living in a biofilm may be a reason for such a marked difference in substrate mineralization, or residual small hydrocarbons that were soluble in the inoculum groundwater may have adsorbed to the Tenax-TA material, in turn increasing their bioavailability. It may also be possible that VOCs present in the air adsorbed to the Tenax-TA material before the pouches were placed in the microcosms, resulting in increased carbon 126

compounds available to the microorganisms (Korpi et al., 1998). As this trend in higher CO2 production was not observed in the sterilized controls, we assume that CO2 production was the result of biotic processes. No aromatic hydrocarbons or small organic acids (acetate, butyrate, or propionate) were detected during the initial water chemistry screening, so the exact reason for the varied CO2 production remains uncertain.

Nitrate reduction differed significantly between microcosms with and without Tenax-TA

(Table 6-3), while the differences in sulfate reduction were not significant. Iron(III)-reducing microcosms, however, demonstrated unusual patterns of iron(II) production. Iron(II) production was substantial in Tenax-TA-free treatments, but nearly negligible in those containing Tenax-TA

(Table 6-3). Tenax-TA seemed to inhibit iron reduction in this experiment, despite equivalent rates of toluene degradation regardless of Tenax-TA presence. This may be due to the low solubility of Fe(OH)3—compared to electron acceptors like O2, nitrate, and sulfate—which encouraged microorganisms to associate with the settled iron(III) at the bottom of the microcosms rather than with the hydrocarbon-associated Tenax-TA. A more soluble form of iron(III) could have resulted in better growth and adhesion of microorganisms to Tenax-TA. It is also worth noting that apparent iron sulfide was formed in the iron(III)-reducing, sulfate-reducing, and no electron acceptor added hydrocarbon-amended microcosms, as the sediment eventually turned black in the live treatments under all of these conditions. This likely resulted from a combination of reduction of natively present (or exogenously added) iron and sulfate (measured at 1.8 mM and

0.7 mM in the inoculum, respectively) and hydrogen sulfide production from sulfate reduction, which together precipitated black iron sulfide (visibly observed, not measured).

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6.4.4 Microbial community analysis

Using Tenax-TA provided a unique opportunity to characterize the microorganisms growing in sessile, aromatic hydrocarbon-associated environments and compare them with associated planktonic communities. The chemical and physical properties of Tenax-TA meant that hydrocarbons added were largely adsorbed and concentrated on the surface of the material, creating a micro-environment of high local concentration within the microcosm (Figure 6-6).

Microorganisms detected were presumed to not only colonize the Tenax-TA material but also be involved in the metabolism of aromatic hydrocarbons. This speaks to the importance of sessile versus planktonic metabolism in hydrocarbon biodegradation. We observed differences in the microbial communities associated with Tenax-TA in the presence of hydrocarbons compared to the unamended treatments. As such we can infer that there is a large range in metabolic capacity within the indigenous groundwater and subsurface microorganisms at the field site examined.

Sequencing analysis of the groundwater prior to its use as the inoculum revealed a native microbial community with many taxa known to be associated with hydrocarbon environments or hydrocarbon-degrading cultures (Figure 6-4). Sediminibacterium, some species of which are facultatively aerobic and associated with degradation of complex carbon compounds (Kim et al.,

2013; Berdugo-Clavijo and Gieg, 2014); and Polaromonas, an aerobe believed to be involved in in situ hydrocarbon degradation (Jeon et al., 2003; Mattes et al., 2008) have previously been detected in cold-climate, hydrocarbon-contaminated environments. Rhodoferax is a facultative aerobe with some species capable of iron reduction (Finneran et al., 2003), and is also commonly found in many contaminated sites (Aburto and Peimbert, 2011; Golby et al., 2012). After 80 days of incubation under aerobic conditions, the sessile communities within hydrocarbon-amended microcosms became enriched in Saprospiraceae (39.8% relative abundance) and Sulfuritalea 128

(15.1% relative abundance). Planktonic fractions from the same incubations were not similarly enriched in these two taxa. The absence of these taxa from hydrocarbon-free treatments (Appendix

Figure B-5) supports the hypothesis that they are involved in hydrocarbon degradation rather than colonization or adhesion. While Saprospiraceae has been implicated in degradation of complex organic compounds (McIlroy and Nielsen, 2014), it has not previously been associated with hydrocarbon degradation. Sulfuritalea on the other hand is commonly found in association with freshwater hydrocarbon contamination. Members of this genus are known to degrade aromatic compounds like benzoate, and have diverse metabolism to use oxygen, arsenate, or sulfur as a terminal electron acceptor (Sperfeld et al., 2018).

Nitrate-reducing, hydrocarbon-amended microbial communities became enriched in

Azoarcus (Figure 6-4) in both the sessile and planktonic phases. Several members of denitrifying

Azoarcus are known hydrocarbon degraders (Beller and Spormann, 1999; Achong et al., 2001).

The relative abundance of Azoarcus in the planktonic fraction (95.4%) outweighed its presence in the sessile fraction (58.4%). In contrast, hydrocarbon-free nitrate-reducing communities were very diverse, with 66.3% of taxa representing 0.2% or less of the total community composition

(Appendix Figure B-6). Iron-reducing, hydrocarbon-amended planktonic communities were mostly dominated by Desulfoprunum (29.5%; Figure 6-4). As iron(II) production was largely limited in Tenax-TA containing microcosms (Table 6-3), it is likely that the Tenax-TA-associated microenvironment was the dominated by sulfate-reducing metabolisms rather than predominance of iron(III) reduction. In fact, the iron(III) treated microbial communities shared many taxa with the sulfate-reducing and no electron acceptor added treatments, and known iron(III)-reducing hydrocarbon-degraders like Geobacter were only found in small abundances (less than 3%, and therefore are part of Other in Figure 6-4). In sulfate-reducing treatments, Desulfoprunum was also 129

present but at higher relative abundances (53.1% planktonic and 37.5% sessile). Desulfoprunum benzoelyticum (a benzoate oxidizer) cannot use Fe(OH)3 as an electron acceptor but instead uses sulfate (Junghare and Schink, 2015). Sulfate from the native groundwater was likely used by this taxon in the iron(III) amended treatments. Desulfoprunum reduces sulfate to sulfide, which at high concentrations can be inhibitory to acetate metabolism (Junghare and Schink, 2015).

The no electron acceptor added microcosms can only be tentatively discussed, as no PCR amplifiable DNA was recovered from the Tenax-TA traps and therefore, we were unable to obtain information regarding the composition of the sessile communities. From Figure 6-4, we saw that the communities from the planktonic no electron acceptor added and sulfate-reducing microcosms were very similar in taxonomic distribution with Desulfoprunum being the most abundant taxon

(36.6%) followed by Rhodoferax (9.2%), both of which fall within the relative taxonomic abundance observed in sulfate-reducing microcosms. The lack of methane production in these microcosms is supported by the absence of methanogenic taxa such as Methanosaeta and

Methanoculleus commonly found in toluene-degrading, methanogenic communities (Fowler et al.,

2012).

Aerobic, nitrate-reducing, and iron(III)-reducing treatments were all moderately enriched (5-

7%) in the candidate phylum Roizmanbacterium. This is a relatively recently discovered taxon however based on genome analysis it is believed to be facultatively anaerobic, involved in interspecies carbon transfer of small fatty acids, and important in overall carbon flow in groundwater systems (Geesink et al., 2020). It also possesses genes for the oxidation of various carbon compounds including necromass and cellulose. As a member of the Candidate Phylum

Radiation (CPR) superphylum, many of which are symbionts or exist in a sessile form,

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Roizmanbacterium was possibly involved in attachment or adherence in our microcosm system

(Geesink et al., 2020).

Aerobic communities diverged in composition considerably from the initial inoculum, as shown in the NMDS analysis (Figure 6-5). There is distinct clustering of hydrocarbon-amended communities (closed circles) compared to those that were hydrocarbon-unamended (triangles).

This indicates selection for different populations of microorganisms (i.e. putative hydrocarbon degraders) as a result of the hydrocarbon treatment. This trend is also readily apparent in the nitrate-reducing communities, where hydrocarbon-amended communities diverged greatly from the hydrocarbon-free communities. This is likely due to the explosive increase in the abundance of Azoarcus, which dominated all nitrate-reducing hydrocarbon-containing treatments.

Communities from iron(III), sulfate-reducing, and no electron acceptor added microcosms group closely with each other and possess more similar microbial taxa (Figure 6-4). Assuming an anoxic state within the subsurface and with minimal input of new nutrients, the microbial communities of the no electron acceptor added control microcosms would most closely match that which we would expect to see in the field.

6.5 Conclusions

Tenax-TA was an effective matrix to trap microorganisms in groundwater systems for analysis when exposed to aerobic, nitrate-reducing, or sulfate-reducing conditions. Iron-reducing treatments resulted in poor DNA recoveries, but this could be due to the insoluble form of iron that was used and predominance of sulfate-reduction in these microcosms instead. This study revealed biomass and DNA recoveries overall correlated with the strength of the associated electron acceptor, the presence of carbon substrates, and the length of experimental incubation. Ecological analysis of community data indicated that while the exact taxonomic composition could change 131

depending on hydrocarbon amendment, the overall diversity of the communities was not impacted by the presence of Tenax-TA. For use in our prototype water quality monitoring device, Tenax-

TA is an efficient, passive sampling matrix for trapping putative hydrocarbon-degrading microorganisms with the added benefits of its hydrocarbon adsorption properties.

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Chapter Seven: Conclusions

7.1 Research objectives and hypotheses

In this thesis, I surveyed two methanogenic cultures for the presence of key aromatic hydrocarbon activation metabolites (Chapters Four and Five) and genes (Chapter Four) while also identifying key microorganisms likely to be involved in hydrocarbon biodegradation. I also examined the feasibility of implementing a sorbent trap for use as a sampling device to recover microorganisms from contaminated groundwater (Chapter Six).

My initial hypothesis that alkyl-substituted aromatic hydrocarbons (other than toluene) would be activated through a fumarate addition mechanism was not supported by the data collected. Biotransformation of toluene and p-xylene was observed; however, the evidence did not indicate that fumarate addition occurred to p-xylene. Toluene did form benzylsuccinic acid as expected, but p-xylene formed only p-toluic acid and ethylbenzene was not transformed into any identifiable product. Naphthalene or 2-methylnaphthalene was transformed into 2-naphthoic acid and possibly a second, currently unknown metabolite. The fate of phenanthrene remains unclear.

Real-time gene expression assays of bssA in TOLDC revealed greater expression with toluene as the sole substrate relative to the unamended, and decreased expression in co-amended treatments with toluene and a second, non-target hydrocarbon, suggesting antagonistic effects on the bss gene when hydrocarbons other than toluene are present. Several microorganisms were identified as likely hydrocarbon-degraders (or associated with downstream hydrocarbon degradation) in all three chapters. Desulfosporosinus sp. Tol M and Desulfovibrio sp. SRL8083 were the dominant microorganisms in toluene-amended treatments in Chapter Four; Pelobacter, Desulfovibrio, and

Clostridium sensu stricto 13 were enriched in PAH-amended treatments in Chapter Five; and several microbes were identified as likely benzene or toluene-degraders with different redox 133

conditions in Chapter Six including Polaromonas, Sediminibacterium, Rhodoferax,

Saprospiraceae, Sulfuritalea, Azoarcus, and Desulfoprunum. The viability of using Tenax-TA as a sorbent microbial trap was thoroughly evaluated and found to exert slight selective pressures on the microbial community in aerobic and iron(III) reducing treatments, but did not influence the rate of benzene or toluene removal. Biomass recoveries were higher in hydrocarbon-amended treatments compared to those that were unamended.

7.2 Conclusions and future directions

7.2.1 Chapter Four

In Chapter Four, p-toluic acid was successfully identified as an intermediate of p-xylene transformation in a methanogenic toluene-degrading enrichment culture, though it may be a dead- end metabolite. Succinate derivatives of p-xylene were not detected in the present experiment, and the formation of p-toluic acid is not believed to have resulted from a carboxylation mechanism.

In co-amended treatments with toluene + p-xylene, p-toluic acid was found at significantly higher concentrations than in those with p-xylene alone, indicating the presence of toluene somehow influenced its production and possible accumulation. Persistence instead of the normally transient production of benzylsuccinic acid from toluene in the same co-amended treatments was observed, suggesting p-xylene or p-toluic acid influenced the downstream transformation of benzylsuccinic acid by some currently unknown mechanism. Desulfovibrio was identified as a key microbe due to its enrichment in toluene-free treatments, while the classical toluene-activator in TOLDC

(Desulfosporosinus) only was present in very small abundance (approximately 1%) in toluene-free treatments.

Future studies with these cultures could include determining the mechanistic role of

Desulfovibrio in TOLDC in toluene degradation, p-xylene transformation, and co-amendment 134

studies. LC or GC-MS/MS could be employed to detect the CoA-associated forms of 4- methylbenzoyl-CoA, to better determine how far metabolically p-xylene was transformed.

Identification of the pathway or enzymes generating 4-methylbenzoyl-CoA, the long-term fate of this metabolite, as well as enzymatic assays of isolated benzylsuccinate synthase and benzoyl-CoA reductase (BCR) should be carried out to determine if BSS and BCR from Desulfosporosinus can act on p-xylene and 4-methylbenzoyl-CoA, respectively. The role of Desulfosporosinus and

Desulfovibrio in p-xylene transformation should be more thoroughly established through stable isotope probing experiments using isotopically heavy p-xylene. Also, the bssA amplicon examined through qPCR analysis should be sequenced with next-generation sequencing technology

(Illumina MiSeq) to confirm if in fact additional sequences encoding benzylsuccinate synthase genes are present in this consortium, either from a second Desulfosporosinus lineage (e.g., one that annotated as clone F5OHPNU07H9HB2 in the BLASTn analysis) or from Desulfovibrio; this will elucidate if more than one microorganism is carrying out toluene degradation in TOLDC.

7.2.2 Chapter Five

While the results of this study indicate that PAH loss occurred, we were unable to conclusively link this process to stoichiometric methane production. We showed that the PAHs were biotransformed to some extent, as 2-naphthoic acid and another metabolite were detected in culture supernatants, however no other products were identified. It is possible that dead-end intermediates formed but we cannot be certain with the data acquired in this experiment. The putative novel metabolite identified in section 5.4.2 was likely not a dead-end product, as its relative abundance varied over the course of the experiment (Figure 5-8). Possibly 2-naphthoic acid was a dead-end product (similar to observations of p-toluic acid in Chapter Four), as it did not increase or decrease significantly over the course of the experiment (p-value 0.166). 135

Pelobacter, Desulfovibrio, and Clostridium were identified as likely being involved in PAH bioconversion reactions due to their increasing relative abundance (Figure 5-9) and observations in other hydrocarbon degradation studies. Ultimately, we have contributed evidence to support carboxylation as the mechanism of activation of naphthalene or 2-methylnaphthalene in a methanogenic enrichment culture.

The results of this experiment have led to numerous other research questions deserving further study. Future experiments with this culture should involve additional attempts to elucidate the structure of the putative metabolite from section 5.4.2 through tandem GC-MS/MS, or LC-MS to retain CoA-associated thioester intermediates. Using the parent culture and incubating with single PAH substrates instead of the cocktail used in this experiment will remove confounding or co-metabolism factors in metabolite analysis while also providing more robust evidence for calculating stoichiometric mass balances. Stable isotope fractionation could be attempted to determine the microorganisms incorporating PAH-derived carbons, however studies by Morasch et al. (2004a) and Meckenstock et al. (2016) identify hydrocarbons with eleven or more carbons as the maximum detection limit of this technique due to minute differences in mass being diluted as the molecular weight of the compound increases. At 10-14 carbons, the PAHs surveyed in the present study would push the limits of detection, therefore hydrogen stable isotope fractionation

(proposed by Meckenstock et al., 2016) could be employed to overcome the limitations of the carbon-based technique. From the results of this eight-month long experiment, future experiments should be run for at least one year to observe full biodegradation of naphthalene and 2- methylnaphthalene, while phenanthrene degradation may take even longer to observe

(approximately 13.5 months at the current rate of removal). Chang et al. (2002) and Christensen et al. (2004) achieved considerable success cultivating methanogenic PAH-degrading cultures at 136

higher temperature (30-40˚C) and slightly alkaline media (pH 8 achieved better naphthalene removal than pH 7) when using soil as the inoculum, which could be explored further in future attempts at cultivating PAH-degraders from soil or sediments.

7.2.3 Chapter Six

In this study we evaluated the use of Tenax-TA for the recovery of microorganisms from contaminated groundwater. Biomass recoveries were improved in hydrocarbon-amended treatments, with aerobic, nitrate-reducing, and sulfate-reducing treatments demonstrating the highest biomass and correlating with the strength of the associated electron acceptor. Benzene was not consumed in the 80 days of monitoring in any anaerobic treatments, but both toluene and benzene were consumed in aerobic treatments. Iron(III) was found to be poorly transformed in the presence of Tenax-TA due to differentiated sessile and planktonic microorganisms. Microbial community analysis revealed the diversity of the communities was not impacted by the presence of Tenax-TA but rather the electron accepting condition. For use as a prototype sampling device,

Tenax-TA has the potential to reduce the amount of material required to assess changes in microbial communities in hydrocarbon-contaminated sites.

Future needs for this project include validation of microcosm experiments by implementing field studies at contaminated sites with more extensive characterization. Examining the effect of

Tenax-TA on biodegradation using microcosm studies with light fraction hydrocarbons and alkanes and assess their removal over time. Varying experimental parameters such as temperature

(to reflect seasonal changes), not reamending with hydrocarbons as they are depleted, and withdrawing samples for sequencing throughout the experiment instead of only the beginning and end will give information on starvation state and how microbial communities change once substrates are depleted. Additional testing with contaminated soil, oil sands produced water, or 137

tailings will give greater information about the efficacy of implementing Tenax-TA samplers in different environments and with different types of samples.

138

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Appendix A: Supplementary Material from Chapter Four

)

s e l Toluene

o 6 m

n Ethylbenzene

(

i p-Xylene

c 4

c Toluene + Ethylbenzene

i o

z 2 Toluene + p-Xylene

n e

b Unamended 0 0 20 40 60 Time (days)

Figure A-2: Benzoic acid detected in phase 2 incubations, error bars ± SEM, n=3.

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161

)

l o

m 4

n Toluene

(

i p-Xylene

i 2 Toluene + p-Xylene

o z

n Unamended

e b 0 0 10 20 30 40 50 Time (days)

Figure A-4: Benzoic acid production from phase 3 incubations, error bars ± SEM, n=3.

162

Toluene + Toluene + Toluene Ethylbenzene p-Xylene Unamended ethylbenzene p-xylene Domain;Phylum;Class;Order;Family;Genus;Species Day Day Day Day Day Day Day Day Day Day Day Day 100 600 100 600 100 600 100 600 100 600 100 600 Archaea;Euryarchaeota;Methanobacteria;Methanobacteriales; Methanobacteriaceae;Methanobacterium;uncultured 0.00 0.00 0.00 0.00 0.00 0.04 0.00 0.11 0.85 0.00 0.00 0.00 Methanobacteriales archaeon Archaea;Euryarchaeota;Methanomicrobia;Methanomicrobiales; 30.00 0.14 14.23 0.00 34.09 0.06 17.14 0.44 20.06 0.24 8.87 0.43 Methanomicrobiaceae;Methanoculleus;NA Archaea;Euryarchaeota;Methanomicrobia;Methanomicrobiales; 0.00 0.00 8.94 0.00 12.12 0.00 0.00 0.00 0.42 0.00 16.13 0.00 Methanoregulaceae;Methanolinea;uncultured archaeon TA02 Archaea;Euryarchaeota;Methanomicrobia;Methanomicrobiales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 2.97 0.00 0.00 0.00 Methanoregulaceae;Methanoregula;uncultured Methanoregula sp. Archaea;Euryarchaeota;Methanomicrobia;Methanomicrobiales; 0.00 0.08 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 Methanoregulaceae;Methanoregula;NA Archaea;Euryarchaeota;Methanomicrobia;Methanomicrobiales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.07 0.00 0.00 0.00 0.00 NA;NA;NA Archaea;Euryarchaeota;Methanomicrobia;Methanosarcinales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 Methanosaetaceae;Methanosaeta;Methanosaeta harundinacea 6Ac Archaea;Euryarchaeota;Methanomicrobia;Methanosarcinales; 70.00 0.20 76.83 0.17 50.00 0.10 82.86 0.20 37.01 0.10 75.00 0.30 Methanosaetaceae;Methanosaeta;uncultured Methanosarcina sp. Archaea;Euryarchaeota;Methanomicrobia;NA;NA;NA;NA 0.00 0.00 0.00 0.00 1.52 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Archaea;Euryarchaeota;Thermococci;Methanofastidiosales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.85 0.00 0.00 0.00 Methanofastidiosaceae;Candidatus Methanofastidiosum;NA Archaea;Nanoarchaeaeota;NA;NA;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.15 0.00 0.00 0.00 0.00 Bacteria;Acidobacteria;Acidobacteriia;Subgroup 2;uncultured Acidobacteriales bacterium;uncultured Acidobacteriales bacterium;uncultured Acidobacteriales 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.08 bacterium Bacteria;Acidobacteria;Aminicenantia;Aminicenantales;uncultured 0.00 2.03 0.00 1.89 0.00 1.13 0.00 1.56 3.53 0.73 0.00 1.23 microorganism;uncultured microorganism;uncultured microorganism Bacteria;Actinobacteria;Actinobacteria;Corynebacteriales;Dietziaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.09 Dietzia;NA Bacteria;Actinobacteria;Actinobacteria;Corynebacteriales;Nocardiaceae; 0.00 0.21 0.00 0.68 0.00 0.00 0.00 0.45 0.00 0.00 0.00 0.57 Rhodococcus;NA Bacteria;Actinobacteria;Actinobacteria;Micrococcales;Microbacteriaceae; 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Leucobacter;NA Bacteria;Actinobacteria;Actinobacteria;Micrococcales;Microbacteriaceae; 0.00 0.00 0.00 0.08 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 Microbacterium;NA Bacteria;Actinobacteria;Actinobacteria;Micrococcales;Micrococcaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 Micrococcus;NA 163

Bacteria;Actinobacteria;Actinobacteria;Micrococcales;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.00 Bacteria;Actinobacteria;Actinobacteria;Propionibacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.05 0.00 0.02 0.00 0.00 Propionibacteriaceae;Cutibacterium;NA Bacteria;Actinobacteria;Coriobacteriia;OPB41;NA;NA;NA 0.00 0.13 0.00 0.75 0.00 0.21 0.00 0.61 0.42 0.18 0.00 0.69 Bacteria;Actinobacteria;Thermoleophilia;Gaiellales;uncultured;uncultured 0.00 0.00 0.00 0.09 0.00 0.04 0.00 0.07 0.00 0.00 0.00 0.07 Solirubrobacter sp.;uncultured Solirubrobacter sp. Bacteria;Actinobacteria;Thermoleophilia;Gaiellales;uncultured;uncultured 0.00 0.54 0.00 0.79 0.00 0.53 0.00 1.06 0.00 0.39 0.00 1.38 organism;uncultured organism Bacteria;Actinobacteria;Thermoleophilia;Gaiellales;uncultured;uncultured soil 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.47 bacterium;uncultured soil bacterium Bacteria;Actinobacteria;WCHB1-81;uncultured actinobacterium;uncultured 0.00 0.04 0.00 0.08 0.00 0.00 0.00 0.10 0.00 0.00 0.00 0.11 actinobacterium;uncultured actinobacterium;uncultured actinobacterium Bacteria;Actinobacteria;WCHB1-81;uncultured bacterium;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.01 0.00 0.00 0.00 0.00 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Aegiribacteria;uncultured bacterium;uncultured bacterium;uncultured 0.00 0.65 0.00 0.17 0.00 0.96 0.00 0.42 0.00 0.45 0.00 0.17 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Armatimonadetes;uncultured;uncultured bacterium SJA- 176;uncultured bacterium SJA-176;uncultured bacterium SJA-176;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.24 0.00 0.00 0.00 0.08 bacterium SJA-176 Bacteria;Armatimonadetes;uncultured;uncultured bacterium;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Armatimonadetes;uncultured;NA;NA;NA;NA 0.00 0.00 0.00 0.17 0.00 0.01 0.00 0.04 0.00 0.00 0.00 0.06 Bacteria;Atribacteria;JS1;Atribacteria bacterium JGI 0000079-L04;Atribacteria bacterium JGI 0000079-L04;Atribacteria bacterium JGI 0000079- 0.00 0.00 0.00 0.07 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 L04;Atribacteria bacterium JGI 0000079-L04 Bacteria;Atribacteria;JS1;uncultured candidate division JS1 bacterium;uncultured candidate division JS1 bacterium;uncultured candidate 0.00 0.04 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 division JS1 bacterium;uncultured candidate division JS1 bacterium Bacteria;BRC1;Omnitrophica bacterium OLB16;Omnitrophica bacterium OLB16;Omnitrophica bacterium OLB16;Omnitrophica bacterium 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 OLB16;Omnitrophica bacterium OLB16 Bacteria;BRC1;uncultured soil bacterium PBS-III-29;uncultured soil bacterium PBS-III-29;uncultured soil bacterium PBS-III-29;uncultured soil bacterium 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.00 0.00 0.00 PBS-III-29;uncultured soil bacterium PBS-III-29 Bacteria;Bacteroidetes;Bacteroidia;Bacteroidales;Bacteroidetes vadinHA17;uncultured Cytophagales bacterium;uncultured Cytophagales 0.00 0.20 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 bacterium Bacteria;Bacteroidetes;Bacteroidia;Bacteroidales;Bacteroidetes 0.00 2.15 0.00 2.30 0.00 1.80 0.00 2.22 3.95 1.26 0.00 2.26 vadinHA17;NA;NA Bacteria;Bacteroidetes;Bacteroidia;Bacteroidales;Dysgonomonadaceae; 0.00 0.37 0.00 0.20 0.00 0.34 0.00 0.23 0.00 0.22 0.00 0.21 Proteiniphilum;NA Bacteria;Bacteroidetes;Bacteroidia;Bacteroidales;Paludibacteraceae;NA;NA 0.00 0.91 0.00 0.50 0.00 0.94 0.00 0.14 0.00 0.58 0.00 0.51 Bacteria;Bacteroidetes;Bacteroidia;Bacteroidales;Prolixibacteraceae;BSV13;NA 0.00 0.00 0.00 0.11 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Bacteria;Bacteroidetes;Bacteroidia;Bacteroidales;Prolixibacteraceae;uncultured; 0.00 0.00 0.00 0.17 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Bacteroidales bacterium 6E

164

Bacteria;Bacteroidetes;Bacteroidia;Bacteroidales;Rikenellaceae;Blvii28 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.08 wastewater-sludge group;NA Bacteria;Bacteroidetes;Bacteroidia;Chitinophagales;Chitinophagaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 Flaviaesturariibacter;uncultured bacterium Bacteria;Bacteroidetes;Bacteroidia;Chitinophagales;Chitinophagaceae; 0.00 0.00 0.00 0.06 0.00 0.01 0.00 0.22 0.00 0.00 0.00 0.00 Sediminibacterium;NA Bacteria;Bacteroidetes;Bacteroidia;Cytophagales;Cyclobacteriaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 Algoriphagus;NA Bacteria;Bacteroidetes;Bacteroidia;Flavobacteriales;Flavobacteriaceae; 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Psychroserpens;NA Bacteria;Bacteroidetes;Bacteroidia;Flavobacteriales;Weeksellaceae;NA;NA 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Bacteria;Bacteroidetes;Bacteroidia;Sphingobacteriales;Lentimicrobiaceae;waste 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 water metagenome;wastewater metagenome Bacteria;Bacteroidetes;Bacteroidia;Sphingobacteriales;Lentimicrobiaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 NA;NA Bacteria;Bacteroidetes;Ignavibacteria;OPB56;uncultured bacterium;uncultured 0.00 0.07 0.00 0.05 0.00 0.10 0.00 0.06 0.00 0.11 0.00 0.04 bacterium;uncultured bacterium Bacteria;Bacteroidetes;Ignavibacteria;SJA-28;uncultured bacterium;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.08 bacterium;uncultured bacterium Bacteria;Chloroflexi;Anaerolineae;ADurb.Bin180;uncultured 0.00 0.07 0.00 0.08 0.00 0.05 0.00 0.08 0.00 0.00 0.00 0.15 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.49 0.00 0.32 0.00 0.17 0.00 0.25 0.00 0.20 0.00 0.37 ADurb.Bin120;uncultured Chloroflexi bacterium Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.14 0.00 0.00 ADurb.Bin120;NA Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.42 0.00 0.00 0.00 0.30 0.00 0.00 0.00 0.28 0.00 0.00 Anaerolinea;uncultured bacterium Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.56 0.00 0.19 0.00 0.48 0.00 0.18 0.00 0.31 0.00 0.13 Leptolinea;uncultured bacterium SHD-254 Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.13 0.00 0.06 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 Leptolinea;NA Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.11 0.00 0.22 0.00 0.11 0.00 0.22 0.00 0.11 0.00 0.20 Ornatilinea;uncultured Chloroflexi bacterium Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.43 0.00 0.36 0.00 0.51 0.00 0.47 0.00 0.26 0.00 0.43 Pelolinea;uncultured bacterium Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.53 0.00 0.70 0.00 0.33 0.00 0.41 0.99 0.51 0.00 0.69 uncultured;uncultured Longilinea sp. Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.57 0.00 0.00 0.00 0.26 0.00 0.47 0.00 0.39 0.00 0.43 uncultured;uncultured anaerobic bacterium Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 uncultured;uncultured delta proteobacterium Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.79 0.00 0.52 0.00 0.57 0.00 0.56 0.00 0.30 0.00 0.67 uncultured;uncultured microorganism Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 uncultured;NA Bacteria;Chloroflexi;Anaerolineae;Anaerolineales;Anaerolineaceae; 0.00 0.79 0.00 0.41 0.00 0.80 0.00 0.21 0.56 0.55 0.00 0.73 165

NA;NA Bacteria;Chloroflexi;Anaerolineae;MSB-5E12;uncultured Chloroflexi bacterium;uncultured Chloroflexi bacterium;uncultured Chloroflexi 0.00 0.00 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 0.00 0.00 bacterium Bacteria;Chloroflexi;Anaerolineae;RBG-13-54-9;NA;NA;NA 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Bacteria;Chloroflexi;Anaerolineae;uncultured;metagenome;metagenome; 0.00 0.00 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.00 metagenome Bacteria;Chloroflexi;Anaerolineae;uncultured;uncultured 0.00 0.00 0.00 0.21 0.00 0.04 0.00 0.24 0.00 0.00 0.00 0.19 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Chloroflexi;Anaerolineae;NA;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.01 0.00 0.00 Bacteria;Chloroflexi;Dehalococcoidia;Dehalococcoidales;uncultured;uncultured 0.00 0.00 0.00 0.06 0.00 0.04 0.00 0.07 0.00 0.02 0.00 0.05 bacterium;uncultured bacterium Bacteria;Chloroflexi;Dehalococcoidia;GIF9;AB-539-J10;NA;NA 0.00 0.62 0.00 0.31 0.00 0.39 0.00 0.44 0.00 0.50 0.00 0.74 Bacteria;Chloroflexi;Dehalococcoidia;NA;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.03 0.00 0.00 Bacteria;Cloacimonetes;Cloacimonadia;Cloacimonadales;Cloacimonadaceae; 0.00 0.79 0.00 0.00 0.00 1.31 0.00 0.00 0.00 0.61 0.00 0.17 uncultured bacterium;uncultured bacterium Bacteria;Cloacimonetes;Cloacimonadia;Cloacimonadales;PBS-18;uncultured 0.00 0.11 0.00 0.22 0.00 0.15 0.00 0.22 0.00 0.22 0.00 0.00 bacterium;uncultured bacterium Bacteria;Cyanobacteria;Oxyphotobacteria;Chloroplast;Chlorella sp. 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 ArM0029B;Chlorella sp. ArM0029B;Chlorella sp. ArM0029B Bacteria;Cyanobacteria;Oxyphotobacteria;Chloroplast;NA;NA;NA 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Bacteria;Cyanobacteria;Sericytochromatia;metagenome;metagenome; 0.00 0.00 0.00 0.02 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 metagenome;metagenome Bacteria;Cyanobacteria;Sericytochromatia;NA;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.03 Bacteria;Epsilonbacteraeota;Campylobacteria;Campylobacterales; 0.00 0.01 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Arcobacteraceae;Arcobacter;Arcobacter cibarius Bacteria;Epsilonbacteraeota;Campylobacteria;Campylobacterales; 0.00 0.03 0.00 0.08 0.00 0.03 0.00 0.00 0.00 0.07 0.00 0.00 Thiovulaceae;Sulfuricurvum;NA Bacteria;Firmicutes;Bacilli;Bacillales;Bacillaceae;Anaerobacillus; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.09 0.00 0.00 0.00 0.00 Anaerobacillus sp. NB2006 Bacteria;Firmicutes;Bacilli;Bacillales;Bacillaceae;Anoxybacillus;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.00 0.00 0.00 Bacteria;Firmicutes;Bacilli;Bacillales;Bacillaceae;Bacillus;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 0.00 0.16 Bacteria;Firmicutes;Bacilli;Bacillales;Bacillaceae;NA;NA 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.08 0.00 0.00 0.00 0.00 Bacteria;Firmicutes;Bacilli;Lactobacillales;Carnobacteriaceae;Trichococcus;NA 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Bacteria;Firmicutes;Bacilli;Lactobacillales;Enterococcaceae;Enterococcus;NA 0.00 0.00 0.00 0.10 0.00 0.00 0.00 0.08 0.00 0.00 0.00 0.00 Bacteria;Firmicutes;Clostridia;Clostridiales;Christensenellaceae;Christensenella 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 ceae R-7 group;NA Bacteria;Firmicutes;Clostridia;Clostridiales;Clostridiaceae 1;Clostridium sensu 0.00 0.03 0.00 0.14 0.00 0.06 0.00 0.06 0.00 0.03 0.00 0.15 stricto 13;NA Bacteria;Firmicutes;Clostridia;Clostridiales;Clostridiaceae 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 1;Youngiibacter;uncultured bacterium Bacteria;Firmicutes;Clostridia;Clostridiales;Clostridiaceae 1;Youngiibacter;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.56 0.00 0.00 0.00 Bacteria;Firmicutes;Clostridia;Clostridiales;Eubacteriaceae;Acetobacterium; 0.00 0.00 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.00 uncultured Acetobacterium sp. Bacteria;Firmicutes;Clostridia;Clostridiales;Eubacteriaceae;Acetobacterium;NA 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 166

Bacteria;Firmicutes;Clostridia;Clostridiales;Eubacteriaceae;Anaerofustis; 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 uncultured bacterium Bacteria;Firmicutes;Clostridia;Clostridiales;Family XI;Sedimentibacter;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.42 0.00 0.00 0.00 Bacteria;Firmicutes;Clostridia;Clostridiales;Family XII;Fusibacter;uncultured 0.00 0.02 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 soil bacterium Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae; 0.00 51.12 0.00 1.16 0.00 53.53 0.00 0.82 4.24 59.70 0.00 1.31 Desulfosporosinus;Desulfosporosinus sp. Tol-M Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae; 0.00 0.11 0.00 0.09 0.00 0.02 0.00 0.00 0.00 0.04 0.00 0.00 Desulfosporosinus;NA Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.00 Desulfotomaculum;NA Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae; 0.00 0.00 0.00 0.23 0.00 0.11 0.00 0.10 0.00 0.05 0.00 0.15 SCADC1-2-3;NA Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae; 0.00 0.50 0.00 2.21 0.00 0.28 0.00 5.22 0.00 0.29 0.00 1.26 uncultured;Clostridium sp. enrichment culture clone 06-1235251-143 Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae; 0.00 0.00 0.00 0.00 0.00 0.10 0.00 0.00 0.00 0.04 0.00 0.00 uncultured;uncultured Clostridiales bacterium Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae;uncultured; 0.00 0.00 0.00 0.00 0.00 0.01 0.00 0.00 0.00 0.00 0.00 0.00 uncultured soil bacterium Bacteria;Firmicutes;Clostridia;Clostridiales;Peptococcaceae;uncultured; 0.00 0.00 0.00 2.88 0.00 0.04 0.00 0.83 0.00 0.21 0.00 0.36 NA Bacteria;Firmicutes;Clostridia;Clostridiales;Peptostreptococcaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.13 0.00 0.00 0.00 0.20 Romboutsia;NA Bacteria;Firmicutes;Clostridia;Clostridiales;Peptostreptococcaceae; 0.00 0.08 0.00 0.20 0.00 0.05 0.00 0.20 0.00 0.03 0.00 0.00 NA;NA Bacteria;Firmicutes;Clostridia;Clostridiales;Ruminococcaceae;Ercella;NA 0.00 0.00 0.00 0.00 0.00 0.04 0.00 0.04 0.00 0.00 0.00 0.00 Bacteria;Firmicutes;Clostridia;Clostridiales;Ruminococcaceae; 0.00 0.05 0.00 0.00 0.00 0.16 0.00 0.00 0.00 0.00 0.00 0.00 Hydrogenoanaerobacterium;uncultured bacterium Bacteria;Firmicutes;Clostridia;Clostridiales;Ruminococcaceae;NA;NA 0.00 0.00 0.00 0.52 0.00 0.10 0.00 0.68 0.00 0.09 0.00 0.56 Bacteria;Firmicutes;Clostridia;Clostridiales;Syntrophomonadaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 Syntrophomonas;NA Bacteria;Firmicutes;Clostridia;Clostridiales;TSAC18;uncultured 0.00 0.19 0.00 1.22 0.00 0.16 0.00 1.21 0.00 0.18 0.00 0.68 bacterium;uncultured bacterium Bacteria;Firmicutes;Clostridia;Clostridiales;NA;NA;NA 0.00 0.04 0.00 0.12 0.00 0.00 0.00 0.09 0.00 0.00 0.00 0.05 Bacteria;Firmicutes;Clostridia;DTU014;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.05 Bacteria;Firmicutes;Clostridia;NA;NA;NA;NA 0.00 0.04 0.00 0.08 0.00 0.04 0.00 0.11 0.00 0.04 0.00 0.05 Bacteria;Firmicutes;Erysipelotrichia;Erysipelotrichales;Erysipelotrichaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.07 0.00 0.00 0.00 0.07 Erysipelothrix;NA Bacteria;Firmicutes;Negativicutes;Selenomonadales;Veillonellaceae;uncultured; 0.00 0.06 0.00 0.16 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 uncultured Peptococcaceae bacterium Bacteria;Gemmatimonadetes;Gemmatimonadetes;Gemmatimonadales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.00 0.00 0.00 Gemmatimonadaceae;uncultured;uncultured bacterium contig00007 Bacteria;Latescibacteria;uncultured bacterium;uncultured bacterium;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.03 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Lentisphaerae;Oligosphaeria;NA;NA;NA;NA 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 167

Bacteria;Omnitrophicaeota;Omnitrophica WOR_2 bacterium RIFCSPLOWO2_12_FULL_51_24;Omnitrophica WOR_2 bacterium RIFCSPLOWO2_12_FULL_51_24;Omnitrophica WOR_2 bacterium 0.00 0.09 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.14 RIFCSPLOWO2_12_FULL_51_24;Omnitrophica WOR_2 bacterium RIFCSPLOWO2_12_FULL_51_24;Omnitrophica WOR_2 bacterium RIFCSPLOWO2_12_FULL_51_24 Bacteria;Omnitrophicaeota;uncultured bacterium;uncultured 0.00 0.00 0.00 0.11 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 bacterium;uncultured bacterium;uncultured bacterium;uncultured bacterium Bacteria;Patescibacteria;Microgenomatia;Candidatus Collierbacteria;uncultured 0.00 0.03 0.00 0.12 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Patescibacteria;Microgenomatia;Candidatus Gottesmanbacteria;uncultured Microgenomates group bacterium;uncultured 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Microgenomates group bacterium;uncultured Microgenomates group bacterium Bacteria;Patescibacteria;Microgenomatia;Candidatus Gottesmanbacteria;uncultured bacterium;uncultured bacterium;uncultured 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 bacterium Bacteria;Patescibacteria;Microgenomatia;Candidatus Roizmanbacteria;Candidatus Roizmanbacteria bacterium RIFCSPLOWO2_01_FULL_44_13;Candidatus Roizmanbacteria bacterium 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 1.13 0.00 0.00 0.00 RIFCSPLOWO2_01_FULL_44_13;Candidatus Roizmanbacteria bacterium RIFCSPLOWO2_01_FULL_44_13 Bacteria;Patescibacteria;Microgenomatia;Candidatus Roizmanbacteria;uncultured bacterium;uncultured bacterium;uncultured 0.00 0.00 0.00 0.27 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.16 bacterium Bacteria;Patescibacteria;Microgenomatia;Candidatus Woesebacteria;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 bacterium;uncultured bacterium;uncultured bacterium Bacteria;Patescibacteria;Parcubacteria;32-520;uncultured bacterium;uncultured 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 bacterium;uncultured bacterium Bacteria;Patescibacteria;WS6 (Dojkabacteria);uncultured candidate division WS6 bacterium;uncultured candidate division WS6 bacterium;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 candidate division WS6 bacterium;uncultured candidate division WS6 bacterium Bacteria;Patescibacteria;WS6 (Dojkabacteria);NA;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.12 0.00 0.00 0.00 0.00 Bacteria;Planctomycetes;Planctomycetacia;Pirellulales;Pirellulaceae;Pir4 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 lineage;NA Bacteria;Proteobacteria;Alphaproteobacteria;Azospirillales;Azospirillaceae; 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Azospirillum;NA Bacteria;Proteobacteria;Alphaproteobacteria;Caulobacterales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 Caulobacteraceae;uncultured;NA Bacteria;Proteobacteria;Alphaproteobacteria;Reyranellales;Reyranellaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 Reyranella;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhizobiales;Rhizobiaceae; 0.00 0.03 0.00 0.19 0.00 0.05 0.00 0.49 0.00 0.02 0.00 0.05 Allorhizobium-Neorhizobium-Pararhizobium-Rhizobium;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhizobiales;Rhizobiaceae; 0.00 0.00 0.00 0.14 0.00 0.00 0.00 0.07 0.00 0.00 0.00 0.05 Aquamicrobium;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhizobiales;Rhizobiaceae; 0.00 0.00 0.00 0.09 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 168

Mesorhizobium;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhizobiales;Rhizobiaceae; 0.00 0.52 0.00 1.23 0.00 0.44 0.00 0.74 0.00 0.46 0.00 0.79 Ochrobactrum;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhizobiales;Xanthobacteraceae; 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.05 0.00 0.02 0.00 0.00 Ancylobacter;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhodobacterales; 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.00 Rhodobacteraceae;Haematobacter;uncultured bacterium Bacteria;Proteobacteria;Alphaproteobacteria;Rhodobacterales; 0.00 0.07 0.00 0.00 0.00 0.12 0.00 0.06 0.00 0.00 0.00 0.17 Rhodobacteraceae;Paracoccus;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhodobacterales; 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.00 0.00 0.00 0.00 0.00 Rhodobacteraceae;Sulfitobacter;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhodobacterales; 0.00 0.04 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Rhodobacteraceae;NA;NA Bacteria;Proteobacteria;Alphaproteobacteria;Rhodospirillales; 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Rhodospirillaceae;BRH-c57;uncultured Marispirillum sp. Bacteria;Proteobacteria;Alphaproteobacteria;Thalassobaculales;uncultured; 0.00 0.00 0.00 0.09 0.00 0.00 0.00 0.03 0.00 0.03 0.00 0.06 uncultured soil bacterium;uncultured soil bacterium Bacteria;Proteobacteria;Deltaproteobacteria;Deltaproteobacteria Incertae 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Sedis;Syntrophorhabdaceae;Syntrophorhabdus;metagenome Bacteria;Proteobacteria;Deltaproteobacteria;Deltaproteobacteria Incertae Sedis;Syntrophorhabdaceae;Syntrophorhabdus;uncultured Syntrophorhabdaceae 0.00 1.68 0.00 4.81 0.00 1.45 0.00 3.91 0.00 0.81 0.00 4.24 bacterium TA12 Bacteria;Proteobacteria;Deltaproteobacteria;Deltaproteobacteria Incertae 0.00 0.54 0.00 0.57 0.00 0.49 0.00 0.41 1.13 0.34 0.00 0.56 Sedis;Syntrophorhabdaceae;Syntrophorhabdus;NA Bacteria;Proteobacteria;Deltaproteobacteria;Desulfarculales;Desulfarculaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.07 Desulfatiglans;uncultured soil bacterium Bacteria;Proteobacteria;Deltaproteobacteria;Desulfarculales;Desulfarculaceae; 0.00 0.03 0.00 0.21 0.00 0.08 0.00 0.19 0.00 0.04 0.00 0.17 Desulfocarbo;uncultured bacterium UASB_TL19 Bacteria;Proteobacteria;Deltaproteobacteria;Desulfobacterales; 0.00 0.01 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Desulfobacteraceae;Desulfobotulus;NA Bacteria;Proteobacteria;Deltaproteobacteria;Desulfobacterales; 0.00 0.44 0.00 1.56 0.00 0.55 0.00 1.50 0.00 0.37 0.00 1.10 Desulfobacteraceae;uncultured;NA Bacteria;Proteobacteria;Deltaproteobacteria;Desulfobacterales; 0.00 0.02 0.00 0.09 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.04 Desulfobacteraceae;NA;NA Bacteria;Proteobacteria;Deltaproteobacteria;Desulfobacterales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.00 Desulfobulbaceae;Desulfoprunum;NA Bacteria;Proteobacteria;Deltaproteobacteria;Desulfovibrionales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.42 0.00 0.00 0.00 Desulfovibrionaceae;Desulfovibrio;Desulfovibrio sp. C1 Bacteria;Proteobacteria;Deltaproteobacteria;Desulfovibrionales; 0.00 21.85 0.00 50.15 2.27 21.85 0.00 54.65 5.37 21.91 0.00 60.39 Desulfovibrionaceae;Desulfovibrio;Desulfovibrio sp. SRL8083 Bacteria;Proteobacteria;Deltaproteobacteria;Desulfovibrionales; 0.00 0.16 0.00 0.55 0.00 0.39 0.00 0.20 1.27 0.29 0.00 0.44 Desulfovibrionaceae;Desulfovibrio;NA Bacteria;Proteobacteria;Deltaproteobacteria;Desulfovibrionales; 0.00 0.00 0.00 0.00 0.00 0.01 0.00 0.00 0.00 0.00 0.00 0.00 NA;NA;NA Bacteria;Proteobacteria;Deltaproteobacteria;Desulfuromonadales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.07 0.00 0.06 0.00 0.00 Desulfuromonadaceae;Desulfuromonas;NA 169

Bacteria;Proteobacteria;Deltaproteobacteria;Desulfuromonadales; 0.00 0.05 0.00 1.02 0.00 0.27 0.00 0.51 0.00 0.00 0.00 0.57 Geobacteraceae;Geobacter;Geobacteraceae bacterium JN18_V95_J Bacteria;Proteobacteria;Deltaproteobacteria;Desulfuromonadales; 0.00 0.26 0.00 0.19 0.00 0.17 0.00 1.09 0.00 0.26 0.00 0.55 Geobacteraceae;Geobacter;NA Bacteria;Proteobacteria;Deltaproteobacteria;MBNT15;NA;NA;NA 0.00 0.00 0.00 0.21 0.00 0.04 0.00 0.21 0.00 0.00 0.00 0.17 Bacteria;Proteobacteria;Deltaproteobacteria;Myxococcales;Phaselicystidaceae; 0.00 0.06 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 Phaselicystis;uncultured organism Bacteria;Proteobacteria;Deltaproteobacteria;PB19;NA;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.07 0.00 0.00 0.00 0.00 Bacteria;Proteobacteria;Deltaproteobacteria;SAR324 clade(Marine group 0.00 0.02 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.03 B);NA;NA;NA Bacteria;Proteobacteria;Deltaproteobacteria;Syntrophobacterales; 0.00 0.00 0.00 0.09 0.00 0.00 0.00 0.38 0.00 0.00 0.00 0.08 Syntrophaceae;Smithella;Smithella sp. SC_K08D17 Bacteria;Proteobacteria;Deltaproteobacteria;Syntrophobacterales; 0.00 0.80 0.00 1.54 0.00 0.69 0.00 1.51 8.62 0.39 0.00 1.43 Syntrophaceae;Smithella;NA Bacteria;Proteobacteria;Deltaproteobacteria;Syntrophobacterales; 0.00 0.00 0.00 0.00 0.00 0.18 0.00 0.00 0.00 0.24 0.00 0.00 Syntrophaceae;Syntrophus;uncultured Syntrophaceae bacterium Bacteria;Proteobacteria;Deltaproteobacteria;Syntrophobacterales; 0.00 0.16 0.00 0.69 0.00 0.06 0.00 0.31 0.00 0.15 0.00 0.38 Syntrophaceae;Syntrophus;NA Bacteria;Proteobacteria;Deltaproteobacteria;Syntrophobacterales; 0.00 0.33 0.00 0.36 0.00 0.25 0.00 0.37 0.00 0.24 0.00 0.73 Syntrophaceae;uncultured;NA Bacteria;Proteobacteria;Deltaproteobacteria;NA;NA;NA;NA 0.00 0.08 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.14 0.00 0.00 Bacteria;Proteobacteria;Gammaproteobacteria;Alteromonadales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.07 0.00 0.00 0.00 0.00 Marinobacteraceae;Marinobacter;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.29 0.00 0.00 0.00 0.27 0.00 0.10 0.00 0.29 Burkholderiaceae;Acidovorax;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.12 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Burkholderiaceae;Aquabacterium;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.10 0.00 0.00 0.00 0.08 0.00 0.00 0.00 0.07 Burkholderiaceae;Burkholderia-Caballeronia-Paraburkholderia;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 1.84 0.00 0.00 0.00 Burkholderiaceae;Hydrogenophaga;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.09 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Burkholderiaceae;Pelomonas;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 1.13 0.00 0.00 0.00 Burkholderiaceae;Polaromonas;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 Burkholderiaceae;Ralstonia;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.38 0.00 1.73 0.00 0.39 0.00 1.21 1.84 0.23 0.00 1.55 Burkholderiaceae;NA;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 Methylophilaceae;Methylotenera;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.05 Methylophilaceae;NA;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.25 0.00 0.08 0.00 0.16 0.00 0.00 0.00 0.28 Neisseriaceae;uncultured;uncultured bacterium

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Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 Neisseriaceae;uncultured;uncultured beta proteobacterium Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.16 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Rhodocyclaceae;Azoarcus;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.04 Rhodocyclaceae;Sulfuritalea;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Rhodocyclaceae;Thauera;NA Bacteria;Proteobacteria;Gammaproteobacteria;Betaproteobacteriales; 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 Rhodocyclaceae;NA;NA Bacteria;Proteobacteria;Gammaproteobacteria;Enterobacteriales; 0.00 0.03 0.00 0.18 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Enterobacteriaceae;Escherichia-Shigella;NA Bacteria;Proteobacteria;Gammaproteobacteria;Enterobacteriales; 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.06 0.00 0.02 0.00 0.00 Enterobacteriaceae;NA;NA Bacteria;Proteobacteria;Gammaproteobacteria;Legionellales;Legionellaceae; 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 0.00 0.00 Legionella;NA Bacteria;Proteobacteria;Gammaproteobacteria;Pseudomonadales; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.07 Moraxellaceae;Acinetobacter;NA Bacteria;Proteobacteria;Gammaproteobacteria;Pseudomonadales; 0.00 0.00 0.00 0.00 0.00 0.07 0.00 0.07 0.00 0.00 0.00 0.32 Pseudomonadaceae;Pseudomonas;NA Bacteria;Proteobacteria;Gammaproteobacteria;Xanthomonadales; 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Rhodanobacteraceae;Dokdonella;NA Bacteria;Proteobacteria;Gammaproteobacteria;Xanthomonadales; 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 Xanthomonadaceae;Arenimonas;uncultured beta proteobacterium Bacteria;Proteobacteria;Gammaproteobacteria;Xanthomonadales; 0.00 0.00 0.00 0.00 0.00 0.02 0.00 0.00 0.00 0.00 0.00 0.00 Xanthomonadaceae;Lysobacter;NA Bacteria;Rokubacteria;NC10;Methylomirabilales;Methylomirabilaceae; 0.00 0.22 0.00 0.46 0.00 0.00 0.00 0.25 0.00 0.07 0.00 0.45 Sh765B-TzT-35;uncultured bacterium Bacteria;Spirochaetes;Spirochaetia;Spirochaetales;Spirochaetaceae; 0.00 1.11 0.00 3.00 0.00 0.80 0.00 1.68 0.00 0.89 0.00 1.47 Sphaerochaeta;NA Bacteria;Spirochaetes;Spirochaetia;Spirochaetales;Spirochaetaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.00 0.00 0.00 Spirochaeta 2;uncultured Spirochaetales bacterium Bacteria;Spirochaetes;Spirochaetia;Spirochaetales;Spirochaetaceae;uncultured; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.09 0.00 0.00 0.00 0.08 uncultured bacterium SJA-102 Bacteria;Spirochaetes;Spirochaetia;Spirochaetales;Spirochaetaceae;uncultured; 0.00 2.87 0.00 3.84 0.00 2.85 0.00 4.64 0.00 2.00 0.00 1.56 NA Bacteria;Synergistetes;Synergistia;Synergistales;Synergistaceae;JGI-0000079- 0.00 0.38 0.00 0.59 0.00 0.33 0.00 0.59 0.00 0.34 0.00 0.33 D21;uncultured Synergistetes bacterium Bacteria;Synergistetes;Synergistia;Synergistales;Synergistaceae; 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.05 0.00 0.06 Thermovirga;uncultured Thermovirga sp. Bacteria;Synergistetes;Synergistia;Synergistales;Synergistaceae;uncultured;NA 0.00 0.07 0.00 0.16 0.00 0.15 0.00 0.05 0.00 0.07 0.00 0.09 Bacteria;Tenericutes;Mollicutes;EUB33-2;uncultured bacterium;uncultured 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.11 bacterium;uncultured bacterium Bacteria;Thermotogae;Thermotogae;Kosmotogales;Kosmotogaceae;Mesotoga; 0.00 0.90 0.00 1.50 0.00 1.09 0.00 0.78 0.42 0.63 0.00 1.05 NA Bacteria;Verrucomicrobia;Verrucomicrobiae;Opitutales;Opitutaceae;NA;NA 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 171

Bacteria;WPS-2;Burkholderiales bacterium Beta_02;Burkholderiales bacterium Beta_02;Burkholderiales bacterium Beta_02;Burkholderiales bacterium 0.00 0.06 0.00 0.06 0.00 0.07 0.00 0.09 0.00 0.06 0.00 0.00 Beta_02;Burkholderiales bacterium Beta_02 Bacteria;WPS-2;bacterium ADurb.Bin236;bacterium ADurb.Bin236;bacterium 0.00 0.06 0.00 0.00 0.00 0.04 0.00 0.00 0.00 0.04 0.00 0.00 ADurb.Bin236;bacterium ADurb.Bin236;bacterium ADurb.Bin236 Bacteria;WS4;uncultured Clostridia bacterium;uncultured Clostridia bacterium;uncultured Clostridia bacterium;uncultured Clostridia 0.00 0.00 0.00 0.08 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 bacterium;uncultured Clostridia bacterium Bacteria;NA;NA;NA;NA;NA;NA 0.00 0.04 0.00 0.74 0.00 0.19 0.00 0.09 0.00 0.09 0.00 0.00 Eukaryota;Centrohelida;Heterophryidae;metagenome;metagenome;metagenome 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.06 0.00 0.00 0.00 0.00 ;metagenome Eukaryota;SAR;Alveolata;Ciliophora;Oligohymenophorea;Peritrichia;NA 0.00 0.00 0.00 0.14 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.05 Eukaryota;SAR;Alveolata;Ciliophora;NA;NA;NA 0.00 0.00 0.00 0.05 0.00 0.00 0.00 0.00 0.00 0.03 0.00 0.00

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Table A-2: Detection of bssA by qPCR from cDNA for expression analysis. Technical replicates were analyzed in triplicate.

Time Toluene Ethylbenzene p-Xylene Toluene + Toluene + Unamended (days) only only only ethylbenzene p-xylene 1 + ++ ++ ++ ++ 4 +++ + ++ ++ ++ 11 ++ +++ + ++ +++ 21 + + ++ ++ 36 + ++ ++ + + ++ 60 ++ +++ + ++ ++ + = detected in 1 replicate, ++ = detected in 2 replicates, +++ = detected in all 3 replicates

Shannon diversity score Hydrocarbon treatment Day 100 Day 600 Toluene 0.6109 1.9789 Ethylbenzene 0.6959 2.5752 Toluene + Ethylbenzene 1.1187 1.8877 p-Xylene 0.4581 2.3574 Toluene + p-Xylene 2.1974 1.6173 Unamended 0.7249 2.1879

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Appendix B: Supplementary Material from Chapter Six

DNA (ng/μL) Activated Diatomaceous Zeolite Mat540 T-carbon carbon earth Positive control 0.45-0.75 0.0 0.0 6.05-6.15 5.5-7.1 (soil + fertilizer) Experimental 0.3 0.0 0.0 3.6-4.0 2.0-2.2 (soil only) Negative control 0.0 0.0 0.0 0.85-0.95 0.9-1.1 (autoclaved soil) Blank 0.0 0.0 0.0 0.0 0.0 (sterile, no soil)

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Table B-2: DNA extraction concentrations from field trials with diatomaceous earth (DE), T- carbon (TC), and Tenax-TA (TA). Trap samplers were deployed into a hydrocarbon-contaminated aquifer and recovered in one-month intervals for a total of three months. Recovered and extracted DNA concentrations are provided for each replicate. Some samples had DNA concentrations higher than the detection limit (> 60 ng/μL) of the instrument and are reported as “too high” or TH. Values for Shannon and Simpson diversity indices were computed in R using vegan.

DNA Average Average Time Trap Sampling concentration Shannon Simpson (months) material depth (m) (ng/μL) diversity diversity 3 1.58-2.92 2.88 0.88 DE 4 3.35-3.72 2.46 0.84 5 3.48-6.76 1.77 0.68 3 3.09-4.16 2.76 0.86 1 TA 4 1.07-1.66 2.48 0.83 5 0.76-0.88 2.69 0.80 3 3.51-4.43 2.37 0.84 TC 4 3.39-3.79 1.66 0.64 5 2.68-3.45 2.34 0.79 3 2.06-8.77 2.84 0.90 DE 4 6.70-8.24 2.70 0.87 5 4.33-6.25 3.25 0.93 3 0.88-2.77 2.92 0.88 2 TA 4 2.07-5.08 2.97 0.92 5 3.88-3.99 3.34 0.94 3 0.42-0.93 2.49 0.83 TC 4 5.87-7.24 2.63 0.87 5 4.19-6.32 2.90 0.91 3 14.0-22.7 2.59 0.81 DE 4 5.00-5.81 2.82 0.88 5 9.42-11.0 3.54 0.95 3 5.50-9.05 3.46 0.93 3 TA 4 2.46-3.97 3.17 0.93 5 9.49-11.5 3.56 0.95 3 37.2-TH 3.21 0.91 TC 4 6.84-7.91 2.83 0.90 5 13.8-19.6 3.05 0.90

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Normalized DNA per g Average Electron Planktonic/ Tenax HCs sample (ng/µL) DNA SEM acceptor Sessile -TA R1 R2 R3 (ng/µL) Inoculum NA P NA 0.335 0.317 0.354 0.335 0.011 + P + 0.076 0.177 0.211 0.155 0.041 + S + 12.300 10.100 8.060 10.153 1.224 - P + 0.062 0.162 0.069 0.098 0.032 O 2 - S + 10.600 11.700 8.000 10.100 0.542 + P - 0.276 0.304 0.208 0.263 0.029 - P - 0.138 0.132 0.059 0.110 0.025 + P + 0.179 0.268 0.330 0.259 0.044 + S + 1.060 1.860 3.650 2.190 0.766 - P + 0.090 0.088 0.153 0.110 0.021 NO - 3 - S + NA 0.057 NA 0.057 0.019 + P - 0.458 0.411 0.481 0.450 0.021 - P - 0.167 0.154 0.147 0.156 0.006 + P + 0.077 0.118 0.079 0.091 0.013 + S + NA NA NA NA NA - P + 0.079 0.119 0.115 0.104 0.013 Fe3+ - S + NA NA NA NA NA + P - 0.038 0.132 0.097 0.089 0.027 - P - 0.113 0.212 0.122 0.149 0.032 + P + 0.091 0.041 0.041 0.058 0.017 + S + NA NA 0.492 0.492 0.164 - P + 0.079 0.068 0.071 0.072 0.003 SO 2- 4 - S + NA NA NA NA NA + P - 0.102 0.098 0.093 0.098 0.003 - P - 0.101 0.106 0.086 0.098 0.006 + P + 0.087 0.072 0.066 0.075 0.006 + S + NA NA NA NA NA No EA - P + 0.116 0.091 0.089 0.099 0.009 added - S + 0.132 NA NA 0.132 NA + P - NA 0.111 0.126 0.118 0.008 - P - 0.200 0.183 0.184 0.189 0.006

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Figure B-2: Design of experimental microcosms. Glass serum bottles were sealed with the Tenax- TA filled pouch suspended into the aqueous phase. Groundwater was added; associated sand settled to the bottom over time. Aerobic treatments received air as the headspace while anoxic treatments were flushed with a headspace of N2 gas.

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Figure B-3: Toluene and benzene degradation profiles in aerobic (panels A and B), nitrate- reducing (C and D), iron(III)-reducing (E and F), sulfate-reducing (G and H), and no electron acceptor-added (I and J) microcosms, respectively, over 80 days of incubation in the presence and absence of Tenax-TA. Both live incubations and heat-killed controls (HK) were established. Error bars represent the standard error of the mean of 3 replicates. Breaks in plot lines represent re- amendment of hydrocarbons after a time point where all previously added hydrocarbons had been consumed.

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Figure B-4: DNA recovered from hydrocarbon-amended (+HCs) and unamended (-HCs) Tenax- TA pouches across different treatments. Tenax-TA samplers from aerobic microcosms with hydrocarbons yielded the most DNA (10.15 ± 1.22 ng/µL) while DNA was below detection limits in iron(III)-reducing microcosms regardless of the presence of hydrocarbons. Error bars represent the standard error of the mean of 3 replicates. Asterisks represent statistically significant differences as calculated by a t-test (** p-value ≤ 0.01).

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Hydrocarbon-amended samples with Tenax-TA (TA) have already been discussed in detail in Figure 4. In the planktonic hydrocarbon-unamended treatments (no HCs), Candidatus Nitrocosmicus was found to be the most abundant.

Figure B-6: Microbial community composition of nitrate-reducing microcosms including the groundwater used as the inoculum as well as all treatments and controls. Samples were analyzed through 16S rRNA gene sequencing in triplicate and the averages of that analysis are shown here. Hydrocarbon-amended samples with Tenax-TA were dominated by Azoarcus and have already been discussed in detail in Figure 4. Hydrocarbon-unamended planktonic communities were dominated by Rhodoferax and Candidatus Roizmanbacteria, while sessile communities were incredibly diverse, with 66% the total relative abundance comprising a variety of taxa each making up less than 0.2% of the overall community. 181

Appendix C: Medium Recipes

Pfennig anoxic minimal freshwater medium (McInerney et al., 1979):

Per 100 mL: MilliQ water – 100 mL Pfennig I solution – 5 mL Pfennig II solution – 5 mL Balch vitamins – 1 mL Wolin trace metals – 1 mL Resazurin (0.1% stock solution) – 0.1 mL NaHCO3 – 0.35 g Na2S (2.5% solution) – 2.5 mL

Pfennig I solution: MilliQ water – 1000 mL K2HPO4 – 10 g

Pfennig II solution: MilliQ water – 1000 mL . MgCl 6H2O – 6.6 g NaCl – 8 g NH4Cl – 8 g . CaCl2 2H2O – 1 g

Balch vitamins: MilliQ water – 1000 mL biotin – 2 mg folic acid – 2 mg pyridoxine-HCl – 10 mg thiamine-HCl – 5 mg riboflavin – 5 mg nicotinic acid – 5 mg DL calcium pantothenate – 5 mg vitamin B12 – 0.1 mg PABA – 5 mg lipoic acid – 5 mg mercaptoethane-sulfonic acid (MESA) – 5 mg

Wolin’s trace metals: MilliQ water – 1000 mL EDTA – 0.5 g . MgSO4 6H2O – 3 g . MnSO4 H2O – 0.5 g 182

NaCl – 1 g . CaCl2 2H2O – 0.1 g . ZnSO4 7H2O – 0.1 g . FeSO4 7H2O – 0.1 g . CuSO4 7H2O – 0.01 g . Na2MoO4 2H2O – 0.01 g H3BO3 – 0.01 g Na2SeO4 – 0.005 g . NiCl2 6H2O – 0.003 g

Pelotomaculum medium (DSMZ 960) for S. aciditrophicus (DSMZ 26646):

MilliQ water – 1000 mL KH2PO4 – 0.14 g . MgCl 6H2O – 0.2 g . CaCl2 2H2O – 0.15 g NH4Cl – 0.54 g Trace element solution (medium 318) – 1 mL Selenite-tungstate solution (medium 386) – 1 mL Resazurin (0.1% stock solution) – 0.5 mL NaHCO3 – 2.5 g Vitamin solution (medium 141) – 10 mL Sodium crotonate – 0.86 g Yeast extract – 0.1 g . L-Cysteine-HCl H2O – 0.5 g . Na2S 9H2O – 0.3 g

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Appendix D: R Scripts for Diversity Analyses

NMDS plots: # Load in required packages and set working directory library(picante) library(vegan) library(ggplot2) setwd("/Users/Nicole/Documents/R/BoxM")

# Import sequencing data files and metadata files data <- read.csv("/Users/Nicole/Documents/R/BoxM/BoxM_Sequencing.csv", header = TRUE, row.names = 1) metadata <- read.csv("/Users/Nicole/Documents/R/BoxM/metadata.csv", header = TRUE, row.n ames = 1)

# Convert raw reads into relative abundance comm <- decostand(data, method = "total") # Combine sequencing and metadata files combined <- match.phylo.comm(comm, metadata) all.equal(rownames(comm), colnames(metadata)) metadata <- metadata[rownames(comm), ]

# Calculate Brad-Curtis distribution of dataset otus_dist = as.matrix((vegdist(comm, "bray")))

# Perform NMDS analysis, then build a data frame with NMDS coordinates and metadata NMDS = metaMDS(otus_dist) NMDS1 = NMDS$points[,1] NMDS2 = NMDS$points[,2] NMDS = data.frame(MDS2 = NMDS2, MDS1 = NMDS1, Hydrocarbons = metadata$hydrocar bons, Electron_Acceptor = metadata$trophic)

# Plot NMDS analysis with ellipses ggplot(NMDS, aes(x=NMDS1, y=NMDS2, col=Electron_Acceptor)) + geom_point(aes(shape = Hydrocarbons, size = 5)) + stat_ellipse(linetype = 1, size = 1) + theme_bw() + theme(axis.text.x = element_text(size = 12, colour = "black"), axis.text.y = element_text(size = 12, colour = "black"), text = element_text(family="serif", size=14, colour = "black"))

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Dendrograms: # Using the previously defined import and convert commands for this dataset: # Calculate Bray-Curtis distance among samples data_dist <- vegdist(comm, method = "bray")

# Cluster communities using average-linkage algorithm data_clust <- hclust(data_dist, method = "average")

# Plot cluster diagram plot(data_clust, ylab = "Bray-Curtis dissimilarity")

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Appendix E: Bioelectrochemical remediation of phenanthrene in a microbial fuel cell using an anaerobic consortium enriched from a hydrocarbon-contaminated site

Citation: Sharma, M., Nandy, A., Taylor, N., Venkatesan. S.V., Ozhukil Kollath, V., Karan, K.,

Thangadurai, V., Tsesmetzis, N., and Gieg, L. (2020) Bioelectrochemical remediation of phenanthrene in a microbial fuel cell using an anaerobic consortium enriched from a hydrocarbon- contaminated site. J Hazard Mater 389: 121845.

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Appendix F: Comparative evaluation of coated and non-coated carbon electrodes in microbial fuel cells for treatment of municipal sludge

Citation: Nandy, A., Sharma, M., Venkatesan, S.V., Taylor, N., Gieg, L., and Thangadurai, V.

(2019) Comparative evaluation of coated and non-coated carbon electrodes in microbial fuel cells for treatment of municipal sludge. Energies 12: 1-15.

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