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.
©2013
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
2
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
Challenges to microbial life in Polar saline water bodies .......................... 11
1.3
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
Applications and astrobiology aspects of the study ................................... 18
1.4
1.4.1
Potential applications of microorganisms from cold saline
environments ..................................................................................................... 18
1.4.2
1.5
Astrobiological aspects ..................................................................... 20
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 CO2 and CH4 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 CO2 and CH4 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 amoA and hcd gene cloning and sequencing and analyses ..... 109
4.2.6 qPCR of Thaumarchaeal 16S/amoA/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
v
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 H2S/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 (amoA 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 amoA 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 amoA 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 H2S/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.
ix
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 amoA, 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.
x
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
xi
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
amoA
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
hcd
Dissolved oxygen content
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
4-hydroxybutyryl-CoA dehydratase gene
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
Alphaproteobacteria; 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
CaCl2 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 34S
(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
δ13CCH4 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 14C-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
inorganic carbon were examined using S2O32- comsuption and
14-
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 (Aw) 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, Aw = 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
13
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
16
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
amoA, hcd and Thaumarchaeal specific 16S rDNA sequences using qPCR;
and to compare the sequences of amoA 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.
23
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 Lay1, Nadia C. S. Mykytczuk1, Thomas D. Niederberger2, Christine
Martineau1,3, Charles W. Greer3 and Lyle G. Whyte1
1
Department of Natural Resource Sciences, McGill University, Canada
2
College of Marine and Earth Studies, University of Delaware, U.S.A.
3
Biotechnology 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 CO2 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
ammonia-oxidizing Thaumarchaeota.
phylotypes
14
most
closely
related
to
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.
25
2.1 Introduction
Cryoenvironments
are
defined
as
permanently frozen
or
subzero
environments including permafrost, glaciers, ice sheets, multi-year sea ice, highelevation 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 CH4 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 30th, 2008; sediment sample
C3 was collected on May 4th, 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
29
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 CO2 and CH4 concentrations and flux measurements
To determine in situ CO2 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 CO2 and
CH4 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. CO2 and
CH4 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 CO2 and CH4 were determined
on a Picarro CRDS (Picarro, California). In order to measure both CO2 and CH4
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
31
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 CO2 and CH4 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 H2O2 were used for the catalyzed reporter deposition for 15 minutes
32
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 Na4P2O7．10H2O, 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 (DifcoTM R2A Agar, Beckton, Dickenson Co., Sparks, MD, USA).
33
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 H2O 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 MgCl2, 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 DH5α cells (Invitrogen,
Carlsbad, CA, USA). Clone screening was carried out using amplified ribosomal
35
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
36
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
mCi ml-1 (100000 disintegrations per min) of [1-14C] 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. CO2 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 CO2 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
37
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
38
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 CO2 and CH4 concentrations and flux measurements
Results for the sediment CO2 and CH4 concentrations and fluxes from the
LH channel and spring are presented in Table 2-3. CH4 concentrations were an
order of magnitude greater in the sediments from the outlet than in the sediments
from the channel (102 v.s. 10 nmol/g), while CO2 concentrations were found to be
similar at both sites (Table 2-3). Estimated CH4 and CO2 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
constant flux rate method CH4 flux was measured as 33.4 mg/m2/day and 15,052
mg/m2/day for CO2 from the LH channel. By using a LiCor-8100, in situ CO2 flux
in the channel was determined at different locations both during spring and
summer and were found to range between 152 mg/m2/day to 38,244 mg/m2/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 CO2 flux rates were measured in spring and summer
39
conditions while sediment temperatures varied from -16°C to 14 °C. Most CO2
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 × 107 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 × 107 cells/g sediment while the mean abundance of archaea was 3.99 ± 0.44
× 106 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 × 105 CFU/g sediment on
R2A media with 7% NaCl and 2.4 ± 0.60 × 103 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 × 103 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 37oC 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
41
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 14C-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
44
(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 O2 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 CH4 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.
49
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 CO2 and CH4 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 CO2 and CH4 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.
52
Table 2-1. Physical and geochemical characteristics for Lost Hammer (LH)
Spring outlet and channel determined during both Arctic winter and summer
LH outleta
Channelb
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
22 to 26
22 to 26
Salinity (%)
Total viable cell count on 7% NaCl R2A media
N.D.
c
1.25±0.59×105
(CFU/ g sediment)
N.D.
2.40±0.60×103
Bacterial cells (CARD-FISH) (cells/ g sediment) d
3.61±0.11×105
4.51±0.65×107
Archaeal cells (CARD-FISH) (cells/ g sediment)
1.63±0.11×104
3.99±0.44×106
Total viable cell count on 12% NaCl R2A media
(CFU/ g sediment)
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
d
N.D., not determined.
Data for CARD-FISH from the LH outlet were converted from the percentage of total
DAPI-stained cells.
53
Table 2-2. Carbon and nitrogen analyses for LH Spring outlet and channel
LH outlet
Channel
0.48a
0.92 to 1.08
Organic Carbon (%)
a
0.45
0.77 to 0.93
Total Nitrogen (%)
N.D.c
0.02 to 0.08
0.13
0.04 to 0.17
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
Total Carbon (%)
Water dissolved Nitrite/Nitrate (mg/ L)
b
Water dissolved Ammonia (mg/ L)
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 Concentrationa
Flux
CH4 (nmol/g)
CO2 (µmol/g)
CH4
CO2
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/m2/dayc
a
Values are means of triplicates; standard error of the mean is presented.
b
Values are an estimate of the CH4 and CO2 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/m2/s and 2.21µmol/m2/s.
54
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)
-5 to
Planococcus sp.
Axel Heiberg
98-99%
Planococcaceae
37
NP 19 (EU196338)
Island, Perennial
(°C )g
7a
0 to 20
(Family, 100%)
Spring
4b
7 to 20
-5 to
Marinobacter sp.
Antarctic Sandy
25
ZS1-16 (FJ889664)
intertidal
97-100%
Marinobacter
(Genus, 100%)
sediments
2c
5
d
1
e
0 to 12
0 to 20
-5 to
Psychrobacter sp., E59
Antarctic sea
25
(DQ667083)
water
5 to 37
Nesterenkonia sp. 35/46,
Antarctica soil
98-99%
(Genus, 100%)
98-100%
(AY571802)
0 to 12
Psychrobacter
Nesterenkonia
(Genus, 100%)
-5 to
Planomicrobium
Mud volcanoes in
37
psychrophilum strain
Xinjian
99%
Planococcaceae
(Family, 100%)
4-5-26 (GQ505362)
1
f
0 to 7
5 to 25
Fulvimarina sp.
Axel Heiberg
NP 28 (EU196328)
Island, Perennial
100%
Aurantimonadace
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
e
This 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).
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
Outleta
Channel
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.
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
(1/D)
a
Data represent the statistics and indices obtained from Niederberger et al., 2010. N.D.: not
determined.
56
(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.
57
(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%.
58
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.
59
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.
60
Fig. 2-5. Mineralization assays of [1-14C] acetate in LH channel sediment
microcosms at different temperatures. Each point represents the mean cumulative
mineralization (%
14
CO2 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.
61
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 Lay1, Nadia C. S. Mykytczuk1, Étienne Yergeau2, Guillaume
Lamarche-Gagnon1, Charles W. Greer2 and Lyle G. Whyte1*
1
Department of Natural Resource Sciences, McGill University, Canada
2
Biotechnology 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
H2S/sulfides (up to 25 ppm), ammonia (≈381 µM), and methane (11.1 g d-1). 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
63
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.
67
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 ddH2O 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
69
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
71
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 iScriptTM 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)
(28F:5’GAGTTTGATCNTGGCTCAG’3,
system
with
bacterial
519R:5’
GTNTTACNGCGGCKGCTG’3) (Handl et al., 2011) and archaeal (ARCH571F:
72
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.
73
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.
74
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
Desulfovibrionales,
and
sulfate
reducers,
such
Desulfobacteriales,
as
were
Desulfuromonadales,
detected
within
the
Deltaproteobacteria and Betaproteobacteria, and whose metabolic activity would
75
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
78
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).
79
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.,
80
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
81
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
82
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
85
(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
87
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.
92
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%
93
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
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
Bacteria
94
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
95
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).
97
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.
98
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, amoA and hcd, in
both channel sediment and the tundra soil. The partial amoA 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 (CO2 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, amoA 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 amoA 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 14C-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 amoA 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 amoA, hcd and Thaumarchaeal specific 16S
rDNA sequences using qPCR, and; 3) to compare the sequences of amoA 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 CO2 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
105
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 amoA, 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
106
(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
(ddH2O) 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 ddH2O 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 iScriptTM 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)
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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
108
al., 2010) based on the GreenGene core dataset.
4.2.5 Archaeal amoA and hcd gene cloning and sequencing and analyses
To clone the partial archaeal ammonia oxidizing gene subunit A (amoA), 2
µL of environmental DNA extracted from the LH channel were applied to
Polymerase chain reaction (PCR) using ImmolaseTM DNA Polymerase (Bioline,
London,
UK).
The
specific
(5’ATGGTCTGGCTWAGACG’3)
primer
and
pair,
CrenamoA23f
CrenamoA616r
(5’GCCATCCATCTGTATGTCCA’3) (Tourna et al., 2008), was applied to
amplify the partial amoA 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
109
final extension of 5 min at 72°C (Zhang et al., 2010). The final PCR products of
both partial amoA 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 DH5α (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/amoA/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 iQTM 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, amoA 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 amoA 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 ddH2O
containing no detectable DNA. We employed previously cloned linear fragments
of amoA, 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 amoA
and hcd genes were 84.9% and 90.2%, respectively. The R2 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 CO2 flux was similar in the channel
sediments in both seasons. On the contrary, the CO2 flux was much less in the
tundra than in the channel. The CO2 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
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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
115
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.
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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
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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 (amoA)
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 amoA revealed that the archaeal amoA grouped
differently with bacterial amoA genes (Figure 4-4a). Additionally, the LH amoA
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 amoA
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 amoA 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 amoA 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 (105). The same trend in copy numbers was observed for the hcd genes,
albeit one magnitude (106) higher than the amoA 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
detect
14
C-CO2 recovery from
14
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.
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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
124
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
128
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 amoA 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 amoA genes we detected at the LH
channel and the adjacent tundra (~ 105) 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 amoA 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 (~106) 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 amoA 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 amoA 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 amoA 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 amoA 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
amoA genes. Although the tree of hcd gene sequences was not identical to the
trees of 16S rRNA or amoA 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 amoA 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, amoA 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, amoA 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 CO2 detection on site.
135
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
4.1
-12.9
16.7
N.D.
Air temperatures °C
15.6
-16.9
15.4
-16.9
Salinity (%)
~2(?)
25.8
N.D.
N.D.
0.38±0.02
0.59±0.12
0.09±0.01
0.05±0.05
Water/soil
temperatures °C
2
CO2 flux µmol/m /s
N.D.: Not determined
136
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
set
Derep
Chao1
Shannon
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
137
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).
Archaea
Bacteria
Extract
Sampling season
Sampling location
F-Value
1.5995
3.0249
0.40359
P-Value
0.2371
0.0259
0.775
F-Value
1.1098
1.841
1.2572
P-Value
0.3152
0.031
0.2531
138
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
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
(B)
139
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.
129
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.
130
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.
131
Fig. 4-4. Maximum-likelihood trees constructed by partial putative amino acid sequences
of (A) amoA (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), amoA (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
H2S/sulfides (up to 25 ppm), ammonia (≈381 µM), and methane (11.1 g d-1). 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 (CO2 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,
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. Comparing these 16S rDNA libraries using
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 (amoA 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 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. Copy
number of Thaumarchaeal amoA 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
microorganisms in the sediments by detecting 14C-CO2. 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, amoA 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
141
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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 librariesa
Bacterial
C1
C2
LH sourceb
C3
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
80
100
171
100
236
100
100
Total
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 librariesa
Archaeal Phylum/Class
C1
C2
LH source b
C3
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
80
100
24
100
80
100
100
Total
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-05 R
P-value
1.00E-10
1.00E-15
1.00E-20
1
0.987563
0.73459
< 2.2e-16
< 2.2e-16
1.93E-05
0.999999
0.995174
0.908828
< 2.2e-16
< 2.2e-16
2.27E-11
Function
1.00E-05 R
P-value
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
Nostoc
8 Cyanobacteria
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
Chromobacterium
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
Herminiimonas
7.9 Betaproteobacteria
Thiobacillus
6.3 Betaproteobacteria
Congregibacter
5.6 Gammaproteobacteria
Janthinobacterium
5.3 Betaproteobacteria
Chlorobaculum
4.3 Bacteroidetes
Polaromonas
3.6 Betaproteobacteria
Bradyrhizobium
3.3 Alphaproteobacteria
Ralstonia
3.0 Betaproteobacteria
Vesicomyosocius
2.6 Gammaproteobacteria
Pelodictyon
2.0 Bacteroidetes
Thioalkalivibrio
1.3 Gammaproteobacteria
Nitrobacter
1.3 Alphaproteobacteria
Acidiphilium
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
Function Protein
names
Oxidative Catalase
Unique Total hits
Phyla
hits
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%),
Firmicutes (28.2%)
hydroperoxide
resistant
protein
194
Table S3-5. Representatives of genes/proteins related to osmotic stress in LH
metagenome
Function
Protein names
Unique Total
hits
Synthesis of
Cyclic beta-1,2-glucan 34
Phyla
hits
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
protein C
Choline and Betaine
Bacteroidetes (95.3%),
Proteobacteria (4.7%)
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
dehydrogenase
2
7
Fungi (85.7%),
Proteobacteria (14.3%)
195
Table S3-6. Representatives of genes/proteins related to cold adaptation in LH
metagenome
Functions
DNA replication
Protein or subsystem
Unique Total hits Phyla
names
hits
gyrA
96
58535
(DNA Gyrase A)
Cyanobacteria (55.6%),
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
initiator Protein)
Cyanobacteria (33.7%),
Bacteroidetes (63.4%),
Actinobacteria (1.0%),
Proteobacteria (1.5%),
Deinococcus-Thermus
(0.4%)
Unsaturated fatty 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
196
55118
Cyanobacteria (57.0%),
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
protein A
Bacteroidetes (49.8%)
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
reads
representative
103
SB-1
Best hit
Accession
Identity
No.
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%
Microlunatus panaciterrae
NR_041517.
94%
strain Gsoil 954
1
Pseudomonas aeruginosa
JF899310.2
99%
SCGC AAA240-C14
5
5
SB-26
SB-27
strain PM389
5
SB-28
Desulfobulbus sp.
AF132865.1
99%
4
SB-29
Staphylococcus aureus subsp.
HE579073.1
99%
NR_041517.
93%
aureus ST228
4
SB-30
Microlunatus panaciterrae
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
Microlunatus panaciterrae
NR_041517.
strain Gsoil 954
1
Microlunatus panaciterrae
NR_041517.
strain Gsoil 954
1
Verrucomicrobia bacterium
HQ675558.1
4
4
3
SB-34
SB-35
SB-36
93%
94%
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%
HQ675558.1
91%
aureus ST228
3
SB-44
Verrucomicrobia bacterium
199
SCGC AAA240-C14
3
SB-45
Verrucomicrobia bacterium
HQ675558.1
97%
HQ675558.1
95%
SCGC AAA240-C14
3
SB-46
Verrucomicrobia bacterium
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%
HE579073.1
97%
JQ900543.1
92%
SL014B-79A
3
SB-51
Staphylococcus aureus subsp.
aureus ST228
2
SB-52
Pseudomonas aeruginosa
strain N83
2
SB-53
Pseudomonas aeruginosa
AB037548.1
96%
2
SB-54
Pseudomonas aeruginosa
DQ666628.1
94%
HE579073.1
99%
Stenotrophomonas maltophilia GU815943.1
96%
strain RsB-29
2
SB-55
Staphylococcus aureus subsp.
aureus ST228
2
SB-56
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
Staphylococcus aureus subsp.
HE579073.1
99%
2
SB-61
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%
HE579073.1
96%
Microlunatus panaciterrae
NR_041517.
91%
strain Gsoil 954
1
aureus ST228
2
SB-76
Staphylococcus aureus subsp.
aureus ST228
2
SB-77
2
SB-78
Acidovorax sp. CNE 29
FR749857.1
97%
2
SB-79
Desulfobulbus sp.
AF132865.1
97%
2
SB-80
Stenotrophomonas sp. NOE8
2
SB-81
Verrucomicrobia bacterium
94%
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%
AB269802.2
90%
SCGC AAA240-C14
1
SB-85
Sphingomonadaceae
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%
HQ018741.1
98%
strain NGKCTS
1
SB-95
Pseudomonas aeruginosa
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%
JF899310.2
97%
AB269802.2
91%
AB269802.2
88%
JQ659749.1
95%
bacterium KF016
1
SB-106
Pseudomonas aeruginosa
strain PM389
1
SB-107
Sphingomonadaceae
bacterium KF016
1
SB-108
Sphingomonadaceae
bacterium KF016
1
SB-109
Enterobacter oryzae strain
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%
HQ675558.1
92%
Microlunatus panaciterrae
NR_041517.
92%
strain Gsoil 954
1
SL014B-79A
1
SB-118
Verrucomicrobia bacterium
SCGC AAA240-C14
1
SB-119
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%
HQ675558.1
96%
strain TERIPS9002
1
SB-132
Verrucomicrobia bacterium
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
Tessaracoccus sp.
GU111568.2
92%
1
SB-136
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%
203
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%
JX155410.1
98%
GS2
1
SB-145
Delftia acidovorans strain
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%
AF132865.1
94%
M-5
1
SB-150
Desulfobulbus sp.
204
Table S3-8. The dereplication hits of active archaeal 16S ribosomal cDNA based on 98%
similarity
No. of
Dereplicated representative
Best hit
reads
206
Accession
Identity
No.
SA-1
Candidatus
EU281334.1
84%
AJ244285.1
94%
HM594677.1
99%
CP002408.1
86%
CP002590.1
98%
AB063641.1
87%
Nitrososphaera
gargensis clone
RHGA41c
172
SA-2
anaerobic
methanogenic
archaeon ET1-9
168
SA-3
Vulcanisaeta sp.
CBA1501
116
SA-4
Candidatus
Nitrososphaera
gargensis
82
SA-5
Thermoproteus
uzoniensis 768-20
49
SA-6
Vulcanisaeta
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%
AJ244285.1
93%
EU239960.1
99%
AB661712.1
86%
methanogenic
archaeon ET1-8
9
SA-11
anaerobic
methanogenic
archaeon ET1-9
8
SA-12
Candidatus
Nitrosocaldus
yellowstonii strain
HL72
6
SA-13
Desulfurococcus
205
amylolyticus
4
SA-14
Methanobacterium sp. CP002551.1
99%
AL-21
3
SA-15
Candidatus
CP002408.1
85%
NR_041513.1
87%
HM594677.1
85%
EU281334.1
83%
HM594677.1
97%
HM594677.1
90%
AM114193.2
92%
HM594677.1
92%
Nitrososphaera
gargensis Ga9.2
3
SA-16
Thermogymnomonas
acidicola strain JCM
13583
3
SA-17
Vulcanisaeta sp.
CBA1501
1
SA-18
Candidatus
Nitrososphaera
gargensis clone
RHGA41c
1
SA-19
Vulcanisaeta sp.
CBA1501
1
SA-20
Vulcanisaeta sp.
CBA1501
1
SA-21
Methanocella
arvoryzae MRE50
1
SA-22
Vulcanisaeta sp.
CBA1501
1
SA-23
Pyrobaculum sp. D11
AJ630373.1
99%
1
SA-24
Candidatus
CP002408.1
85%
AB063641.1
86%
Nitrososphaera
gargensis Ga9.2
1
SA-25
Vulcanisaeta
distributa
206
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
SB-46
99
99
SB-118
HQ675558.1|_Verrucomicrobia_bacterium_SCGC_AAA240-C14_from_dark_ocean
50
76
55
AJ633938.1|_Uncultured_Gram-positive_clone_50ANG1_epibiotic_bacteria_of_nidamentalglands_of_squids
gHQ675471.1|_Verrucomicrobia_bacterium_SCGC_AAA007-J17_from_dark_ocean
25
Coraliomargarita_akajimensis_DSM_45221_strain_DSM_45221
99
26
EU050946.1|_Uncultured_bacterium_clone_SS1_B_07_50_from_Arctic_sediment
93
AB073978.1|_Fucophilus_fucoidanolyticus
X99392.1|_Opitutus_sp._VeSm13
AB372851.1|_Cerasicoccus_frondis_YM31-067_from_sea
35
EF591088.1|_Methylacidiphilum_fumariolicum_strain_SolV
75
46
HQ625077.1|_Uncultured_Verrucomicrobia_from_LH_channel
SB-117
SB-99
gi|99160149|gb|DQ521126.1|_Uncultured_bacterium_clone_G73_from_Gypsum_Hill_Springs
AJ309733.1|_Aquifex_aeolicus
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