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 Copyright © 2018 Guðný Vala Þorsteinsdóttir All rights reserved 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.

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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

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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).

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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

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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

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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!

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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

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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).

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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

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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

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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

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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

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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

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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

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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.

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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.

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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).

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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; Stra̧poć 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; Stra̧poć 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; Stra̧poć 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

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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

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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.

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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).

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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

0 30 6015Kilometers

DL K

Th

N

A

KF Vatnajökullglacier

H

TJ

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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.

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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).

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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

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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?

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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, oddurv@unak.is

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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

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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

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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

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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)

AEX-A Skógakíll 66°10'2.250 N 16°33'59.687 W

Gas seepage sediment

6,4 35,2

AEX-B Skógakíll 66°10'8.014 N 16°33'56.422 W

Gas seepage water

6,4 53,3

AEX-C Skógakíll 66°10'12.029 N 16°33'56.998 W

Mud-mire ND 30,4

AEX-D Skógakíll 66°10'9.554 N 16°33'57.793 W

Sand-mud 6,4 19,3

SX-A Skógalón 66°09.222 N 016°37.089 W

Lagoon sediment

8,5 12,1

SX-B Skógalón 66°09.244 N 016°37.240 W

Puddle water 8,5 13,4

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.

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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

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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

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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

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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 (µScm-1)

8945 ± 728 773 ± 392 (p = 0.005)

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,

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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) Methane (CH4) [%THCG]

46.1×10-3 (±48.3×10-3) 6.78 (±1.66)

Ethane (C2H6) [%THCG]

2.60×10-3 (±4.50×10-3) 0 (±0)

Propane (C3H8) [%THCG]

1.03×10-3 (±1.79×10-3) 0 (±0)

iso-Pentane (i-C5H12) [%THCG]

18.7×10-3 (±32.3×10-3) 8.00×10-3 (±11.3×10-3)

Pentane (n-C5H12) [%THCG]

73.7×10-3 (±127×10-3) 49.5×10-3 (±70.0×10-3)

Benzene (C6H6) [%THCG]

3.80×10-3 (±6.24×10-3) 7.15×10-3 (±5.45×10-3)

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.

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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.

54

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 seqs

Reads in OTUs

PredictOTUs 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

55

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)

56

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

57

Tabl

e 6.

Bac

teri

al is

olat

es a

nd th

eir t

axon

omic

cla

ssifi

catio

n by

par

tial 1

6S rR

NA g

ene

sequ

enci

ng.

Stra

ins

Gen

Ban

k ac

cess

ion

num

bers

Ez

Taxo

n cl

assi

ficat

ion

Cla

ss

Ord

er

Fam

ily

Gen

us

SX02

06, S

X12

05

MG

5759

48, M

G57

5949

Ac

tinob

acte

ria

Cory

neba

cter

iale

s No

card

iace

ae

Rhod

ococ

cus

OX

0615

M

G57

5950

´´

M

icro

cocc

ales

Ce

llulo

mon

adac

eae

Oer

skov

ia

OX

0315

, OX

0319

M

G57

5951

, MG

5759

52

´´

´´

Mic

roba

cter

iace

ae

Cryo

bact

eriu

m

OX

0117

, OX

0308

, OX

0313

M

G57

5953

-MG

5759

55

´´

´´

Mic

roco

ccac

eae

Arth

roba

cter

O

X01

07

MG

5759

56

´´

´´

´´

Pseu

darth

roba

cter

O

X06

14

MG

5759

57

´´

´´

Sang

uiba

cter

acea

e Sa

ngui

bact

er

OX

0316

M

G57

5958

´´

St

rept

omyc

etal

es

Stre

ptom

ycet

acea

e St

rept

omyc

es

OX

0104

, OX

0118

M

G57

5959

, MG

5759

60

Cyto

phag

ia

Cyto

phag

ales

Cy

clob

acte

riac

eae

Algo

ripha

gus

OX

0108

, OX

0126

, OX

0312

M

G57

5961

-MG

5759

63

´´

´´

´´

Aqui

flexu

m

OX

0623

, O

X01

25,

OX

0129

, O

X03

11,

OX

0122

, O

X01

23,

OX

0314

, OX

0127

M

G57

5964

-MG

5759

71

Flav

obac

teri

ia

Flav

obac

teri

ales

Fl

avob

acte

riac

eae

Flav

obac

teri

um

OX

0625

M

G57

5972

Sp

hing

obac

teri

a Sp

hing

obac

teri

ales

Sp

hing

obac

teri

acea

e Pe

doba

cter

O

X12

13,

OX

2513

, O

X15

05,

OX

2509

, O

X10

11,

OX

1604

, O

X18

05,

OX

1004

, O

X10

06,

OX

1007

, O

X12

10,

OX

1212

, O

X23

08, O

X23

09, O

X25

06

MG

5759

73 -

MG

5759

87

Baci

lli

Baci

llale

s Ba

cilla

ceae

Ba

cillu

s

58

OX

0317

M

G57

5988

´´

´´

´´

Ps

ychr

obac

illus

O

X22

05,

OX

0307

, O

X23

10,

OX

0301

, OX

0302

M

G57

5989

- M

G57

5993

´´

´´

Pa

enib

acill

acea

e Pa

enib

acill

us

OX

0310

M

G57

5994

´´

´´

Pl

anoc

occa

ceae

Je

otga

libac

illus

O

X06

26

MG

5759

95

´´

´´

Stap

hylo

cocc

acea

e St

aphy

loco

ccus

O

X11

07,

OX

1109

, O

X25

15,

OX

2106

, OX

2313

M

G57

5996

-MG

5760

00

´´

Lact

obac

illal

es

ince

rtae

sedi

s Ex

iguo

bact

eriu

m

OX

0102

, OX

2008

, OX

0309

M

G57

6001

-MG

5760

03

Alph

apro

teob

acte

ria

Caul

obac

tera

les

Caul

obac

tera

ceae

Br

evun

dim

onas

O

X01

06,

OX

1214

, O

X01

19,

SX12

14

MG

5760

04-M

G57

6007

´´

Rh

izobi

ales

Rh

izobi

acea

e Rh

izobi

um

OX

0632

M

G57

6008

´´

Rh

odob

acte

rale

s Rh

odob

acte

race

ae

Cere

ibac

ter

OX

1314

M

G57

6009

´´

´´

´´

Pa

raco

ccus

SX

0604

M

G57

6010

´´

Sp

hing

omon

adal

es

Eryt

hrob

acte

race

ae

Porp

hyro

bact

er

OX

0620

M

G57

6011

´´

´´

Sp

hing

omon

adac

eae

Sphi

ngob

ium

O

X13

13,

OX

1403

, O

X17

02,

OX

1216

M

G57

6012

-MG

5760

15

Beta

prot

eoba

cter

ia

Burk

hold

eria

les

Burk

hold

eria

ceae

Pa

rabu

rkho

lder

ia

OX

0105

, OX

0120

, OX

0124

M

G57

6016

-MG

5760

18

´´

´´

Com

amon

adac

eae

Acid

ovor

ax

OX

0630

, OX

0321

M

G57

6019

, MG

5760

20

´´

´´

´´

Rhod

ofer

ax

OX

0130

M

G57

6021

´´

´´

´´

Va

riovo

rax

OX

0611

M

G57

6022

´´

´´

in

certa

e se

dis

Pauc

ibac

ter

OX

0627

M

G57

6023

´´

Rh

odoc

ycla

les

Rhod

ocyc

lace

ae

Dec

hlor

omon

as

59

OX

0617

, O

X06

22,

SX03

03,

OX

0110

, OX

0112

, OX

0612

M

G57

6024

-MG

5760

29

Gam

map

rote

obac

teri

a Ae

rom

onad

ales

Ae

rom

onad

acea

e Ae

rom

onas

O

X06

19

MG

5760

30

´´

Alte

rom

onad

ales

Sh

ewan

ella

ceae

Sh

ewan

ella

O

X19

09, O

X12

08

MG

5760

31-M

G57

6032

´´

Ch

rom

atia

les

Chro

mat

iace

ae

Rhei

nhei

mer

a O

X10

12

MG

5760

33

´´

Ente

roba

cter

iale

s En

tero

bact

eria

ceae

Es

cher

ichi

a O

X01

03, O

X01

01

MG

5760

34-M

G57

6035

´´

´´

´´

Ra

hnel

la

OX

0604

, OX

0606

, OX

0607

M

G57

6036

-MG

5760

38

´´

´´

´´

Shig

ella

O

X06

01, O

X06

21

MG

5760

39-M

G57

6040

´´

Ps

eudo

mon

adal

es

Mor

axel

lace

ae

Acin

etob

acte

r O

X18

07,

OX

1808

, O

X03

06,

OX

0322

, O

X11

10,

OX

0613

, SX

0305

, SX

0307

, SX

1216

, SX

0304

, O

X06

31,

SX12

13,

OX

0304

M

G57

6041

-MG

5760

53

´´

´´

Pseu

dom

onad

acea

e Ps

eudo

mon

as

60

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

61

(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

62

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

63

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-proteobacteria

0.43 0.23

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-proteobacteria

0.48 0.63

Rhizobiales 0.88 0.92 Rhodobacterales 0.06 0.02 Others 0.06 0.05 Gamma-proteobacteria

0.11 0.06

Beta-proteobacteria

0.06 0.07

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

64

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.

65

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ls

SX12

09

Hex

22

°C

Ligh

t ora

nge,

roun

d O

X22

09

Hex

22

°C

Whi

te, s

mal

l, cr

awls

SX

1210

H

ex

22°C

Li

ght-w

hite

, sm

all

OX

2210

H

ex

22°C

W

hite

, sm

all

SX12

12

Hex

22

°C

Whi

te, r

ound

O

X23

05

R2A

22

°C

Ligh

t-tra

nspa

rent

, cra

wls

SX

1213

H

ex

22°C

W

hite

, ver

y sl

imy

OX

2307

R

2A

22°C

O

rang

e, li

ghte

r edg

e,

irreg

ular

SX

1214

H

ex

22°C

W

hite

-tran

spar

ent,

smal

l

OX

2308

R

2A

22°C

O

rang

e, ro

und

SX12

15

Hex

22

°C

Ligh

t-dra

b, ro

und

74

Stra

in

no.

Med

ia

T (°C

) D

escr

iptio

n St

rain

no

. M

edia

T (°C

) D

escr

iptio

n

OX

2309

R

2A

22°C

R

ed-p

ink,

roun

d, li

ghte

r ed

ge

SX20

1 R

2A

22°C

W

hite

-tran

spar

ent,

craw

ls a

lot

OX

2310

R

2A

22°C

W

hite

w/ z

one

SX

202

R2A

22

°C

Ligh

t-tra

nspa

rent

-yel

low

, cra

wls

O

X23

11

R2A

22

°C

Ora

nge,

shin

es, c

raw

ls

SX20

3 R

2A

22°C

Pi

nk, c

raw

ls, s

limy

O

X23

12

R2A

22

°C

Ligh

t-tra

nspa

rent

, cra

wls

SX

204

R2A

22

°C

Ligh

t-pin

k, b

ig tr

ansp

aren

t edg

e O

X23

13

Hex

22

°C

Ora

nge,

slim

y, tr

ansp

aren

t SX

301

R2A

22

°C

Ligh

t-pin

k, b

ig, s

limy

OX

2314

B

S4

(R2A

)*

22°C

W

hite

-tran

spar

ent,

trans

pare

nt e

dge

SX50

4 R

2A

22°C

W

hite

, irr

egul

ar, c

raw

ls a

lot

OX

2404

R

2A

22°C

Li

ght-y

ello

w, c

raw

ls

SX60

1 R

2A

22°C

W

hite

, lig

hter

edg

e, c

raw

ls a

lot

OX

2405

R

2A

22°C

Li

ght s

emi-t

rans

pare

nt,

craw

ls

SX60

2 R

2A

22°C

W

hite

, cra

wls

OX

2406

H

ex

22°C

Li

ght-o

rang

e, ir

regu

lar

SX05

05

Hex

22

°C

Pink

, rou

nd

OX

2514

H

ex

22°C

Li

ght o

rang

e SX

1201

R

2A

22°C

W

hite

, cra

wls

a lo

t O

X25

15

Hex

22

°C

Ora

nge

SX12

02

R2A

22

°C

Shin

y w

hite

, cra

wls

OX

2516

9K

(R

2A)*

15

° V

ery

light

-pin

k, sm

all,

roun

d SX

0501

R

2A

22°C

Li

ght-p

ink,

ligh

ter e

dge,

slim

y

SX03

02

R2A

22

°C

Ligh

t-whi

te, c

raw

ls (l

ong

bran

ches

) SX

0502

R

2A

22°C

Pi

nk, c

raw

ls

SX03

03

Hex

22

°C

Tran

spar

ent

SX05

03

R2A

22

°C

Dar

k-ye

llow

, cra

wls

a lo

t, ve

ry

slim

y SX

0304

H

ex

22°C

W

hite

, sm

all ,

roun

d

75

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

Strain EzTaxon assignemnt

Seq. length

GenBank accession number

%ID Order Site1

Alphaproteobacteria

OX0102 Brevundimonas bullata

636 MG576001 99.8 Caulobacterales SX

OX0309 Brevundimonas bullata

1394 MG576003 99.9 Caulobacterales SX

OX2008 Brevundimonas halotolerans

1423 MG576002 98.9 Caulobacterales AEX

SX0604 Porphyrobacter colymbi

349 MG576010 98.6 Sphingomonadales SX

OX0620 Sphingobium xenophagum

1451 MG576011 99.4 Sphingomonadales SX

OX0106 Rhizobium selenitireducens

804 MG576004 99.9 Rhizobiales SX

OX0119 Rhizobium selenitireducens

1131 MG576007 99.9 Rhizobiales SX

OX1214 Rhizobium sphaerophysae

872 MG576006 98.1 Rhizobiales AEX

SX1214 Rhizobium selenitireducens

1447 MG576005 97.3 Rhizobiales SX

OX1314 Paracoccus homiensis

721 MG576009 98.1 Rhodobacterales AEX

OX0632 Cereibacter changlensis

1396 MG576008 99.9 Rhodobacterales SX

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 toxinivorans

1441 MG576022 97.9 Burkholderiales SX

OX1216 Paraburkholderia fungorum

1404 MG576015 99.9 Burkholderiales AEX

76

OX1313 Paraburkholderia fungorum

1489 MG576012 99.9 Burkholderiales AEX

OX1403 Paraburkholderia fungorum

1492 MG576013 100 Burkholderiales AEX

OX1702 Paraburkholderia fungorum

959 MG576014 100 Burkholderiales AEX

OX0630 Rhodoferax fermentans

1360 MG576019 98.5 Burkholderiales AEX

OX0321 Rhodoferax saidenbachensis

1440 MG576020 99.3 Burkholderiales AEX

OX0130 Variovorax ginsengisoli

1465 MG576021 99.4 Burkholderiales AEX

OX0627 Dechloromonas hortensis

1501 MG576023 98.8 Rhodocyclales SX

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 hydrophila

960 MG576025 100 Aeromonadales SX

SX0303 Aeromonas piscicola

876 MG576026 100 Aeromonadales SX

OX0617 Aeromonas cavernicola

721 MG576024 100 Aeromonadales SX

OX0631 Pseudomonas pictorum

1517 MG576051 99.2 Pseudomonadales SX

OX1807 Pseudomonas aestusnigri

671 MG576041 99.4 Pseudomonadales AEX

OX1808 Pseudomonas anguillisepticum

1536 MG576042 97.9 Pseudomonadales AEX

OX0306 Pseudomonas extremaustralis

1429 MG576043 99.7 Pseudomonadales AEX

OX0322 Pseudomonas extremaustralis

1342 MG576044 99.7 Pseudomonadales AEX

OX1110 Pseudomonas guineae

1502 MG576045 99.8 Pseudomonadales AEX

OX0613 Pseudomonas linyingensis

1450 MG576046 99.9 Pseudomonadales AEX

77

SX0305 Pseudomonas mandelii

1416 MG576047 99.9 Pseudomonadales SX

SX0307 Pseudomonas mandelii

1503 MG576048 99.9 Pseudomonadales SX

SX1216 Pseudomonas mandelii

1503 MG576049 99.9 Pseudomonadales SX

SX0304 Pseudomonas peli 702 MG576050 99.5 Pseudomonadales SX

SX1213 Pseudomonas vancouverensis

1500 MG576052 99.7 Pseudomonadales SX

OX0304 Pseudomonas veronii

702 MG576053 99.6 Pseudomonadales SX

OX0601 Acinetobacter pakistanensis

1501 MG576039 100 Pseudomonadales SX

OX0621 Acinetobacter pakistanensis

839 MG576040 100 Pseudomonadales SX

OX0619 Shewanella putrefaciens

1511 MG576030 99.5 Alteromonadales SX

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 fergusonii

1504 MG576033 99.7 Enterobacteriales AEX

OX1208 Rheinheimera soli 1515 MG576032 99.0 Chromatiales AEX

OX1909 Rheinheimera aestuari

977 MG576031 99.1 Chromatiales AEX

Bacilli

OX0301 Paenibacillus xylanexedens

1214 MG575992 98.8 Bacillales SX

OX0302 Paenibacillus xylanexedens

1510 MG575993 99.7 Bacillales SX

OX2310 Paenibacillus tundrae

1497 MG575991 99.3 Bacillales AEX

OX0307 Paenibacillus terrae

1502 MG575990 98.9 Bacillales SX

OX2205 Paenibacillus alba 961 MG575989 99.7 Bacillales AEX

78

OX0310 Jeotgalibacillus campisalis

1484 MG575994 99.4 Bacillales SX

OX0626 Staphylococcus argentus

1409 MG575995 99.9 Bacillales SX

OX0317 Psychrobacillus psychrodurans

1496 MG575988 99.5 Bacillales SX

OX1213 Bacillus aquimaris 1507 MG575973 99.5 Bacillales AEX

OX2513 Bacillus halmapalus

872 MG575974 98.8 Bacillales AEX

OX1505 Bacillus hwajinpoensis

1262 MG575975 99.0 Bacillales AEX

OX2509 Bacillus hwajinpoensis

1282 MG575976 99.4 Bacillales AEX

OX1011 Bacillus oceanisediminis

937 MG575977 100 Bacillales AEX

OX1604 Bacillus oceanisediminis

363 MG575978 99.2 Bacillales AEX

OX1805 Bacillus safensis 885 MG575979 99.6 Bacillales AEX

OX1004 Bacillus vietnamensis

1434 MG575980 99.9 Bacillales AEX

OX1006 Bacillus vietnamensis

1351 MG575981 99.9 Bacillales AEX

OX1007 Bacillus vietnamensis

1508 MG575982 99.9 Bacillales AEX

OX1210 Bacillus vietnamensis

1394 MG575983 99.8 Bacillales AEX

OX1212 Bacillus vietnamensis

1509 MG575984 99.9 Bacillales AEX

OX2308 Bacillus vietnamensis

1205 MG575985 99.8 Bacillales AEX

OX2309 Bacillus vietnamensis

1059 MG575986 99.7 Bacillales AEX

OX2506 Bacillus vietnamensis

1374 MG575987 99.9 Bacillales AEX

OX2515 Exiguobacterium oxidotolerans

861 MG575998 99.8 Lactobacillales AEX

OX2106 Exiguobacterium profundum

754 MG575999 99.9 Lactobacillales AEX

79

OX2313 Exiguobacterium profundum

1053 MG576000 99.4 Lactobacillales AEX

OX1107 Exiguobacterium profundum

809 MG575996 99.5 Lactobacillales AEX

OX1109 Exiguobacterium profundum

1463 MG575997 98.2 Lactobacillales AEX

Flavobacteria

OX0623 Flavobacterium glaciei

992 MG575964 99.0 Flavobacteriales SX

OX0125 Flavobacterium granuli

1446 MG575965 97.8 Flavobacteriales SX

OX0129 Flavobacterium granuli

1449 MG575966 97.8 Flavobacteriales SX

OX0311 Flavobacterium hydatis

1506 MG575967 97.6 Flavobacteriales SX

OX0314 Flavobacterium succinians

1496 MG575970 98.4 Flavobacteriales SX

OX0122 Flavobacterium succinians

1451 MG575968 98.5 Flavobacteriales SX

OX0123 Flavobacterium succinians

1402 MG575969 98.4 Flavobacteriales SX

OX0127 Flavobacterium terrigena

1447 MG575971 98.6 Flavobacteriales SX

Cytophagia

OX0104 Algoriphagus alkaliphilus

1481 MG575959 97.9 Cytophagales SX

OX0118 Algoriphagus alkaliphilus

1471 MG575960 97.5 Cytophagales SX

OX0108 Aquiflexum balticum

1060 MG575961 94.7 Cytophagales SX

OX0126 Aquiflexum balticum

1459 MG575962 95.5 Cytophagales SX

OX0312 Aquiflexum balticum

1459 MG575963 95.5 Cytophagales SX

Sphingobacteria

OX0625 Pedobacter ruber 1452 MG575972 98.5 Sphingobacteriales SX

80

Actinobacteria

OX0615 Oerskovia paurometabola

1353 MG575950 98.4 Micrococcales SX

OX0315 Cryobacterium arcticum

1463 MG575951 99.3 Micrococcales SX

OX0319 Cryobacterium arcticum

932 MG575952 100 Micrococcales SX

OX0117 Arthrobacter alpinus

1471 MG575953 99.0 Micrococcales SX

OX0308 Arthrobacter humicola

1460 MG575954 99.9 Micrococcales SX

OX0313 Arthrobacter oryzae

1494 MG575955 99.3 Micrococcales SX

OX0107 Pseudarthrobacter siccitolerans

1487 MG575956 99.0 Micrococcales SX

OX0614 Sanguibacter suarezii

1243 MG575957 99.1 Micrococcales SX

OX0316 Streptomyces clavifer

1156 MG575958 99.6 Streptomycetales SX

SX1205 Rhodococcus globerulus

1294 MG575949 99.7 Corynebacteriales SX

SX0206 Rhodococcus globerulus

1482 MG575948 99.7 Corynebacteriales SX

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.

81

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

82

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

83

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).

84

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

85

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

86

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) can be subjected to volatilization while it is necessary to biodegrade high-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.

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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

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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

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(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.

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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

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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.

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Appendix A

Figure 10. Poster presentation at the 7th Congress of European Microbiologists

(FEMS) in Valencia, Spain, 9-13 July 2017.

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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.