Microbial Diversity, Activity, and Ecology of a Hypersaline High Arctic Spring System

Chih-Ying Lay

Department of Natural Resource Sciences

McGill University, Montreal

August, 2013

A thesis submitted to McGill University in partial fulfillment of the requirements of the

degree of PhD.

Although we have no rational grounds for believing in an objective reality, we also have no choice but to act as if it is true. - David Hume

ACKNOWLEDGEMENTS First of all, I would like to thank Dr. Lyle G. Whyte, my supervisor, who let me study in his lab and supported me with his resources to complete my PhD research. From his supervision, I acquired a lot of knowledge of experiment design, planning field trips, interpreting research results, and building academic networks. His enthusiasm on unique microbiology topics and desires of using newly-developed technologies always encouraged me to face all the trendiest topics in the frontline of environmental microbiology. He let me to attend the unforgettable field trip to the Canadian High Arctic to perform field works. Without his help, I will not finish this thesis. I also would like to thank all the support from Dr. Charles Greer, Dr. Brian Driscoll, Dr. Donald Niven, and Dr. Sébastien Faucher. They always kindly gave me many useful and professional advices to overcome research problems. I would like to thank Dr. Thomas Niederberger, Dr. Nadia Mykytczuk and Dr. Étienne Yergeau. When I had in situ questions or problems for my research, they always offered me the most immediate helps. My lab mates, Guillaume Lamarche-Gagnon, Sara Sheibani, Roland Wilhelm, Kris Radtke, Jen Allan, Jackie Goordial, Diana Popa, Sara Klemm, Dr. Christine Martineau, Dr. Ofelia Ferrera Rodriquez, Dr. Jennifer Ronholm, Dr. Olga Onyshchenko, and Dr. Helen Vrionis were very friendly and helpful to me. In the last five years, when I needed their friendship, advices, or help, they always offered me much more than I expected. They helped me to adapt to the life and work style at Macdonald campus. I also would like to thank all the members of the Microbiology Division. They enriched my microbiology knowledge through seminars and conversations. I would also like to thank NRS support staff, Dave Meek, Marie Kubecki, Ann Gossage, and Marlene Parkinson. They tried their best to keep me in line and guided me when I feel confused with the school system. I would like to thank Dr. Joann Whalen and Hélène Lalande for the soil/sediment analyses. I also would like to thank Dr. Anthony Cushing for doing a final English edition for my thesis and Patricia Görner-Potvin for translating my abstract into French. The Polar Continental Shelf Project, the Canadian Astrobiology Training Program, National Sciences and Engineering Research Council of Canada, Canadian Space Agency, and Northern Scientific Training Program all contributed to making this thesis possible. During the past five years, I had enormous friendship support from Ting-Heng Yu, Chia-Chen Chang, Ming-Yueh Wu, James Wang, Dr. Eric Huang, Li-Jen Chen, Gengrui Wang, Chen Chen, Seamus McClare, Nathaniel Fink, Timothy Schwinghamer, Arturo Mayorga, Claude Gravel, and members from Sainte Anne Singers, Musica Orbium, and McGill Taiwanese Graduate Student Association. Finally, I would like to thank my parents, Jiunn-Yuan Lay and Dr. Pen-Ho Yeh, who encouraged me to study in Canada. Thank you! i

TABLE OF CONTENTS ABSTRACT ...... vi RÉSUMÉ ...... viii CONTRIBUTIONS TO KNOWLEDGE ...... x LIST OF TABLES ...... xi LIST OF FIGURES ...... xii LIST OF ABBREVIATIONS ...... xiii CHAPTER 1 ...... 1 Introduction and Literature Review ...... 1 1.1 Introduction ...... 1 1.2 Terrestrial saline water body ecosystems in Polar regions ...... 2 1.2.1 Definitions of terrestrial saline water bodies ...... 2 1.2.2 Saline lakes in the Polar regions and the microbiology studies on them ……………… ...... 4 1.2.3 Saline springs in Polar regions ...... 7 1.3 Challenges to microbial life in Polar saline water bodies ...... 11 1.3.1 The availability of liquid water in cryoenvironments ...... 11 1.3.2 The adaptation of microorganisms to cryoenvironments ...... 12 1.3.2.1 Cold adaptations of microorganisms ...... 13 1.3.2.2 Saline adaptation of microorganisms ...... 16 1.4 Applications and astrobiology aspects of the study ...... 18 1.4.1 Potential applications of microorganisms from cold saline environments ...... 18 1.4.2 Astrobiological aspects ...... 20 1.5 Objectives ...... 22 CONNECTING TEXT ...... 24 CHAPTER 2 ...... 24 Microbial Diversity and Activity in Hypersaline High Arctic Spring Channels ...... 24 ABSTRACT ...... 25 2.1 Introduction ...... 26 2.2 Materials and Methods ...... 29 2.2.1 Sample site description and geochemical analyses ...... 29

2.2.2 CO 2 and CH 4 concentrations and flux measurements ...... 31 2.2.3 Microscopy and catalyzed reporter deposition fluorescence in situ hybridization (CARD-FISH) ...... 32 2.2.4 Microbial cultivation and characterization ...... 33 ii

2.2.5 Bacterial and Archaeal 16S rRNA gene clone libraries ...... 35 2.2.6 Biodiversity indices and statistical analysis of 16S rRNA gene clone libraries ...... 36 2.2.7 Microbial activity at cold temperatures ...... 37 2.2.8 Nucleotide accession numbers ...... 37 2.3 Results ...... 38 2.3.1 Geochemical analyses ...... 38

2.3.2 CO 2 and CH 4 concentrations and flux measurements ...... 39 2.3.3 Cell enumeration ...... 40 2.3.4 Identification and characterization of isolates ...... 40 2.3.5 Bacterial and Archaeal 16S rRNA gene clone libraries ...... 41 2.3.6 Microbial activity at cold temperatures ...... 44 2.4 Discussion ...... 44 2.5 Acknowledgements ...... 52 CONNECTING TEXT ...... 62 CHAPTER 3 ...... 62 Defining the Functional Potential and Active Community Members of a Sediment Microbial Community in a High Arctic Hypersaline Subzero Spring ...... 62 ABSTRACT ...... 63 3.1 Introduction ...... 64 3.2 Materials and Methods ...... 68 3.2.1 Study site and sample collection ...... 68 3.2.2 Metagenomic DNA extraction and sequencing ...... 68 3.2.3 Metagenomic DNA analyses ...... 69 3.2.4 Statistical analyses ...... 71 3.2.5 RNA extraction and 16S ribosomal cDNA analyses ...... 72 3.2.6 Nucleotide and metagenome sequence accession numbers ...... 73 3.3 Results and Discussion ...... 74 3.3.1 Metagenomic sequencing statistics ...... 74 3.3.2 Metagenomic microbial community composition ...... 75 3.3.3 Functional gene profiles of the LH metagenome ...... 78 3.3.4 Methane metabolism ...... 79 3.3.5 Nitrogen metabolism ...... 80 3.3.6 Sulfur Metabolism ...... 81 3.3.7 Stress response ...... 82 3.3.8 Comparison with other metagenomes ...... 85 iii

3.3.9 Active profiling of LH based on 16S ribosomal cDNA pyrosequencing . 87 3.4 Conclusion ...... 90 3.5 Acknowledgements ...... 91 CONNECTING TEXT ...... 99 CHAPTER 4 ...... 99 Seasonal Changes in Microbial Communities at a Hypersaline Spring Channel and the Adjacent Tundra ...... 99 ABSTRACT ...... 99 4.1 Introduction ...... 100 4.2 Materials and methods ...... 105 4.2.1 Sample collection and geochemical analyses ...... 105 4.2.2 DNA and RNA extraction, cDNA synthesis, pyrosequencing and analyses...... 106 4.2.4 UniFrac analysis of the LH libraries ...... 108 4.2.5 Archaeal amo A and hcd gene cloning and sequencing and analyses ..... 109 4.2.6 qPCR of Thaumarchaeal 16S/ amo A/ hcd genes in LH channel sediments and tundra ...... 110 4.3 Results ...... 112 4.3.1 Geochemical analyses of the LH channel and tundra sampling sites .... 112 4.3.2 Pyrosequencing library statistics ...... 113 4.3.3 Microbial compositions in LH spring channel and tundra in the summer ...... 115 4.3.4 Microbial compositions in LH spring channel and the tundra in the winter ...... 117 4.3.5 Archaeal functional genes for ammonia oxidation and carbon fixation 119 4.4 Discussion ...... 120 4.4.1 Seasonal changes in active microbial components in LH channel area . 120 4.4.2 Microbial biodiversity and richness in LH channel sediments ...... 124 4.4.3 Thaumarchaeal signature functional genes in the LH channel sediment and the adjacent tundra ...... 128 4.5 Conclusion ...... 133 4.6 Acknowledgements ...... 134 CHAPTER 5 ...... 134 Discussion and Conclusions ...... 134 5.1 Microbial diversity and activity in the hypersaline spring channel ...... 134 5.2 Functional potential and the active components at LH outlet ...... 135 iv

5.3 Seasonal changes in microbial communities at a hypersaline spring channel and the adjacent tundra ...... 136 5.4 Conclusions ...... 138 References ...... 142 APPENDIX: Supporting Information ...... 182

ABSTRACT

The Lost Hammer (LH) spring, located on Axel Heiberg Island in the Canadian

High Arctic, is the coldest and saltiest terrestrial spring discovered to date. It is characterized by perennial discharges of subzero temperatures (-5°C), hypersalinity (24% salinity), along with reducing ( ≈-165 mV), microoxic, and oligotrophic conditions. It is rich in sulfates (10.0% w/w), dissolved H 2S/sulfides (up to 25 ppm), ammonia ( ≈381 µM), and methane (11.1 g d -1). The LH spring system contains the outlet and the outflow channel. In the initial study of the LH channel sediment, the results determined the microbial abundance by using fluorescent microscopy in the channel sediment; also, the study characterized the cultured representatives and confirmed that most of these isolates are halotolerant and psychrotolerant microorganisms. The mineralization assays on the

LH channel sediment revealed that the heterotrophic microorganisms remained activity down to -20°C. To determine the total microbial communities inhabiting the LH spring system, the study demonstrated the microbial 16S rRNA genes and the active 16S rDNA profiles for different sampling locations, including the outlet, channel and the adjacent tundra. We identified that the Bacteria from the five phyla ( Bacteroidetes , Proteobacteria ,

Actinobacteria , Firmicutes , and Cyanobacteria ) were the dominant bacterial groups at the

LH spring system. In the archaeal communities, microorganisms affiliated with three phyla (Euryarchaeota , Crenarchaeota , and Thaumarchaeota ) were identified. To determine its total functional and genetic potential, we performed metagenomic analysis of the LH spring outlet microbial community. Reconstruction of the enzyme pathways responsible for bacterial nitrification/denitrification/ammonification and sulfate reduction appeared nearly complete in the metagenomic dataset. Stress-response genes for adapting to cold, osmotic stress, and oxidative stress were also abundant in the metagenome. vi

Comparing functional community composition of the LH spring to metagenomes from other saline/subzero environments revealed a close association between LH and another

Canadian High Arctic permafrost environment, particularly in genes related to sulfur metabolism and dormancy. To identify the abundance and the presence of the featured genes ( amo A and hcd ) of Thaumarchaea at the LH spring system, we performed qPCR to assess their abundance. A phylogenetic analysis was performed using the putative amino acid sequences of these genes to identify their phylogenetic affiliation. The copy numbers of Thaumarchaeal amo A and hcd genes in LH channel sediment and the adjacent tundra were roughly 10 to a hundred-folds less than those reported in other environments. The phylogenetic tree of amo A showed similar patterns of grouping as the analysis done by

16S rRNA. This thesis demonstrates the microbial ecology, diversity and activity at the

LH spring system and provides knowledge for the microbiology studies on cryo- and hypersaline environments.

vii

RÉSUMÉ

La source de Lost Hammer (LH), située sur l’ile d’Axel Heiberg dans le Grand

Nord Canadien, est la source la plus froide et la plus salée découverte à ce jour sur terre.

Elle est caractérisée par des décharges pérennes de températures sous zéro (-5°C), par ses conditions hyper salines (salinité de 24%), réductrices ( ≈-165 mV), microoxiques, et oligotrophiques. Elle est aussi riche en sulfates (10.0% w/w), en H 2S/sulfites dissouts, en ammoniac ( ≈381 µM), et en methane (11.1 g d -1). Le système de la Source de LH contient la sortie et le canal de sortie. L’étude originale des sédiments du canal détermina l’abondance de microbes avec des techniques de microscopie fluorescente. L’étude caractérisa aussi les cultures représentatives et confirma que la majorité des isolats sont des microorganismes halotolérants et psychrotolérants. Les tests de minéralisation des sédiments du canal de LH ont révélés que les microorganismes hétérotrophes restent actifs jusqu’à -20°C. Pour determiner la communauté totale vivant dans le système de la source de LH, l’étude investigua les profils de l’ARNr 16S ainsi que l’ARNr 16S actif microbien pour différents endroits d’échantillonnage, incluant la sortie, le canal, et la toundra adjacente. Nous avons identifié que les bactéries de cinq phylums ( Bacteroidetes, protéobactéries, Actinobactéries, Firmicutes et Cyanobactéries ) étaient les groupes de bactéries dominantes dans ce système. Dans les communautés archées, des microorganismes affiliés avec trois phylums ( Euryarchaeota, Crenarchaeota et

Thaumarchaeota ) ont été identifiés. Pour déterminer son potentiel fonctionel et génétique total, nous avons performé l’analyse métagénomique de la communauté microbienne de la sortie de la source de LH. La reconstruction des voies enzymatiques responsables pour la nitrification/dénitrification/ammonification bactériennes et pour la réduction du sulfate apparut presque complète dans la banque de donnés métagnénomique. Les gènes de viii réponse au stress pour l'adaptation au froid, pour les chocs osmotiques et pour le stress oxydatif étaient aussi abondants dans le métagénome. En comparant la composition des communautés fonctionnelles du métagénome de la Source de LH, avec d’autres environnements salins et sous zéro, a révélé une association entre LH et un autre environnement du permafrost du Grand Nord Canadien, particulièrement dans les gènes reliés au métabolisme de sulfure et de dormance. Pour identifier l’abondance et la présence de gènes d’intérêt ( amoA et hcd ) de Thaumarchaea dans le système, nous avons performé les expériences qPCR. Une analyse phylogénique a aussi été faite pour identifier leur affiliation phylogénique en utilisant les séquences probables d’acides aminés de ces gènes. Le nombre de copies des gènes amoA et hcd de Thaumarchaea dans les sédiments du canal et dans la toundra adjacente étaient environ dix à cent fois moins que ceux rapportés dans d’autres environnements. L’arbre phylogénique de amoA a démontré des motifs similaires de regroupement à ceux de l’analyse faite avec r16S rADN. Cette thèse démontre l’écologie microbienne, la diversité et l’activité du système de la Source de LH, et apporte un savoir pour les études de microbiologie sur des environnements froids et hypersalins.

CONTRIBUTIONS TO KNOWLEDGE

The work presented in this thesis contributes to the advancement of knowledge in several ways: 1. The study contributes to the knowledge of the microbial ecology of Polar hypersaline

spring systems, which is a rarely-studied field of environmental microbiology.

2. This is the first study on Thaumarchaea inhabiting subzero hypersaline environment.

The study confirmed the presence of its amo A, hcd , and 16S rDNA gene sequences in

the hypersaline environment. Based on the 16S cDNA pyrosequencing library

analyses, Thaumarchaea was active in the hypersaline spring system.

3. Bulk heterotrophic microbial activity was detected at -20°C using mineralization

assays in this study. To date, it is the lowest temperature record of the evidence of

microbial activity of hypersaline environments.

4. This is the first study to present sequences of a group of thermophilic archaea,

Thermoprotei , detected in the active 16S cDNA pyrosequencing library of a perennial

subzero environment.

5. This study presents the first metagenome of a hypersaline subzero spring, Lost

Hammer Spring. We analyzed the genetic functional potential based on the

metagenome, which provides a comparable genetic database for similar microbial

studies.

LIST OF TABLES Table 2-1. Physical and geochemical characteristics for Lost Hammer (LH) Spring outlet and channel ...... 53 Table 2-2. Carbon and nitrogen analyses for LH Spring outlet and channel ..... 54 Table 2-3. CO2 and CH4 sediment concentrations and fluxes from LH Spring outlet and channel ...... 54 Table 2-4. Characteristics of 22 bacterial strains isolated from LH channel sediments ...... 55 Table 2-5. Summary of the range of statistics and indices for the 16S rRNA gene clone libraries of LH channel and outlet sediments ...... 56 Table 3-1. The statistical analyses of LH metagenome ...... 93 Table 3-2. The composition of organisms detected in the LH metagenome ...... 94 Table 3-3. Numbers of different gene variants retrieved in the LH metagenomic data sets for different functions ...... 95 Table 4-1. Geochemical measurement of the LH channel sediments and the adjacent tundra ...... 136 Table 4-2. Statistics and the indices of richness and diversity of the libraries. 137 Table 4-3. The F and P values of Permanova analyses for the samples ...... 138 Table 4-4. Unifrac distance ...... 139 Table 4-5. The comparison of the primer pairs used in the Lay et al. 2012 study and the present study ...... 128

LIST OF FIGURES Fig. 2-1. The images of LH channel ...... 57 Fig. 2-2. Phylogenetic composition of sequences ...... 58 Fig. 2-3. Phylogenetic relationships of representative bacterial 16S rRNA gene sequences obtained from the LH Spring channel clone libraries and strains ...... 59 Fig. 2-4. Phylogenetic relationships of the archaeal 16S rRNA gene sequences obtained from the LH channel clone libraries ...... 60 Fig. 2-5. Mineralization assays of [1-14C] acetate in LH channel sediment microcosms at different temperatures...... 61 Fig. 3-1. Phylogenetic profiles for key enzymes ...... 96 Fig. 3-2. Functional community composition of the LH spring sediment and other extremely cold or saline environments ...... 97 Fig. 3-3. The proportions of different clades ...... 98 Fig. 4-1. PCoA analyses for the microbial compositions ...... 129 Fig. 4-2. Summer 16S rDNA libraries of the channel and tundra...... 130 Fig. 4-3. Winter 16S rDNA libraries of the channel and tundra ...... 131 Fig. 4-4. Maximum-likelihood trees constructed by partial putative amino acid sequences ...... 132 Fig. 4-5. The copy numbers of Thaumarchaeal genes ...... 133

xii

LIST OF ABBREVIATIONS

AHI Axel Heiberg Island amo A Ammonia monooxygenase subunit A gene ANME Anaerobic methane oxidizing archaea AOA Ammonia oxidizing archaea AOB Ammonia oxidizing bacteria

Aw Water activity BLAST Basic Local Alignment Search Tool CARD-FISH Catalyzed reporter deposition fluorescent in situ hybridization CFU Colony formation unit DO Dissolved oxygen content hcd 4-hydroxybutyryl-CoA dehydratase gene LH Lost Hammer Spring MG-RAST Metagenomics Rapid Annotation using Subsystem Technology ORP Redox potential OTU Operational taxonomy unit PCR Polymerase chain reaction qPCR Quantitative polymerase chain reaction RDP Ribosomal Database Project TDS Total dissolved solids

xiii

CHAPTER 1

Introduction and Literature Review

1.1 Introduction

The Lost Hammer (LH) spring system is located on Axel Heiberg Island in the Canadian High Arctic. It is famous for its hypersalinity (~ 25% of salts, mainly sulfate salts) and perennial subzero temperatures at the spring outlet (~

-5°C) (Niederberger et al., 2010). Along with these two main properties, it is also a methane seep (~50% of gas discharged from the sediments) (Niederberger et al.,

2010), with relatively high content of ammonia present in the spring water (381

µM) (Lay et al., 2012). The spring outlet and the channel differ mainly in their reducing and less reducing waters (ORP = -187.4 to -154.0 mV in the outlet and

-29.9 to 125.5 mV in the channel) (Lay et al., 2012), as well as in the stability of their environments over the years, the former being more constantly stable than the latter in terms of temperatures (perennial subzero, oxygen content, salinity and location). Three microbiological studies of the LH system have been published thus far (Lay et al., 2012; Lay et al., 2013; Niederberger et al., 2010); two of them are included in this thesis, approaching the LH system from different perspectives.

To support the basis of these studies, the definition of the background environmental concepts of degree of salinity, terrestrial saline water bodies, and water activity will be introduced in this chapter, along with a discussion of previous microbiological studies on polar saline water bodies, including polar lakes and polar saline springs. Since salinity and low/subzero temperatures are the 1

two main environmental properties limiting microbial communities in these environments, we present microbial adaptations to low temperatures and hypersalinity in terms of DNA, RNA and protein, as well as the strategies microorganisms employ to deal with hypersaline environments. This information, along with the discussion of the industrial and astrobiological applications derived from studying microorganisms from saline and cold environments, mentioned in the introduction should be sufficient for the reader to understand the subseuquent chapters.

1.2 Terrestrial saline water body ecosystems in Polar regions

1.2.1 Definitions of terrestrial saline water bodies

Terrestrial saline water bodies exist on every continent, especially in arid areas exposed to low precipitation and high evaporation rates. The Caspian Sea and Black Sea are the largest saline water bodies in Asia, while the Dead Sea is well-known for its hypersalinity (with about 27 to 34% of salts) (Oren, 2002b). In

North America, the Great Salt Lake is known for its effect on weather, and Mono

Lake is known for the controversial study of the bacteria with arsenic-based DNA backbones (Wolfe-Simon et al., 2011). In Africa, Lake Retba in Senegal is famous for its pink color due to the rich community of phototrophic microorganisms present in the water (Sime-Ngando et al., 2011). In Australia, numerous saline lakes are present in the arid inland area. Terrestrial saline water bodies also include saline lakes, saline ponds, saline springs and salt marshes. Terrestrial saline water bodies occur mainly in arid areas, but these can occur under diverse temperature regimes. Their common feature is high salinity (>5%). They provide 2

habitats and shelters for the life forms inhabiting such areas; on the other hand, they also limit and shape the biodiversity within them. Understanding the relationship of the living organisms and the environments of these saline water bodies broadens our knowledge in the fields of biology and ecology. However, since salinity is the most common feature of these water bodies, the classification terminology for these saline water bodies must first be established.

Several different saline water classification systems, based on different salinity criteria have been proposed by limnologists and geologists since 1926.

Zoologists defined “brackish” water as that present in water bodies whose salinity ranged between that of fresh water and that of sea water. However, this term was ambiguous and not sufficient to describe saline water bodies. In addition, the maximum salinity of fresh water is clearly defined, either 0.05% (w/v) or 0.1%

(w/v), occasionally up to 0.3% (w/v), but the ranges of different degrees of salinity vary. Every classification system is based on the specific purpose of the particular study. Among these classification systems, that of Hammer’s, adjusted from Beadle’s, is widely acceptable in biology (Hammer, 1986). He defined that fresh water had a salinity of less than 0.05% (w/v), and that saline waters bore a salinity equal or in excess of 0.3% (w/v). Water with a salinity level between 0.05 and 0.3% is considered ‘subsaline’ water. Thus, according to Hammer (1986), there are three major groups into which saline water bodies can be categorized: hyposaline (0.3-2%), mesosaline (2-5%), and hypersaline ( ≥5%). As a basis for comparison, sea water salinity is approximately 3%, in the mesosaline range. The definition of different halophilic microorganisms based on the tolerance of salt concentrations has two main categories, which are moderate halophiles and 3

extreme halophiles (Rodriguez-Valera et al., 1981). The moderate halophiles are able to grow on the media with 3 – 15% of NaCl in the media (about 0.5 to 2.5 M); microorganisms, which are able to grow on media with 15 – 30% (about 2.5 to 5.2

M), are considered as extreme halophiles (Rodriguez-Valera et al., 1981).

1.2.2 Saline lakes in the Polar regions and the microbiology studies on them

Saline lakes and ponds are prevalently distributed in the Polar Regions, including saline lakes (No name recorded) in the North Great Plain of Greenland

(Hansen, 1969; Ryves et al., 2006), Lake Garrow and Lake Sophia in Canadian

High Arctic (Ouellet et al., 1987; Ouellet et al., 1989), and Levinson-Lessing Lake in Siberia (Boike et al., 1998), as well as the Vestfold Hill lakes [Deep Lake

(Bowman et al., 2000a; Ferris and Burton, 1988), Ekho Lake (Bowman et al.,

2000a; Labrenz et al., 1998), Organic Lake (Bowman et al., 2000a), Ace Lake

(Lauro et al., 2011; Ng et al., 2010), Burton Lake, Clear Lake, Pendant Lake,

Scale Lake (Bowman et al., 2000b)], Syowa oasis lakes [ Lake Nurume, Lake

Suribati, Lake Hunazoko (Tominaga and Fukui, 1981)], McMurdo Dry Valley

[Vida Lake (Mosier et al., 2007)], and Wright Valley [Don Juan Pond (Samarkin et al., 2010; Siegel et al., 1983)] in Antarctica. These saline water bodies are mostly meromictic (non-mixing) and relatively stable. Thus, interchangeable materials can diffuse from the adjacent soils or be carried by seasonal streams from distant sources. Although many saline lakes have been recognized in the

Arctic, microbiological surveys in Antarctica are more complete than those undertaken in the Arctic. The salinity of these polar lakes ranges from 1.4% (Clear

Lake) to 40% (Don Juan Pond) (Bowman et al., 2000b; Samarkin et al., 2010); 4

temperatures and limnology vary according to the geological properties of each location.

Microbial research in polar lakes revealed many facets of microbial diversity, ecology and activities in these locations. Methods applied to polar lakes in Antarctica are diverse, but basically, non-cultural methods (clone library,

DGGE or microscopy) and cultural methods (enrichment or isolation) have both been applied in such studies; however, the detection of in situ activity was not performed in the saline lakes. Given the methodological limitations, it is worth keeping in mind that these studies only reveal partial microbial communities from the habitats. Although the saline water beneath Antarctic Lake Vida has been known to researchers, initial microbiological studies of Lake Vida focused on the thick layer of ice on top of the lake. More recently, microbial activity was detected using cDNA libraries in the -13°C briny water in the ice of Lake Vida (Murray et al., 2012). The Lake Vida briny microbiota was isolated in the ice without external source of energy; the energy source for microorganisms was originated from the water-rock reactions based on the geochemical analyses of the environment

(Murray et al., 2012).

Several studies examine the microbial diversity by culture-independent methods for the Vestfold Hill lakes and the Syowa Oasis lakes (Bowman et al.,

2000a; Bowman et al., 2000b; Kurosawa et al., 2010). In comparing the microbial diversity of Ekho Lake, Organic Lake and Deep Lake (Bowman et al., 2000a), the diversity profile shifted noticeably according to the differences in salinity. The most saline (32%), Deep Lake was dominated by halophilic archaea with a lower microbial diversity and richness (Shannon index = 0.94 and Chao1 index = 15) 5

than the other lakes. The halophiles in Ekho lake were mostly bacteria. The community structures of the other five saline lakes, Clear Lake, Pendent Lake,

Scale Lake, Ace Lake, and Burton Lake, in Antarctica also show different patterns from low to high salinity; however, low G+C Gram positive bacteria, which includes psychrophilic, fermentative, and acetogenic species, are the commonly dominant (>31% in each lake) microorganisms in these saline lakes.

In the Syowa Oasis area in the Antarctica, the survey for archaeal and bacterial compositions in anoxic sediments was based on 16S rDNA clone libraries from Lake Nurume. The archaea in the archaeal clone library was composed of only two phylotypes, marine benthic group D (MBG-D; 93%) and another unknown euryarchaea. The most abundant bacteria are

Alphaproteobacteri a; however, they represented only 20% of total bacterial clones, which means that, although it was not demonstrated in the report, the evenness of species of the bacteria in Lake Nurume should be high. In other saline lakes of

Syowa Oasis, the microbial diversity was not completely studied, but the DMSO utilizing bacteria, Marinobacter , were detected from a 16S rDNA clone library from Lake Suribati (Matsuzaki et al., 2006). As Lake Suribati is a meromictic lake with different salinity (increasing by the depth) in each water layer, the bacterial halophiles, Halomonas , Idiomarina and Marinobacter , were detected in different distributions in each water layer (Naganuma et al., 2005).

The presence of microbial life in Don Juan Pond, which is likely the most constantly extreme place on Earth (~40% salinity and temperatures as low as

-52°C), is still under debate. Three studies reported cultivated microorganisms

(Meyer et al., 1962), observation of microflora (by microscopy) (Siegel et al., 6

1979), and an algal mat (Siegel et al., 1983) on Don Juan Pond. More recently, no microbial activity was detected, and only abiotic N2O emissions were observed

(Samarkin et al., 2010). Lack of presence of microorganisms might be due to high

CaCl 2 concentration (> 470 g/L), which lower down the Aw to 0.45 and represses the growth of any possible life (Oren, 2013). Several saline lakes also exist in the

Arctic, including saline lakes in Greenland (Hansen, 1969; Ryves et al., 2006),

Lake Garrow, Lake Sophia in the Canadian High Arctic (Ouellet et al., 1987;

Ouellet et al., 1989), and Levinson-Lessing Lake in Arctic Siberia (Boike et al.,

1998); Scientific studies of these Arctic saline lakes have, in most cases, been conducted for geochemical analyses. In contrast to the saline lakes in the Antarctic, studies of the microbiology of these Arctic saline lakes are conspicuous by absence.

1.2.3 Saline springs in Polar regions

Polar saline springs are defined as salty water bodies that have a continuous water discharge from permafrost, with adjacent glaciers or some other water bodies, such as lakes or underground waters as the water source. Saline spring systems include outflow channels allowing water to flow out to rivers or the sea. So far, with some from the Arctic, and only one from the Antarctic, few reports or studies of Polar region saline springs have been published. Some Arctic saline springs include the Gypsum Hill (GH) Springs, Colour Peak (CP) Springs

(Perreault et al., 2007; Perreault et al., 2008), and Lost Hammer (LH) Spring (Lay et al., 2012; Lay et al., 2013; Niederberger et al., 2010) on Axel Heiberg (AH)

Island; ten sulfur (TS) springs (they are not named formally) on Ellesmere (EL) 7

Island (Grasby et al., 2003); as well as Fisosen and Trollosen springs on Svalbard

(SB) Island (Lauritzen and Bottrell, 1994; Reigstad et al., 2011). The Antarctic

Blood Falls in McMurdo Dry Valley is visually astonishing (Bakermans, 2008;

Mikucki et al., 2004; Mikucki et al., 2009; Mikucki and Priscu, 2007) because of its rusty red color. All of these springs can be considered as sulfate/sulfite/sulfur springs due to their richness in substrates, such as sulfate, sulfide or sulfur, related to sulfur cycle. In addition, methane or methane-related activity evidence has been observed in all these springs (except TS springs and the Blood Falls).

The microbiology studies on polar saline springs focus primarily on the

Arctic. The methods applied to microbiology of polar saline springs are quite diverse. Besides culture-independent and -dependent methods, some methods of measuring microbial activity in situ have been applied. The use of sulfur stable isotopes is one such method. This method, namely measuring the depletion of 34 S

(23.6 to 29.5 ‰), was performed at Trollosen, Fisosen and TS springs and indicated that bacterial sulfate reduction occurred (Grasby et al., 2003; Lauritzen and Bottrell, 1994). In addition, a stable isotope carbon study, which found a

13 δ CCH4 value of 71.2‰ indicated in situ microbial methanogenesis in the GH springs (Perreault et al., 2008) Radioactive mineralization assays were applied to the LH spring outlet and channel sediments using 14 C-labeled glucose or acetate as carbon substrate, which also indicated heterotrophic microbial activities in the

LH sediments (0.34 % of cumulative percentage of recovery of mineralization at

-10°C for the outlet sediment with 14 C-labeled glucose (Steven et al., 2007b), and

0.17% of cumulative percentage of recovery of mineralization at -20°C for the channel sediment with 14 C-labeled acetate) (Lay et al., 2012). The microbial 8

activity of Blood Falls was measured by 3H-labeled thymidine and leucine for doubling time of microorganisms under temperatures of 0°C to 30°C. The doubling time ranged from 37 to 54 days (Mikucki et al., 2004). Unfortunately, there is not yet a subzero incubation study for Blood Falls.

To identify the microbial communities in these springs, non-culture methods, i.e . 16S rRNA clone library or DGGE, were applied in some studies.

Betaproteobacteria and Gammaproteobacteria are commonly present in these springs as part of the bacterial community, and perhaps it is because these classes include many halophilic, sulfur-related compound metabolizers, and methanotrophs/methylotrophs (Lay et al., 2012; Niederberger et al., 2010;

Perreault et al., 2007; Perreault et al., 2008), most of which are better adapted to cold salty environments. Deltaproteobacteria and Epsilonproteobacteria are detected usually in these springs because of the sulfur-cycle related metabolizers

(Lay et al., 2012; Niederberger et al., 2010; Perreault et al., 2007; Perreault et al.,

2008). In addition, Bacteroidetes and Firmicutes are common in these springs as well. The special methane oxidizers, Verrucomicrobia, were detected in the LH channel and GH springs (Lay et al., 2012; Perreault et al., 2007). However, the archaeal communities are more diverse than the bacterial ones. Halophiles are common in the AH springs (Lay et al., 2012; Niederberger et al., 2010; Perreault et al., 2007; Perreault et al., 2008), which is related to the salinity, but evidence is lacking regarding their presence in the SB and EL springs. The anaerobic methane oxidizing archaea (AMNE-1) were detected in the LH spring outlet and Fisosen spring (Niederberger et al., 2010; Reigstad et al., 2011). The presence of ANME-1 might be correlated to the methane emissions at LH outlet (Niederberger et al., 9

2010). In addition, Thaumarchaea, which is involved in inorganic carbon fixation and ammonia oxidation, were identified in the LH spring channel (as well as in the outlet, see chapter 4), Fisosen, and Trollosen springs (Lay et al., 2012;

Niederberger et al., 2010; Reigstad et al., 2011). In the studies regarding the springs on AH, heterotrophic and autotrophic bacteria were cultured from the spring sediments. Most of the isolates from LH spring areas could deal with polyextreme growth conditions, i.e ., growth at -5°C and relatively high salinity (>

10%) (Lay et al., 2012; Niederberger et al., 2010). However, isolates from GH springs have similar patterns of their saline tolerance as the isolates from the LH spring system but lack of information to the subzero culture (Perreault et al.,

2008). In GH spring, a chemolithoautotrophic streamer, Thiomicrosipra sp., was identified as a sulfide/thiosulfate oxidizer and carbon fixer (Niederberger et al.,

2010). The ability of GH Thiomicrosipra sp. for oxidizing sulfide and utilizing

2- 14- inorganic carbon were examined using S2O3 comsuption and C bicarbonate uptake methods (Niederberger et al., 2010).

The only reported saline spring in the Antarctic is the Blood Falls (~1,375 mM Cl –), which is rich in sulfate (50 mM) and iron [(3.45 mM, >97% is Fe (II)]

(Mikucki et al., 2009). The Blood Falls is the outflow of an unknown and deep subglacial water system under the Tayler Glacier in Antarctica. The oxidized iron

(Fe III) converted from Fe II in the water caused the rusty red color on the channel of the falls. The water of the falls subsequently flows into the adjacent

Lake Bonne (Mikucki et al., 2009). The microbial community of Blood Falls is mainly composed by Gammaproteobacteria , especially Thiomicrospira arctica

(46%) (Mikucki and Priscu, 2007). Thiomicrospira arctica , also cultivated, is a 10

sulfur-oxidizing species, and reflects the relatively high sulfate contents in Blood

Falls. Betaproteobacteria , Deltaproteobacteria and Bacteroidetes were detected and cultivated, including sulfate reducing bacteria and halophilic bacteria

(Mikucki and Priscu, 2007). However, the archaeal clone library was not done yet for Blood Falls.

1.3 Challenges to microbial life in Polar saline water bodies

1.3.1 The availability of liquid water in cryoenvironments

Liquid water is the basic element for organisms to maintain their metabolism and growth. Water provides an environment for biochemical and enzymatic reactions, as well as being a solvent for dissolving molecules, such that they are able to interact with bio-molecules. In cold environments, especially subzero environments, liquid water is not always accessible. In these cryoenvironments, once the water freezes, the microorganisms have difficulty in obtaining liquid water to utilize for maintaining their viability. The most common reagents in nature to drop freezing points are solutes. Freezing points differ when water contains different concentrations of solutes, e.g., pure water freezes at 0°C; sea water (3.5% salinity) freezes at -1.9°C, and a saturated NaCl solution (6.15M) drops the freezing point down to -21°C (Bakermans, 2008). Liquid water in cryoenvironments are not only present in bulk water reservoirs, such as saline lakes and springs, they are sometimes preserved in some microhabitats, e. g., brine channels in sea ice, glacier, permafrost, or are present as a thin layer (less than 1 µm) of water on an ordered surface due to ordering effects at subzero temperatures (Drost-Hansen, 2001). In macro- or micro- cryoenvironments, such 11

liquid water in cryoenvironments is usually maintained by high solute concentrations. Water is not always available to microorganisms even though it is in liquid form because of the effect called ‘water activity.’

Water activity is affected by multiple environmental factors. The concept of water activity (A w) was originally used for food science, and serves as a useful parameter to predict the possible microbial viability in arid, saline or cold environments. The Aw decreases as solute concentrations increase or temperatures decrease. The Aw is determined by the vapour pressure of the solution divided by the pure water at the same temperature. The parameter of Aw has been studied for testing harsh conditions in terms of the limit of life, thus serving as an indicator for the living appropriateness for the microorganisms in environments containing water. So far, A w = 0.62 is the minimum value at which a microorganism (yeast) can reproduce (Beaty et al., 2006). The high salinity, extreme temperatures, or other factors introduced by the environment are able to perturb the hydration structure of water molecules and limit the activities of the biomolecules, e.g. proteins and enzymes, which may reduce the interaction with the water. The ionic interactions with water molecules may inhibit microorganisms from obtaining water in such environments, even though the water is still in liquid form. Thus, microbial adaptations to cryoenvironments must include the ability to resist low temperatures and high solute concentrations.

1.3.2 The adaptation of microorganisms to cryoenvironments

The two main critical problems with which microorganisms cope in cryoenvironments are subzero temperatures and salinity, both of which repress 12

water activity. Consequently, microorganisms must make an effort to use available water. For cold adaptation, Bakermans et al. (2009) provided several factors facing microbial cells: 1) control of molecular motion inside/outside the cells to maintain the normal processes of metabolism, 2) efficiently using resources for metabolism, due to the difficulty of energy generation, and 3) expression of temperature-adapted enzyme alleles to adapt to the cold temperature environment.

Similarly, Oren (2011) provided several points regarding life in a high salinity environment: 1) environments are more energy-consuming for microorganisms to inhabit, 2) bioenergetic constraints are the main factors to determine dissimilatory processes, and 3) the total energy-generation and the mode of osmotic adaptation are the main factors that limit growth of microorganisms in saline environments.

Terrestrial saline water bodies represent environments combining both these parallel and simultaneous challenges for microorganisms. In the following sections, cold and saline adaptations of microbial life will be separately introduced and discussed.

1.3.2.1 Cold adaptations of microorganisms

At the molecular level, the structures of biomolecules show some adaptive differences between cold or saline environments and mild-temperature and non-saline environments. Cold adaptations happen in several different biomolecules in microbial cells, including nucleic acids, proteins, and membrane lipids. The rationale for modification is to make the bio-molecular structure flexible and maintain functions at cold temperatures. Cold temperatures stabilize

the structures of DNA, RNA, proteins and membrane lipids (Bakermans, 2012;

D'Amico et al., 2006). In this situation, these biomolecules stop functioning and the whole metabolisms of microbial cells will be affected. For example, cells will not function when more than 50% of membrane lipids are solidified due to cold temperatures (Jackson and Cronan, 1978; Melchior, 1982). These biomolecules are necessary to sustain microbial viability; therefore, maintaining their functioning at cold temperatures is necessary for microorganisms inhabiting cold environments. Without these molecules, no biochemical reaction can occur and the microorganisms cannot survive.

The adaptation of nucleic acids of microorganisms are not only restricted to differences in G+C content in DNA, though a lower G+C content would lower hydrogen bonding in DNA or RNA molecules and allow them to more easily unwind. In terms of the content of other nucleic acids, Khachane et al. discovered that uracil content inversely corresponds with the optimum growth temperature of psychrophiles (Khachane et al., 2005). An increase (40 to 47% higher than the average) in dihydrouridine content (dihydrouridine is reduced from uridine by tRNA-dihydrouridine synthase) of psychrophile tRNAs observed in both bacteria and archaea, indicates a cold adaptations at the RNA level (Dalluge et al., 1997;

Noon et al., 2003). Protein adaptation to cold environments, include fewer hydrogen bonds, reduced Arg/Lys ratio, decreased prolines in loops, fewer salt bridges, reduced ion pairs, higher accessibility to active sites, increased hydrophobicity in enzyme or protein cores, increased interactions with solvent, and fewer disulfide bonds. All of these features contribute to an internal protein structural flexibility but also result in decreased stability of higher temperatures 14

(Bowman et al., 2000b; D'Amico et al., 2006; Feller, 2003; Feller and Gerday,

2003; Mykytczuk et al., 2013).

Membrane lipids may solidify at low temperatures. Once the membrane is solid, microorganisms cannot maintain proper membrane functions. Maintaining fluidity of the membrane is necessary for microorganisms inhabiting cryoenvironments. The rationale for keeping the fluidity of the membranes is to loosen the lipid structure. Several strategies are observed for maintaining fluidity of microbial cell membranes, including increasing unsaturated fatty acid content, methyl branching modification on membrane lipids (bacteria), and anteiso /iso -branched ratio of membrane lipids (bacteria); decreasing in average fatty acid chain length (all kinds of microbes) and sterol/phospholipid ratio

(eukaryotes) (Nichols et al., 1997; Russell, 1990, 1997).

In environments of subzero temperatures, reduced water activity is attributable to crystallization of water molecules. Cryoprotectants, which are compatible solutes, including sugars, amino acids alcohols or cryoprotective proteins, may depress non-specifically the freezing points of water and increase the water availability to microorganisms (Bakermans, 2012; Chin et al., 2010).

Psychrophiles commonly use these compounds, especially glycine betaine, glycerol, trehalose, sucrose, proline and dimethylsufoniopropionate (Welsh, 2000), to prevent ice formation inside cells. The cryoprotectants also help to remain a water layer on the protein surface to maintain their structures and functions

(Welsh, 2000). Microorganisms can maintain their metabolisms at low temperatures. Some of these compatible solutes, e.g. glycine betaine, also balance osmotic pressures under hypersaline environments. 15

Cryoprotective proteins are bio-molecules that serve to maintain liquid water inside/outside microbial cells. Cryoprotective proteins include anti-nucleating, ice-binding and anti-freezing proteins. Anti-nucleating proteins may inhibit the formation of ice nuclei and cooperate with other anti-nucleating materials, such as polyglycerol, to decrease the freezing point of water

(Bakermans, 2012). Ice-binding and anti-freezing proteins are able to bind ice crystals and prevent them from growing. These proteins have been detected in several microorganisms, such as Pseudomonas putida GR12-2, Antarctic isolate

Marinomonas primoryensis , Micrococcus , Rhodococcus , Sphingomonas ,

Halomonas , Pseudoalteromonas , and Psychrobacter (Gilbert et al., 2005;

Kawahara, 2008; Xu et al., 1998).

1.3.2.2 Saline adaptation of microorganisms

Halophilc microorganisms are usually classified into two types, moderate and extreme halophiles. They are only able to grow in the environments with high concentration of salt. The moderate halophiles are able to grow on the media with

3 – 15% of NaCl in the media (about 0.5 to 2.5 M); microorganisms, which are able to grow on media with 15 – 30% (about 2.5 to 5.2 M), are considered as extreme halophiles (Rodriguez-Valera et al., 1981). Some microorganisms are defined as halotolerant microorganisms, which can grow also with the presence of high salt but the optimal growth concentration of salt is relatively low. Living in high salt concentrations, halophiles adapt to the environments in many molecular levels. Halophilic adaptations occur at the physiological, genomic and proteomic

levels in microorganisms inhabiting saline/hypersaline environments. At the genomic level, halophilic microorganisms seem to have special codon usage, with

GA/TC, AC/GT and CG appearing at a greater than normal frequency in the first two positions of codons (Paul et al., 2008). The GA, AC, and GT pairs are linked especially to translation of aspartate, glutamate, threonine and valine residues

(Paul et al., 2008). Protein features of halophiles include a greater proportion of aspartate and glutamate residues (Paul et al., 2008), a higher valine residue content, indicative of an increased propensity towards coiled structures, (Li and

Hermans, 1993), as well as low hydrophobicity, low propensity of helix formation, under representation of cysteine, low lysine content, and decreased aliphatic residues (Madern et al., 2000; Paul et al., 2008). Although some of these features appear to conflict, for example, valine is a hydropholic amino acid but in general, the halophilic proteins have low hydrophobicity, the presence of these features also depends on the locations and functions of these proteins/enzymes in cells.

Besides the modification of biomolecules, microorganisms inhabiting saline environments have two strategies to balance the osmotic stress inside and outside the cell, i.e ., salt-in vs. compatible-solute strategies (McGenity and Oren, 2012).

Salt-in strategy only occurs in the halophilic archaeal family “Halobacteriaceae ”, and two bacterial groups, Salinibacter spp. and members of Halanaerobiales

(Oren, 2002a; Oren and Mana, 2002). They pump KCl molecules into cells to balance the osmotic pressure of their environments. In this case, the protein surfaces contain acidic and negatively charged residues, which serve to maintain a hydration layer around proteins to keep them functional. Thus, all the biomolecules in these microorganisms must be adapted to high ion concentrations 17

inside the cells. Such an adaptation limits their ability to survive at low salt environments.

The strategy of using compatible solutes for microorganisms to deal with hypersaline environments is relatively common. Compatible solutes are organic compounds, e.g ., glycine betaine, ectoine, trehalose, sarcosine, glycerol, sugars, amino acids, and some other organic molecules (Kempf and Bremer, 1998;

Mykytczuk et al., 2013), which may also serve as cryoprotectants for cold adaptation. Some of these compounds are species specific. These compounds are synthesized or taken up to balance the osmotic pressure inside cells. This approach is more energy-consuming than the salt-in strategy but allows microorganisms the relative versatility to survive in different salt concentrations

(Oren, 1999), as their protein structures do not need much modification when using this strategy. This strategy has been observed in most halotolerant bacteria, fungi, algae and methanogenic archaea (McGenity and Oren, 2012).

1.4 Applications and astrobiology aspects of the study

1.4.1 Potential applications of microorganisms from cold saline

environments

In general, studies on extremophiles may be applied to several different biotechnological orientations, such as environmental remediation, detoxification, agriculture, chemical industry, detergent/leather processing, biofuel/bioplastic production, cancer detection, biosensors, pharmaceutical industry, and biomining/biorefining (Arora and Bell, 2012). Because of cold adaptation, extremophiles in cryoenvironments play important roles for environmental 18

bioremediations. For example, hydrocarbon contamination is a pressing issue in

Arctic soil, cold marine waters, and sea ice due to mining and military activities.

It is difficult for contaminants in these areas to degrade naturally. Cold-adapted microorganisms, under proper biostimulation conditions, have the proven capacity to degrade hydrocarbon contaminants on site (Brakstad et al., 2010; Greer, 2010).

This includes the degradation of naphthalene, polyaromatic hydrocarbons (PAHs), and hexadecane at temperatures lower than 5°C. Microorganism-produced cryoprotectant materials can be isolated from the EPS (Extracellular polysaccharides substances) of psychrophiles (Boonsupthip and Lee, 2003; Marx et al., 2009). The cryoprotectants extracted from psychrophiles, such as

Pseudoalteromonas spp. from Antarctic marine and Colwellia psychrerythraea from Arctic sediment, may be applied to food production for preservation purposes (Boonsupthip and Lee, 2003; Marx et al., 2009). There are four potential fields of applications of cold adapted enzymes: 1) detergents and personal care, 2) food, pharmaceutical/chemical and cosmetic industries, 3) biofuels, and 4) molecular biology (Huston, 2008). Regarding detergents and personal care, genetically-modified cold-adapted cellulase and protease have been applied to commercial products to increase the cleansing agency of detergents under cold and moderate temperatures (Huston, 2008). In the food industry, cold-adapted proteases and lipases are applied to cheese production to accelerate maturation rates; polygalacturonases and pectate lyases are added in fruit and vegetable procedures to degrade pectin compounds at low temperatures; the addition of cold-adapted β-galactosidases to dairy products can attenuate the effects of lactose-intolerance in humans (Huston, 2008; Nakagawa et al., 2004; Shahidi and 19

Kamil, 2001). In the pharmaceutical/chemical industry, cold active lipases isolated from Candida antarctica has been found to have applications in modifying polysaccharides, and desymmetrizing intermediate products of drugs

(Suen et al., 2004). In cosmetic products, cold-adapted proteolytic enzymes can be added to gels to enhance treatment of scarring, infection, and heal wounds

(Huston, 2008). Regarding biofuel generation, cold-adapted α-amylases and glucoamylases have the potential to reduce the reaction temperatures of bio-ethanol production, i.e. , cold hydrolysis, to lower energy demands and reduce unwanted side products during starch fermentation (Lin and Tanaka, 2006).

Cold-adapted enzymes are also applied to biotechnology. An alkaline phosphatase cloned from an Antarctic psychrophilic strain, TAB5, was proven to remove 5’ phosphoryl groups from nucleic acids, which may help nucleic acid to process self-ligation under low temperatures (Rina et al., 2000).

1.4.2 Astrobiological aspects

Considering water is one of the necessary elements for life (Bartik et al.,

2010), a trace of water may indicate the presence of life. The existence of large quantities of water on Mars’ South Polar Region’s massive ice cap (Bibring et al.,

2004), suggests the possibility for extraterrestrial life. Some geological evidence shows the possibility of the current presence of water on Mars (Bibring, 2010), i.e . water frost (Carrozzo et al., 2009), newly forming evaporate deposits (Malin et al.,

2006), ancient spring remains (Allen and Oehler, 2008), hydrated minerals

(silicates and sulfates) (Bibring et al., 2005; Mustard et al., 2008), water-ice clouds (Whiteway et al., 2009), and water absorbing salts (perchlorates) (Hecht et 20

al., 2009; Zorzano et al., 2009). Water frost may imply the existence of permafrost underneath the Martian surface (Farmer and Doms, 1979). Similar geological structures or geochemical properties observed in the Polar Regions on the Earth are considered as analogous sites to Mars.

Terrestrial cold saline springs are considered to be analogous to Martian sites due to their geophysical and geochemical properties. The water of cold saline springs are discharging from the permafrost and contain high salinity. Thus, the spring water may remain unfrozen at subzero temperatures. The perennial springs on Axel Heiberg Island and Ellesmere Island are examples of Martian analogues because 1) they are rich in sulfate (AH and Ellesmere springs) which was also detected on Mars; 2) the gully structures with evaporate deposits, i.e. GH and CP springs, are similar to those on Mars; 3) the saline liquid water, which may be the type of liquid water present on Mars, decreases water’s freezing point. In addition, methane emissions were also detected on Mars in 2003 (Mumma et al., 2009).

Though the observation of methane is still under debate (Zahnle et al., 2011), the methane may be one of the potential biosignatures of Martian habitats. The methane emissions of the LH spring make it a Martian analogue site on the Earth

(Niederberger et al., 2010). Moreover, the structure of the remains of an ancient spring, showing liquid-carried deposit diffusion in Arabia Terra on Mars (Allen and Oehler, 2008), exhibits similar geological patterns as the LH spring.

Regarding other extraterrestrial bodies, two ice covered moons, Jupiter’s

Europa and Saturn’s Enceladus, are both prime targets for investigating life due to possible oceans underneath the thick ice cover (Carr et al., 1998) and water vapor plumes (Hansen et al., 2006), respectively. On Europa, the subsurface ocean may 21

sustain an environment for generating hydrogen and oxygen (Hall et al., 1995). In the water vapor plume of Enceladus, nitrogen, carbon dioxide, methane, propane, acetylene, and ammonia (Matson et al., 2007; Waite Jr et al., 2009) were detected.

Vida Lake and Lake Vostok are the astrobiological analogous sites of Europa.

These lakes have perennial ice/glacier covers on tops of lake water (Doran et al.,

2003; Jouzel et al., 1999), which are completely isolated environments protected by ice as the ocean on Europa. The LH spring system, the primary subject of this dissertation, is an analogous site for Enceladus (Lay et al., 2012). It contains gas emissions of nitrogen, carbon dioxide, and methane, as well as ammonia in the liquid. These properties are similar to the features that scientists observed from the plume of Enceladus.

1.5 Objectives

This dissertation includes three major studies of the Lost Hammer cold saline spring system in the Canadian High Arctic. The objectives were:

1) To characterize the microbial biodiversity, ecology, and activity of the

microbial communities present in runoff channel sediment of the LH spring

using culture-dependent, molecular-based (CARD-FISH—catalyzed reporter

deposition fluorescence in situ hybridization and 16S rRNA gene clone

libraries), and activity analyses. Our primary goals were to determine if the

LH channel sediments were microbially active at subzero, hypersaline

conditions (down to -20 °C) and to compare the more heterogeneous channel

features with the outlet to fully describe the range of geochemical and 22

microbial characteristics that exist within the LH spring system.

2) To map the microbial metabolic pathways driving biogeochemical cycles,

focusing on methane, ammonia, and sulfur cycling, which were expected to

play key roles in shaping LH communities based on previous investigations of

the LH system (Lay et al., 2012; Niederberger et al., 2010); to identify the

dominant genes involved in adaptations to cold and high salt concentrations

that would allow autochthonous populations to cope with the extreme natural

conditions of the site; to compare the functional potential of the LH

metagenome to metagenomes from other cold or saline environments; and to

identify the bacterial and archaeal taxa that may be active in situ .

3) To characterize the seasonal microbial components at the LH channel and the

adjacent tundra based on 16S rDNA and rRNA libraries and evaluate the

significant differences between factors of those samples, i.e., sampling

seasons, locations, and sample types, based on OTUs; to assess the amount

archaeal ammonia oxidizers (Thaumarchaea) at the LH channel and the

adjacent tundra based on the sequences of the featured functional genes of

amo A, hcd and Thaumarchaeal specific 16S rDNA sequences using qPCR;

and to compare the sequences of amo A and hcd cloned from LH channel

samples with other published sequences for identifying the relationship the

LH Thaumarchaea with the ones present in other environments.

CONNECTING TEXT

Due to the dearth of research on the LH channel sediment, we designed a preliminary microbial study on the channel sediment using culture-dependent and culture-independent methods to describe the indigenous microbial communities.

We needed to establish a background microbial profile of the channel environment for future comparative studies.

CHAPTER 2

Microbial Diversity and Activity in Hypersaline High Arctic Spring Channels

Chih-Ying Lay 1, Nadia C. S. Mykytczuk 1, Thomas D. Niederberger 2, Christine

Martineau 1,3 , Charles W. Greer 3 and Lyle G. Whyte 1

1Department of Natural Resource Sciences, McGill University, Canada 2College of Marine and Earth Studies, University of Delaware, U.S.A.

3Biotechnology Research Institute, National Research Council Canada, Montreal,

Canada

Published in: Extremophiles, March 2012. 16 (2): 177-191

CONTRIBUTION OF AUTHORS Dr. Niederberger chose the site for sampling and collected the samples for analyses. He also measured on site data. Dr. Martineau performed the measurements of CO 2 and methane concentrations. All of the rest experiments were designed and performed by myself under the consultation of Dr. Whyte and

Dr. Greer. The manuscript were written by myself and Dr. Mykytczuk. 24

ABSTRACT

Lost Hammer (LH) spring is a unique hypersaline, subzero, perennial high

Arctic spring arising through thick permafrost. In the present study, the microbial and geochemical characteristics of the LH outflow channels, which remain unfrozen at ≥-18°C and are more aerobic/less reducing than the outlet, were examined and compared to the previously characterized spring outlet. LH channel sediments contained greater microbial biomass (~100 fold) and greater microbial diversity, as reflected in the different species abundances in 16S rRNA clone libraries. Phylotypes related to methanogenesis, methanotrophy, sulfur reduction and oxidation were detected in the bacterial clone libraries while the archaeal community was dominated by phylotypes most closely related to ammonia-oxidizing Thaumarchaeota . 14 C-acetate mineralization rates in channel sediment microcosms exceeded ~30 % and ~10 % at 5°C and -5°C, respectively, but sharply decreased at -10°C ( ≤ 1%). Most bacterial isolates, ( Marinobacter,

Planococcus , and Nesterenkonia spp.), were psychrotrophic, halotolerant, and capable of growth at -5°C. Overall, the hypersaline, subzero Lost Hammer spring channel has higher microbial diversity and activity than the outlet, and supports a variety of niches in which diverse and metabolically active microbial communities exist.

2.1 Introduction

Cryoenvironments are defined as permanently frozen or subzero environments including permafrost, glaciers, ice sheets, multi-year sea ice, high- elevation Antarctic dry valleys, and glaciers as well as their associated microhabitats such as brine veins in sea ice and permafrost (Bakermans, 2008,

2012; Priscu and Christner, 2004; Steven et al., 2006; Wells and Deming, 2006).

In addition to prolonged exposure to subzero temperatures, microbial communities existing in such cryoenvironments must overcome extremely low rates of nutrient and metabolite transfer, high solute concentrations, low water activity, and potentially high background radiation (Ayala-del-Rio et al., 2010;

Bakermans, 2008; Steven et al., 2006). Nevertheless, microbial diversity, ecology and activity have been recently described in numerous cryoenvironment habitats and generally indicate that viable microbial communities consisting of Bacteria,

Archaea, viruses, and eukaryotes exist in these extreme habitats (Bakermans,

2008, 2012; Priscu and Christner, 2004; Steven et al., 2006; Wells and Deming,

2006) and are capable of both growth and metabolic activity at ambient subzero temperatures (Anesio et al., 2007; Bakermans, 2012; Bottos et al., 2008; D'Amico et al., 2006; Niederberger et al., 2010; Steven et al., 2008). Cold-adapted microorganisms inhabiting such environments exhibit a variety of modifications to their proteins, nucleic acids, and membranes, which allow them to maintain their fluidity and flexibility and associated activity at low temperatures, as well as other adaptations including cryoprotectant production, and highly efficient regulation of growth (Ayala-del-Rio et al., 2010; Bakermans, 2008). However, the 26

means by which microorganisms survive and even sustain active metabolism, despite the extreme challenges presented by these cryoenvironments, warrants further investigation. For example, it is still not clear what the cold temperature limits of microbial life are in terms of growth, metabolism/maintenance and survivability (Price and Sowers, 2004), whether the microbial communities inhabiting cryoenvironments are active microbial ecosystems or merely microbial survivors, and what contributions these micro-organisms make to global biogeochemical cycles (Bakermans, 2012; Price and Sowers, 2004; Steven et al.,

2009).

The cold saline springs on Axel Heiberg Island (AHI) in the Canadian High

Arctic are among the only known cold springs in permafrost cryoenvironments on

Earth and represent a unique opportunity for expanding our knowledge of microbial life in extreme cold environments. The microbial communities of two moderately extreme High Arctic spring systems, Gypsum Hill (GH) and Colour

Peak (CP) were found to contain active microbial communities capable of existing in an extreme environment that experiences prolonged periods of continuous light or darkness, low temperatures (-1ºC to 8ºC), and moderate salinity (~8 to 15%), and where life seems to rely on sulfur-based chemolithoautotrophy (Niederberger et al., 2009; Perreault et al., 2007; Perreault et al., 2008). These streams occur in an area with an average annual air temperature of -15ºC and with air temperatures below -40ºC common during the winter months.

We recently described the microbial communities inhabiting in Lost Hammer

(LH) spring, a hypersaline (24 % salinity), subzero (-5°C) perennial spring that is the only known terrestrial CH 4 seep in a cryoenvironment on Earth arising 27

through thick permafrost (Niederberger et al., 2010). Our initial microbial characterization of LH spring sediments revealed a novel low diversity, low biomass microbial community capable of metabolic activity at in situ subzero, saline conditions. Molecular analyses (bacterial and archaeal 16S rRNA gene clone libraries, CARD-FISH) detected Bacteria phylotypes related to microorganisms previously recovered from cold, saline habitats. Archaeal phylotypes were related to signatures from hypersaline deep-sea methane-seep sediments and were dominated by the anaerobic methane group 1a (ANME-1a) clade of anaerobic methane oxidizing archaea, indicating that the thermogenic methane exsolving from the Lost Hammer spring outlet may act as an energy and carbon source for sustaining anaerobic oxidation of methane-based microbial metabolism under ambient hypersaline, subzero conditions (Niederberger et al.,

2010).

The Axel Heiberg springs are regarded as Martian analogue sites due to their unique geology, climate and geomorphology which mimic conditions once existing, or currently existing, on Mars (Pollard et al., 2009). For example, a gully which formed during the past decade on Mars provides compelling evidence that liquid water (or brine) may exist on Mars (Malin et al., 2006), while the trace amounts of methane in the Mars atmosphere (Formisano et al., 2004) may originate from localized ‘hot spots’ or ‘plumes’ of methane arising from the frozen terrestrial Martian surface (Mumma et al., 2009). The origin of Martian atmospheric methane is under extensive debate (Lefevre and Forget, 2009) and could be attributable to either geological or biological (methanogenesis) sources.

In 2005, during our first winter expedition to the LH spring site, we 28

discovered that the outflow spring channels downstream from the LH spring outlet remained unfrozen, due to high salt concentration in the sediment pore water, and contained evidence of active microbial activity in the form of gas bubbles exsolving from the channel sediments despite ambient sediment temperatures as low as -18°C (Fig. 2-1a). The objectives of the present work were to characterize the microbial biodiversity, ecology, and activity of the microbial communities present in runoff channel sediment of the LH spring using culture–dependent, molecular-based (CARD-FISH – catalyzed reporter deposition fluorescence in situ hybridization and 16S rRNA gene clone libraries), and activity analyses. Our primary goals were to determine if the LH channel sediments were microbially active at subzero, hypersaline conditions (down to

-20°C) and to compare the channel features with the outlet to verify the range of geochemical and microbial characteristics that exist within the LH spring system.

2.2 Materials and Methods

2.2.1 Sample site description and geochemical analyses

A total of three sediment samples (C1-C3) were collected from the outflow channel (79°04.608N; 90°12.739W) of Lost Hammer Spring (Figure 2-1.a): sediment samples C1 and C2 were collected on April 30 th , 2008; sediment sample

C3 was collected on May 4 th , 2007. Samples were obtained using sterile scoopula and material down to 5 cm depth was collected. Sediments were placed into 500 mL sterile sample bottles and the remaining volume filled with

LH spring channel water. The samples were transported under 4°C and stored at

-20°C for future analyses. When possible, parallel geochemical measurements were taken. Due to the remoteness of the LH spring site, logistical challenges and difficult weather often either prevented adequate time at the site or prevented planned field investigations completely. Therefore, it was not possible to acquire complete in situ geochemical data in each sampling campaign on an annual basis for both later winter (early May) and summer (July). Multiple geochemical parameters including temperature, pH, salinity, total dissolved solids and redox potential (ORP) were measured using the YSI 556 Multi Probe System (YSI

Incorporated, Yellow Springs, OH, USA). Hydrogen sulfide and dissolved oxygen concentrations were measured by colorimetric assay, as per manufacturer’s instructions (CHEMetrics, Calverton, VA, USA). For geochemical analysis sediments were dried at 60°C and finely ground to pass through a 1-mm-mesh sieve. Carbonate content was determined using a subsample of each oven-dried sediment that was acidified using 1M HCl and then dried at 50°C to remove the carbonates (Hedges and Stern, 1984). The original sediments were analyzed for total carbon and total nitrogen, and the acidified sediments were analyzed for organic carbon by combustion at 900 °C with a Carlo Erba Flash EA NC Soils

Analyzer (Carlo Erba, Milan, Italy; (Lim and Jackson, 1982). Ammonia, nitrite and nitrate concentrations were measured in the aqueous phase extracted from sediments following centrifugation at 2000 g for 10 minutes. Sediment-bound ammonia concentrations were determined by washing 30 g of sediment with 30 mL milli-Q water and then extracting with 30 mL of 2 M KCl (Maynard and

Kalra, 1993). Ammonia and nitrate/nitrite concentrations were analyzed on a 30

multi-channel Lachat AE Quik-Chem auto-analyser (Lachat Instruments;

Milwaukee, WI, USA).

2.2.2 CO 2 and CH 4 concentrations and flux measurements

To determine in situ CO 2 flux from LH channels, a Li-Cor Li-8100 (Li-Cor

Bioscience, Lincoln, Nebraska, USA) was used as described by the manufacturer.

Static chambers (Hoover et al., 2008) were also used to measure in situ CO 2 and

CH 4 flux rates from the LH spring outlet and channel sediments. Existing methodologies were adapted as follows: collars with a diameter of 24 cm were installed in the channel sediments or over bubbling hot spot/loci in the spring and

7-L chambers were placed on the collars and the system was allowed to equilibrate ~20 min-1h. For outlet measurements, the air in the chambers was mixed prior to sampling using a 50-mL syringe and 40-mL samples were collected and stored in 20-mL evacuated vials every 5 min for a 20-min period. CO 2 and

CH 4 concentrations in the gas samples were determined by gas chromatography as previously described (Roy and Greer, 2000) and fluxes were calculated based on linear regression. For the channel, flux chambers were set up with fixed sampling rates of 60 mL/min for 20 min, and then the total CO 2 and CH 4 were determined on a Picarro CRDS (Picarro, California). In order to measure both CO 2 and CH 4 concentrations within the sediments from LH spring and channel, the protocol described by Wagner et al . (Wagner et al., 2003), with some modifications was employed. Briefly, 10 g of sediment were added to a 60 mL vial containing 20 mL of saturated NaCl solution. The vial was crimp-sealed, vortexed for 30 s and

incubated for one hour at 80°C to allow for the transfer of gases from the sample to the headspace of the vial. The resulting CO 2 and CH 4 concentration in the headspace of the vial was determined by gas chromatography as described by Roy and Greer (Roy and Greer, 2000).

2.2.3 Microscopy and catalyzed reporter deposition fluorescence in situ hybridization (CARD-FISH)

To determine the numbers of bacterial and archaeal cells, CARD-FISH was applied to the samples. Sediment samples were prepared according to Pernthaler et al., (Pernthaler et al., 2001). In brief, 0.5 g of sediment from each sample was fixed using 4% para-formaldehyde overnight at 4°C, and then the samples were washed in PBS buffer 3 times and then stored in PBS/ethanol (1:1) solution at

-20°C. The fixed samples were filtered through polycarbonate filters of 0.22 µm pore size which were then embedded in 0.1% (w/v) low melting point agarose.

The dried filters were then treated with lysozyme solution for 1 hour at 37°C to increase the permeability of the microbial cell walls. Subsequently, the filters were hybridized with horseradish peroxidase (HRP) labeled probes (50 ng/ µl)

EUB338 (Amann et al., 1990), ARCH915 (Medina-Sanchez et al., 2005),

ANME-350 (Boetius et al., 2000), and NON338 (Wallner et al., 1993)] for bacteria, archaea, ANME-1, and a negative control, respectively. Filters were treated with formamide - 55% for EUB338 and ANME-350, 35% for ARCH915,

20% for NON338, and incubated at 35°C over-night. Fluorescently labelled tyramide and H 2O2 were used for the catalyzed reporter deposition for 15 minutes

at 46°C (Niederberger et al., 2010). Filters were viewed (10 fields of cell counting per slide) using a fluorescent Nikon Eclipse E600 microscope (Nikon, Melville,

NY, USA) at an excitation wave length of 568 nm (Furukawa et al., 2006) under a

100X immersion oil objective.

To determine the total cell numbers in the sediments, DAPI

(4’,6-diamidino-2-phenylindole) staining was performed. The protocol was modified based on Porter (Porter and Feig, 1980). In brief, 0.5 g of sediment was fixed using 3.7% formaldehyde at 25°C for 1 hour. The fixed samples were then washed by PBS buffer and then diluted 100 fold in PBS. The diluted sediment was filtered through polycarbonate 0.22 µm filters. Dried filters were then washed in water and 100% ethanol. Each slice of the cut filters was incubated with 10 µL of 20 µg/mL DAPI solution at 25°C for 10 minutes. The filters were again washed in PBS for 30 minutes at 25°C and then dehydrated with 100% ethanol. The filters were viewed (20 fields of cell counting per slide) using a fluorescent Nikon

Eclipse E600 microscope (Nikon, Melville, NY, USA) at an excitation wavelength of 350 nm under a 100X immersion oil objective.

2.2.4 Microbial cultivation and characterization

To evaluate culturable heterotrophic microorganisms, a total of 5 g of sample was used to prepare serial dilutions in tetrasodium pyrophosphate solution

(0.1w/v Na 4P2O7．10H 2O, pH 7.0) followed by spreading of 100 µL of each suspension (undiluted, 10 -1, 10 -2 and 10 -3 dilutions) onto R2A plates with 7% and

12% NaCl (Difco TM R2A Agar, Beckton, Dickenson Co., Sparks, MD, USA).

This medium has been successfully used for culturing and enumerating heterotrophic microorganisms from arctic hypersaline springs as described in previous studies (Niederberger et al., 2010; Perreault et al., 2008). All plate counts were performed in triplicate. The plates were incubated at room temperature for 2 weeks and 5°C for 2 months followed by colony counts. From the colonies that appeared and subsequent re-streaking, 22 isolates exhibiting unique colony morphology from R2A plates with 7% NaCl were selected. Cold temperature tolerance was tested by sub-culturing onto R2A plates supplemented with 7%,

12%, 20% and 25% NaCl incubated at 37°C, 25°C, 5°C, -5°C, and -10°C for 2 to

6 months.

For culturing of archaea, undiluted suspensions were also streaked onto plates containing DSMZ 371 media as modified by Walsh (Niederberger et al.,

2010) Versions of this media supplemented with 7% and 12% NaCl for selective isolation of archaea and evaluation of salt-tolerance were also tested.

DNA from isolated cells was extracted using a phenol/chloroform DNA extraction method (Barrett et al., 2006). Partial 16S rRNA fragments of the isolates were amplified by polymerase chain reaction (PCR) using the primer pair

27F (5' AGAGTTTGATCCTGGCTCAG 3’) and 758R (5'

CTACCAGGGTATCTAATCC 3’) (Bottos et al., 2008; Lane, 1991; Woese,

1987). The conditions for PCR reactions were as described by Steven et al .

(Steven et al., 2008). PCR products were sequenced using a 16-capillary genetic analyzer, ABI Prism 3130XL at the University Laval Sequencing Facility

(Plate-forme d’Analyses Biomoléculaires, Laval, QC, Canada). The sequences were compared against the Genbank database using the BLASTn algorithm and 34

Classifier tool of the RDP II (Cole et al., 2003).

2.2.5 Bacterial and Archaeal 16S rRNA gene clone libraries

Total genomic DNA was isolated from 0.5 g of sediment from LH channel samples using the Ultraclean Soil DNA Isolation Kit (MoBio Laboratories,

Carlsbad, CA) as per manufacturer’s instructions. DNA was eluted in 50 µL of sterile distilled H 2O and stored at -20°C. The 16S rRNA gene was amplified from the total isolated genomic DNA by PCR using primer pairs 27F and 758R for bacteria, and 109F (5' ACKGCTCAGTAACACGT 3’) and 934R (5'

GTGCTCCCCCGCCAATTCCT 3’) for archaea (Baker et al., 2003; Whitehead and Cotta, 1999). Each PCR contained 25 µl volumes with 1X PCR buffer, 0.2 mM of each dNTP, 3.5 mM MgCl 2, 0.5 µM of each primer, 6.25 µg bovine serum albumin, 1U of Taq polymerase and 2 µl of template DNA. Thermo-cycling conditions for archaeal PCR consisted of 94°C for 5 minutes followed by 20 cycles of 94°C for 30 seconds, 62°C for 30 seconds decreasing 1°C per cycle until

52°C, 72°C for 1 minute and 30 seconds followed by 15 cycles of 94°C for 1 minute, 52°C for 30 seconds, 72°C for 1 minute and 30 seconds and a final extension of 5 minutes at 72°C. For bacterial 16S rRNA genes, PCR conditions were the same as those used in the amplification of partial fragments of the 16S rRNA genes of the 22 isolates above (Steven et al., 2008). PCR products were cloned into the pGEM-T easy vector system (Promega, Madison, WI, USA) and the ligation products transformed into competent DH5 α cells (Invitrogen,

Carlsbad, CA, USA). Clone screening was carried out using amplified ribosomal

DNA restriction analyses (ARDRA) (Niederberger et al., 2010; Steven et al.,

2007a). Identical ARDRA patterns were considered as one OTU (operational taxonomic unit) and one or two representative clones were selected for sequencing. 16S rRNA sequence taxonomic affiliations were determined using the

Classifier tool of the RDP II (Cole et al., 2003). Sequences were also compared with the GenBank database using the BLASTn algorithm. All sequences from each clone library were aligned using ClustalW software and neighbor-joining phylogenetic trees built within the MacVector 7.2 software package (Oxford

Molecular Ltd., Oxford, UK) using Jukes-Cantor modeling with 1000 bootstrap re-samplings. The clone libraries were examined both in terms of total species richness for the channel communities as a whole (the unique sum of all clone libraries) and as individual profiles depicting changes in the relative species abundance between samples.

2.2.6 Biodiversity indices and statistical analysis of 16S rRNA gene clone libraries

Sampling coverage of clone libraries was calculated as defined by Good

(Good, 1953) using the formula C=(1-nl /N)×100, where nl is the number of phylotypes which only appeared once in the sample, and N is the size of the library. To determine the biodiversity, richness and evenness, Shannon index,

Simpson’s index, Chao1 (Chao, 1984; Perreault et al., 2007) and evenness were estimated using DOTUR software (Schloss and Handelsman, 2005). The reciprocal value of Simpson’s index (1/D) was used in this study for showing the

numbers of the most abundant phylotypes. The evenness was calculated by the formula: E=eH’ /N , where H’ is the value of Shannon index, and N is the total numbers of the phylotypes (Krebs, 1989).

2.2.7 Microbial activity at cold temperatures

To examine microbial activity at cold temperatures, microcosms containing

5 g of sediment from the Lost Hammer Spring channel were prepared as described by Steven et al. (Steven et al., 2008). Each microcosm was performed in triplicate.

Sterile controls were autoclaved twice for 30 min at 120°C and 1.0 atm, with a 24 h period between sterilizations. Each microcosm was supplemented with 0.045

14 mCi ml-1 (100000 disintegrations per min) of [1- C] acetate (specific activity

57.0 mCi/mmol; Amersham Biosciences, NJ, USA) and incubated at 5ºC, -5ºC,

-10ºC, -15 ºC and -20ºC in temperature-monitored incubators with +/- 1ºC temperature control. CO 2 traps in microcosms consisted of 1 M KOH (for 5°C and

-5°C) or 1M KOH + 15% v/v ethylene glycol (for -10°C, -15 ºC and -20ºC) to prevent freezing during incubation. The CO 2 traps were sampled at timed intervals (1 month) and radioactive counts determined by liquid scintillation spectrometry on a Beckman Coulter (CA, USA) LS 6500 Multi-purpose

Scintillation Counter (Steven et al., 2007b).

2.2.8 Nucleotide accession numbers

Partial 16S rRNA sequences were obtained from all clones and strains for

building phylogenetic trees as described above and have been deposited in the

NCBI database under accession numbers HQ444225-HQ444250 and HQ625077

(bacterial clones), HQ444251-HQ444262 (archaeal clones), and

HQ625055-HQ625076 (bacterial isolates).

2.3 Results

2.3.1 Geochemical analyses

Geochemical characteristics of the LH channel water and sediments varied seasonally and showed some distinct features compared to the LH outlet characteristics (Table 2-1). The water pH was near-neutral and had ~ 25 % salinity which are similar to the water characteristics previously reported for the LH outlet in 2005-2008 (Table 2-1). Due to seasonal periods of both flowing water and dry conditions in the channel, the temperature of these sediments experienced much more pronounced variation than LH outlet sediments, ranging from -18°C to above 0°C and as depicted in Figure 1b, the channel sediments remained unfrozen at -18°C. The channel is also more exposed to ambient air temperatures that range from -40°C in the winter to 15°C in summer, compared to the outlet that is more insulated within the salt dome. The total carbon content of the channel samples was also considerably higher than the total carbon content in the LH outlet (Table

2-1). The ammonia content of LH outelt water (6.87 mg/kg) was as concentrated as the upper range determined from channel water (6.57 mg/kg) sampled during low flow conditions in July 2009 (Table 2-1). However, samples analyzed from a

period of high precipitation as observed in July 2010, exhibited much lower ammonia content in the channel waters at (0.615 mg/kg). The range of carbon and nitrogen values for the LH channel indicated a seasonally variable nutrient supply.

The reduction potential in the channel sediments ranged between moderately reducing to moderately oxidizing conditions (-29.9 to 125.5 mV) while the outelt remained highly reducing. These values correspond to higher dissolved oxygen levels in the channel water (> 1.0 ppm) compared to the outlet.

2.3.2 CO 2 and CH 4 concentrations and flux measurements

Results for the sediment CO 2 and CH 4 concentrations and fluxes from the

LH channel and spring are presented in Table 2-3. CH 4 concentrations were an order of magnitude greater in the sediments from the outlet than in the sediments

2 from the channel (10 v.s. 10 nmol/g), while CO 2 concentrations were found to be similar at both sites (Table 2-3). Estimated CH 4 and CO 2 fluxes for the LH outlet as a whole (based on 4 actively bubbling seep spots), using the static chamber technique were 11,124 mg/day and 11,924 mg/day, respectively. Using the

2 constant flux rate method CH 4 flux was measured as 33.4 mg/m /day and 15,052

2 mg/m /day for CO 2 from the LH channel. By using a LiCor-8100, in situ CO 2 flux in the channel was determined at different locations both during spring and summer and were found to range between 152 mg/m 2/day to 38,244 mg/m 2/day.

The variability in flux within the channels appeared spatially heterogeneous and possibly driven by the degree of saturation of the sediments and not temperature, as both low and high CO 2 flux rates were measured in spring and summer

conditions while sediment temperatures varied from -16°C to 14 °C. Most CO 2 flux rates for the channel were lower than those estimated for the outlet with the exception of occasional bursts of gas exceeding outlet values.

2.3.3 Cell enumeration

Three methods were used to enumerate sediment microbial populations:

DAPI, CARD-FISH, and viable plate counts. The total abundance of microbial cells in the channel sediments had a mean value of 4.14 ± 1.58 × 10 7 cells/g sediment according to DAPI counts. The abundance of bacteria in channel sediments as determined by CARD-FISH enumeration had a mean value of 4.51 ±

0.65 × 10 7 cells/g sediment while the mean abundance of archaea was 3.99 ± 0.44

× 10 6 cells/g sediment. The ratio of bacteria to archaea was approximately 9:1.

The bacterial and archaeal numbers showed no significant differences between the different samples used in the analyses (p > 0.05). ANME-1 viable cells were below detection by CARD-FISH.

Plate count enumeration was used to determine the numbers of viable heterotrophic colonies with an average of 1.25 ± 0.59 × 10 5 CFU/g sediment on

R2A media with 7% NaCl and 2.4 ± 0.60 × 10 3 CFU/g sediment on R2A media with 12% NaCl at room temperature. Viable counts on R2A media with 20% NaCl incubated at 5°C were more variable with a mean of 1.05 ± 1.4 × 10 3 CFU/g of channel sediment.

2.3.4 Identification and characterization of isolates 40

A total of 22 unique bacterial strains were isolated from the R2A plates with

7% NaCl, identified by 16S rRNA sequencing, and characterized in terms of growth temperature ranges and salinity tolerance (Table 2-4). Isolated strains were grouped within four different phyla: the Firmicutes , Actinomycetes , Alpha - and

Gammaproteobacteria . The isolates were all related to known halophilic or psychrophilic representatives (Table 2-4). The majority of the strains (15/22) were growing on R2A media with 7% NaCl at -5°C (Table 2-4, supplemental Table

S2-1). None of these cold-adapted isolates grew at 37 oC on R2A media with 0% and 20% NaCl. All of the isolates grew on R2A media with 7% NaCl at 25°C and

5°C. A total of 6 strains were considered obligate halophiles and grew with 20%

NaCl but were unable to grow on media without a minimum of 7% NaCl.

However, no strain grew on the media with 25% NaCl, at any temperatures and no strain grew at -10°C (data not shown). Generally, increasing NaCl concentration and decreasing temperature inhibited growth of the isolates

(supplemental Table S2-1). Most of the strains (19/22) were pigmented

(supplemental Table S2-1). No archaea were successfully isolated.

2.3.5 Bacterial and Archaeal 16S rRNA gene clone libraries

A total of 486 bacterial clones and 184 archaeal clones were obtained from

LH channel samples and then analyzed as combined clone libraries for bacteria and archaea, respectively; the microbial composition of the individual clone libraries are shown in Table S2-2 and S2-3. The clone libraries were examined both in terms of total species richness for the channel community as a whole (the

unique sum of all clone libraries) and as individual profiles depicting natural variation in the relative species abundance between samples. According to the statistical analyses conducted in DOTUR (OTUs > 97%), these clone libraries indicated large variation in the bacterial diversity ranging between 16 and 76 unique bacterial phylotypes and less variation for the archaea having between 3 to

6 unique phylotypes, respectively (Table 2-5). The most abundant groups in the bacterial clone library were the Bacteroidetes (46.1 % of all clones), followed by similar amounts of Actinomycetes (18.3 %), Alphaproteobacteria (16.5 %) , and

Gammaproteobacteria (11.1%) (Figure 2-2a). The most common genus among the Bacteroidetes clones was related to Gillisia spp. (32.3%). Among the other phylotypes, species involved in methanotrophy/methylotrophy and sulfur cycling were present in the 16S rRNA gene clone library and their phylogenetic comparisons are shown in Figure 3. Among the methanotrophs, one clone

(LHCbac-24) had a top BLASTn match (91% similarity) with Crenothrix polyspora (DQ295898), a filamentous aerobic methane oxidizer (Stoecker et al.,

2006). Several clones affiliated with methylotrophs were also detected including close matches to Methylophaga sulfidovorans (91% identity to NR_026313) and

Methylophaga thiooxidans (90% identity to DQ660915), which are both able to oxidize dimethylsulfide (DMS) (Boden et al., 2010; de Zwart et al., 1996). Two other clones, LHCbac-15 and LHCbac-19, were affiliated with methylotrophic bacteria Methylobacterium sp. and Methylibium sp., respectively. Sulfur-cycling phylotypes, such as sulfur reducing species including Desulfuromonas

(LHCbac-25) and sulfur oxidizing bacteria including Thiobacillus (LHCbac-16), were also detected and are shown in Figure 2-3. 42

Amongst the archaeal 16S rRNA clone libraries, representatives were classified within four phyla with the most abundant being among the newly defined archaeal phylum Thaumarchaeota (70.2 %) (Figure 2-2b)

(Brochier-Armanet et al., 2008). The remainders were all classified within

Euryarchaeal phylotypes including Halobacteria (15.9 %), Methanobacteria

(12.4 %) , and a small percentage of unclassified Euryarchaea (1.5 %). According to the phylogenetic analyses, several clones (LHCarc-9, LHCarc-10, LHCarc-7,

LHCarc-8) grouped with the ammonia-oxidizing Thaumarchaea species

Candidatus Nitrososphaera gargensis (Spang et al., 2010) (Figure 2-4). Although the classification of two clones, LHCarc-11 and LHCarc-12, was not entirely conclusive, they appeared to match Thaumarchaea more than any other archaeal phyla with greater than 94% identity (Figure 2-4). Phylotypes representing methanogenic archaea, which were all related to the genus Methanobrevibacter

(12.4 %), were also found in the clone library. No clones related to the anaerobic methane oxidizing ANME-1 clade were detected in the clone libraries.

The biodiversity indices (Table 2-4) for the bacterial and archaeal 16S rRNA clone libraries from the LH channel indicated relatively low diversity of the archaeal clone libraries (Shannon index = 0.78 – 1.37) compared to the higher diversity within the bacterial clone libraries (Shannon index: 1.69 – 3.80). Library characteristics were described using multiple indices, indicating that a large proportion of the expected bacteria (Chao1= 44 – 104, evenness = 34 – 59%) and archaea (Chao1= 3 – 6, evenness = 36 – 98%) were successfully sampled. The most abundant phylotypes defined by the inverse of the Simpson’s index were 3 to 25 bacterial phylotypes, and 1 to 4 archaeal phylotypes. The Good’s coverage 43

ranged between 67.8 to 80.0 for the bacterial and 87.5 to 95.0 for the archaeal libraries. Compared to LH outlet clone libraries, the channel depicts higher bacterial diversity but comparable archaeal diversity in the sediment microbial communities.

2.3.6 Microbial activity at cold temperatures

In order to detect microbial respiration and activity at cold temperatures, mineralization of 14 C-acetete was evaluated within channel sediment microcosms.

Following 6 months of incubation, mineralization was observed in triplicate microcosms compared to sterile controls (Figure 2-5). The highest mineralization rates occurred at 5°C (~ 30%) but microbial respiration was also detected at -5°C

(~10%), -10°C (0.19%), -15°C (1.21%) and -20°C (0.17%) (Figure 2-5).

Although rates dropped significantly at temperatures ≤ -10 °C, levels of microbial respiration were above background levels measured in the -10°C, -15°C and

-20°C sterile controls.

2.4 Discussion

Lost Hammer Spring represents the first described example of a subzero terrestrial methane seep ecosystem (Niederberger et al., 2010). We have previously reported that the LH spring outlet discharge conditions and temperature remain stable throughout the year and that the majority of the methane exsolving from the outlet appears to be thermogenic in origin

(Niederberger et al., 2010). The physical and geochemical characteristics of the

LH outlet correspond well with the types of microbial metabolism (including anaerobic methane oxidation AOM, by ANME-1 archaea) that were indicated from molecular microbiological based analyses that showed low diversity but viable bacterial and archaeal communities (Niederberger et al., 2010). The LH outflow channel described here, represents a distinct and more heterogeneous and stochastic environment occurring downstream of this subzero, hypersaline methane seep.

The LH channel exhibits many of the extreme conditions found in the outlet but experiences greater seasonal oscillations in physical and geochemical characteristics including water level, much broader temperature ranges, changes in O 2 content, and different carbon and nitrogen concentrations than observed in the outlet. Most notable are the channels’ large variations in temperatures (-18 to

14 °C), and their less reducing, more aerobic conditions compared to the seasonally stable outlet (consistently ~ -5°C). There is a sharp decrease in methane concentration in the channel sediments with only ~1/10 of the methane present than in the outlet sediments and only moderate CH 4 flux detection over a short time period but remain higher than surrounding permafrost and atmospheric levels. These variations create a more dynamic and heterogeneous environment for the channel microbial communities and the ranges in channel geochemical data were reflected by changes in relative abundance of different bacterial and archaeal phyla from the individual channel clone libraries, as described below

(Supplementary Table S2-2). In similar investigations of gradients along marine methane seeps, differences in geochemical conditions have been shown to 45

correlate with niches occupied by distinct microbial communities including different spatial abundance of ANME archaea (Arakawa et al., 2006; Knittel et al.,

2005). Although, inferences of metabolism determined by 16S rRNA gene phylogenies alone must be taken with some degree of caution, the changing composition of the LH channel microbial communities between samples and over time suggest that species abundance is in fact dynamic and that as with diversity, would likely be driven by environmental conditions.

With the exception of LH spring, all methane seeps studied to date are located at deep sea marine margins where methane hydrates occur (Valentine,

2002; Valentine and Reeburgh, 2000). In these cold marine seep ecosystems, simple assemblages of key functional groups including methanotrophs, hydrocarbon degraders, sulfate-reducing and sulfide-oxidizing are typically found

(Jorgensen and Boetius, 2007). Compared to the LH outlet, the microbial communities inhabiting the LH channel depicts a similarly simple archaeal composition (Shannon diversity index of 1.39 (outlet) and 0.76 - 0.98 (channel), respectively) which is common in different marine methane seep communities

(Knittel et al., 2005). Notably, the bacterial distribution shows a more diverse assemblage occurring in the channel compared to the outlet (Shannon diversity index of 1.64 (outlet) and 1.69 - 3.80 (channel), respectively). The microbial biomass is also 10 to 100 times greater in the channel than in the outlet. Both these observations suggest that the channel community provides a more favourable environment for microbial growth and has a greater diversity of niches than are present in the outlet. When compared with similar terrestrial cryoenvironments, the biodiversity estimates indicate that the LH channel 46

sediments (Chao1 = 44 – 104) have a slightly more diverse bacterial community than two other studied low-temperature saline springs found on AHI, namely

Gypsum Hill (GH: Chao1 = 71) and Colour Peak (CP; Chao1 = 50) (Perreault et al., 2007). However, the biodiversity of archaea in the LH channel (Chao1 = 3-6) reveals this community is simpler than those identified in CP (Chao1 = 68) and

GH (Chao1 = 23) (Perreault et al., 2007).

Steep geochemical gradients that are naturally present along seep systems, including the LH ecosystem, are important drivers of niche formation (Knittel et al., 2005). The presence of numerous species in the LH channel environment not found in the LH outlet reflects a greater potential metabolic diversity utilizing a variety of nitrogen and sulfur substrates in addition to methane. The phylotypes previously found within the LH outlet sediments include Bacteroidetes ,

Alphaproteobacteria , Betaproteobacteria , Gammaproteobacteria , Cyanobacteria, and Firmicutes (Niederberger et al., 2010). With the exception of Cyanobacteria , all LH outlet phyla were represented in the channel as well as several additional phyla, including an abundance of Actinobacteria, that were not detected in the LH outlet sediments. Phylotypes for Halomonas , Gillisia , and Marinobacter, which are common bacterial genera in cold Arctic and Antarctic environments (Bowman et al., 1997; Bowman and Nichols, 2005; Brinkmeyer et al., 2003; Franzmann et al., 1987; Guan et al., 2009; Zhang et al., 2008), were detected in both the LH outlet (Niederberger et al., 2010) and channel sediments. Representatives of sulfur cycling bacteria were also detected in channel sediments that were not found in the outlet sediments. The presence of diverse bacterial, sulfur related phylotypes indicate that various sulfur intermediates were important metabolic substrates 47

within the LH channel including sulfite ( Sulfitobacter sp. and Desulfitibacter sp.), elemental sulfur ( Desulfuromonas sp.) and various reduced sulfur compounds for sulfur oxidizers ( Thiobacillus sp., Sulfuricurvum sp., Sulfurovum sp.). Most of the closest relatives for these identified phylotypes were found in cold and/saline environments including Desulfuromonas sp. identified in sub-permafrost saline fracture water at the Lupin mine in the Canadian arctic (Onstott et al., 2009).

In contrast to finding several similar species in the bacterial profiles of the

LH outlet and channel sediments, the archaeal 16S rRNA clone libraries showed large differences in detected phylotypes. The most abundant and notable phylotype in the outlet was related to the anaerobic methane oxidizing archaea group 1 (ANME-1), however, this group was not detected in the channel clone libraries. A few putative cells of ANME-1 homologues (statistically below the detection limit) were detected in the channel sediments by CARD-FISH using an

ANME-1 specific probe (ANME-1 350) (Losekann et al., 2007), suggesting that the much lower concentration of methane and increased oxygen content may not favour ANME-1 in the channel sediments. However, unlike in the outlet sediments, methanogenic archaea, predominantly Methanobrevibacter were detected in the channel suggesting that at least part of the methane produced in the channel, albeit at significantly lower concentrations than the outlet, may be of biogenic origin. Further analyses are currently underway to determine the biogenic/thermogenic signature of LH channel methane. The potential for methane cycling in the channel sediments is also supported by an abundance of aerobic methylotrophic/methanotrophic species including Methylobacterium , suggesting that methane may be used as both carbon and energy outlet under the 48

moderately aerobic conditions in the LH channel system.

A notable feature of the channel communities is abundance of the phylum,

Thaumarchaeota (Brochier-Armanet et al., 2008), which comprised the largest proportion of the archaeal clone libraries but was not detected in the LH outlet sediments. To date the Thaumarchaeota , which are found in a wide range of environments, are thought to be associated with pathways involved in nitrification, and are often referred to as ammonia-oxidizing archaea (AOA) (Spang et al.,

2010). Marine AOA species are able to convert ammonia to nitrite in environments of low ammonia concentrations, such as the open sea (< 0.03–1 μM) and coastal waters (< 0.03–100 μM) (Konneke et al., 2005) both of which are lower than the ammonia concentrations measured in the LH channel sediments.

The majority of the currently known species of Thaumarchaeota have been identified within mesophilic and thermophilic environments (de la Torre et al.,

2008; Hatzenpichler et al., 2008a; Konneke et al., 2005; Schouten et al., 2008), however some phylotypes have also been detected in cold Antarctic bathypelagic sediments (Gillan and Danis, 2007). Four channel clones grouped closely with

Candidatus Nitrososphaera gargensis and Nitrosopumilus maritimus , the latter being a moderately psychrophilic and halophilic species (Konneke et al., 2005).

The presence of Thaumarchaeota in the LH channel seems plausible as suitable conditions (moderately aerobic, high salinity and low sediment ammonia concentrations) for their autotrophic ammonia oxidizing and nitrification activities exist within the LH channel. This is the first description of Thaumarchaeota identified within a either a subzero and/or hypersaline environment.

All the strains isolated in this study were related to known cold and salt tolerant species. Interestingly, we were able to culture a portion of the bacterial phylotypes detected within the 16S clone libraries including Marinobacter ,

Planococcus and Nesterenkonia with all isolates capable of growth at LH in situ temperatures of (-5°C) and 13 of 22 isolates capable of growth at the high in situ salinity concentrations (20%). Only Marinobacter spp. has been also isolated from the LH outelt this sentence is not clear? (Niederberger et al., 2010) but most of the isolated strains have also been found in nearby GH and CP springs

(Perreault et al., 2008), as well as permafrost from Eureka (Steven et al., 2008), and in Antarctic sea ice brine (Junge et al., 1998). The majority of the isolated channel strains were also pigmented; a common adaptive strategy in many cold environment species that may serve several functions including cryo- and solar radiation protection, light-harvesting, and anti-oxidative activity (Dieser et al.,

2010; Mueller et al., 2005).

Due to the long term stability of DNA at high ionic concentration, low temperatures and anoxic environmental conditions (Inagaki et al., 2005) it is difficult to conclude which of the microbial phylotypes detected in the LH channel are active or dormant under in situ conditions. However, the high rate of

14 C-acetete mineralization suggests that a significant portion of the microbial biomass could be active in situ . The high viable cell counts suggest that a large proportion of the cells were in fact alive, and could be active under in situ conditions as were all of the isolated strains. The mineralization rates reported at 5

°C and -5 °C within the same days (60 to 120 days) are similar to mineralization rates (around 20 to 30%, and 10 to 15%, respectively) reported for other Arctic 50

samples, such as Eureka permafrost active layer (Steven et al., 2008), the

Markham ice shelf, and the Ward Hunt Ice shelf (Steven et al., 2007b). In contrast, microbial activity at -5°C, -10°C, -15°C and -20°C were lower than permafrost soils but remained similar to ice shelf mineralization rates (Steven et al., 2007b).

The mineralization rates of 14 C-glucose by sediments from the LH outlet (< 2% for 5°C , -5°C and -10°C after 6 months (Steven et al., 2007b) were not as high as rates of 14 C-acetete utilization by the LH channel sediments. However, the very low rate of 14 C mineralization at -10 °C in the channel sediments indicate that microbial activity and growth may be restricted at temperatures ≤ -10 °C in this hypersaline environment. However, detection of CO 2 and CH 4 flux from the channel sediments does suggest that microbial respiration occurs both in winter and summer within the channel sediments under ambient conditions. It is also likely that some of the measured gas flux represents pockets of gas periodically released from saturated and/or partially frozen sediments. The higher activity may be due in part to the 10 to 100 times greater biomass present in the channel compared to the LH outlet. Additional analyses are underway to determine how much of the CO 2 and CH 4 efflux is biotic in origin.

In conclusion, our results illustrate that the LH channel sediments present a dynamic yet extreme habitat supporting microbial community with greater biomass, diversity, and activity than previously found in the LH spring outlet.

Overall, the findings further our understanding of extreme microbial ecosystems within the context of unique niche formation along chemical gradients within the same spring environment. Future investigations will probe which species are active under in situ conditions through deep sequencing of cDNA libraries of 16S 51

rRNA coupled with metatranscriptomic analyses to identify the active metabolic pathways occurring within the channel relative to the LH outlet environment.

Lastly, examination of the unique and diverse microbial communities present in such a relatively cold, hypersaline cryoenvironment both will increase our knowledge of microbial life in extremely cold habitats on Earth as well as lead to a more realistic understanding for the potential of microbial life to exist in other very cold solar system bodies such as Mars, Europa, or Enceladus.

2.5 Acknowledgements

This project is supported by NSERC CREATE Astrobiology Training

Program, CSA Canadian Analogue Research Network Grant Program, Polar

Continent Shelf Program, NSERC Discovery Grant Program, and Department of

Natural Resource Sciences, McGill. We thank Dr. M. Wagner for the information regarding Thaumarchaeota , Dr. A. Chao for the questions regarding the indices,

Dr. J. Whalen and H. Lalande for the help of C/N/nitrite/nitrate/ammonia analyses, and thanks the members of the Whyte and Greer Labs for helpful discussions and to Dr. H. Vrionis for critical review of the manuscript.

Table 2-1. Physical and geochemical characteristics for Lost Hammer (LH) Spring outlet and channel determined during both Arctic winter and summer

LH outlet a Channel b Temperature (°C) -5.9 to -4.7 -18 to 9.2 pH 5.96 to 7.38 6.52 to 7.28 DO (ppm) 0.1 to 1.0 >1.0

H2S (ppm) 0 to 50 0 to 20 ORP (mV) -187.4 to -154.0 -29.9 to 125.5 TDS (g/L) 175.0 to 241.7 61.5 to 95.7 Salinity (%) 22 to 26 22 to 26 Total viable cell count on 7% NaCl R2A media N.D. c 1.25±0.59×10 5 (CFU/ g sediment) Total viable cell count on 12% NaCl R2A media N.D. 2.40±0.60×10 3 (CFU/ g sediment) Bacterial cells (CARD-FISH) (cells/ g sediment) d 3.61±0.11×10 5 4.51±0.65×10 7 Archaeal cells (CARD-FISH) (cells/ g sediment) 1.63±0.11×10 4 3.99±0.44×10 6 a Data represent the range of measurements from the outlet determined between 2005 to 2008 described in Niederberger et al., 2010. b Data represent the range of measurements from the channel region determined between 2008 to 2010. c N.D., not determined. d Data for CARD-FISH from the LH outlet were converted from the percentage of total DAPI-stained cells.

Table 2-2. Carbon and nitrogen analyses for LH Spring outlet and channel LH outlet Channel Total Carbon (%) 0.48 a 0.92 to 1.08 Organic Carbon (%) 0.45 a 0.77 to 0.93 Total Nitrogen (%) N.D. c 0.02 to 0.08 Water dissolved Nitrite/Nitrate (mg/ L)b 0.13 0.04 to 0.17 Water dissolved Ammonia (mg/ L) 6.87 0.62 to 6.57 (381 µM) (91.6 to 365 µM) Sediment Nitrite/Nitrate (mg/ kg) 2.87 0.09 to 0.12 Sediment Ammonia (mg/ kg) 2.55 0.76 to 0.88 a These values are taken from Niederberger et al. 2010 b The nitrite/nitrate/ammonia concentrations in the water and sediments were analyzed separately. c N.D., not determined.

Table 2-3. CO2 and CH4 sediment concentrations and fluxes from LH Spring outlet and channel determined during both Arctic winter and summer Site Sediment Concentration a Flux

CH 4 (nmol/g) CO 2 (µmol/g) CH 4 CO 2 LH outlet 102 ± 17.7 41.1 ± 0.53 11.1 g/day b 11.9 g/day b Channel 9.36 ± 1.70 35.5 ± 4.82 N.D. 18.3 to 84.0 g/m 2/day c a Values are means of triplicates; standard error of the mean is presented. b Values are an estimate of the CH 4 and CO 2 fluxes for the entire spring, based on the assumption that four hotspots are continuously bubbling; N.D.: not determined. c The values were determined using the LiCor 8100. These values were converted from 0.48 µmol/m 2/s and 2.21 µmol/m 2/s.

Table 2-4. Characteristics of 22 bacterial strains isolated from LH channel sediments Numbers Salinity Temper Closest cultured Origin of Similarity RDP classifier of unique range ature BLAST hit BLAST to BLAST (> 80% strains (% NaCl) range (Accession #) relative sequence confidence)

(°C )g 7a 0 to 20 -5 to Planococcus sp. Axel Heiberg 98-99% Planococcaceae 37 NP 19 (EU196338) Island, Perennial (Family, 100%)

Spring

4b 7 to 20 -5 to Marinobacter sp. Antarctic Sandy 97-100% Marinobacter 25 ZS1-16 (FJ889664) intertidal (Genus, 100%)

sediments

2c 0 to 12 -5 to Psychrobacter sp., E59 Antarctic sea 98-99% Psychrobacter 25 (DQ667083) water (Genus, 100%) 5d 0 to 20 5 to 37 Nesterenkonia sp. 35/46, Antarctica soil 98-100% Nesterenkonia

(AY571802) (Genus, 100%)

1e 0 to 12 -5 to Planomicrobium Mud volcanoes in 99% Planococcaceae 37 psychrophilum strain Xinjian (Family, 100%)

4-5-26 (GQ505362)

1f 0 to 7 5 to 25 Fulvimarina sp. Axel Heiberg 100% Aurantimonadace

NP 28 (EU196328) Island, Perennial ae (Family, 100%)

Spring a These 7 strains include CY-C3-1, CY-C3-3, CY-C3-4, CY-C3-5, CY-C3-6, CY-C1-11, CY-C2-19-2 b These 4 strains include CY-C3-2, CY-C3-9, CY-C2-15, CY-C2-17-1 c These 2 strains include CY-C3-7, CY-C3-8 d These 5 strains include CY-C1-10, CY-C2-13, CY-C2-14, CY-C2-17-2, CY-C2-19-1 eThis 1 strain includes CY-C1-12 f This 1 strain includes CY-C2-18 g No strain grew at 37 °C on R2A media with 0% and 20% NaCl. All the strains grew at 25 and 5 °C on R2A media with 7% NaCl. Specific characteristics for each strain are presented in the supplementary files (Table S2-1).

Table 2-5. Summary of the range of statistics and indices for the 16S rRNA gene clone libraries of LH channel and outlet sediments Channel Outlet a Bacteria Archaea Bacteria Archaea No. of Clones 80 to 236 24 to 80 61 66 No. of Phylotypes 16 to 76 3 to 6 9 7 Shannon Index (H’) 1.69 to 3.80 0.78 to 1.37 1.65 1.39 Simpson Index 3 to 25 2 to 4 N.D. N.D. (1/D) Chao1 44 to 104 3 to 6 N.D. N.D. Evenness (E) 0.34 to 0.59 0.36 to 0.98 N.D. N.D. Coverage (%) 67.8 to 80.0 87.5 to 95.0 98.5 95.1 a Data represent the statistics and indices obtained from Niederberger et al., 2010. N.D.: not determined.

(A)

(B)

Fig. 2-1. The images of LH channel (A) Photograph showing the position of Lost

Hammer Spring outlet and channel. (Scale bar: 2 m.) (B) Measurements being taken in LH Spring channel; the sediments remained unfrozen at -18°C.

(A)

(B)

Fig. 2-2. Phylogenetic composition of sequences from (A) Bacterial and (B)

Archaeal 16S rRNA gene clone libraries constructed from samples from Lost

Hammer spring channel sediments. Sequences were grouped using the RDP

Classifier function of the Ribosomal Database Project-II release 9 with a confidence threshold of 80%.

Fig. 2-3. Phylogenetic relationships of representative bacterial 16S rRNA gene sequences obtained from the LH Spring channel clone libraries and strains. The tree was inferred by neighbor-joining analysis of 460 homologous positions of sequence from each organism or clone. Numbers on the nodes are the bootstrap values based on 1,000 replicates. Scale bar indicates the estimated number of base changes per nucleotide sequence position. Percentages indicate the prevalence of the clone types within the clone library with the number of clones indicated in parentheses. The titles starting with LHCbac indicate LH channel clone representatives, and the titles starting with CY indicate the strains.

Fig. 2-4. Phylogenetic relationships of the archaeal 16S rRNA gene sequences obtained from the LH channel clone libraries. The tree was inferred by neighbor-joining analysis of 576 homologous positions of sequence from each clone. Numbers on the nodes are the bootstrap values based on 1,000 replicates.

The scale bar indicates the estimated number of base changes per nucleotide position. Percentages indicate the prevalence of clone types within the clone library with the number of clones indicated in parentheses. The titles starting with

LHCbac indicate LH channel clone representatives.

Fig. 2-5. Mineralization assays of [1-14C] acetate in LH channel sediment microcosms at different temperatures. Each point represents the mean cumulative

14 mineralization (% CO 2 recovered) from triplicate assays. Curves are shown for

5°C ( ◆) and -5°C ( ■) on the primary axis, and for -10°C ( ▲), -15°C ( ●), -20°C

(＊) and the sterile controls on the secondary axis. The curves of the samples and the sterile controls are shown in the solid lines and the dotted lines, respectively.

CONNECTING TEXT

To understand the functional genetic potential of the LH system, we changed our focus to the outlet sediment. Using the metagenomic strategy, it reveals a relatively complete map for understanding the pathways driven by microorganisms within it. To corresponding with the environmental properties, the study focused the analyses on the pathways related to sulfate, nitrogen, and methane metabolisms, as well as the genes involved in stress response.

CHAPTER 3

Defining the Functional Potential and Active Community Members of a Sediment Microbial Community in a High Arctic Hypersaline Subzero Spring

Chih-Ying Lay 1, Nadia C. S. Mykytczuk 1, Étienne Yergeau 2, Guillaume

Lamarche-Gagnon 1, Charles W. Greer 2 and Lyle G. Whyte 1*

1Department of Natural Resource Sciences, McGill University, Canada 2Biotechnology Research Institute, National Research Council Canada, Montreal,

Canada

Published in: Applied and Environmental Microbiology, June 2013. 79 (12):3637-3648

CONTRIBUTION OF AUTHORS The metagenome sample was collected by myself and Dr. Whyte; as well as the total RNA sample was collected by Dr. Mykytczuk. All of the experiments were designed and performed by myself with the consultation of Dr. Whyte and Dr. 62

Greer. The PCoA analysis was performed by Dr. Yergeau. Most analysis related to methanogen was done by G. Lamarche-Gagnon. Dr. Mykytczuk analyzed most genes related to sulfur cycle. The rests of the analyses were processed by myself.

The manuscript was written by myself with critical review provided by Dr. Whyte,

Dr. Mykytczuk, and G. Lamarche-Gagnon.

ABSTRACT

The Lost Hammer (LH) spring is the coldest and saltiest terrestrial spring discovered to date and is characterized by perennial discharges of subzero temperatures (-5°C), hypersalinity (24% salinity), along with reducing ( ≈-165 mV), and microoxic, conditions. It is rich in sulfates (10.0% w/w), dissolved

-1 H2S/sulfides (up to 25 ppm), ammonia ( ≈381 µM), and methane (11.1 g d ). To determine its total functional and genetic potential and to identify its active microbial components, we performed metagenomic analyses of the LH Spring outlet microbial community and pyrosequencing analyses of the cDNA of its 16S rRNA genes. Reads related to Cyanobacteria (19.7%), Bacteroidetes (13.3%), and

Proteobacteria (6.6%) represented the dominant phyla identified among the classified sequences. Reconstruction of the enzyme pathways responsible for bacterial nitrification/denitrification/ammonification and sulfate reduction appeared nearly complete in the metagenomic dataset. In the LH 16S ribosomal cDNA active community profile, ammonia oxidizers ( Thaumarchaeota ), denitrifiers ( Pseudomonas spp.), sulfate reducers ( Desulfobulbus spp.), and other sulfur oxidizers ( Thermoprotei ) were present, highlighting their involvement in nitrogen and sulfur cycling. Stress-response genes for adapting to cold, osmotic

stress, and oxidative stress were also abundant in the metagenome. Comparing functional community composition of the LH spring to metagenomes from other saline/subzero environments revealed a close association between LH and another

Canadian High Arctic permafrost environment, particularly in genes related to sulfur metabolism and dormancy. Overall, this study provides insights into the metabolic potential and the active microbial populations that exist in this hypersaline cryoenvironment and contributes to our understanding of microbial ecology in extreme environments.

3.1 Introduction

Cryoenvironments are defined as permanently subzero or frozen environments, such as permafrost, glaciers, ice sheets, multi-year sea ice, high-elevation Antarctic dry valleys, and some cold saline springs (Bowman et al.,

2012; Cary et al., 2010; Lay et al., 2012; Niederberger et al., 2010; Steven et al.,

2008; Varin et al., 2012). Microorganisms inhabiting cryoenvironments must face the challenges of subzero temperatures, low water activity, and often, high solute concentrations to sustain their viability. The cold saline springs on Axel Heiberg

Island (AHI) in the Canadian high Arctic discharge through 500-600 m of thick permafrost, maintain a liquid state at subzero temperatures, and offer a unique opportunity to assess microbial adaptations to extremes of both high salinity and subzero temperatures (Lay et al., 2012; Niederberger et al., 2010; Perreault et al.,

2007; Perreault et al., 2008; Pollard et al., 2009). These springs occur in an area with an average annual air temperature of -15ºC, reaching below -40ºC during the 64

winter months, and probably originate from sub-permafrost groundwater flow through carboniferous evaporites in areas of diapiric uplift on AHI (Andersen et al., 2002; Pollard et al., 1999). Other Arctic cold springs have been described on

Ellesmere Island in the Canadian high Arctic, as well as on the Norwegian high

Arctic Svalbard archipelago (Gleeson et al., 2011; Grasby et al., 2003; Reigstad et al., 2011), although the discharges from these springs are not subzero. Viable microbial communities were described in all of these Arctic springs (Gleeson et al., 2011; Lay et al., 2012; Niederberger et al., 2010; Perreault et al., 2007;

Perreault et al., 2008; Reigstad et al., 2011; Steven et al., 2007b).

The Lost Hammer (LH) spring, located in the central west region of AHI

(79°07’ N; 90°21 W’) is the coldest and saltiest of all described Arctic springs to date. LH is characterized by hypersaline (24%) perennial discharge at subzero temperatures (~ -5ºC) flowing to the surface through a hollow, 2m high cone-shaped salt tufa structure. The discharge waters are microoxic (dissolved oxygen 0.1 to 1 ppm), highly reducing ( ≈ -165 mV), and neutral (pH ≈ 7), and contain ammonia (6.87 mg kg -1), and high concentrations of sulfate (10.0% w/w).

During the summer months, the spring waters empty from the dome structure partially exposing the spring sediments to ambient conditions; however, the sediments remain anoxic and highly reducing. Continuous gas emissions from the spring indicate a thermogenic methane source underlying LH (Lay et al., 2012;

Niederberger et al., 2010). Based on these properties, this spring is considered a significant astrobiology analogue site to possible habitats currently present on

Mars and the cold moons Europa and Enceladus. For example, the widespread distribution of chloride and sulfate minerals on Mars (Gendrin et al., 2005; Hecht 65

et al., 2009), reports of spring-like structures on the Martian surface (Allen and

Oehler, 2008; Rossi et al., 2008), recent images indicating that liquid brines flowed on Mars during the past decade under mean surface temperatures of -60°C and extensive permafrost (Malin et al., 2006; McEwen et al., 2011), and the potential detection of atmospheric methane on Mars (Keppler et al., 2012;

Mumma et al., 2009; Zahnle et al., 2011), highlight the importance of terrestrial cold hypersaline environments, such as LH, as analogue sites for Mars as well as for the icy Saturnian moon Enceladus where methane, ammonia, and simple organics have been detected in the saline plume features erupting from its surface

(Postberg et al., 2011).

In our initial studies of the sediments of the LH spring outlet (Steven et al.,

2007b) and outflow channels (Lay et al., 2012), microbial activity was detected using mineralization assays. We also isolated halophilic and cryophilic microbial strains from the sediments of both the spring outlet and outflow channels.

Microbial community profiling (16S rRNA clone libraries) of LH revealed phylotypes related to halophilic bacteria/archaea, sulfate reducing archaea/bacteria, methylotrophic/methanotrophic bacteria, and methanogenic archaea (Lay et al.,

2012; Niederberger et al., 2010). The ANME-1, a clade of anaerobic methane-oxidizing archaea, dominated the archaeal community in the spring outlet sediments (Niederberger et al., 2010), while sequences related to

Thaumarchaeota dominated the spring channel sediments (Lay et al., 2012).

Metagenomic analyses of other extremely cold or saline environments have revealed the importance of genes involved in carbon cycling operating in permafrost (Mackelprang et al., 2011; Yergeau et al., 2010), genes related to stress 66

responses of microorganisms colonizing ice shelves (Varin et al., 2012), evidence of lateral gene transfer in deep-sea hydrothermal vent biofilms (Brazelton and

Baross, 2009, 2010; Xie et al., 2011), and the microbial ecology of an Antarctic meromictic lake (Lauro et al., 2011; Ng et al., 2010). Surveys of complementary

DNA (cDNA) of 16S ribosomal RNA are currently used to identify potentially active microorganisms in diverse environments (Burow et al., 2012; Jones and

Lennon, 2010; Murray et al., 2012; Rodriguez-Blanco et al., 2009) including a subzero, briny ice sealed lake in the Antarctic (Murray et al., 2012). In the present study, we combined a metagenomic approach with bacterial and archaeal 16S ribosomal cDNA pyrosequencing analyses to assess the functional potential of the

LH microbial community and to identify the active community members in LH and consequently infer their possible ecological functions.

The specific objectives for analyzing the LH metagenomic and 16S ribosomal cDNA datasets of the spring outlet sediment were to (i) map the microbial metabolic pathways driving biogeochemical cycles, focusing on methane, ammonia, and sulfur cycling which were expected to play key roles in shaping LH communities based on previous investigations of the LH system (Lay et al., 2012; Niederberger et al., 2010); (ii) identify the dominant genes involved in adaptations to cold and high salt concentrations that would allow autochthonous populations to cope with the natural extreme conditions of the site;

(iii) compare the functional potential of the LH metagenome to metagenomes from other cold or saline environments, and (iv) identify the bacterial and archaeal taxa that may be active in situ .

3.2 Materials and Methods

3.2.1 Study site and sample collection

The LH spring (79°07’ N; 90°21’W) is located on Axel Heiberg Island in a valley off the south shore of Strand Fjord. A ~1.7 m high dome-like structure composed of precipitated mineral salts surrounds the spring outlet.

Our in situ analyses of geochemical/physical parameters recorded from

2005 to 2012 indicates that the LH spring outlet sediment and water environment has remained very consistent (-5ºC, ~ 25% salinity, -160 mV ORP, 0.1 to 1 ppm

D.O.) in both late winter (April / May) and midsummer (July) sampling points.

Therefore, we considered that LH samples collected from different years are comparable.

To extract total environmental DNA, about 250 g of sediment was collected

(July 2009) approximately 50 mm below the surface, using an ethanol-sterilized scoopula, and placed in a sterile plastic sampling bottle. LH sediment (15 g) designated for RNA analysis were collected (July 2010) using an ethanol-sterilized spatula and stored in sterile 50 mL conical tubes filled with

LifeGuard™ Soil Preservation Solution (Mobio Laboratories, Inc, Carlsbad, CA) to a final volume of 50 mL. Both DNA and RNA samples were transported to

Montreal at temperatures below 5°C, where they were then stored at -20°C until further analyses.

3.2.2 Metagenomic DNA extraction and sequencing 68

DNA was extracted from 5 g of LH sediment using the PowerMax® Soil

DNA Isolation Kit (Mobio Laboratories, Inc, Carlsbad, CA, USA) following the manufacturers instructions. To obtain sufficient DNA (minimum of 500 ng DNA) for metagenomic pyrosequencing, the purified meta-DNA was amplified via multiple displacement amplification (MDA) using a GenomiPhi V2 DNA

Amplification Kit (GE Healthcare, Piscataway, NJ, USA) following the manufacturers instructions. One µL of DNA (concentration of less than 10 ng/ µL) was used as template and mixed with 9 µL of buffer. The mixed DNA was heated at 95 °C for 3 mins and then cooled to 4 °C followed by an incubation at 30 °C for

90 mins with 1 µL of enzyme mix and 9 µL of reaction buffer. To terminate the reaction, the sample was heated at 65°C for 10 mins. Control reactions were also performed in parallel using the provided kit positive control and sterile ddH 2O as a negative control. No DNA band was detected in the negative control sample following MDA amplification. The amplified samples were pooled and purified using Amicon Ultra-0.5 mL Centrifugal Filters (Millipore Corporation, MA, USA) to a final volume of 21 µL of solution containing 253.4 ng DNA µL-1. The purified sample was sequenced using a Roche 454 GSFLX Titanium sequencer (454 Life

Sciences, Branford, CT, USA), located at the Centre for Applied Genomics,

Hospital for Sick Children, Toronto, ON, Canada.

3.2.3 Metagenomic DNA analyses

To analyze and annotate the metagenomic data, all LH reads were uploaded to the online metagenomic annotation server, MetaGenome Rapid Annotation

with Subsystem Technology (MG-RAST) (Meyer et al., 2008). Based on the

BLAST-like alignment tool (BLAT algorithm) (Kent, 2002), metagenomic sequences were compared to those of gene and protein-coding databases; the

Genbank ( http://www.ncbi.nlm.nih.gov/genbank/ ) taxonomic database was used for the LH metagenome, while the SEED protein-coding gene database

(http://www.theseed.org/wiki/ index.php/Home_of_the_SEED ) served to compare with the putative coding proteins in the metagenome. Metabolic pathways were mapped using the Kyoto Encyclopedia of Genes and Genomes (KEGG; http://www.genome.jp/kegg/ ) database. For all used databases, only matches of over 50 nucleotides and 50% similarity with an E value ≤ 10 -5, were included for both taxonomy and function analyses. The cut-off stringency was tested with different E values (down to 10 -15 ) at the phylum and subsystem levels for taxonomy and functional classifications, respectively (Fig. S2-1). The correlations between different E values were tested by Pearson product-moment correlation coefficient (Table S2-1).

In addition to automated annotations by MG-RAST, the complete LH metagenome was subjected to additional screenings targeting marker genes of

(reverse) methanogenesis (i.e., the alpha subunit of the methyl coenzyme M reductase, mcrA), and methane oxidation (i.e., the alpha subunit of the particulate and soluble methane monooxigenase, pmoA and mmoX). Amino acid sequences of MCRA were recovered from the NCBI protein database (on February 16 2013) and used as target databases for alignments with the LH metagenome. BLASTX alignments were performed using the BLAST command line application (version

2.2.27+) with default algorithm parameters and an E-value cut-off of 10 -5. Results 70

were then visualised and proofread in MEGAN (version 4.70.4) and hits with Bit

Scores higher than 50 were considered significant (Huson et al., 2011). Reads of significant hits were then extracted and subjected to a second set of BLASTX alignments against the complete GenBank non-redundant (nr) database to ascertain their function and were finally re-annotated in MEGAN.

3.2.4 Statistical analyses

We selected 9 other metagenomes publically available in MG-RAST, generated from these habitats: the Markham ice shelf, the Ward Hunt ice shelf, an estuary of the Bay of Fundy, the Lost City hydrothermal system, a hypersaline lagoon in the Galapagos, a microbial mat from the McMurdo Ice shelf in the Ross

Sea sector of Antarctica, an Antarctic saline lake (Ace Lake), and high Arctic permafrost and active layer soils from Eureka, Ellesmere Island (MG-RAST ID:

4445126.3, 4445129.3, 4441582.3, 4461585.3, 4441599.3, 4445845.3, 4443684.3,

4443232.3 and 4443231.3, respectively). We also re-annotated the assembled

Alaskan permafrost metagenome (IMG ID: 1618) from Mackelprang and colleagues (Mackelprang et al., 2011) in MG-RAST to make it comparable in the

MG-RAST subsystem. The relative abundance at the “function” level of the Seed hierarchy were used to calculate Bray-Curtis distances between sample pairs using the “vegdist” function of the “vegan” package

(http://vegan.r-forge.r-project.org/) in R ( version 2.9.0, The R Foundation for

Statistical Computing). Principal coordinate analyses (PCoA) were then performed using the ‘cmdscale’ function. Arrows representing the relative

abundance at “level 1” of the Seed hierarchy were then superimposed on the ordination as supplementary variables, not involved in the calculation of the ordination (Yergeau et al., 2010).

3.2.5 RNA extraction and 16S ribosomal cDNA analyses

To obtain total RNA, sediment samples (2 g) from LH were processed with an RNA PowerSoil® Total RNA Isolation Kit (Mobio Laboratories, Inc, Carlsbad,

CA, USA) according to the manufacturer’s instructions with minor modifications as follows: (i) an additional 1.0 g of 0.1 mm glass beads (Mobio Laboratories, Inc,

Carlsbad, CA, USA) were added to each reaction tube, (ii) bead-beating time was doubled, and (iii) nucleotide precipitation was performed overnight. The extracted

RNA was then treated with amplification grade DNase I (Invitrogen, Carlsbad,

CA, USA) at room temperature for 15 minutes following the manufacturer’s instructions and then inactivated by the addition of EDTA at 65°C for 20 minutes.

The treated sample was concentrated and purified using Amicon Ultra-0.5 mL

Centrifugal Filters (Millipore Corporation, MA, USA). To synthesize cDNA, we used an iScript TM Select cDNA synthesis kit (Bio-Rad, Hercules, CA, USA) to process the purified RNA samples, using random primers provided in the kit. The

16S ribosomal cDNA was sequenced at the Research and Testing Laboratory

(Lubbock, Texas, USA) using a Roche 454 GSFLX Titanium sequencer (454 Life

Sciences, Branford, CT, USA) system with bacterial

(28F:5’GAGTTTGATCNTGGCTCAG’3, 519R:5’

GTNTTACNGCGGCKGCTG’3) (Handl et al., 2011) and archaeal (ARCH571F:

5’GCYTAAAGSRNCCGTAGC’3 (Baker et al., 2003), ARCH909R (a.k.a.

890aR): 5’TTTCAGYCTTGCGRCCGTAC3’ (Burggraf et al., 1997) primers. The tag-encoded pyrosequencing were performed following established protocols

(Bailey et al., 2010).

The ribosomal cDNA sequences were trimmed, aligned, and dereplicated using the RDP pyrosequencing pipeline (Cole et al., 2009). The minimum quality score was set to 20 and sequences shorter than 150 bp were excluded from downstream analyses. The trimmed sequences were aligned using the pyrosequencing aligner using Bacteria/Archaea model. The aligned sequences were then clustered using the complete-linkage clustering method with a maximum distance of 15% and step size of 1.0. Dereplicated sequences were generated using the representative sequence method with 98% similarity and then analyzed via the BLASTn algorithm against the online GenBank nr database.

(Altschul et al., 1997). Neighbor-joining phylogenetic trees of selected sequences were generated with MEGA 5 (Tamura et al., 2011), using a bootstrap method with 1,000 replications and a Jukes-Cantor model.

3.2.6 Nucleotide and metagenome sequence accession numbers

The 16S ribosomal cDNA gene sequences obtained in this study have been deposited in the GenBank database under accession numbers KC470367 -

KC470541 . The ID number of the LH metagenome deposited in MG-RAST is

4478244.3.

3.3 Results and Discussion

3.3.1 Metagenomic sequencing statistics

In total, sequencing resulted in 1,032,783 reads containing 341,472,858 bases, with an average length of 330 bp (Table 3-1). Among all sequences,

751,870 reads (72.8%) passed the quality control (QC) criteria. The predicted protein and rRNA features were 403,739 (39.1%) and 50,589 (4.9%) reads, respectively. Out of these putative protein and rRNA related reads, 187,609

(18.2%) and 220 (2.1%) matched known protein and rRNA sequences, respectively. However, 434,847 (42.1%) reads remained unidentified due to a lack of comparable reference sequences highlighting the need to further isolate, characterize, and genome sequence new strains from extreme environments such as the LH spring to complement existing databases making the annotation of the reads from extreme environments more reliable and informative. The average GC content was 45% for the sequences that passed quality control. The total Duplicate

Read Inferred Sequencing Error Estimation (DRISEE) error was 0.7% (Keegan et al., 2012), which was deemed acceptable to proceed in further analyses. The

Pearson product-moment correlation coefficients of the cut-off E-value (≤ 10 -5) with lower E-values (10 -10 , 10-15, and 10-20 ) showed that the correlations were highly significant down to 10-15 and 10 -20 for taxonomic and functional classifications at phylum and subsystem levels, respectively (Table S3-1). Thus, it proved that the E-value cut-off we set was appropriate for applying to this metagenome study.

3.3.2 Metagenomic microbial community composition

Among the total 751,870 sequences which passed the QC criteria, 50.8%,

5.1% and 0.8% were identified as Bacteria-, Eukaryota- and Archaea-originating fragments, respectively. A total of 719,330 sequences (95.7% of the total sequences which passed QC) could be classified and assigned to different phyla by MG-RAST. Sequences related to Cyanobacteria (19.7%), Bacteroidetes

(13.3%), and Proteobacteria (6.6%) were the dominant phyla among the classified sequences (Table 3-2). Over 90% of Cyanobacteria hits belonged to the orders Nostocales (35.5%), Oscillatoriales (23.0%), and Chlorococcales (38.4%).

The abundance of Cyanobacteria related sequences suggest that these microorganisms could potentially carry out photosynthesis, carbon fixation, and nitrogen fixation metabolism at LH. All five classes of Proteobacteria were also detected in the LH metagenome with Gammaproteobacteria (47.6%),

Betaproteobacteria (30.7%) and Alphaproteobacteria (12.4%) being the major groups present. Methylophilic and methanotrophic genera were detected in both the Gammaproteobacteria and Betaproteobacteria . Gene fragments related to the ammonia-oxidizing orders Nitrosomonadales were identified in the metagenome, which may provide evidence for bacterial ammonia oxidizing activity at LH in accordance to the presence of ammonia. In terms of sulfur metabolism, clades related to bacterial sulfate reducers, such as Desulfuromonadales ,

Desulfovibrionales , and Desulfobacteriales , were detected within the

Deltaproteobacteria and Betaproteobacteria , and whose metabolic activity would

be favourable with the high abundance of sulfate and sulfide, as well as the reducing conditions, detected at the site.

The small proportion of total archaeal reads within the metagenome were represented by Euryarchaeota (0.5%, 4,037 hits), Crenarchaeota (0.05%, 351 hits), and Thaumarchaeota (0.01%, 75 hits) (Table 3-2). Among the

Crenarchaeota , only Thermoprotei (343 hits) and unclassified Crenarchaeota (8 hits) were detected, while for Thaumarchaeota , sequences related to unclassified

Thaumarchaeota (75 hits) were found. These hits for Crenarchaeota and

Thaumarchaeota were primarily related to genes involved in DNA duplication, transcription, translation, and electron transport. The hits for Euryarchaeota were more varied; Archaeoglobi (213 hits), Halobacteria (420 hits), Methanomicrobia

(1,878 hits), Methanobacteria (420 hits), Thermoccoci (323 hits), Methanococci

(381 hits), Methanopyri (14 hits), Thermoplasmata (85 hits) and unclassified

Euryarchaota (242 hits) were identified. As well, the functional gene categories associated with these hits were more diverse, with the most abundant ones being related to potassium channel proteins (Potassium metabolism; 145 hits), cold-shock DEAD-box protein A (RNA metabolism; 84 hits),

O-phosphoseryl-tRNA:cysteinyl-tRNA synthase (Protein metabolism; 76hits), and lysyl-tRNA synthetase (Protein metabolism; 41 hits).

Diverse methanogenic genera were present within the Euryarchaeota dataset, including Methanobrevibacter (29 hits), Methanothermobacter (342 hits),

Methanothermus (21 hits), Methanococcus (226 hits), and Methanocorpusculum

(39 hits). The detection of these methanogen-related sequences support the idea that at least a small portion of the methane exsolving from LH spring may be 76

partly biogenic although previous carbon and hydrogen isotope analyses indicated that the LH methane is thermogenic in origin (Niederberger et al., 2010). The presence of Methanobrevibacter was also reported in the LH outflow channel area using 16S rRNA clone library (Lay et al., 2012). Methanogenic populations have also been found in other cold extreme environments, e.g., Canadian High Arctic and Alaskan permafrost, melting glaciers, and other AHI saline springs (Perreault et al., 2007; Walter-Anthony et al., 2012; Yergeau et al., 2010), and have been shown to remain active down to -16.5°C in Siberian permafrost (Rivkina et al.,

2004). Although hypersaline conditions are known to inhibit acetoclastic and hydrogenotrophic methanogenesis above ~12% NaCl, methanogens relying on

“non-competitive” substrates (i.e., methylated amines, methanol, or dimethylsulfide) can withstand higher salt concentrations, with reports of methanogenesis at salinities of 30% in endoevaporite communities (Oren, 2011;

Tazaz et al., 2012). Methanogenesis typically requires a lower redox potential than most other anaerobic bioreactions; considering the anoxic and highly reducing conditions in the LH spring sediments, methanogens can be expected to be present and active in this ecosystem.

In our previous study of the LH spring (Niederberger et al., 2010), the anaerobic methane-oxidizing archaea group 1 (ANME-1) was the dominant archaeal clade (46.8%) detected, based on archaeal 16S rRNA clone library results.

Although we detected archaeal 16S rRNA sequences identical to the previously identified LH ANME-1 sequences in the MDA-amplified LH DNA prior to metagenome pyrosequencing, no ANME-1 16S rRNA was detected after the metagenomic analysis which we suspect was due to their population’s low 77

abundance. Searches for ANME-1-related DNA fragments using unordered contigs of an ANME-1 genome (Genebank database, accession No.: FP565147)

(Meyerdierks et al., 2010). BLAST analyses in MG-RAST (version 2.0) resulted in 1,000 hits within the LH metagenome with sequence similarities of 80% to

97% to ANME-1-related sequences. Functional annotation by BLASTx against the NCBI nr database classified these ANME-1 related hits as genes encoding integrase core domain, FDA (flavin adenine dinucleotide)-containing dehydrogenase, Fe-S oxidoreductase related to Leucyl-tRNA synthetase, and hypothetical proteins. Since the identity of the matches between the LH metagnome sequences and the ANME-1 genome (FP565147) was generally low,

LH ANME-1s most likely belong to a different subgroup than the one with the published genome (FP565147). However, we still need more evidence to support this assumption and to better understand the ANME-1 previously detected at the

LH spring sediments.

3.3.3 Functional gene profiles of the LH metagenome

The functional gene profile revealed that among the 259,557 annotated protein sequences (25.1% of the total reads), the most abundant functional groups were related to house-keeping functions, such as carbohydrate metabolism

(10.1%), amino acid biosyntheses (10.0%), and vitamin and pigment metabolism

(6.6%). Stress-response-related sequences comprised 2.3% of all annotated reads and these included a high proportion of oxidative stress (53.1%) and osmotic stress (11.9%)-related sequences. The abundance of these genes may reflect

adaptations to the high salinity and possibly, the high salinity induced oxidative stress at LH, indicating that the LH microorganisms have the potential to survive and remain viable under these prevailing conditions. Descriptions of genes involved in methane, nitrogen, and sulfur metabolism, as well as stress responses are discussed in following paragraphs.

3.3.4 Methane metabolism

Several functional genes directly related to methanogenesis were detected

(Table 3-3). These include: an F420-dependent methylene-H4 MPT reductase (EC

1.5.99.11; 1 hit), formylmethanofuran dehydrogenases (fmd) (EC 1.2.99.5; 2 hits),

CoB-CoM heterodisulfide reductases (EC 1.8.98.1; 2 hits), F420-reducing hydrogenases (EC 1.12.98.1; 30 hits), and methylenetetrahydromethanopterin dehydrogenases (mer) (EC 1.5.99.9; 2 hits). We could not confirm the presence of the gene encoding methyl-coenzyme M reductase (MCR), which dominates the last step of methanogenesis, in the LH metagenome. The potential (MCR) homologs were identified by an additional search of the metagenome against an

MCR target database, but their potential functions mostly were close to ABC transporters/ATP-binding proteins. With the low abundance of other methanogenesis-related genes in the LH metagenome, the absence of mcr gene may be due to insufficient sequencing coverage. Another metagenomic study regarding deep subsurface marine sediments described similar results, which reported the recovery of genes involved in methanogenesis without detected mcr sequence (Teske and Biddle, 2008).

3.3.5 Nitrogen metabolism

Most genes involved in nitrogen cycling pathways were detected and mainly related to Cyanobacteria (Figure 3-1a, Table S3-2). The nifH gene (EC

1.18.6.1), which encodes a nitrogenase that converts nitrogen gas to ammonia, was detected and matched sequences from Cyanothece and Nostoc , cyanobacterial species. Sequences related Burkholderia , typical denitrifiers in saline environments (Ferrer et al., 2011; Steward et al., 2004), were detected for two enzymes, narG (EC 1.7.99.4) and nirS (EC 1.7.2.1). Denitrifiers usually found in saline and fresh water environments including Kangiella spp. and Flavobacterium spp. (Auclair et al., 2012; Qu et al., 2009), respectively, were also detected in the

LH metagenome in genes involved in denitrification pathways. Two enzymes (EC

1.7.1.4 and 1.7.7.1) involved in the reduction of nitrite to ammonia were matched to Cyanobacteria , such as Synechocystis spp., Cyanothece spp. and Nostoc spp.

Synechocystis and Cyanothece have been reported to undergo heterotrophic metabolism during the dark phase of their life cycle (Reddy et al., 1993; Vernotte et al., 1992), which may be advantageous to maintain activity during long term darkness of the Arctic winter. Three enzymes that play key roles in nitrogen cycling, nitric oxide reductase (EC 1.7.99.7), ammonia monooxygenase (EC

1.13.12.4), and hydroxylamine oxidase (EC 1.7.3.4), were absent from the LH metagenome. As has been observed in other environments, the function of nitric oxide reductase may be replaced by abiotic processes (Venterea, 2007). Although

16S phylogenetic evidence of thaumarchaea were detected previously (Lay et al.,

2012) and in the 16S ribosomal cDNA (see below), the absence of ammonia monooxygenase and hydroxylamine oxidase indicates that the complete ammonia oxidation pathway could not be reconstructed from the LH metagenome and our analyses cannot yet confirm AOA/B (ammonia oxidizing archaea/bacteria) metabolic activity within the LH spring sediments. We have also not been able to detect ammonia oxidation in flask enrichments of LH outlet and channel sediments but we have cloned Thaumarchaeal amoA genes from the LH spring channel sediments (unpublished data). Thus, the occurrence of in situ ammonia oxidation at LH has not yet been experimentally confirmed.

3.3.6 Sulfur Metabolism

A complete sulfur cycle through reduction and oxidation between sulfur end members and intermediates was identified in the LH metagenome over a high diversity of taxa (Figure 3-1b, Table S3-3). The enzymes driving sulfate reduction, including sulfate adenylyltransferase (EC 2.7.7.4; 222 hits), adenylyl-sulfate kinase (EC 2.7.1.25; 172 hits), phosphoadenylyl-sulfate reductase (EC 1.8.4.8; 41 hits), and sulfite reductase (EC 1.8.7.1; 112 hits) were all detected; however, pathways were not completely reconstructed for all of the species identified

(Figure 3-1b). A large number of sulfur oxidation genes (soxB/D/H/R genes) were recovered (302 hits) with dominant taxa including Thiomicrospira (20% of all sox reads), Thiobacillus , Nitrosococcus , and Roseiflexus . The enzymes for both assimilatory (EC 1.8.99.1) and dissimilatory (EC 1.8.99.3) sulfate reduction were also found, with assimilatory pathways appearing more abundant. The role of

anoxygenic photosynthetic green and purple sulfur bacteria was prominent with the abundance of hits to Chlorobium spp., a typical anoxygenic phototrophic sulfide oxidizer, Roseiflexus spp., a filamentous low concentration sulfide oxidizer, and Chloroflexus spp., a species containing metabolic features of both purple and green sulfur bacteria, all of which have been found in spring, hypersaline, or sulfur-rich ecosystems (Kompantseva et al., 2005; Nubel et al., 2001). Several potential metabolic linkages between the LH nitrogen and sulfur cycles are plausible with the abundance of hits to species including Thiobacillus denitrificans (54 hits), Alkaliminicola sp. (9 hits), and Nitrosococcus (7 hits).

Thiobacillus denitrificans couples the oxidation of inorganic sulfur compounds to the reduction of oxidized nitrogen compounds (such as nitrate, nitrite) to dinitrogen (Kelly et al., 1997). Alkaliminicola sp., an anaerobic facultatively autotrophic arsenite oxidizing bacterium, respires nitrate or nitrite, or alternatively, uses sulfide or thiosulfate as electron donors. Nitrosococcus is a group of ammonium-oxidizing purple sulfur bacteria (Table S3-3), which oxidizes ammonia to nitrite and reduces sulfate to sulfide (Klotz et al., 2006). The detection of these genes and species involved in both nitrogen and sulfur cycles provides evidence that these two cycles may be synergistically linked by similar species in the LH system.

3.3.7 Stress response

The presence of stress-response-related gene fragments (5,690 hits), which were all associated with bacterial taxa, likely reflected the potential of LH bacteria

to deal with or adapt to stressors in this hypersaline and subzero habitat. Given the stable LH-spring-sediment environment, these genes may be more adaptive than being typical “stress-response” genes, i.e., cold or heat shock . For example, many cold shock proteins can also be characterized as cold acclimation proteins, i.e., present at relatively high levels during growth at constant cold temperatures(Feller and Gerday, 2003).

The three most abundant groups corresponded to oxidative stress (2,255 hits), heat shock (1,491 hits) and osmotic stress (987 hits), all possibly linked to natural stressors at LH. LH outlet sediments are highly reducing and microoxic but are exposed to periodic exposure to the air during the summer months. The genes related to oxidative stress in the LH metagenome were mainly associated with Bacteroidetes , Proteobacteria and Cyanobacteria (Table S3-4).

Anti-oxidative stress genes may also help in responding to sudden changes in oxygen concentrations under predominantly anoxic conditions (Briolat and

Reysset, 2002; Jean et al., 2004; Kawakami et al., 2004; Rocha et al., 1996).

Many of the identified genera encoding these enzymes were aerobes, indicating that they may be dormant, or periodically active, in the LH spring. The presence of genes related to anti-oxidative stress may also be attributed to salinity-induced anti-oxidative defense responses as high salinity may also provoke the formation of reactive oxygen species (ROS) as a byproduct of energy metabolism, including photosynthesis (Srivastava, 2010). For example, the expression of antioxidant enzymes in response to salinity has been observed in Cyanobacteria in Nostoc and Synechocystis species (Latifi et al., 2009; Srivastava, 2010), and ROS genes related to these two genera were indeed present in the LH metagenome. Lastly, 83

antioxidant defense against ROS is a requirement of growth for some species at low temperatures due to a decrease in requirement of ATP, which results in the electron accumulation of respiratory chain and therefore increase in ROS at cold temperatures (Chattopadhyay, 2002).

Most hits related to osmotic stress involved compatible-solute adaptations, typical in halophilic bacteria. Most hits (767 hits out of 978 hits) were related to the synthesis of the osmoregulated periplasmic glucan; the gene related to this synthesis may respond to sudden changes in salinity (Bohin, 2000), and 131 hits were affiliated to choline and betaine biosyntheses (Table S3-5). Two enzymes involved in betaine synthesis, choline dehydrogenase (from Cyanobacteria ,

Actinobacteria , and Gammaproteobacteria ) and betaine-aldehyde dehydrogenase

(from fungi and Gammaproteobacteria ), were detected, suggesting that betaine might be the main osmolyte used by the LH microbial community. Betaine is a well-known osmolyte that plays an important role to balance the high osmotic pressure exerted on microbial cells in hypersaline environments (Sleator and Hill,

2002). Other typical adaptive responses to osmotic stress, such as sodium transporters, which are usually used by halophilic archaea to balance the osmotic pressure inside and outside the cells, were not detected.

The heat-shock-protein genes present in the metagenomic dataset probably do not reflect heat-shock responses in the permanently cold LH spring as these genes were more related to the general chaperone protein dnaK (640 hits) and its interacting protein, dnaJ (31 hits); these proteins are prevalent in microorganisms in cold environments for assisting with protein folding (Panoff et al., 1995; Ting et al., 2010). 84

Microorganisms that sustain metabolic processes at cold temperatures produce cold acclimation and cold shock proteins involved in DNA replication

(gyrA, recA, and dnaA) (Atlung and Hansen, 1999; Merrin et al., 2011; Yamanaka,

1999), transcription (nusA), RNA unwinding (cold shock DEAD-box protein A

(csdA) and cold shock protein A (cspA) (Bakermans et al., 2009; Hunger et al.,

2006; Jones et al., 1996; Py et al., 1996; Ray et al., 2010), protein folding (prolyl isomerase) (Cavicchioli et al., 2000; de la Cruz et al., 1999; Suzuki et al., 2004;

Yamanaka, 1999), pyruvate metabolism (aceE and aceF) (Scherer and Neuhaus,

2006; Wouters et al., 2001), and unsaturated fatty acid metabolism (fatty acid desaturases, dnaJ) (Kenny et al., 2009; Rodrigues and Tiedje, 2008; Thieringer et al., 1998; Varin et al., 2012). Genes encoding for such proteins were identified as containing features of cold adaptation within the LH metagenome, mostly originating from Proteobacteria , Bacteroidetes and Cyanobacteria (Table S3-6).

These genes were also detected in these three phyla in both Antarctic and Arctic metagenomic samples from the McMurdo Ice Shelf, the Ward Hunt Ice Shelf, and the Markham Ice Shelf (Varin et al., 2012). The ubiquity of these genes among similar taxa in other polar habitat metagenomes strongly suggests that such adaptive genetic systems are a global feature in cryoenvironments.

3.3.8 Comparison with other metagenomes

Based on the relative abundance of functional genes in different MG-RAST subsystems, we created an ordination of the LH metagenome together with other metagenomes of cold and salty environments. The ordination produced by PCoA

(Figure 3-2) shows the similarity between samples (the closer the sample, the more functionally similar they are). The subsystem arrows are pointing toward the samples that have the highest relative abundance of this particular subsystem. The

PCoA revealed that the LH metagenome clustered with the metagenomes from permafrost and active layer samples from the Canadian high Arctic and was clearly distinct from the other samples. These patterns can perhaps be explained by higher relative abundance of genes related to dormancy and sporulation, reflecting possible adaptations to the extreme conditions of the LH spring and

Arctic permafrost environments at Eureka (Nicholson et al., 2000;

Sachidanandham and Yew-Hoong Gin, 2009), which are from the same geological region, about 70 to 80 km away from each other. A total of 82,711 genetic hits

(Cyanobacteria , 66.3%; Bacteroidetes , 30.0%; Proteobacteria , 3.5%; Firmicutes ,

0.2%) related to dormancy and sporulation were present in the LH metagenome.

In addition, the LH spring water flows through permafrost before reaching the surface, which would result in the transfer of permafrost microbial communities to the LH spring outlet sediments. Although the Markham ice shelf and the Ward

Hunt ice shelf are also located in the same geographical region, the biomat metagenomes from these two ice shelves were not closely related to the LH metagenome, probably due to the the difference in habitat conditions between the ice shelve mat habitats (freshwater, oxic) being different than the LH spring system. The metagenome originating from the biofilm of the deep-sea-hydrothermal field Lost City, was also associated relatively closely with

LH, perhaps reflecting similarities with respect to methane and oxygen concentrations, and sulfur metabolism (Brazelton and Baross, 2009; Brazelton et 86

al., 2006; Niederberger et al., 2010).

3.3.9 Active profiling of LH based on 16S ribosomal cDNA pyrosequencing

In an attempt to reveal which microorganisms are active at LH spring, we pyrosequenced the reverse-transcribed bacterial and archaeal 16S ribosomal RNA gene from extracted sediment RNA and compared the obtained 16S ribosomal cDNA profile with the LH metagenomic dataset. Comparing both datasets revealed similar trends for bacterial and archaeal abundance at the phylum and class levels (Figure 3-3, Figure S3-4 and S3-5). Regarding bacterial abundance,

Bacteroidetes , Proteobacteria , Firmicutes , Actinobacteria , and Verrucomicrobia were the most abundant phylotypes detected in the 16S ribosomal cDNA dataset

(Figure 3-3a, Figure S3-4). Whereas the Gammaproteobacteria , Actinobacteria, and Verrucomicrobia were over-represented by more than a 50 % increase in relative abundance in the 16S ribosomal cDNA dataset when compared to the metagenome , the Bacteroidetes and Firmicutes showed a relative decrease (Figure

3-3a). The Gammaproteobacteria consisted of four genera, ( Marinobacter,

Stenotrophomonas, Pseudomonas, and Enterobacter ) in the 16S ribosomal cDNA library (Table S3-7). The reads belonging to the same orders as these four genera

(i.e., orders Alteromonadales , Pseudomonadales , Xanthomonadales , and

Enterobacteriales ) comprised 60.0% of the total gammaproteobacterial reads in the LH metagenome. The high proportion of gammaproteobacterial reads present in the 16S ribosomal cDNA library, and the taxonomic similarities between those reads and the ones present in the metagenome, suggest that the LH

Gammaproteobacteria may be adapted to the hypersaline and cold conditions of the site and as such, active in situ . It should be pointed out that Marinobacter and

Pseudomonas species have also been detected and isolated at other perennial cold saline springs on AHI (Niederberger et al., 2010; Perreault et al., 2007; Perreault et al., 2008).

A striking difference between both datasets, however, was that no

Cyanobacteria reads were detected with cDNA pyrosequencing, while they were the most abundant phylum in the LH metagenome. This difference might have been caused by amplification biases as the primer pair used for cDNA pyrosequencing only matched 61% of the Cyanobacteria sequences present in the

RDP database used for annotation. Yet since this primer pair can amplify reads of

Synechococcus , Cyanothece and Trichodesmium (which accounted for 41.3% of the total cyanobacterial reads in the LH metagenome), the results obtained in the

16S ribosomal cDNA library suggest that Cyanobacteria were probably dormant at LH, which would be supported by the finding that up to 66.3% of the genes related to dormancy found in the LH metagenome were of cyanobacterial origin.

The bacterial 16S ribosomal cDNA dataset also contained microorganisms involved in nitrogen cycling. For example, Pseudomonas and

Stenotrophomonas -related sequences detected in the 16S ribosomal cDNA dataset suggest that denitrification may be driven by these microorganisms (Figure S3-2).

Sequences associated with Desulfobulbus suggest that active sulfate reduction occurs within the LH outlet sediments (Figure S3-2).

Verrucomicrobia sequences were also detected in both datasets (Figure 3-3 and S3-2) and occupied an important/large proportion (16.8%) of the bacterial 88

cDNA library (Figure S3-4). However, their potential function could not be determined; no sequence of verrucomicrobial pmoA gene was present in the LH metagenome, nor did the verrucomicrobial sequences from the 16S cDNA library cluster with any known culture representatives (Figure S3-6). Therefore, the ecological status of the LH Verrucomicrobia is unknown.

Regarding archaeal abundance, the metagenome dataset included all five known archaeal phyla, including Korarchaeota and Nanoarchaeota . On the other hand, the 16S ribosomal cDNA dataset only comprised sequences related to

Euryarchaeota (20.1%), Crenarchaeota (42.4%), and Thaumarchaeota (37.5%)

(Fig. 3-3b and S3-5; Table S3-8). Although Euryarchaeota sequences were the most abundant ones in the archaeal metagenome dataset, Crenarchaeota and

Thaumarchaeota related sequences dominated the archaeal 16S ribosomal cDNA library, supporting some metagenomic interpretations of active S, N, and C biogeochemical cycles in situ . The detection of methanogenic archaea in both the metagenomic and the 16S archaeal ribosomal cDNA datasets suggests that at least some of the methane exsolving from the LH outlet is biogenic (Table S3-8).

The high proportion of Crenarchaoeta in the 16S ribosomal cDNA library was unexpected, considering that they usually represent hyperthermophilic archaea. These sequences had greater sequence identities to the thermophilic crenarchaea than to the “mesophilic crenarchaea”, which were re-classified as

Thaumarchaeaota after 2008 (Brochier-Armanet et al., 2008). Thus, the detection of these thermophilic crenarchaea at LH provides evidence of cold-adapted crenarchaea at LH. The crenarchaeal phylotypes recovered from 16S ribosomal cDNA were most closely related to Thermoprotei species [Vulcanisaeta sp. 89

(Identity = 97%), Thermoproteus sp. (Identity = 98%), and Pyrobaculum sp.

(Identity = 99%); (Table S3-8, Figure S3-5)]. All of these genera are anaerobic sulfur/sulfide oxidizing microorganisms and may participate in sulfur cycling at

LH (Itoh et al., 2002; Selig and Schönheit, 1994; Strauss et al., 1992). Among these, Thermoproteus , a sulfide oxidizer, may also be involved in inorganic carbon fixation; Pyrobaculum , a sulfur oxidizer and denitrifier, may utilize sulfur as an electron acceptor anaerobically (Selig and Schönheit, 1994; Strauss et al.,

1992). In addition, it was surprising that no halophilic archaea were detected in the active community profile considering the hypersalinity in situ , and that they were previously detected (Lay et al., 2012; Niederberger et al., 2010). This result perhaps reflects extraction and/or amplification biases. On the other hand, halophilic archaea are mostly aerobic microorganisms and might not be suitable of inhabiting the microoxic sediments of the LH outlet, indicating that the halophilic archaea were inactive in the LH sediments.

3.4 Conclusion

Here we report the first metagenomic and 16S ribosomal cDNA pyrosequencing study of the unique Lost Hammer hypersaline subzero spring.

Metagenomic data combined with 16S ribosomal cDNA analyses provides insights into the complex metabolic and functional genes present in this extreme hypersaline cryoenvironment where active C, N, and S cycling appear to be functioning in situ and probably synergistically. The metagenomic dataset also contains a large diversity of genes belonging to taxa not known to inhabit 90

cryoenvironments and may reflect a large pool of dormant microorganisms or novel cold-adapted representatives of these taxa, such as in Crenarchaeota , capable of surviving in halophilic cryoenvironments. An abundance of active aerobic taxa also supports that seasonal or microoxic environments may support the activity of allochtonous bacteria and archaea that are deposited in the anoxic

LH spring sediments. Genes involved in oxidative and osmotic stress, present in relatively high abundance as respond to other stress-related genes, probably represent adaptations to the multiple stressors intrinsic of subzero temperatures and high salinity in this habitat. Overall, investigations of the hypersaline subzero

LH spring system expand our knowledge of microbial life in extreme cryoenvironments on Earth (Lay et al., 2012; Niederberger et al., 2010) and provide evidence of how microbial life could inhabit subzero briny environments thought to exist on Mars (McEwen et al., 2011) and Enceladus (Postberg et al.,

2011).

3.5 Acknowledgements

This work was supported by the Canadian Astrobiology Training Program

(CATP), National Sciences and Engineering Research Council of Canada

(NSERC), Canadian Space Agency (CSA), Fonds de Recherche du Québec -

Nature et Technologies (FQRNT), Canada Foundation of Innovation (CFI), Polar

Continent Shelf Program (PCSP), and Northern Scientific Training Program

(NSTP). The Alaskan permafrost sequence data was produced by the US

Department of Energy Joint Genome Institute (http://www.jgi.doe.gov/) in 91

collaboration with the user community.

Table 3-1. The statistical analyses of LH metagenome

Total no. of sequences 1,032,783

Total sequence size (bp) 341,472,858

Shortest sequence length (bp) 40

Longest sequence length (bp) 918

Average sequence length (bp) 330

Sequences passed QC 751,870

Predicted/identified protein features 403,739/187,609

Predicted/identified rRNA features 50,589/220

GC content 45%

Table 3-2. The composition of organisms detected in the LH metagenome Hits %

Unclassified reads 322,498 42.9

Archaea 6,020 0.8

Euryarchaeota 4,037 0.5

Crenarchaeota 351 <0.1

Thaumarchaeota 75 <0.1

Other Archaea 1,557 0.2

Bacteria 382,205 50.8

Cyanobacteria 148,325 19.7

Bacteroidetes 99,693 13.3

Proteobacteria 49,803 6.6

Firmicutes 4,803 0.6

Actinobacteria 3,469 0.6

Verrucomicrobia 1,615 0.2

Other bacteria 74,470 9.9

Eukaryota 38,233 5.1

Others 2,914 0.4

Total 751,870

Table 3-3. Numbers of different gene variants retrieved in the LH metagenomic data sets for different functions Function Hits

Methane

F420-Dependent methylene- H4 MPT reductase (EC 1.5.99.11) 1

F420-reducing hydrogenase (EC 1.12.98.1) 30

CoB-CoM Heterodisulfide reductase (EC 1.8.98.1) 2

Formylmethanofuran dehydrogenase (EC 1.2.99.5) 2

Methylenetetrahydromethanopterin dehydrogenases (EC 1.5.99.5) 2

Nitrogen

Nitrogenase (EC 1.18.6.1) 2

Copper-containing nitrite reductase (EC 1.7.2.1) 45

Nitric-oxide reductase (EC 1.7.99.7) 14

Nitrite reductase (NAD(P)H) (EC 1.7.1.4) 48

Nitrous-oxide reductase (EC 1.7.99.6) 16

Respiratory nitrate reductase (EC 1.7.99.4) 4

Sulfur

Phosphoadenylyl-sulfate reductase (EC 1.8.4.8) 41

Sulfite reductase (ferredoxin) (EC 1.8.7.1) 112

Sulfate adenylyltransferase (EC 2.7.7.4) 222

Adenylyl-sulfate kinase (EC 2.7.1.25) 172

Sulfur oxidation genes (sox BDHR) 302

Fig. 3-1. Phylogenetic profiles for key enzymes in (A) nitrogen cycling and (B) sulfur reduction and oxidation. Bar graphs depict percent abundance of genera for each category of enzymes. The number of reads annotated from the metagenome is shown in brackets. The solid and hollow arrows indicate the presence and absence of the enzymes, respectively. The dotted line showed in Sox pathways indicated that the steps are more complicated than shown. 96

Fig. 3-2. Functional community composition of the LH spring sediment and other extremely cold or saline environments, based on principal coordinate analysis of the relative abundance of all MG-RAST subsystems. Comparative sites include the Markham, the Ward Hunt (Arctic) and the McMurdo ice shelves (Antarctica);

Alaskan and Eurekan (Canadian Arctic) permafrost, the Lost City, a hypersaline lagoon in the Galapagos Island, the Bay of Fundy, and Ace Lake (Antarctica).

Fig. 3-3. The proportions of different clades of (A) bacterial and (B) archaeal reads from LH metagenome and 16S ribosomal cDNA libraries. The classification was based on phyla with exception of the Proteobacteria shown at the class level.

The total reads of bacteria were 662,411 and 1,092 in the LH metagenome and

16S ribosomal cDNA library, respectively. The total reads of archaea were 943 and 8,604 in the LH metagenome and 16S ribosomal cDNA library, respectively.

CONNECTING TEXT

After establishing the background microbial profiles for the LH channel sediment, in the chapter 4, we tried to characterize the active microbial communities inhabiting the channel and the adjacent tundra. Further, we quantified the abundance of the Thaumarchaeal featured genes, amo A and hcd , in both channel sediment and the tundra soil. The partial amo A and hcd were sequenced and then translated into amino acid sequences for phylogenetic study.

CHAPTER 4

Seasonal Changes in Microbial Communities at a Hypersaline Spring Channel and the Adjacent Tundra

ABSTRACT

The Lost Hammer (LH) spring channel is notable for the hypersalinity and the unfrozen stream flow at -18°C. In our previous study, we detected heterotrophic microbial metabolic activity (CO 2 mineralization recovery of 0.17%) at -20°C. The pyrosequencing libraries of the channel sediments and adjacent tundra sampled from summer (July) and winter (Early Arctic Spring, April) were present using total microbial RNA and DNA. We analyzed the microbial compositions of active (RNA) and DNA communities inhabiting the system. In the summer, the LH channel sediment was dominated by the active groups,

Alphaproteobacteria and Betaproteobacteria . In the winter, Cyanobacteria ,

Gammaproteobacteria , Verrucomicrobia , and Firmicutes were the highly expressed bacteria in the LH channel. The results showed that the bacterial community shift happened in between the two seasons. From the 16S rDNA 99

pyrosequencing library comparison using UniFrac, the analysis showed sampling seasons to be the most significant variant affecting the microbial composition in the sediments and tundra. Signature genes, amo A and hcd , of ammonia oxidizing archaea were sequenced and analyzed for the phylogenetic affiliations with other published ones using their putative amino acid sequences from other environments. The result of phylogenetic tree showed similar patterns of grouping as 16S rDNA, especially using amoA. The amoA, hcd and the Thaumarchaeal 16S rDNA genes were quantified by qPCR in both sediment and tundra samples to support the genetic information regarding the LH Thaumarchaea. The copy numbers of Thaumarchaeal amo A and hcd genes in LH channel sediment and the adjacent tundra were roughly 10 to 100 folds less than those reported in other similar environments. Besides providing knowledge regarding microorganisms present in extreme environments, especially hypersaline and cold ones, this study presents an analogous model for studying potential microbial life forms on extraterrestrial bodies.

4.1 Introduction

The Lost Hammer (LH) spring, located on Axel Heiberg Island, is characterized by its hypersalinity, perennial subzero temperatures, microoxic and sulfate-rich environment, in addition to its methane emissions. Compared to the microoxic spring outlet itself, the oxic downstream channel differs in terms of temperature as well as having greater redox potential and dissolved oxygen value

(> 1 ppm) (Lay et al., 2012). In addition, the methane emissions were not detected 100

at the channel, but it contains relatively low methane concentrations in the sediments (about 1/10 of the outlet) (Lay et al., 2012). The channel passes through the tundra and leads the spring water from the outlet to an adjacent river. In the winter, the channel water is known to remain unfrozen down to -18°C as a result of the hypersaline outflow from the spring outlet (Lay et al., 2012). Based on analyses using 16S rDNA clone libraries, these similar but non-identical environments, the outlet and the channel, have distinct microbial profiles and microbial activities (Lay et al., 2012). In the previous study, based on

CARD-FISH microscopy, both bacterial and archaeal abundances were 100 times higher in the channel sediments than in the outlet. Bacterial diversity in the LH channel, evaluated with the Shannon index, was observed to exceed that of the outlet (Shannon indices of bacteria, channel: 1.69 to 3.8, outlet: 1.65). However, the archaeal diversity at the channel sediment was lower or close to the outlet sediments (Shannon indices of archaea, channel: 0.78 to 1.37, outlet, 1.39) (Lay et al., 2012; Niederberger et al., 2010). The dominant microbial species of channel sediments differed from those at the spring outlet particularly in terms of the archaeal community. The Thaumarchaea was the dominant group in channel sediments while ANME-1 was dominant at the outlet (Lay et al., 2012;

Niederberger et al., 2010). The result suggests that, in the channel sediments,

Thaumarchaea may play a key role for driving nitrogen and carbon cycles due to its ability for oxidizing ammonia and fixing inorganic carbon (Dang et al., 2013;

Zhang et al., 2010). For the bacterial communities, the channel sediments hosted similar bacterial groups to the outlet (Lay et al., 2012; Niederberger et al., 2010).

In view of the channel flow remaining unfrozen at -18°C, heterotrophic microbial 101

activity of LH channel sediments was measured at -20°C using 14 C-labeled acetate

(Lay et al., 2012). As an environment with these unique properties, the LH spring system is not only a model site for understanding microbial ecology at extreme saline and subzero environments, but also an extraterrestrial analog. The physical properties of the channel are similar to those leading putatively to the formation of salt deposits at a gully site on Mars. The salt deposits on Mars would imply the prior presence of liquid water (Malin et al., 2006) having provided an adequate environment for microorganisms to inhabit. Also, the presence of methane and ammonia with a saline liquid shows properties similar to the plume on Saturn’s moon Enceladus (Kerr, 2011; McKay et al., 2012). These characteristics support the working hypothesis of the LH system as an analogue to extraterrestrial potential ecosystems. The LH channel is important for comparison with the

Martian gully structures due to the salt deposits.

According to the previous study on the LH channel (Lay et al., 2012), we know the microbial compositions and the bulk heterotrophic activities at subzero temperatures at the LH channel. However, questions remain about the LH channel sediments: 1) the channel water flows above the tundra soils; what is the similarity of the microbial communities of the channel sediments and the tundra?

2) Most of the samples we obtained were from summertime (July); what is the difference of the channel samples from the wintertime (April, early Arctic spring)?

3) Before, we obtained only DNA samples, thus, the results in the previous studies only represent the potential microbial communities at the LH channel (Lay et al.,

2012) due to the long term preservation of DNA in the high salt environment. The active microbial components are still unknown at the LH channel. 4) As showed 102

in the previous study, Thaumarchaea occupied a large proportions in the channel archaeal communities (Lay et al., 2012); what is their real abundance in the channel sediments? Are their featured genes involved in ammonia oxidation and inorganic carbon fixation present in the LH channel sediments as well?

To answer the above questions, we sampled the channel sediments and the adjacent tundra soils in summer 2011 and winter 2012. In each season, we collected samples for DNA and RNA analyses. Using these samples, we processed them with 16S rDNA pyrosequencing to obtain the pyrosequencing libraries, which was used for exhibiting the microbial communities in different environments, including human microbiota (Keijser et al., 2008), fresh water samples (Vaz-Moreira et al., 2011), benthic coral reef microbiota (Gaidos et al.,

2011), soils (Hirsch et al., 2010; Youssef et al., 2009), and dust samples (Abed et al., 2012). With the pyrosequencing libraries, we compared the biodiversity indices of microbial communities to assess their basic bacterial and archaeal diversity (Shannon index) and richness (Chao1 index). The bacterial and archaeal affiliated sequences were classified using RDP database and Genbank to identify the proportions based on the microbial taxons, phyla or classes (for

Proteobacteria ), in each library. Using the sequences in each library, we applied them to UniFrac analysis (Hamady et al., 2010; Lozupone and Knight, 2005) to assess the possible factors shaping the different microbial communities in each library. In this instance, we calculated two factors, sampling seasons and locations, along with the difference of extracts from samples (DNA v.s. RNA). With the results of these analyses, we sought to provide a greater insight into the microbial components present in different seasons and locations in the channel and the 103

adjacent tundra soils. To extend our knowledge of the LH channel Thaumarchaea, we cloned and sequenced their featured genes, ammonia oxidizing genes ( amoA ) and 4-hydroxybutyryl-CoA dehydratase genes (hcd ) (Dang et al., 2013; Zhang et al., 2010), and Thaumarchaeal specific 16S rDNA sequences to verify their ability to oxidize ammonia and fix inorganic carbon, respectively. The hcd gene is one of the genes involved in the 3-hydroxypropionate/4-hydroxybutyrate cycle, which is used in some archaea or bacteria to fix inorganic carbon (Berg et al., 2010;

Zarzycki et al., 2009). Detecting the presence of hcd genes may prove that the

Thaumarchaea in the LH area has the ability to be a potential primary producer in this ecosystem. Further, we used the putative amino acid sequences of amo A and hcd genes to compare the phylogenetic affiliations with Thaumarchaeal sequences from other environments. These genes were quantified by qPCR for understanding the potential of Thaumarchaea driving the nitrogen and carbon cycles at the LH channel sediments.

Our objectives for this study are: 1) to characterize the seasonal microbial components at the LH channel and the adjacent tundra based on 16S rDNA and rRNA libraries and evaluate the significant differences between factors of those samples, i.e., sampling seasons, locations, and sample types (DNA v.s. RNA), based on OTUs; 2) to assess the amount of archaeal ammonia oxidizers

(Thaumarchaea) at the LH channel and the adjacent tundra based on the sequences of the featured functional genes of amo A, hcd and Thaumarchaeal specific 16S rDNA sequences using qPCR, and; 3) to compare the sequences of amo A and hcd cloned from LH channel samples with other published sequences for identifying the relationship of the LH Thaumarchaea with the ones present in other 104

environments.

4.2 Materials and methods

4.2.1 Sample collection and geochemical analyses

Designated for RNA analysis, sediment samples (15 g each) of both the main outflow channel (79°04.608N; 90°12.739W) and the adjacent tundra

(79°04.614N, 090°12.666W) at the LH spring were collected in July 2011 and

April 2012. Using an ethanol-sterilized scoopula, these summer and winter samples were placed in a sterile 50 mL Falcon tube filled with LifeGuard™

(Mobio Laboratories, Inc, Carlsbad, CA) solution to a final volume of 50 mL to fix RNA and its related activities. For DNA extraction and geochemical analyses, channel and tundra sediment samples (approx. 250 g) collected approximately 50 mm below the surface with an ethanol-sterilized scoopula, were placed in sterile plastic sampling bottles (except the winter 2012 tundra sample, collected in a

Whirlpak bag). All samples were maintained below 5°C during their transport to

Montreal, and then stored at -20°C until analysis. Due to the logistic difficulty of attending this remote location, we obtained the samples as a snap shot with no true replicate.

To determine in situ CO 2 flux from LH channel and the tundra, an automated static-chamber Li-Cor model Li-8100 (Li-Cor Bioscience, Lincoln,

Nebraska, USA) was used as described by the manufacturer. The collars with a diameter of 24 cm were installed in the channel sediments and tundra soil; the

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system was allowed to equilibrate ~20 min to 1 h. The salinity and the water temperatures were measured using the YSI 556 Multi Probe System (YSI

Incorporated, Yellow Springs, OH, USA).

For the analysis of total nitrogen/total carbon/inorganic carbon content, sediments were oven-dried at 60°C, and then finely ground. Carbonate content was determined using subsamples of oven-dried sediments acidified using 1 M

HCl to remove the carbonates and then dried at 50°C (Hedges and Stern, 1984).

The original and acidified sediments were analyzed for total carbon, total nitrogen, organic carbon by combustion at 900°C with a Carlo Erba Flash EA NC Soils

Analyzer [Carlo Erba, Milan, Italy (Lim and Jackson, 1982)]. All carbon and nitrogen analyses were performed in triplicate. Ammonia, nitrite and nitrate concentrations were measured in the pore water extracted from sediments following centrifugation at 2,000g for 10 min. Tundra samples’ soil-bound ammonia were determined by washing 30 g of sediment with 30 ml milli-Q water, and then extracted with 30 ml of 2 M KCl (Maynard and Kalra, 1993). Ammonia and nitrate/nitrite concentrations were analyzed on a multi-channel Lachat AE

Quik-Chem auto-analyser (Lachat Instruments; Milwaukee, WI, USA).

4.2.2 DNA and RNA extraction, cDNA synthesis, pyrosequencing and analyses.

To obtain total DNA for qPCR analyses to quantify amo A, hcd and

Thaumarchaeal 16S rDNA genes, sediment and soil samples (1 g) from the LH channel and tundra were processed with an UltraClean® Soil DNA Isolation Kit

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(Mobio Laboratories, Inc, Carlsbad, CA, USA) according to the manufacturer’s instructions, with a phenol/chloroform extraction added after the first bead shaking step. Extracted DNA was dissolved in 50 µL of deionized distilled water

(ddH 2O) for each sample and stored at -20°C until further analysis.

To obtain total RNA and parallel DNA, LH channel sediment and the tundra soil samples (2 g) were processed with an RNA PowerSoil® Total RNA Isolation

Kit (Mobio Laboratories, Inc, Carlsbad, CA, USA). The manufacturer’s recommended protocol was slightly modified as follows: (i) an additional 1 g of

0.1 mm glass beads (Mobio Laboratories, Inc, Carlsbad, CA, USA) were added to each reaction tube, (ii) bead-beating time was doubled up to 30 min, and (iii) nucleotide precipitation proceeded overnight. The parallel DNA sample was extracted using an RNA PowerSoil® DNA Elution Accessory Kit (Mobio

Laboratories, Inc, Carlsbad, CA, USA) and then dissolved in 50 µL of ddH 2O for each sample and stored at -20°C. Total RNA was treated with amplification grade

DNase I (Invitrogen, Carlsbad, CA, USA) at room temperature for 15 min based on the manufacturer’s recommendations and then inactivated by the addition of

EDTA incubation at 65°C for 20 mins. The DNase I-treated RNA samples were concentrated and purified using Amicon Ultra-0.5 mL Centrifugal Filters

(Millipore, Tullagreen, Cork, Ireland). To synthesize cDNA, an iScript TM Select cDNA synthesis kit (Bio-Rad, Hercules, CA, USA) was applied to process the purified RNA samples, using random primers provided by the kit.

To determine the 16S ribosomal DNA and cDNA, the samples were sequenced at the Research and Testing Laboratory (Lubbock, Texas, USA) using a

Roche 454 GSFLX Titanium sequencer (454 Life Sciences, Branford, CT, USA) 107

system with bacterial (28F:5’GAGTTTGATCNTGGCTCAG’3,

519R:5’GTNTTACNGNGGNKGCTG’3) (Handl et al., 2011) and archaeal

[ARCH571F: 5’GCYTAAAGSRNCCGTAGC’3 (Baker et al., 2003), ARCH909R:

5’TTTCAGYCTTGCGRCCGTAC3’ (Burggraf et al., 1997)] primers. The tag-encoded pyrosequencing was performed following established protocols

(Bailey et al., 2010). The ribosomal 16S DNA and cDNA sequences were trimmed, aligned, and dereplicated using the RDP pyrosequencing pipeline (Cole et al., 2009). In brief, the sequences were screened for their quality using the following parameters: a minimum quality score of 20, and a minimum sequence length of 150 bp; sequences that did not match these parameters were excluded from the downstream analyses. Sequences were aligned and then clustered using a maximum distance of 15% and step size of 1.0. The dereplicated representative sequences, which had a minimum similarity of 98% to those they represented, were analyzed via the BLASTn algorithm against the online GenBank database

(Altschul et al., 1990).

4.2.4 UniFrac analysis of the LH libraries

To analyze the genetic distance of different LH samples (Beta-diversity), we processed the libraries using UniFrac analysis (Lozupone and Knight, 2005). The libraries were pretreated using NCBI blastall based on commandline.

Weighted-normalized Unifrac distances between each sample pair were calculated using the FastUnifrac website (http://bmf2.colorado.edu/fastunifrac/ ) (Hamady et

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al., 2010) based on the GreenGene core dataset.

4.2.5 Archaeal amo A and hcd gene cloning and sequencing and analyses

To clone the partial archaeal ammonia oxidizing gene subunit A ( amo A), 2

µL of environmental DNA extracted from the LH channel were applied to

Polymerase chain reaction (PCR) using Immolase TM DNA Polymerase (Bioline,

London, UK). The specific primer pair, CrenamoA23f

(5’ATGGTCTGGCTWAGACG’3) and CrenamoA616r

(5’GCCATCCATCTGTATGTCCA’3) (Tourna et al., 2008), was applied to amplify the partial amo A gene of ~600 bp, under thermocycler conditions of:

95°C 10 min, followed by 35 cycles of 94°C 30 sec, 55°C 30 sec and 72°C 1 min; with a final extension of 5 min at 72°C. With the same amount of LH channel

DNA, the partial 4-hydroxybutyryl-CoA dehydratase (hcd ) gene was amplified in a nested PCR reaction using Immolase TM DNA Polymerase, with a first primer pair, hcd-120F-SCM1 (5’AGCCTGTAGACCACCCAATG’3) and hcd-1367R-SCM1 (5’ TATTCTTTGGGCCTGTGGAG’3), under thermocycler conditions of: 94°C 10 min, followed by 30 cycles of 94°C 30 sec, 58°C 30 sec and 72°C 1 min; with a final extension is 5 min at 72°C. An aliquot (1 µL) of the first PCR product was then applied to the second PCR reaction using the primer pair, hcd-911F (5’AGCTATGTBTGCAARACAGG’3) and hcd-1267R (5’

CTCATTCTGTTTTCHACATC’3), under thermocycler conditions of: 94°C 10 min, followed by 30 cycles of 94°C 30 sec, 58°C 30 sec and 72°C 30 sec; with a

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final extension of 5 min at 72°C (Zhang et al., 2010). The final PCR products of both partial amo A and hcd genes were purified using QIAquick Gel Extraction

Kit (Qiagen, MD, USA) followed by ligation to pGEM-T easy vector at 4°C overnight (Promega, Madison, WI, USA). The vectors with the partial gene insertions were transformed into competent cells DH5 α (Invitrogen, Carlsbad, CA,

USA) using heat shock for 90 seconds at 42°C. The competent cells were then cultured at 37°C on ampicillin (final concentration 100 µg/mL) containing

Luria-Bertani agar plates with 20 µL X-Gal (50 mg/mL) on each plate. The selected white colonies were verified by colony PCR using SP6-T7 primer pairs.

Positive PCR products were sequenced using a 16-capillary genetic analyzer (ABI

Prism 3130XL) at the Université Laval Sequencing Facility (Plateforme d’Analyses Biomoléculaires, Québec, QC, Canada). DNA sequences of the functional genes were translated into 3 amino acid sequences using Bio-edit (Hall,

1999). For the partial gene sequences, these without stop codons were considered as the putative sequences of the partial proteins. Maximum-likelihood phylogenetic trees of selected sequences were generated with MEGA 5 (Tamura et al., 2011), using a bootstrap method with 1,000 replications and a Jukes-Cantor model.

4.2.6 qPCR of Thaumarchaeal 16S/ amo A/ hcd genes in LH channel sediments and tundra

To quantify the copy numbers of Thaumarchaeal 16S/amoA/hcd genes in the LH channel sediments and tundra, qPCRs were performed in triplicate on an

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iCycler IQ5 System (Bio-Rad, Hercules, CA, USA) using iQ TM SYBR® Green supermix (Bio-Rad, Hercules, CA, USA) as reagent. For each reaction, 2 µL of

DNA was applied to the final volume of 25 µL reaction mixture, resulting in final primer concentrations of 1 µM. Primer pairs detecting 16S, amo A and hcd genes were 771F (5’ ACGGTGAGGGATGAAAGCT ‘3)/957R(5’

CGGCGTTGACTCCAATTG ’3) (Ochsenreiter et al., 2003), CrenamoA23f/

CrenamoA616r, and hcd-911F/ hcd-1267R, respectively. For all three genes, the qPCR program began with 95°C for 3 min and 30 cycles at 94°C for 30 s and continued with 50°C for 30 s, and 72°C for 30 s for the 16S gene, 3 min and 30 cycles at 94°C for 30 s, 55°C for 30 s, and 72°C for 1 m for the amo A gene, and

58°C for 30 s, and 72°C for 1 m for the hcd gene. After these cycles, for each gene set point temperatures were increased by 0.5°C increments from 55°C to

95°C for collecting melt curve data. The negative controls were ddH 2O containing no detectable DNA. We employed previously cloned linear fragments of amo A, hcd and Thaumarchaeal 16S rDNA genes to generate standard curves.

They were diluted to 5 concentrations for generating the standard curve. The PCR efficiency of 16S was 96.9% and 121.2% for two separate reactions, and of amo A and hcd genes were 84.9% and 90.2%, respectively. The R 2 values of standard curves were 98.4% and 92.8%, for the two reactions of the 16S gene, and 98.1% and 99.8% for the amoA and hcd genes respectively.

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4.3 Results

4.3.1 Geochemical analyses of the LH channel and tundra sampling sites

The background nutrient supply and environmental properties of the sampling sites were documented (Table 4-1). Winter season outflow channel water salinity was 25%, similar to that found in the outlet. However, due to unknown reasons (It might be higher levels of precipitations before sampling.

There is no long term weather monitoring station at the LH spring area or any other places on the Axel Heiberg Island) prior to obtaining the summer samples, channel sediment salinity was low. Based on past records and samples in following years (e.g., winter sample used in this study), this low salinity may be temporary. Water temperatures varied from 4.1°C in the summer down to -12.9°C in the winter, which was consistent with records of air temperatures. With the exception of higher ammonia concentrations in winter 2012, nitrogen and carbon contents in spring sediments were lower than in the adjacent tundra. Ammonia concentrations in the winter channel and tundra were both higher than in the summer. Less microbial activity in the winter utilizing ammonia may result in greater ammonia accumulation. The CO 2 flux was similar in the channel sediments in both seasons. On the contrary, the CO 2 flux was much less in the tundra than in the channel. The CO 2 flux may result from microbial activities. The records of the flux implied that the channel was a relatively suitable environment for microorganisms to inhabit.

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4.3.2 Pyrosequencing library statistics

The pyrosequencing results showed reasonable original reads numbers (>

1K) in most DNA and cDNA bacterial libraries from summer and winter samples

(Table 4-2). However, one summer and four winter archaeal libraries, i.e. TASC,

CAWC, CAWD, TAWC and TAWD (Table 4-2; each letter of sample’s names stands for sampling site, primer set for bacteria or archaea, sampling season, and extract’s type in order), showed a low output of read numbers (< 1K). Among these libraries, the CAWD and TAWC libraries were particularly low (4 and 29 reads, respectively; Table 4-2); thus, we did not calculate their indices of richness and diversity. The sequenced reads of TASC, CAWC, and TAWD libraries were fewer (271, 707 and 194 reads, respectively); however, the original quality of cDNA sent for sequencing was sufficient to generate reliable sequences: the

TASC, CAWC and TAWD cDNA concentrations were 16.2 ng/ µL (ratio of 16/280 of 1.36), 19.3 ng/ µL (ratio of 260/280 of 1.89), and 8.1 ng/ µL (ratio of 260/280 of

1.66), respectively.

The Chao1 index indicates the richness (the potential numbers of the phylotypes at the sampling site) of samples. In the LH libraries, for each

DNA/cDNA pairing sample, the richness of the DNA libraries were generally greater than that of the cDNA libraries, except in the case of the tundra winter

Bacteria libraries, TBWD and TBWC. The Shannon index, representing biodiversity, showed higher numbers in all bacterial libraries compared to their parallel archaeal libraries (1 to 2 folds). As with the Chao1 index, the Shannon indices of the channel libraries were lower for the winter than summer libraries.

However, the tundra libraries, particularly the bacterial ones, did not change by 113

season. Moreover, for the two Bacterial library pairs, the summer channel libraries and the winter tundra libraries, the Shannon indices revealed lower diversities in the DNA libraries than in the cDNA libraries.

A Unifrac analysis of LH samples, visualized through PCoA analysis, compared all libraries to create a more comprehensive picture of the

Beta-diversity. Most summer samples were grouped into adjacent areas, while winter samples showed clustering with different extract’s types (DNA v.s. RNA;

Figure 4-1). However, sampling site differences were not significant. Permanova analyses based on the same matrix drew the same conclusions (Table 4-3). For archaeal libraries, the only significant difference was based on seasons ( P ≤ 0.05), whilst for two other factors (molecular type and sampling site), no effect was seen

(P > 0.05).

The beta-diversity of the Bacterial libraries showed that the summer and winter samples were mostly grouped together according to their sampling seasons of the PCoA figure (Figure 4-1a). Although the two libraries, CBWC and TBSD, were away from the winter samples and summer samples, respectively, the

Permanova analysis still showed that the significant factor for these samples was the season ( P ≤ 0.05; Table 4-3). The bacterial libraries did not show significance based on extract’s type or sampling site (P > 0.05). A similar trend appeared on the Archaeal summer libraries. In the Archaeal figure (Figure 4-1b), summer samples grouped together but the winter samples were relatively separate.

However, based on the Permanova analysis for identifying significant factors, season was still the primary factor in shaping the differences in each Archaeal library (P ≤ 0.05; Table 4-3). Neither of the other two factors (sampling site and 114

extract’s type) had significant effect on the difference of the samples.

According to the Unifrac distance, the bacterial samples were similar to each other based on the distances were generally less than 0.2. The distances of similarity in between some samples, i.e., CBWC v.s. CBSD, CDSC, and TBSD, were higher than others (Table 4-4 a), which made the dissimilarity mainly focusing on the sampling seasons. In general, the distances of similarity were lower for the samples from the same seasons than the locations (Table 4-4 a). The

Unifrac distances of archaeal samples were higher than these of bacteria (Table

4-4 b). The archaeal samples from different seasons showed high distances among them (more than 0.3) (Table 4-4 b). The archaeal samples from summer showed relatively low distance among themselves than the samples from the winters. It was probably due to the less efficient sequencing results for the winter samples.

4.3.3 Microbial compositions in LH spring channel and tundra in the summer

In general, the pyrosequencing reads for the summer samples were sufficient for further analyses, because most of the analyses returned between 3K and 6K of raw reads. Only the TASC has relatively low numbers of raw reads

(271) being generated. Comparing the DNA/cDNA pyrosequencing pairs to each other, cDNA libraries usually had fewer reads sequenced than DNA libraries. The only exceptions were the summer channel bacterial libraries, where the numbers of CBSC reads slightly exceeded those of CBSD (Figure 4-2), In the 16S rDNA libraries, bacterial diversity in both the summer channel and tundra was higher

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than archaeal diversity (Table 4-2, See the Shannon indices). Five active bacterial phyla/classes ( Alphaproteobacteria , Verrucomicrobia , Deltaproteobacteria ,

Betaproteobacteria , and Firmicutes ) had higher proportions of reads than their

DNA libraries in the tundra, while only two phyla/classes ( Alphaprteobacteria and Deltaproteobacteria ) appeared in the channel. However, the total proportions of the reads from these active phyla/classes in the channel (57.54%) were almost equal to those active phyla/classes in tundra (58.58%), implying that in the tundra, the active phyla were more selective in the channel than in the tundra. According to the DNA libraries, Bacteroidetes and Alphaproteobacteria were among the most abundant bacterial phyla in the channel and tundra. The proportions of

Bacteroidetes in DNA (30 to 40%) and cDNA (20 to 30%) libraries were similar in the tundra and in the channel. Gammaproteobacteria and Actinobacteria were also abundant in both habitats, but their DNA/cDNA proportions indicated they were not as active. Cyanobacteria was not abundant in the channel libraries (less than 5%), but it increased in the tundra library, especially in the DNA library (10 to 15%). The proportions of Verrucomicrobia in the tundra were greater than in the channel (5 to 10% in the tundra, less than 5% in the channel). As we addressed, based on the cDNA library, the Verrucomicrobia were highly active in the tundra.

The archaeal libraries in both channel and tundra did not contain many phyla/classes (Figure 4-2b and 4-2d). Halobacteria and Thaumarchaeota were present at both sites; however, the Halobacteria seemed to be suppressed in the tundra, whereas they were active in the channel, suggesting they were more suitable inhabiting the channel than the tundra due to the hypersalinity. Regarding

Thaumarchaeota , at both sites, the cDNA was proportionately less than the DNA. 116

This may indicate that the LH environment, either in the channel or the tundra, was not quite suitable for Thaumarchaea to live and metabolize. However, this environment may preserve their DNA for a long time after they appear. Only reads related to methanogens were detected in the DNA library of the summer channel. In the tundra, there were a high proportion of unclassified archaea both from the cDNA and the DNA libraries, indicating some other unknown species may be present at this sampling site.

4.3.4 Microbial compositions in LH spring channel and the tundra in the winter

The winter libraries were less reliable than the summer ones (Figure 4-3) based on the total numbers of reads (Table 4-2). Not all the winter libraries were well-sequenced; for example, the archaeal DNA library of the channel (CAWD) and the archaeal cDNA library of the tundra (TAWC) contained too few reads (4 and 29, respectively). Therefore, these two libraries may not reflect the true archaeal compositions of these sites and will not appear in most parts of this study.

Most bacterial phyla present in the summer channel DNA library were also present in the winter channel DNA library. The channel bacterial DNA and cDNA libraries differed clearly, especially for the four most active phyla, i.e.

Cyanobacteria , Firmicutes , Gammaproteobacteria and Verrucomicrobia .

Comparing to the summer channel libraries, these seemed to be more active in the winter than in the summer. It implies that only few phyla/classes of bacteria would be better adapted to the winter channel environment. The winter tundra

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bacterial libraries showed lesser differences between DNA and cDNA. Although

Cyanobacteria , Actinobacteria , Verrucomicrobia , Deltaproteobacteria , and

Planctomycetes showed higher read proportions in the cDNA than in the DNA libraries, only reads of Verrucomicrobia and Deltaproteobacteria were more than

50% higher in the cDNA than in the DNA libraries. Bacteria of these two phyla were active in the tundra soil in both the winter and summer.

In the winter archaeal libraries (Figure 4-3b and 4-3d), CAWD and TAWC had too few reads (4 and 29, respectively). Since in the summer channel DNA library, we detected more diverse reads than just related to Halobacteria , we do not think the channel only contained Haolobacteria ’s DNA during winter in the channel. Similar reasons for the tundra archaeal cDNA library, TAWC, where the representative reads only related to Thaumarchaea and other unclassified archaea.

However, due to the few numbers of few reads, we found it unlikely that it reflected the true active archaeal communities. These two libraries, CAWD and

TAWC, may exhibit only bias. In the winter archaeal libraries, the CAWC and

TAWD had relatively higher raw reads (707 and 194, repectively) comparing to

CAWD and TAWC. CAWC reflected the active archaeal community in the channel during the winter time. It contained reads related to Thaumarchaeota ,

Methanobacteria , and unclassified archaea. It implies that ammonia oxidizing and methane metabolizing may happen at the winter channel sediment. In the winter tundra DNA library (TAWD), only reads related to halobacteria were detected.

Considering the summer tundra archaeal DNA library (TASD) containing more diverse phylotypes than TAWD, there are two possible reasons for this results: 1) the tundra soil was not a good environment to reserve DNA for a long time; the 118

DNA decomposed while the microbial activities decreased during the winter; 2) this is also a sequencing or extraction bias.

4.3.5 Archaeal functional genes for ammonia oxidation and carbon fixation

Three partial sequences of the ammonia oxygenase alpha subunit ( amo A) gene and four partial sequences of the 4-hydroxybutyryl-CoA dehydratase (hcd ) gene were cloned and translated into amino acid sequences. The phylogenetic tree based on amino acid sequences of amo A revealed that the archaeal amo A grouped differently with bacterial amo A genes (Figure 4-4a). Additionally, the LH amo A clustered with other Thaumarchaeota sequences into two groups: one was closest to the genus, Nitrosoarchaeum , and the other was closest to the genus,

Nitrososphaera .

The phylogenetic tree based of hcd amino acid sequences showed three groups: Crenarchaeota , Thaumarchaeota and Bacteria (Figure 4-4b). The cloned sequences from LH fit into the Thaumarchaeota, and were closely related to each other. The clustering of the hcd gene did not show a pattern similar to the amo A gene. The closest sequences to the LH hcd were from a known symbiotic uncultured Thaumarchaea , Cenarchaeum symbiosum and an uncultured soil archaeon (Genbank accession number: ADO79756.1).

The qPCR detected Thaumarchaeal amo A and hcd genes in both the channel and the tundra in summer and winter (Figure 4-5). To compare the abundance of copy numbers, we also quantified Thaumarchaeal 16S genes as references. The copy numbers of amo A genes in the summer samples were

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slightly higher than in the winter samples, but all within the same level of magnitude (10 5). The same trend in copy numbers was observed for the hcd genes, albeit one magnitude (10 6) higher than the amo A gene. Except in summer tundra samples, hcd copy numbers exceeded those of Thaumarchaeal 16S genes.

4.4 Discussion

4.4.1 Seasonal changes in active microbial components in LH channel area

LH spring system is an ecosystem with many extreme properties, including hypersalinity, cold/subzero temperatures, and rich in sulfate, methane, and ammonia. The microbial communities at LH system relate to its properties in many different facets. The previous studies mainly focused on the microbial communities based on their DNA (Lay et al., 2012; Niederberger et al., 2010), which revealed the potential microorganisms inhabiting LH system. In the study of Lay et al. in 2012, the mineralization assays in microcosms were performed to

14 14 detect C-CO 2 recovery from C-acetate (Lay et al., 2012), It was the first to detect microbial activity in the LH channel sediments, but this finding only revealed the bulk heterotrophic microbial activity, at temperatures as low as -20°C

(Lay et al., 2012). The active microorganisms were still unknown. The Lay et al. implemented the metagenomic study of the LH outlet for knowing the functional genetic potential, and well as the active microbial components (Lay et al., 2013).

In this study, more than 70% of the active bacterial component at the LH site was found to be Gammaproteobacteria , Verrucomicrobia , Actinobacteria and

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Betaproteobacteria , while Thaumarchaeota and Crenarchaeota comprised over

80% of the active archaeal component. These results are the references for the active microbial components at the downstream channel based on 16S rDNA pyrosequencing libraries. In this study, 16S rDNA pyrosequencing libraries served to assess microbial diversity not only from the DNA but also from the cDNA, indicating active microbial components of the bulk sediment’s biomass.

Here, the cDNA libraries further showed the active components partially comprised of autotrophic microorganisms, e.g., Cyanobacteria and Chloroflexi

(Figure 4-2 and 4-3), including genera of Nostoc , Synecchococcus , and Caldilinea , which may be the basic primary producers in the system. The pyrosequencing libraries showed similar trends with several main phyla and classes, i.e.,

Bacteroidetes , Alphaproteobacteria , Gammaproteobacteria and Actinobacteria , but also indicated a greater abundance of other minor phyla than in the clone libraries (e.g., Verrucomicrobia , Cyanobacteria , Betaproteobacteria and

Deltaproteobacteria ) from the Lay et al. 2012 study. The prior study did not determine if the Thaumarchaea were active or if only their DNA fragments were present in the sediments. We confirmed that Thaumarchaea were active both in the channel sediments and in the adjacent tundra. However, a second large component, the methanogenic archaea, which was detected in the previous study (Lay et al.,

2012), did not appear much in either 16S cDNA or DNA pyrosequencing libraries.

Reads related to methanogens were only detected in the summer channel DNA and winter channel cDNA libraries. Although it showed relatively high proportions (up to 41.2%) in the winter channel cDNA library (CAWD), as we mentioned in the results, the CAWD library only contained 4 raw reads. The result 121

of CAWD was not representative at all. Comparing this and the last study on the channel sediments (Lay et al., 2012), there was probably a microbial community shift in terms of methanogenic archaea happened in this ecosystem; however, we did not have more replicate samples to confirm this observation.

Based on the P-value from the UniFrac analysis, the key factor, besides molecular type and sampling site, affecting the difference between libraries was the season (Table 4-3). The same trend was supported by the Unifrac distance

(Table 4-4). The samples showed relatively less distances in terms of seasons than other factors (Table 4-4). We do not understand why the sampling sites did not make a significant difference for the microbial compositions. The most important difference of the channel sediment and the tundra is salinity. The tundra soil is possibly less saline than the channel sediment. The salinity is also the most important environmental property to shape the microbial communities (Lozupone and Knight, 2007). However, this factor was not reflected in our pyrosequencing libraries. The reasons for this result may be due to low salinity while we obtained the channel sample in the summer. We suggest that this low saline condition in the channel was only temporary, but we do not know if the lasting low saline condition was sufficiently long to change the microbial community within the channel sediment. It needs confirmation by more timely samples and future studies.

Based on the seasons, an important difference in the summer and winter bacterial components was that the proportions of Verrucomicobia , Actinobacteria , and Cyanobacteria in the cDNA libraries increased significantly in the winter samples compared to summer samples, both in the channel and tundra. 122

Verrucomicrobia were also reported as the dominating microorganisms during the winter season in an alpine lake covered by snowpack (Lake Redon, Pyrenees)

(Llorens-Marès et al., 2012); in this study, the abundance of these dominant microorganisms decreased as the winter season passed. The study of a sub-Antarctic marine area (Kerguelen Islands and Antarctic Peninsula areas) also showed some trends of Verrucomicrobia as slightly more abundant in the winter than in the summer, though the mean water temperatures in that area did not vary widely (2.9 in the winter and 5.2 °C in the summer, respectively) (Ghiglione and

Murray, 2012). The dominance of active Actinobacteria in a winter Arctic tundra soil, detected using bromodeoxyuridine labeling, was reported by McMahon et al. in 2011 (McMahon et al., 2011). Similarly, here, cDNA indicated Actinobacteria as being one of the dominant active phyla during the high Arctic winter. We observed relatively high cyanobacterial proportions in the active bacterial communities. Although the Arctic winter we sampled the sediment and soil was still cold, the eternal day light had started there. It may be the reason caused the high proportional cyanobacterial reads in the winter libraries. The temperatures when we sampled was subzero (-12.9 to -16.9°C), and this results may hint that the photosynthesis occurred at subzero temperatures at LH channel sediment and the tundra even though we did not have direct evidence regarding photosynthesis.

Subzero photosynthesis was observed and reported on Cyanobacterial species, which was able to reactivate photosynthesis down to -18°C (Sand-Jensen and

Jespersen, 2012). Thus, the Cyanobacteria probably had photosynthesis activity during the winter. Also, this was not the first time that high proportions of

Cyanobacteria have been reported in the Arctic region (Chukchi Sea) in winter. 123

The cyanobacterial genus, Synechococcus , exhibited an elevated activity during the winter in Chukchi Sea (Cottrell and Kirchman, 2009). This might be attributable to the Cyanobacteria ’s consumption of dissolved organic matters, e.g., amino acids, cyanate and organic sulfur compounds, as an alternative pathway of maintaining its metabolic activities (Malmstrom et al., 2005; Palenik et al., 2003).

The high proportions of these microorganisms in winter might be caused also by the decrease of other microbial species that share similar ecological niches at the

LH system.

4.4.2 Microbial biodiversity and richness in LH channel sediments

The channel sediment biodiversity and richness indices for both bacterial and archaeal libraries present at the LH site were far higher than those measured in the previous study (Lay et al., 2012), particularly in terms of richness. The richness (Chao1 ~ 1000 to 3000) of some of the libraries, e.g., CBSC, CBSD,

CASD, RBSD, TASD, and TBWC (Table 4-2), were as high as or even higher than the Chao1 indices of soils in Oklahoma (~1000 to 2000) (Youssef et al.,

2009). Using the same methods (pyrosequencing), the benthic coral microbiota

(Gaidos et al., 2011) exhibited even higher Chao1 (1000 to 8000) and Shannon (5 to 8) indices in the libraries while the LH libraries had the numbers of Shannon index ranging from 3 to 6.5 (Table 4-2). However, a study on the microbial communities of the coast of Andaman Sea showed relatively low richness (Chao =

372) and diversity (Shannon = 3.04) (Sundarakrishnan et al., 2012). These numbers revealed that the microbial communities among LH system were as

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diverse and rich as a temperate soil ecosystem but less rich and diverse than the coral reefs. Compared to the previous study on the channel sediment (Chapter 2), the increases in the numbers of biodiversity and richness of the LH libraries may indicate that the microbial communities in the LH channel have expanded during the past few years; however, are they as true as they represent? The huge differences of numbers in the previous and current studies made this statement controversial. There are three reasons which might cause the differences in the estimation of biodiversity: the capability of primers, the sequencing region, and changes in methodology.

Each 16S rDNA primer pair for implementing PCR or amplicon pyrosequencing reactions have diverse capacities to amplify bacterial rDNA from each taxonomic group. The capacities can be checked using the Probe Match tool provided by RDP (http://rdp.cme.msu.edu/probematch/search.jsp ) (Table 4-5).

The forward primers for cloning bacterial 16S ribosomal RNA sequences in the current study and the one used in the 2012 study were almost the same (28F and

27F, respectively). However, it still showed significantly different coverage (46.1 and 75.7%, respectively) for matching the bacterial sequences in the RDP database. Conversely, the reverse primers were quite dissimilar (758R and 519R for the 2012 and this study, respectively). They targeted different regions of 16S rDNA and generated different lengths of PCR products. Although, in theory, they could amplify 16S ribosomal RNA sequences of all bacterial phyla, these two reverse primers matched 77.1% (758R) and 90.8% (519R) of 16S ribosomal RNA sequences in the RDP bacterial database, based on Probe match tool estimates

(Table 4-5). The primer 758R could not even cover all the bacterial phyla 125

(sequences related to Dictyoglomi and Thermodesulfobacteria were missing.).

Since both forward and reverse primers of the present study had higher coverage for the bacterial database, it is reasonable to amplify more diverse reads in this study than in the 2012 study. For the archaeal libraries, the forward primers showed similar percentages for cloning archaeal reads in the database for both studies (109F for 49.3% in the 2012 study and 571F for 51.5% in the present study). The reverse primer used in the 2012 study showed a much higher coverage

(934R, 88.0%) while the reverse primers used in this study showed lower coverage (909R, 61.0%). Also, the prediction for the four primers showed that none of the sequences related to Thaumarchaeota can be amplified by using them.

However, in both of our studies, we used these two archaeal primer pairs to amplify sequences affiliated with Thaumarchaeota . The archaeal database of RDP is not as good as the bacterial database. This disadvantage was also mentioned by another research group, i.e. Gaidos et al. indicated that RDP database was inadequate for archaea classification in their study of benthic coral reef microbiota

(Gaidos et al., 2011). The results of the probe match tool may not be appropriate for elucidating the phylogenetic capacity of archaeal primers.

A second reason might be related to the amplified regions. Here, these were

V1-V3 for bacteria and V4-V5 region for archaea, whereas in 2012 study, they were V1-V4 and V1-V5, respectively (Table 4-5). Amplifying variable regions of ribosomal RNA may result in different richness of OTUs (Youssef et al., 2009).

The study of Youssef et al. revealed that using the V1+V2 regions may overestimate the microbial richness because of their relatively high variability; furthermore, the V3 region is hypervariable. In the contrary, the richness based on 126

V4 and V5+V6 regions are relatively comparable to the richness estimated by using the nearly full length 16S rDNA sequences. The bacterial sequences used in this study only contained V1 - V3 regions, which are considered as high variable to hypervariable regions. In the 2012 study, the bacterial sequences contained

V1 – V4 regions, which included the less variable region, V4. It may reduce the estimation of richness. Thus, for the bacterial 16S libraries, the difference of diversity and richness may be caused by this reason. However, the same assumption is not practical for the archaeal libraries. In this study, the archaeal

16S rDNA sequences contained V4 –V5 regions, which were supposed to be less variable than the archaeal 16S rDNA regions, V1- V5, used in the 2012 study.

Comparing the archaeal richness and diversity of these two studies, this study got returned higher numbers than the 2012 study. Thus, there may be another reason to explain these results.

Thirdly, in the present study, we used pyrosequencing to generate high numbers of reads (usually, more than 1,000 reads for each library). In the 2012 study, a clone library gave a maximum ~200 clones per library. The size of DNA pools was very different and affected the statistical results. The output lengths of

DNA were shorter from pyrosequencing (mean length approx. 300 bp) than the clone library (700 bp and 800 bp for bacteria and archaea, repectively). These two factors, which may be caused by the efficiency of sequencing/cloning techniques, and then strongly affect the results. Additionally, two studies indicate that using different culture-independent methods for analyzing microbiota in the same samples may yield different results (Dowd et al., 2008) or higher richness and diversity (Vaz-Moreira et al., 2011). Proportions obtained from amplicon 127

pyrosequencing may result in bias (Hirsch et al., 2010; Tedersoo et al., 2010).

This remains under debate and more research on comparisons of methods is needed. Thus, pyrosequencing may reflect true or relative trends of the microbial diversity and richness at the LH channel sediment site.

4.4.3 Thaumarchaeal signature functional genes in the LH channel sediment and the adjacent tundra

To extend our knowledge of the LH Thaumarchaeota from our 2012 study on Thaumarchaea-related phylotypes detected in the LH channel sediments (Lay et al., 2012), a further survey was executed on the sediments’ ammonia oxidizing archaea (AOA), which usually constitute a large proportion of the archaeal community across diverse environments. Thaumarchaeota , formerly a crenarchaeal marine group, is a newly-classified phylum, which, as far as we know, includes mostly ammonia-oxidizing archaea (AOA). The ammonia detected at both the LH outlet and in the channel is consistent to supply as nutrients for

Thaumarchaea (Lay et al., 2012). Members of this phylum have been detected all over the ocean and soil environments, including agricultural soils, grass lands, fresh water, forest soils, sea water, hot springs, and cold springs (Dang et al., 2013;

Francis et al., 2005; Hatzenpichler, 2012; Hatzenpichler et al., 2008b; Lay et al.,

2012; Lay et al., 2013; Ochsenreiter et al., 2003; Tourna et al., 2008; Yao et al.,

2011; Zhang et al., 2010), and are among the most abundant microorganisms in the ocean (DeLong, 2003). However, only few species have been isolated or enriched in labs (Zhalnina et al., 2012). A large proportion of the AOA remains

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unknown and uncultured. Thus far, all cultured Thaumarchaeal representatives are aerobes and have the potential to oxidize ammonia (Spang et al., 2010).

Compared to the ability of these bacteria (AOB) to oxidize ammonia, AOA can adapt to the environments with wider range of ammonia concentration than AOB

(Verhamme et al., 2011), including the conditions with relatively low ammonia concentrations (Schleper, 2010). Besides ammonia oxidation, Thaumarchaeota are also able to utilize inorganic carbon, such as carbonate, placing them in a group of carbon fixers as primary producers in their environments. Recent studies suggest that, when the environment is lacking in ammonia, Thaumarchaeota may decompose urea, if it is present, to generate both ammonia and carbonate for further use (Alonso-Saez et al., 2012). The whole process of Thaumarchaea using and consuming carbon and nitrogen may, therefore, be more complicated than previously thought.

In previous studies of the LH system, Thaumarchaeal DNA and RNA were in both LH channel and outlet sediments, respectively (Lay et al., 2012; Lay et al.,

2013). However, we did not detect DNA in any of AOA’s signature functional genes, e.g., the ammonia monooxidizing gene ( amoA ) or the

4-hydroxybutyryl-CoA dehydratase gene ( hcd ) in the LH outlet metagenome.

These genes are considered as signatures for the presence of Thaumarchaeota in diverse environments (Dang et al., 2013; Zhang et al., 2010). In this study, we cloned and sequenced Thaumarchaeal amo A and hcd genes to understand their phylogenetic affiliations to different Thaumarchaeal groups, and then quantified their abundance using qPCR to compare the abundance of these genes in other environments. 129

We found that the copy numbers of amo A genes we detected at the LH channel and the adjacent tundra (~ 10 5) were 1 numerical magnitude less than in agricultural soils, seashores, and the ocean (Dang et al., 2013; Trias et al., 2012;

Yao et al., 2011). This might relate to the low amount of the total nitrogen in this environment (0.01-0.1% in the channel and 0.1-0.3% in the tundra; Table 4-1).

The total nitrogen was relatively low in the LH channel sediment, whereas at most agricultural soils it ranges from 0.1 to 0.5% (Yao et al., 2011), which is similar to the levels found in the tundra adjacent to the LH system. Thus, the overall impact of the lower copy numbers of the amo A genes in the LH channel and the tundra might be due to other environmental impacts, not just due to the nitrogen supply.

The hcd gene is one of the genes involved in the

3-hydroxypropionate/4-hydroxybutyrate cycle, which is used in some archaea or bacteria to fix inorganic carbon (Berg et al., 2010; Zarzycki et al., 2009). As being a potential primary producer, the Thaumarchaea-driven inorganic carbon fixation does not require light. In fact, some reports indicate that Thaumarchaea is able to fix carbon in dark (Bergauer et al., 2013; Yakimov et al., 2011; Zhang et al., 2010).

It makes Thaumarchaea more suitable for fixing inorganic carbon to supply the ecosystem than the Cyanobacteria since the Arctic has no sun light for several months during the winter. In a study on assessing the inorganic carbon fixation in the deep ocean driven by the 3-hydroxypropionate/4-hydroxybutyrate cycle indicates that the rate was 50 –60 mgCm -3 per day (Yakimov et al., 2011).

Considering the harsher conditions at LH system, we expected the rate of carbon fixation using the same cycle driven by Thaumarchaea would be lower than this number. However, we did not apply any experiment to prove it. 130

In the study, we observed that the copy numbers of hcd genes from both sites exceeded those of Thaumarchaeota 16S ribosomal RNA genes, except in the summer tundra. The ratio of the copy numbers of hcd/16S rDNA genes in the summer tundra was dramatically different than the other three samples (0.62 of the summer tundra compared to 5.03, 6.68 and 11.73 from summer channel, winter channel and winter tundra, respectively). It may indicates that during the summer, the potential of Thaumarchael inorganic carbon fixation in the tundra was not as high as in other sampling times and locations. Their copy numbers were about 1 numerical magnitude higher (~10 6) than the copy numbers of the

Thaumarchaeal 16S ribosomal DNA genes. In general, the copy numbers of 16S ribosomal RNA genes should exceed those of functional genes. However, Dang et al. reported similar observation regarding the ratio of Thaumarchaeal specific 16S ribosomal RNA genes and hcd genes, i.e., in some of their sea water samples

(Dang et al., 2013), where the copy numbers of hcd genes were slightly higher than those of 16S ribosomal RNA genes. This may be caused by non-specific amplifications of the primers. However, the reason behind this phenomenon remains unknown. Overall, the high copy numbers of Thaumarchaeal hcd genes indicates the potential of inorganic carbon fixation and the potential ability of primary producers in the LH spring system and in the adjacent area.

In attempt to know the phylogenetic affiliation based on the amo A and hcd genes, we cloned and sequenced several genes of them from our environmental samples using Sanger’s sequencer. In addition, we also tried to apply our samples to a new method, amplicon pyrosequencing, to amplify the archaeal amo A from the samples. However, the results showed non-specific amplifications and did not 131

return valid sequences (data not shown). Thus, we could only show the results from the Sanger’s seuquencing in this study. In a previous study, an analysis of the

Thaumarchaeal 16S ribosomal RNA sequences in the LH channel were used to develop phylogenetic trees and revealed two closed clusters of Thaumarchaeal phylotypes (Lay et al., 2012). One related to the hot spring Thaumarchaea,

Nitrososphaera gargensis , and the other to an uncultured group, separate from the branches of Nitrosopumilus maritimus and Nitrosocaldus yellowstonii (Lay et al.,

2012). In the neighbor-joining phylogenetic tree based on the partial putative amino acid sequences in this study (Figure 4-4), our amo A representative sequences were also grouped into two branches, one with Nitrososphaera gargensis , and the other one was with the newly characterized Nitrosoarchaeum koreensis , also distinct from the branches of the amo A sequences of

Nitrosopumilus maritimus and Nitrosocaldus yellowstonii . This suggests that two similar clades of Thaumarchaea exist at the LH site. However, the neighbor-joining phylogenetic tree of amino acids of hcd genes only showed one clustered group (Figure 4-4b), which was dissimilar to any cultured or enriched

Thaumarchaea. The sequences of hcd genes might lack the resolution to distinguish different phylotypes to the same degree as 16S ribosomal RNA or amo A genes. Although the tree of hcd gene sequences was not identical to the trees of 16S rRNA or amo A genes, the Thaumarchaeal hcd genes were distinctly different than the crenarchaeal hcd branch, which supported the divergence of the two phyla, Crenarchaeota and Thaumarchaeota . Sampling from cold/subzero and hypersaline environments, however, we could not analyze the cold or saline adaptations of these Thaumarchaeal amo A and hcd enzymes based on their partial 132

putative amino acid sequences. Analyses of adaptations according to amino acid sequences require full length sequences of amino acids. Molecular level adaptations to cold and salinity must be expected in Thaumarchaeal functional genes at LH.

4.5 Conclusion

In this study, we tried to analyze DNA and RNA samples from the LH channel sediment and the adjacent tundra soil, both of which were collected in the summer and the Arctic winter (early spring). By analyzing the 16S rRNA libraries, we could determine the active microbial components and compare them with the parallel 16S rDNA libraries. Also, we undertook a further survey of the LH channel Thaumarchaea based on the two primary genes, amo A and hcd genes. Our study demonstrated seasonal microbial communities based on their active components (RNA) and potential components (DNA) in the LH channel and the adjacent tundra, exhibiting the greater diversity and richness of the microbial communities in different libraries than the previous study on the LH channel.

Overall, our findings indicated that the changes in the abundances of individual microbial clades in different libraries may reflect the certain microbiota responding to the environmental properties. For examples, the Halobacteria was present much more in the salty channel than in the tundra. However, the most significant variable of shaping different microbial communities was not the obvious; sampling locations, which were hypersaline in the channel and less in the tundra. The expected results of the channel sediment might be overtaken by the 133

consequence of temporary low salinity; nevertheless, we confirmed that the sampling season was a significant factor to shape the microbial communities. This confirmed factor may be due in part to the differences of temperatures, water activities, or the strength of sunlight. The detailed factors must be examined in further studies. With the sequences of the featured genes, amo A and hcd , we provided the potential partial metabolic pathways involved in nitrogen and carbon cycles and confirmed that the Thaumarchaea in the LH system has the ability to metabolize ammonia and inorganic carbon at the LH channel and the adjacent tundra. It enriched the knowledge regarding this five archaeal phylum in a relatively cold and salty environment and provided more genetic information for further studies. Studying microbial communities in such a hypersaline and cold environment will provide comparable knowledge as an analogue site for the microbial life search on other extraterrestrial bodies, including Mars, Europa, and

Enceladus.

4.6 Acknowledgements

This work was supported by the Canadian Astrobiology Training Program

(CATP), National Sciences and Engineering Research Council of Canada

(NSERC), Canadian Space Agency (CSA), Fonds de Recherche du Québec -

Nature et Technologies (FQRNT), Canada Foundation of Innovation (CFI), Polar

Continent Shelf Program (PCSP), and Northern Scientific Training Program

(NSTP). I especially thank the help from Dr. Yergeau for the UniFrac analysis. I also would like to thank that Dr. Mykytczuk and G. Lamarche-Gagnon collected 134

the samples and performed the CO 2 detection on site.

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Table 4-1. Geochemical measurement of the LH channel sediments and the adjacent tundra

Channel Channel Tundra Tundra (Summer 2011) (winter 2012) (Summer 2011) (Winter 2012) Total Nitrogen (%) 0.17±0.03 0.01±0.002 0.31±0.11 0.13±0.005 Total Carbon (%) 1.25±0.15 0.30±0.03 1.09±0.85 2.54±0.20 Organic Carbon (%) 0.77±0.08 0.20±0.02 0.51±0.14 2.48±0.14 Inorganic Carbon (%) 0.48±0.22 0.09±0.04 0.59±0.41 0.06±0.28 Ammonia (Liqiud) 0.26 mg/L 7.59 mg/L 0.66 mg/kg 4.62 mg/kg Nitrite/Nitrate (Liquid) 0.03 mg/L 0.12 mg/L 1.27 mg/kg 0.99 mg/kg Water/soil 4.1 -12.9 16.7 N.D. temperatures °C Air temperatures °C 15.6 -16.9 15.4 -16.9 Salinity (%) ~2(?) 25.8 N.D. N.D. CO2 flux µmol/m 2/s 0.38±0.02 0.59±0.12 0.09±0.01 0.05±0.05 N.D.: Not determined

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Table 4-2. Statistics and the indices of richness and diversity of the libraries. The four letters of the sample’s names indicate the sampling site, primer set, sampling season, and the extract’s type (pool) in order. The Derep reads indicates the numbers of dereplicated reads, which were the unique phylotypes, in the samples. Chao1 index indicates the potential numbers of phylotypes in the samples, as well as the Shannon index indicates the biodiversity.

Sample Site Primer Season Pool Reads Derep Chao1 Shannon set reads (98%) CBSC Channel Bacteria Summer cDNA 5817 816 1876 6.5 CBSD Channel Bacteria Summer DNA 5626 1150 2935 6.3 CASC Channel Archaea Summer cDNA 3965 183 305 2.9 CASD Channel Archaea Summer DNA 4546 943 1320 4.5 TBSC Tundra Bacteria Summer cDNA 2843 554 737 5.6 TBSD Tundra Bacteria Summer DNA 5283 1175 2479 6.4 TASC Tundra Archaea Summer cDNA 271 125 150 3.0 TASD Tundra Archaea Summer DNA 5738 761 1002 4.4 CBWC Channel Bacteria Winter cDNA 1454 309 373 4.7 CBWD Channel Bacteria Winter DNA 1487 356 472 5.3 CAWC Channel Archaea Winter cDNA 707 36 48 2.1 CAWD Channel Archaea Winter DNA 4 3 ND ND TBWC Tundra Bacteria Winter cDNA 7804 1704 2260 6.4 TBWD Tundra Bacteria Winter DNA 1726 342 408 5.3 TAWC Tundra Archaea Winter cDNA 29 4 ND ND TAWD Tundra Archaea Winter DNA 194 10 10 2.0

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Table 4-3. The F and P values of Permanova analyses for the samples based on their extract’s types (DNA and RNA), sampling season (Summer and Winter), and the locations (Channel and Tundra).

Extract Sampling season Sampling location

Archaea F-Value 1.5995 3.0249 0.40359

P-Value 0.2371 0.0259 0.775

Bacteria F-Value 1.1098 1.841 1.2572

P-Value 0.3152 0.031 0.2531

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Table 4-4. Unifrac distance based on weighted, normalized pairwise comparisons between bacterial communities from different (A) bacterial and (B) archaeal samples.

(A)

CBSC CBSD TBSC TBSD CBWC CBWD TBWC TBWD CBSC 0 0.116 0.146 0.201 0.227 0.21 0.2 0.183 CBSD 0.116 0 0.158 0.165 0.217 0.182 0.164 0.162 TBSC 0.146 0.158 0 0.159 0.188 0.188 0.165 0.161 TBSD 0.201 0.165 0.159 0 0.221 0.187 0.125 0.147 CBWC 0.227 0.217 0.188 0.221 0 0.195 0.18 0.199 CBWD 0.21 0.182 0.188 0.187 0.195 0 0.174 0.149 TBWC 0.2 0.164 0.165 0.125 0.18 0.174 0 0.156 TBWD 0.183 0.162 0.161 0.147 0.199 0.149 0.156 0

(B)

CASC CASD TASC TASD CAWC CAWD TAWC TAWD CASC 0 0.233 0.246 0.263 0.401 0.285 0.537 0.258 CASD 0.233 0 0.301 0.219 0.397 0.414 0.57 0.401 TASC 0.246 0.301 0 0.313 0.386 0.397 0.507 0.394 TASD 0.263 0.219 0.313 0 0.481 0.464 0.625 0.446 CAWC 0.401 0.397 0.386 0.481 0 0.455 0.394 0.452 CAWD 0.285 0.414 0.397 0.464 0.455 0 0.541 0.085 TAWC 0.537 0.57 0.507 0.625 0.394 0.541 0 0.538 TAWD 0.258 0.401 0.394 0.446 0.452 0.085 0.538 0

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Table 4-5. The comparison of the primer pairs used in the Lay et al. 2012 study and the present study . The abbreviation “V” indicates the variable regions of 16S rDNA sequences. The coverage indicates the ratio of the matched numbers of the primers and the numbers in the RDP database. The parentheses show the matched ratios of numbers/total numbers in the database.

2012 study Bacteria (V1 – V4) Archaea (V1 – V5)

Names of the primers 27F 758R 109F 934R

Sequences AGAGTTTGATCCTGGCTCAG CTACCAGGGTATCTAATCC ACKGCTCAGTAACACGT GTGCTCCCCCGCCAATTCCT

Coverage (%) 46.1 (254689/552836) 77.1 (1581485/2051347) 49.3 (32228/65386) 88.0 (61856/70293)

Numbers of Covered 35 (all) 33 (out of 35) a 3 (out of 5) b 3 (out of 5) b Phyla

Present study Bacteria (V1 – V3) Archaea (V4 – V5)

Names of the primers 28F 519R 517F 909R

Sequences GAGTTTGATCNTGGCTCAG GTNTTACNGNGGNKGCTG GCYTAAAGSRNCCGTAGC TTTCAGYCTTGCGRCCGTA

Coverage (%) 75.7 (418475/552836) 90.8 (1927529/2123836) 51.5 (60432/117233) 61.0 (59106/96881)

Numbers of Covered 35 (all) 35 (all) 4 (out of 5) c 4 (out of 5) c Phyla

a The sequences related to Dictyoglomi and Thermodesulfobacteria cannot be detected by 758R. b The sequences related to Nanoarchaeota and Thaumarchaeota cannot be detected by 109F and 934R. c The sequences related to Thaumarchaeota cannot be detected by 517F and 909R

128

Fig. 4-1. PCoA analyses for the microbial compositions of (A) bacterial and (B) archaeal reads of the 16S rDNA pyrosequencing libraries.

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Fig. 4-2. Summer 16S rDNA libraries of the channel and tundra. (A) Channel bacteria and (B) archaea, as well as the tundra (C) bacteria and (D) archaea. The bars in black and white represent RNA (cDNA) and DNA, respectively. The names of each library are followed by the numbers of raw reads.

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Fig. 4-3. Winter 16S rDNA libraries of the channel and tundra (A) Channel bacteria and (B) archaea, as well as the tundra (C) bacteria and (D) archaea. The bars in black and white represent RNA (cDNA) and DNA, respectively. The names of each library are followed by the numbers of raw reads.

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Fig. 4-4. Maximum-likelihood trees constructed by partial putative amino acid sequences of (A) amo A (173 positions) and (B) hcd (95 positions) cloned from LH channel sediments with other published related amino acid sequences. The cloned gene representatives were marked with the initials of “LH”. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) are shown next to the branches. Branches corresponding to partitions reproduced in less than 50% bootstrap replicates are collapsed.

132

Fig. 4-5. The copy numbers of Thaumarchaeal genes, including 16S ribosomal RNA (black), amo A (grey), and hcd (white) genes of the summer and winter samples of LH channel sediments and the adjacent tundra. The mean values of each gene are shown in the table under the figure. The values on top of the bars indicate the standard deviations.

133

CHAPTER 5

Discussion and Conclusions

The LH Spring system is a unique habitable environment for microorganisms. The spring outlet is a hypersaline (salinity = 24 – 26%), subzero ( -5°C), micro-oxic, highly-reduced, methane- and sulfate-rich environment. The spring channel is less extreme than the outlet, but the hypersalinity in the channel is generally as high as in the outlet. In the channel, the temperatures vary due to the ambient temperature. It is also more oxic than the outlet. In this study, we present different facets of microbiology approaches on the LH spring system in terms of microbial ecology, diversity and activities.

5.1 Microbial diversity and activity in the hypersaline spring channel

The LH spring channel is notable for the hypersalinity and the unfrozen stream flow at -18°C. We examined the microbial and geochemical characteristics of the LH outflow channels and then compared it to the previously characterized LH spring outlet. LH channel sediments contained greater microbial biomass (~100 fold) and greater microbial diversity, as reflected in the different species abundances in 16S rRNA clone libraries.

Phylotypes related to methanogenesis, methanotrophy, sulfur reduction and oxidation were detected in the bacterial clone libraries while the archaeal community was dominated by phylotypes most closely related to ammonia-oxidizing Thaumarchaeota .

14 C-acetate mineralization rates in channel sediment microcosms exceeded ~30 % and

~10 % at 5°C and -5°C, respectively, but sharply decreased at -10°C ( ≤ 1%). However, 134

we detected slight mineralization rate (0.17%) at -20°C. Most bacterial isolates

(Marinobacter, Planococcus , and Nesterenkonia spp.) were psychrotrophic, halotolerant, and capable of growth at -5°C. The physical and geochemical characteristics of the LH outlet were strongly related to the types of microbial metabolism found there, including anaerobic methane oxidation by ANME-1 archaea (Niederberger et al. 2010). The LH outflow channel described in this thesis, represents a distinct, heterogeneous, and stochastic environment occurring downstream of this subzero, hypersaline methane seep.

It contains the phylotypes: Halomonas , Gillisia , and Marinobacter, which are common bacteria in cold Arctic and Antarctic environments (Bowman et al. 1997; Bowman and

Nichols 2005; Brinkmeyer et al. 2003; Franzmann et al. 1987; Guan et al. 2009; Zhang et al. 2008). The ANME-1 group, however, was not detected in the channel sediments as it was in the outlet. We were the first to report the presence of Thaumarchaeota in an exreme hypersaline environment. A previous study published by Niederberger et al. in

2010 did not report this finding of Thaumarchaea. Overall, the LH spring channel has higher microbial diversity and activity than the outlet and supports a variety of niches in which diverse and metabolically active microbial communities exist.

5.2 Functional potential and the active components at LH outlet

The LH spring is the coldest and most saline terrestrial spring discovered to date and is defined by perennial discharges of subzero (-5°C), hypersaline (24% salinity), reducing

(≈-165 mV), and oligotrophic water. It is rich in sulfates (10.0% w/w), dissolved

-1 H2S/sulfides (up to 25 ppm), ammonia ( ≈381 µM), and methane (11.1 g d ). To determine its total functional potential and elucidate its active microbial components, 135

metagenomic and 16S ribosomal cDNA pyrosequencing analyses of the LH-spring outlet microbial community were preformed. Cyanobacteria (19.7%), Bacteroidetes (13.3%), and Proteobacteria (6.6%) were the dominant phyla identified in the spring outlet.

Reconstruction of the enzyme pathways responsible for bacterial nitrification/denitrification/ammonification and sulfate reduction appeared nearly complete in the metagenomic dataset. The key genes involved in methanogenesis/reverse-methanogenesis were of interest, but they were not obvious in the LH metagenome. In the LH 16S ribosomal cDNA active community profiles, ammonia oxidizers ( Thaumarchaeota ), denitrifiers ( Pseudomonas spp.), sulfate reducers

(Desulfobulbus spp.), and other sulfur oxidizers ( Thermoprotei ) were present, highlighting their involvement in nitrogen and sulfur cycling. The phylotypes of

Thermoprotei present in the active cDNA library from a perennial subzero spring is unique to this study. This is the first reported evidence of Thermoprotei being active in a cold environment.

Stress-response genes for adapting to cold, osmotic, and oxidative stress were abundant in the metagenome. Comparing functional community composition of the LH spring to metagenomes from other saline/subzero environments revealed a close association between LH and another Canadian High Arctic permafrost environment, the permafrost soils in Eureka, particularly in genes related to sulfur metabolism and dormancy. Overall, this study provides insights into the metabolic potential and the active microbial populations that exist in this hypersaline cryoenvironment and contributes to our understanding of microbial ecology in extreme environments.

5.3 Seasonal changes in microbial communities at a hypersaline spring channel and 136

the adjacent tundra

In our initial study on the LH channel sediment, we detected heterotrophic microbial metabolic activity (CO 2 mineralization recovery of 0.17%) at -20°C. Sixteen S rDNA pyrosequencing of the channel sediment and adjacent tundra sampled from summer (July) and winter (April) was performed using total microbial RNA and DNA. Microbial compositions of active (RNA) and DNA communities inhabiting the system were analyzed. In the summer, the LH channel sediment was dominated by the active groups,

Alphaproteobacteria and Betaproteobacteria . In the winter, Cyanobacteria ,

UniFrac, the analysis showed sampling seasons to be the most significant variant affecting the microbial composition in the sediments and tundra.

Signature genes of ammonia oxidizing archaea ( amo A and hcd ) were sequenced and analyzed for the phylogenetic affiliations with other published ones using their putative amino acid sequences from other environments. The result of phylogenetic tree showed similar patterns of grouping as 16S rDNA, especially using amo A. The amo A, hcd and the Thaumarchaeal 16S rDNA genes were quantified by qPCR in both sediment and tundra samples to support the genetic information regarding the LH Thaumarchaea. Copy number of Thaumarchaeal amo A and hcd genes in LH channel sediment and the adjacent tundra were roughly 10 to100 folds less than those reported in other similar environments.

Overall, this study provided the knowledge of the changes in active microbial communities existing in the extreme environment in two contrary seasons. It also enriched the insights of Thaumarchaea at a hypersaline and cold environment. 137

5.4 Conclusions

LH spring system is a fascinating site for microbiological study, since it is characterized by hypersalinity (~25%), subzero temperatures (perennial at the outlet and various in the channel), low redox potential (mainly in the outlet), microoxic (mainly in the outlet), and being rich in methane, sulfate and ammonia. However, it is a difficult location to reach for logistic reasons. Thus, this study is an important milestone for enriching the knowledge of environmental microbiology in extreme conditions. Here we demonstrate how the LH channel sediment is an environment with high microbial diversity and richness, responding to the environmental properties, including sulfate, methane, ammonia, hypersalinity, and cold/subzero temperatures. The mineralization assays showed that the LH channel sediment contained bio-active microorganisms down to -20°C. This result corresponded to that the heterotrophic microorganisms might be active and the biogeochemical cycling was occurring, when we recorded the LH channel water temperature was -18°C on site. The pyrosequencing libraries of the channel sediments originating from the winter samples revealed a wide range of microorganisms that maintained their metabolisms, including heterotrophic microorganisms, and the autotrophic Cyanocbacteria and Chloroflexi . The active 16S rDNA libriaries originating from the winter samples compensated for the deficiency of mineralization assays. The mineralization assays could only confirm the bulk activities of heterotrophic

14 microorganisms in the sediments by detecting C-CO 2. They were not able to reveal the detailed communities being activated among the sediments.

The study’s second important contribution was assembling a metageome for the LH outlet sediment. This demonstrated a detailed genetic potential for biogeochemical 138

cycling, including the genes related to sulfur, nitrogen and methane metabolisms. The LH metagenome revealed that metabolic pathways might be completed by several microorganisms in the environment. As the metagenome was based on the environmental

DNA, which implied the genetic potential at LH outlet, we also analyzed the active 16S rDNA libraries of LH outlet. It indicated fewer species might be active at the LH outlet.

Thus, for understanding the expressed genes at the LH outlet, a metatranscriptomic study should be considered in the future.

Thaumarchaea has been highlighted in all three studies in this thesis and in each chapter. We detected the 16S rDNA sequences in the clone library of LH channel sediment, which was the first report of the evidence of Thaumarchaea’s existence in the

LH spring system. The relatively high content of ammonia opens the possibility for the

LH spring system to be considered as an Enceladus’ analogous site for astrobiology research.

As in other environments, Thaumarchaea is a common archaea inhabiting the LH spring system, and is active either in the outlet or in the channel sediments based on the active 16S rDNA libraries. The results of cloning and sequencing the two featured functional genes, amo A and hcd , from the channel sediment, implied that Thaumarchaea might be one of the important primary producers at the LH spring system. This statement is not just based on confirmations of the functional genes, but also based on their abundance assessed by qPCR.

Our understanding regarding the microbiology of the LH spring system is limited to three published studies, which are two studies of the outlet sediments by Niederberger et al. in 2010 and Lay et al. in 2013; as well as one focused on the channel sediment by Lay et al. in 2012. In the future, research must address profiles of the active genes and 139

metabolic pathways, i.e., metatranscriptiomic study. This will elucidate the in situ biogeochemical pathways driven by certain types of microorganisms. This study may be extended to include samples from the outlet, channel and the adjacent tundra for a more complete comparison. There remain many questions to address about the LH

Thaumarchaea. Enrichment and isolation is the most direct way to study the adaptations of this kind of archaea inhabiting subzero and hypersaline environments. As long as obtaining the isolate, genomic study on LH Thaumarchaea will unveil all its adaptation strategies in different perspectives, including genomic, protein, and cell membranes. LH

Cyanobacteria is another microbial clade for examination, as it was present in the 16S libraries from both outlet and channel samples, and in the LH outlet metagenome, but only active in the channel sediment. These two groups of microorganisms may contribute a significant portion of the primary production to support the LH ecosystem. This study recommends long-term monitoring of the LH system pending the budgetary and logistical support. An ongoing study will establish a continuous environmental database regarding temperature, precipitations, and other geological properties. Based on this database, we may correspond to the geological changes to the microbial communities, especially for the relatively variable channel area. It also establishes a knowledge base for other types of studies, including environmental chemistry, geology, hydrology, and meterology.

The microbial study on extreme environment like the LH spring system may be applied to environmental remediation, pharmaceutical industry, and biomining/biorefining. Also, it also provides a model for other microbiological studies on similar hypersaline and cold/subzero environments; as well as it is an analogous site for extraterrestrial bodies for microbial life search.

140

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APPENDIX: Supporting Information Table S2-1. Growth in various NaCl concentrations and temperatures for 22 isolates strains Colors R2A with 0% NaCl R2A with 7% NaCl R2A with 12% NaCl R2A with 20% NaCl

37°C 25°C 5°C -5°C 37°C 25°C 5°C -5°C 37°C 25°C 5°C -5°C 37°C 25°C 5°C -5°C

CY-C3-1 Orange - + + - + + + + - + + - - + - -

CY-C3-2 White - - +/- - - + + + - + + + - + - -

CY-C3-3 Orange - - - - - + + + - + + + - + - -

CY-C3-4 Orange - + + - + + + + + + + - - - - -

CY-C3-5 Orange - + + - + + + + + + + - - - +/- -

CY-C3-6 Orange - + + + + + + + - + + + - + - -

CY-C3-7 White - + + - - + + + - - + - - - - -

CY-C3-8 White - - - - - + + + ------

CY-C1-9 White/Red - - - - - + + + - + + + - + - -

CY-C1-10 Light Red - + + - - + + - - + + - - + - -

CY-C1-11 Orange - + + - + + + + - + + - - - - -

CY-C1-12 Orange - + + - + + + + - + + - - - - -

CY-C2-13 Light Red - + + - - + + - - + + - - + - -

CY-C2-14 Light Red - + + - - + + - - + + - - + - -

CY-C2-15 Deep Red - - - - - + + + - + + + - + - -

CY-C2-16 Orange - + + - - + + + - + ------

CY-C2-17-1 White/Red - - - - - + + + - + - - - + + -

CY-C2-17-2 Light Red - + + - + + + - - + - - - + - -

CY-C2-18 Yellow - + + - - + + ------

CY-C2-19-1 Orange - + + - - + + - - + - - - + + -

CY-C2-19-2 Yellow - + + - + + + + - + ------

CY-C2-20 Orange - + + - + + + - + + + - - - - -

182

Table S2-2 –Relevant taxonomic proportions of sequences from Bacterial 16S rRNA gene clone libraries a Bacterial C1 C2 C3 LH source b

Phyla/ Class No. % No. % No. % %

Bacteroidetes 51 64 96 56 77 33 44

Alphaproteobacteria 17 21 11 6 52 22 21

Betaproteobacteria - - - - 17 7 2

Gammaproteobacteria - - 13 7 41 17 29

Deltaproteobacteria - - 1 1 3 1 -

Epsilonproteobacteria - - - - 2 1 -

Actinomycetes 7 9 49 29 33 14 -

Firmicutes 4 5 1 1 10 4 2

Deinococci 1 1 - - - - -

Verrucomicrobia - - - - 1 1 -

Cyanobacteria ------2

Total 80 100 171 100 236 100 100 a Libraries constructed from samples from Lost Hammer spring channel sediments. Sequences were grouped using the RDP Classifier function of the Ribosomal Database Project-II release 9 with a confidence threshold of 80%. b The data from the LH outlet sediment were taken from Niederberger et al. 2010.

183

Table S2-3 - Relevant taxonomic proportions of sequences from archaeal 16S rRNA gene clone libraries a

Archaeal Phylum/Class C1 C2 C3 LH source b

No. % No. % No. % %

Halobacteria 7 12.5 - - 22 27.5 8

Unclassified Euryarchaeota 3 3.6 - - - - -

Methanobacteria - - - - 25 31.2 -

Thaumarchaeota 70 83.9 24 100 33 41.3 -

Archaeoglobaceae ------2

ANME-1a ------46.8

KTK 4A (93% similarity) c ------9

ABBA-25 (91% similarity) d ------34.2

Total 80 100 24 100 80 100 100 a Libraries constructed from samples from Lost Hammer spring channel sediments. Sequences were grouped using the RDP Classifier function of the Ribosomal Database Project-II release 9 with a confidence threshold of 80%. b The data from the LH outlet sediment were taken from Niederberger et al. 2010. c KTK 4A was obtained from a highly saline sediment in the Red Sea. (Eder et al. 1999) d ABBA-25 was from a deep anoxic hypersaline basin. (van der Wielen et al. 2005)

Table S3-1. Pearson product-moment correlation coefficients of the LH metagenome taxonomic and functional reads under different cutoff E values ≤ 10 -5, 10 -10 , 10 -15, and 10 -20

Taxonomy 1.00E-10 1.00E-15 1.00E-20 1.00E-05 R 1 0.987563 0.73459 P-value < 2.2e-16 < 2.2e-16 1.93E-05 Function 1.00E-05 R 0.999999 0.995174 0.908828 P-value < 2.2e-16 < 2.2e-16 2.27E-11

184

Table S3-2. The percentages of genera related to the genes encoding enzymes facilitating nitrogen cycles in LH metagenome Genus/Enzyme/EC % Phylum/Class of Proteobacteria Copper-containing nitrite reductase (EC 1.7.2.1) Burkholderia 2.4 Betaproteobacteria Bdellovibrio 15 Deltaproteobacteria Propionibacterium 1.2 Actinobacteria Gramella 3.6 Bacteroidetes Kangiella 23 Gammaproteobacteria Neisseria 7.1 Betaproteobacteria Maribacter 13 Bacteroidetes Flavobacterium 32 Bacteroidetes Leeuwenhoekiella 1.2 Bacteroidetes Flavobacteria 1.2 Bacteroidetes Nitrous-oxide reductase (EC 1.7.99.6/1.7.2.4) Magnetospirillum 11 Alphaproteobacteria Campylobacter 68 Deltaproteobacteria Nitratiruptor 3.6 Deltaproteobacteria Sulfurovum 11 Deltaproteobacteria Magnetospirillum 3.6 Alphaproteobacteria Gramella 3.6 Bacteroidetes Nitrogenase (EC 1.18.6.1) Nostoc 50 Cyanobacteria Cyanothece 50 Cyanobacteria Nitrite reductase [NAD(P)H] (EC 1.7.1.4) Thermobaculum 1.4 Bacteroidetes Pseudomonas 1.4 Gammaproteobacteria Cytophaga 10 Cyanobacteria Escherichia 2.9 Gammaproteobacteria Spirosoma 7.2 Bacteroidetes Klebsiella 2.9 Bacteroidetes

185

Polaromonas 1.4 Betaproteobacteria Geobacillus 1.4 Firmicutes Polaribacter 11.6 Bacteroidetes Saccharophagus 1.4 Gammaproteobacteria Acinetobacter 2.8 Gammaproteobacteria Chitinophaga 7.2 Bacteroidetes Dyadobacter 2.9 Bacteroidetes Psychrobacter 10 Gammaproteobacteria Mycobacterium 1.4 Bacteroidetes Rhodoferax 1.4 Betaproteobacteria Dokdonia 8.7 Bacteroidetes Leptothrix 2.9 Betaproteobacteria Psychromonas 4.3 Gammaproteobacteria Methylococcus 2.8 Gammaproteobacteria Thiobacillus 1.4 Betaproteobacteria Sorangium 5.8 Deltaproteobacteria Croceibacter 5.8 Bacteroidetes Ferredoxin--nitrite reductase (EC 1.7.7.1) Synechocystis 24 Cyanobacteria Cyanothece 16 Cyanobacteria Trichodesmium 8 Cyanobacteria Nostoc 44 Cyanobacteria Thermosynechococcus 4 Cyanobacteria Microcystis 4 Cyanobacteria Assimilatory nitrate reductase (EC 1.7.99.4) RhodoPseudomonas 0.7 Alphaproteobacteria Flavobacterium 0.7 Bacteroidetes Psychrobacter 30 Gammaproteobacteria Thermobaculum 0.7 Unclassidied Bacteria Streptosporansium 0.7 Actinobacteria Bacillus 5.7 Firmicutes Trichodesmium 1.4 Cyanobacteria Mycobacterium 0.7 Bacteroidetes Methylococcus 1.4 Gammaproteobacteria

186

Burkholderia 2.8 Betaproteobacteria Pectobacterium 0.7 Gammaproteobacteria Cyanothece 4.3 Cyanobacteria Shewanella 0.7 Gammaproteobacteria Pseudomonas 0.7 Gammaproteobacteria Bradyrhizobium 2.2 Gammaproteobacteria Azoarcus 0.7 Betaproteobacteria Alcanivorax 1.4 Gammaproteobacteria Anabaena 6.5 Cyanobacteria Chitinophaga 0.7 Bacteroidetes Cytophaga 2.9 Bacteroidetes Rhodococcus 0.7 Actinobacteria Marinomonas 0.7 Gammaproteobacteria Roseobacter 1.4 Alphaproteobacteria Nostoc 12 Cyanobacteria Methylibium 2.2 Betaproteobacteria Pseudoalteromonas 0.7 Gammaproteobacteria Microcystis 2.2 Cyanobacteria Polaromonas 0.7 Betaproteobacteria Novosphingobium 0.7 Alphaproteobacteria Crocosphaera 0.7 Cyanobacteria Thiobacillus 2.2 Betaproteobacteria Pseudomonas 0.7 Gammaproteobacteria Azorhizobium 2.2 Alphaproteobacteria Acidovorax 1.4 Betaproteobacteria Maribacter 0.7 Bacteroidetes Acinetobacter 2.2 Gammaproteobacteria Mesorhizobium 0.7 Alphaproteobacteria Rhizobium 1.4 Alphaproteobacteria

187

Table S3-3. The percentages of genera related to the genes encoding enzymes facilitating Sulfate reduction in LH metagenome Genus/Enzyme/EC % Phylum/Class of Proteobacteria Adenylylsulfate kinase (EC 2.7.1.25) Roseiflexus 16.3 Chloroflexi Aquifex 16.3 Aquificae Chloroflexus 5.8 Chloroflexi Schizosaccharomyces 5.2 Opisthokonta Caulobacter 4.6 Alphaproteobacteria Halothermothrix 4.7 Firmicutes Xanthomonas 3.5 Gammaproteobacteria Paracoccus 2.9 Alphaproteobacteria Zymomonas 2.9 Alphaproteobacteria Trichodesmium 2.3 Cyanobacteria Cyanothece 2.3 Cyanobacteria Bacteroides 1.7 Bacteroidetes Erythrobacter 1.7 Alphaproteobacteria Magnetococcus 1.7 Alphaproteobacteria Microcystis 1.7 Cyanobacteria Solibacter 1.7 Acidobacteria Francisella 1.8 Gammaproteobacteria Pseudomonas 1.8 Gammaproteobacteria Vibrio 1.2 Gammaproteobacteria Burkholderia 1.2 Betaproteobacteria Chlorobium 1.2 Bacteroidetes Deinococcus 1.2 Deinococcus Frankia 1.2 Actinobacteria Hydrogenobaculum 1.2 Aquificae Oceanobacillus 1.2 Firmicutes Rhodopseudomonas 1.2 Alphaproteobacteria Synechocystis 1.2 Cyanobacteria Ashbya 0.6 Opisthokonta Aurantimonas 0.6 Alphaproteobacteria

188

Campylobacter 0.6 Deltaproteobacteria Crocosphaera 0.6 Cyanobacteria Delftia 0.6 Betaproteobacteria Geobacter 0.6 Deltaproteobacteria Methylococcus 0.6 Gammaproteobacteria Mycobacterium 0.6 Actinobacteria Neurospora 0.6 Opisthokonta Oceanicaulis 0.6 Alphaproteobacteria Parvibaculum 0.6 Alphaproteobacteria Pelobacter 0.6 Deltaproteobacteria Prochlorococcus 0.6 Cyanobacteria Roseobacter 0.6 Alphaproteobacteria Shewanella 0.6 Gammaproteobacteria Sphingomonas 0.6 Alphaproteobacteria Thermobispora 0.6 Actinobacteria Thermosynechococcus 0.6 Cyanobacteria Phosphoadenylyl-sulfate reductase [thioredoxin] (EC 1.8.4.8)/ Adenylyl-sulfate reductase [thioredoxin] (EC 1.8.4.10) Marinobacter 29.2 Gammaproteobacteria Acinetobacter 17 Gammaproteobacteria Azotobacter 9.8 Gammaproteobacteria Reinekea 9.8 Gammaproteobacteria Cyanothece 4.8 Cyanobacteria Pseudomonas 4.8 Gammaproteobacteria Crocosphaera 4.9 Cyanobacteria Hahella 4.9 Gammaproteobacteria Alkalilimnicola 2.4 Gammaproteobacteria Anabaena 2.4 Cyanobacteria Bordetella 2.4 Betaproteobacteria Cellvibrio 2.4 Gammaproteobacteria Croceibacter 2.4 Bacteroidetes Nostoc 2.4 Cyanobacteria Sulfate adenylyltransferase,

189

dissimilatory-type (EC 2.7.7.4) Aquifex 12.6 Aquificae Roseiflexus 12.7 Chloroflexi Anabaena 5.9 Cyanobacteria Chloroflexus 4.5 Chloroflexi Cytophaga 3.6 Cyanobacteria Synechococcus 4.2 Cyanobacteria Caulobacter 3.7 Alphaproteobacteria Nitrosococcus 1.8 Gammaproteobacteria Xanthomonas 3.3 Gammaproteobacteria Nostoc 2.7 Cyanobacteria Maribacter 3.7 Bacteroidetes Paracoccus 2.3 Alphaproteobacteria Microcystis 1.8 Cyanobacteria Thermosynechococcus 1.8 Cyanobacteria Pseudomonas 1.4 Gammaproteobacteria Trichodesmium 1.4 Cyanobacteria Erythrobacter 1.4 Alphaproteobacteria Francisella 1.4 Gammaproteobacteria Leeuwenhoekiella 2.3 Bacteroidetes Magnetococcus 1.4 Alphaproteobacteria Pseudoalteromonas 1.9 Gammaproteobacteria Robiginitalea 2.3 Bacteroidetes Streptosporangium 1.4 Actinobacteria Aeropyrum 0.9 Crenarchaeota Bacillus 0.9 Firmicutes Deinococcus 0.9 Deinococcus Hydrogenobaculum 0.9 Aquificae Pyrococcus 0.9 Euryarchaeota Frankia 0.9 Actinobacteria RhodoPseudomonas 0.9 Alphaproteobacteria Saccharophagus 0.9 Gammaproteobacteria Cyanothece 0.5 Cyanobacteria Ferroplasma 0.5 Euryarchaeota Neurospora 0.5 Opisthokonta

190

Petrotoga 0.5 Thermotogae Roseobacter 0.5 Alphaproteobacteria Staphylococcus 0.5 Firmicutes Staphylothermus 0.5 Crenarchaeota Thermobispora 0.5 Actinobacteria Flavobacterium 0.5 Bacteroidetes Geobacter 0.5 Deltaproteobacteria Hahella 0.5 Gammaproteobacteria Kytococcus 0.5 Actinobacteria Opitutus 1 Verrucomicrobia Psychromonas 1 Gammaproteobacteria Aurantimonas 0.5 Alphaproteobacteria Chromobacteriu m 0.5 Betaproteobacteria Leptospira 0.5 Spirochaetes Methylococcus 0.5 Gammaproteobacteria Mycobacterium 0.5 Actinobacteria Nocardioides 0.5 Actinobacteria Oceanicaulis 0.5 Alphaproteobacteria Parvibaculum 0.5 Alphaproteobacteria Pelobacter 0.5 Deltaproteobacteria Salmonella 0.5 Gammaproteobacteria Shewanella 0.5 Gammaproteobacteria Sphingomonas 0.5 Alphaproteobacteria Nitrosomonas 1.8 Betaproteobacteria Sulfite reductase (EC 1.8.99.1)/ Ferredoxin--sulfite reductase (EC 1.8.7.1)/ Sulfite reductase [NADPH] hemoprotein (EC 1.8.1.2)/ Sulfite reductase, dissimilatory-type (EC 1.8.99.3) Thiobacillus 31.3 Betaproteobacteria Nostoc 10.8 Cyanobacteria Cyanothece 9.9 Cyanobacteria Synechocystis 8.9 Cyanobacteria Alkalilimnicola 6.3 Gammaproteobacteria

191

Anabaena 5.4 Cyanobacteria Thioalkalivibrio 4.5 Gammaproteobacteria Microcystis 3.6 Cyanobacteria Xanthomonas 2.7 Gammaproteobacteria Magnetospirillum 2.7 Alphaproteobacteria Xylella 1.8 Gammaproteobacteria Vesicomyosocius 1.8 Gammaproteobacteria Trichodesmium 1.8 Cyanobacteria Shewanella 0.9 Gammaproteobacteria Polaribacter 0.9 Bacteroidetes Nitrosospira 0.9 Betaproteobacteria Myxococcus 0.9 Deltaproteobacteria Halorhodospira 0.9 Gammaproteobacteria Flavobacterium 0.9 Bacteroidetes Crocosphaera 0.9 Cyanobacteria Chromobacterium 0.9 Betaproteobacteria Acinetobacter 0.9 Gammaproteobacteria Sulfur oxidation protein SoxB Thiomicrospira 20.2 Gammaproteobacteria Chlorobium 15.9 Bacteroidetes Dechloromonas 12.6 Betaproteobacteria Herminiimona s 7.9 Betaproteobacteria Thiobacillus 6.3 Betaproteobacteria Congregibacter 5.6 Gammaproteobacteria Janthinobacterium 5.3 Betaproteobacteria Chlorobac ulum 4.3 Bacteroidetes Polaromonas 3.6 Betaproteobacteria Bradyrhizobium 3.3 Alphaproteobacteria Ralstonia 3.0 Betaproteobacteria Vesicomyo socius 2.6 Gammaproteobacteria Pelodictyon 2.0 Bacteroidetes Thioalkalivibrio 1.3 Gammaproteobacteria Nitrobacter 1.3 Alphaproteobacteria Acidi philium 1.3 Alphaproteobacteria Polynucleobacter 1.0 Betaproteobacteria

192

Methylobacterium 0.7 Alphaproteobacteria Arcobacter 0.7 Deltaproteobacteria Oligotropha 0.3 Alphaproteobacteria Anaeromyxobacter 0.3 Deltaproteobacteria

193

Table S3-4. Representatives of genes/proteins related to oxidative stress in LH metagnome Func tion Protein Unique Total hits Phyla names hits Oxidative Catalase 67 29592 Bacteroidetes (74.0%), stress Cyanobacteria (11.0%), Proteobacteria (11.6%), Acidobacteria (1.1%), Chlorobi (0.3%), Actinobacteria (1.3%), Firmicutes (0.5%), Deinococcus-Thermus (0.2%) Superoxide 46 57074 Cyanobacteria (66.9%), dismutase Bacteroidetes (29.4%) , Chloroflexi (0.4%), Proteobacteria (2.9%), Bacteria (0.3%), Firmicutes (0.3%), Thermotoga (0.1%) Peroxidase 41 18062 Bacteroidetes (65.8%), Cyanobaceria (18.0%), Proteobacteria (11.9%), Acidobacteria (1.7%), Chlorobi (0.5%), Firmicutes (0.5%), Actinobacteria (1.6%) Iron-binding 17 19709 Cyanobacteria (28.4%), ferritin-like Bacteroidetes (66.2%), antioxidant Proteobacteria (5.4%) protein Organic 11 609 Verrucomicrobia (71.8%), hyd roperoxide Firmicutes (28.2%) resistant protein

194

Table S3-5. Representatives of genes/proteins related to osmotic stress in LH metagenome Function Protein names Unique Total Phyla hits hits Synthesis of Cyclic beta-1,2-glucan 34 4475 Acidobacteria (12.1%), osmoregulated periplasmic synthase Proteobacteria (57.8%), glucan Chloroflexi (15.6%), Planctomycetes (5.4%), Firmicutes (7.3), Euryarchaeota (1.8) Glucans biosynthesis 3 2563 Bacteroidetes (95.3%), protein C Proteobacteria (4.7%) Choline and Betaine Sarcosine oxidase 30 10504 Acidobacteria (7.4%), Uptake and Betaine Actinobacteria (30.0%), Biosynthesis Proteobacteria (62.6%) L-proline glycine 14 4233 Acidobacteria (64.6%), betaine ABC transport Proteobacteria (30.0%), system permease Actinobacteria (3.2%), protein Firmicutes (2.2%) Choline dehydrogenase 12 8386 Cyanobacteria (88.4%), Proteobacteria (6.9%), Actinobacteria (4.7%) Betaine aldehyde 2 7 Fungi (85.7%), dehydrogenase Proteobacteria (14.3%)

195

Table S3-6. Representatives of genes/proteins related to cold adaptation in LH metagenome Functions Protein or subsystem Unique Total hits Phyla names hits DNA replication gyrA 96 58535 Cyanobacteria (55.6%), (DNA Gyrase A) Bacteroidetes (40.8%), Proteobacteria (2.3%), Actinobacteria (0.6%), Firmicutes (0.7%) recA 36 28954 Cyanobacteria (48.4%), (Recombination factor Bacteroidetes (46.5%), A) Proteobacteria (2.6%), Chlorobi (0.3%), Deinococcus-Thermus (0.3%), Actinobacteria (1.8%), Chloroflexi (0.1%) dnaA (Replication 30 28928 Cyanobacteria (33.7%), initiator Protein) Bacteroidetes (63.4%), Actinobacteria (1.0%), Proteobacteria (1.5%), Deinococcus-Thermus (0.4%) Unsaturated fat ty acids Fatty acid desaturases 36 54030 Cyanobacteria (99.7%), Proteobacteria (0.3%) dnaJ 53 57229 Cyanobacteria (75.3%), Bacteroidetes (21.3%), Proteobacteria (2.9%), Firmicutes (0.4%), Actinobacteria (0.1%) Protein folding Prolyl-isomerase 64 7063 Bacteroidetes (60.5%), Proteobacteria (33.3%), Acidobacteria (4.4%), Cyanobacteria (1.5%), Synergistetes (0.3%) Nucleosides and aceE 79 55118 Cyanobacteria (57.0%),

196

Nucleotides Bacteroidetes (38.5%), Acidobacteria (0.6%), Proteobacteria (3.0%), Chloroflexi (0.4%), Actinobacteria (0.3%), Deinococcus-Thermus (0.1%), Chlamydiae (0.1%) Pyruvate metabolism aceF 72 52841 Cyanobacteria (49.6%), Bacteroidetes (43.3%) Proteobacteria (6.1%), Acidobacteria (0.6%), Actinobacteria (0.2%), Deinococcus-Thermus (0.1%), Firmicutes (0.1%) Transcription nusA 26 28035 Cyanobacteria (49.4%), Bacteroidetes (45.3%), Bacteria (0.5%), Firmicutes (1.2%), Actinobacteria (3.6%) RNA helicase Cold-shock DEAD-box 61 25563 Bacteroidetes (49.8%) protein A Cyanobacteria (48.4%) Proteobacteria (1.6%) Methanococcales (0.1%) Firmicutes (0.1%) Verrucomicrobia (1.0%) cspA 9 14 Bacteroidetes (71.4%) Proteobacteria (21.4%) Firmicutes (7.2%)

197

Table S3-7. Dereplication hits of active bacterial 16S ribosomal cDNA based on 98% similarity No. of Dereplicated Best hit Accession Identity reads representative No. 103 SB-1 Verrucomicrobia bacterium HQ675558.1 94% SCGC AAA240-C14 84 SB-2 Stenotrophomonas sp. NOE8 JX842835.1 100% 73 SB-3 Microlunatus panaciterrae NR_041517. 93% strain Gsoil 954 1 73 SB-4 Rhodothalassium sp. PHT1 HE806302.1 91% 65 SB-5 Pseudomonas aeruginosa JF899310.2 99% strain PM389 63 SB-6 Roseateles sp. R-45571 FR775142.1 98% 60 SB-7 Verrucomicrobia bacterium HQ675558.1 95% SCGC AAA240-C14 55 SB-8 Pedobacter heparinus AB680215.1 98% 54 SB-9 Pseudomonas aeruginosa JF899310.2 99% strain PM389 43 SB-10 Enterobacter sp. DHL-02 AB714445.1 99% 38 SB-11 Staphylococcus aureus subsp. HE579073.1 99% aureus ST228 31 SB-12 Desulfobulbus sp. AF132865.1 99% 30 SB-13 Stenotrophomonas sp. NOE8 JX842835.1 99% 22 SB-14 Microlunatus panaciterrae NR_041517. 93% strain Gsoil 954 1 17 SB-15 Delftia lacustris HE861943.1 100% 9 SB-16 Acidovorax sp. CNE 29 FR749857.1 99% 7 SB-17 Enterobacter sp. DHL-02 AB714445.1 99% 7 SB-18 Desulfobulbus sp. AF132865.1 98% 6 SB-19 Pseudomonas aeruginosa JF899310.2 98% strain PM389 6 SB-20 Enterobacter sp. DHL-02 AB714445.1 99% 6 SB-21 Stenotrophomonas sp. NOE8 JX842835.1 99% 6 SB-22 Pseudomonas sp. RB5-M5 JN019027.1 97% 5 SB-23 Staphylococcus aureus CP003808.1 99%

198

08BA02176 5 SB-24 Marinobacter sp. V3H-008 JN106689.1 98% 5 SB-25 Verrucomicrobia bacterium HQ675558.1 95% SCGC AAA240-C14 5 SB-26 Microlunatus panaciterrae NR_041517. 94% strain Gsoil 954 1 5 SB-27 Pseudomonas aeruginosa JF899310.2 99% strain PM389 5 SB-28 Desulfobulbus sp. AF132865.1 99% 4 SB-29 Staphylococcus aureus subsp. HE579073.1 99% aureus ST228 4 SB-30 Microlunatus panaciterrae NR_041517. 93% 1 4 SB-31 Stenotrophomonas sp. NOE8 JX842835.1 99% 4 SB-32 Pedobacter heparinus AB680215.1 98% 4 SB-33 Microlunatus panaciterrae NR_041517. 93% strain Gsoil 954 1 4 SB-34 Microlunatus panaciterrae NR_041517. 93% strain Gsoil 954 1 4 SB-35 Microlunatus panaciterrae NR_041517. 94% strain Gsoil 954 1 3 SB-36 Verrucomicrobia bacterium HQ675558.1 96% SCGC AAA240-C14 3 SB-37 Stenotrophomonas maltophilia JF431276.1 98% strain BXCC-58 3 SB-38 Pseudomonas sp. PC IW 25 FM164626.1 98% 3 SB-39 Pseudomonas aeruginosa JF899310.2 99% strain PM389 3 SB-40 Stenotrophomonas sp. NOE3 JX842830.1 99% 3 SB-41 Pseudomonas aeruginosa JN969597.1 92% strain 9Cit 3 SB-42 Pedobacter heparinus AB680215.1 97% 3 SB-43 Staphylococcus aureus subsp. HE579073.1 99% aureus ST228 3 SB-44 Verrucomicrobia bacterium HQ675558.1 91%

199

SCGC AAA240-C14 3 SB-45 Verrucomicrobia bacterium HQ675558.1 97% SCGC AAA240-C14 3 SB-46 Verrucomicrobia bacterium HQ675558.1 95% SCGC AAA240-C14 3 SB-47 Delftia lacustris HE861943.1 99% 3 SB-48 Pseudomonas aeruginosa JF899310.2 96% strain PM389 3 SB-49 Mitsuaria sp. H29L1B EU714912.1 96% 3 SB-50 Tessaracoccus sp. GU111568.2 93% SL014B-79A 3 SB-51 Staphylococcus aureus subsp. HE579073.1 97% aureus ST228 2 SB-52 Pseudomonas aeruginosa JQ900543.1 92% strain N83 2 SB-53 Pseudomonas aeruginosa AB037548.1 96% 2 SB-54 Pseudomonas aeruginosa DQ666628.1 94% strain RsB-29 2 SB-55 Staphylococcus aureus subsp. HE579073.1 99% aureus ST228 2 SB-56 Stenotrophomonas maltophilia GU815943.1 96% strain GGI-22 2 SB-57 Enterobacter sp. DHL-02 AB714445.1 98% 2 SB-58 Roseateles sp. AM989118.1 95% AKB-2008-KU7 2 SB-59 Rhodothalassium salexigens FR682008.1 90% 2 SB-60 Microlunatus panaciterrae NR_041517. 92% strain Gsoil 954 1 2 SB-61 Staphylococcus aureus subsp. HE579073.1 99% aureus ST228 2 SB-62 Afifella marina strain P530 GU370095.1 88% 2 SB-63 Roseateles sp. R-45571 FR775142.1 95% 2 SB-64 Klebsiella oxytoca E718 CP003683.1 99% 2 SB-65 Verrucomicrobia bacterium HQ675558.1 96% SCGC AAA240-C14

200

2 SB-66 Stenotrophomonas maltophilia GU420674.1 97% clone FH030 2 SB-67 Pseudomonas aeruginosa AY499109.1 99% 2 SB-68 Sphingomonadaceae AB269802.2 91% bacterium KF016 2 SB-69 Pedobacter heparinus AB680215.1 94% 2 SB-70 Stenotrophomonas sp. NOE8 JX842835.1 98% 2 SB-71 Desulfobulbus sp. AF132865.1 91% 2 SB-72 Stenotrophomonas sp. NOE8 JX842835.1 97% 2 SB-73 Alcaligenes faecalis strain N8 EU567029.1 98% 2 SB-74 Pseudomonas sp. AMAAS232 JN391539.1 98% 2 SB-75 Staphylococcus aureus subsp. HE579073.1 99% aureus ST228 2 SB-76 Staphylococcus aureus subsp. HE579073.1 96% aureus ST228 2 SB-77 Microlunatus panaciterrae NR_041517. 91% strain Gsoil 954 1 2 SB-78 Acidovorax sp. CNE 29 FR749857.1 97% 2 SB-79 Desulfobulbus sp. AF132865.1 97% 2 SB-80 Stenotrophomonas sp. NOE8 94% 2 SB-81 Verrucomicrobia bacterium HQ675558.1 93% SCGC AAA240-C14 1 SB-82 Pedobacter heparinus AB680215.1 98% 1 SB-83 Alcaligenes faecalis strain N8 EU567029.1 98% 1 SB-84 Verrucomicrobia bacterium HQ675558.1 94% SCGC AAA240-C14 1 SB-85 Sphingomonadaceae AB269802.2 90% bacterium KF016 1 SB-86 Mitsuaria sp. H29L1B EU714912.1 89% 1 SB-87 Pseudomonas aeruginosa AB037548.1 94% 1 SB-88 Alcaligenes faecalis strain N8 EU567029.1 97% 1 SB-89 Pseudomonas aeruginosa JF513140.1 94% strain S85R 1 SB-90 Roseateles sp. R-45571 FR775142.1 94% 1 SB-91 Enterobacter sp. 2391 JX174268.1 92%

201

1 SB-92 Afifella marina strain P530(0) GU370095.1 88% 1 SB-93 Stenotrophomonas sp. NOE8 JX842835.1 94% 1 SB-94 Pseudomonas aeruginosa FJ556919.1 95% strain NGKCTS 1 SB-95 Pseudomonas aeruginosa HQ018741.1 98% strain ASFP-38 1 SB-96 Roseateles sp. MC12 AB013425.1 92% 1 SB-97 Desulfobulbus sp. AF132865.1 97% 1 SB-98 Pseudomonas aeruginosa AY499109.1 97% strain TERIPS9002 1 SB-99 Porphyrobacter cryptus FR774566.1 87% 1 SB-100 Porphyrobacter cryptus FR774566.1 89% 1 SB-101 Roseateles sp. R-45571 FR775142.1 97% 1 SB-102 Pedobacter heparinus AB680215.1 97% 1 SB-103 Pseudomonas aeruginosa GU212673.1 99% strain XRF-6 1 SB-104 Desulfobulbus sp. AF132865.1 99% 1 SB-105 Sphingomonadaceae AB269802.2 89% bacterium KF016 1 SB-106 Pseudomonas aeruginosa JF899310.2 97% strain PM389 1 SB-107 Sphingomonadaceae AB269802.2 91% bacterium KF016 1 SB-108 Sphingomonadaceae AB269802.2 88% bacterium KF016 1 SB-109 Enterobacter oryzae strain JQ659749.1 95% R5-362 1 SB-110 Stenotrophomonas sp. NOE8 JX842835.1 99% 1 SB-111 Pseudomonas aeruginosa JF899310.2 98% strain PM389 1 SB-112 Sphingomonas sp. V3M21 FN794222.1 93% 1 SB-113 Pedobacter heparinus AB680215.1 96% 1 SB-114 Acidovorax sp. CNE 29 FR749857.1 91% 1 SB-115 Microlunatus sp. M5_21 AB468984.1 90% 1 SB-116 Desulfobulbus sp. AF132865.1 97%

202

1 SB-117 Tessaracoccus sp. GU111568.2 89% SL014B-79A 1 SB-118 Verrucomicrobia bacterium HQ675558.1 92% SCGC AAA240-C14 1 SB-119 Microlunatus panaciterrae NR_041517. 92% strain Gsoil 954 1 1 SB-120 Pedobacter heparinus AB680215.1 98% 1 SB-121 Desulfobulbus sp. AF132865.1 91% 1 SB-122 Porphyrobacter cryptus FR774566.1 91% 1 SB-123 Pseudomonas aeruginosa JF899310.2 99% strain PM389 1 SB-124 Roseateles sp. R-45571 FR775142.1 95% 1 SB-125 Staphylococcus aureus subsp. HE579073.1 96% aureus ST228 1 SB-126 Roseateles sp. R-45571 FR775142.1 94% 1 SB-127 Enterobacter sp. SP1 JQ001784.1 99% 1 SB-128 Pseudomonas sp. PC IW 25 FM164626.1 97% 1 SB-129 Pseudomonas aeruginosa AB062598.1 96% 1 SB-130 Musa acuminata EU017026.1 99% 1 SB-131 Pseudomonas aeruginosa AY499109.1 97% strain TERIPS9002 1 SB-132 Verrucomicrobia bacterium HQ675558.1 96% SCGC AAA240-C14 1 SB-133 Pedobacter heparinus AB680215.1 91% 1 SB-134 Pseudomonas stutzeri EU520400.1 90% 1 SB-135 Microlunatus panaciterrae NR_041517. 93% strain Gsoil 954 1 1 SB-136 Tessaracoccus sp. GU111568.2 92% SL014B-79A 1 SB-137 Pedobacter heparinus AB680215.1 93% 1 SB-138 Pseudomonas aeruginosa JQ927361.1 100% strain M10 1 SB-139 Sneathiella sp. BFLP-8 FN687912.1 87% 1 SB-140 Pseudomonas sp. RB5-M5 JN019027.1 97% 1 SB-141 Pedobacter sp. 9-15 HM151618.1 96%

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1 SB-142 Pseudomonas aeruginosa AB680503.1 95% 1 SB-143 Mitsuaria sp. RV4 JQ433927.1 91% 1 SB-144 Pantoea agglomerans strain GQ374474.1 93% GS2 1 SB-145 Delftia acidovorans strain JX155410.1 98% IAC/BECa-020 1 SB-146 Pseudomonas sp. INBio2893C HM771055.1 91% 1 SB-147 Rhodothalassium salexigens FR682008.1 90% 1 SB-148 Pedobacter heparinus AB680215.1 96% 1 SB-149 Enterobacter cloacae strain HM030748.1 99% M-5 1 SB-150 Desulfobulbus sp. AF132865.1 94%

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Table S3-8. The dereplication hits of active archaeal 16S ribosomal cDNA based on 98% similarity No. of Dereplicated representative Best hit Accession Identity reads No. 206 SA-1 Candidatus EU281334.1 84% Nitrososphaera gargensis clone RHGA41c 172 SA-2 anaerobic AJ244285.1 94% methanogenic archaeon ET1-9 168 SA-3 Vulcanisaeta sp. HM594677.1 99% CBA1501 116 SA-4 Candidatus CP002408.1 86% Nitrososphaera gargensis 82 SA-5 Thermoproteus CP002590.1 98% uzoniensis 768-20 49 SA-6 Vulcanisaeta AB063641.1 87% distributa 36 SA-7 Aeropyrum pernix AB263905.1 86% 33 SA-8 Thermogladius CP003531.1 85% cellulolyticus 1633 24 SA-9 Sulfolobus sp. JP3 AY907890.1 95% 13 SA-10 anaerobic AJ244284.1 95% methanogenic archaeon ET1-8 9 SA-11 anaerobic AJ244285.1 93% methanogenic archaeon ET1-9 8 SA-12 Candidatus EU239960.1 99% Nitrosocaldus yellowstonii strain HL72 6 SA-13 Desulfurococcus AB661712.1 86%

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amylolyticus 4 SA-14 Methanobacterium sp. CP002551.1 99% AL-21 3 SA-15 Candidatus CP002408.1 85% Nitrososphaera gargensis Ga9.2 3 SA-16 Thermogymnomonas NR_041513.1 87% acidicola strain JCM 13583 3 SA-17 Vulcanisaeta sp. HM594677.1 85% CBA1501 1 SA-18 Candidatus EU281334.1 83% Nitrososphaera gargensis clone RHGA41c 1 SA-19 Vulcanisaeta sp. HM594677.1 97% CBA1501 1 SA-20 Vulcanisaeta sp. HM594677.1 90% CBA1501 1 SA-21 Methanocella AM114193.2 92% arvoryzae MRE50 1 SA-22 Vulcanisaeta sp. HM594677.1 92% CBA1501 1 SA-23 Pyrobaculum sp. D11 AJ630373.1 99% 1 SA-24 Candidatus CP002408.1 85% Nitrososphaera gargensis Ga9.2 1 SA-25 Vulcanisaeta AB063641.1 86% distributa

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Fig. S3-1 The proportions of (A) taxonomic and (B) functional classifications of the total metagenomic reads under E values ≤ 10 -5, 10 -10 , 10 -15, and 10 -20 at phylum (except

Eukaryota) and subsystem levels.

207

208

Fig. S3-2. Neighbor-joining phylogenetic tree of the bacterial representative sequences from the LH 16S ribosomal cDNA pyrosequencing dataset with the number of representing reads indicated in parentheses. Bootstrap values ≧ 60% of 1000 replicates are indicated at the nodes.

209

FIG S3-3. Neighbor-joining phylogenetic tree of the archaeal representative sequences from the LH 16S ribosomal cDNA pyrosequencing dataset with the number of representing reads indicated in parentheses. Bootstrap values ≧ 60% of 1000 replicates are indicated at the nodes.

210

Fig. S3-4. The active bacterial composition of LH spring sediments based on 16S ribosomal cDNA pyrosequencing analyses.

Fig. S3-5. The active archaeal composition of LH spring sediments based on 16S ribosomal cDNA pyrosequencing analyses.

211

SB-1 SB-84 47 SB-7 17 SB-25 95 SB-45 SB-81

86 99 SB-46 99 SB-118 HQ675558.1|_Verrucomicrobia_bacterium_SCGC_AAA240-C14_from_dark_ocean 50 76 EU050946.1|_Uncultured_bacterium_clone_SS1_B_07_50_from_Arctic_sediment 55 AJ633938.1|_Uncultured_Gram-positive_clone_50ANG1_epibiotic_bacteria_of_nidamentalglands_of_squids

25 gHQ675471.1|_Verrucomicrobia_bacterium_SCGC_AAA007-J17_from_dark_ocean

0.1 Fig. S3-6. Neighbor-joining phylogenetic tree of the verrucomicrobial representative sequences (phylotypes with the initial of SB) from the LH 16S ribosomal cDNA pyrosequencing dataset based on 437 bp. EF591088 is a sequence of methanotrophic Verrucomicrobia and its clustering sequence (HQ625077.1) originates from a clone of our previous study on LH channel sediment.

212