Geomicrobiology of Natural Gas Seepage Pockmarks in Öxarfjörður, NE Iceland

Geomicrobiology of natural gas seepage pockmarks in Öxarfjörður, NE Iceland

Guðný Vala Þorsteinsdóttir

Master thesis for 90 credit Magister Scientiarum in Natural Resource Sciences

Supervisor Oddur Vilhelmsson

Co-Supervisors Anett Blishcke Kristinn Pétur Magnússon Margrét Auður Sigurbjörnsdóttir

Faculty of Natural Resource Sciences School of Business and Science University of Akureyri Akureyri, March 2018

Geomicrobiology of natural gas seepage pockmarks in Öxarfjörður, NE- Iceland

90 credit thesis for Magister Scientiarum in Natural Resource Sciences

Faculty of Natural Resource Sciences School of Business and Science University of Akureyri Sólborg, Norðurslóð 2 600 Akureyri

Tel: 460 8000

Registration information: Guðný Vala Þorsteinsdóttir, 2018, Geomicrobiology of natural gas seepage pockmarks in Öxarfjörður, NE-Iceland, Master thesis, Faculty of Natural Resource Sciences, School of Business and Science, University of Akureyri, 103 p.

Printed by Stell Akureyri, March 2018

Abstract

Natural gas seepage is the emission of gaseous hydrocarbon from the subsurface of the Earth, commonly found in sea sediments and along tectonic plate boundaries. Natural gas seepage pockmarks are found offshore and onshore in the Öxarfjörður graben. Previous measurements of hydrocarbon content and carbon isotope ratio of methane, suggested that the gas originates from coal bearing strata in the graben. No previous studies have been made on natural gas seepage microbiota in Iceland, but natural gas seeps are likely to harbour hydrocarbon-utilising microbes that can be extremely valuable for bioremediation of polluted environments. This thesis describes a study that aims towards characterizing the microbiota of two distinct natural gas seepage pockmark sites in Öxarfjörður. Geochemical analysis of the sites included chemical analysis of the water submerging the pockmarks, hydrocarbon content and carbon isotope composition of methane. Analysis of the bacterial community structure was performed with amplicon sequencing of variable regions V3-V4 in the 16S rRNA gene from environmental samples of the gas seepage pockmarks. Bacterial strains from the pockmarks were isolated on selective and differential media to screen for bacterial strains with bioremediative potential. The carbon isotope composition and hydrocarbon content at the two study sites revealed biogenic generation of methane with more complex hydrocarbon composition and thermogenic origin with less organic matter, respectively. Both sites had Proteobacteria as the most abundant bacterial phyla, where the Deltaproteobacteria were more abundant at the geothermal site, and the Alphaproteobacteria at the biogenic site. The Dehalococcoidia class of the Chloroflexi phylum was abundant at the geothermal site while the Anaerolineae class was more abundant at the biogenic site. Bacteroidetes were more abundant at the biogenic site. A total of 106 strains were isolated to purity and characterised, including representatives from the abundant phyla Proteobacteria, Bacterioidetes, Firmicutes and Actinobacteria, many of which have potential as hydrocarbon bioremediators in cold aquatic and sedimentary environments.

Keywords: Bacterial community analysis, coal bed-associated microbiota, bioprospecting, environmental microbiology, microbial ecology.

Útdráttur

Útstreymi vetniskolefna úr jarðlögum er jafnan kallað jarðgas. Jarðgas telst til jarðefnaeldsneytis og er gjarnan að finna í setlögum sjávar og við flekaskil jarðar. Gasútstreymisaugu er að finna bæði ofan- og neðansjávar á Austursandi í Öxarfirði, en greiningar kolefnissamsæta hafa gefið til kynna að jarðgasið eigi rætur sínar að rekja til surtarbrandslaga í setlögum sandsins. Engar rannsóknir hafa áður verið gerðar á örverulífríki slíkra gasútstreymisaugna, en áætla má að þar sé að finna bakteríur sem nýta megi í lífhreinsun á menguðu umhverfi. Þetta verkefni miðar að því að lýsa örverulífríki gasútstreymisaugna í Öxarfirði með því að skoða tvö mismunandi svæði þar sem gasuppstreymi finnst. Mælt var innihald vetniskolefna og framkvæmd ísótópagreining metans til að meta uppruna gassins, ásamt efnagreiningu vatns. Greining á örverusamfélagi gasútstreymisaugnanna var framkvæmd með háafkasta raðgreiningu á breytilega svæðinu V3-V4 innan 16S rRNA gensins úr umhverfissýnum gasaugnanna. Bakteríustofnar voru einangraðir á valæti og skimað fyrir niðurbroti og örðum eiginleikum sem nýta mætti í lífhreinsun. Jarðefnamælingar leiddu í ljós að á öðrum staðnum mátti finna jarðgas en á hinum staðnum var gasið af lífrænum uppruna. Örverusamsetning á jarðgassvæðinu sýndi að meirihluti bakteríubíótunnar tilheyrir Proteobacteria fylkingunni þar sem flokkur Deltaproteobacteria var ráðandi. Fylking Chloroflexi ásamt Aminicenantes og Firmicutes voru einnig ráðandi þar sem flokkur Dehalococcoidia innan Chloroflexi fylkingarinnar var mest áberandi. Þar sem gasið var af lífrænum uppruna var fylking Proteobacteria einnig í mestum mæli þar sem flokkur Alphaproteobacteria var ráðandi. Þar var einnig fylking Bacteroidetes stór hluti bíótunnar, ásamt Chloroflexi þar sem Anaerolineae flokkurinn var ráðandi. Alls voru 106 stofnar einangraðir og kennigreindir, en í stofnasafninu mátti finna fulltrúa flestra algengustu fylkinganna og þar með talið stofna sem mögulega væri hægt að nýta til lífhreinsunar.

Lykilorð: Örveruþýðisgreining, örverur tengdar surtarbrandi, örveruauðlindaleit, umhverfisörverufræði, örveruvistfræði.

A biologist and a geochemist went out on a date. They didn’t know what to talk about, so instead they ended up figuring out the origin of life.

Table of Contents

Figures ...... xi

Tables ...... xiii

Acknowledgements ...... xv

1 Introduction ...... 17 1.1 Geomicrobiology ...... 17 1.1.1 Metals and minerals ...... 19 1.2 Methane and microbes ...... 20 1.2.1 Natural gas seepage ...... 24 1.2.2 Cold seeps and gas hydrates ...... 26 1.2.3 Coal bed methane and associated bacteria ...... 26 1.2.4 Identifying bacterial communities in natural gas seepage with amplicon sequencing ...... 29 1.3 Geological settings of Öxarfjörður ...... 31 1.4 Aims of the present study...... 35 References ...... 37

2 Manuscript I - Gas seepage pockmark microbiomes suggest the presence of sedimentary coal seams in the Öxarfjörður graben of NE-Iceland ...... 43

3 Manuscript II – Bioremediative potential of bacteria in cold desert environments ...... 81

4 Conclusions and future perspectives ...... 99

Appendix A ...... 101

Appendix B ...... 103

ix x Figures

Figure 1. Biogeochemical processes of metals and minerals that are studied in geomicrobiology, from Gadd (2010)...... 18 Figure 2. Schematic representation of the biogeochemistry in marine sediments (Jørgesen & Kasten, 2006). The figure shows the biochemical zones in the relation to pore water chemistry and mineralization processes...... 23 Figure 3. Plots from Xu (2010) to distinguish the origin of methane. (a) The stable isotope ratio plotted against carbon ratios to determine microbial, geothermal or thermogenic origin of methane. (b) Stable isotope ratios of hydrogen and carbon used to determine the microbial, thermogenic and geothermal origin of methane...... 25 Figure 4. Formation of methane gas in coalification processes in relation to organic material, vitrinite reflectance and dryness, figure from Tim A Moore (2012)...... 27 Figure 5. The northeast Iceland plate bondaries and rift systems, and location of the study area within the Northern Volcanic Zone (modified from Einarsson et al., 2008; Jakobsson et al., 2008; Magnúsdóttir et al., 2015; Hjartarson & Sæmundsson, 2014). Abbreviations are: TJ – Tjörnes peninsular sediment outcrops; structural lineament segments DL – Dalvíkur lineament and the HFF – Húsavík- Flatey-Fault system; volcanic rift complexes A – Askja, H - Hrúthálsar , K – Krafla, KF – Kverkfjöll, N – Fremri- Námar, and Th – Theistareykir...... 32 Figure 6. Sample site locations within the Öxarfjörður graben system, existing boreholes, inferred extensional fault locations, and surface depositional environments of the area (Ólafsson et al. 1993 and Sæmundsson et al. 2012)...... 34 Figure 7. A map of the study area showing the AEX sampling sites (blue squares) and the SX sites (orange circles). Black diamonds indicate geothermal boreholes. Faults are inferred from the works of Sæmundsson et al. (31) and Ólafsson et al. (18).

xi The insert shows the location of the study area in Iceland and the volcanic rift zone, bounded by the solid lines...... 46 Figure 8. Hydrocarbon content in the natural gas seepage pockmarks. Extractable organic matter (EOM) concentration as determined by gas chromatography is compared between the AEX (dark columns) and SX (light columns) study sites. Error bars are omitted for clarity...... 53 Figure 9. Bacterial community structure in natural gas seepage pockmarks at Skógakíll (AEX) and Skógalón (SX) sites, presented as the relative abundance of bacterial phyla from amplicon sequencing of V3-V4 in 16S rDNA. Operational taxanomic units (OTUs) with relative abundance lower than 0.1 were omitted...... 55 Figure 10. Poster presentation at the 7th Congress of European Microbiologists (FEMS) in Valencia, Spain, 9-13 July 2017...... 101

xii Tables

Table 1. Location and description of sampling sites. Temperature and pH were measured in situ with a hand-held probe...... 47 Table 2. Major physicochemical characteristics1 of water from the two study sites...... 51 Table 3. Headspace gas analysis1 on sediment samples from seepage pockmarks at the two study sites...... 52 Table 4. Number of predicted operational taxanomic units (OTUs) and alpha diversity metrics as calculated at 25,000 sequences from the two study sites...... 54 Table 5. Colony-forming units per gram sample after 7 days at 22°C on selective media...... 56 Table 6. Bacterial isolates and their taxonomic classification by partial 16S rRNA gene sequencing...... 57 Table 7. Fractional abundance of Chloroflexi classes and orders in the seepage pockmarks microbiomes as determined with amplicon sequencing...... 61 Table 8. Fractional abundance of Proteobacteria classes and orders in the seepage pockmarks microbiomes as determined with amplicon sequencing...... 63 Table 9. Description of strains, cultured and isolated on differential and selective media, and conserved in 30% glycerol at -70 °C...... 71 Table 10. Taxanomic assignment of strains identified by 16S rDNA sequencing and their GenBanka ccession numbers...... 75

xiii

xiv Acknowledgements

I would like to start by thanking my supervisor, professor Oddur Vilhelmsson, for being an outstanding mentor. Thank you for believing in me and giving me the opportunity to grow as a researcher. I want to thank my thesis committee for their contribution and help during the project; Anett Blischke geologist, professor Kristinn Pétur Magnússon and dr. M. Auður Sigurbjörnsdóttir.

Thanks to Orkusjóður from Orkustofnun and Orkurannsóknarsjóður Landsvirkjunar for financially supporting the research.

Many thanks to Þórarinn Sveinn Arnarson at Orkustofnun for his interest, input and guidance. Thanks to ÍSOR for sampling and analysis, especially Finnbogi Óskarson geochemist at ÍSOR for the help during sampling and interpreting chemical data. Thanks to everyone who helped with sampling and labwork, especially M. Auður Sigurbjörnsdóttir and Helga Helgadóttir. I also want to thank my co-workers at Náttúrufræðistofnun Íslands for the support and understanding.

Last but not least, I want to thank my family for all the support and patience during these years. Helgi and our boys Hlynur and Hilmir, my parents, my sister Linda and my in-laws: I couldn’t have done this without you!

xvi 1 Introduction

Extremophiles are organisms that can live in environments that push the limits of biological and physicochemical tolerance in form of extreme temperature, pressure, pH, water availability or any other factor harmful to living organisms. These environments can range from hydrothermal vents to polar ice, the deep subsurface or even nuclear reactors. Bacteria and archaea make up the most diverse group of extremophiles and their adaptations to intense conditions are quite remarkable. Not only is it the understanding of limits to life that interest researchers of extreme environments, but by investigating habitats of extreme organisms one can presumably find novel organisms, new pathways or other unique biochemical features. Or on the other hand, find out the origin of various biochemical features. These extreme organisms can then possibly be used for biotechnological processes. This procedure of investigating specific environments with the hope of finding useful traits using systematic methods is often referred to as bioprospecting. Bioprospecting for specific traits calls for the exploration of environments that are likely to be characterised by the physical and chemical conditions that encourage growth of the microbes that have evolved that trait. In geomicrobiology the biogeochemically active roles are studied, not only for scientific purposes such as for the study of biogeochemical cycles, but also for biotechnological purposes, as many geomicrobiological processes have features exploitable as methods for environmental biotechnology.

1.1 Geomicrobiology

For a long time, microbiologists have been seeking answers to questions regarding how microbes can live and grow in different environments and how the environment shapes the microbial community, and conversely, how do microbes shape their environments? These questions can be answered in a relatively new and important field of interest: geomicrobiology. Geomicrobiology touches the fields of microbiology and geology, and is concerned with elucidating the dynamic relationships between microbes and the earth through chemistry. Hence, biogeochemistry. The microbes play a larger role in shaping our environment than we often realise. In fact, soil is a good example. Soil can be divided into three major components: solid

17 particles, gas, and liquid. The solid particles are mostly minerals and organic matter from former living organisms. The microbiota can assist in the breakdown of large rocks into the smaller particles that form the soil, and are largely responsible for the degradation of organic matter. Bacteria, along with fungi, are therefore critical for the formation of soil. Apart from that, bacteria are also a significant part of the soil where they fix nitrogen, mineralise and mobilise both organic and inorganic nutrients that are vital for, for example, plant growth (Van Der Heijden et al., 2008). Among the most important groups of bacteria in the context of biogeochemistry are those bacteria that are capable of iron-oxidation and reduction, sulphur-oxidation and reduction, sulfate- reduction as well as formation or degradation of carbonates, phosphates and silicates (Gadd, 2010) (Figure1.). These elements are necessary for living organisms and the extraction of metal and minerals from inorganic matter is vital to growth and maintenance of cells. Metal and mineral transformation is therefore a highly important process for the availability of nutrients, and concomitantly, in the shaping of nature.

Figure 1. Biogeochemical processes of metals and minerals that are studied in geomicrobiology, from Gadd (2010).

18 1.1.1 Metals and minerals

Together, metals and minerals make up the Earth´s crust. Minerals are inorganic solids that have definite chemical composition, the most common ones being silicates, carbonates, oxides, sulphides and phosphates (Sparks, 2005). Metals are often a part of mineral structure so the transformation of minerals and metals are closely related mechanisms and can be described as the foundation of geomicrobiology (Gadd & Pan, 2016). While most metals are regarded as toxic in high concentrations, some metals are important for living organisms in low concentrations and may be described as bio-elements (Wächtershäuser, 2010). These metals serve physiological purposes and are highly relevant to biochemistry and the function of a living organism. The transition metals, vanadium, molybdenum, tungsten, manganese, iron, cobalt, nickel, copper and zinc are part of metallo-enzymes, where they act as catalysts. Sodium, magnesium, potassium and calcium are however used as counter-ions for the forming of organic structure with the non-metal bio- elements of hydrogen, carbon, nitrogen, oxygen, phosphorus, sulfur and selenium (Wächtershäuser, 2010). Many of these metals are bound in minerals in the Earth´s crust, and just like the metals, minerals are vital for the growth of living organisms. Thus, the microbes need to “have a plan” for acquiring the minerals they need. Since minerals are usually major components of rocks, the effect on the environment during this dissolution process is noticeable, resulting in rocks breaking and the formation of soil and sediment. This mineral deterioration can be referred to as bio-weathering. Bio-weathering can be the result of different mechanisms, oxidoreduction, acidolysis and chelation (Uroz et al., 2009). Mineral oxidoreduction is when bacteria use minerals, or their components, for an electron acceptor. In general, aerobic organisms use oxygen as an electron acceptor when driving the proton motive force. When oxygen is not present the organisms must use another electron acceptor, like sulfate, nitrate or iron (Uroz et al., 2009). However, the weathering is not only a by-product of the process of bacterial metabolism. The bacterial biota can also benefit from the bio-weathering of specific minerals (Bennett et al., 2001). Bio-weathering can also occur when by-products of bacterial metabolism affects the minerals in their environment, like organic acids and protons. These acids or chelating agents can be responsible for three major roles in the weathering process: extracting nutrients from the minerals with electron transfer, braking oxygen links in the minerals, and chelating ions for faster dissolution of the minerals (Uroz et al., 2009). Mineral deterioration of microbes is often driven by the need to access the building blocks forming the minerals, including metals. That need of accessing metals can also result in biomineralisation, the process of precipitating minerals by living organisms from inorganic matter. The process can either be induced by the bacteria or controlled, that is: directly

19 carried out by bacteria. When biomineralisation is controlled, the organism precipitates the mineral inside the cell, a feature that is well known amongst magnetotactic bacteria that precipitate magnetite (Blakemore, 1982; Jogler & Schüler, 2009). Mineralisation that is induced by bacteria is, however, much more common in nature. Induced mineralisation is when the bacteria manipulate or modify their closest environment in any way, such as by releasing organic acids, that results in the precipitation of minerals (Gadd, 2010). This process is highly related to metal transformation where the metal is solubilised (metal mobilisation) and the minerals precipitate on the bacterial surface. The bacterial pathways studied in the field of geomicrobiology, such as metal and mineral biotransformation, decomposition and bioweathering, can easily be exploited for biotechnological uses, such as in bioremediation (Gadd, 2010; Gadd & Pan, 2016), a process often used for environmental cleanup as discussed in detail in my review article for "Biotechnological Applications of Extremophilic Mircroorganisms (Life in Extreme Environments)" (Manuscript II).

1.2 Methane and microbes

As outlined above, bacteria and other microorganisms play an important role in the formation and deterioration of minerals and mobilisation and immobilisation of metals in the environment, which sets the basis for other organisms to be able to take up metals in the right form. Since metals are necessary for living organisms to function, one can wonder why in fact that is, and how did the “pioneer organisms” form these kinds of organic compounds. This question is the foundation of the Iron-Sulfur World Hypothesis about the origin of life by Wächtershäuser (2010). The theory is derived from biogeochemical studies in volcanic flow settings and centres upon, among other things, the dependency on transition metals, synthesis of organic compounds from inorganic nutrients and the chemoautotrophic origin of life. The chemoautotrophic origin is basically referring to the ability of carbon fixation or carbon assimilation, the process of turning inorganic carbon to an organic compound. The structure of a carbon atom forces the atom to form bonds and fill up its valences with hydrogen, so C-C bonds are highly stable and ideally suited for structural purposes of life. Carbon forms stable bonds with oxygen, nitrogen, sulfur and selenium, and together they form the functional groups of organic compounds (Wächtershäuser, 2010). The start of biological evolution begins with this process, and pathways for carbon fixation have been extensively studied in archaea and bacteria (Berg et al., 2010; Fuchs, 2011). For the fixation of carbon, archaea and bacteria can use six different pathways to assimilate carbon dioxide: (1) The Calvin cycle, (2) the reductive

20 citric acid cycle, (3) the reductive acetyl-coenzyme A pathway, (4) the 3- Hydroxypropionate bicycle, (5) hydroxypropionate-hydroxybutyrate cycle and (6) the dicarboxylate-hydroxybutyrate cycle. The reductive acetyl-coenzyme A pathway, also known as the Wood-Ljungdahl pathway, allows the bacteria to reduce carbon dioxide to carbon monoxide and formic acid, which is then reduced to a methyl group for the formation of Acetyl-CoA. This is a process that is well known in the group of “acetogens” and archaeal group of “methanogens” and is the only autotrophic pathway that can fix carbon dioxide and generate ATP simultaneously (Fuchs, 2011). When a pathway of carbon fixation results in the formation of methane as byproduct, the process is called methanogenesis and is performed by archaea. However, methanogens usually need other bacteria to oxidise complex hydrocarbons to short chain hydrocarbons that the archaea can use as substrates for methanogenesis (Stams, 1994; Berg et al., 2010). The progenote of all extant life on Earth is commonly known as LUCA, which is short for “last universal common ancestor”. It refers to the last common ancestor of bacteria and archaea, and represents the link between the abiotic phase and the first signs of life (Weiss et al., 2016). The study that Weiss et al. (2016) carried out, supports the theory of autotrophic origin in hydrothermal settings, where the Wood-Ljungdahl pathway is involved and transition metals and Iron-Sulfur pathways are the basics of metabolism. This suggests that the formation of biologically produced methane could possibly be as old as microbial life. Whether this is in fact the case is uncertain, but hydrothermal or thermogenic methane has been available in excessive amounts since before the origin of life (Wächtershäuser, 2010). If so, one can imagine that progenotes of bacteria needed to tolerate methane or utilise the methane, even prefer it as a carbon source. Which leads us to the next subset of physiological microbial groups: the methanotrophs. This group of methanotrophic bacteria consists of bacteria that can utilise methane as their main carbon source, whether aerobically or anaerobically. Most methanotrophs are said to be obligate methanotrophs. However, a few facultative methanotrophic bacteria have been described (Hanson & Hanson, 1996). When bacteria can utilise methane they are defined by the use of methane monooxygenases that catalyse the oxidation of methane to methanol (Anthony, 1986). The methane oxidation is initiated by methane monooxygenase which can be cytoplasmic and soluble (sMMO) or particulate and membrane bound (pMMO). A membrane bound methane monooxygenase is found in all methanotrophs, at least the ability to express it, but a soluble methane monooxygenase has only been found in two types of methanotrophic bacteria, including Methylococcus species, Methylosinus species and Methylobacterium species (Hanson & Hanson, 1996). Methanotrophs are not fastidious and present low specificity to substrates. This is the reason why this

21 group of methanotrophic bacteria are very interesting for environmental biotechnology, because of their broad metabolism for various compounds and the ability to degrade persistent chemicals. However, the soluble methane monooxygenase has a broader substrate specificity than the membrane bound ones, and can for example oxidise naphthalene to 1- and 2-naphthols that can be used to distinguish sMMO expression, while the pMMO cannot oxidise aromatics, only low-chain alkanes consisting of five carbons or lower (Higgins, Best, & Hammond, 1980; Lipscomb, 1994). The gene expression and regulation of methane monooxygenases has been studied extensively (Murrell et al., 2000). The membrane bound pMMO is encoded in three genes in the pmo gene cluster, each coding for the PmoA, PmoB and PmoC subunits respectively. A copper-binding polypeptide is also expressed along with the pMMO (Lieberman & Rosenzweig, 2004). The soluble sMMO is encoded in six genes that are expressed as an operon; the mmo operon. The enzyme consists of three proteins; a reductase, a hydroxylase where the active site has a di-iron centre, and a component that couples electron and dioxygen consumption (Merkx et al., 2001). Studies on the gene regulation of methane monooxygenases have revealed that copper is the key element to gene expression of methane monooxygenases. As mentioned above, all methanotrophs have pMMO but in those bacteria that also have sMMO gene cluster, the regulation is perfomed with a “copper-switch” where copper-ion starvation initiates the sMMO expression. However, when the copper concentration gets higher in the environment, the copper-ions inhibit the sMMO and the bacteria only express the pMMO for methane oxidation (Murrell et al., 2000). The soluble methane monooxygenase can therefore possibly be a defence mechanism for low nutrient concentration. In anaerobic environments, methane is the most stable of all carbon compounds and is highly important for biogeochemical reactions that lead to mineralization of organic matter (Dagley, 1978; Hanson & Hanson, 1996). The utilization of methane in anaerobic environment is one of the most significant pathways in the earth’s carbon cycle, where microbes in anoxic marine sediments are responsible for the largest amount of methane conversion to carbon dioxide (Boetius et al., 2000). Methanotrophic bacteria are sometimes described as the exact opposite of methanogens, in fact, it has been suggested that methanogenesis and oxidation of methane in anaerobic conditions is the same pathway, only reversed (Scheller, Goenrich, Boecher, Thauer, & Jaun, 2010), even though the pathways for anaerobic oxidation of methane (AOM) are not fully understood. Almost all the methane that is produced by methanogenesis in marine sediments, is also converted to carbon dioxide in the sediments, through anaerobic oxidation of methane coupled with sulfate reduction (Jørgensen & Kasten, 2006). In geothermal sites the gas composition differs from non-geothermal sites depending on the temperature of the geothermal systems. High-temperature

22 geothermal sites have carbon dioxide, hydrogen sulphide and hydrogen in abundant measures while low-temperature geothermal sites have gas with abundant nitrogen. In these systems, natural hydrocarbon gases can occur (Ólafsson et al., 1993). The methane concentration in geothermal systems has been used as parameter for the temperature, and an indicator for low- geothermal systems and high-geothermal systems. High methane concentrations in geothermal gasses also seem to correlate with the gas coming in contact with marine sediments (Arnórsson et al., 2007). This could be linked to the microbial activity in marine sediments where methanogenesis occurs with the reduction of carbon dioxide, resulting in higher methane concentration and lower carbon dioxide concentration.

Figure 2. Schematic representation of the biogeochemistry in marine sediments (Jørgesen & Kasten, 2006). The figure shows the biochemical zones in the relation to pore water chemistry and mineralization processes.

Marine sediments can be sub-divided into biogeochemical zones, where specific groups of bacteria are responsible for specific mineralization processes as reviewed by Jørgensen & Kasten (2006) (see Figure 2). In the uppermost layer of marine sediments, the oxic zone, oxygen respiration is described as the primary process, however since the availability of oxygen is limited, facultative anaerobes are usually more abundant than aerobic bacteria in this environment. Below this oxic zone is the suboxic zone, where reduction of

23 nitrate, manganese and iron is thermodynamically more favourable than sulfate and these processes are dominating in mineralization of organic matter. When the sediment gets anoxic, sulfate reduction becomes the primary process. Sulfate reduction is very important for the oxidation of hydrocarbons in marine sediments, and is constantly fuelled by the methanogenesis in deeper sediments. As sediment depth increases, sulfate supplies become exhausted and are not available for organic carbon oxidation anymore and the anoxic sediment zone reaches into the “methanic” zone, where methanogenesis is the main mineralization process. The methane and carbon dioxide is utilised as a carbon source by bacteria and archaea present in these deep layers of marine sediment, and since methane is the most stable carbon compound, it tends to accumulate and slowly seep up to the sulfidic zone to be utilised (Jørgensen & Kasten, 2006; Xu, 2010). If methane production and accumulation is far greater than can be oxidised by sulfate reduction processes, the methane can seep up all the way through the sediment, resulting in what we call methane gas seepage.

1.2.1 Natural gas seepage

Natural gas seepage is the flow of gaseous hydrocarbons from the subsurface of the Earth. Gaseous bubbles or steam are used as clear indicators of natural gas seepage and the surface gas or near surface sediment gas is generally studied to evaluate if hydrocarbon accumulation is present in underlying layers (Abrams & Dahdah, 2010). Natural gas is mostly compiled of alkanes, with up to 90% methane, but also contains smaller amounts of other gaseous alkanes like ethane, propane and butane. Non-hydrocarbon elements in natural gas can be, for example, carbon dioxide, hydrogen, oxygen, nitrogen, hydrogen sulphide and traces of rare gases, like argon and helium. Natural hydrocarbon gas can therefore be divided in terms of origin, whether it is formed by biochemical reactions (Biogenic), thermochemical reactions (Thermogenic), or by magmatic or post-magmatic processes (Abiotic) of gas formation characterized by the absence of life or living organisms within the earth´s crust (Etiope & Sherwood Lollar, 2013). Thus, when natural hydrocarbon gas is produced by living organisms, it is referred to as “biogenic”. As discussed previously, methane gas is produced by living organisms, including microorganisms through methanogenesis, and is the most common form of natural gas. When natural hydrocarbon gas is formed by a “thermogenic” process, organic matter is matured under high temperature and pressure in deep sedimentary layers as a function of time. Specifically the breakdown of terrestrial organic matter is thought to result in natural gas formation, while organic matter originated from marine environments primarily results in the formation of oil or condensates (Schoell, 1988). However, such formed deep marine oil reservoirs can be the source of gas

24 formation as well, if they are moved into increased temperature and pressure conditions, which is referred to as secondary maturation or petroleum alteration process (Tissot & Welte, 1978; McKenzie & Quigley, 1988). The origin of natural occurring gas can be distinguished by stable isotope ratios, the gas dryness and the ratio of methane to the sum of ethane and propane, since the processes result in different chemical compositions and isotopic signatures, as shown in Figure 3 (Schoell, 1988; Xu, 2010).

Figure 3. Plots from Xu (2010) to distinguish the origin of methane. (a) The stable isotope ratio plotted against carbon ratios to determine microbial, geothermal or thermogenic origin of methane. (b) Stable isotope ratios of hydrogen and carbon used to determine the microbial, thermogenic and geothermal origin of methane.

25 1.2.2 Cold seeps and gas hydrates

Cold seeps are found in marine sediments along active continental margins and consist of hydrocarbon seeps that have no obvious source of heat. Cold seeps are often linked to gas hydrates, which are crystallised gas that form ice-like deposits, usually methane crystals. For the past two decades, cold seeps and methane gas hydrates have attracted interest in the field of geomicrobiology (Zhang & Lanoil, 2004). These environments seem to be ideal to study anaerobic oxidation of methane and the link to sulfate reduction, and how biogeochemical processes contribute to the macrofauna present at these sites (Levin, 2005). Biogeochemical studies on cold methane seeps in the Gulf of Mexico indicated that the link between anaerobic oxidation of methane and sulfate reduction might not be that critical, but other hydrocarbons could be more important than first thought. The utilisation of other hydrocarbon compounds than methane are likely to fuel sulfate reduction as well (Joye et al., 2004), and sulfate reducing bacteria are capable of oxidising various hydrocarbons, like short and long chain alkanes and aromatic compounds, by using sulfate as electron acceptor. This could suggest that sulfate reducing bacteria are possibly the primary hydrocarbon utilisers at methane gas seepage sites. Methanotrophic bacteria and sulfate-reducing bacteria have also been studied in relation to methane seeps and gas emission from submarine mud volcanos, where both aerobic and anaerobic oxidation of methane seems to be relevant (Niemann et al., 2006). The methanotrophic bacteria found at gas seepage sites largely consist of members of the Proteobacteria phylum, like Methylobacter and Methylophaga (Lösekann et al., 2007) and Desulfosarcina/Desulfococcus (Knittel et al., 2005), that have been reported as important alkane degraders in marine seepage sites (Kleindienst et al., 2014).

1.2.3 Coal bed methane and associated bacteria

Coal beds and petroleum reservoirs are found in deeper organic sediment layers. Methane that is trapped in coal beds is referred to as coal bed methane and can be produced either biogenically or thermogenically from coal beds in organic layers (Moore, 2012). The biogenically formed coal bed methane is the result of bacteria converting the coal to carbon dioxide or acetate, where archaea take over and reduce it to methane The thermogenically formed coal bed methane is the result or the byproduct of coalification, the process where coal beds are formed by heat and pressure in organic sediment layers. When coal beds are forming, it starts by transforming organic material into peat through a process called peatification, where organic matter in wetlands does not fully decompose before it gets buried. At this stage, the methane production is largely of biogenic origin. After that, heat and pressure cause dehydration of the peat and result in the formation of lignite, also referred to as brown-coal.

26 With continued burial, higher temperature and pressure, the coalification continues with bituminisation where lignite is transformed to sub-bituminous and bituminous black coal, and later anthracite (see Figure 4). With deeper burial, heat and pressure, the coal material becomes denser with higher concentrations of carbon, lower concentrations of oxygen, and condensed with polycyclic aromatic ring system (Strąpoć et al., 2011). The bituminisation is believed to be very important process in the coalification of bituminous coals where petroleum-like material is generated (Teichmüller, 1987). During this process, when the coals are becoming more concentrated, methane gas is formed and tends to get trapped in the coal beds, hence, coal bed methane (Moore, 2012). Methane that is bound in coal beds lacks hydrogen sulphide and is often composed of lighter alkanes and a low ratio of heavier gaseous hydrocarbons like propane and butane, compared to other natural gasses, and can therefore be used as a relatively clean and inexpensive fuel. Because of this, coal bed methane has been unconventionally produced as fuel production by burying organic material fast and heating it up.

Figure 4. Formation of methane gas in coalification processes in relation to organic material, vitrinite reflectance and dryness, figure from Tim A Moore (2012).

27 The organic material in coals has different aromatic composition where anthracite, for example, consists of highly aromatic structures compared to bituminous coal (Yang et al., 2017). When bituminous coals are being transformed into anthracite, it is not only a thermogenic process (see Figure 4.) but also results in methane formation of biogenic origin. As mentioned above, coal bed methane can be produced by microorganisms and for the past two decades, accumulation of methane of microbial origin has been detected in several coal beds, so the presence of microbes might possibly be important to later coalification processes, or at least some part of it (Strąpoć et al., 2011). Studies on the microbes present in coal beds have mainly been focused on the methanogenic archaea and the microbial production of methane. Enrichment studies on bacteria from coal beds can be very useful when bioprospecting for special biochemical traits, and has been used for the characterisation of methanogenic microbes (Green et al., 2008; Penner et al., 2010; Stra̧ poć et al., 2008). Even though enrichments and isolation studies serve a meaningful function in studies of coal bed microbes, the enrichments tend to get very selective and favour bacteria that are easily grown in vitro, in that way missing out on other interesting bacteria is a matter of concern. For this reason, molecular genetic analysis on microbes from environmental samples is important to gather information about what bacteria can be found in a specific environment, even though it is not possible to enrich or isolate them. With cloning techniques and DNA sequencing, the microbiota of several coal beds have been described and suggest the abundancy of methanogenic archaea, but also a number of other bacteria with possibly different roles (Beckmann et al., 2011; Penner et al., 2010; Stra̧ poć et al., 2008). When the microbial community in coal beds is studied as a whole with molecular methods, as opposed to enrichments, the presence of Comamondaceae and Geobacter families within the Proteobacteria phylum is commonly reported along with the Clostridiales and more members of the Firmicutes phylum, as well as members of the Actinobacteria and Bacteroidetes phyla (Li et al., 2008; Midgley et al., 2010; Shimizu et al., 2007; Stra̧ poć et al., 2008). These studies rely on phylogenetic information to predict the role of non-methanogenic bacteria in the microbial community of coal beds and associating groundwater, and the presumptive metabolism of these bacteria shows that hydrocarbon degradation is possibly highly important and suggest denitrification and fermentation to be abundant metabolism. The Proteobacteria phylum harbours a variety of nitrate-reducing species and hydrocarbon utilisers that seem to be abundant in coal beds (Barnhart et al., 2013; Penner et al., 2010) and if the coals are influenced by marine water it could be expected to find Deltaproteobacteria involved in sulphur cycling or use ferric iron as terminal electron acceptor like Geobacter species (Midgley et al., 2010). Bacteria in the Bacteroidetes and Firmicutes phyla are considered to be fermenters in coal beds (Penner et al., 2010) but in

28 these phyla are acetogenic bacteria that can possibly be fuelling methanogenesis instead of hydrogenotrophic methanogens (Beckmann et al., 2011). Most of the Actinobacteria found in coal beds are in the suborder Micrococcineae, and are believed to be involved in anaerobic metal reduction and can therefore be assisting the formation of coals (Midgley et al., 2010).

1.2.4 Identifying bacterial communities in natural gas seepage with amplicon sequencing

For the past few years, the interest in amplicon sequencing of microbial DNA from any kind of environment has grown, since it can provide an understanding of the microbial communities without culturing limitations, and can be of great importance to microbial ecology. Microbial communities change across different environments depending on geographic distances, temperature, oxygen, salinity, pH and other biotic factors (Thompson et al., 2017). To identify an entire microbial community in a specific environment, the environmental DNA is extracted and a bacterial-specific DNA sequence is amplified and compared. By amplifying variable regions within the ribosomal gene 16S rRNA it is possible to sequence the same DNA region of almost all the bacteria present in that environment. The ribosomal 16S gene in bacteria codes for the small subunit of ribosomes that engage in translation, and is therefore a highly conserved gene within the bacterial domain. However, the subunit has active sites and non-active sites, and these non-active sites do not need to be conserved between genera or species. The mutations in the DNA sequence of these non-active sites can be used to differentiate between bacterial genera with considerable certainty. Bacterial community composition studies described in the previous chapter, mainly consist of molecular genetic studies based on cloning and Sanger sequencing. Since high-throughput sequencing (HTS) is constantly becoming faster and less expensive, this method is becoming more widespread and commonly used in microbial ecology and geomicrobiology. For example, the Earth Microbiome Project is a project set to map microbial communities all around the planet with standardised methods for more accurate comparison between environments (Gilbert et al., 2014). Great advances have been made in high throughput sequencing for the past decade and many different sequencing technologies have been made commercially available, for example the most dominant technology from Illumina and the most recent technology from Oxford Nanopore Technologies, but high throughput sequencing technologies have been reviewed multiple times (Mardis, 2017; Metzker, 2010; Reuter et al., 2015; Shendure & Ji, 2008; Shokralla et al., 2012). Even though HTS technologies were originally aimed towards genome sequencing, they are very well suited for amplicon sequencing of bacterial genes, since they generate millions of short sequences and enable sequencing of multiple

29 samples in one run. This has practically revolutionised microbial ecology studies. The basis of the advancement in sequencing technology is rapid developments in computer sciences. High throughput sequencing relies on computer algorithms to analyse and interpret the data generated during sequencing, and numbers of different platforms specifically for the analysis bacterial communities have been made available, like QIIME (Caporaso et al., 2010), Mothur (Schloss et al., 2009) and UPARSE (Edgar, 2013). The analysis of amplicon sequencing data consists of quality filtering and denoising of sequences, eliminating chimeric sequences and clustering the sequences together into “operational taxonomic unit” or OTU, that represent a bacterial taxon. The OTU sequences are then compared to databases, for example Greengenes (DeSantis et al., 2006) or SILVA (Pruesse et al., 2007), that consist of sequences of already identified genera and statistical analysis used to estimate abundance and diversity, within or between bacterial communities. The bacterial communities in natural gas seepage and coal beds are no exception from this spiking interest in bacterial communities, and recently there was published a reference set for microbes found in coal seams (Vick et al., 2018). Amplicon sequencing of the microbes present in this environment has been studied where the community has abundant anaerobic taxa (Case et al., 2015; Kirk et al., 2015; Ruff et al., 2016) in agreement with cloning-based studies as discussed in previous chapter. Kirk et al. (2015) hypothesised that the bacterial composition in coal bearing strata change regarding to the coal maturity, because the chemical structure of the coal changes with maturity, and therefore bacteria harbouring other enzymes are needed for degradation, resulting in different bacterial community structure. Community analysis with amplicon sequencing has also revealed the presence of aerobic hydrocarbon degrading bacteria associated with coal beds and other hydrocarbon rich environments even though the environment has commonly been noted as anoxic (An et al., 2013). This could suggest that the biogeochemistry related to coal bed methane is not entirely dependent on anaerobic processes, and that the microbial community in coal beds serve multiple different biogeochemical roles. Microbial communities in methane seeps have been shown to be more similar to one another than the microbial community in surrounding sediment (Ruff et al., 2016) indicating that the natural gas flow has an effect on the community structure. This means that the gas origin is likely to be a factor in the microbial community structure, leading to the possibility of the bacterial community in coal gas seepage being different from methane seeps of biogenic origin. In environments like marine methane seeps, the Chloroflexi phylum is repeatedly reported, especially the Anaerolinea class (Case et al., 2015; Kirk et al., 2015; Ruff et al., 2016) and that the Anaerolinea play a key role in sediment carbon cycling (Hug et al., 2013). However, Ruff et al. (2016) conclude that shallow methane seeps are very different from deep sea methane seeps further encouraging studies on shallow methane seeps.

30 1.3 Geological settings of Öxarfjörður

Iceland’s geology in general is characterized by the complex processes along its plate boundary between the North-American and the Eurasian lithospheric plates, traversing across Iceland from the southwest to the northeast. The boundaries are far from being simple, featuring both diverging margins and conservative plate margins, meaning that the plate boundaries alternate between dividing the island along with formation of new crust, and moving beside one another in friction-like movements along fracture zones (Sæmundsson, 1974). This results in the formation of complex volcanic rift systems and frequently occurring earthquakes. In contrast to the high temperature processes connected to active volcanism and tectonics, is the islands northern location straddling the arctic circle that results in glacial formations and glacial river systems, which also have massively contributed to Iceland’s geology and landscaping. Along the obliquely extending plate margins form onshore and offshore rift graben valleys, such as the Northern Volcanic Zone (NVZ) and the Grímsey oblique rift system (GORS) of northeast Iceland that is formed by overlapping multiple rift systems, such as the Theistareykir (Th), the Krafla, (K), or the Fremrí-Námar (N) volcanic rift system onshore (e.g. Sæmundsson, 1974; Jakobsson et al., 2008; Einarsson et al. 2008; Magnúsdóttir et al. 2015) (Figure 5). These rift valleys are filled by extrusive and intrusive igneous and erosional material, and can form complex and deep sedimentary graben fills structures. The Öxafjörður graben is this study´s field site and represents one of the deeper graben structures in Iceland, and is located at the onshore to offshore oblique rift transition within the NVZ (Figure 5 & Figure 6). Here, specifically, is a connection between the Krafla and Theistareykir volcanic rift systems directly to the Grímsey oblique rift system, which is part of the Tjörnes Fracture Zone (Ólafsson et al., 1993; Sæmundsson, 1974) (Figure 5). As this region is tectonically very active and extending, rapid subsidence of these graben systems follows, which is simultaneously filled in by extrusive and intrusive rocks of variable composition, but primarily by sediment input from local river systems, here specifically the Jökulsá á Fjöllum glacial river system. These volcanic rift systems are comprised of central volcanic complexes and fissure swarms, which are basically long fault systems and cracks in the earth´s crust that form pathways for rising magma and hot fluids, venting gases and fluids vertically. Three major swarms, cross-section the Öxarfjörður graben from west to east. The largest fissure swarm is tens of kilometres long and extends from the Krafla central volcano through the central area of the Öxarfjörður, while the Theistareykir fissure swarm straddles the western margin, and the Fremri-námar fissure swarm forms the eastern rift valley graben boundary (Figure 5).

31 0 15 30 60 Kilometers

Th DL K

N H

KF Vatnajökull glacier

Figure 5. The northeast Iceland plate bondaries and rift systems, and location of the study area within the Northern Volcanic Zone (modified from Einarsson et al., 2008; Jakobsson et al., 2008; Magnúsdóttir et al., 2015; Hjartarson & Sæmundsson, 2014). Abbreviations are: TJ – Tjörnes peninsular sediment outcrops; structural lineament

32 segments DL – Dalvíkur lineament and the HFF – Húsavík-Flatey-Fault system; volcanic rift complexes A – Askja, H - Hrúthálsar , K – Krafla, KF – Kverkfjöll, N – Fremri-Námar, and Th – Theistareykir.

The Öxarfjörður as a low geothermal area and the nature of the fissure swarms have been studied for the geothermal manifestations since the 1970´s (e.g. Sæmundsson, 1974; Ólafsson et al., 1993; Georgsson et al., 2000). In 1991 the deepest well in the Öxarfjörður section of the Krafla fissure swarm was drilled at the Skógarlón site, where the water temperature reached up to 100 °C at a shallow level (Ólafsson et al., 1993) (Figure 6). For geothermal and gas exploration it is important to have detailed knowledge of the present and underlying sedimentary stratification (Ólafsson et al., 1993), and in the Öxarfjörður area the sedimentary stratification is atypical for Iceland´s geology, as older thick sediment and coal seam layers are preserved within the deep graben structure. West of Öxarfjörður is the Tjörnes peninsula (TJ on Figure 5), where the thickest marine sediments in Iceland can be found. Sedimentation is a result of many different factors besides tectonics, including climate, biological factors and weathering. On diverging plate boundaries new crust is formed so Öxarfjörður can be regarded as a presently developing and young basin in Iceland. Even though this might be true, the occurrence of older sediments in the deeper layers of the graben is quite possible. After the deep drilling at Skógarlón, a better understanding of the sediments in Öxarfjörður was revealed. The marine sediments measured up to 1.000 m thick and the uppermost 350 m of the sediment have been developing since 10.000 years ago (Ólafsson et al., 1993). Natural gas seepages have been detected in the Öxarfjörður geothermal field, in accordance with the fissure swarms. These hydrocarbon rich gases are believed to be emitted through the sediment along the Öxarfjörður area, resulting in bubbling pockmarks, both offshore and onshore (Ólafsson et al., 1993; Richter & Gunnarsson, 2010). In 1989-1991 geothermal holes were drilled by Skógalón and the origin and composition of the gas was analysed to determine the hydrocarbon potential of the detected gases. The gasses detected were over 20,000 years old thermogenic gasses, both wet and dry, with fairly high concentrations of evolved hydrocarbon gases (methane through hexane) (Ólafsson et al., 1993; Richter & Gunnarsson, 2010). The stable isotope ratio of the natural gas suggests that the sediment layers underneath were bituminous, and the gas is a mixture between biogenic and thermogenic coal gas (Ólafsson et al., 1993). Polycyclic aromatic hydrocarbons (PAH) have also been detected in the same area, suggesting accumulation of hydrocarbons in the sediment (Geptner et al., 2006). PAH are amongst the geochemical constituents that indicate hydrocarbon flow, whether it is a natural hydrocarbon flow or induced by humans, and can suggest hydrocarbon seepage from petroleum deposits or active hydrothermal vents.

Figure 6. Sample site locations within the Öxarfjörður graben system, existing boreholes, inferred extensional fault locations, and surface depositional environments of the area (Ólafsson et al. 1993 and Sæmundsson et al. 2012).

34 1.4 Aims of the present study

Geomicrobiology and environmental biotechnology are fields that have thus far received very little attention from the Icelandic research community. The microbiota in Öxarfjörður´s natural gas seepage pockmarks is entirely undiscovered and the possibility of finding bacterial taxa of interest is substantial. It has been pointed out that Iceland is actually an ideal place to study natural PAH flow of hydrothermal origin since there is very little industrial influence on the environment that can interfere with PAH measurements (Geptner et al., 2006). However, the associated microbiota have not recieved much attention. Studying this kind environment is of great importance since Iceland is always becoming more of a target to environmental pollution due to ever-growing transportation. To be able to make use of bioremediation processes in Iceland, it is critical to know what kind of microbes inhabit our environment and in what way they interact with the nature and each other in terms of biochemistry and geochemistry. No previous studies have been conducted on the microbial communities of natural hydrocarbon seepage sites within the Icelandic rift zone even though the Icelandic rift zone provides a continuous source of low-chain, high-chain and polycyclic aromatic hydrocarbon flow to the environment through natural hydrocarbon seepage (Geptner et al., 2006; Ólafsson et al., 1993). This study is therefore the first geomicrobiological study of natural hydrocarbon seepage sites in Icelandic rift zone area and will provide insight into the microbial community of natural gas seepage pockmarks, located in Öxarfjörður. The study will also provide isolated bacterial strains that can be further investigated for bioremediative purposes and is vital for better understanding of the biogeochemistry of the environment.

This study is based on the concepts of geomicrobiology and bioprospecting extreme environments for bacteria that can be useful in environmental biotechnology. The thesis consists of two distinct but related manuscripts. The first manuscript is a research article about the bacterial community composition of natural gas seepage pockmarks at two distinct sampling sites in Öxarfjörður and the isolation of bacterial strains from that environment. The second manuscript is a review chapter about the bioremediative potential of bacteria in cold desert environments.

Manuscript I: Gas seepage pockmark microbiomes suggest the presence of sedimentary coal seams in the Öxarfjörður graben of NE- Iceland

35 Manuscript II: “Bioremediative potential of bacteria in cold desert environments” in Biotechnological Applications of Extremophilic Microorganisms (Life in Extreme Environments) (In press)

The aim of manuscript I was to: (1) Identify the bacterial taxa located in the natural gas seepage pockmarks of two distinct sites in Öxarfjörður, (2) detect biochemically relevant taxa within that community and (3) isolate bacterial strains that can possibly be useful in environmental biotechnology and bioremediation. The following research questions were used for the research:

• Is the natural gas in the gas seepage pockmarks of the same origin at both sampling sites? • Is the microbial community of the natural gas seepage pockmarks biochemically relevant to the composition of the natural methane gas detected from the pockmarks? • Are there taxa within the microbial community of the gas seepage pockmarks that can possibly be used for environmental biotechnological purposes? • Is it possible to isolate hydrocarbon-degrading or methane-utilising bacteria from the natural gas seepage pockmarks in Öxarfjörður?

36 References

Abrams, M. A., & Dahdah, N. F. (2010). Surface sediment gases as indicators of subsurface hydrocarbons - examining the record in laboratory and field studies. Marine and Petroleum Geology, 27(1), 273–284. An, D., Caffrey, S. M., Soh, J., Agrawal, A., Brown, D., Budwill, K., … Voordouw, G. (2013). Metagenomics of Hydrocarbon Resource Environments Indicates Aerobic Taxa and Genes to be Unexpectedly Common. Environmental Science & Technology, 47(18), 10708–10717. Anthony, C. (1986). Bacterial Oxidation of Methane and Methanol. Advances in Microbial Physiology, 27, 113–210. Arnórsson, S., Stefánsson, A., & Bjarnason, J. O. (2007). Fluid-Fluid Interactions in Geothermal Systems. Reviews in Mineralogy and Geochemistry, 65(1), 259–312. Barnhart, E. P., De León, K. B., Ramsay, B. D., Cunningham, A. B., & Fields, M. W. (2013). Investigation of coal-associated bacterial and archaeal populations from a diffusive microbial sampler (DMS). International Journal of Coal Geology, 115, 64–70. Beckmann, S., Krüger, M., Engelen, B., Gorbushina, A. A., & Cypionka, H. (2011). Role of bacteria, archaea and fungi involved in methane release in abandoned coal mines. Geomicrobiology Journal, 28(4), 347–358. Bennett, P. C., Rogers, J. R., Choi, W. J., & Hiebert, F. K. (2001). Silicates, silicate weathering, and microbial ecology. Geomicrobiology Journal, 18(1), 3–19. Berg, I. A., Kockelkorn, D., Ramos-Vera, W. H., Say, R. F., Zarzycki, J., Hügler, M., … Fuchs, G. (2010). Autotrophic carbon fixation in archaea. Nature Reviews Microbiology. Nature Publishing Group. Blakemore, R. P. (1982). Magnetotactic Bacteria. Annual Review of Microbiology, 36(1), 217–238. Boetius, A., Ravenschlag, K., Schubert, C. J., Rickert, D., Widdel, F., Gleseke, A., … Pfannkuche, O. (2000). A marine microbial consortium apparently mediating anaerobic oxidation methane. Nature, 407(6804), 623–626. Caporaso, J. G., Kuczynski, J., Stombaugh, J., Bittinger, K., Bushman, F. D., Costello, E. K., … Knight, R. (2010). QIIME allows analysis of high- throughput community sequencing data. Nature Methods, 7(5), 335–336. Case, D. H., Pasulka, A. L., Marlow, J. J., Grupe, B. M., Levin, L. A., & Orphan, V. J. (2015). Methane seep carbonates host distinct, diverse, and dynamic microbial assemblages. mBio, 6(6), e01348-15. Dagley, S. (1978). Determinants of biodegradability. Quarterly Reviews of Biophysics, 11(4), 577–602. DeSantis, T. Z., Hugenholtz, P., Larsen, N., Rojas, M., Brodie, E. L., Keller,

37 K., … Andersen, G. L. (2006). Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Applied and Environmental Microbiology, 72(7), 5069–72. Edgar, R. C. (2013). UPARSE: Highly accurate OTU sequences from microbial amplicon reads. Nature Methods, 10(10), 996–998. Einarsson, P. (2008). Plate boundaries, rifts and transforms in Iceland. Jökull, 58(12), 35-58. Etiope, G., & Sherwood Lollar, B. (2013). Abiotic methane on Earth. Reviews of Geophysics, 51(2), 276-299. Fuchs, G. (2011). Alternative Pathways of Carbon Dioxide Fixation: Insights into the Early Evolution of Life? Annual Review of Microbiology, 65(1), 631–658. Gadd, G. M. (2010). Metals, minerals and microbes: Geomicrobiology and bioremediation. Microbiology. Gadd, G. M., & Pan, X. (2016). Biomineralization, bioremediation and biorecovery of toxic metals and radionuclides. Geomicrobiology Journal, 33(3–4), 175–178. Georgsson, L. S., Friðleifsson, G. Ó., Ólafsson, M., & Flóvenz, Ó. G. (2000). The geothermal exploration of the Öxarfjördur high-temperature area, NE-Iceland. In Proceedings of the World Geothermal Congress 2000, Kyushu-Tohoku, Japan(pp. 1157-1162). Geptner, A. R., Richter, B., Pikovskii, Y. I., Chernyansky, S. S., & Alexeeva, T. A. (2006). Polycyclic aromatic hydrocarbons as evidence of hydrocarbon migration in marine and lagoon sediments of a recent rift zone (Skjálfandi and Öxarfjörður), Iceland. Chemie Der Erde - Geochemistry, 66(3), 213–225. Gilbert, J. A., Jansson, J. K., & Knight, R. (2014). The Earth Microbiome project: Successes and aspirations. BMC Biology. BioMed Central. Green, M. S., Flanegan, K. C., & Gilcrease, P. C. (2008). Characterization of a methanogenic consortium enriched from a coalbed methane well in the Powder River Basin, U.S.A. International Journal of Coal Geology, 76(1–2), 34–45. Hanson, R. S., & Hanson, T. E. (1996). Methanotrophic bacteria. Microbiological Reviews, 60(2), 439–71. Higgins, I. J., Best, D. J., & Hammond, R. C. (1980). New findings in methane- utilizing bacteria highlight their importance in the biosphere and their commercial potential. Nature, 286(5773), 561–564. Hjartarson, Á. & Sæmundsson, K. (2014). Geological Map of Iceland. Bedrock. 1:600 000. Iceland GeoSurvey. Hug, L. A., Castelle, C. J., Wrighton, K. C., Thomas, B. C., Sharon, I., Frischkorn, K. R., … Banfield, J. F. (2013). Community genomic analyses constrain the distribution of metabolic traits across the Chloroflexi phylum and indicate roles in sediment carbon cycling.

38 Microbiome, 1(1), 22. Jakobsson, M., Macnab, R., Mayer, L., Anderson, R., Edwards, M., Hatzky, J., ... & Johnson, P. (2008). An improved bathymetric portrayal of the Arctic Ocean: Implications for ocean modeling and geological, geophysical and oceanographic analyses. Geophysical Research Letters, 35(7). Jogler, C., & Schüler, D. (2009). Genomics, Genetics, and Cell Biology of Magnetosome Formation. Annual Review of Microbiology, 63(August), 501–521. Jørgensen, B. B., & Kasten, S. (2006). Sulfur cycling and methane oxidation. In Marine Geochemistry (pp. 271–309). Berlin/Heidelberg: Springer- Verlag. Joye, S. B., Boetius, A., Orcutt, B. N., Montoya, J. P., Schulz, H. N., Erickson, M. J., & Lugo, S. K. (2004). The anaerobic oxidation of methane and sulfate reduction in sediments from Gulf of Mexico cold seeps. Chemical Geology, 205(3–4), 219–238. Kirk, M. F., Wilson, B. H., Marquart, K. A., Zeglin, L. H., Vinson, D. S., & Flynn, T. M. (2015). Solute concentrations influence microbial methanogenesis in coal-bearing strata of the Cherokee basin, USA. Frontiers in Microbiology, 6(NOV), 1287. Kleindienst, S., Herbst, F.-A., Stagars, M., von Netzer, F., von Bergen, M., Seifert, J., … Knittel, K. (2014). Diverse sulfate-reducing bacteria of the Desulfosarcina/Desulfococcus clade are the key alkane degraders at marine seeps. The ISME Journal, 8(10), 2029–2044. Knittel, K., Lösekann, T., Boetius, A., Kort, R., & Amann, R. (2005). Diversity and distribution of methanotrophic archaea at cold seeps. Applied and Environmental Microbiology, 71(1), 467–479. Levin, L. A. (2005). Ecology of cold seep sediments: Interactions of fauna with flow, chemistry and microbes. An Annual Review, 43, 1–46. Li, D., Hendry, P., & Faiz, M. (2008). A survey of the microbial populations in some Australian coalbed methane reservoirs. International Journal of Coal Geology, 76(1–2), 14–24. Lieberman, R. L., & Rosenzweig, A. C. (2004). Biological methane oxidation: Regulation, biochemistry, and active site structure of particulate methane monooxygenase. Critical Reviews in Biochemistry and Molecular Biology. Lipscomb, J. D. (1994). Biochemistry of the soluble methane monooxygenase. Annu Rev Microbiol, 48(1), 371–399. Lösekann, T., Knittel, K., Nadalig, T., Fuchs, B., Niemann, H., Boetius, A., & Amann, R. (2007). Diversity and abundance of aerobic and anaerobic methane oxidizers at the Haakon Mosby Mud Volcano, Barents Sea. Applied and Environmental Microbiology, 73(10), 3348–3362. Magnúsdóttir, S., Brandsdóttir, B., Driscoll, N., & Detrick, R. (2015). Postglacial tectonic activity within the Skjálfandadjúp Basin, Tjörnes

39 Fracture Zone, offshore Northern Iceland, based on high resolution seismic stratigraphy. Marine Geology, 367, 159-170. Mardis, E. R. (2017). DNA sequencing technologies: 2006-2016. Nature Protocols. Nature Publishing Group. Mackenzie, A. S., Price, I., Leythaeuser, D., Muller, P., Radke, M., & Schaefer, R. G. (1987). The expulsion of petroleum from Kimmeridge Clay source- rocks in the area of the Brae oilfield, UK continental shelf. Petroleum geology of north west Europe, 2, 865-877. Merkx, M., Kopp, D. A., Sazinsky, M. H., Blazyk, J. L., Muand̈ ller, J., & Lippard, S. J. (2001). Dioxygen activation and methane hydroxylation by soluble methane monooxygenase: A tale of two irons and three proteins. Angewandte Chemie - International Edition. WILEY‐VCH Verlag GmbH. Metzker, M. L. (2010). Sequencing technologies the next generation. Nature Reviews Genetics. Nature Publishing Group. Midgley, D. J., Hendry, P., Pinetown, K. L., Fuentes, D., Gong, S., Mitchell, D. L., & Faiz, M. (2010). Characterisation of a microbial community associated with a deep, coal seam methane reservoir in the Gippsland Basin, Australia. International Journal of Coal Geology, 82(3–4), 232– 239. Moore, T. A. (2012). Coalbed methane: A review. Int. J. Coal Geol., 101, 36– 81. Murrell, J. C., McDonald, I. R., & Gilbert, B. (2000). Regulation of expression of methane monooxygenases by copper ions. Trends in Microbiology. Elsevier Current Trends. Niemann, H., Lösekann, T., de Beer, D., Elvert, M., Nadalig, T., Knittel, K., … Boetius, A. (2006). Novel microbial communities of the Haakon Mosby mud volcano and their role as a methane sink. Nature, 443(7113), 854–858. Ólafsson, M., Friðleifsson, G. Ó., Eiríksson, J., & Ármannsson, H. (1993). On the origin of organic gas in Öxarfjörður, NE-Iceland. Orkustofnun. Penner, T. J., Foght, J. M., & Budwill, K. (2010). Microbial diversity of western Canadian subsurface coal beds and methanogenic coal enrichment cultures. International Journal of Coal Geology, 82(1–2), 81–93. Pruesse, E., Quast, C., Knittel, K., Fuchs, B. M., Ludwig, W., Peplies, J., & Glockner, F. O. (2007). SILVA: a comprehensive online resource for quality checked and aligned ribosomal RNA sequence data compatible with ARB. Nucleic Acids Research, 35(21), 7188–7196. Reuter, J. A., Spacek, D. V., & Snyder, M. P. (2015). High-Throughput Sequencing Technologies. Molecular Cell. Cell Press. Richter, B., & Gunnarsson, K. (2010). Overview of hydrocarbon related research in Tjörnes. Gagnavefsja.Is. Retrieved from

40 http://www.gagnavefsja.is/media/olia/2010_007_Hydrocarbon_research _Tjornes.pdf Ruff, S. E., Kuhfuss, H., Wegener, G., Lott, C., Ramette, A., Wiedling, J., … Weber, M. (2016). Methane seep in shallow-water permeable sediment harbors high diversity of anaerobic methanotrophic communities, Elba, Italy. Frontiers in Microbiology, 7(MAR), 374. Sæmundsson, K. (1974). Evolution of the axial rifting zone in Northern Iceland and the Tjörnes fracture zone. Bulletin of the Geological Society of America, 85(4), 495–504. Sæmundsson, K., Sigurgeirsson, M. Á. & Grönvold, K. (2012). Þeistareykir. Jarðfræðirannsóknir 2011. Iceland GeoSurvey, ÍSOR2012/024. Scheller, S., Goenrich, M., Boecher, R., Thauer, R. K., & Jaun, B. (2010). The key nickel enzyme of methanogenesis catalyses the anaerobic oxidation of methane. Nature, 465(7298), 606–608. Schloss, P. D., Westcott, S. L., Ryabin, T., Hall, J. R., Hartmann, M., Hollister, E. B., … Weber, C. F. (2009). Introducing mothur: open-source, platform-independent, community-supported software for describing and comparing microbial communities. Applied and Environmental Microbiology, 75(23), 7537–41. Schoell, M. (1988). Multiple origins of methane in the Earth. Chemical Geology, 71(1–3), 1–10. Shendure, J., & Ji, H. (2008). Next-generation DNA sequencing. Nature Biotechnology. Nature Publishing Group. Shimizu, S., Akiyama, M., Naganuma, T., Fujioka, M., Nako, M., & Ishijima, Y. (2007). Molecular characterization of microbial communities in deep coal seam groundwater of northern Japan. Geobiology, 5(4), 423–433. Shokralla, S., Spall, J. L., Gibson, J. F., & Hajibabaei, M. (2012). Next- generation sequencing technologies for environmental DNA research. Molecular Ecology. Blackwell Publishing Ltd. Sparks, D. L. (2005). Toxic Metals in the Environment: The Role of Surfaces. Elements, 1(4), 193–197. Stams, A. J. (1994). Metabolic interactions between anaerobic bacteria in methanogenic environments. Antonie van Leeuwenhoek, 66(1-3), 271- 294. Strąpoć, D., Mastalerz, M., Dawson, K., Macalady, J., Callaghan, A. V., Wawrik, B., … Ashby, M. (2011). Biogeochemistry of Microbial Coal- Bed Methane. Annual Review of Earth and Planetary Sciences, 39(1), 617–656. Stra̧ poć, D., Picardal, F. W., Turich, C., Schaperdoth, I., Macalady, J. L., Lipp, J. S., … Schimmelmann, A. (2008). Methane-producing microbial community in a coal bed of the Illinois Basin. Applied and Environmental Microbiology, 74(8), 2424–2432. Teichmüller, M. (1987). Recent advances in coalification studies and their

41 application to geology. Geological Society, London, Special Publications, 32(1), 127–169. Thompson, L. R., Sanders, J. G., McDonald, D., Amir, A., Ladau, J., Locey, K. J., … Zhao, H. (2017). A communal catalogue reveals Earth’s multiscale microbial diversity. Nature, 551(7681), 457–463. Tissot, B. P., & Welte, D. H. (1978). Petroleum Formation and Occurrence. Berlin, Heidelberg: Springer Berlin Heidelberg. Uroz, S., Calvaruso, C., Turpault, M. P., & Frey-Klett, P. (2009). Mineral weathering by bacteria: ecology, actors and mechanisms. Trends in Microbiology. Van Der Heijden, M. G. A., Bardgett, R. D., & Van Straalen, N. M. (2008). The unseen majority: Soil microbes as drivers of plant diversity and productivity in terrestrial ecosystems. Ecology Letters. Blackwell Publishing Ltd. Vick, S. H. W., Greenfield, P., Tran-Dinh, N., Tetu, S. G., Midgley, D. J., & Paulsen, I. T. (2018). The Coal Seam Microbiome (CSMB) reference set, a lingua franca for the microbial coal-to-methane community. International Journal of Coal Geology, 186, 41–50. Wächtershäuser, G. (2010). Chemoautotrophic origin of life: The iron-sulfur world hypothesis. In Geomicrobiology: Molecular and Environmental Perspective (pp. 1–35). Dordrecht: Springer Netherlands. Weiss, M. C., Sousa, F. L., Mrnjavac, N., Neukirchen, S., Roettger, M., Nelson-Sathi, S., & Martin, W. F. (2016). The physiology and habitat of the last universal common ancestor. Nature Microbiology, 1(9), 16116. Xu, H. (2010). Synergistic roles of microorganisms in mineral precipitates associated with deep sea methane seeps. In Geomicrobiology: Molecular and Environmental Perspective (pp. 325–346). Dordrecht: Springer Netherlands. Yang, F., Hou, Y., Wu, W., & Liu, Z. (2017). Structures of Aromatic Clusters of Different Coals Based on Benzene Carboxylic Acids from Coal via Oxidation. Energy & Fuels, 31(11), 12072–12080. Zhang, C. L., & Lanoil, B. (2004). Geomicrobiology and biogeochemistry of gas hydrates and cold seeps. Chemical Geology, 205(3–4), 187–194.

42 2 Manuscript I - Gas seepage pockmark microbiomes suggest the presence of sedimentary coal seams in the Öxarfjörður graben of NE-Iceland

Guðný Vala Þorsteinsdóttir1,2, Anett Blischke3, M. Auður Sigurbjörnsdóttir1, Finnbogi Óskarsson4, Bjarni Gautason3, Þórarinn Sveinn Arnarson5, Kristinn P. Magnússon1,2,6, and Oddur Vilhelmsson1,6

1University of Akureyri, Faculty of Natural Resource Sciences, Borgir v. Nordurslod, 600 Akureyri, Iceland. 2Icelandic Institute of Natural History, Borgir v. Nordurslod, 600 Akureyri, Iceland 3Íslenskar orkurannsóknir / Iceland GeoSurvey (ISOR), Akureyri Branch, Rangarvollum, 600 Akureyri, Iceland 4Íslenskar orkurannsóknir / Iceland GeoSurvey (ISOR), Department of Geothermal Engineering, Grensasvegi 9, 108 Reykjavik, Iceland 5Orkustofnun / The Icelandic Energy Authority, Grensasvegi 9, 108 Reykjavik, Iceland 6Biomedical Center, University of Iceland, Vatnsmyrarvegur 16, 101 Reykjavik, Iceland

Correspondence: Oddur Vilhelmsson, [email protected]

43 Abstract Natural gas seepage pockmarks present ideal environments for bioprospecting for alkane and aromatic degraders, and investigation of microbial populations with potentially unique adaptations to the presence of hydrocarbons. On-shore seepage pockmarks are found at two disparate sites in the Jökulsá-á- Fjöllum delta in NE Iceland. The origin and composition of headspace gas samples from the pockmarks were analysed by GC-MS and stable isotope analysis, revealing a mixture of thermogenic and biogenic gases with considerable inter-site variability. The warmer, geothermally impacted site displayed a more thermogenic character, comprising mostly methane and CO2 with minor amounts of higher alkanes. The water chemistry of the pockmark sites was determined, revealing considerable heterogeneity between sites. The geothermally impacted site water contained higher amounts of calcium and zinc, and lower amounts of iron than the more biologically impacted site. Microbial communities were analysed by 16S rDNA amplicon sequencing of extracted DNA from the same pockmarks. The bacterial community of the thermogenic gas site was mostly composed of the phyla Proteobacteria, Chloroflexi and Atribacteria, while the bacterial community of the more biologically impacted site was mostly made of Proteobacteria, Bacteriodetes and Chloroflexi.

Introduction Natural gas seepage, the emission of gaseous hydrocarbons from the subsurface, has been studied extensively in the context of petroleum exploration because it can be used as an indicator of petroleum generation in subsurface sediments (1–3). Natural methane gas seepage is the result of subsurface generation or accumulation of methane and the methane concentration in the gas varies according to its source (4). At geothermal and hydrothermal sites, methane is generated by thermogenic processes and seeps up to the surface through cracks and pores, however, the accumulation of methane in deep sea sediments can result in cold seeps or methane hydrates where no direct input of heat is found. This is often linked to biogenic methane which is a product of microbial processes in various anaerobic environments, like bog lakes and sea sediments (5, 6). In many cases the methane generation is of mixed origin, that is both thermogenic and biogenic. For example, methane that is formed during early coalification processes (coal bed methane) is not only of thermogenic origin but also produced by microbes utilizing the

44 lignite (7). In these environments it is expected to find bacteria that participate in methanogenesis and are capable of methane oxidation, respectively. Where natural methane gas seeps up to the surface, pockmarks can develop, that are a habitat for diverse microorganisms (8) and can be regarded as hotspots for anaerobic oxidation of methane (AOM). AOM is often dependent on archaea and sulphate-reducing bacteria, but can in some cases be driven by bacteria through intra-aerobic-denitrification (9) or possibly reductive dehalogenation (10). Microbial communities of hydrocarbon gas seepage environments have been studied around the world, including the Gulf of Mexico (11), Pacific Ocean Margin (12), Cascadia Margin (13) and Barents Sea (14), mainly because of their sulfate-reducing capabilities and AOM. In Öxarfjörður bay, NE Iceland, natural gas seepage pockmarks are found both on the seafloor and on shore. Öxarfjörður is located along the lithospheric boundaries of the North-American and the Eurasian plates and forms a graben bounded by the Tjörnes Fracture Zone in the west and the eastern rim of the North Iceland Volcanic Zone in the east. Geothermal activity in Öxarfjörður bay is confined to three major fissure swarms, cross-sectioning the volcanic zone. The area is prevailed by the river delta of Jökulsá-á-Fjöllum, causing the Öxarfjörður bay to be even more dynamic in nature. Geological settings of the Öxarfjörður area were studied extensively in the 1990s (15–18), leading to the discovery that the methane-rich seepage gas likely originates due to thermal alteration of lignite and coal seams from beneath the 1 km thick sediment (18). Taken together, these studies strongly suggest the presence of sedimentary lignite in the Öxarfjörður graben (19). Very little geomicrobiological work has been conducted in Iceland, with most environmental microbiology work thus far being bioprospective in nature, often paying little attention to community structures or biogeochemical activity. Notable exceptions include the recent attention to basalt glass bioweathering (20–23), as well as investigations into the microbiota of various geothermally impacted environments such as smectite cones (24, 25), subglacial lakes (26, 27), and various kinds of hot springs and geothermal sinters (28–30). Natural gas seeps such as are found in Öxarfjörður, Iceland have thus far not been investigated from a microbiological standpoint despite their unique character which makes them ideal for geomicrobiological studies as both sparsely vegetated geothermal gas seepage pockmarks and colder, more vegetated seepages are found in close proximity to one another. Each methane seep system is thought to be unique in terms of the composition of geological and biological features (8), so taking a snapshot of the microbial community at a methane gas seepage site can provide valuable insight into the dynamics of the system and initiate biological discoveries. In this article, we report the first microbial analysis of the natural gas seepage pockmarks in Öxarfjörður, providing a platform for future

45 geomicrobiological studies in the area as well as displaying the potential of geomicrobiological studies in Iceland.

Materials and methods Sampling and in-field measurements

Figure 7. A map of the study area showing the AEX sampling sites (blue squares) and the SX sites (orange circles). Black diamonds indicate geothermal boreholes. Faults are inferred from the works of Sæmundsson et al. (31) and Ólafsson et al. (18). The insert shows the location of the study area in Iceland and the volcanic rift zone, bounded by the solid lines.

Samples were collected at Skógalón (site SX, 66°09'N, 16°37'W) on August 21st, 2014, and on September 11th, 2015, and at Skógakíll (site AEX, 66°10'N, 16°34'W) on August 13th, 2015 (Figure 7). At site SX, where the natural gas seepage pockmarks are somewhat difficult to distinguish from ordinary marsh gas pockmarks, sites were selected where pockmarks were visibly active and appeared to form straight lines extending NW-SE. Temperature, pH and conductivity were measured in-situ during sampling with hand-held meters. Sediment samples were collected from shallow cores obtained using a corer constructed from a 3-cm diameter galvanized-iron pipe that was hammered into the ground using a sledgehammer, and transferred aseptically to sterile IsoJars (IsoTech laboratories, Champaign, Illinois). Surface soil samples were collected aseptically directly into sterile IsoJars. Water samples were collected

46 aseptically into sterile glass bottles. Gas samples were collected into evacuated double-port glass bottles by means of an inverted nylon funnel connected to silicone rubber tubing. All samples for microbial analysis were immediately put on dry ice where they were kept during transport to laboratory facilities at University of Akureyri where they were either processed immediately or stored in a freezer at -18°C until processing. Samples collected, along with in-situ measurements and types of sample are listed in Table 1.

Table 1. Location and description of sampling sites. Temperature and pH were measured in situ with a hand-held probe (Orion 4-Star, Thermo Scientific).

Site Location Coordinates Description pH T (°C) 66°10'2.250 N Gas seepage AEX-A Skógakíll 6,4 35,2 16°33'59.687 W sediment 66°10'8.014 N Gas seepage AEX-B Skógakíll 6,4 53,3 16°33'56.422 W water 66°10'12.029 N AEX-C Skógakíll Mud-mire ND 30,4 16°33'56.998 W 66°10'9.554 N AEX-D Skógakíll Sand-mud 6,4 19,3 16°33'57.793 W 66°09.222 N Lagoon SX-A Skógalón 8,5 12,1 016°37.089 W sediment 66°09.244 N SX-B Skógalón Puddle water 8,5 13,4 016°37.240 W

Chemical analysis of geothermal fluids Dissolved sulphide in the water samples was determined on-site by titration with mercuric acetate using dithizone in acetone as indicator (Arnórsson et al., 2006). Major components in the water samples were determined at the laboratories of Iceland GeoSurvey (ÍSOR) in Reykjavík: Dissolved inorganic carbon was determined by alkalinity titration (pH 8.2 to 3.8), purging with nitrogen gas and back-titration (pH 3.8 to 8.2) as described by Arnórsson et al. (2006). Silica was analysed by colorimetric determination of a silica- molybdate complex at 410 nm using a Jenway 6300 spectrophotometer. Total dissolved solids were determined by gravimetry. Anions were determined by suppressed ion chromatography on a ThermoScientific ICS-2100 with an AS- 20 column. Major metals were analysed by atomic absorption spectrometry on a Perkin Elmer 1100B spectrometer. The composition of dry gas was also determined at the ÍSOR laboratories by gas chromatography on a Perkin Elmer Arnel 4019 light gas analyser equipped with HayeSep and MolSieve columns and three TCDs.

47 The concentration of trace elements in water samples were determined by ICP methods at the ALS Laboratories, Luleå, Sweden. Stable water isotopes (2H and 18O) were determined by mass spectrometry using a Delta V Advantage IRMS coupled with a Gasbench II at the Institute of Earth Sciences, University of Iceland. Headspace gas analysis from sediment samples was performed at Applied Petroleum Technologies, Kjeller, Norway, using standard techniques. Briefly as follows: Sample preparation and extraction. Sediment samples were washed in water to remove mud before extraction using a Soxtec Tecator instrument. Thimbles were pre-extracted in dichloromethane with 7% (vol/vol) methanol, 10 min boiling and 20 min rinsing. The crushed sample was weighed accurately in the pre-extracted thimbles and boiled for 1 hour and rinsed for 2 hours in 80 cc of dichloromethane with 7% (vol/vol) methanol. Copper blades activated in concentrated hydrochloric acid were added to the extraction cups to cause free sulphur to react with the copper. An aliquot of 10% of the extract was transferred to a pre-weighed bottle and evaporated to dryness. The amount of extractable organic matter (EOM) was calculated from the weight of this 10% aliquot. Deasphaltening. Extracts were evaporated almost to dryness before a small amount of dichloromethane (3 times the amount of EOM) was added. Pentane was added in excess (40 times the volume of EOM/oil and dichloromethane). The solution was stored for at least 12 hours in a dark place before the solution was filtered or centrifuged and the weight of the asphaltenes measured. GC analysis of gas components. Aliquots of the samples were transferred to exetainers. 0.1-1ml were sampled using a Gerstel MPS2 autosampler and injected into a Agilent 7890 RGA GC equipped with Molsieve and Poraplot Q columns, a flame ionisation detector (FID) and 2 thermal conductivity detector (TCD). Hydrocarbons were measured by FID. H2, CO2, N2, and O2/Ar by TCD. Carbon isotope analysis of hydrocarbon compounds and CO2. The carbon isotopic composition of the hydrocarbon gas components was determined by a GC-C-IRMS system. Aliquots were sampled with a syringe and analysed on a Trace GC2000, equipped with a Poraplot Q column, connected to a Delta plus XP IRMS. The components were burnt to CO2 and water in a 1000 °C furnace over Cu/Ni/Pt. The water was removed by Nafion membrane separation. Repeated analyses of standards indicate that the reproducibility of δ13C values is better than 1 ‰ PDB (2 sigma). Carbon isotope analysis of low concentration methane using the Precon. The carbon isotopic composition of methane was determined by a Precon-IRMS system. Aliquots were sampled with a GCPal autosampler. CO2, CO and water were removed on chemical traps. Other hydrocarbons than CH4 and remaining traces of CO2 were removed by cryotrapping. The methane was burnt to CO2 and water in a 1000 °C furnace over Cu/Ni/Pt. The water was

48 removed by Nafion membrane separation. The sample preparation system described (Precon) was connected to a Delta plus XP IRMS for δ 13C analysis. Repeated analyses of standards indicate that the reproducibility of δ13C values is better than 1 ‰ PDB (2 sigma). GC of EOM fraction. A HP7890 A instrument was used. The column was a CP-Sil-5 CB-MS, length 30 m, i.d. 0.25 mm, film thickness 0.25 m. C20D42 is used as an internal standard. Temperature programme: 50°C (1 min), -4 °C/min, -320 °C (25 min).

Metataxonomic community analysis Total DNA was extracted from sediment samples in duplicates, using the PowerSoil kit (MoBio laboratories) following the manufacturer’s protocol. The DNA isolated was measured with Qubit fluorometer (Invitrogen, Carlsbad, CA) to confirm dsDNA in the samples, and a PCR carried out with 8F/1522R primers (32) to determine that the bacterial 16S rDNA could be amplified. Paired-end library of the 16S rDNA hypervariable region V3/V4, was sequenced on Illumina MiSeq platform by Macrogen, Netherlands, from 8 samples in total, four from AEX-A and four from SX-A. The data was processed and analysed using CLC Genomics Workbench 10.1.1 (https://www.qiagen bioinformatics.com/) and the CLC Microbial Genomics Module 2.5.1, with default parameters. Operational taxonomic units (OTU) were clustered by reference based OTU clustering and tree alignment was performed by using the GreenGenes v.15.5 database for 97% similarity. For statistical analysis only alpha-diversity of samples was performed since the sequencing data only contained technical replicates, which does not allow analyses of beta-diversity. Differential abundance analysis (Likelihood Ratio test) was performed to see statistically significant differences in taxa between sampling sites.

Initial culturing and isolation of bacteria Samples were serially diluted to 10-6 in sterile Butterfield's buffer (250 mM KH2PO4, pH 7.2) and all dilutions plated in duplicate onto Reasoner's agar 2A (Difco) and several selective and differential media including medium 9K for iron oxidizers (33), Mn medium for manganese oxidizers (990 mL basal agar B [0.42 g NaOAc, 0.1 g peptone, 0.1 g yeast extract, 15 agar, 990 mL sample water, autoclaved and cooled to 50°C], 10 mL pre-warmed filter-sterilized 1 M HEPES at pH 7.5, 100 µL filter-sterilized 100 mM Mn(II)SO4), Gui medium for laccase producers (990 mL basal agar B, 0,01% guiaicol), Hex medium for hexane degraders (990 mL basal agar B, 1.3 mL filter-sterilized 99 parts hexane/1 part dishwashing detergent), Naph medium for naphthalene degraders (basal agar B with several crystals of naphthalene added to the lid of inverted plates and then sprayed with fast blue for degradation indication), and 2,4-D medium for dichlorophenoxyacetate degraders (basal agar B

49 supplemented with 2 mM 2,4-D). Three atmospheric incubation conditions were used: an unmodified atmosphere in sealed plate bags, a propane-enriched aerobic atmosphere in sealed plate bags flushed daily with propane, and an anaerobic, propane-supplemented atmosphere in anaerobic jars scrubbed of oxygen with a palladium catalyst (BBL GasPak) and monitored for anaerobicity with a resazurin strip. The jars were injected with 100 mL propane through a septum. Plates were incubated in the dark at 5, 15, or 22°C until no new colonies appeared (up to 4 weeks). Colonies were isolated based on isolation medium and colony morphology and streaked onto fresh media and recultured up to three times or until considered pure. Stocks of purified isolates were prepared by suspending a loopful of growth in 1.0 mL 28% (v/v) glycerol and are stored at -70°C in the University of Akureyri culture collection.

16S rRNA gene-based identification of cultured strains For each strain, a minute colony mass or 1 µl of freezer stock was suspended in 25 µl of lysis buffer (1% Triton x-100, 20 mM Tris, 2 mM EDTA, pH 8,0) and incubated for 10 minutes at 95°C in the thermocyler (MJR PTC-200 thermocycler, MJ Research Inc. Massachusetts, USA). The lysis buffer solution (1 µl), or 1 µl of extracted DNA (using UltraClean® Microbial DNA Isolation Kit (MoBio Laboratories, Carlsbad, California, USA)), was used as a DNA template for Polymerase chain reaction (PCR) using Taq DNA- polymerase to amplify the DNA using the ‘universal‘ bacterial primers 8F (5'- AGTTTGATCCTGGCTCAG'3) and 1522R: (5'-AAGGAGGTGATCCAGC CGCA-'3). The PCR reaction was conducted as follows: 95°C for 3 min, followed by 35 cycles of 95°C for 30 sec, 50°C for 30 sec and 68°C for 90 sec, then final extension at 68°C for 7 min. The PCR products were loaded on 0,8% agarose gel with TBE buffer and run at 100 V for approximately 30 minutes to verify the presence of approximately 1500-bp amplicons. The PCR products were purified for sequencing using 22 µl of sample in 10 µl ExoSap Mix (mix for 50 reactions; 1.25 µl Exonuclease I [20 U/µl], 2.5 µl Antarctic phosphatase [5 U/µl] (New England BioLabs Inc.), 496.5 µl distilled H2O). The products were incubated at 37°C for 30 minutes and heated to 95°C for 5 minutes to inactivate the enzymes in the ExoSap. The purified PCR products were sequenced with BigDye terminator kit on Applied Biosystem 3130XL DNA analyser (Applied Biosystems, Foster City, California, USA) at Macrogen Europe, Amsterdam, the Netherlands. Two sequencing reactions were run for each strain, using the primers 519F (5'- CAGCAGCCGCGGTAATAC-'3) and 926R (5'-CCGTCAATTCCTTTGAG TTT-'3). The resulting sequences were trimmed using ABI Sequence Scanner (Applied Biosystems), the forward sequence and the reverse complement of

50 the reverse sequence aligned and combined, and taxonomic identities obtained using the EzTaxon server (34). Results Water chemical analysis revealed several differences in major components at the two sites (Table 2). Although the pH of the water did not differ significantly as judged by a two-tailed Student’s t-test, electrical conductivity was nearly 12-fold higher at the AEX site, significant at the 99.5% confidence level. The AEX site water contained more than 420-times as much silica as did the SX site water, and several other ions, such as sodium, potassium chloride and bromide were also found to be present at significantly higher levels at the AEX site, underscoring the more geothermal character of the environment (Table 2).

Table 2. Major physicochemical characteristics1 of water from the two study sites. AEX SX pH at 21°C 7.57 ± 0.31 8.64 ± 0.73 (p = 0.19) Conductivity at 25°C 8945 ± 728 773 ± 392 (p = 0.005) (µScm-1)

DIC (as ppm CO2) 44.8 ± 31.0 94.5 ± 43.1 (p = 0.32)

SiO2 (ppm) 147.5 ± 2.1 0.35 ± 0.07 (p = 0.0001) Na (ppm) 1900 ± 56.6 107.4 ± 57.5 (p = 0.001) K (ppm) 104.9 ± 18.6 4.01 ± 1.96 (p = 0.02) Mg (ppm) 24.0 ± 15.9 17.8 ± 10.5 (p = 0.69) Ca (ppm) 311.5 ± 111.0 17.1 ± 9.0 (p = 0.06) F (ppm) 0.45 ± 0.13 0.21 ± 0.01 (p = 0.11) Cl (ppm) 3355 ± 276 169.4 ± 105.5 (p = 0.004) Br (ppm) 12.2 ± 0.8 0.34 ± 0.31 (p = 0.002)

SO4 (ppm) 215 ± 76.4 5.13 ± 0.60 (p = 0.06) 1Dissolved sulphide was determined on-site by titration with mercuric acetate using dithizone as indicator (Arnórsson et al., 2006). Dissolved inorganic carbon was determined by alkalinity titration and back-titration (Arnórsson et al., 2006). Silica was analysed by colorimetric determination of a silica-molybdate complex at 410 nm. Total dissolved solids were determined by gravimetry. Anions were determined by suppressed ion chromatography. Major metals were analysed by atomic absorption spectrometry.

Headspace gas analysis revealed similar amounts of hydrocarbon gas at the two sites (Table 3). Although methane content was consistently (n=3) found to be lower at the AEX site than at the SX site, the difference was not deemed significant by a two-tailed Student’s t-test (not shown). Isotope composition suggests a thermogenic origin of the AEX-site headspace gas,

51 whereas a biogenic origin is suggested for the SX-site headspace gas ((Figure 8). Table 3). Thermogenic origin of the AEX gas was further supported by a high methane/ethane ratio (17.8). Both the composition of the headspace gas and the methane isotope composition were similar to those reported by Ólafsson et al. for borehole gasses in the Skógalón area (18). EOM fractions showed a difference in hydrocarbon content in terms of lower-chain hydrocarbons in AEX and higher amounts of higher-chain hydrocarbons at the SX site (Figure 8).

Table 3. Headspace gas analysis1 on sediment samples from seepage pockmarks at the two study sites.

AEX SX

N2 [%total] 39.2 (±31.5) 81.2 (±2.62)

O2 + Ar [%total] 8.46 (±8.53) 14.10 (±5.66) ppm THCG 5.09×105 (±3.88×105) 0.389×105 (±0.296×105)

CO2 [%THCG] 99.9 (±0.23) 93.2 (±1.77) -3 -3 Methane (CH4) 46.1×10 (±48.3×10 ) 6.78 (±1.66) [%THCG] -3 -3 Ethane (C2H6) 2.60×10 (±4.50×10 ) 0 (±0) [%THCG] -3 -3 Propane (C3H8) 1.03×10 (±1.79×10 ) 0 (±0) [%THCG] -3 -3 -3 -3 iso-Pentane (i-C5H12) 18.7×10 (±32.3×10 ) 8.00×10 (±11.3×10 ) [%THCG] -3 -3 -3 -3 Pentane (n-C5H12) 73.7×10 (±127×10 ) 49.5×10 (±70.0×10 ) [%THCG] -3 -3 -3 -3 Benzene (C6H6) 3.80×10 (±6.24×10 ) 7.15×10 (±5.45×10 ) [%THCG] Methane δ¹³C [‰] -26.6 -63.2 CO₂ δ¹³C [‰] -3.3 nd 1Samples were extracted in dichloromethane/methanol, deasphaltened in pentane, and analysed by gas chromatography.

Figure 8. Hydrocarbon content in the natural gas seepage pockmarks. Extractable organic matter (EOM) concentration as determined by gas chromatography is compared between the AEX (dark columns) and SX (light columns) study sites. Error bars are omitted for clarity.

53 Metataxanomic analysis. Four technical replicates from each study site were used to compare the microbial communities in the gas seepage pockmarks of Skógalón (SX) and Skógakíll (AEX). The DNA extraction yielded on average 2,8 g/ml ( 0,3 g/ml) and the amplicon library generated on average 42,8 ng/L ( 1,3 ng/L) of amplicons with the length of 5875 bp. Over 4 million paired sequences, were analysed and trimmed to the average of 523.241 reads per sample with the length of 301 bp. A total of 595.137 reads generated the OTU table after filtering out chimeric sequences, where predicted OTUs were in total 26.786 OTUs (Table 4). Alpha diversity metrics were measured at the depth of 60,000 sequences per sample. The number of OTUs had reached plateau at 25,000 sequences, meaning the dataset was fully sufficient to estimate the diversity of the bacterial communities in the natural seepage pockmarks. The species richness as estimated with Chao1 index and Shannon´s diversity index are shown in Table 4.

Table 4. Number of predicted operational taxanomic units (OTUs) and alpha diversity metrics as calculated at 25,000 sequences from the two study sites.

Alpha diversity metrics Paired Reads in Predict seqs OTUs OTUs OTUs Shannon Chao1 AEX 2.064.234 307.384 14.019 3.461 ± 623 8,3 ± 0,5 3.980 ± 751 SX 2.121.996 287.753 12.767 3.762 ± 333 9,1 ± 0,2 4.203 ± 504

Taxonomic composition The focus was set on analysing the most abundant taxa, so after filtering out chloroplast OTUs, the OTUs with the lowest combined abundance (<=1000) were omitted. A total of 14 bacterial phyla was observed as the most abundant at AEX and SX sites, divided into 23 classes and 45 observed genera (Figure 9). Proteobacteria was the most abundant phylum at both AEX and SX sites, 28% and 30%, respectively. At the AEX site, Proteobacteria was followed by Chloroflexi (22%) and Aminicenantes (10%) at phylum level. At the SX site, the Bacteroidetes had significantly higher abundance than in the AEX site, with relative abundance of 24%, followed by Chloroflexi (13%). The bacterial community structure differed between sites, but only one class, within the phylum of Aminicenantes was found by likelihood ratio analysis to be significantly more abundant at AEX than at the SX site. An order within Bacteriodetes was found to be significantly more abundant at the SX site compared to the AEX. The family of Syntrophaceae under the class of Deltaproteobacteria was more abundant at the SX site with 2,6-fold higher

54 abundance. Three genera within the Clostridia class had over 3,0-fold higher abundance at AEX than SX site.

Figure 9. Bacterial community structure in natural gas seepage pockmarks at Skógakíll (AEX) and Skógalón (SX) sites, presented as the relative abundance of bacterial phyla from amplicon sequencing of V3-V4 in 16S rDNA. Operational taxanomic units (OTUs) with relative abundance lower than 0.1 were omitted.

Cultured microbial load. Plate counts after 7 days at 22°C (Table 5) of samples from site SX indicated the presence of substantial communities of naphthalene and hexane degraders, particularly under aerobic conditions. One hundred and eighty-six colonies were restreaked for isolation in pure culture (Table 9 in Supplements).

Isolates. Putatively facultative chemoautotrophs were surprisingly numerous judging by growth on Mn media, but the extremely restrictive medium 9K only yielded a few colonies, all from sample OX06. Spraying colonies with fast blue confirmed the presence of alpha-naphthol in some of the colonies on Naph-agar, but not all. Strains OX0102 and OX0103 tested positive for naphthalene degradation by fast blue; strains OX0304 and OX0306 tested positive for 2,4-diphenoxyacetate degradation by fast blue. One hundred and six strains have been identified by partial 16S rDNA sequencing using the Sanger method and found to comprise 38 genera in 8 classes (Table 6, Table 10 in Supplements)

55 Table 5. Colony-forming units per gram sample after 7 days at 22°C on selective media.

OX01 OX01 OX03 OX03 OX06 OX06 Aerobic Anaerobic Aerobic Anaerobic Aerobic Anaerobic R2A 1.4×105 2.8×104 5.0×105 2.4×105 1.7×107 5.0×106 Naph 2.0×103 <1×103 <1×103 <1×103 2.5×105 1.0×105 Hex 7.5×104 <1×103 2.0×105 <1×103 >2.5×105 2.0×105 2,4-D nd nd <1×103 <1×103 nd nd Mn 9.3×104 5.0×103 2.0×105 <1×103 >2.5×105 7.5×104 9K <1×103 <1×103 <1×103 <1×103 1.0×103 3.0×103 Gui nd nd nd nd 1.0×106 3.0×106

56 Table 6. Bacterial isolates and their taxonomic classification by partial 16S rRNA gene sequencing.

GenBank accession EzTaxon classification Strains numbers Class Order Family Genus SX0206, SX1205 MG575948, MG575949 Actinobacteria Corynebacteriales Nocardiaceae Rhodococcus OX0615 MG575950 ´´ Micrococcales Cellulomonadaceae Oerskovia OX0315, OX0319 MG575951, MG575952 ´´ ´´ Microbacteriaceae Cryobacterium OX0117, OX0308, OX0313 MG575953-MG575955 ´´ ´´ Micrococcaceae Arthrobacter OX0107 MG575956 ´´ ´´ ´´ Pseudarthrobacter OX0614 MG575957 ´´ ´´ Sanguibacteraceae Sanguibacter OX0316 MG575958 ´´ Streptomycetales Streptomycetaceae Streptomyces OX0104, OX0118 MG575959, MG575960 Cytophagia Cytophagales Cyclobacteriaceae Algoriphagus OX0108, OX0126, OX0312 MG575961-MG575963 ´´ ´´ ´´ Aquiflexum OX0623, OX0125, OX0129, OX0311, OX0122, OX0123, OX0314, OX0127 MG575964-MG575971 Flavobacteriia Flavobacteriales Flavobacteriaceae Flavobacterium OX0625 MG575972 Sphingobacteria Sphingobacteriales Sphingobacteriaceae Pedobacter OX1213, OX2513, OX1505, OX2509, OX1011, OX1604, OX1805, OX1004, OX1006, OX1007, OX1210, OX1212, OX2308, OX2309, OX2506 MG575973 - MG575987 Bacilli Bacillales Bacillaceae Bacillus

57 OX0317 MG575988 ´´ ´´ ´´ Psychrobacillus OX2205, OX0307, OX2310, OX0301, OX0302 MG575989 - MG575993 ´´ ´´ Paenibacillaceae Paenibacillus OX0310 MG575994 ´´ ´´ Planococcaceae Jeotgalibacillus OX0626 MG575995 ´´ ´´ Staphylococcaceae Staphylococcus OX1107, OX1109, OX2515, OX2106, OX2313 MG575996-MG576000 ´´ Lactobacillales incertae sedis Exiguobacterium OX0102, OX2008, OX0309 MG576001-MG576003 Alphaproteobacteria Caulobacterales Caulobacteraceae Brevundimonas OX0106, OX1214, OX0119, SX1214 MG576004-MG576007 ´´ Rhizobiales Rhizobiaceae Rhizobium OX0632 MG576008 ´´ Rhodobacterales Rhodobacteraceae Cereibacter OX1314 MG576009 ´´ ´´ ´´ Paracoccus SX0604 MG576010 ´´ Sphingomonadales Erythrobacteraceae Porphyrobacter OX0620 MG576011 ´´ ´´ Sphingomonadaceae Sphingobium OX1313, OX1403, OX1702, OX1216 MG576012-MG576015 Betaproteobacteria Burkholderiales Burkholderiaceae Paraburkholderia OX0105, OX0120, OX0124 MG576016-MG576018 ´´ ´´ Comamonadaceae Acidovorax OX0630, OX0321 MG576019, MG576020 ´´ ´´ ´´ Rhodoferax OX0130 MG576021 ´´ ´´ ´´ Variovorax OX0611 MG576022 ´´ ´´ incertae sedis Paucibacter OX0627 MG576023 ´´ Rhodocyclales Rhodocyclaceae Dechloromonas

58 OX0617, OX0622, SX0303, OX0110, OX0112, OX0612 MG576024-MG576029 Gammaproteobacteria Aeromonadales Aeromonadaceae Aeromonas OX0619 MG576030 ´´ Alteromonadales Shewanellaceae Shewanella OX1909, OX1208 MG576031-MG576032 ´´ Chromatiales Chromatiaceae Rheinheimera OX1012 MG576033 ´´ Enterobacteriales Enterobacteriaceae Escherichia OX0103, OX0101 MG576034-MG576035 ´´ ´´ ´´ Rahnella OX0604, OX0606, OX0607 MG576036-MG576038 ´´ ´´ ´´ Shigella OX0601, OX0621 MG576039-MG576040 ´´ Pseudomonadales Moraxellaceae Acinetobacter OX1807, OX1808, OX0306, OX0322, OX1110, OX0613, SX0305, SX0307, SX1216, SX0304, OX0631, SX1213, OX0304 MG576041-MG576053 ´´ ´´ Pseudomonadaceae Pseudomonas

59 Discussion The study sites, AEX and SX, were found to be distinct in terms of geochemistry. The AEX site contained higher concentrations of silica, very similar to the concentrations of previous studies on geothermal activity in Öxarfjörður (18) indicating geothermal water coming from the pockmarks. The sodium chloride concentration was higher than in previous studies which implies a mixture of seawater with the geothermal water, however, the pockmarks at AEX are located in a river delta and water samples were taken at the pockmark surface so the intermixture of seawater is not surprising. The water chemistry at SX site shows low concentration of silica indicating no geothermal activity and less intermixture of seawater than at AEX. The stable isotope ratio δ¹³C of methane also indicates a biogenic origin of the methane at SX, while at AEX the δ¹³C suggests a mixture of thermogenic and biogenic origin of methane, which can most likely be linked to microbial lignite utilization at the site as well as the geothermal activity previously described (18). The hydrocarbon content also shows more complex and higher chain hydrocarbons at SX, that can be related to more vegetation and organic matter accumulation, in contrast with lower-chain hydrocarbons at the AEX site with less vegetation (Figure 8). These geochemical factors underline how disparate the two sites are: the AEX pockmarks containing geothermal groundwater with thermogenic methane generation and the SX pockmarks the result of biogenic natural gas accumulation. The location of SX site and the lining up of the pockmarks can easily suggest thermogenic methane seepage at the site, but the pockmarks explored in this study were identified as marsh gas seepage. The seepage pockmarks were found to harbour diverse microbiotas consisting largely of anaerobic heterotrophs. Given the lack of visible vegetation at the AEX site, available organic matter seems likely to be to a large extent restricted to the gas seep itself. This kind of environment is thus likely to contain a microbial community composed largely of facultative chemolithotrophs and oxidizers of methane, lighter alkanes, and aromatics. These organisms can be valuable for bioremediation of petroleum contamination in basaltic oligotrophic environments such as Icelandic beach environments.The inter-site diversity of both sampling sites was notable, however, several groups of bacteria were shown to vary in relative abundance between the two sampling sites:

Chloroflexi and methyl halide metabolism The high relative abundance of Dehalococcoidia in the microbial consortia at the study sites, particularly site AEX, is noteworthy and underscores the profound effect that petrochemical seepage has on the composition of the local microbiota. The class Dehalococcoidia contains at the present time only one validly described order (Dehalococcoidales), one family

60 (Dehalococcoidaceae), and three genera (Dehalococcoides, Dehalobium and Dehalogenimonas), comprising a total of four species, all of which are capable of anaerobic reductive dehalogenation (35–39).

Table 7. Fractional abundance of Chloroflexi classes and orders in the seepage pockmarks microbiomes as determined with amplicon sequencing. Class AEX1 SX1 Order AEX2 SX2 Anaerolineae 0.42 0.63 Anaerolineales 0.31 0.22 ‘envOPS12’ 0.21 0.43 ‘GCA004’ 0.09 0.13 ‘SHA-20’ 0.20 0.04 Others 0.19 0.19 Dehalococcoidia 0.53 0.27 Dehalococcoidales 0.18 0.48 ‘GIF9’ 0.76 0.32 ‘FS117-23B-02’ 0.02 0.06 Others 0.04 0.14 ‘Ellin6529’ 0.03 0.06 ‘S085’ 0.02 0.02 Others 0.00 0.01 1Fraction of Chloroflexi paired-end sequences (total of 86,729 paired-end reads). Figures in bold indicate a significant difference between the AEX and SX microbiota by a two-tailed two-sample Student‘s t-test (p<0.05). 2Fraction of class-level paired-end sequences (total of 44,384 paired-end Anaerolineae reads and 36,549 paired-end Dehalococcoidia reads). Figures in bold indicate a significant difference between the AEX and SX microbiota by a two-tailed two-sample Student‘s t-test (p<0.05).

A large fraction of the dehalococcoidal OTUs in this study were found to be closely related to the described, dehalorespiring members of this class, with 18% of dehalococcoidal paired-end reads from the AEX site and 48% from the SX site being assigned to the order Dehalococcoidales (Table 7), the majority (9120 of 9579) being further assignable to the family Dehalococcoidaceae. In further support of dehalorespiration being an important process in the seepage pockmark microbiotas, genera known to contain facultative dehalorespirers, like the betaproteobacterial genus Dechloromonas (40) the deltaproteobacterial genus Anaeromyxobacter (41), were statistically more abundant at the AEX site. Furthermore, cultured bacteria from the Öxfjörður seeps, while not including Dehalococcoidia, do include isolates assigned to genera known to include aerobic facultative dechlorinators, such as Dechloromonas and Shewanella (Table 7). Dechloromonas isolates are

61 capable of anaerobic oxidation of benzene (42) and could possibly be used for bioremediation. The as-yet unnamed and uncharacterized order GIF9, suggested by several environmental studies and highly abundant in the AEX and SX microbiomes (Table 7), may consist of bacteria that posess other metabolic processes than just organohalide respiration. Thus, a recent metagenomic study indicated that some members of this group may be homoacetogenic fermenters that possess a complete Wood-Ljungdahl CO2 reduction pathway (43). It should thus be stressed that the presence of a large contingent of Dehalococcoidia, as was found to be the case in the present study, need not necessarily be indicative of dehalorespiration constituting a major metabolic activity in the environment under study. Indeed, considerable variation in metabolic characteristics occurs in most well-characterized bacterial classes and hence it must be considered likely that other, perhaps non-dehalorespiring taxa remain to be characterized within this class. Nevertheless, the high abundance of Dehalococcoidaceae, as discussed above, does strongly indicate organohalide respiration as an important process in the seepage pockmarks. Recently, it was suggested, in part because of the notable abundance of Dehalococcoides, that in certain Antarctic lakebeds, anaerobic methane oxidation may be fuelled by reductive dehalogenation (10). The results of the present study are suggestive of the presence of such an ecosystem in the methane seeps in the Öxarfjörður graben. Methyl halides are often associated with coal combustion (44), further suggesting subsurface interaction of geothermal matter with lignite as a source of chloromethane. Another highly abundant class within the Chloroflexi was the Anaerolineae (Table 7), a class originally described as consisting of strictly anaerobic chemo-organotrophs (45), and frequently detected in subsurface environments (46–49). However, due to the scarcity of cultured representatives, the metabolic capabilities of this class have remained elusive. A recent study of seven single-cell genomes from deep submarine hydrothermal vent sediments indicated the presence of a Wood-Ljungdahl CO2 reduction pathway, as well as a number of ABC transporters, and in one case a putative reductive dehalogenase (50). In the present study, the Anaerolineae appear fairly diverse, with 81% of Chloroflexi paired-end reads being assigned to four orders: the Anaerolineales and three putative orders without cultured representation, envOPS12, GCA004, and SHA-20 (Table 7).

Proteobacteria and the sulfur cycle The Proteobacteria phylum mainly consisted of Deltaproteobacteria and Alphaproteobacteria (Table 8). The alphaproteobacterial fraction was fairly homogeneous, consisting mostly of reads assigned to the order Rhizobacteriales, of which 77% could be assigned to a single genus, Bosea, a genus of chemolithoheterotrophs noted for their ability to oxidize inorganic

62 sulfur compounds (51). The deltaproteobacterial fraction was found to be more diverse although most of the OTUs could be assigned to either of two orders, the Syntrophobacterales and the Desulfobacterales (Table 8), both of which contain mostly, albeit not exclusively, sulfate-reducing organisms. Table 8. Fractional abundance of Proteobacteria classes and orders in the seepage pockmarks microbiomes as determined with amplicon sequencing. Class AEX1 SX1 Order AEX2 SX2 Delta- 0.43 0.23 proteobacteria Syntrophobacterales 0.27 0.55 Desulfobacterales 0.26 0.07 Myxococcales 0.07 0.08 Desulfarculales 0.06 0.06 Desulfuromonadales 0.06 0.04 ‘BCP076’ 0.14 0.06 Others 0.15 0.12 Alpha- 0.48 0.63 proteobacteria Rhizobiales 0.88 0.92 Rhodobacterales 0.06 0.02 Others 0.06 0.05 Gamma- 0.11 0.06 proteobacteria Beta- 0.06 0.07 proteobacteria Others 0.02 0.01 1Fraction of Proteobacteria paired-end sequences (total of 152,153 paired- end reads). Figures in bold indicate a significant difference between the AEX and SX microbiota by a two-tailed two-sample Student‘s t-test (p<0.05). 2Fraction of class-level paired-end sequences (total of 83,764 paired-end Deltaproteobacteria reads and 43,823 paired-end Alphaproteobacteria reads). Figures in bold indicate a significant difference between the AEX and SX microbiota by a two-tailed two-sample Student‘s t-test (p<0.05).

The Syntrophobacterales, known to be frequently associated with anoxic aquatic environments (52), are significantly enriched in the SX marshland site as compared to the AEX site, perhaps reflecting an influx of marshland- associated bacteria into the seepage pockmark environment. Most of the Syntrophomonadales reads (77%) can be assigned to the family Syntrophaceae, which contains both sulfate-reducing and non-sulfate-reducing bacteria (53). Many of the Syntrophaceae reads (59%) could not be confidently assigned to genera, rendering the question of the importance of sulfate reduction of this taxon in the seepage pockmarks unresolved. However, taken together with the high abundance of the sulfate-reducing order

63 Desulfobacterales, sulfate reduction is likely to be a major process in the seepage pockmarks, likely supporting AOM consortia. Families within Desulfobacterales have been reported to actively oxidize short and long chain alkanes and are suggested to be the key alkane degraders in marine seeps (54). Furthermore, considering the high abundance of the sulfur-oxidizing Bosea, we can surmise that Proteobacteria consitute an important driver of sulfur cycling within the seepage pockmark microbiota.

Is anaerobic oxidation of methane carried out by Atribacteria in the seepage pockmarks? Atribacteria (group ‘OP9’) are often found to be predominant in methane- rich anaerobic environments such as marine sediments and subseafloor “mud volcanoes” (55, 56). Although they have not been directly linked to AOM in these environments, they have been suggested to mediate AOM in some cold seep environments (10). In general, the Atribacteria are thought to play heterotrophic roles, likely fermentative (55, 57), but single-cell genomics studies on representative Atribacteria suggests that these organisms may be indirectly responsible for methane production through the production of acetate or CO2 (55, 58).

Concluding remarks This study reveals natural gas seeps of biogenic origin in Öxarfjörður in addition to known geothermal gas seepage pockmarks in the Jökulsá-á-Fjöllum river delta. The microbial communities associated with the pockmarks show higher biodiversity at biogenic gas seepage than in thermogenic gas seepage pockmarks. The abundant taxa in the pockmarks indicate that the microbial community is most likely involved in hydrocarbon degradation linked to sulfur cycling and AOM, and the abundance of Dehalococcoidia suggests the presence of anaerobic reductive dehalogenation in natural gas seepage pockmarks of thermogenic origin. Further studies are needed to demonstrate the connection between the gas origin and the pockmark microbiota, establishing the need for further geomicrobiological research in Icelandic natural gas seeps.

Acknowledgments This work was funded by Orkustofnun and Orkurannsoknarsjodur Landsvirkjunar. Thanks to Geir Hansen & Co. at Applied Petroleum technology.

64 References 1. Link WK. 1952. Significance of oil and gas seeps in world oil exploration. Am Assoc Pet Geol Bull 36:1505–1540. 2. Jones, V. T., & Drozd RJ. 1983. Predictions of oil or gas potential by near-surface geochemistry. Am Assoc Pet Geol Bull 67:932–952. 3. Abrams MA, Dahdah NF. 2010. Surface sediment gases as indicators of subsurface hydrocarbons - examining the record in laboratory and field studies. Mar Pet Geol 27:273–284. 4. Etiope G, Feyzullayev A, Baciu CL. 2009. Terrestrial methane seeps and mud volcanoes: A global perspective of gas origin. Mar Pet Geol 26:333–344. 5. Zengler K, Richnow HH, Rosselló-Mora R, Michaelis W, Widdel F. 1999. Methane formation from long-chain alkanes by anaerobic microorganisms. Nature 401:266–269. 6. Martini AM, Budai JM, Walter LM, Schoell M. 1996. Microbial generation of economic accumulations of methane within a shallow organic-rich shale. Nature 383:155–158. 7. Ritter D, Vinson D, Barnhart E, Akob DM, Fields MW, Cunningham AB, Orem W, McIntosh JC. 2015. Enhanced microbial coalbed methane generation: A review of research, commercial activity, and remaining challenges. Int J Coal Geol 146:28–41. 8. Zhang CL, Lanoil B. 2004. Geomicrobiology and biogeochemistry of gas hydrates and cold seeps. Chem Geol 205:187–194. 9. Ettwig KF, Butler MK, Le Paslier D, Pelletier E, Mangenot S, Kuypers MMM, Schreiber F, Dutilh BE, Zedelius J, de Beer D, Gloerich J, Wessels HJCT, van Alen T, Luesken F, Wu ML, van de Pas-Schoonen KT, Op den Camp HJM, Janssen-Megens EM, Francoijs K-J, Stunnenberg H, Weissenbach J, Jetten MSM, Strous M. 2010. Nitrite- driven anaerobic methane oxidation by oxygenic bacteria. Nature 464:543–548. 10. Saxton MA, Samarkin VA, Schutte CA, Bowles MW, Madigan MT, Cadieux SB, Pratt LM, Joye SB. 2016. Biogeochemical and 16S rRNA gene sequence evidence supports a novel mode of anaerobic methanotrophy in permanently ice-covered Lake Fryxell, Antarctica. Limnol Oceanogr 61:S119–S130. 11. Mills HJ, Martinez RJ, Story S, Sobecky PA, Sobecky A. 2005. Characterization of Microbial Community Structure in Gulf of Mexico Gas Hydrates : Comparative Analysis of DNA- and RNA-Derived Clone Libraries Characterization of Microbial Community Structure in Gulf of Mexico Gas Hydrates : Comparative Analysis of DNA-. Appl Environ Microbiol 71:3235–3247. 12. Inagaki F, Nunoura T, Nakagawa S, Teske A, Lever M, Lauer A,

65 Suzuki M, Takai K, Delwiche M, Colwell FS, Nealson KH, Horikoshi K, D’Hondt S, Jørgensen BB. 2006. Biogeographical distribution and diversity of microbes in methane hydrate-bearing deep marine sediments on the Pacific Ocean Margin. Proc Natl Acad Sci U S A 103:2815–20. 13. Knittel K, Boetius A, Lemke A, Eilers H, Lochte K, Pfannkuche O, Linke P, Amann R. 2003. Activity, Distribution, and Diversity of Sulfate Reducers and Other Bacteria in Sediments above Gas Hydrate (Cascadia Margin, Oregon). Geomicrobiol J 20:269–294. 14. Niemann H, Lösekann T, de Beer D, Elvert M, Nadalig T, Knittel K, Amann R, Sauter EJ, Schlüter M, Klages M, Foucher JP, Boetius A. 2006. Novel microbial communities of the Haakon Mosby mud volcano and their role as a methane sink. Nature 443:854–858. 15. Flóvenz ÓG, Gunnarsson K. 1991. Seismic crustal structure in Iceland and surrounding area. Tectonophysics 189:1–17. 16. Gunnarsson K. 1998. Sedimentary basins of the N-Iceland shelf. Draft Version for Discussion (April–May 1998). Report OS-98014. Reykjavík. 17. Friðleifsson GÓ. 1994. Geothermal gradient and hydrothermal systems off North Iceland. Statement GÓF-94-05. Reykjavík. 18. Ólafsson M, Friðleifsson GÓ, Eiríksson J, Sigvaldason H, Ármansson H. 1993. On the origin of organic gas in Öxarfjörður, NE-Iceland. Report No. OS-93015/JHD-05. Reykjavík. 19. Richter B, Gunnarsson K. 2010. Overview of hydrocarbon related research in Tjörnes. GagnavefsjaIs. 20. Kelly LC, Cockell CS, Herrera-Belaroussi A, Piceno Y, Andersen G, DeSantis T, Brodie E, Thorsteinsson T, Marteinsson VV, Poly F, LeRoux X. 2011. Bacterial Diversity of Terrestrial Crystalline Volcanic Rocks, Iceland. Microb Ecol 62:69–79. 21. Cockell CS, Olsson K, Knowles F, Kelly L, Herrera A, Thorsteinsson T, Marteinsson V. 2009. Bacteria in Weathered Basaltic Glass, Iceland. Geomicrobiol J 26:491–507. 22. Cockell CS, Kelly LC, Marteinsson V. 2013. Actinobacteria—an ancient phylum active in volcanic rock weathering. Geomicrobiol J 30:706–720. 23. Kelly LC, Cockell CS, Thorsteinsson T, Marteinsson V, Stevenson J. 2014. Pioneer Microbial Communities of the Fimmvörðuháls Lava Flow, Eyjafjallajökull, Iceland. Microb Ecol 68:504–518. 24. Marteinsson V, Kristjánsson JK, Kristmannsdóttir H, Dahlquist M, Sæmundsson K, Hannington M, Pétursdóttir SK, Geptner A, Stoffers P. 2001. Discovery and description of giant submarine smectite cones on the seafloor in Eyjafjordur, Northern Iceland, and a novel thermal microbial habitat. Appl Environ Microbiol 67:827–833.

66 25. Geptner A, Kristmannsdóttir H, Kristjánsson J, Marteinsson V. 2002. Biogenic saponite from an active submarine hot spring, Iceland. Clays Clay Miner 50:174–185. 26. Gaidos E, Marteinsson V, Thorsteinsson T, Johannesson T, Runarsson AR, Stefansson A, Glazer B, Lanoil B, Skidmore M, Han S, Miller M, Rusch A, Foo W. 2009. An oligarchic microbial assemblage in the anoxic bottom waters of a volcanic subglacial lake. Isme J 3:486–497. 27. Gaidos E, Lanoil B, Thorsteinsson T, Graham A, Skidmore M, Han SK, Rust T, Popp B. 2004. A viable microbial community in a subglacial volcanic crater lake, Iceland. Astrobiology 4:327–344. 28. Hobel CF V, Marteinsson V, Hreggviðsson GÓ, Kristjánsson JK. 2005. Investigation of the microbial ecology of intertidal hot springs by using diversity analysis of 16S rRNA and chitinase genes. Appl Environ Microbiol 71:2771–2776. 29. Tobler DJ, Benning LG. 2011. Bacterial diversity in five Icelandic geothermal waters: temperature and sinter growth rate effects. Extremophiles 15:473. 30. Pétursdóttir SK, Björnsdóttir SH, Hreggviðsson GÓ, Hjörleifsdóttir S, Kristjánsson JK. 2009. Analysis of the unique geothermal microbial ecosystem of the Blue Lagoon. FEMS Microbiol Ecol 70:425–432. 31. Sæmundsson K, Hjartarson Á, Kaldal I, Sigurgeirsson MA, Kristinsson SG, S V. 2017. Jarðfræðikort af Norðurgosbelti. Nyrðri hluti. 1:100.000. Íslenskar Orkurannsóknir and Landsvirkjun., Reykjavík. 32. Lane DJ. 1991. 16S/23S rRNA sequencing. En: Stackebrandt E., Goodfellow M. (ed), Nucleic acid techniques in bacterial systematics. John Wiley Sons, New York, NY 115–175. 33. Silverman MP, Lundgren DG. 1959. Studies on the chemoautotrophic iron bacterium Ferrobacillus ferrooxidans. J Bacteriol 77:642–647. 34. Kim O-S, Cho Y-J, Lee K, Yoon S-H, Kim M, Na H, Park S-C, Jeon YS, Lee J-H, Yi H, Won S, Chun J. 2012. Introducing EzTaxon-e: a prokaryotic 16S rRNA gene sequence database with phylotypes that represent uncultured species. Int J Syst Evol Microbiol 62:716–21. 35. Löffler F, Yan J, Ritalahti K, Adrian L, Edwards E, Konstantinidis K, Müller J, Fullerton H, Zinder S SA. 2013. Dehalococcoides mccartyi gen. nov., sp. nov., obligately organohalide-respiring anaerobic bacteria relevant to halogen cycling and bioremediation, belong to a novel bacterial class, Dehalococcoidia classis nov., order Dehalococcoidales ord. nov. and famil. Int J Syst Evol Microbiol 63:625–635. 36. Bowman KS, Nobre MF, da Costa MS, Rainey F a, Moe WM. 2012. Dehalogenimonas alkenigignens sp. nov., a chlorinated alkane dehalogenating bacterium isolated from groundwater. Int J Syst Evol Microbiol.

67 37. Moe WM, Yan J, Nobre MF, da Costa MS, Rainey FA. 2009. Dehalogenimonas lykanthroporepellens gen. nov., sp. nov., a reductively dehalogenating bacterium isolated from chlorinated solvent-contaminated groundwater. Int J Syst Evol Microbiol 59:2692– 2697. 38. May HD, Sowers KR. 2016. “Dehalobium chlorocoercia” DF-1-from discovery to application, p. 563–586. In Organohalide-Respiring Bacteria. 39. Nuzzo A, Negroni A, Zanaroli G, Fava F. 2017. Identification of two organohalide-respiring Dehalococcoidia associated to different dechlorination activities in PCB-impacted marine sediments. Microb Cell Fact 16:127. 40. Achenbach LA, Michaelidou U, Bruce RA, Fryman J, Coates JD. 2001. Dechloromonas agitata gen. nov., sp. nov. and Dechlorosoma suillum gen. nov., sp. nov., two novel environmentally dominant (per)chlorate- reducing bacteria and their phylogenetic position. Int J Syst Evol Microbiol 51:527–533. 41. Sanford RA, Cole JR, Tiedje JM. 2002. Characterization and description of Anaeromyxobacter dehalogenans gen. nov., sp. nov., an aryl-halorespiring facultative anaerobic myxobacterium. Appl Environ Microbiol 68:893–900. 42. Coates JD, Chakraborty R, Lack JG, O’Connor SM, Cole KA, Bender KS, Achenbach LA. 2001. Anaerobic benzene oxidation coupled to nitrate reduction in pure culture by two strains of Dechloromonas. Nature 411:1039–1043. 43. Hug LA, Castelle CJ, Wrighton KC, Thomas BC, Sharon I, Frischkorn KR, Williams KH, Tringe SG, Banfield JF. 2013. Community genomic analyses constrain the distribution of metabolic traits across the Chloroflexi phylum and indicate roles in sediment carbon cycling. Microbiome 1:22. 44. McCulloch A, Aucott ML, Benkovitz CM, Graedel TE, Kleiman G, Midgley PM, Li Y-F. 1999. Global emissions of hydrogen chloride and chloromethane from coal combustion, incineration and industrial activities: Reactive Chlorine Emissions Inventory. J Geophys Res 104:8391–8403. 45. Yamada T, Sekiguchi Y, Hanada S, Imachi H, Ohashi A, Harada H, Kamagata Y. 2006. Anaerolinea thermolimosa sp. nov., Levilinea saccharolytica gen. nov., sp. nov. and Leptolinea tardivitalis gen. nov., sp. nov., novel filamentous anaerobes, and description of the new classes Anaerolineae classis nov. and Caldilineae classis nov. in the . Int J Syst Evol Microbiol 56:1331–1340. 46. Schippers A, Kock D, Höft C, Köweker G, Siegert M. 2012. Quantification of Microbial Communities in Subsurface Marine

68 Sediments of the Black Sea and off Namibia. Front Microbiol 3:16. 47. Galand PE, Potvin M, Casamayor EO, Lovejoy C. 2010. Hydrography shapes bacterial biogeography of the deep Arctic Ocean. ISME J 4:564–576. 48. Kato S, Kobayashi C, Kakegawa T, Yamagishi A. 2009. Microbial communities in iron-silica-rich microbial mats at deep-sea hydrothermal fields of the Southern Mariana Trough. Environ Microbiol 11:2094–2111. 49. Cleary DFR, Coelho FJRC, Oliveira V, Gomes NCM, Polónia ARM. 2017. Sediment depth and habitat as predictors of the diversity and composition of sediment bacterial communities in an inter-tidal estuarine environment. Mar Ecol 38:e12411. 50. Fullerton H, Moyer CL. 2016. Comparative Single-Cell Genomics of Chloroflexi from the Okinawa Trough Deep-Subsurface Biosphere. Appl Environ Microbiol 82:3000–3008. 51. Das SK. 2015. BoseaBergey’s Manual of Systematics of Archaea and Bacteria. John Wiley & Sons, Ltd. 52. Kuever J, Rainey FA, Widdel F. 2015. Syntrophobacterales ord. novBergey’s Manual of Systematics of Archaea and Bacteria. John Wiley & Sons, Ltd. 53. Kuever J, Rainey FA, Widdel F. 2015. Syntrophaceae fam. nov.Bergey’s Manual of Systematics of Archaea and Bacteria. John Wiley & Sons, Ltd. 54. Kleindienst S, Herbst F-A, Stagars M, von Netzer F, von Bergen M, Seifert J, Peplies J, Amann R, Musat F, Lueders T, Knittel K. 2014. Diverse sulfate-reducing bacteria of the Desulfosarcina/Desulfococcus clade are the key alkane degraders at marine seeps. ISME J 8:2029– 2044. 55. Carr SA, Orcutt BN, Mandernack KW, Spear JR. 2015. Abundant Atribacteria in deep marine sediment from the Adélie Basin, Antarctica. Front Microbiol 6:872. 56. Hoshino T, Toki T, Ijiri A, Morono Y, Machiyama H, Ashi J, Okamura K, Inagaki F. 2017. Atribacteria from the Subseafloor Sedimentary Biosphere Disperse to the Hydrosphere through Submarine Mud Volcanoes. Front Microbiol 8:1135. 57. Dodsworth JA, Blainey PC, Murugapiran SK, Swingley WD, Ross CA, Tringe SG, Chain PSG, Scholz MB, Lo C-C, Raymond J, Quake SR, Hedlund BP. 2013. Single-cell and metagenomic analyses indicate a fermentative and saccharolytic lifestyle for members of the OP9 lineage. Nat Commun 4:1854. 58. Nobu MK, Dodsworth JA, Murugapiran SK, Rinke C, Gies EA, Webster G, Schwientek P, Kille P, Parkes RJ, Sass H, Jørgensen BB, Weightman AJ, Liu W-T, Hallam SJ, Tsiamis G, Woyke T, Hedlund BP. 2016.

69 Phylogeny and physiology of candidate phylum “Atribacteria” (OP9/JS1) inferred from cultivation-independent genomics. ISME J 10:273–8

70 Supplements Table 9. Description of strains, cultured and isolated on differential and selective media, and conserved in 30% glycerol at -70 °C. Strain T Strain T Media Description Media Description no. (°C) no. (°C) OX0101 Naph 22 White OX0306 2,4-D 22 Tan OX0102 Naph 22 White OX0307 2,4-D 22 Greyish white OX0103 Naph 22 White OX0308 2,4-D 22 White OX0104 HEX 22 Pink OX0309 2,4-D 22 White OX0105 HEX 22 Clear-white OX0310 HEX 22 Tan OX0106 HEX 22 Yellow, shiny OX0311 HEX 22 Pink OX0107 HEX 22 White OX0312 HEX 22 Pale pink, flat, somewhat swarmy OX0108 HEX 22 Pink-clear OX0313 HEX 22 Yellow OX0109 HEX 22 Pink OX0314 HEX 22 White OX0110 HEX 22 White, small OX0315 HEX 22 Clear-white, swarming OX0111 HEX 22 Yellow OX0316 Mn 22 White with a grey centre OX0112 HEX 22 Clear-white, swarming OX0317 Mn 22 Buff OX0113 HEX 22 White OX0318 Mn 22 Red OX0116 Mn 22 White with a grey centre OX0319 R2A 22 Bright yellow OX0117 Mn 22 bright yellow OX0320 R2A 22 White, irregular OX0118 Mn 22 Pink OX0321 R2A 22 Bright red OX0119 Mn 22 snowy white OX0322 R2A 22 Light yellowish brown, irregular OX0120 Mn 22 Clear-yellow "fried egg" OX0323 R2A 22 Very slightly buff OX0121 Mn 22 Yellow, shiny OX0601 Naph 22 White OX0122 R2A 22 Bright orange OX0602 Naph 22 Orange OX0123 R2A 22 Bright orange OX0603 Naph 22 White, flat OX0127 R2A 22 Dark yellow OX0607 Naph 22 White

71 Strain T Strain T Media Description Media Description no. (°C) no. (°C) White w. dark red peak, OX0128 R2A 22 OX0608 LAS 22 Snowy white coherent OX0129 R2A 22 Bright yellow OX0611 HEX 22 White, shiny/oily OX0130 R2A 22 Pale yellow OX0612 HEX 22 White, shiny/oily OX0620 Gui 22 bright yellow OX1215 Hex 22°C Orange White, runny consistency BS4 Transparent-white, transparent OX0621 Gui 22 OX1216 15°C (R2A)* edge OX0622 Gui 22 Buff, lobate margins OX17B03 R2A 22°C Very transparent OX0623 Gui 22 bright yellow OX17B05 R2A 22°C Light-pink OX0624 Gui 22 Buff OX17B06 R2A 22°C Transparent OX0625 R2A 22 Bright red OX1804 R2A 22°C Red, crawls a lot OX0626 R2A 22 Pastel yellow OX1805 R2A 22°C White-transparent, crawls OX0627 R2A 22 Yellow "fried egg" OX1806 Hex 22°C White, round OX0628 R2A 22 White OX1807 Hex 22°C Light yellow, slimy OX0629 R2A 22 Clear-white OX1808 Hex 22°C Pink Reddish brown BS4 White-transparent, transparent OX0630 R2A 22 OX1809 15° (R2A)* edge Bright yellow with a clear, OX0631 R2A 22 OX1905 R2A 22°C Transparent-White, crawls flat halo OX0632 R2A 22 Pink OX1906 R2A 22°C White, irregular, crawls OX0633 R2A 22 Red OX1907 R2A 22°C Yellow, lighter edge, crawls OX1004 R2A 22°C Red, irregular OX1908 Hex 22°C White, small, irregular OX1005 R2A 22°C Small light, crawl much OX1909 Hex 22°C Transparent, slimy OX1006 R2A 22°C Yellow, small, very irregular OX2005 R2A 22°C Very light-yellow, crawls, small Drab (yellow-ish even), OX1007 R2A 22°C OX2006 R2A 22°C Light-drab, small, crawls alot irregular OX1008 R2A 22°C Light-orange, crawls OX2007 R2A 22°C Orange, crawls a lot, lighter edge

Strain T Strain T Media Description Media Description no. (°C) no. (°C) OX1011 Hex 15°C White OX2103 R2A 22°C Light-transparent BS4 OX1012 35°C White-transparent, small OX2104 R2A 22°C White-transpaeant, crawls a lot (R2A)* OX1105 R2A 22°C Small, transparent, round OX2105 R2A 22°C Yellow, crawls OX1106 R2A 22°C Small, drab (yellow-ish) OX2106 Hex 22°C Orange, round Light-orange, lighter OX1107 R2A 22°C OX2107 Hex 22°C Light-orange, round irregular edge Yellow, very irregular, OX1211 R2A 22°C OX1502 R2A 22°C Drab, regular, thin edge lighter edge OX1212 R2A 22°C White, round OX1503 R2A 22°C Creamy, irregular edge Orange, very irregular, OX1213 R2A 22°C OX1504 R2A 22°C Drab, crawls lighter edge OX1214 Hex 22°C White, small OX1505 Hex 15°C Very light orange, small OX1602 R2A 22°C White-transparent, thin edge SX1204 R2A 22°C Light-tranparent, transparent edge Yellow, regular, stuck to the OX1603 R2A 22°C SX1205 R2A 22°C Light-pink, cramped, crawls a lot agar OX1604 Hex 15°C White, small SX1206 R2A 22°C Light-pink, small Creamy, transparent, OX1702 R2A 22°C SX1208 Hex 22°C Light-transparent irregular edge OX2208 R2A 22°C Light-orange, crawls SX1209 Hex 22°C Light orange, round OX2209 Hex 22°C White, small, crawls SX1210 Hex 22°C Light-white, small OX2210 Hex 22°C White, small SX1212 Hex 22°C White, round OX2305 R2A 22°C Light-transparent, crawls SX1213 Hex 22°C White, very slimy Orange, lighter edge, OX2307 R2A 22°C SX1214 Hex 22°C White-transparent, small irregular OX2308 R2A 22°C Orange, round SX1215 Hex 22°C Light-drab, round

73 Strain T Strain T Media Description Media Description no. (°C) no. (°C) Red-pink, round, lighter OX2309 R2A 22°C SX201 R2A 22°C White-transparent, crawls a lot edge OX2310 R2A 22°C White w/ zone SX202 R2A 22°C Light-transparent-yellow, crawls OX2311 R2A 22°C Orange, shines, crawls SX203 R2A 22°C Pink, crawls, slimy OX2312 R2A 22°C Light-transparent, crawls SX204 R2A 22°C Light-pink, big transparent edge OX2313 Hex 22°C Orange, slimy, transparent SX301 R2A 22°C Light-pink, big, slimy BS4 White-transparent, OX2314 22°C SX504 R2A 22°C White, irregular, crawls a lot (R2A)* transparent edge OX2404 R2A 22°C Light-yellow, crawls SX601 R2A 22°C White, lighter edge, crawls a lot Light semi-transparent, OX2405 R2A 22°C SX602 R2A 22°C White, crawls crawls OX2406 Hex 22°C Light-orange, irregular SX0505 Hex 22°C Pink, round OX2514 Hex 22°C Light orange SX1201 R2A 22°C White, crawls a lot OX2515 Hex 22°C Orange SX1202 R2A 22°C Shiny white, crawls 9K Very light-pink, small, OX2516 15° SX0501 R2A 22°C Light-pink, lighter edge, slimy (R2A)* round Light-white, crawls (long SX0302 R2A 22°C SX0502 R2A 22°C Pink, crawls branches) Dark-yellow, crawls a lot, very SX0303 Hex 22°C Transparent SX0503 R2A 22°C slimy SX0304 Hex 22°C White, small , round

74 Table 10. Taxanomic assignment of strains identified by 16S rDNA sequencing and their GenBanka ccession numbers.

Strain EzTaxon Seq. GenBank %ID Order Site1 assignemnt length accession number Alphaproteobacteria OX0102 Brevundimonas 636 MG576001 99.8 Caulobacterales SX bullata OX0309 Brevundimonas 1394 MG576003 99.9 Caulobacterales SX bullata OX2008 Brevundimonas 1423 MG576002 98.9 Caulobacterales AEX halotolerans SX0604 Porphyrobacter 349 MG576010 98.6 Sphingomonadales SX colymbi OX0620 Sphingobium 1451 MG576011 99.4 Sphingomonadales SX xenophagum OX0106 Rhizobium 804 MG576004 99.9 Rhizobiales SX selenitireducens OX0119 Rhizobium 1131 MG576007 99.9 Rhizobiales SX selenitireducens OX1214 Rhizobium 872 MG576006 98.1 Rhizobiales AEX sphaerophysae SX1214 Rhizobium 1447 MG576005 97.3 Rhizobiales SX selenitireducens OX1314 Paracoccus 721 MG576009 98.1 Rhodobacterales AEX homiensis OX0632 Cereibacter 1396 MG576008 99.9 Rhodobacterales SX changlensis Betaproteobacteria OX0105 Acidovorax radicis 744 MG576016 98.8 Burkholderiales SX OX0120 Acidovorax radicis 1472 MG576017 99.7 Burkholderiales SX OX0124 Acidovorax radicis 1380 MG576018 99.5 Burkholderiales SX OX0611 Paucibacter 1441 MG576022 97.9 Burkholderiales SX toxinivorans OX1216 Paraburkholderia 1404 MG576015 99.9 Burkholderiales AEX fungorum

75 OX1313 Paraburkholderia 1489 MG576012 99.9 Burkholderiales AEX fungorum OX1403 Paraburkholderia 1492 MG576013 100 Burkholderiales AEX fungorum OX1702 Paraburkholderia 959 MG576014 100 Burkholderiales AEX fungorum OX0630 Rhodoferax 1360 MG576019 98.5 Burkholderiales AEX fermentans OX0321 Rhodoferax 1440 MG576020 99.3 Burkholderiales AEX saidenbachensis OX0130 Variovorax 1465 MG576021 99.4 Burkholderiales AEX ginsengisoli OX0627 Dechloromonas 1501 MG576023 98.8 Rhodocyclales SX hortensis Gammaproteobacteria OX0110 Aeromonas popoffii 1122 MG576027 99.8 Aeromonadales SX OX0112 Aeromonas popoffii 1506 MG576028 99.7 Aeromonadales SX OX0612 Aeromonas popoffii 1451 MG576029 99.7 Aeromonadales SX OX0622 Aeromonas 960 MG576025 100 Aeromonadales SX hydrophila SX0303 Aeromonas 876 MG576026 100 Aeromonadales SX piscicola OX0617 Aeromonas 721 MG576024 100 Aeromonadales SX cavernicola OX0631 Pseudomonas 1517 MG576051 99.2 Pseudomonadales SX pictorum OX1807 Pseudomonas 671 MG576041 99.4 Pseudomonadales AEX aestusnigri OX1808 Pseudomonas 1536 MG576042 97.9 Pseudomonadales AEX anguillisepticum OX0306 Pseudomonas 1429 MG576043 99.7 Pseudomonadales AEX extremaustralis OX0322 Pseudomonas 1342 MG576044 99.7 Pseudomonadales AEX extremaustralis OX1110 Pseudomonas 1502 MG576045 99.8 Pseudomonadales AEX guineae OX0613 Pseudomonas 1450 MG576046 99.9 Pseudomonadales AEX linyingensis

76 SX0305 Pseudomonas 1416 MG576047 99.9 Pseudomonadales SX mandelii SX0307 Pseudomonas 1503 MG576048 99.9 Pseudomonadales SX mandelii SX1216 Pseudomonas 1503 MG576049 99.9 Pseudomonadales SX mandelii SX0304 Pseudomonas peli 702 MG576050 99.5 Pseudomonadales SX SX1213 Pseudomonas 1500 MG576052 99.7 Pseudomonadales SX vancouverensis OX0304 Pseudomonas 702 MG576053 99.6 Pseudomonadales SX veronii OX0601 Acinetobacter 1501 MG576039 100 Pseudomonadales SX pakistanensis OX0621 Acinetobacter 839 MG576040 100 Pseudomonadales SX pakistanensis OX0619 Shewanella 1511 MG576030 99.5 Alteromonadales SX putrefaciens OX0103 Rahnella aquatilis 576 MG576034 94.4 Enterobacteriales SX OX0101 Rahnella inusitata 1507 MG576035 99.7 Enterobacteriales SX OX0604 Shigella flexneri 1505 MG576036 99.7 Enterobacteriales SX OX0606 Shigella flexneri 1510 MG576037 99.7 Enterobacteriales SX OX0607 Escherichia coli 858 MG576038 99.5 Enterobacteriales SX OX1012 Escerichia 1504 MG576033 99.7 Enterobacteriales AEX fergusonii OX1208 Rheinheimera soli 1515 MG576032 99.0 Chromatiales AEX OX1909 Rheinheimera 977 MG576031 99.1 Chromatiales AEX aestuari Bacilli OX0301 Paenibacillus 1214 MG575992 98.8 Bacillales SX xylanexedens OX0302 Paenibacillus 1510 MG575993 99.7 Bacillales SX xylanexedens OX2310 Paenibacillus 1497 MG575991 99.3 Bacillales AEX tundrae OX0307 Paenibacillus 1502 MG575990 98.9 Bacillales SX terrae OX2205 Paenibacillus alba 961 MG575989 99.7 Bacillales AEX

77 OX0310 Jeotgalibacillus 1484 MG575994 99.4 Bacillales SX campisalis OX0626 Staphylococcus 1409 MG575995 99.9 Bacillales SX argentus OX0317 Psychrobacillus 1496 MG575988 99.5 Bacillales SX psychrodurans OX1213 Bacillus aquimaris 1507 MG575973 99.5 Bacillales AEX OX2513 Bacillus 872 MG575974 98.8 Bacillales AEX halmapalus OX1505 Bacillus 1262 MG575975 99.0 Bacillales AEX hwajinpoensis OX2509 Bacillus 1282 MG575976 99.4 Bacillales AEX hwajinpoensis OX1011 Bacillus 937 MG575977 100 Bacillales AEX oceanisediminis OX1604 Bacillus 363 MG575978 99.2 Bacillales AEX oceanisediminis OX1805 Bacillus safensis 885 MG575979 99.6 Bacillales AEX OX1004 Bacillus 1434 MG575980 99.9 Bacillales AEX vietnamensis OX1006 Bacillus 1351 MG575981 99.9 Bacillales AEX vietnamensis OX1007 Bacillus 1508 MG575982 99.9 Bacillales AEX vietnamensis OX1210 Bacillus 1394 MG575983 99.8 Bacillales AEX vietnamensis OX1212 Bacillus 1509 MG575984 99.9 Bacillales AEX vietnamensis OX2308 Bacillus 1205 MG575985 99.8 Bacillales AEX vietnamensis OX2309 Bacillus 1059 MG575986 99.7 Bacillales AEX vietnamensis OX2506 Bacillus 1374 MG575987 99.9 Bacillales AEX vietnamensis OX2515 Exiguobacterium 861 MG575998 99.8 Lactobacillales AEX oxidotolerans OX2106 Exiguobacterium 754 MG575999 99.9 Lactobacillales AEX profundum

78 OX2313 Exiguobacterium 1053 MG576000 99.4 Lactobacillales AEX profundum OX1107 Exiguobacterium 809 MG575996 99.5 Lactobacillales AEX profundum OX1109 Exiguobacterium 1463 MG575997 98.2 Lactobacillales AEX profundum Flavobacteria OX0623 Flavobacterium 992 MG575964 99.0 Flavobacteriales SX glaciei OX0125 Flavobacterium 1446 MG575965 97.8 Flavobacteriales SX granuli OX0129 Flavobacterium 1449 MG575966 97.8 Flavobacteriales SX granuli OX0311 Flavobacterium 1506 MG575967 97.6 Flavobacteriales SX hydatis OX0314 Flavobacterium 1496 MG575970 98.4 Flavobacteriales SX succinians OX0122 Flavobacterium 1451 MG575968 98.5 Flavobacteriales SX succinians OX0123 Flavobacterium 1402 MG575969 98.4 Flavobacteriales SX succinians OX0127 Flavobacterium 1447 MG575971 98.6 Flavobacteriales SX terrigena Cytophagia OX0104 Algoriphagus 1481 MG575959 97.9 Cytophagales SX alkaliphilus OX0118 Algoriphagus 1471 MG575960 97.5 Cytophagales SX alkaliphilus OX0108 Aquiflexum 1060 MG575961 94.7 Cytophagales SX balticum OX0126 Aquiflexum 1459 MG575962 95.5 Cytophagales SX balticum OX0312 Aquiflexum 1459 MG575963 95.5 Cytophagales SX balticum Sphingobacteria OX0625 Pedobacter ruber 1452 MG575972 98.5 Sphingobacteriales SX

79 Actinobacteria OX0615 Oerskovia 1353 MG575950 98.4 Micrococcales SX paurometabola OX0315 Cryobacterium 1463 MG575951 99.3 Micrococcales SX arcticum OX0319 Cryobacterium 932 MG575952 100 Micrococcales SX arcticum OX0117 Arthrobacter 1471 MG575953 99.0 Micrococcales SX alpinus OX0308 Arthrobacter 1460 MG575954 99.9 Micrococcales SX humicola OX0313 Arthrobacter 1494 MG575955 99.3 Micrococcales SX oryzae OX0107 Pseudarthrobacter 1487 MG575956 99.0 Micrococcales SX siccitolerans OX0614 Sanguibacter 1243 MG575957 99.1 Micrococcales SX suarezii OX0316 Streptomyces 1156 MG575958 99.6 Streptomycetales SX clavifer SX1205 Rhodococcus 1294 MG575949 99.7 Corynebacteriales SX globerulus SX0206 Rhodococcus 1482 MG575948 99.7 Corynebacteriales SX globerulus 1 Site SX is at Skógaeyralón (66.15°N, 16.62°W). Vegetated, wetland area. Some suspected gas seepage, but little or no evidence of geothermal influence. Site AER is at Skógakíll near Ærlækjarsel (66.17°N, 16.57°W). Mostly barren, sandy, estuarine area characterized by geothermal activity.

80 3 Manuscript II – Bioremediative potential of bacteria in cold desert environments

In: Biotechnological Applications of Extremophilic Microorganisms (Life in Extreme Environments) (In press)

Guðný Vala Þorsteinsdóttir and Oddur Vilhelmsson Department of Natural Resource Sciences, University of Akureyri, Iceland

Contents

Bioremediation – general considerations ...... 82 Hydrocarbon degradation ...... 83 Aerobic degradation ...... 84 Anaerobic degradation ...... 84 Effects of environmental conditions ...... 85

Microbial life in cold deserts ...... 87 Bioprospecting cold desert soils for hydrocarbon-degrading microbes ...... 88

Concluding remarks ...... 89

References ...... 90

81 Bioremediation – general considerations

With the opening up of the Arctic to shipping, petroleum extraction, tourism, and other human activities, the risk of pollutant release into Arctic environments is steadily increasing (1). This has in recent years led to increased awareness among the northern countries of the need for environment-friendly processes for pollution control and cleanup, which are now being sought after to replace less eco-friendly practices in many industries, as well as on the municipal level (2). Ironically, the act of cleaning up pollutants from the environment can in some cases be harmful to the environment due to physical and/or chemical disturbance. Bioremediation, that is, the degradation, neutralization or removal of contamination by living organisms, is potentially a less environmentally harmful way of cleaning up environmental pollution. It should be stressed, however, that bioremediation is even under the best of circumstances a very slow process compared with physical methods, and is thus almost always used in conjunction with physical pollution removal in real-world situations (1).

There are multiple technical strategies with which one can approach bioremediation, which can be broadly sorted into in situ techniques and ex situ techniques. Bioremediation in situ is when the pollution is attacked at the polluted site in its “natural environment”, whereas ex situ bioremediation is handled offsite at purpose-built facilities such as biopiles or bioreactors. For in situ bioremediation, the two main strategies thought to be the most promising ways to bioremediate soil and groundwater are bioaugmentation and biostimulation techniques (3). Biostimulation refers to the techniques employed when the indigenous microbial community is stimulated to overcome barriers in metabolic pathways and limitations of degradation. That can be attained by providing nutrients or other compounds that enhance the biodegradation in any way, such as oxygen or other electron receptors. Bioaugmentation is the process of introducing allochthonous microorganisms into the environment, usually because they are better degraders than those of the autochthonous microbiota. These technical considerations for bioremediation in general have been reviewed multiple times (3–8) and we refer the reader to these reviews for an in-depth discussion.

Bioremediation fundamentally rests on the process of biodegradation, whereby an organism depolymerizes and oxidizes an environmental organic compound or macromolecule with the terminal products being carbon dioxide and water, a very common process and a major component of the global carbon cycle (9). This process of microbially mediated decomposition of substances can result either from the microorganism´s utilization of a certain compound as a carbon and energy source, or of the organism´s co-metabolism of that

82 compound (10), leading to a staggering versatility of biodegradation activities within the microbial world. Indeed, the maxim often attributed to C. B. van Niel (11), that for any naturally occurring compound there exists an organism capable of degrading it can, broadly speaking, be taken as true. Or, to put it another way: “given favourable environmental conditions, all natural organic compounds degrade in the end“ (12) and biodegradation by naturally occurring microbial populations is indeed important ecologically. Nevertheless, bioremediation efforts are often confounded by a disparity in the biodegradation behaviour of organisms in natural environments versus in laboratory culture, or between natural environments of differing characteristics (13). When initiating bioremediation of a certain pollutant, either in the laboratory or in the natural environment, the degradation of that pollutant needs to be favourable, that is, the intermediates and products of the degradation pathway must be at least less toxic than the substrate, preferably neutralized or completely degraded.

Hydrocarbon degradation

Hydrocarbons are the most widespread pollutants and are of great concern worldwide because of their toxicity and persistence in the environment (5). Due to their chemical inertness and the high energy barrier that needs to be overcome in order to cleave the apolar C-H bond, microbial degradation of hydrocarbons is intrinsically challenging. Nevertheless, degradation of these very abundant organic molecules occurs in a variety of habitats under either aerobic or anaerobic conditions and organisms capable of utilizing hydrocarbons as a sole energy source are present in almost every environment. The ability of the microbiota in a given environment to degrade the hydrocarbon pollutants present is chiefly dependent on the structure of the hydrocarbon compounds and on oxygen availability, as both of these variables determine which pathways are required for degradation of that specific structure. The degradation efficiency however, is strongly dependent on various environmental and physical factors, for example temperature, pH, amount of nutrients and bioavailability of the hydrocarbons. Many bacteria can use hydrocarbons as an energy source but most of the time that source of energy is not the preferred one, as compounds such as sugars or amino acids that are more readily convertible into substrates in the central energy metabolism pathways such as glycolysis or the tricarboxylic acid cycle are used in preference. However, hydrocarbonoclastic bacteria are highly specialized in utilizing hydrocarbons and play a key role in degrading hydrocarbons in polluted sites (14,15).

83 Aerobic degradation Oxygenase-mediated hydrocarbon degradation is quite commonly encountered in aerobic habitats such as surface soils where it is carried out by a wide variety of bacteria, fungi, and algae. Aliphatic hydrocarbons (n-alkanes) are most commonly degraded through α- and ω-hydroxylation of the alkane to alcohlol, catalyzed by alkane monooxygenase (16). In pseudomonads, electrons for the reaction are relayed through an electron transport chain that involves rubredoxin and an FAD-dependent rubredoxin reductase (17). Aerobic alkane metabolism is exemplified by the Pseudomonas oleovorans alkBFGHJKL operon, which encodes the enzymes required for conversion of alkanes to acetyl-coenzyme A (16). The degradation pathways of alkenes and alkynes are highly similar to alkane-degrading pathways but they can also undergo other additional reactions because of their unsaturated nature (18). Degradation of aromatic hydrocarbons is similarly initiated by oxygenase- mediated hydroxylation, resulting in the formation of catechols, followed by ring cleavage and aldehyde or carboxylic acid formation (16). Typically, the initial reaction is mediated by a dioxygenase (19). For polycyclic aromatics, naphthalene degradation by Pseudomonas putida serves as a well studied model. It involves three plasmid-borne operons, with one (nahAaAbAcAdBFCED) encoding the enzymes required for the conversion of naphthalene to salicylate, the second (nahGTHINLOMKJ) those for the conversion of salicylate to tricarboxylic acid cycle intermediates, and the third (nahR) encodes a regulator for the other two (16,20). Anaerobic degradation

Due to the critical role of O2-dependent oxygenases in biodegradation of both aromatic and aliphatic hydrocarbons, it was generally assumed until about the 1990s that hydrocarbon biodegradation was an exclusively aerobic process. Multiple studies have since demonstrated the presence of hydrocarbon oxidation in anaerobic environments. Under anaerobic conditions, the O2- dependent oxygenase-catalyzed reactions are for the most part rendered irrelevant, although there are a few examples in the literature of intra-aerobic anaerobes capable of deriving oxygen from chlorate or nitrate and can thus employ monooxygenases for hydrocarbon degradation even in anaerobic environments (21,22). For the most part, however, anaerobic hydrocarbon oxidation requires alternative means of C-H bond activation through a variety of less well known reactions using electron acceptors such as sulphate or nitrate (16,23,24). A comparatively well studied example is the oxidation of aromatics through the action of toluene-activating benzylsuccinate synthase and related glycyl radical-bearing alkyl or arylalkylsuccinate synthases. This enzyme adds toluene to a fumarate co-substrate, forming a benzyl-substituted succinate (25,26). Among other examples are the O2-independent

84 hydroxylation of ethylbenzene and ATP-dependent and ATP-independent dearomatization of benzoyl-CoA (26). Anaerobic hydrocarbon oxidation by various members of the Betaproteobacteria, Deltaproteobacteria and Clostridia is now recognized to be a significant contributor to hydrocarbon turnover in nature, occurring wherever hydrocarbon loads exceed oxygen availability, such as in contaminated groundwater and other subsurface environments (27).

Effects of environmental conditions

Since every environment is unique, the bioremediation process sometimes needs to be tailor-made for that environment. The remediation possibilities of polluted sites are determined by the composition of the hydrocarbons, the microbial community composition and the environmental conditions (28). For effective bioremediation, it is important to know the physical and chemical composition of the contaminated soil and the composition of the microbiota present because the soil composition, along with the microbiome composition, dictates the biochemistry predominant at the contaminated site. Thus, the choice and the design of bioremediation techniques has to be in accordance with the biochemical processes, the bioavailability and bioactivity at the polluted site (29,30). However, the environmental factor is often confounded by various complications since it can be difficult to tamper with. In the context of the Arctic environment, the most obvious physicochemical variable to consider is temperature. Low temperature has a profound effect on biodegradation, not only in terms of biological activity, which is naturally hampered by lower reaction rates as dictated by basic thermodynamics, but it also affects various physicochemical properties of both the environment itself and the pollutant present (13). Hydrocarbon degradation is therefore more problematic in cold environments because the viscosity of the pollutant is higher and the solubility is lower (31). Because of this, the physicochemical features of the environment are of primary concern in bioremediation. So, the bottleneck of bioremediation is usually not the biological pathway potential of the bacteria, but rather limitation by the bioavailability and solubility of the targeted pollutant, that both can be adversely affected by lower temperature. Nevertheless, it has been demonstrated that microbial activity in Icelandic soil is governed by substrate availability rather that temperature (32,33), supporting the viable idea of biostimulation in colder areas. Studies have shown that bioremediation on petroleum hydrocarbons with psychrotolerant bacteria can result in degradation of pollutants at 10°C that are similar to the efficiency of bacteria at 30°C (34). Freeze-thaw cycles in polar soil have even been reported to possibly stimulate the degradation of hydrocarbons in arctic soil (35,36). Therefor the use of autochthonous microbial community is often the favoured approach, since these microbes are already adapted to the cold environment and fluctuations in temperature. Putting the indigenous bacterial

85 population to work through intrinsic bioremediation or biostimulation has been suggested to be the most feasible way of remediating hydrocarbons from cold environment (37). Intrinsic bioremediation is the process of letting the environment “heal itself”, but observing and managing the indigenous microbial community while biodegrading the pollutant without enhancing the process. Biostimulation refers to the closely related techniques of when the indigenous microbial community is stimulated to overcome barriers in metabolic pathways and limitations of degradation. That can be attained by providing nutrients or other compounds that enhance the biodegradation in any way, like oxygen or other electron receptors. Arctic soil is usually limited in nitrogen as well as phosphorous and fertilizing arctic soil with nitrogen can enhance the biodegradation of hydrocarbons tremendously (38,39). A study on sub- Antarctic soils showed that 95% of the total hydrocarbon pollution was degraded by indigenous microorganisms within a year at 0-7°C (40). In that study, the use of fertilizer was more effective to stimulate the assemblage of hydrocarbon-degrading bacteria in a desert Antarctic soil than in a vegetated soil. Over-fertilizing can however result in low water activity and therefore inhibition of microbial hydrocarbon degradation (Braddock et al. 1997). Sanscartier et al. (2009) studied the potential for on-site bioremediation and different hydrocarbon removal processes in a polar desert in high arctic soils, where the annual mean in temperature is -15°C. They suggest that in dry, cold deserts, remediation of low-molecular hydrocarbons (nC16). In that study, biostimulation with the addition of surfactant was found to be the most effective treatment of the hydrocarbon contaminated soil (42). The bioavailability of the targeted chemicals is another very important feature regarding bioremediation of pollutants. The fact that most major hydrocarbon pollutants are insoluble or poorly soluble in water makes it difficult for organisms to access the target chemical in order to degrade it. In that case, measures to alter the bioavailability of the pollutants may be important (4,30). In addition to supporting the mixing of water- insoluble hydrocarbons, biosurfactants have been shown to slow down the dispersion of hydrocarbon pollutants during freeze-thaw cycles and by that favouring further bioremediation (43). In fact, biosurfactants are thought to be a promising solution to limited bioavailability in contaminated environments due to chemical structure of the pollutant or other physicochemical barriers of the environment.

86 Microbial life in cold deserts

The environmental conditions in Arctic desert soils, such as in the Icelandic highlands, the Canadian high Arctic or in glacial moraines and forefields in Svalbard and Greenland, are highly restrictive, characterized by year-round low temperatures, freeze-thaw cycles, low water retention, regolith-like “soil“ and largely abiotic sediments, low organic carbon content, and high salinity, resulting in a near total lack of visible vegetation (44). There are pronounced phylogenetic dissimilarities between bacterial communities in unvegetated soil versus vegetated developed soil (45), possibly because of the different role they play in harvesting energy and nutrients. The microbial communities found in these desert environments are thus to a large extent borne by the rock- weathering actions of chemolithotrophic bacteria such as iron and sulphur oxidizers and by carbon and nitrogen fixation by cyanobacteria (46). Indeed, Actinobacteria and Cyanobacteria are far more abundant in unvegetated soils than in vegetated developed soil (Zumsteg et al., 2012). Nevertheless, complex microbial communities have been described in these environments in a number of studies. For example, glacial drift sheets in the Darwin-Hatherton glacier region of Antarctica were found to harbour a community comprising members of eight phyla: Actinobacteria, Bacteroidetes, Deinococcus-Thermus, Proteobacteria, Gemmatimonadetes, Firmicutes, Verrucomicrobia, and Planctomycetes (47). In the Arctic region, microbial desert communities tend to be even more complex than those in comparable alpine or Antarctic environments (48). Nevertheless, a characteristic community structure can be elucidated, strongly dominated by Proteobacteria, Actinobacteria and Chloroflexi (49), whereas the Acidobacteria found to be characteristic of Arctic tundras (50), are generally not found in large numbers in Arctic deserts (49). In recent years, bacterial communities in glacial forefields have been studied in terms of understanding the bacterial communities in poor developed soil and bacterial community succession in the Arctic. Even though low cell number and low activity of bacterial cells can be found in young forefields, the diversity of the bacterial community in poor developed and desert soil is surprisingly high (51). Proteobacteria, Actinobacteria, Acidobacteria, Firmicutes and Cyanobacteria showed high diversity in arctic forefields and the abundance of Alphaproteobacteria increases while the abundance of Betaproteobacteria decreases in unvegetated soils (52). Indeed, even the ice caps themselves are home to complex communities as evidenced by a comparative study of the microbial communities present in the Icelandic glaciers Snæfellsjökull, Langjökull, Eyafjallajökull, Vatnajökull, Drangajökull, and Hofsjökull. In this study, members of the classes Betaproteobacteria, Alphaproteobacteria, Sphingobacteria, and Saprospirae were found to be the most abundant (53). Furthermore, a recent metagenomic study of Greenland cryoconite communities found surprising abundance of

87 genes involved in the degradation of polyaromatic hydrocarbons and polychlorinated biphenyls, indicating that the accumulation of these pollutants in Greenland glacier cryoconite has impacted its microbiota composition (54). There is thus clearly a diverse autochthonous biota present in cold desert and ice environments that can be expected to be affected by polluting input, resulting in increased natural degradation. Nevertheless, several limiting factors can also be expected to limit natural biodegradation of pollutants in these environments.

Bioprospecting cold desert soils for hydrocarbon- degrading microbes

The Arctic environment has attracted considerable attention from bioprospectors in recent years (55–58). Arctic soil microbiotas are recognized as valuable sources of cold-active enzymes, biosurfactants, and various metabolites, as well as for cold-adapted bioremediation (59–63). While the numbers of hydrocarbon degraders is usually low or undetectable in pristine Arctic soils, contaminated soils often display high numbers of hydrocarbon degraders that may persist for decades after the initial spillage (64). Although only a small part of the resident microbiota can be expected to be easily culturable, the aerobic bacteria most readily utilizable as bioremediators can be expected to grow with relative ease in many commercially available growth media. It therefore seems feasible to prospect contaminated cold soils for biodegraders of alkanes and aromatics and, indeed, a large number of such efforts can be found in the literature. Hydrocarbon-degrading members of the genera Rhodococcus, Pseudomonas and Sphingomonas appear most often in these studies (64), although several other taxa have been reported, such as Pedobacter (65), Arthrobacter (66), Acidovorax, and Variovorax (67), among others. An obvious requirement for microorganisms to be used for in situ bioremediation in Arctic environments is that it be cold-adapted. While it seems reasonable to expect isolates from cold deserts to be adapted to the cold, it is important to remember that summertime temperatures in surface soils may periodically approach 20°C and thus some of the bacteria thriving in that environment will have optimal growth rates above the psychrophilic range (15°C). Nevertheless, many of the hydrocarbon-degrading bacteria isolated from cold desert environments have indeed been shown be psychrotrophic and oxidize hydrocarbons at low temperatures. For example, Sphingomonas Ant 17 degraded phenanthrene at 4°C, although higher degradation rates were observed at 28°C (68), and Rhodococcus Q15 mineralized alkanes at 0°C (69), which is traditionally considered to be at the lower temperature threshold for significant hydrocarbon biodegradation, meaning that bioremediation in terrestrial polar environments is likely limited to the warmer summer season

88 (13). However, more recent work has shown this traditional view to be too simplistic as wintertime bacterial activity under snow cover, often at sub-zero temperatures, has been found to be an important factor in Arctic ecology (70,71). It should be borne in mind, however, that there are more limiting factors to hydrocarbon degradation rates in cold climates than simply the microorganism‘s psychrotolerance. As discussed above, temperature will also affect the physical properties of petrochemicals. At lower temperature, the viscosity of oil is increased, the volatilization of short-chain alkanes is reduced and water solubility decreased. The bioavailability of hydrocarbons is thus generally decreased at lower temperatures (72). Nevertheless, hydrocarbon degradation has been demonstrated in situ to occur in soils at sub-zero temperatures in Svalbard (73). Tolerance of wide temperature fluctuations and freeze-thaw cycles are also among requirements for potential bioremediators in Arctic soils. Indeed, some of the hydrocarbon-degrading isolates from cold soils have been found to be well adapted to freeze-thaw cycles. For example, Sphingomonas Ant 17 was found to be more tolerant to freeze-thaw cycles than was the mesophilic Sphingomonas WPO-1 (68) and freeze-thaw microcosm experiments in the presence of diesel fuel indicated that the freeze-thaw regime led to the predominance of hydrocarbon-degrading rhodococci in the microcosms (35). Similarly, a study of hydrocarbon-contaminated Resolution Island-soil communities and their responses to freezing-thawing cycles showed an increase in 14C-hexadecane mineralization and the emergence of Corynebacterineae (of which Rhodococcus is a member) during the freezing phase (36).

Concluding remarks

Although the Arctic deserts and glacial moraines appear barren to the casual observer, they are in fact home to a diverse, abundant, and active microbiota, within which easily culturable hydrocarbon-degrading bacteria can be found. These organisms can be expected to be adapted to harsh Arctic conditions, including low temperature, freeze-thaw cycles, and low organic carbon content and may thus be utilizable for partial bioremediation of contaminated sites in the delicate Arctic environment, although large scale testing of their applicability to real-world situations is mostly lacking. The autochthonous microbiota of Arctic desert and ice environments has also been shown to respond to pollutant input, suggesting that in situ biostimulation techniques may be considered when contemplating bioremediation in these environments.

89 References

1. Atlas RM. Microbial Bioremediation in Polar Environments: Current Status and Future Directions. In: Bej AK, Aislabie J, Atlas RM, editors. Polar Microbiology: The Ecology, Biodiversity and Bioremediation Potential of Microorganisms in Extremely Cold Environments. Boca Raton, FL: Taylor & Francis; 2009. p. 373–91. 2. Gunnarsdóttir R, Jenssen PD, Erland Jensen P, Villumsen A, Kallenborn R. A review of wastewater handling in the Arctic with special reference to pharmaceuticals and personal care products (PPCPs) and microbial pollution. Ecol Eng [Internet]. 2013 Jan [cited 2017 Jul 18];50:76–85. Available from: http://linkinghub.elsevier.com/retrieve/pii/S0925857412001528 3. Tyagi M, da Fonseca MMR, de Carvalho CCCR. Bioaugmentation and biostimulation strategies to improve the effectiveness of bioremediation processes. Biodegradation. 2011;22(2):231–41. 4. Speight JG, Arjoon KK. Bioremediation of Petroleum and Petroleum Products. Bioremediation of Petroleum and Petroleum Products. 2012. 1-567 p. 5. Singh P, Jain R, Srivastava N, Borthakur A, Pal D, Singh R, et al. Current and emerging trends in bioremediation of petrochemical waste: A review. Crit Rev Environ Sci Technol [Internet]. Taylor & Francis; 2017 Apr 18;0. Available from: http://dx.doi.org/10.1080/10643389.2017.1318616 6. Herrero M, Stuckey DC. Bioaugmentation and its application in wastewater treatment: A review. Chemosphere [Internet]. 2015 Dec [cited 2017 Jun 30];140:119–28. Available from: http://linkinghub.elsevier.com/retrieve/pii/S0045653514012181 7. Trujillo ME, Kroppenstedt RM, Fernandez-Molinero C, Schumann P, Martinez-Molina E. Micromonospora lupini sp nov and Micromonospora saelicesensis sp nov., isolated from root nodules of Lupinus angustifolius. Int J Syst Evol Microbiol. [Trujillo, Martha E.; Fernandez-Molinero, Carmen; Martinez-Molina, Eustoquio] Univ Salamanca, Dept Microbiol & Genet, Lab 209, Edificio Dept, E- 37008 Salamanca, Spain. [Kroppenstedt, Reiner M.; Schumann, Peter] Deutsch Sammlung Mikroorganismen & Zellkultu; 2007;57:2799–804. 8. Fernández-Luqueño F, López-Valdez F, Sarabia-Castillo CR, García-

90 Mayagoitia S, Pérez-Ríos SR. Bioremediation of Polycyclic Aromatic Hydrocarbons-Polluted Soils at Laboratory and Field Scale: A Review of the Literature on Plants and Microorganisms. In: Anjum NA, Gill SS, Tuteja N, editors. Enhancing Cleanup of Environmental Pollutants: Volume 1: Biological Approaches [Internet]. Cham: Springer International Publishing; 2017. p. 43–64. Available from: http://dx.doi.org/10.1007/978-3-319-55426-6_4 9. Stevenson FJ, Cole MA. Cycles of soils: carbon, nitrogen, phosphorus, sulfur, micronutrients. New York: John Wiley & Sons; 1999. 427 pp. 10. Horvath RS. Microbial co-metabolism and the degradation of organic compounds in nature. Bacteriol Rev [Internet]. 1972 Jun;36(2):146– 55. Available from: http://www.ncbi.nlm.nih.gov/pmc/articles/PMC408321/ 11. Staley JT, Gunsalus RP, Lory S, Perry JJ. Microbial Life. Sunderland, MA: Sinauer Associates; 2007. 1066 p. 12. Markúsdóttir M. Anthropogenic impact on the microbiota of seashore and freshwater environments in Northern Iceland: preliminary assessment and surfactant-degrader bioprospecting. Vol. M.Sc. thes. [Akureyri]: The University of Akureyri; 2011. 13. Atlas RM. Microbial degradation of petroleum hydrocarbons: an environmental perspective. Microbiol Rev [Internet]. 1981 Mar;45(1):180–209. Available from: http://www.ncbi.nlm.nih.gov/pmc/articles/PMC281502/ 14. McKew BA, Coulon F, Yakimov MM, Denaro R, Genovese M, Smith CJ, et al. Efficacy of intervention strategies for bioremediation of crude oil in marine systems and effects on indigenous hydrocarbonoclastic bacteria. Environ Microbiol. 2007;9(6):1562–71. 15. Rojo F. Degradation of alkanes by bacteria: Minireview. Vol. 11, Environmental Microbiology. 2009. p. 2477–90. 16. Brakstad OG. Natural and stimulated biodegradation of petroleum in cold marine environments. In: Psychrophiles: from Biodiversity to Biotechnology. 2008. p. 389–407. 17. Smits THM, Witholt B, van Beilen JB. Functional characterization of genes involved in alkane oxidation by Pseudomonas aeruginosa . Antonie Van Leeuwenhoek [Internet]. 2003;84(3):193–200. Available from: http://dx.doi.org/10.1023/A:1026000622765 18. Bouwer EJ, Zehnder AJB. Bioremediation of organic compounds -

91 putting microbial metabolism to work. Trends Biotechnol. 1993;11(8):360–7. 19. Cerniglia CE. Biodegradation of polycyclic aromatic hydrocarbons. Biodegradation. 1992;3(2–3):351–68. 20. Li W, Shi J, Wang X, Han Y, Tong W, Ma L, et al. Complete nucleotide sequence and organization of the naphthalene catabolic plasmid pND6-1 from Pseudomonas sp. strain ND6. Gene [Internet]. 2004 [cited 2017 May 20];336(2):231–40. Available from: http://www.sciencedirect.com/science/article/pii/S0378111904001994 21. Zedelius J, Rabus R, Grundmann O, Werner I, Brodkorb D, Schreiber F, et al. Alkane degradation under anoxic conditions by a nitrate- reducing bacterium with possible involvement of the electron acceptor in substrate activation. Environ Microbiol Rep. 2011;3(1):125–35. 22. Mehboob F, Oosterkamp MJ, Koehorst JJ, Farrakh S, Veuskens T, Plugge CM, et al. Genome and proteome analysis of Pseudomonas chloritidismutans AW-1 T that grows on n -decane with chlorate or oxygen as electron acceptor. Environ Microbiol [Internet]. 2015;n/a- n/a. Available from: http://www.ncbi.nlm.nih.gov/pubmed/25900248%5Cnhttp://doi.wiley .com/10.1111/1462-2920.12880 23. Kropp KG, Davidova IA, Suflita JM. Anaerobic oxidation of n- dodecane by an addition reaction in a sulfate-reducing bacterial enrichment culture. Appl Environ Microbiol. 2000;66(12):5393–8. 24. Rabus R, Wilkes H, Behrends A, Armstroff A, Fischer T, Pierik AJ, et al. Anaerobic initial reaction of n-alkanes in a denitrifying bacterium: Evidence for (1-methylpentyl)succinate as initial product and for involvement of an organic radical in n-hexane metabolism. J Bacteriol. 2001;183(5):1707–15. 25. Heider J, Szaleniec M, Martins BM, Seyhan D, Buckel W, Golding BT. Structure and function of benzylsuccinate synthase and related fumarate-adding glycyl radical enzymes. Vol. 26, Journal of Molecular Microbiology and Biotechnology. 2016. p. 29–44. 26. Rabus R, Boll M, Heider J, Meckenstock RU, Buckel W, Einsle O, et al. Anaerobic Microbial Degradation of Hydrocarbons: From Enzymatic Reactions to the Environment. J Mol Microbiol Biotechnol [Internet]. 2016;26(1–3):5–28. Available from: http://www.karger.com/DOI/10.1159/000443997 27. von Netzer F, Kuntze K, Vogt C, Richnow HH, Boll M, Lueders T. Functional Gene Markers for Fumarate-Adding and Dearomatizing

92 Key Enzymes in Anaerobic Aromatic Hydrocarbon Degradation in Terrestrial Environments. J Mol Microbiol Biotechnol [Internet]. 2016;26(1–3):180–94. Available from: http://www.karger.com/DOI/10.1159/000441946 28. Atlas RM. Effects of temperature and crude oil composition on petroleum biodegradation. Appl Microbiol. American Society for Microbiology; 1975 Sep;30(3):396–403. 29. Dua M, Singh A, Sethunathan N, Johri A. Biotechnology and bioremediation: Successes and limitations. Appl Microbiol Biotechnol. 2002;59(2–3):143–52. 30. Blackburn J, Hafker WR. The impact of biochemistry, bioavailability and bioactivity on the selection of bioremediation techniques. Trends Biotechnol. 1993 Aug;11(8):328–33. 31. Margesin R, Schinner F. Biodegradation and bioremediation of hydrocarbons in extreme environments. Appl Microbiol Biotechnol. 2001;56(5–6):650–63. 32. Guicharnaud R, Arnalds O, Paton GI. Short term changes of microbial processes in Icelandic soils to increasing temperatures. Biogeosciences. 2009;7:671–82. 33. Lehtinen T, Mikkonen A, Sigfusson B, Ólafsdóttir K, Ragnarsdóttir KV, Guicharnaud R. Bioremediation trial on aged PCB-polluted soils—a bench study in Iceland. Environ Sci Pollut Res [Internet]. 2014;21(3):1759–68. Available from: http://dx.doi.org/10.1007/s11356-013-2069-z 34. Margesin R. Potential of cold-adapted microorganisms for bioremediation of oil-polluted Alpine soils. Int Biodeterior Biodegrad. 2000;46(1):3–10. 35. Eriksson M, Ka J-O, Mohn WW. Effects of Low Temperature and Freeze-Thaw Cycles on Hydrocarbon Biodegradation in Arctic Tundra Soil. Appl Environ Microbiol [Internet]. 2001 Nov 1;67(11):5107–12. Available from: http://aem.asm.org/content/67/11/5107.abstract 36. Chang W, Klemm S, Beaulieu C, Hawari J, Whyte L, Ghoshal S. Petroleum Hydrocarbon Biodegradation under Seasonal Freeze−Thaw Soil Temperature Regimes in Contaminated Soils from a Sub-Arctic Site. Environ Sci Technol [Internet]. American Chemical Society; 2011 Feb 1;45(3):1061–6. Available from: http://dx.doi.org/10.1021/es1022653

93 37. Van Stempvoort D, Biggar K. Potential for bioremediation of petroleum hydrocarbons in groundwater under cold climate conditions: A review. Cold Reg Sci Technol. 2008;53(1):16–41. 38. Mohn WW, Stewart GR. Limiting factors for hydrocarbon biodegradation at low temperature in Arctic soils. Soil Biol Biochem. 2000 Aug;32(8–9):1161–72. 39. Braddock JF, Ruth ML, Catterall PH, Walworth JL, Mccarthy KA. Enhancement and inhibition of microbial activity in hydrocarbon- contaminated arctic soils: Implications for nutrient-amended bioremediation. Environ Sci Technol. 1997;31(7):2078–84. 40. Delille D, Coulon F, Pelletier E. Effects of temperature warming during a bioremediation study of natural and nutrient-amended hydrocarbon-contaminated sub-Antarctic soils. Cold Reg Sci Technol. 2004;40(1–2):61–70. 41. Braddock JF, Ruth ML, Catterall PH, Walworth JL, McCarthy KA. Enhancement and Inhibition of Microbial Activity in Hydrocarbon- Contaminated Arctic Soils: Implications for Nutrient-Amended Bioremediation. Environ Sci Technol [Internet]. American Chemical Society; 1997 Jul 1;31(7):2078–84. Available from: http://dx.doi.org/10.1021/es960904d 42. Sanscartier D, Laing T, Reimer K, Zeeb B. Bioremediation of weathered petroleum hydrocarbon soil contamination in the Canadian High Arctic: Laboratory and field studies. Chemosphere [Internet]. 2009 Nov;77(8):1121–6. Available from: http://www.sciencedirect.com/science/article/pii/S0045653509010558 43. Yao Y, Huang GH, An CJ, Cheng GH, Wei J. Effects of freeze?thawing cycles on desorption behaviors of PAH-contaminated soil in the presence of a biosurfactant: a case study in western Canada. Environ Sci Process Impacts. 2017; 44. Rhodes M, Knelman J, Lynch RC, Darcy JL, Nemergut DR, Schmidt SK. Alpine and Arctic Soil Microbial Communities. In: Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F, editors. The Prokaryotes: Prokaryotic Communities and Ecophysiology [Internet]. Berlin, Heidelberg: Springer Berlin Heidelberg; 2013. p. 43–55. Available from: http://dx.doi.org/10.1007/978-3-642-30123-0_37 45. Knelman JE, Legg TM, O?Neill SP, Washenberger CL, Gonz?lez A, Cleveland CC, et al. Bacterial community structure and function change in association with colonizer plants during early primary succession in a glacier forefield. Soil Biol Biochem. 2012

94 Mar;46:172–80. 46. Borin S, Ventura S, Tambone F, Mapelli F, Schubotz F, Brusetti L, et al. Rock weathering creates oases of life in a High Arctic desert. Environ Microbiol [Internet]. Blackwell Publishing Ltd; 2010 Feb [cited 2017 May 21];12(2):293–303. Available from: http://doi.wiley.com/10.1111/j.1462-2920.2009.02059.x 47. Aislabie JM, Lau A, Dsouza M, Shepherd C, Rhodes P, Turner SJ. Bacterial composition of soils of the Lake Wellman area, Darwin Mountains, Antarctica. Extremophiles [Internet]. 2013;17(5):775–86. Available from: http://dx.doi.org/10.1007/s00792-013-0560-6 48. Schütte UME, Abdo Z, Foster J, Ravel J, Bunge J, Solheim B, et al. Bacterial diversity in a glacier foreland of the high Arctic. Mol Ecol. 2010;19(SUPPL. 1):54–66. 49. McCann CM, Wade M, Gray ND, Roberts JA, Hubert CRJ, Graham DW. Microbial communities in a High Arctic polar desert landscape. Front Microbiol [Internet]. 2016;7(March):1–10. Available from: http://www.frontiersin.org/Journal/Abstract.aspx?s=420&name=extre me_microbiology&ART_DOI=10.3389/fmicb.2016.00419 50. Männistö MK, Kurhela E, Tiirola M, Häggblom MM. Acidobacteria dominate the active bacterial communities of Arctic tundra with widely divergent winter-time snow accumulation and soil temperatures. FEMS Microbiol Ecol. 2013;84(1):47–59. 51. Sigler WV, Zeyer J. Microbial Diversity and Activity along the Forefields of Two Receding Glaciers. Microb Ecol. 2002 Jun;43(4):397–407. 52. Zumsteg A, Luster J, Göransson H, Smittenberg RH, Brunner I, Bernasconi SM, et al. Bacterial, Archaeal and Fungal Succession in the Forefield of a Receding Glacier. Microb Ecol. 2012 Apr;63(3):552–64. 53. Lutz S, Anesio AM, Edwards A, Benning LG. Microbial diversity on Icelandic glaciers and ice caps. Name Front Microbiol. 2015;6:307. 54. Stibal ALH and TS-P and KAC and JB and DRP and MD and M. Contamination of the Arctic reflected in microbial metagenomes from the Greenland ice sheet. Environ Res Lett [Internet]. 2017;12(7):74019. Available from: http://stacks.iop.org/1748- 9326/12/i=7/a=074019 55. de Pascale D, De Santi C, Fu J, Landfald B. The microbial diversity of Polar environments is a fertile ground for bioprospecting. Mar

95 Genomics [Internet]. 2012 [cited 2017 Jul 18];8:15–22. Available from: http://www.sciencedirect.com/science/article/pii/S1874778712000384 56. Ferrera-Rodríguez O, Greer CW, Juck D, Consaul LL, Martínez- Romero E, Whyte LG. Hydrocarbon-degrading potential of microbial communities from Arctic plants. J Appl Microbiol [Internet]. 2013 Jan 1 [cited 2017 Jul 18];114(1):71–83. Available from: http://doi.wiley.com/10.1111/jam.12020 57. Crisafi F, Giuliano L, Yakimov MM, Azzaro M, Denaro R. Isolation and degradation potential of a cold-adapted oil/PAH-degrading marine bacterial consortium from Kongsfjorden (Arctic region). Rend Lincei [Internet]. 2016 Sep;27(1):261–70. Available from: http://dx.doi.org/10.1007/s12210-016-0550-6 58. Garneau M-È, Michel C, Meisterhans G, Fortin N, King TL, Greer CW, et al. Hydrocarbon biodegradation by Arctic sea-ice and sub-ice microbial communities during microcosm experiments, Northwest Passage (Nunavut, Canada). FEMS Microbiol Ecol [Internet]. 2016 Oct 1;92(10):fiw130-fiw130. Available from: http://dx.doi.org/10.1093/femsec/fiw130 59. Srinivas TNR, Nageswara Rao SSS, Vishnu Vardhan Reddy P, Pratibha MS, Sailaja B, Kavya B, et al. Bacterial Diversity and Bioprospecting for Cold-Active Lipases, Amylases and Proteases, from Culturable Bacteria of Kongsfjorden and Ny-{Å}lesund, Svalbard, Arctic. Curr Microbiol [Internet]. 2009;59(5):537–47. Available from: http://dx.doi.org/10.1007/s00284-009-9473-0 60. Jóelsson JP, Frijónsdóttir H, Vilhelmsson O. Bioprospecting a glacial river in Iceland for bacterial biopolymer degraders. Cold Reg Sci Technol [Internet]. Elsevier B.V.; 2013;96:86–95. Available from: http://dx.doi.org/10.1016/j.coldregions.2013.03.001 61. Markúsdóttir M, Heidmarsson S, Eythórsdóttir A, Magnússon KP, Vilhelmsson O. The natural and anthropogenic microbiota of Glerá, a sub-arctic river in northeastern Iceland. Int Biodeterior Biodegrad [Internet]. Elsevier Ltd; 2013;84:192–203. Available from: http://dx.doi.org/10.1016/j.ibiod.2012.04.001 62. Singh SM, Singh SK, Yadav LALS, Singh PN, Ravindra R. Filamentous Soil Fungi from Ny-Ålesund, Spitsbergen, and Screening for Extracellular Enzymes. Arctic [Internet]. Arctic Institute of North America; 2012;65(1):45–55. Available from: http://www.jstor.org/stable/23187223

96 63. Park HJ, Kim D. Isolation and characterization of humic substances- degrading bacteria from the subarctic Alaska grasslands. J Basic Microbiol [Internet]. 2015 Jan 1 [cited 2017 Jun 30];55(1):54–61. Available from: http://doi.wiley.com/10.1002/jobm.201300087 64. Aislabie J, Foght J. Hydrocarbon-degrading bacteria in contaminated cold soils. In: Bioremediation of Petroleum Hydrocarbons in Cold Regions. New York: Cambridge University Press; 2008. p. 69–83. 65. Margesin R, Spröer C, Schumann P, Schinner F. Pedobacter cryoconitis sp. nov., a facultative psychrophile from alpine glacier cryoconite. Int J Syst Evol Microbiol [Internet]. 2003;53(5):1291–6. Available from: http://ijs.sgmjournals.org/content/53/5/1291.abstract 66. Margesin R, Gander S, Zacke G, Gounot AM, Schinner F. Hydrocarbon degradation and enzyme activities of cold-adapted bacteria and yeasts. Extremophiles [Internet]. 2003;7(6):451–8. Available from: http://dx.doi.org/10.1007/s00792-003-0347-2 67. Eriksson M, Sodersten E, Yu Z, Dalhammar G, Mohn WW. Degradation of Polycyclic Aromatic Hydrocarbons at Low Temperature under Aerobic and Nitrate-Reducing Conditions in Enrichment Cultures from Northern Soils. Appl Environ Microbiol [Internet]. 2003 Jan 1;69(1):275–84. Available from: http://aem.asm.org/content/69/1/275.abstract 68. Baraniecki CA, Aislabie J, Foght JM. Characterization of Sphingomonas sp. Ant 17, an Aromatic Hydrocarbon-Degrading Bacterium Isolated from Antarctic Soil. Microb Ecol. 2002;43:44–54. 69. Whyte LG, Hawari J, Zhou E, Bourbonnière L, Inniss WE, Greer CW. Biodegradation of Variable-Chain-Length Alkanes at Low Temperatures by a Psychrotrophic Rhodococcussp. Appl Environ Microbiol [Internet]. 1998 Jul 1;64(7):2578–84. Available from: http://aem.asm.org/content/64/7/2578.abstract 70. Schmidt SK, Lipson DA. Microbial growth under the snow: Implications for nutrient and allelochemical availability in temperate soils. Plant Soil [Internet]. 2004;259(1):1–7. Available from: http://dx.doi.org/10.1023/B:PLSO.0000020933.32473.7e 71. Nikrad MP, Kerkhof LJ, Häggblom MM. The subzero microbiome: microbial activity in frozen and thawing soils. FEMS Microbiol Ecol [Internet]. 2016 Jun 1;92(6):fiw081-fiw081. Available from: http://dx.doi.org/10.1093/femsec/fiw081 72. Aislabie J, Saul DJ, Foght JM. Bioremediation of hydrocarbon- contaminated polar soils. Extremophiles. 2006;10:171–9.

97 73. Rike AG, Haugen KB, Engene B. In situ biodegradation of hydrocarbons in arctic soil at sub-zero temperatures—field monitoring and theoretical simulation of the microbial activation temperature at a Spitsbergen contaminated site. Cold Reg Sci Technol [Internet]. 2005 [cited 2017 May 23];41(3):189–209. Available from: http://www.sciencedirect.com/science/article/pii/S0165232X04001

98 4 Conclusions and future perspectives

This thesis binds together the fields of geomicrobiology and environmental biotechnology and describes the first microbial community research related to gas seepage pockmarks in Iceland. In chapter 1.4, the aims of the research presented in Manuscript I was described and research questions presented. All aims were met during the research, and research questions were answered as follows:

• Is the natural gas in the gas seepage pockmarks of the same origin at both sampling sites? The natural gas seeping up from the two sampling sites at Öxarfjörður turned out to be of different origin. The gas seepage at Skógakíll (AEX) site was of thermogenic origin and surrounded by geothermal impacted water as described previously, while the gas seepage at Skógalón (SX) turned out to be of mixed origin with more biogenic character and no geothermal activity was detected.

• Is the microbial community of the natural gas seepage pockmarks biochemically relevant to the composition of the natural methane gas detected from the pockmarks? The bacterial community composition was described at both sampling sites revealing notable differences in abundancy at phylum, class and family-level, indicating that the bacterial taxa composition differs between the two sites. However, additional geochemical studies would be needed to determine the relevance of the bacterial composition to the geochemical characteristics of the two sampling sites with statistical significance. Further, the microbial community of only one gas seepage at each site was described and therefore not feasible to infer from that data the biochemical relevance of bacterial taxa to the gas composition.

• Are there taxa within the microbial community of the gas seepage pockmarks that can possibly be used for environmental biotechnological purposes? Community composition at the natural gas seepage pockmarks revealed the presence of bacterial taxa known to have various capabilities that can be of value to environmental biotechnology and bioremediation. Bacterial taxa at the natural gas seepage pockmarks are highly likely to be able to degrade simple

99 and complex hydrocarbons and possibly conduct dehalogenation of halogenated compounds. These bacteria have the potential to be used for the bioremediation of hydrocarbon polluted environments.

• Is it possible to isolate hydrocarbon-degrading or methane-utilising bacteria from the natural gas seepage pockmarks in Öxarfjörður? The strain collection generated during culturing and isolation revealed bacterial strains capable of utilising naphthalene and degradation of 2,4- diphenoxyacetate. Also many of the isolated bacterial strains were characterised to be closely related to known hydrocarbon-degrading bacteria even though futher studies are needed to confirm the degradation capabilities.

Bacterial strains in the strain collection could potentially be used for bioremediative purposes and give the opportunity to continue physiological screening of various degradation pathways. Also, the data generated during amplicon sequencing could be used for further analysis of the bacterial community where specific taxonomic groups could be analysed and compared. In addition the technical replecates in amplicon sequencing of the study sites can be useful for establishing the importance of replication in sequencing- based community analysis. For future research it should be noted that the Skógakíll site (AEX) is a meaningful study site for microbial relations to natural gas seeps at tectonic boundaries, since gas seepage pockmarks are located onshore and therefore easily accessed with low cost, as opposed to gas seepage sites that are commonly found in seafloor sediments. The abundance of Dehalococcoidia class could be interesting to study further in releation to whether or how these bacteria use reductive dehalogenation for energy production, but orders within the Dehalococcoidia class are not very well known and the culturing and isolation of novel representatives could be of considerable scientific value. It is concluded that the natural gas seepage pockmarks in Öxarfjörður harbour potentially valuable microbial taxa for environmental biotechnology. Bioprospecting natural gas seepage environments and wetlands for bacteria with bioremediative potential is therefore feasible. In addition the results of this thesis encourages further geomicrobiological studies in Iceland, related to natural gas seepage environments and fissure swarms.

100 Appendix A

Figure 10. Poster presentation at the 7th Congress of European Microbiologists (FEMS) in Valencia, Spain, 9-13 July 2017.

101

102 Appendix B

Abstract for oral presentation at the 7th International Conference on Polar & Alpine Microbiology (PAM2017) in Nuuk, Greenland, 8-12 September 2017

The microbiota of on-shore gas seepage pockmarks in northern Iceland

Oddur Vilhelmsson1,2, Guðný Vala Þorsteinsdóttir1 og Anett Blischke3 1The University of Akureyri 2The University of Iceland Biomedical Center 3Iceland Geosurvey

Natural gas seepage pockmarks, thought to be at least partly derived from thermogenic gas, are found on shore at two sites by Öxarfjörður bay in northeastern Iceland. These pockmarks are thought to harbour a microbiota adapted to continuous flushing with natural gas composed of short-chain alkanes and simple aromatics. They thus present a rich environment for investigating natural populations of microbial degraders of alkanes and aromatic compounds.

The two sites differ widely in terms of vegetation and geothermal input, with the Skógalón site being mostly barren and strongly impacted by geothermal activity, whereas the Skógakíll site is densely vegetated with no apparent evidence of geothermal activity. Community analysis by 16S rDNA tag sequencing of DNA extracted from seepage pockmark samples reveal a strong dominance of anaerobic, dehalorespiring bacteria from the class Dehalococcoidia, suggesting a microbial ecosystem characterized primarily by anaerobic methane oxidation fuelled by reductive dehalogenation. Several facultatively dehalorespiring bacteria are also present among the cultured bacteria, which also comprise a large number of facultatively lithotrophic Alphaproteobacteria and Betaproteobacteria, suggesting considerable biogeochemical activity in the pockmarks, such as bioweathering and biodeposition activities. Preliminary investigations on the bioremediation potential of the isolated bacteria from the seepage pockmarks at both sites have revealed degraders of naphthalene, hexane and propane.

103