picoplankton community analysis along the atlantic meridional … · 2016-11-04 · i picoplankton...
TRANSCRIPT
i
Picoplankton Community Analysis along the Atlantic
Meridional Transect AMT22
Greta Reintjes
Master Thesis
Bremen 2013
Max Planck International Research School for Marine Microbiology
Max Planck Institute for Marine Microbiology
&
The University of Bremen
ii
Master Thesis proposed by Greta Reintjes
Bremen, March 2013
1. Examiner: Prof.Dr.Rudolf Amann
2. Examiner: PD. Dr.Bernhard Fuchs
3
Affidavit
Statement
Hiermit versichere ich dass ich diese Arbeit selbständig verfasst und keine anderen als die
angegebenen Quellen und Hilfsmittel verwendet habe.
I hereby declare that the following master thesis has been written by myself and that no
sources have been used in the preparation of this thesis other than those referenced in the
thesis itself. All figures which are not self made but gathered from external sources are
marked as such.
Bremen, March 2013 ……………………….......…
Greta Reintjes
4
Summary
The purpose of this study was to analyse the marine picoplankton community along the
Atlantic Meridional Transect AMT22. The diversity and abundance of dominant taxonomic
groups, potential biogeographical patterns in their distribution and their relation to
environmental parameters were investigated. We applied catalyzed reporter deposition-
fluorescence in situ hybridisation (CARD-FISH) and massive parallel tag sequencing of the
16S rRNA gene.
Firstly, the bacterioplanktonic diversity in distinct biogeographical provinces (Longhurst et al.,
2007) was analysed using tag sequencing. A high microbial diversity was found in all
provinces. Eight phyla represented 97% of the relative sequence abundance of all provinces.
The Proteobacteria, Cyanobacteria and Bacteroidetes have an average abundance of 39.5 ±
11.5, 28.5 ± 10.5% and 12 ± 7% respectively in all provinces. Other phyla Planctomycetes,
Chloroflexi, Deferribacteres and Planctomycetes made up < 10% of the total sequence
abundance. Of the Proteobacteria the alphaproteobacterial clade SAR11 made up nearly half
of the abundance with 41.5 ± 15.5%. The Gammaproteobacteria were the second most
abundance proteobacterial class with 39 ± 14%. A large portion of this (15 ± 5%) was made
up of the SAR86 clade.
Secondly the abundance and distribution of the dominate taxon along an Atlantic transect
was analysed. The absolute picoplankton abundance (cells ml-1) changed with latitude and
with depth and was positively correlated with high chlorophyll a concentrations (r2 0.76).
Bacteria dominate the surface waters of all provinces with 72 ± 16%. Archaea had an
absolute abundance of 6.1E+04 ± 2.1E+04 cells ml-1 throughout the water column. However,
their relative abundance showed a significant change from 1% at the surface to 40% at 150
m depth.
This study found that the taxonomic groups which dominate the sequence data could also be
found and enumerated using CARD-FISH. These groups made up a significant fraction of the
total cellular abundance of the bacterioplankton community. Only a few groups (SAR11,
SAR86 and Bacteroidetes) could be found at all latitudes and depths and although they were
ubiquitous in the sequencing data, their relative abundances did vary.
The diversity analysis showed that at different taxonomic level biogeographic distribution
patterns were present. They depended on distance (km) or environmental variation between
the provinces. The combined analysis of the diversity and abundance of microorganisms
based on biogeographic patterns resulted in a more accurate interpretation of the
biogeography of microorganisms in the Atlantic Ocean. Understanding the biogeography of
microorganisms is a key step in understanding the ecosystem function of specific microbial
assemblages.
5
Contents
1. Introduction
2. Methods
2.1. Sampling
2.2. Classification of Biogeographical Provinces
2.3. Catalyzed Reporter Deposition-Fluorescence in situ Hybridisation (CARD-FISH) for
Planktonic Samples on Membrane Filters According to Pernthaler et al.(2004)
2.4. Statistical Analysis of Abundances
2.5. Sampling for Massive Parallel Tag Sequencing
2.6. DNA Extraction
2.7. Fusion Primer Design
2.7.1. Theoretical Background to PCR and Fusion Primers
2.7.2. Procedure
2.8. Roche 454 Pyro- Sequencing
2.9. Analysis of 454 Sequences
3. Results and Discussion
3.1. Classification of Ocean Biogeographic Provinces along the AMT22
3.2. Sequencing Statistics and Diversity Analysis
3.3. Relative Abundance of Phyla
3.4. Relative Abundance of Dominant Phyla
3.5. Absolute and Relative Abundances of Picoplankton Along the AMT22
3.6. Absolute and Relative Abundances of the Dominant Bacterial Taxon Along the
AMT22
3.6.1. Cyanobacteria
3.6.2. SAR11 and SAR86
3.6.3. Bacteroidetes
3.7. Other Groups of the Bacterioplankton Community
4. Conclusion
5. Acknowledgments
6. References
7. Appendices
6
Abbreviations
AAIW Antarctic Intermediate Water
AMT Atlantic Meridional Transect
BLAST Basic local alignment search tool
bp Basepair
CARD-FISH Catalysed Reporter Deposition-Fluorescence in situ
Hybridisation
CCD charge coupled device
CTD Conductivity/ Temperature/ Depth Profiler
DAPI 4',6-diamidino-2-phenylindole
DCM Deep Chlorophyll Maximum
DNA Deoxyribonucleic acid
dNTPs deoxyribonucleotide triphosphates
EB Elution buffer
EDTA Ethylene diamine tetra acetic acid
emPCR Emulsion Polymerase-Chain-Reaction
FISH Fluorescence in situ Hybridisation
HCL Hydrochloric Acid
NADR North Atlantic Drift
NAST North Atlantic Subtropical Gyre
NATR North Atlantic Tropical Gyre
NGS Next generation sequencing
nt Nucleotide
ODV Ocean Data View
OTU Operation Taxonomic Unit
PAST Palaeontological Statistics
PBS Phosphate Buffer Solution
PCR Polymerase-Chain-Reaction
PE Ethanol Wash Buffer
PTP Picotiterplate
QG Solubilisation and Binding Buffer
rDNA Ribosomal deoxyribonucleic acid
RNA Ribonucleic acid
rRNA Ribosomal ribonucleic acid
RRV Royal Research Vessel
7
RT Room temperature
SATL South Atlantic Gyre
SDS Sodium dodecyl sulphate
SSTC South Atlantic Subtropical Gyre
SSU Small ribosomal subunit
TAE Tris-acetate-EDTA
TAQ Thermus aquaticus
Tris HCL Tris (hydroxymethyl) aminomethane Hydrochloric Acid
V3, V4 Hyper variable Region
WTRA Western Tropical Gyre
8
Introduction
The Atlantic Meridional Transect (AMT) is a multidisciplinary ocean observatory program
started in 1995 (Robinson et al., 2006). Its aim is to understand the biological, chemical and
physical parameters of the Atlantic Ocean (www.amt-uk.org). The Atlantic Ocean is the
world’s second largest ocean and covers approximately 20 percent of the earth’s surface. It
is highly variable in its physical oceanography and biogeography. It is not homogeneous but
has varying nutrient concentrations partly due to seasonal mixing and heat stratification of
water masses. This heterogeneity causes variations in its primary production which is
dependent on solar irradiance and nutrient availability. Correspondingly it has areas of high
primary production in high temperate and equatorial regions, where seasonal mixing occurs,
and areas of low primary production in gyral regions where there is a high amount of thermal
stratification (Longhurst, 2007).
One of the major findings of the AMT was that the Atlantic Ocean can be sectioned into
provinces with characteristic biotic (phytoplankton and zooplankton) and abiotic features
(Longhurst et al., 1995). Longhurst et al., (1995) defined the distinct provinces based mainly
on the biodiversity and abundance of primary producers (phytoplankton). However marine
microbial communities are known to make up a considerable amount of biomass and
contribute significantly to the biogeochemistry of the oceans (Azam & Malfatti, 2007). They
are key organisms in the degradation of organic matter derived from primary production but
can also make up a significant fraction of the primary production (Wetz et al., 2008;
Zwirglmaier et al., 2008).
Marine microorganisms were long believed to have a ubiquitous distribution due to their
large population size and because normal geographical isolation is thought to have little
effect on their dispersal rates. This is especially true for the marine microorganisms which
find themselves in an environment where there are few geographical barriers and a
continuous circulation (Martiny et al., 2006). Previous studies have shown that the
composition of the Atlantic Ocean surface water bacterioplankton differs with latitude
(Schattenhofer et al., 2009). This has also been shown for other environments and a large
body of research now supports the idea that marine microorganisms are not randomly
distributed but exhibit biogeographical patterns (Fuhrman et al., 2008; Green & Bohannan,
2006; Lomolino et al., 2006; Martiny et al., 2006).
Currently there are three main hypotheses attempting to explain the biogeography of
microorganisms. The first is based on the Baas-Becking and Beijerinck hypothesis
“everything is everywhere: but the environment selects”. In this sense, if the environment
presented a suitable ecological niche then any microorganisms could exist there (O'Malley,
9
2007). This indicates that the biogeography of microorganisms reflects the influence of
environmental factors and variations. The second is based on spatial and temporal
variations due to historic events. This hypothesis explains biogeography based on speciation
events due to past environmental conditions or influences. Additionally the hypothesis
incorporates historic geographical isolation events. The third hypothesis is a mix of the two
previous. It bases the biogeography of microorganisms on environmental and historical
factors (Martiny et al., 2006). However, the attempts to define universal laws for the
distribution and biogeography of microorganisms are still far from finished.
This is due, in part, to the lack of understand in the factors that control the distribution and
diversity of microbes. The complexity of the microbial environment can be represented by an
n-dimensional hyperspace. With n being the number of possible environmental parameters
effecting a microorganism (Whittaker et al., 1973). However the classification of distinct
biogeographic provinces (Longhurst, 2007) was a key step in defining the environment to
which marine microorganisms are exposed and enable a more detailed analysis of the
biogeography of microorganisms in the Atlantic Ocean.
Another difficulty in the definition of the biogeography of microorganisms is the problem
associated with the microbial species definition. A large fraction of microbes are resistant to
cultivation and due to their lack of distinguishing features and small size are hard to
differentiate. Microbiologists alternatively use molecular tool to distinguish a microbial
species. The development of cultivation independent techniques such as the polymerase
chain reaction (PCR) and DNA sequencing allows for the identification of microorganisms
directly from the environment without the need for isolation. The main method associated
with the definition of a microbial species is the use of universal genes for a phylogenetic
based taxonomic classification. The 16S ribosomal DNA (rDNA) gene is currently the most
used gene for phylogenetic based classification. The 16S rDNA gene has a ubiquitous
presence in prokaryotic and eukaryotic cells, has a relative slow evolution, and functional
homology (Nübel et al., 1997) making it ideal for phylogenetic comparisons. The
comparative analysis of the 16S rDNA gene is a well established approach for the
phylogenetic classification of Bacteria and Archaea, with extensive databases available
(Huse et al., 2008; Tamaki et al., 2011; Woese, 1987; Zuckerkandl & Pauling, 1965).
The DNA based phylogenetic classification has been further enhanced by the development
of next generation sequencing techniques (NGS) (Frias-Lopez et al., 2008; Huse et al.,
2008). The combination of PCR with NGS yields microbial diversity data up to three orders
of magnitudes greater than previous methods (Ghiglione et al., 2012; Kirchman et al., 2010).
The massive parallel sequencing of the 16s rDNA to characterise a microbial population was
initiated by Sogin et al., (2006). They reported bacterial communities with up to two orders of
magnitude higher complexity than ever reported. Additionally the barcoding of samples
10
enables the simultaneous sequencing of several samples in single sequencing run (Tamaki
et al., 2011) making it cost effective.
The sequencing of the 16s rRNA gene has additionally allowed for the identification of
signature sequences which are unique for different taxonomic groups of microorganisms.
These signature sequences can be targeted by complimentary oligonucleotides (probes) for
identification purposes. Fluorescence in situ hybridisation (FISH) uses probes which are
labelled with a fluorochrome allowing for, with the use of fluorescent microscopy, a
phylogenetic staining (Amann et al., 2001; DeLong et al., 1989). Using this method, specific
bacterial taxonomic groups can be targeted and enumerated in their environment. The
application of this method to analyse the abundance of specific microbial groups has given
new insight into the relative abundance of specific taxonomic groups in different habitats
(Grossart et al., 2005; Morris et al., 2012; Pérez et al., 2005; Thiele et al., 2012).
With the development of new methods our understanding of the diversity and abundance of
marine microorganisms has significantly increased. This is important because they dominate
the marine environment in terms of abundance, diversity and metabolic activates (Azam &
Malfatti, 2007).Additionally they are key mediators of the biogeochemical cycling of elements
in the ocean (Arrigo, 2004; Falkowski et al., 2008).
The application of these molecular techniques has given a more accurate insight into
microbial community diversities which can now be applied to analysis of biogeography.
In this study we applied massive parallel tag sequencing of the 16S rDNA gene and catalytic
reporter deposition-fluorescence in situ hybridisation (CARD-FISH) to analyse the microbial
community composition of the Atlantic Ocean. Additionally we used biotic, environmental
and geographical similarity matrixes to analyse the biogeography of the dominant taxonomic
groups (Martiny et al., 2006).
11
2. Methods
2.1 Sampling
During the Atlantic Meridional Transect AMT22 from 50°N to 50°S, upon the RRV James
Cook (Southampton, UK, to Punta Arenas, Chile, 10 Oct to 24 Nov 2012), seawater samples
were taken using a Sea Bird CTD (Sea Bird Electronics Inc., U.S.A). The CTD was deployed
twice daily between 20 and 400 m water depth at predawn and solar noon intervals. For all
74 sampling stations (Figure 1) three individual depths were sampled (surface; 20 m, DCM;
various and mesopelagic; 150 m). If the DCM was 150 m or deeper, the mesopelagic
sample was taken from > 200 m. From each sample 100 ml of seawater was fixed in 1%
formaldehyde for 1 hr at RT. Subsequently, 20 ml subsamples were filtered (200 mbar) onto
47 mm polycarbonate filters with a 0.2 µm pore size in triplicates. These filters were then
stored at -20°C until further analysis.
12
Figure 1:Map of the 74 sampling points along the AMT22. The individual dot colours indicate the biogeographic provinces in which the sample was taken. Red: North Atlantic Drift (NADR), Orange: North Atlantic Subtropical Gyre (NAST), Yellow: North Atlantic Tropical Gyre (NATR), Green: western Tropical Gyre (WTRA), White: South Atlantic Gyre (SATL), Pink: South Atlantic Subtropical Gyre (SSTC) (Longhurst et al., 1995). The map colours indicate the average chlorophyll a concentrations (mg m
-3) during the AMT22 time period
(www.oceancolor.gstc.nansa.gov). Red circles indicate sites analysis using pyro- sequencing.
13
2.2 Classification of Biogeographic Provinces
The AMT22 passed through six ocean provinces the North Atlantic Drift (NADR), North
Atlantic Subtropical Gyre (NAST), North Atlantic Tropical Gyre (NATR), Western Tropical
Gyre (WTRA), South Atlantic Gyre (SATL), and the South Atlantic Subtropical Gyre (SSTC)
according to(Longhurst, 2007). The ocean biogeographical provinces were identified using
their physical, chemical and biological characteristics. These were measured at every CTD
sampling station from 0 m to 500 m depth. Temperature (°C) was measured using a SEA-
BIRD 3 premium temperature sensor. Dissolved oxygen (ml l-1) was measured using the
Sea-Bird 43 dissolved oxygen sensor and calibrated against Winkler titration measurements
from 9 samples collected from the pre-dawn CTD. Conductivity (S m-1) was measured using
a Sea-Bird4 conductivity sensor. Fluorescence (mg m-3) was measured using a CTG FAST
track Fast Repetition Rate fluorimeter and calibrated against extracted chlorophyll-a
measurements made on seawater samples collected from 9 depths at each station.
Pressure (mbar) was measured using a Digiquartz pressure sensor suspended below the
CTD. Salinity (PSU) was measured using a Guildline Autosal 8400B salinometer and
calibrated against bench salinometer measurements from 4 samples collected from each
cast.
2.3 Catalyzed Reporter Deposition-Fluorescence in situ Hybridisation
(CARD-FISH) for planktonic samples on membrane filters according to
(Pernthaler et al., 2004).
Figure 2: Schematic representation of CARD-FISH method from Arb-Silva (Quast et al., 2013a)
14
All stations and depths were analysed using CARD-FISH (Figure 2). The filters were first
embedded in 0.1 % LE agarose (Biozym Scientific, Germany) and subsequently
permeabilised using lysozyme (10 mg ml-1 in 0.05 M EDTA, pH 8.0; 0.1 M Tris-HCL, pH 8.0).
Next endogenous peroxidases were inactivated by incubating the filters in 0.01 M HCL for 15
min. Then the samples were hybridised with 16s rRNA probes chosen based on previous
literature and the study by (Schattenhofer et al., 2009) (Table 1). Hybridisation was carried
out in humidity chambers at 46 °C for 2.5 hrs. The hybridisation buffer (formamide
concentration varied according to probe used, see Appendix 1) was mixed with a probe
concentration of 8.42 ng µl-1. After hybridisation the filters were washed in pre heated (48 °C)
wash buffer for 10 min at 48 °C and then incubated in 1 X PBS for 15 min at RT. Next the
amplification was carried out using an amplification buffer H2O2 solution at a ratio of 100:1
with 1 mg ml-1 fluorescently labelled tyramide (Appendix 1.). Amplification was carried out at
46 °C for 45 min. After the CARD-FISH process the filters were counter stained with DAPI
and mounted using a citiflour/vector shield (4:1) mounting solution. CARD-FISH and DAPI
staining the cells were visualised and enumerated on a Zeiss Axioskop 2 motplus
fluorescent microscope.
Table 1: List of probes with sequence and specific formamide (FA) concentration applied during this study.
Probe Target organisms Sequence (5′→3′) FA
(%)
Reference
Arch915 Almost all Archaea GTGCTCCCCCGCCAATTCCT 35 (Amann et al.,
1990)
Cren554 Crenarchaeota marine
group I
TTAGGCCCAATAATCMTCCT 0 (Massana et
al., 1997)
Eury806 Euryarchaeota marine
group II
CACAGCGTTTACACCTAG 0 (Teira et al.,
2004)
Eub338 I-III Almost all Bacteria GCWGCCWCCCGTAGGWGT 35 (Amann et al.,
1990)
Non338 Control ACTCCTACGGGAGGCAGC 35 (Wallner et al.,
2005)
Gam42a γ-Subgroup of
Proteobacteria
GCCTTCCCACATCGTTT 35 (Manz et al.,
1992)
Ros537 Roseobacter-clade CAACGCTAACCCCCTCC 35 (Eilers et al.,
2001)
SAR86-1249 SAR86 clade TTAGCGTCCGTCTGTAT 35 (Eilers et al.,
2000)
SAR86-1245-
h3
Helper to SAR86 1245 GGATTRGCACCACCTCGCGGC 35 (Zubkov et al.,
2001)
15
SAR86-1245-
h5
Helper to SAR86 1245 CCATTGTAGCACGTGTGTAGC 35 (Zubkov et al.,
2001)
CF319a most Flavobacteria, some
Bacteroidetes, some
Sphingobacteria
TGGTCCGTGTCTCAGTAC 35 (Manz et al.,
1996)
SAR11 MIX SAR11 clade GGACCTTCTTATTCGGGT 25 (Morris et al.,
2002)
Pro405 Prochlorococcus AGAGGCCTTCGTCCCTCA 40 (West et al.,
2001)
SYN405 Synechococcus AGAGGCCTTCATCCCTCA 40 (West et al.,
2001)
SAR202–
312R
SAR202 clade TGTCTCAGTCCCCCTCTG 40 (Morris et al.,
2002)
SAR324-1412 SAR324 clade GCCCCTGTCAACTCCCAT 35 (Schattenhofer
et al., 2009)
POL740 Polaribacter CCCTCAGCGTCAGTACATACGT 35 (Malmstrom et
al., 2004)
EUK516 Eukarya ACCAGACTTGCCCTCC
0 (Amann et al.,
1990)
2.4 Statistical Analysis of Abundances
Absolute (cell ml-1) and relative cell abundances (% of total DAPI)were enumerated at every
sampling point for all probes. The results were graphed as contours plots using the ocean
data view 4 (ODV) software (www.odv.awi.de). The relative abundance of bacterial groups in
biogeographical provinces were visualised with pie charts using Sigmaplot
(www.sigmaplot.co.uk). Additionally the relative abundance data was correlated to physical
parameters using Spearman Rank Order correlations. This correlation was applied because
it does not define variable as dependent or independent and measure the strength of the
association between all variables. The correlation results are indicated with the correlation
coefficient r, which indicates the strength of the correlation (-1 strong negative correlation to
+1 strong positive correlation).
2.5 Sampling for Massive Parallel Tag Sequencing
Large volumes of seawater (10 - 45 l) were sampled from 20 m depth at every solar noon
station for DNA analysis. The water was sequentially filtrated onto 142 mm polycarbonate
filters with pore sizes of 10 µm, 3 µm and 0.2 µm. These filters were stored at -80 °C until
further analysis. The samples analysed by pyro- sequencing are indicated with red circles in
Figure 1.
16
2.6 DNA Extraction
Microbial DNA was extraction using the MoBio Ultra Clean Soil DNA Extraction Kit (MoBio
Laboratories, Inc., Carlsbad, CA,) as recommended by the manufacturer with the alteration
that instead of soil a fixed size (150mm x 250mm) polycarbonate filter piece was directly
added to the Bead Solution Tubes. The method comprise of an initial chemical lyses of cells
using an SDS based solution and the subsequent mechanical lyses of cells. SDS is a
detergent which aids the breakdown of cell membranes specifically fatty acids and lipids.
The mechanical lysis was done by vortexing the cells in a bead solution. After cell lyses
impurities such as proteins and salts, which would interfere with further analysis, were
removed by precipitation and centrifugation steps. The DNA was then bond to a silica
membrane within a column in the presence of a high salt solution. Further washing to
remove salts was done using > 70% ethanol to avoid any elution of the DNA. Finally the
DNA was eluted using a Tris-HCL buffer, pH 8.0.
2.7 Fusion Primer Design
2.7.1 Theoretical Background to PCR and Fusion Primers
The polymerase chain reaction (PCR) enables the exponential amplification of a targeted
strand of DNA. In the reaction the heat stable polymerase (Taq), derived from the bacterium
Thermus aquaticus, is used to amplify single stranded DNA fragments with a primer, short
complementary oligonucleotide DNA fragment (Innis et al., 1988). In a series of temperature
cycles the DNA is denatured, the primer is annealed and elongation occurs to create a
complimentary DNA strand of the targeted DNA sequence (Schochetman et al., 1988).
Gene specific primers consist of a short oligonucleotide fragment (20 bp length)
complimentary to the targeted genes DNA sequence. In this study “fusion primers” were
used to amplify selected regions of bacterial SSU rDNA. Our fusion primers had a Sfi-A or B
site and a barcode sequence attached to the 5’- prime end of the gene specific primer
(Figure 3). After PCR the Sfi-A or B site is digested using the Sfil enzyme to generate a 3-
base single stranded overhang to which the 454 sequencing adaptors are ligated. This
allows for direct sequencing of the PCR product. The barcode consists of a unique 6
nucleotide sequence which allows for the pooling of multiple PCR products onto one
sequence run (Binladen et al., 2007).
17
Figure 3: Schematic representation of “Fusion Primers” layout. The Sif A and B are located on the 5’ end indicated in blue. The gene specific primer is on the 3’ end indicated in red. In the centre is a barcode (yellow).
The fusion primers used in this study targeted the V3 – V4 region of the 16S rDNA of
Bacteria and Archaea (Table 2). The comparative sequence analysis of the whole 16S rRNA
or hyper variable regions such as V3 or V6 are well established approach for the
phylogenetic classification of Bacteria and Archaea, with extensive databases available
(Huse et al., 2008; Tamaki et al., 2011; Woese, 1987). The primers used in this study were
tested and evaluated elsewhere (Klindworth et al., 2012).
Together with gene specific primers, phusion polymerase (New England, Bioloabs, UK) was
used to amplify the DNA. Phusion polymerase is a high fidelity polymerase (Frey &
Suppman, 1995). It has high speed and yield amplification with an error rate of 4.4 X 10-7
errors per base pair, which is 50 times lower than the regular Taq polymerase. This is due to
the 3´- 5´ exonuclease or “proofreading” activity, which results in a 3-fold increase in the
fidelity of DNA synthesis (Li et al., 2006).
Table 2:Sequence of Primers applied in this study, primers were names after (Alm et al., 1996)
Primer Name Sequence 5’ – 3’ Fragment
size (bp)
Annealing
temperature
(°C)
16S_D_BAC_0341_A_s CCTACGGGNGGCWGCAG 444 55
16S_D_BAC_0785_A_a ACTACHVGGGTATCTAATCC 444 55
16S_D_ARCH_0340_A_s CCCTACGGGGYGCASCAG 660 57
16S_D_ARCH_1000_A_a GGCCATGCACCWCCTCTC 660 57
2.7.2 Procedure
PCR was carried out in a total volume of 20 µl using the primers indicated inTable 2.The
master mix components and concentrations are shown in Table 3. The master mix and
18
DNAwas incubated in a thermocycler (Mastercycler Tm gradient, Eppendorf, Hamburg,
Germany) with the program indicated inTable 3. Subsequently, the PCR products were
visualized by gel electrophoresis with a 1% LE agarose (Biozyme, Oldendorf, Germany)
dissolved in 100 ml 1X TAE Buffer (For 2 l; 50 X TAE Buffer; 484 g 2 M Tris, 114.2 ml glacial
acetic acid and 200 ml 0.5 M EDTA pH 8.0). A total of 19 µl of the PCR product was mixed
with 5 µl 6 X loading dye (30%Glycerol, 0.25% Bromophenol blue in Milli-Q® water) and
loaded onto the gel. 5 µlpeqGOLD DNA ladder (PEQLAB Biotechnologie GMBH, Erlangen,
Germany) was used as a standard. Electrophoresis was carried out for 1 hr at 100 V. The
gel was stained for 30 min in ethidium bromide nd visualised using a documentation system
(Vilber-Lourmat, Eberhardzell, Germany).
Table 3:PCR reaction mix and thermoycler conditions applied in this study. X is the temperature used for annealing which varied according to the primers used.
PCR Reaction Thermocycler conditions
Molecular grade H2O
5 x HF buffer
dNTP’s mix (2,5 mM)
Forward Primer ( 100 µM)
Reverse Primer (100 µM)
Phusion Polymerase (0,02 units µl-1
)
DNA (10 ngµL-1
)
DMSO
12 µl
4 µl
1.6 µl
0.4 µl
0.4 µl
0.2 µl
1 µl
0.3 µl
1 cycle:
94°C for 5 min
35 cycles:
94°C for 30 sec
X °C for 30 sec
72°C for 2 min
1 cycle:
72°C for 10 min
Amplicon bands were visualized using a transilluminator DR-45M (Clare Chemical
Research, Göttingen, Germany) and cut out with a sterile scalpel. The cut out gel slices were
transferred in pre weighed 1.5 ml tubes and the DNA was purified using the Qiagen
MinElute® kit (Qiagen, Hilden, Germany). Briefly the weight of the slice was determined and
three volumes of QG buffer were added to one volume gel. The mix was incubated at 50 °C
for 10 min. Then one volume isopropanol was added and mixed by inverting. This was
applied to a MinElute spin column and centrifuged at 12,000 x g for 1 min. The flow through
was discarded and 500 µl QG buffer was added to the column followed by centrifugation at
13,200 x g for 1 min. Again the flow through was discarded and 750 µl of PE buffer was
added and this was centrifugation at 12,000 x g for 1 min. The column was placed in a new
collection tube and the wash step was repeated with 96% ethanol. Finally the column was
place in a new clean 1.5 ml tube and 10 µl EB buffer was added to the centre of the
19
membrane without touching the wall of the column and incubated for 5 min at RT. The DNA
was eluted by centrifugation at 12,000 x g for 1 min yielding 8 - 9 µl purified DNA.
After purification the PCR products were pooled into libraries containing several samples but
only one replicate of each barcode. The libraries had a minimum DNA concentration of 1 µg
DNA which was measure using a Qubit assay (Invitrogen, Darmstadt, Germany). The
libraries were then sent to the Max-Planck Institute for Plant Genomics in Cologne, for
sequencing on a ROCHE 454 titanium FLX (ROCHE, Germany).
2.8 ROCHE 454 Pyro- Sequencing
The Roche 454 sequencing approach is a sequencing by synthesis method. The DNA used
during sequencing can be genomic DNA fragments or amplicons. The DNA with attached
sequence adaptors is immobilized on capture beads. One piece of DNA is immobilized on
one capture bead (Margulies et al., 2005). These DNA fragments are them amplified on the
bead in an emulsion PCR (emPCR) reaction (Roche, 2010c). After the emPCR a single
capture bead will have multiple copied of a single fragment of DNA attached to it. Now the
beads are transferred to a picotiter plate (PTP). The PTP has a very small valve diameter
(44 µm) allowing for only one capture bead per valve (Dressman et al., 2003; Ghadessy et
al., 2001) (Figure 4 A-F). Then enzymes (polymerase, apyrase, ATP sulfuryase, luciferase)
are added to each valve for the sequencing reaction. The DNA fragment is synthesised
using a polymerase. The PTP is sequentially flooded with one of the four
deoxyribonucleotide triphosphates (dNTPs) (Margulies et al., 2005), if a dNTP is
incorporated then pyrophosphate (PPi) is released. The ATP sulfurase quantitatively
converts the PPi to ATP. When ATP is formed a light signal is produced by the luciferase
catalysed reaction. This light is detected by a charge coupled device (CCD) camera and
integrated into a pyrogram. The process continues with the addition of the next dNTP and a
complementary DNA strand is synthesised. The sequence of which can be deduced from the
pyrogram signal intensities (Figure 4 G). This is done for each valves and each DNA
fragment which is bound to the initial bead. This allows for a high number of individual
sequencing reactions to be performed simultaneously(Roche, 2010d).
20
(G)
Figure 4 A-G :Principle of 454-Sequencing, images modified from (Mardis, 2008). (A) Top Left: The DNA is fragmented and adaptors are added to the 5’ and 3’ ends. (B) The DNA fragment is incorporated into a bead, then the beads are captured in droplets of the oil from the emulsion PCR mixture. During the emPCR the DNA on the individual beads is amplified exponentially resulting in millions of copies of a single DNA fragment on every bead. (C) The beads are purified out of the emPCR mixture and beads are deposited into valves of the PTP. (D) Additional enzyme beads are added to each valve. (E) Microscope picture of emPCR process showing empty droplets and droplets with beads. The small arrow is indicated a bead and the large arrow a droplet. (F) Scanning electron microscope picture of PTP valves. (G)During sequencing the DNA-polymerase incorporates the complementary nucleotide and sets free inorganic pyrophosphate. The pyrophosphate is converted by the action of sulfurylase to ATP. During the action of luciferase energy in form of ATP is uses to create a flash of light. The light is detected by a CCD Camera and the Nucleotide sequence is given out as a pyrogram by the software. Apyrase degrades surplus molecules of ATP and e.g. dGTP before the next nucleotide is washed over the picotiter plate. Images modified from (Fabrice & Didier, 2009).
2.9 Analysis of 454 Sequences
The sequence reads from the 454 sequencer were further processed using the
bioinformatics pipeline of the SILVA project (Quast et al., 2013b). This involved quality
controls for sequence length (> 200 bp) and the presences of ambiguities (< 2%) and
homopoymers (<2%). All sequences which did not meeting these criteria were removed and
not considered in further analysis. The remaining reads were aligned against the SSU rRNA
seed of the SILVA database release 108 (http://www.arb-
silva.de/documentation/background/release-108). Reads which were not aligned were not
further considered in this study. This allowed for the removal of putative non SSU rRNA
gene reads and other artefacts. The remaining reads were dereplicated, clustered and
classified. Dereplication was done with cd-hit-est of the cd-hit package 3.1.2
(http://www.bioinformatics.org/cd-hit) using an identity criterion of 1.00 and a word size of 8.
21
The reads remaining after dereplication were clustered with cd-hit-est using an identity
criterion of 0.98. The longest read of each cluster was used as a reference for taxonomic
classification. The classification was done by a local BLAST search against the SILVA
SSURef 108 NR database (http://www.arb-silva.de/projects/ssu-ref-nr/) using blast-2.2.22+
(http://blast.ncbi.nlm.nih.gov/Blast.cgi) with standard settings. The obtained full SILVA
taxonomic path of the best blast hit was assigned to the read when the values for
(% sequence identity + %alignment coverage)/2
were at least 93.0. Finally the taxonomic path of each cluster reference read was mapped
onto the other reads in the cluster and the corresponding replicates. This gave (semi-)
quantitative information (number of individual reads representing in a taxonomic pool) on the
composition of the original PCR amplicon pool. The relative abundance of sequences
belonging to a taxonomic level was represented in 100% stack column graphs. The analysis
of ecological distance between the samples was computed with a Bray-Curtis similarity
analysis with 1000 times bootstrapping in the statistical software package
PASThttp://folk.uio.no/ohammer/past/).
Richness and diversity analyses were done using OTU (operational taxonomic unit, from
98% similarity clustering of sequences) as the basic unit. Due to the variability in the
bacterial species definition and the bias applied to the method by PCR direct abundance
analysis of the sequences was not used to do diversity analysis. Rarefraction curves were
calculated for each site based on the OTU number.
22
3. Results & Discussion
3.1 Classification of Ocean Biogeographic Provinces along the AMT22
Oceanic biogeographic provinces are dynamic with varying boundary locations (Oliver &
Irwin, 2008). This is due to the variation of oceanographic and atmospheric pressures acting
on the oceans at any given time. Biogeographic provinces are classified based on their
physical, chemical and biological parameters. The AMT22 covered 6 biogeographic
provinces according to their definition by Longhurst, (2007). The boundary of each province
was determined using the change in temperature (°C), salinity (PSU), oxygen (µmol l-1) and
chlorophyll a (mg m-3) shown in Figure 5 and Figure 6.
Figure 5: Temperature (°C) and salinity (PSU) section plot along the AMT22 by latitude (50°N– 45°S).
23
Figure 6: Oxygen concentration and chlorophyll a (calibrated against Winkler titrations) section plot along the AMT22 by latitude (50 °N – 45 °S).
The first province crossed during the cruise was the North Atlantic Drift (NADR) extending
from 50°N to 43°N. It had constant temperatures of 10 – 15 °C along a vertical and latitudinal
gradient. The salinity had a constant value of 35.5 PSU. Oxygen had a high concentration in
the surface region (260 - 270 µmol l-1) but stayed at a consistent 240 µmol l-1with depth. High
surface chlorophyll a values of 1 mg m-3 were measured which decreased with depth.
Next the North Atlantic Subtropical Gyre (NAST), which covered latitude from 43°N to 32°N,
was crossed. The surface water temperature was 20 °C which decrease rapidly to 15 °C by
100 m. Salinity increased to 36.5 PSU along a latitudinal gradient and was highest in surface
waters at 32°N. Oxygen concentrations were at a maximum of 240 µmol l-1 at 100 m. The
highest chlorophyll a concentrations were measured at 100 m depth (1 – 1.2 mg m-3). The
area of maximum chlorophyll a deepened along a latitudinal gradient (50 to 150 m).
The next province transverse during the AMT22 was the North Atlantic Tropical Gyre
(NATR) from 32°N to 17°N. It was defined based on a characteristic increase in water
temperature to 25 °C at the surface. The temperature decreased with depth and gradually
reaching 15 °C at 400 m. A high broad band of salinity (36 PSU) is also characteristic of the
province and was observed during this study. Oxygen concentrations had a maximum of 230
µmol l-1 at a depth of 100 to 200 m throughout the province. The chlorophyll a maximum (>
1mg m-3) was at 120 m depth. The Western Tropical Gyral (WTRA) province spanned from
24
17 °N to 8 °S. It had high surface temperatures of 25 - 27 °C, which were the highest
seawater temperatures observed during this study. Temperature decreased quickly with
depth to 15 °C within 200 m. Salinity was highest at 70 m depth (36 PSU) and decreased to
35 PSU above and below. Oxygen was highest at the surface (200 µmol l-1) but decrease
considerably with depth (100 µmol l-1). There was a wide band of chlorophyll a (0.5 – 1.5 mg
m-3) between 70 – 150 m depth.
The last two provinces transverse during the AMT22 were the South Atlantic Gyre (SATL)
and the South Atlantic Subtropical Gyral (SSTC) province. The SATL was the largest
province and spanned from 8 °S to 35 °S. It had surface temperatures of 20 -25 °C which
decreased slowly with depth. Salinity was high (37 PSU) at the surface to 200 m but
decreased below that. The oxygen concentration was 220 µmol l-1 throughout the water
column with a slight decrease with depth. Chlorophyll a was low with a maximum of < 0.5mg
m-3 between 150 – 200 m. The most rapid change in the physical parameters was observed
during the transition between the SATL and the SSTC provinces. The SSTC is defined by
characteristic steep gradients in temperature and salinity. These were also observed during
the AMT22 where surface temperatures drop to 15 °C and decreased even further to 5 °C by
120 m depth. These were the lowest temperatures measured along the AMT22. The same
rapid change in salinity was observed with the lowest salinity concentrations of the cruise
being observe (34.5 PSU) in this province. A high oxygen concentration was observed
throughout the water column (270 µmol l-1). In the surface waters of the SSTC the highest
chlorophyll a concentrations (> 1.5mg m-3) were measured between 0 and 70 m.
The NADR and SSTC provinces showed constant temperature and salinity values
throughout the water column, indicating a deep mixing process. Deep mixing increases the
nutrient availability in surface waters and results in high primary productivity (Longhurst,
2007). This was also observed during this study by the high chlorophyll a and oxygen values
in the water column, corresponding to oxygen production by photosynthesis.
In the central provinces (NAST, NATR, WTRA, SATL) a thermocline was observed. A
thermocline is an area in the water column with an abrupt change in temperature. Often
associated with a thermocline is a deep chlorophyll maximum (DCM) layer (Mann & Lazier,
1996). This was also observed in this study through the deepening of the chlorophyll a layer
(Figure 6). A DCM layer forms because a thermocline acts as a barrier trapping nutrients in
deeper waters. Phototrophic organisms aggregate near the thermocline to obtain essential
nutrients required for photosynthesis. In the NAST and southern SATL regions the depth of
the thermocline varied. The formation of a thermocline is dependent on the heating of
surface waters by solar irradiance. When the solar irradiance decreases sea surface
temperatures decrease and the thermocline diminishes. Subsequently mixing can occur and
25
nutrients are made available in surface waters allowing for primary production (Mann &
Lazier, 1996).
The gyral provinces (NATR and SATL) have deeper warm water and salinity layers due to
the down welling of water from the Ekman transport. The Ekman transport is the net motion
of fluid resulting from the balance of the Coriolis force and wind turbulence. It causes a
convergence of water in the gyre regions and the subsequent down welling due to the
excess mass of water converged (Mann & Lazier, 1996). This down welling causes a
depression in the thermocline which is usually deeper in the SATL province (Maranön et al.,
2000). The high surface salinity observed in the gyral regions is due to the characterised net
evaporation at these latitudes (Robinson et al., 2006; Weisse & Storch, 2010).
The WTRA province is characterised by high precipitation rates (Longhurst et al 2007),
which explained the low salinity values measured at the surface during this study. The
Ekman transport also acts on this region but causes an upwelling of Antarctic Intermediate
Water (AAIW). This was observed by the influx of low oxygen waters from depth. These
waters are low in oxygen due to their prolonged absence from the atmosphere (Robinson et
al., 2006).
3.2 Sequencing Statistics and Diversity Analysis
The 16S rRNA hypervariable regions V3-V4 was amplified because it has previously been
shown to be 99% accurate when classified to genus level (Huse et al., 2008). The six
sampling sites are referred to by the specific name of the province in which they were taken.
A large amount of sequences were produced from a single sequencing run (summary
statistics are shown in Table 4). Sample richness varied from 5806 to 2276 OTUs.
Rarefraction analysis of the bacterial diversity (Appendix 2) indicated that only part of the
total richness of each site was observed in our samples although previous studies have
indicated an asymptotic curve could be obtained for Atlantic planktonic samples (Friedline et
al., 2012).
26
Table 4: Sequencing statistics for 6 bacterial communities along the AMT22. Sequences indicates the total number of sequences obtained for a specific sample. Checked sequences are sequences which passed the quality controls. Clustering of sequences into OTUs was done using cd-hit-est with a criterion of 0.98. Replicates indicate sequences with 100% identity.
Site /Province NADR NAST NTRA WTRA SATL SSTC
Statistics
Number of sequences 29505 17134 25429 15662 18310 20912
Average length (bp) 458 510 452 458 458 458
Checked sequences 18633 10409 14147 9472 11785 14465
Clusters (98%) (OTUs) 5806 2276 2956 3227 3504 3557
Replicates 5027 4419 8294 2932 2984 3041
The sequencing resulted in 22583.5 ± 6921.5 sequences reads per site which fell into 4041
± 1765 OTUs (clustered at 98% identity). The average sequence length was 465 bp covering
the complete 16S rRNA gene hypervariable regions V3 and V4. The variability in number of
sequences and number of OTU was taken into account in the diversity analysis.
Microbial community diversity indices for all 6 sites based on a 98% sequence similarity cut
off (OTU) showed high microbial diversity (Table 5). There was no significant spatial pattern
observed in the diversity analysis. However the highest bacterial richness (number of OTUs)
and diversity (Shannon’s Index) were found in the NADR and SSTC provinces. These
provinces have characteristic high primary production associated with a large amount of
organic matter production which could result in an increase in the diversity of
microorganisms (Teeling et al., 2012). Simpson’s dominance index indicated that all sites
were dominated by specific organisms. This correlated to the evenness values which were
low indicating that there were rare and abundance organisms present.
Table 5: Diversity indices results for 6 bacterial communities along the AMT22. Individual OTU indicated the number of OTUs present at each site. Dominance was calculated using the 1+ Simpson’s index.
Diversity Indices NADR NAST NTRA WTRA SATL SSTC
Individual OTUs 5806 2276 3956 3227 3504 3557
Dominance Index 0.29 0.47 0.39 0.34 0.35 0.25
Shannon’s Index 1.57 1.15 1.27 1.44 1.39 1.57
Evenness (Shannon’s H Equitability) 0.54 0.35 0.41 0.47 0.45 0.60
27
3.3 Relative Abundance of Phyla
Eight phyla represented 97% of the relative sequence abundance of all provinces (Figure 7
a). The Proteobacteria had an average abundance of 39.5 ± 11.5% (Figure 7 a) and were
most abundant in the NAST and NATR provinces (51%). The Cyanobacteria had an average
abundance of 28.5 ± 10.5% in all provinces. Additionally the Bacteroidetes were present at
all station with 12 ± 7%. The abundance of Bacteroidetes was higher in the NADR and
SSTC provinces which had higher chlorophyll a concentrations. Bacteroidetes are known to
play an important role in the degradation or organic matter and have been previously
associated with phytoplankton blooms (Pereira, 2010; Teeling et al., 2012).
Actinobacteria made up a similar abundance with a maximum (15%) in the WRTA province.
This high abundance of Actinobacteria has previously been shown by (Morris et al., 2012)
however their functional role is still unknown because there are no isolated representatives.
The other phyla (Planctomycetes, Chloroflexi, Deferribacteres and Planctomycetes)
represent <10% abundance in all regions. Not indicated in Figure 7a. are phyla which had
relative sequence abundance below 1%. The results obtained in this study was consistent
with other studies of marine ecosystems (Friedline et al., 2012; Ghiglione et al., 2012;
Kirchman et al., 2010; Schmitt et al., 2012). These studies found similar abundance at the
phylum level, although they targeted various regions of the 16S rRNA.
The relative abundance of phyla in each province was analysed using a Bray-Curtis
similarity cluster (Figure 7 b). It compares the presence, absence and relative abundance of
each phylum between each province. There was a higher similarity between the sites
located in areas of low chlorophyll a, high temperature and high salinity (NAST, SATL,
NATR). Sites which had a low similarity exhibited high chlorophyll a measurements. Sites
with comparable physical and chemical conditions showed a high similarity in the abundance
of different phyla. This is evidence of biogeographical distribution at the phylum level. The
SSTC province showed the lowest similarity to all other provinces. From the relative
abundance it is apparent that a change in the dominance from Proteobacteria to
Cyanobacteria occurred in this province.
28
(a)
(b)
0.72
0.76
0.80
0.84
0.88
0.92
0.96
Sim
ilarity
SS
TC
NA
ST
SA
TL
NA
TR
WT
RA
NA
DR
Figure 7: Comparison of bacterial abundance at phylum level between the different sampling sites. (a) 100% stack column graph representing relative sequence abundances of the dominant phyla at different stations. (b) Bray-Curtis cluster analysis of the bacterial diversity at the phylum level using PAST.
29
3.4 Relative Abundance of Dominant Phyla
The relative abundance of Proteobacteria, Cyanobacteria and Bacteroidetes represented a
significant fraction (76%) of all sequences. Their high abundance could indicate a significant
function in the environment (Hunt et al., 2012) therefore they were individually analysed to a
deeper taxonomic resolution.
The Cyanobacteria were represented by two main genera. The Prochlorococcus
represented 83% of all cyanobacterial sequences in the subtropical and tropical provinces
and the genus Synechococcus represented 10 ± 5% in all provinces.
The Proteobacteria made up the highest abundance (51%)at all sites. Of this 99% consists
of the classes Alphaproteobacteria (55 ± 15%), Gammaproteobacteria (39 ± 14%),
Deltaproteobacteria (8 ± 5%) and Betaproteobacteria (2 ± 1%). Other Proteobacteria made
up less than 0.5% abundance together (Figure 8 a). Alphaproteobacteria had the highest
abundance in all provinces. The Alphaproteobacteria SAR11 made up nearly half of this
abundance with 41.5 ± 15.5%. SAR11 was present to a high abundance in all provinces but
was highest in the NATR and SATL provinces. These are the two gyral provinces which
have characteristic low chlorophyll a concentrations and are considered to be oligotrophic
environments. SAR11 is known for its ability to strive under oligotrophic conditions (Morris et
al., 2002) which could explain the high abundance in these regions. The wide distribution of
SAR11 shown in this study has been previously shown and it is believe to be ubiquitous in
the marine environment (Giovannoni et al., 2005; Hunt et al., 2012; Rappé et al., 2002).
The Gammaproteobacteria were the second most abundance class (39 ± 14%). Their
abundance was highest in the NADR, NAST and SSTC provinces which are characterised
by high primary production. Gammaproteobacteria have been associated with the
degradation of organic matter from phytoplankton which may explain their high abundance in
these provinces (Tada et al., 2011). Additionally a large portion of the gammaproteobacterial
abundance was made up of the SAR86 clade (15 ± 5%). Previous studies have shown
similar high abundances in this clade throughout the water column (Dupont et al., 2011;
Molloy, 2012).
Bray-Curtis similarity analysis (Figure 8 b) showed that there was a high similarity between
the Proteobacteria in all sites (r2 0.72). There was no distinguishable biogeographical
distribution within the Proteobacteria. The high similarity could be due to the dominance of
single subgroups such as SAR11 and SAR86, which appear to have omnipresence
throughout all provinces.
30
(a)
(b)
0.75
0.78
0.81
0.84
0.87
0.90
0.93
0.96
0.99
Sim
ilarity
WT
RA
NA
DR
SA
TL
SS
TC
NA
TR
NA
ST
Figure 8: Comparison of proteobacterial abundance, at class and order level, between the different sampling sites. (a) 100% stack column graph representing relative sequence abundances of the dominant classes at different stations. (b) Bray-Curtis cluster analysis of the proteobacterial diversity at the class level using PAST.
The Bacteroidetes were analysed to genus level for each biogeographical province because
they were represented at class and order level to 92% by the Flavobacteria,
Flavobacteriales. The relative abundances of Bacteroidetes was analysed by Bray-Curtis
31
similarity clustering (Figure 9). The similarity between sites was much less (r2 0.26) than
previously seen in the Proteobacteria. Geographically closer sites were clustered together. A
high similarity between geographically adjacent site such as NTRA and WTRA was
observed. Between the NAST and NADR province there was a low similarity (r2 0.32) but
they were still more similar to each other than more distant provinces. The clustering of
adjacent provinces indicates a similarity in the diversity in geographically closer regions. This
also leads to the inference that there is a latitudinal biogeographical distribution of distinct
Bacteroidetes genera.
The sequence abundances indicated that at a phylum level there was a high similarity
between all biogeographical provinces. The main differences that were observed could not
be related to geographical location but to the physical and chemical structure of the
environment. At a phylum level there was a clustering correlated to the level of primary
production within the environment.
The Proteobacteria did not exhibit a distinct clustering based on geographical location
because they are dominate by groups such as SAR11 or SAR86 which had an
omnipresents. The Bacteroidetes however had a more diverse assemblage with no clear
dominating genera and show distinct geographical distributions.
32
0.24
0.32
0.40
0.48
0.56
0.64
0.72
0.80
0.88
0.96
Sim
ilarity
NT
RA
WT
RA
SA
TL
SS
TC
NA
ST
NA
DR
Figure 9: Bray-Curtis cluster analysis of the Bacteroidetes diversity at the genus level using PAST.
High through-put pyrosequencing of the PCR amplified 16S rRNA gene provides an in depth
analysis of the bacterial diversity in distinct biogeographical provinces. The diversity analysis
lead to the conclusion the bacterial communities are not randomly but biogeographically
separated. However high through-put sequencing is semi quantitative and limited due to
PCR bias (Engelbrektson et al., 2010; Lee et al., 2012; Sergeant et al., 2012). To overcome
this limitation CARD-FISH was used to analyse the in situ abundance of the organisms
which dominated the sequencing data. This data was subsequently correlated to the
physical and chemical measurements taken at each sampling point, to see if there was a
biogeographical distribution pattern.
3.5 Absolute and Relative Abundances of Picoplankton along AMT22
To get an indication of the latitudinal and vertical distribution of the picoplanktonic community
of the Atlantic Ocean the three domains of life was analysed for their absolute and relative
33
abundances. There was a considerable difference when looking at the absolute (cell ml-1)
and relative cell abundances (% of DAPI). This is indicated in Figures 15 a & b and 16 a & b.
Figure 10:(a) Latitudinal-depth contours of Picoplankton cell abundances determined by DAPI (DNA) staining from 50°N to 50°S from 20 to 300m.(b) Latitudinal-depth contours of bacterial cell abundances determined by CARD-FISH using the Eub338 I-III probe.
The absolute picoplankton abundance (cells ml-1) changed with latitude and with depth
(Figure 10 a). In high latitudes and at the equator there were >1.5E+06 cells ml-1, whereas
gyre regions had <1.0E+06 cell ml-1. With depth there was a change from > 1.5E+06 cells
ml-1 to < 5E+05 cells ml-1. The picoplankton abundance was positively correlated high
chlorophyll a concentrations (r2 0.76).The highest cell numbers were observed in the surface
waters below 40 °S, shown in red figure 11 a. This was also the area of highest chlorophyll
a. Not hybridized absolute cell numbers did not reflect a specific latitudinal or depth
distribution (2.9E+05 ± 1.2E+05 cells ml-1). The correlation of picoplankton abundance with
34
high chlorophyll a can be accounted for by the fact that a large part of the picoplankton is
itself phototrophic. Eukaryotic primary production in the ocean is nutrient limited (Margalef,
1987). In high latitudes and at the equator there are higher mixing rates which results in the
availability of nutrients (Robinson et al., 2006). Subsequently, there is a significant increase
the amount of primary production (Yooseph et al., 2010) and in the absolute cell numbers,
which was also observed during this study. The increase in primary production results in a
high amount of organic matter production due to fixation of inorganic carbon by phototrophs
(Falkowski et al., 1998). This increase in organic matter causes an increase in the
heterotrophic prokaryotic community (Teeling et al., 2012; Yooseph et al., 2010). This was
also seen in the difference in the bacterial cell abundances in this study (Figure 10 b).
Bacteria showed a decrease in absolute cell abundances with depth from 1.1E+06 cells ml-1
to 8.2E+04 cells ml-1. Additionally, there were higher cell numbers in high and central
latitudes, corresponding with the picoplankton high abundance areas. Bacteria were
positively correlation to high light (r2 0.43) and chlorophyll a (r20.71). Although there was a
higher absolute abundance of Bacteria in high and central latitudes, they made up a larger
proportion of the picoplankton in gyre regions (up to 98%) (Figure 11). Bacteria dominate the
surface waters of all provinces with an average abundance of 72 ± 16%. The relative
abundance of Bacteria decreased to 59 ± 15% in the DCM and to 51 ± 9% in the
mesopelagic layer. The high abundance of Bacteria in marine systems has resulted in a
large amount of research to deduce the role which they play and their significance in higher
food webs and element cycles (DeLong, 2009).
There was a change in the community from Bacteria dominated at the surface to a near
mixed community of Bacteria and Archaea in mesopelagic depths in all biogeographical
provinces.
35
Figure 11: Latitudinal-depth contours of bacterial and archaeal relative cell abundances (% DAPI counts).
Archaea had an absolute abundance of 6.1E+04 ± 2.1E+04 cells ml-1 throughout the water
column, however their relative abundance (Figure 11)showed a significant change from 0 to
40%. Archaea had an abundance of 2 ± 1% at the surface, with an increase to 7% in the
NADR province. This increased to 10 ± 5% in the DCM layer and reached a maximum of 37
± 3% in mesopelagic depths. The increase in the relative abundance of Archaea has also
been reported by (Herndl et al., 2005). They measured the archaeal production and found it
contributed to up to 37% of the total prokaryotic production in meso- and bathypelagic
depths. This indicates that Archaea are an active part of the deep ocean prokaryotic
community. There was a clear difference in the distribution of the two domains but unlike the
Bacteria the absolute abundance of Archaea stays relatively constant with depth.
3.6 Relative Abundances and Biogeography of the Dominant Bacterial
Taxon along the AMT22
The dominate groups found in the tag sequence (Prochlorococcus, Synechococcus, SAR11,
SAR86 and Bacteroidetes) were analysis for relative cellular abundances and distribution.
Their relative cellular abundances were enumerated by using groups specific CARD-FISH
probes. Results for each biogeographical region are shown in appendix 3.
36
3.6.1 Cyanobacteria
The Cyanobacteria are photoautotrophic prokaryotes (Whitton & Potts, 2012). Two classes,
Prochlorococcus and Synechococcus, dominated the picoplankton community in the
euphotic zones (<300 m) (Figure 12). Prochlorococcus is unicellular and has a considerable
impact on global carbon cycles. Marine primary production accounts for 40% of the global
carbon fixation, of which Prochlorococcus contributes up to 50% (Malmstrom et al., 2013;
Partensky et al., 1999). This study found a 20% abundance of Prochlorococcus in tropical
and subtropical regions which was also shown in the study by (Schattenhofer et al., 2009).
At the surface it represented 13.5 ± 5.5% abundance and in the DCM 9 ± 6%. It was also
present at 150 m in the SATL provinces at 3%. In the gyral provinces (NATR, SATL) it was
the second most abundant group with 19% at the surface and 13.5 ± 1.5% in the DCM layer.
Previously have shown it to be ubiquitous in tropical and subtropical regions (40° N to 40°S)
and can occupies a considerable area of the water column (20 -200 m) (Partensky et al.,
1999), which was also observed in our study (2% at 200 m). Spearman Rank correlations
indicated a positive relationship between Prochlorococcus and temperature (r2 0.68) and
light (r2 0.63). This is consistent with its phototrophic metabolisms and temperature
dependent distribution (Malmstrom et al., 2013). Previous genomic and chemical analysis of
different Prochlorococcus strains showed that it has evolved divinyl derivatives of chlorophyll
a and chlorophyll b. These pigments absorb the fluorescence emission of the blue part of the
light spectrum and are characteristic of the Prochlorococcus. Blue light penetrates deepest
into the oceans and these pigments allow Prochlorococcus to exist in areas of less than 1%
surface irradiance (Zwirglmaier et al., 2008).
Synechococcus had a lower abundance than Prochlorococcus but had a wider latitudinal
distribution. It highest abundance was in high latitudes where it made up 5 ± 5% of the
picoplankton. Although Prochlorococcus and Synechococcus were present throughout the
water column there was a higher overall abundance of Prochlorococcus especially in lower
latitudes. Previous studies have shown that Synechococcus exists at about one order or
magnitude lower than Prochlorococcus but still makes a considerable contribution to the
carbon fixation of the oceans(Palenik et al., 2003).Its higher abundance in areas of higher
bacteria abundances (NADR, SSTC) could be related to its growth dependence on lysis of
heterotrophic Bacteria. (Weinbauer et al., 2011) showed higher growth rates of
Synechococcus in the presence of viruses due to their ability to utilize essential nutrients in
organic forms (amino acids, oligopeptides, phosphonates).
37
Figure 12: Latitudinal-depth contours of Prochlorococcus and Synechococcus (SYN) relative cell abundances (% DAPI counts).
3.6.2SAR11 and SAR 86 Clade
The Alphaproteobacterium SAR11 represented up to 50% of the picoplankton community
(Figure 13).It made up 30.5 ± 5.5% of all cells in the surface, 20.5 ± 14.5% in the DCM and
26.5 ± 11.5% in the mesopelagic layer. It had a high abundance (>20%) across all latitudes
and with depth. It was more abundant in the North Atlantic gyre region (NTRA, NAST) which
was also shown by (Schattenhofer et al., 2009). SAR11 was first isolated by (Rappé et al.,
2002) and is likely the most abundant heterotrophic bacterial group in the marine
environment. It dominates in both oligotrophic and copiotrophic conditions and has been
shown to have photoheterotrophic nutrient uptake abilities (Gomez-Pereira et al., 2012).
It had a lower abundance in the SATL region, which was also shown on the AMT20 (25%)
and AMT13 (22%) (Mary et al., 2006; Schattenhofer et al., 2009). It also showed a low
abundance in the SSTC province with only 6% in the DCM layer. Previous reasons for a low
abundance were related to the variability in nutrient availability (Morris et al., 2002), however
the SSTC region has high mixing rates and nutrient availability. Another reason for their low
abundance could be the decrease in light penetration due to high primary production at the
surface resulting in a decreases in nutrient uptake. SAR11 have been shown to use
38
proteorhodopsins for nutrient and amino acid uptake (Gomez-Pereira et al., 2012; Mary et
al., 2008). Additionally (Morris et al., 2012) have recently indicated that the SAR11 clade is
comprised of multiple ecotypes which may have varying nutrient requirements. They have
shown that a fraction of the SAR11 community responds rapidly to the input of organic
matter by phytoplankton and are thus phytoplankton dependent, whereas the North Atlantic
ecotypes is more dependent on amino acid and reduced sulphur uptake. This variation in
ecotypes nutrient dependence could indicate variations in the abundance.
The Gammaproteobacterium SAR86 was present along all latitudes up to 200 m depth
(Figure 13). The abundance of SAR86 was one order of magnitude lower than SAR11.
SAR86 was most abundance (5%) in the surface waters of the gyral latitudes and high
southern latitudes. The uncultured SAR86 clade has previously been reported
(Schattenhofer et al., 2009) at much lower relative abundances (0.5%). In this study two
helper probes were applied which has been shown to increase the probe binding ability
(Fuchs et al., 2000; Zubkov et al., 2001). These results indicated that this group maybe more
abundance in the surface waters than previously thought.
Both SAR11 and SAR86 are heterotrophic Bacteria which coexist in the water column of the
Atlantic Ocean. The reason for their coexistence could be due to their carbon compound
specialisation. SAR86 has specialised for the uptake of lipids and carbohydrates (Dupont et
al., 2011) where as SAR11 is specialised in the uptake of C1 compounds (Morris et al.,
2012; Rappé et al., 2002).
39
Figure 13: Latitudinal-depth contours of SAR11 and SAR86 relative cell abundances (% DAPI counts). The x-axis scale varies between the two plots (SAR11 100%, SAR86 10%).
3.6.3 Bacteroidetes
The members of the Bacteroidetes represented an average abundance of 5% (Figure 14).
They were present throughout the water column but were most abundance in the surface.
Their highest abundance was in high latitudes (NADR, SSTC) where they had up to 25 %
abundance. They were positively correlated to chlorophyll a concentrations (0.77) which has
been previously shown by (Schattenhofer et al., 2009). The Bacteroidetes have been
extensively studies due to their direct association with primary production in the marine
environment. They are key players in the degradation of phytoplankton derived organic
matter in coastal systems (Tada et al., 2011). They have also been shown to exhibit
succession patterns associated with the variability in available organic matter (Fernandez-
Gomez et al., 2013; Pereira, 2010; Teeling et al., 2012). The apparent selection by different
organic matter compounds makes them an interesting for the study of biogeography.
However in this and other studies (Schattenhofer et al., 2009) Bacteroidetes also made up a
significant fraction in the low nutrient and chlorophyll a provinces (NTRA, SATL). These
Bacteroidetes seem to cope with limiting conditions in contrast to their costal counterparts.
40
This data also corresponds to the genus level Bray-Curtis similarity analysis of the 454
sequences of Bacteroidetes which indicated variability between different provinces.
Figure 14: Latitudinal-depth contours of Bacteroidetes relative cell abundances (% DAPI counts).
3.6.4 Other Groups of the Bacterioplankton Community
In addition to these groups which were singled out due to their high abundance in the 454
tag sequencing data, other bacterial groups were also analysed by CARD-FISH. These
groups (Roseobacter, SAR202 and SAR324) have previously been shown by (Schattenhofer
et al., 2009) to have a high cellular abundance in the Atlantic ocean. They represented minor
abundance in the tag sequence data. However two of these groups are known for their
abundance in the meso- and bathypelagic water column (Giovannoni & Vergin, 2012; Swan
et al., 2011) which were not analysed by tag sequencing.
The group Roseobacter was present in the surface waters of all provinces and the DCM of
NADR, representing 3.5 ± 0.5% of the total abundance (Appendix 3). Their distribution was
positively correlated with chlorophyll a (r2 0.763) and negatively with temperature and salinity
(r2 0.293, 0.345 respectively). The Cloroflexi like SAR202 was contrastingly only present in
the meopelagic depth representing 11.5 ± 4.5% of the total abundance. This has previously
been shown by Giovannoni & Vergin, (2012) and Morris et al., (2004) who showed that
SAR202 accounted for 10% of all bacterioplankton from 500- 4000m.
SAR324 exhibited the same mesopelagic distribution and was present at 4 ± 1%. Similar
wide distribution patterns have been shown previously in the Eastern Atlantic Ocean by
Schattenhofer et al., (2009). Swan et al., (2011) showed that SAR324 could be
chemolithoautotrophs which fix inorganic carbon at significant rates in the deep ocean.
41
Conclusion
This study has increased our understanding of the bacterioplanktonic diversity, distribution
and abundance in the Atlantic Ocean. It complements previous diversity and abundance
studies of the bacterioplankton of the Eastern and Central Atlantic Ocean (Friedline et al.,
2012; Morris et al., 2012; Schattenhofer et al., 2009).
Firstly, the bacterioplanktonic diversity in different biogeographical provinces was analysed
using massive parallel tag sequencing and subsequently, the abundance and distribution
patterns of the dominate bacterioplanktonic groups throughout the Atlantic Ocean were
enumerate. The dominant community made up a significant fraction of the total cellular
abundance of the picoplankton community. There were few groups (SAR11, SAR86 and
Bacteroidetes) which could be found at all latitudes and depths and although they were
ubiquitous in the 454 sequences, their relative abundances did vary. There was no direct
comparison between the relative sequence abundance and relative cellular abundance due
to the different nature of the methods applied. The distribution and abundance of the
dominant bacterioplankton groups could be correlated with the presence of strong
environmental drivers such as chlorophyll a and temperature. Additionally this could be
associated with previously described distribution patterns and assumed metabolic and
physiological capabilities of the organisms. The results presented here are consistent with
the emerging picture of the distribution patterns of the dominant microorganisms within the
Atlantic Ocean.
The methods applied in this study differ from previous studies because the diversity,
abundance and distribution were analysed and related to environmental parameters. When
analysing diversity using 454 sequencing a high taxonomic resolution is obtained but there is
no direct relation to the relative abundance, because there is a PCR step in the method (Lee
et al., 2012). The sequencing results can only be used for the analysis of presence and
absence or for a comparative analysis between the diversity and relative sequence
abundance of different sites, as shown in this study. The change in relative sequence
abundance can then be related to the change in diversity with latitude and correlated to
environmental parameters. However it is difficult to deduce from this, which organisms are
abundance or activity in the environment. The use of CARD-FISH allows for the analysis of
abundance which can be related to activity.
From the diversity results presented here it becomes apparent that the taxonomic resolution
is important when looking at the biogeography of microorganisms. Distinct biogeographical
trends may not be apparent or accurate at a broader taxonomic resolution. In this study at a
phylum level there was an apparent biogeography distribution based on environmental
42
parameters. Bray Curtis similarity analysis showed that sites with low chlorophyll a and high
temperatures were more similarly to each other. This indicated that at a phylum level there
was a difference in the abundance, presence and absence of different phyla depending on
primary production and temperature.
Within the classes of the Proteobacteria however there was no apparent biogeographical
distribution. This could have been due to the dominance of ubiquitious groups in all
provinces (SAR11, SAR86), or due to the fact that the Proteobacteria are a highly diverse
group of organisms with varying metabolic capabilities.
Within the Bacteroidetes there was a distinct biogeographical with latitude. Geographically
closer samples were more similar indicating a limited distribution or geographical isolation of
specific genera. From these three examples it became apparent that it is important to
consider the taxonomic resolution when looking for a biogeographical pattern due to the
ecophysiological and metabolic flexibility within different taxonomic groups.
When looking at diversity based on a high taxonomic level there is no ecological similarity
considered. Microorganisms of the same order can have extremely different physiologies.
This can be shown by the apparent ubiquitous nature of SAR11. This Alphaproteobacterium
was present to a high abundance throughout the Atlantic Ocean and showed no
biogeographical distribution in this study. However recent analysis have indicated that there
are specific ecotypes of SAR11 with varying nutrient requirements (Morris et al., 2012). It
may be that specific ecotypes of SAR11 do show a biogeographical distribution however this
is not reflected at a broad taxonomic level. The lineages microdiversity could determine its
success in the marine environment (Friedline et al., 2012) but could also give an indication of
it biogeographic distribution and possible reasons for its variations in abundance.
When looking at the biogeography of microorganisms it is important to consider that a
biogeographical distribution pattern is only expected if there is ecological coherence within a
lineage (Philippot et al., 2010). Then an association between the environment and the
biodiveristy can be defined. If there is no ecological coherence then a pattern of distribution
based on the chemical and physical environment is hard to assume because the
ecophysiologies can vary.
The Cyanobacteria are one of the few phyla which show ecological coherence (all
photoautotrophs). The diversity and distribution of cyanobacteria can therefore be correlated
with light because this is an essential requirement. They are known for having specific
temperature optimums (Zwirglmaier et al., 2008) which can be analysed and in this study
we saw a difference between the distribution of Synechococcus and Prochlorococcus, with
Prochlorococcus showing a correlation to higher temperatures. This was also reflected in
their abundance and distribution patterns from CARD-FISH analysis. The ecological
43
coherence and high abundance of Cyanobacteria may also account for the biogeographical
patterns observed in the 454 data at the phylum level.
The importance of taxonomic resolution could also be shown in the biogeographical pattern
in the Bacteriodetes shown in this study. They have been associated with high chlorophyll a
areas in this and previous studies using the relative cellular abundance. They did occur to
higher abundance in regions of high chlorophyll a in this study but not exclusively. The
diversity varied between different regions based on latitudinal distance not on the chlorophyll
a concentrations. Provinces with high chlorophyll a were not more similar in diversity. This
indicates that when interpreting microbial biogeography it is important to incorporate the
ecology of the specific taxonomic level not just the diversity or abundance.
The combined analysis of the diversity and abundance of microorganisms based on
biogeographical patterns resulted in a more accurate interpretation of the biogeography of
microorganisms in the Atlantic Ocean. Understanding the biogeography of groups of
microorganisms is a key step in understanding the ecosystem function of specific microbial
assemblages and interpreting and prediction variations to this pattern.
44
Acknowledgements
This master’s thesis was sponsored by the Max Planck International Research School for
Marine Microbiology. Thanks to the Master and crew of the RRV James Cook as well as the
principle scientist Glen Tarren from Plymouth Marine Laboratories. Thank you to my
supervisor Rudolf Amann who made this master possible and to the Department of
Molecular Ecology for hosting me. Additional thanks goes to Jörg Wulf, Martha
Schattenhofer, Mike Zubkov and Sara Cregeen for their technical support and insightful
discussions on cruise. Thank you also to Bernhard Fuchs, Christin Bennke and Marion
Stagars.
My final thanks goes to my mother Susanne Zühlke and the Marmic 15 (sunshine) class.
You have been a constant supportive ear and ever helping hand, thank you.
45
References
Alm, E. W., Oerther, D. B., Larsen, N., Stahl, D. A. & Raskin, L. (1996). The oligonucleotide probe database. Applied and Environmental Microbiology 62, 3557. Amann, R., Fuchs, B. M. & Behrens, S. (2001). The identification of microorganisms by fluorescence in situ hybridisation. Current Opinion in
Biotechnology 12, 231236. Amann, R. I., Binder, B. J., Olson, R. J., Chisholm, S. W., Devereux, R. & Stahl, D. A. (1990). Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Applied and environmental microbiology 56, 1919-1925. Arrigo, K. R. (2004). Marine microorganisms and global nutrient cycles. Nature 437,
349355. Azam, F. & Malfatti, F. (2007). Microbial structuring of marine ecosystems. Nature
Reviews Microbiology 5, 782791. Binladen, J., Gilbert, M. T. P., Bollback, J. P., Panitz, F., Bendixen, C., Nielsen, R. & Willerslev, E. (2007). The use of coded PCR primers enables high-throughput sequencing of multiple homolog amplification products by 454 parallel sequencing. PloS one 2, e197. DeLong, E. F., Wickham, G. S. & Pace, N. R. (1989). Phylogenetic stains: ribosomal RNA-based probes for the identification of single cells. Science (New York, NY) 243, 1360. DeLong, E. F. (2009). The microbial ocean from genomes to biomes. Nature 459, 200-206. Dressman, D., Yan, H., Traverso, G., Kinzler, K. W. & Vogelstein, B. (2003). Transforming single DNA molecules into fluorescent magnetic particles for detection and enumeration of genetic variations. Proceedings of the National Academy of
Sciences 100, 88178822. Dupont, C. L., Rusch, D. B., Yooseph, S. & other authors (2011). Genomic insights to SAR86, an abundant and uncultivated marine bacterial lineage. The ISME
journal 6, 11861199. Eilers, H., Pernthaler, J., Glöckner, F. O. & Amann, R. (2000). Culturability and in situ abundance of pelagic bacteria from the North Sea. Applied and Environmental Microbiology 66, 3044-3051. Eilers, H., Pernthaler, J., Peplies, J. r., Glöckner, F. O., Gerdts, G. & Amann, R. (2001). Isolation of novel pelagic bacteria from the German Bight and their seasonal
46
contributions to surface picoplankton. Applied and environmental microbiology 67, 5134-5142. Engelbrektson, A., Kunin, V., Wrighton, K. C., Zvenigorodsky, N., Chen, F., Ochman, H. & Hugenholtz, P. (2010). Experimental factors affecting PCR-based
estimates of microbial species richness and evenness. ISME J 4, 642647. Fabrice, A. & Didier, R. (2009). Exploring microbial diversity using 16S rRNA high-
throughput methods. J Comput Sci Syst Biol 2, 074092. Falkowski, P. G., Barber, R. T. & Smetacek, V. (1998). Biogeochemical controls and feedbacks on ocean primary production. Science 281, 200-206. Falkowski, P. G., Fenchel, T. & Delong, E. F. (2008). The microbial engines that
drive Earth's biogeochemical cycles. Science 320, 10341039. Fernandez-Gomez, B., Richter, M., Schuler, M., Pinhassi, J., Acinas, S. G., Gonzalez, J. M. & Pedros-Alio, C. (2013). Ecology of marine Bacteroidetes: a comparative genomics approach. ISME J.
Frey, B. & Suppman, B. (1995). BioChemical Journal 2, 3435. Frias-Lopez, J., Shi, Y., Tyson, G. W., Coleman, M. L., Schuster, S. C., Chisholm, S. W. & DeLong, E. F. (2008). Microbial community gene expression in ocean surface waters. Proceedings of the National Academy of Sciences 105,
38053810. Friedline, C. J., Franklin, R. B., McCallister, S. L. & Rivera, M. C. (2012). Bacterial assemblages of the eastern Atlantic Ocean reveal both vertical and latitudinal
biogeographic signatures. Biogeosciences 9, 21772193. Fuchs, B. M., Glöckner, F. O., Wulf, J. & Amann, R. (2000). Unlabeled Helper Oligonucleotides Increase the In Situ Accessibility to 16S rRNA of Fluorescently Labelled Oligonucleotide Probes. Applied and Environmental Microbiology 66, 3603–3607. Fuhrman, J. A., Steele, J. A., Hewson, I., Schwalbach, M. S., Brown, M. V., Green, J. L. & Brown, J. H. (2008). A latitudinal diversity gradient in planktonic
marine bacteria. Proceedings of the National Academy of Sciences 105, 77747778. Ghadessy, F. J., Ong, J. L. & Holliger, P. (2001). Directed evolution of polymerase function by compartmentalized self-replication. Proceedings of the National Academy
of Sciences 98, 45524557. Ghiglione, J. F., Galand, P. E., Pommier, T. & other authors (2012). Pole-to-pole biogeography of surface and deep marine bacterial communities. Proceedings of the
National Academy of Sciences 109, 1763317638.
47
Giovannoni, S. J., Bibbs, L., Cho, J. C. & other authors (2005). Proteorhodopsin in the ubiquitous marine bacterium SAR11. Nature 438, 82-85. Giovannoni, S. J. & Vergin, K. L. (2012). Seasonality in Ocean Microbial Communities. Science 335, 671-676. Gomez-Pereira, P. R., Hartmann, M., Grob, C., Tarran, G. A., Martin, A. P., Fuchs, B. M., Scanlan, D. J. & Zubkov, M. V. (2012). Comparable light stimulation of organic nutrient uptake by SAR11 and Prochlorococcus in the North Atlantic subtropical gyre. ISME J. Green, J. & Bohannan, B. J. (2006). Spatial scaling of microbial biodiversity.
Trends in ecology & evolution 21, 501507. Grossart, H.-P., Levold, F., Allgaier, M., Simon, M. & Brinkhoff, T. (2005). Marine diatom species harbour distinct bacterial communities. Environmental Microbiology
7, 860873. Herndl, G. J., Reinthaler, T., Teira, E., Van Aken, H., Veth, C., Pernthaler, A. & Pernthaler, J. (2005). Contribution of Archaea to total prokaryotic production in the
deep Atlantic Ocean. Applied and environmental microbiology 71, 23032309. Hunt, D. E., Lin, Y., Church, M. J., Karl, D. M., Tringe, S. G., Izzo, L. K. & Johnson, Z. I. (2012). Relationship between Abundance and Specific Activity of Bacterioplankton in Open Ocean Surface Waters. Applied and Environmental
Microbiology 79, 177184. Huse, S. M., Dethlefsen, L., Huber, J. A., Welch, D. M., Relman, D. A. & Sogin, M. L. (2008). Exploring Microbial Diversity and Taxonomy Using SSU rRNA Hypervariable Tag Sequencing. PLoS Genet 4, e1000255. Innis, M. A., Myambo, K. B., Gelfand, D. H. & Brow, M. (1988). DNA sequencing with Thermus aquaticus DNA polymerase and direct sequencing of polymerase chain reaction-amplified DNA. Proceedings of the National Academy of Sciences 85,
94369440. Kirchman, D. L., Cottrell, M. T. & Lovejoy, C. (2010). The structure of bacterial communities in the western Arctic Ocean as revealed by pyrosequencing of 16S
rRNA genes. Environmental Microbiology 12, 11321143. Klindworth, A., Pruesse, E., Schweer, T., Peplies, J., Quast, C., Horn, M. & Glöckner, F. O. (2012). Evaluation of general 16S ribosomal RNA gene PCR primers for classical and next-generation sequencing-based diversity studies. Nucleic Acids Research. Lee, C. K., Herbold, C. W., Polson, S. W., Wommack, K. E., Williamson, S. J., McDonald, I. R. & Cary, S. C. (2012). Groundtruthing Next-Gen Sequencing for Microbial Ecology Biases and Errors in Community Structure Estimates from PCR Amplicon Pyrosequencing. PLoS ONE 7, e44224.
48
Li, M., Diehl, F., Dressman, D., Vogelstein, B. & Kinzler, K. W. (2006). BEAMing up for detection and quantification of rare sequence variants. Nature methods 3,
9597. Lomolino, M. V., Riddle, B. R. & Brown, J. H. (2006).Biogeography. Longhurst, A., Sathyendranath, S., Platt, T. & Caverhill, C. (1995). An estimate of global primary production in the ocean from satellite radiometer data. Journal of
Plankton Research 17, 12451271. Longhurst, A. R. (2007). Chapter 9 - The Atlantic Ocean. In Ecological Geography
of the Sea (Second Edition), pp. 131273. Burlington: Academic Press. Malmstrom, R. R., Kiene, R. P., Cottrell, M. T. & Kirchman, D. L. (2004). Contribution of SAR11 bacteria to dissolved dimethylsulfoniopropionate and amino acid uptake in the North Atlantic ocean. Applied and environmental microbiology 70, 4129-4135. Malmstrom, R. R., Rodrigue, S., Huang, K. H. & other authors (2013). Ecology of uncultured Prochlorococcus clades revealed through single-cell genomics and biogeographic analysis. The ISME Journal. Mann, K. H. & Lazier, J. R. N. (1996). Dynamics of Marine Ecosystems 2nd edn. Cambridge, Massachusetts.: Blackwell Science, . Manz, W., Amann, R., Ludwig, W., Wagner, M. & Schleifer, K.-H. (1992). Phylogenetic oligodeoxynucleotide probes for the major subclasses of proteobacteria: problems and solutions. Systematic and Applied Microbiology 15, 593-600. Manz, W., Amann, R., Ludwig, W., Vancanneyt, M. & Schleifer, K.-H. (1996). Application of a suite of 16S rRNA-specific oligonucleotide probes designed to investigate bacteria of the phylum Cytophaga-Flavobacter-Bacteroides in the natural environment. Microbiology 142, 1097-1106. Maranön, E., Holligan, P. M., Varela, M., Mouriäo, B. & Bale, A. J. (2000). Basin-scale variability of phytoplankton biomass, production and growth in the Atlantic
Ocean. Deep Sea Research Part I: Oceanographic Research Papers 47, 825857. Mardis, E. R. (2008). The impact of next-generation sequencing technology on
genetics. Trends in Genetics 24, 133141. Margalef, R. (1987). Life-forms of phytoplankton as survival alternatives in an unstable environment. . Oceanologica acta 1, 493–509. Margulies, M., Egholm, M., Altman, W. E. & other authors (2005). Genome
sequencing in microfabricated high-density picolitre reactors. Nature 437, 376380.
49
Martiny, J. B. H., Bohannan, B. J., Brown, J. H. & other authors (2006). Microbial biogeography: putting microorganisms on the map. Nature Reviews Microbiology 4,
102112. Mary, I., Heywood, J. L., Fuchs, B. M., Amann, R., Tarran, G. A., Burkill, P. H. & Zubkov, M. V. (2006). SAR11 dominance among metabolically active low nucleic acid bacteriplankton in surface water along an Atlantic Meridional Transect. Aquatic Microbial Ecology 45, 107–113. Mary, I., Tarran, G. A., Warwick, P. E., Terry, M. J., Scanlan, D. J., Burkill, P. H. & Zubkov, M. V. (2008). Light enhanced amino acid uptake by dominant bacterioplankton groups in surface waters of the Atlantic Ocean. FEMS Microbiology
Ecology 63, 3645. Massana, R., Murray, A. E., Preston, C. M. & DeLong, E. F. (1997). Vertical distribution and phylogenetic characterization of marine planktonic Archaea in the Santa Barbara Channel. Applied and Environmental Microbiology 63, 50-56. Molloy, S. (2012). Marine microbiology: SAR86: streamlined for success. Nature
Reviews Microbiology 10, 8283. Morris, R. M., Rappé, M. S., Connon, S. A., Vergin, K. L., Siebold, W. A., Carlson, C. A. & Giovannoni, S. J. (2002). SAR 11 clade dominates ocean surface
bacterioplankton communities. Nature 420, 806810. Morris, R. M., Rappe, M. S., Urbach, E., Connon, S. A. & Giovannoni, S. J. (2004). Prevalence of the Chloroflexi-Related SAR202 Bacterioplankton Cluster throughout the Mesopelagic Zone and Deep Ocean. Applied and Environmental Microbiology 70, 2836-2842. Morris, R. M., Frazar, C. D. & Carlson, C. A. (2012). Basin-scale patterns in the abundance of SAR11 subclades, marine Actinobacteria (OM1), members of the Roseobacter clade and OCS116 in the South Atlantic. Environmental Microbiology
14, 11331144. Nübel, U., Garcia-Pichel, F. & Muyzer, G. (1997). PCR primers to amplify 16S rRNA genes from cyanobacteria. Applied and Environmental Microbiology 63,
33273332. O'Malley, M. A. (2007). The nineteenth century roots of'everything is everywhere'.
Nature Reviews Microbiology 5, 647651. Oliver, M. J. & Irwin, A. J. (2008). Objective global ocean biogeographic provinces. Geophys Res Lett 35, L15601. Palenik, B., Brahamsha, B., Larimer, F. W. & other authors (2003). The genome of a motile marine Synechococcus. Nature 424, 1037-1042.
50
Partensky, F., Hess, W. R. & Vaulot, D. (1999). Prochlorococcus, a marine photosynthetic prokaryote of global significance. Microbiology and Molecular Biology Reviews 63, 106-127. Pereira, P. R. G. (2010).Marine Bacteroidetes: distribution patterns and role in the degradation of organic matter. Pérez, V., Fernández, E., Marañón, E. & other authors (2005). Latitudinal distribution of microbial plankton abundance, production, and respiration in the Equatorial Atlantic in autumn 2000. Deep Sea Research Part I: Oceanographic
Research Papers 52, 861880. Pernthaler, A., Pernthaler, J. & Amann, R. (2004). Sensitive multi-color fluorescence in situ hybridization for the identification of environmental
microorganisms. Molecular microbial ecology manual 3, 711726. Philippot, L., Andersson, S. G. E., Battin, T. J., Prosser, J. I., Schimel, J. P., Whitman, W. B. & Hallin, S. (2010). The ecological coherence of high bacterial taxonomic ranks. Nat Rev Micro 8, 523-529. Quast, C., Pruesse, E., Yilmaz, P., Gerken, J., Schweer, T., Yarza, P., Peplies, J. & Glöckner, F. O. (2013a). The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Research 41, (D1):
D590D596. . Quast, C., Pruesse, E., Yilmaz, P., Gerken, J., Schweer, T., Yarza, P., Peplies, J. & Glöckner, F. O. (2013b). The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Research 41,
D590D596. Rappé, M. S., Connon, S. A., Vergin, K. L. & Giovannoni, S. J. (2002). Cultivation of the ubiquitous SAR 11 marine bacterioplankton clade. Nature 418, 630-633. Robinson, C., Poulton, A. J., Holligan, P. M. & other authors (2006). The Atlantic Meridional Transect (AMT) programme: a contextual view 1995–2005. Deep Sea
Research Part II: Topical Studies in Oceanography 53, 14851515. Roche (2010c). Method manual: EmPCR amplification Lib-A. . Roche (2010d). Method manual: Sequencing. . Schattenhofer, M., Fuchs, B. M., Amann, R., Zubkov, M. V., Tarran, G. A. & Pernthaler, J. (2009). Latitudinal distribution of prokaryotic picoplankton populations
in the Atlantic Ocean. Environmental Microbiology 11, 20782093. Schmitt, S., Tsai, P., Bell, J. & other authors (2012). Assessing the complex sponge microbiota: core, variable and species-specific bacterial communities in
marine sponges. ISME J 6, 564576.
51
Schochetman, G., Ou, C.-Y. & Jones, W. K. (1988). Polymerase chain reaction.
The Journal of infectious diseases, 11541157. Sergeant, M. J., Constantinidou, C., Cogan, T., Penn, C. W. & Pallen, M. J. (2012). High-Throughput Sequencing of 16S rRNA Gene Amplicons: Effects of Extraction Procedure, Primer Length and Annealing Temperature. PLoS ONE 7, e38094. Sogin, M. L., Morrison, H. G., Huber, J. A., Welch, D. M., Huse, S. M., Neal, P. R., Arrieta, J. M. & Herndl, G. J. (2006). Microbial diversity in the deep sea and the underexplored “rare biosphere”. Proceedings of the National Academy of Sciences
103, 1211512120. Swan, B. K., Martinez-Garcia, M., Preston, C. M. & other authors (2011). Potential for Chemolithoautotrophy Among Ubiquitous Bacteria Lineages in the Dark Ocean. Science 333, 1296-1300. Tada, Y., Taniguchi, A., Nagao, I., Miki, T., Uematsu, M., Tsuda, A. & Hamasaki, K. (2011). Differing growth responses of major phylogenetic groups of marine bacteria to natural phytoplankton blooms in the western North Pacific Ocean. Applied
and environmental microbiology 77, 40554065. Tamaki, H., Wright, C. L., Li, X., Lin, Q., Hwang, C., Wang, S., Thimmapuram, J., Kamagata, Y. & Liu, W.-T. (2011). Analysis of 16S rRNA Amplicon Sequencing Options on the Roche/454 Next-Generation Titanium Sequencing Platform. PLoS ONE 6, e25263. Teeling, H., Fuchs, B. M., Becher, D. & other authors (2012). Substrate-Controlled Succession of Marine Bacterioplankton Populations Induced by a
Phytoplankton Bloom. Science 336, 608611. Teira, E., Reinthaler, T., Pernthaler, A., Pernthaler, J. & Herndl, G. J. (2004). Combining catalyzed reporter deposition-fluorescence in situ hybridization and microautoradiography to detect substrate utilization by bacteria and archaea in the deep ocean. Applied and environmental microbiology 70, 4411-4414. Thiele, S., Fuchs, B. M., Ramaiah, N. & Amann, R. (2012). Microbial Community Response during the Iron Fertilization Experiment LOHAFEX. Applied and
Environmental Microbiology 78, 88038812. Wallner, G. n., Amann, R. & Beisker, W. (2005). Optimizing fluorescent in situ hybridization with rRNA―targeted oligonucleotide probes for flow cytometric identification of microorganisms. Cytometry 14, 136-143. Weinbauer, M. G., Bonilla-Findji, O., Chan, A. M., Dolan, J. R., Short, S. M., Šimek, K., Wilhelm, S. W. & Suttle, C. A. (2011). Synechococcus growth in the ocean may depend on the lysis of heterotrophic bacteria. Journal of Plankton Research 33, 1465-1476.
52
Weisse, R. & Storch, H. (2010). Climate and climate variability. In Marine Climate and Climate Change, pp. 1-25: Springer Berlin Heidelberg. West, N. J., Schönhuber, W. A., Fuller, N. J., Amann, R. I., Rippka, R., Post, A. F. & Scanlan, D. J. (2001). Closely related Prochlorococcus genotypes show remarkably different depth distributions in two oceanic regions as revealed by in situ hybridization using 16S rRNA-targeted oligonucleotides. Microbiology 147, 1731-1744. Wetz, M. S., Hales, B. & Wheeler, P. A. (2008). Degradation of phytoplankton-derived organic matter: Implications for carbon and nitrogen biogeochemistry in
coastal ecosystems. Estuarine, Coastal and Shelf Science 77, 422432. Whittaker, R. H., Levin, S. A. & Root, R. B. (1973). Niche, habitat, and ecotope.
American Naturalist, 321338. Whitton, B. A. & Potts, M. (2012). Introduction to the cyanobacteria. Ecology of Cyanobacteria II, 1-13. Woese, C. R. (1987). Bacterial evolution. Microbiological reviews 51, 221. Yooseph, S., Nealson, K. H., Rusch, D. B. & other authors (2010). Genomic and functional adaptation in surface ocean planktonic prokaryotes. Nature 468, 60-66. Zubkov, M. V., Fuchs, B. M., Archer, S. D., Kiene, R. P., Amann, R. & Burkill, P. H. (2001). Linking the composition of bacterioplankton to rapid turnover of dissolved dimethylsulphoniopropionate in an algal bloom in the North Sea. Environmental microbiology 3, 304-311. Zuckerkandl, E. & Pauling, L. (1965). Molecules as documents of evolutionary
history. Journal of Theoretical Biology 8, 357366. Zwirglmaier, K., Jardillier, L., Ostrowski, M. & other authors (2008). Global phylogeography of marine Synechococcus and Prochlorococcus reveals a distinct partitioning of lineages among oceanic biomes. Environmental Microbiology 10, 147–161.
Websites www.oceancolor.gstc.nansa.gov
www.odv.awi.de
www.sigmaplot.co.uk
http://www.arb-silva.de/documentation/background/release-108
http://www.bioinformatics.org/cd-hit
http://www.arb-silva.de/projects/ssu-ref-nr/
http://folk.uio.no/ohammer/past
53
Appendices
Apendix 1.
Corresponding NaCl concentrations in wash buffer for formamide concentration in hybridisation buffer.
% formamide in hybridization
buffer
[NaCl] in Mol μl 5 M NaCl
0 0.900 8900
5 0.636 6260
10 0.450 4400
15 0.318 3080
20 0.225 2150
25 0.159 1490
30 0.112 1020
35 0.080 700
40 0.056 460
45 0.040 300
50 0.028 180
55 0.020 100
60 0.014 40
Dye labels used in this study and their characteristic excitiation (+/- 10 nm)
Dye Excitiation (+/- 10 nm) Emission (+/-10nm)
DAPIb (for DNA
counterstaining)
358 350
Alexa 488 495 519
Alexa 594 590 617
54
Appendix 2.
Rarefraction analysis for each sample representing a biogeographical province.
55
56
57
Appendix 3.
58
Relative abundance of the dominant bacterioplankton groups (represented by the CARD-FISH hybridization with probes Eub338 I-III (bacteria), ARCH915 (Archaea), SAR11- MIX (SAR 11 clade), ROS537 (Roseobacter), SAR86-1245 (SAR 86 clade), CF319a (Bacteroidetes), SAR202-312R (SAR 202 clade), CYA664 (Cyanobacteria), SYN405 (Synechococcus); Pro405 (Prochlorococcus), SAR324-1412 (SAR324 clade). Abundance is shown for different biogeographical provinces (from 50 °N to 50°S, NADR, NAST, NATR, WTRA, SATL, SSTC) and across a vertical gradient (20m, DCM and 150m representing the surface, DCM and mesopelagic water column). Cell which did not hybridize with one of the applied group specific groups are indicated in green.