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Seventh Framework Programme
Theme 6 Environment FP7-ENV.2010.2.2.1-2
Global plankton data set building in view of modeling
Grant agreement for: Collaborative Project. Small- or medium scale focused research
project for specific cooperation actions (SICA) dedicated to
international cooperation partner countries
Project acronym:
GREENSEAS
Project title:
Development of global plankton data base and model
system for eco-climate early warning
Grant agreement no.
Start date of project:
Duration:
Project coordinator:
265294
01.01.11
36 months
Nansen Environmental and Remote Sensing Center, Bergen,
Norway
D3.3: Report on province-specific microbial stocks,
rates of group-specific production and trophic
interactions in the Atlantic Ocean
Due date of deliverable: 31.03.2013
Actual submission date: 22.04.2014
Organization name of lead contractor for this deliverable: NERC
Authors: Manuela Hartmann and Mikhail V. Zubkov
Project
co-funded
by
the
European
Commission
within the Seventh Framework Programme, Theme 6 Environment
Dissemination Level
PU
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X
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Restricted to a group specified by the consortium (including the Commission)
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ISSUE
DATE
CHANGE RECORDS
AUTHOR
0
1
2
17/09/2013
16/10/2013
27/03/2014
Report first draft
Report second draft
Report third draft
3
22/04/2014
Included DOI’s for datasets
M. Hartmann
M. Hartmann
M. Hartmann/
M. Zubkov
M. Hartmann
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SUMMARY
Report on province-specific microbial stocks, rates of group-specific production and trophic
interactions in the Atlantic Ocean.
In line with the GreenSeas deliverable 3.3 the advanced measurements of microbial stocks, growth
rates based on carbon fixation as well as the control of the bacterioplankton by small protist predators
are reported. Microbial stocks were measured by flow cytometry. The flow cytometric sorting was
combined with prior radiotracer labelling of microbial plankton samples to determine CO2 fixation,
amino acid and phosphate uptake rates of dominant microbial groups, which were phylogenetically
affiliated using molecular tools.
The collected data directly feeds into GreenSeas WP5 and 6 for indicator development and ecosystem
modelling, respectively. The report is organised in three chapters:
1) Province-specific microbial stocks and bacterial production
2) Group-specific carbon fixation rates
3) Trophic interactions
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GREENSEAS CONSORTIUM
Participant no.
Participant organisation name
Short name
Country
1 (Coordinator)
Nansen Environmental and Remote Sensing Center
NERSC
NO
2
3
4
5
6
7
Plymouth Marine Laboratory
PML
UNI Research
UNI Research
National Environment Research Council
NERC
Murmansk Marine Biological Institute
MMBI
Council for Scientific and Industrial Research
CSIR
University of Cape Town
UCT
Centro Euro-Mediterraneo per i Cambiamenti Climatici
CMCC
SCARL
Universidade Federal do Rio Grande
FURG
8
9
UK
NO
UK
RU
ZA
ZA
IT
BR
No part of this work may be reproduced or used in any form or by any means (graphic,
electronic, or mechanical including photocopying, recording, taping, or information storage and
retrieval systems) without the written permission of the copyright owner(s) in accordance with
the terms of the GREENSEAS Consortium Agreement (EC Grant Agreement 265294).
All rights reserved.
This document may change without notice.
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Table of Contents
Table of Contents ............................................................................................................................ 5
1
2
3
Province-specific microbial stocks ........................................................................................... 8
1.1
Introduction ..................................................................................................................... 8
1.2
Materials and Methods ................................................................................................... 8
1.3
Results and Discussion..................................................................................................... 8
1.4
Conclusion ..................................................................................................................... 12
1.5
References ..................................................................................................................... 12
Carbon fixation and growth rates of specific populations in Atlantic Ocean ........................ 14
2.1
Introduction ................................................................................................................... 14
2.2
Materials and Methods ................................................................................................. 14
2.3
Results and Discussion................................................................................................... 17
2.4
Conclusion ..................................................................................................................... 19
2.5
References ..................................................................................................................... 19
Trophic interactions - Mixotrophic basis of the Atlantic Ocean (Hartmann et al. 2012) ...... 22
3.1
Introduction ................................................................................................................... 22
3.2
Materials and Methods ................................................................................................. 22
3.3
Results ........................................................................................................................... 27
3.4
Discussion ...................................................................................................................... 31
3.5
Conclusion ..................................................................................................................... 31
3.6
References ..................................................................................................................... 32
List of Figures
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Figure 1-1. Microbial cell abundance along an Atlantic Meridional Transect cruise.
Populations are indicated by colors: low nucleic acid bacteria (grey),
Prochlorococcus (yellow), Synechococcus (red), plastidic eukaryotes (green) and
aplastidic eukaryotes (blue) ...................................................................................... 9
Figure 1-2. Contribution of Prochlorococcus cyanobacteria (yellow), Synechococcus
cyanobacteria (red) and small and large plastidic eukaryotes (light and dark green
respectively) to the microbial biomass along an Atlantic Meridional transect .......... 10
Figure 1-3. Example of the bioassay technique using 3H-labelled leucine to measure
microbial uptake rates, ambient concentration and turnover time ........................... 11
Figure 1-4. Microbial uptake rates (blue) and ambient concentrations of leucine (yellow)
along an Atlantic Meridional Transect. .................................................................... 12
Figure 2-1 Cruise track of the RRS James Cook on AMT-20, 2010. Different oceanic
regions are indicated. NG=Northern Subtropical Gyre, EQ=equatorial waters,
SG=Southern Subtropical Gyre, ST=Southern temperate waters ........................... 15
Figure 3-1 A schematic map of the Atlantic Ocean showing the area sampled in the
2007, 2008, 2009 and 2010 cruises ....................................................................... 23
Figure 3-2. Characteristic flow cytometric signatures of SYBR Green I - DNA stained
bacterioplankton (a-b) and smallest planktonic protists (c-f). The groups were
differentiated according to light scattering properties (90° or side light scatter, SSC),
relative concentration of SYBR Green I stain per particle (green fluorescence, FL1,
530±30nm), and chlorophyll content (red fluorescence, FL3, >650nm).
Bpl=Bacterioplankton, Plast-S=small, plastidic protists, Plast-L=large, plastidic
protists, Aplast=aplastidic protists .......................................................................... 26
Figure 3-3. A comparison of mean rates of cell bacterivory by the flow-sorted aplastidic
(Aplast), large plastidic (Plast-L) and small plastidic (Plast-S) protists in the five
Atlantic regions (NT=Northern temperate, NG=North subtropical gyre,
EQ=equatorial waters, SG=South subtropical gyre, ST=Southern temperate). The
numbers next to the region abbreviations indicate the year of sampling, and then the
numbers in brackets indicate the number of separate experiments performed in
each region. The rates were calculated using 35S-methionine pulse-chase tracing.
Error bars show single standard deviations to indicate the variance of rates within
regions. .................................................................................................................. 29
Figure 3-4 A comparison of mean absolute (a) and relative (b) population biomass and
mean absolute (c) and relative (d) population bacterivory of aplastidic (Aplast), large
plastidic (Plast-L) and small plastidic (Plast-S) protists in the five Atlantic regions
(NT=Northern temperate, NG=North subtropical gyre, EQ=equatorial waters,
SG=South subtropical gyre, ST=Southern temperate). The numbers next to the
region abbreviations indicate the year of sampling. The numbers in brackets
indicate the number of experiments done in each region. The rates were calculated
using 35S-methionine pulse-chase tracing. Error bars show single standard
deviations to indicate the variance of biomass and rates within regions. ................ 30
List of Tables
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Table 2-1 Regional differences of photosynthetic microbial communities in average per
cell CO2 fixation rates ............................................................................................. 18
Table 2-2. Regional differences of photosynthetic microbial communities of populationspecific CO2 fixation rates. ..................................................................................... 18
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Development of global plankton data base and model
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1 Province-specific
production rates
microbial
stocks
and
Ref: D.3.3
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bacterial
1.1 Introduction
At the core of the open Atlantic Ocean ecosystems are microbial communities, which are
numerically dominated by tiny cells with diameter less than 5 µm. The photosynthetic part of
these communities comprise two types of cyanobacteria belonging to the genera Synechococcus
and Prochlorococcus complemented by the smallest eukaryotic phytoplankton of the classes
Pelagophyceae, Chlorophyceae, Prymnesiophyceae and Prasinophyceae (Grob et al., 2011). The
rest of the microbial communities depend on the primary production of these phytoplankton,
particularly the numerically dominant heterotrophic bacteria of the SAR11 cluster
(Alphaproteobacteria), whose growth is controlled by the amount of dissolved organic
molecules released by phytoplankton. Therefore accurate separation of heterotrophic bacteria
from their phototrophic counterparts is crucial for adequate modelling of biogeochemical cycles
mediated by the oceanic microbial communities. Within the framework of the GreenSeas
project and the Atlantic Meridional Transect programme, such separation was achieved by flow
cytometry using cellular pigmentation, DNA content and size. The pronounced changes in
microbial community structure along the Atlantic Meridional Transect allowed to separate five
major oceanic provinces: Northern temperate waters (NT), Northern subtropical gyre (NG),
equatorial waters (EQ), Southern subtropical grey (SG) and the southern temperate waters (ST).
1.2 Materials and Methods
Determination of microbial cell numbers
Microbial populations were differentiated and enumerated using the FACSort or FACS Calibur
flow cytometer (Becton-Dickinson, UK). Prochlorococcus cyanobacteria (Pro) were enumerated
in unstained, live samples on the basis of chlorophyll pigmentation and size and in stained, fixed
samples using DNA content and size as defining parameters. Synechococcus cyanobacteria (Syn)
cell numbers were determined in unstained, live samples using phycoerythrin as a specific
marker. Small plastidic eukaryotes (2µm, Plast-S), large plastidic eukaryotes (3µm, Plast-L)
and large, aplastidic eukaryotes (3µm, Aplast) were enumerated
in stained,
paraformaldehyde-fixed samples. Briefly, subsamples were taken from a 20L carboy and fixed
with 1% v/v paraformaldehyde (PFA, final concentration, Sigma-Aldrich, Germany) for 1h in the
dark at room temperature and stained with SybrGreen I dye (Sigma-Aldrich, Germany )(Marie, et
al., 1997). As internal standard a mixture of 0.5m and 1.0m multi-fluorescent beads
(Polysciences, USA) at a defined concentration was added to the samples prior to analyses.
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Determination of uptake rates, turnover time and biologically available concentrations of
inorganic and organic substrates
Bioassays with radiolabelled leucine, methionine, inorganic and organic phoshorate were used
to determine bacterial production and uptake rates of phosphorus in the Atlantic Ocean. Briefly,
the samples were incubated with different concentrations of 3H-labelled leucine, 35S-labelled
methionine or 33P-labelled phosphoric acid (inorganic phosphate) or adenosine-tri-phosphate
(organic phosphate) and subsamples fixed with 1% w/v paraformaldehyde at certain time points
(10, 20, 30 and 40 min). Samples were then filtered on 0.2 µm polycarbonate filters to collect
microbial cells. Radioactivity of labelled substrates taken up by microbial cells was measured
using a liquid scintillation counter (TriCarb3100, PerkinElmer). For detailed description of
method please refer to Wright and Hobbie (1966) and Zubkov et al. (2007 and 2008).
1.3 Results and Discussion
The microbial communities of the open Atlantic Ocean are numerically dominated by
Prochlorococcus cyanobacteria and low nucleic acid containing bacteria (Figure 1-1).
Synechococcus cyanobacteria and small plastidic and aplastidic eukaryotes (< 3µm) represent
only a minor fraction of the total community (Figure 1-1).
Cell numbers, ml-1
107
106
105
104
103
102
-40
-20
0
20
40
Latitude, oN
Figure 1-1. Microbial cell abundance along an Atlantic Meridional Transect cruise. Populations are
indicated by colors: low nucleic acid containing bacteria (grey), Prochlorococcus (yellow), Synechococcus
(red), plastidic eukaryotes (green) and aplastidic eukaryotes (blue)
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Prochlorococcus cyanobacteria dominate the biomass of the phototrophic part of the microbial
communities in the oligotrophic regions while Synechococcus cyanobacteria and small plastidic
eukaryotes dominate in temperate waters (Figure 1-2).
Figure 1-2. Contribution of Prochlorococcus cyanobacteria (yellow), Synechococcus cyanobacteria (red)
and small and large plastidic eukaryotes (light and dark green respectively) to the microbial biomass along
an Atlantic Meridional transect
Biogeochemical cycling of organic and inorganic substrates within the bacterioplankton
populations were determined using a concentration dependent bioassay technique developed
first by Wright and Hobbie (1966). The bioassay method reveals bioavailable ambient
concentrations, microbial uptake rates and turnover times of the chosen substrates (Figure 1-3).
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Bioassay computation
40
4x103
3x103
Leu turnover time, h
0.05 nM
0.1 nM
0.2 nM
0.4 nM
0.6 nM
0.8 nM
5x103
2x103
1x103
30
20
10
3
H-Leupart, DPM per sample
Concentration series
0
0
10
20
Incubation time, min
30
-0.2
0.0
0.2
0.4
0.6
0.8
added Leu, nM
Figure 1-3. Example of the bioassay technique using 3H-labelled leucine to measure microbial uptake
rates, ambient concentration and turnover time
Comparison of leucine uptake rates in different regions showed that uptake rates are not
significantly different in the Northern and Southern subtropical gyres, while they are slightly
higher in the temperate and equatorial regions (see Figure 1-4 for an example and also Hill et al.
2011).
Especially in the North Atlantic gyre, where phosphate concentrations are particularly low, it is
important to understand phosphate dynamics. Using the bioassay technique in combination
with flow cytometrical cell sorting we determined that the major consumers of bio-available
phosphate were low nucleic acid bacteria and Prochlorococcus cyanobacteria, whilst
Synechococcus (7%) and picoeukaryotic (0.3%) phytoplankton played minor roles in direct
phosphate uptake.
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Figure 1-4. Microbial uptake rates (blue) and ambient concentrations of leucine (yellow) along an Atlantic
Meridional Transect.
1.4 Conclusion
The relatively uniform distribution of Prochlorococcus in the photic layer of the oligotrophic
Atlantic gyres suggests that these regions are balanced, i.e. consumer’s demand can be
sustained by phytoplankton via production of organic matter. Growth of heterotrophic bacteria
might be restricted in these regions due to the economical growth of Prochlorococcus releasing
only limited amounts of dissolved organic matter. Biomass dominance in conjunction with high
carbon fixation rates (see Section 2) and their unique photoheterotrophic life style make
Prochlorococcus a key player in the Atlantic Ocean ecosystems. Therefore we advise ecosystem
modellers to parameterise Prochlorococcus in their novel model developments.
As a result of our bioassay measurements, GreenSeas WP5 was advised that it is valid to use an
average microbial uptake rate for inorganic phosphorus or leucine for the Atlantic Ocean gyres
to parameterise the model since no significant regional differences were observed. Variability of
microbial uptake rates is much more pronounced in coastal and upwelling regions than in the
vast oligotrophic areas of the Atlantic Ocean (Hill et al., 2011).
The intriguingly low phosphate uptake rates of plastidic eukaryotes pose a question of how
these eukaryotes can meet their phosphate demands. We hypothesize that mixotrophy, i.e.
feeding on bacterial prey; can supply enough phosphate for growth of plastidic eukaryotes with
the additional advantage of reducing numbers of potential competitors such as Prochlorococcus
cyanobacteria (for details see Section 3).
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1.5 Publications
Journals
Hill PG, Mary I, Purdie DA, Zubkov MV (2011). Similarity in microbial amino acid uptake in
surface waters of the North and South Atlantic (sub-)tropical gyres. Prog. Oceanogr. 91: 437446.
Data series:
Holland R.J.; Zubkov M.V. (2014). Abundance and composition of microbial plankton
communities using flow cytometry on cruise AMT20 (JC053). British Oceanographic Data Centre
- Natural Environment Research Council, UK. doi:10/sc3.
Gomez-Pereira P.; Hartmann M.; Zubkov M.V. (2014). Ambient concentrations, microbial
turnover rates and turnover times of methionine, leucine and organic phosphate measured on
cruise AMT20 (JC053). British Oceanographic Data Centre - Natural Environment Research
Council, UK. doi:10/sc4.
1.6 References
Marie D, Partensky F, Jacquet S & Vaulot D (1997) Enumeration and cell cycle analysis of
natural populations of marine picoplankton by flow cytometry using the nucleic acid stain SYBR
Green I. Appl Environ Microbiol 63: 186-193.
Wright RT & Hobbie JE (1966) Use of glucose and acetate by bacteria and algae in aquatic
ecosystems. Ecol 47: 447-464.
Zubkov MV, Tarran GA, Mary I & Fuchs BM (2008) Differential microbial uptake of dissolved
amino acids and amino sugars in surface waters of the Atlantic Ocean. J Plankton Res 30: 211220.
Zubkov MV, Mary I, Woodward EMS, Warwick PE, Fuchs BM, Scanlan DJ & Burkill PH (2007)
Microbial control of phosphate in the nutrient-depleted North Atlantic subtropical gyre. Environ
Microbiol 9: 2079-2089.
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2 Carbon fixation and growth
populations in Atlantic Ocean
rates
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of
specific
2.1 Introduction
Oceans phytoplankton are only a minute fraction of the photosynthetic biomass on Earth, but
they contribute almost half of the primary production (45-50 billion tonnes vs. 52 billion tonnes,
marine vs. terrestrial systems). Prochlorococcus cyanobacteria (Pro) are the most abundant
photosynthetic organism on Earth occupying niches in the low latitude and oligotrophic marine
environments. Small plastidic eukaryotes are the other key primary producers in these
ecosystems, while Synechococcus cyanobacteria, preferring more nutrient replenished
environments, abundances are very low and hence their contribution to carbon fixation is
minute. Oligotrophic gyres cover >40% of the ocean surface and are predicted to expand
because of climatic changes (Polovina, et al., 2008), hence the global distribution of Pro might
change. Due to their high abundances it is generally assumed, that Pro is a major primary
producer in these systems, but due to difficulties with separating Pro, because of their low
pigment content, proof remains focused on the deep Pro (Chisholm, et al., 1988) or small areas
of the ocean (Li, 1994, Jardillier, et al., 2010). Hence, actual information on the contribution of
Pro to primary production is very limited.
Here, we provide for the first time in detail analyses of surface Prochlorococcus,
Synechococcocus and small plastidic eukaryotes CO2 fixation rates and their respective
contribution to primary production in the Atlantic subtropical gyres and adjacent regions. We
compare Prochlorococcus uptake rates to their phototrophic competitors. Moreover, we show
that flow cytometrical sorting of surface Pro according to pigmentation can skew CO 2 fixation
rates and suggest that DNA-content as a sorting parameter is more precise and reliable.
Furthermore, the data proves beyond doubt that Pro dominates primary production in the
surface layer of the low latitude Atlantic Ocean.
2.2 Materials and Methods
Sampling
20L sea water sample were taken from the pre-dawn CTD casts into an acid-rinsed
polycarbonate carboy on the AMT-20 cruise in 2010 (Atlantic Meridional Transect) on board the
UK RRS James Cook (Figure 2-1). To prevent exposure of cells to artificial light on board, the
carboy was covered completely with two layers of dark plastic. Samples were immediately
processed.
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Figure 2-1 Cruise track of the RRS James Cook on AMT-20, 2010. Different oceanic regions are indicated.
NG=Northern Subtropical Gyre, EQ=equatorial waters, SG=Southern Subtropical Gyre, ST=Southern
temperate waters
Catalysed reporter deposition fluorescence in situ hybridisations (CARD-FISH) on flow
cytometrically sorted cells
In order to confirm that the targeted bacterial population consists mainly of Pro cells, catalysed
reporter deposition fluorescence-in-situ-hybridisations were performed on sorted cells using the
Prochlorococcus-specific probe PRO405 (West, et al., 2001) at selected stations covering each
province (NG=Northern subtropical gyre, EQ=equatorial waters, SG=Southern subtropical gyre).
Flow cytometric cell sorting and CARD-FISH were carried out according to Gomez-Pereira et al.
(2013).
Total and group-specific CO2 fixation
Prior to each experiment, 60ml Pyrex glass bottles (Fisher Scientific, UK) were acid-soaked over
night (10% HCl) and rinsed twice with 30ml sample seawater. After washing, 60ml of seawater
sample were added to each bottle and spiked with tracemetal-clean 14C radiolabelled sodium
bicarbonate (DHI, Denmark). Samples were then incubated at ambient temperatures (regulated
by a refrigerated water bath (Grant Instruments, UK)) in a 6L water tank illuminated by a warm
white light-emitting diode array (Photon Systems Instruments, Czech Republic) adjusted to a
constant output of 500 µmol photons m-2 s-1. The chosen light intensity equals half the
irradiance reaching the water surface noon-time in the equatorial region. In contrast to
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incubations at ambient light, the constant light output makes it possible to compare different
stations.
Two different concentrations of 14C sodium bicarbonate were used in order to determine total
CO2 fixation during a time series and to measure CO2 fixation of flow cytometrically sorted
phytoplankton populations, respectively. Time series were carried out to ensure linear uptake of
label and to ensure that the small volumes of sorted cells are representative for the whole
community.
For time series CO2 fixation measurements, to six 60ml Pyrex glass bottles 3.7 kBq ml-1 14C
sodium bicarbonate was added and bottles were incubated for 2, 4, 6, 8, 10 h in the light, plus
one bottle incubated for 10h in the dark. At each discrete time point, the whole bottle was
sacrificed by adding 1% v/v PFA (final concentration). After incubation for 1h at room
temperature the complete sample was filtered on a 0.2 µm polycarbonate filter (Nuclepore,
Whatman, UK), washed three times with ultra-clean water (MQ system, Millipore, Whatman,
UK) and placed in a scintillation vial. Before addition of 5ml scintillation cocktail (Goldstar,
Meridian, UK), 1ml of 10% v/v HCl was added, the vial gently swirled and incubated for 1030min to fume out remaining unbound 14C sodium bicarbonate.
Due to the small size of the organisms higher 14C sodium bicarbonate concentrations were used
to determine group-specific CO2 fixation rates. To 60ml seawater sample in a Pyrex glass bottle
246kBq ml-1 of 14C sodium bicarbonate were added, the sample incubated for 10h and then fixed
with 1% PFA (final concentration). Three 1.6ml subsamples were taken directly to determine
total CO2 fixation and to sort stained Pro cells. 20ml of the sample were concentrated on a
0.6µm polycarbonate filter (Nuclepore, Whatman, UK) mounted in a filtration unit (Swynnex,
Millipore, UK) using a syringe pump (KD Scientific, USA) at a flow rate of 2.5ml min-1 in order to
concentrate and sort cyanobacteria. The remaining sample was concentrated on a 0.8µm
polycarbonate filter (Nuclepore, Whatman, UK) the same way to enrich eukaryotic
phytoplankton. Apart from the 0.6µm concentrated fraction all samples were stained with
SybrGreen I according to the protocol described above, stored at 4°C and sorted flow
cytometrically within 10h.
Flow cytometrical cell sorting
Different phytoplankton populations were sorted according to light-scattering properties (90° or
side light scatter), relative concentration of SYBR Green I stain per particle (green fluorescence;
FL1, 530±30nm), phycoerythrin-content (orange fluorescence, FL2 580±30nm) and chlorophyll
content (red fluorescence; FL3, >650nm) using a FACSort instrument (Becton Dickinson). Due to
their low pigmentation in surface waters we sorted Pro cells according to side scatter and FL1
properties from unconcentrated, SybrGreen I stained samples, where a distinct bacterial
population could be observed, which was verified to consist mainly of Pro by CARD-FISH
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analyses (Mary et al. (2008) and our own data), Syn cells were sorted according to phycoerythrin
content from 0.6µm concentrated samples. Eukaryotic Plast-S and Plast-L populations were
sorted from 0.8µm concentrated, stained samples using side scatter, SybrGreen I stain and
chlorophyll content as defining parameters. For each population 4-6 replicates of different cell
numbers were sorted. Bacterial and eukaryotic cells were collected on 0.2µm and 0.8µm
polycarbonate filters respectively and treated following the same procedure as for total CO2
fixation measurements (see above) before counting. Radio-assaying of samples was carried out
using an ultra-low-level liquid scintillation counter (1220 Quantulus, Wallac).
Data analyses
Cell specific carbon fixation rates were determined from average per cell values of each of the
sorted replicates and converted to fg C cell-1 h-1 according to Parsons et al. (1984). CO2 fixation
rates of different populations were calculated by multiplying the cell-specific CO2 fixation rates
with the group’s abundance. A conversion factor of 200 fg C µm-3 (Waterbury, et al., 1986) was
used to calculate biomass-specific CO2 fixation rates assuming spherical cell shape and average
cell diameters of 2.0 and 3.1µm for Plast-S and Plast-L cells respectively (Zubkov, et al., 2000,
Hartmann, et al., 2012). For Pro and Syn cells conversion factors of 220fg C µm-3 (Li, 1994) and
cell diameters of 0.52 and 0.95µm (Zubkov, et al., 1998, Zubkov, et al., 2000) were applied.
Statistical analyses were carried out using SigmaPlot 12.5, in case of normal distribution and
equal variance, t-tests were carried out for comparison. If the data was non-normally distributed
or the equal variance test failed, t-tests were replaced by Mann-Whitney Rank sum tests.
2.3 Results and Discussion
Prochlorococcus CO2 fixation efficiency in comparison to other phototrophic groups
Understanding primary production in an ecosystem necessitates knowledge of the different
groups contributing to it. Pro cyanobacteria, are a major part of the phototrophic microbial
community in the low latitude Atlantic Ocean, yet remain quite elusive to study due to their
small size and low chlorophyll content. Because of their extremely low chlorophyll content, it is
virtually impossible to confidently count or separate these cells in the upper part of the photic
zone based on cellular pigment alone (Chisholm, et al., 1988). It even became a modern
engineering challenge to enhance sensitivity of flow cytometers to unambiguously enumerate
Pro cells by autofluorescence (Olson, et al., 1990).
Average per cell CO2 fixation rates were measured for cyanobacterial (Pro and Syn) and
eukaryotic (Plast-S and Plast-L) organisms (Table 2-1). As expected, a strong positive correlation
of cell size and CO2 fixation was observed. Surprisingly, biomass normalised values did not follow
the expected negative correlation between cell volume and biomass-specific CO2 uptake. It is
true, that the larger eukaryotic organisms were significantly less efficient in CO2 uptake than
their cyanobacterial counterparts (Mann-Whitney, p<0.001). But within the groups biomass-
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specific CO2 fixation was actually positively correlated to cell size, i.e. Plast-S were less efficient
than Plast-L (t-test, p=0.01), whilst for the cyanobacteria Syn showed on average slightly higher
CO2 fixation than Pro (t-test, p=0.04). The addition of nutrients in the form of deep water (300m
depth) had no significant effect on the Pro or Syn CO2 fixation rates (t-test, p>0.5)
CO2 fixation
Plast-L
s.d.
Plast-S
s.d.
Pro
s.d.
Syn
s.d.
NG
63.13
11.06
12.88
1.72
0.37
0.02
4.95
1.71
EQ
94.94
21.45
15.84
3.89
0.85
0.21
6.67
1.18
SG
61.97
14.88
12.85
7.78
0.44
0.22
4.54
3.24
ST
59.46
41.66
7.50
4.21
0.50
0.24
2.24
2.16
[fg C cell-1 h-1]
Table 2-1 Regional differences of photosynthetic microbial communities in average per cell CO 2 fixation
rates
Pro CO2 uptake rates in the two gyres were similar, but more than doubled in the EQ (t-test,
p=0.002). Plast-L uptake rates followed the same pattern (t-test, p<0.001). In contrast, Plast-S as
well as Syn cells showed similar rates for NG, SG, EQ, but significantly lower carbon fixation in
the ST region (t-test, p=0.042 and p=0.032 respectively). See Table 2-1 for details.
Prochlorococcus contribution to community CO2 fixation
Despite their lower chlorophyll content Pro dominate CO2 fixation throughout the low latitude
Atlantic Ocean (on average 55±12%, Fig. 4), followed by Plast-L (29±9%), Syn (10±9%) and Plast-S
(6±6%) (Figure 2-2). Not surprisingly, gyre regions were less productive than EQ (t-test, p<0.001)
and ST (t-test, p<0.001) (Table 2-2). Although the NG and SG were in different seasons (boreal
autumn and austral spring) during the sampling period, CO2 fixation rates on the population
level were similar.
CO2 fixation
Plast-L
s.d.
Plast-S
s.d.
Pro
s.d.
Syn
s.d.
NG
0.45
0.11
0.04
0.01
0.54
0.10
0.13
0.15
EQ
1.11
0.29
0.10
0.02
2.94
0.54
0.51
0.14
SG
0.57
0.24
0.16
0.15
1.30
0.95
0.12
0.08
ST
1.92
1.29
1.05
0.61
0.65
0.01
2.93
2.51
[mg C m-3 d-1]
Table 2-2. Regional differences of photosynthetic microbial communities of population-specific CO2 fixation
rates.
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Our results show that the combination of high biomass and CO2 fixation rates ensures the
dominance of Pro contribution to primary production (Figure 2-). The presented measurements
of average CO2 fixation rates by these smallest phytoplankton cells are within a range of values,
reported in literature for Pro living in deeper habitats (Chisholm, et al., 1988, Li, 1994) or less
oligotrophic habitats (Jardillier, et al., 2010). The seminal study in the Sargasso Sea reported that
Pro contributed 15-60% to total primary production at the bottom of the photic layer (Chisholm,
et al., 1988). Pro contribution of 60% in the EQ Atlantic Ocean (Figure 2-2) favourably compared
to their estimated contribution of up to 48% of total primary production in the Equatorial Pacific
(Vaulot, et al., 1995).
Figure 2-2. Relative contribution of different microbial groups to CO2 fixation (a) and biomass (b) in the
Atlantic Ocean (Prochlorococcus (Pro), Synechococcus (Syn) and smaller and larger, plastidic eukaryotes
(<2µm, Plast-S and 2-3µm, Plast-L).
2.4 Conclusion
Despite their low chlorophyll content, Pro cyanobacteria are key players of primary production
in the surface layer of the low latitude Atlantic Ocean contributing more than 55% of the total
CO2 fixation. Based on satellite radiometer data Longhurst et al. (1995) estimated that
oligotrophic regions could contribute >30% to the total marine primary production.
Conservatively, Pro contribution could be then translated to 16% of the marine primary
production or 8% of Earth’s total primary production, a significant contribution for the smallest
phototrophic organism. Moreover, keeping in mind that the oligotrophic regions might expand
due to climate change Pro might intrude into new areas. These findings indicate that
Prochlorococcus should be specially taken into account when photosynthetic pigmentation data
is used for deducing biological CO2 fixation in the oligotrophic open ocean. Hence, it is of utmost
importance to fully understand the constraints and demands of these tiny organisms.
2.5 Publications
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Journal
Hartmann, M., Gomez-Pereira, P., Grob, C., Ostrowski, M., Scanlan, D.J., Zubkov, M.V. (2014).
Efficient CO2 fixation by surface Prochlorococcus in the Atlantic Ocean. ISME J. Accepted
Data series
Hartmann M.; Gomez-Pereira P.; Grob C.; Zubkov M.V. (2014). CO2 fixation of dominant
bacterial and eukaryotic groups for cruise AMT20 (JC053). British Oceanographic Data Centre Natural Environment Research Council, UK. doi:10/sc5.
2.6 References
Chisholm SW, Olson RJ, Zettler ER, Goericke R, Waterbury JB & Welschmeyer NA (1988) A
novel free-living prochlorophyte abundant in the oceanic euphotic zone. Nature 334: 340-343.
Gomez-Pereira PR, Hartmann M, Grob C, et al. (2013) Comparable light stimulation of organic
nutrient uptake by SAR11 and Prochlorococcus in the North Atlantic subtropical gyre. ISME J 7:
603-614.
Hartmann M, Grob C, Tarran GA, Martin AP, Burkill PH, Scanlan DJ & Zubkov MV (2012)
Mixotrophic basis of Atlantic oligotrophic ecosystems. Proc Nat Acad Sci USA 109: 5756-5760.
Jardillier L, Zubkov MV, Pearman J & Scanlan DJ (2010) Significant CO2 fixation by small
prymnesiophytes in the subtropical and tropical northeast Atlantic Ocean. ISME J 4: 1180-1192.
Li WKW (1994) Primary production of prochlorophytes, cyanobacteria, and eukaryotic
ultraphytoplankton - measurements from flow cytometric sorting. Limnol Oceano 39: 169-175.
Mary I, Tarran GA, Warwick PE, Terry MJ, Scanlan DJ, Burkill PH & Zubkov MV (2008) Light
enhanced amino acid uptake by dominant bacterioplankton groups in surface waters of the
Atlantic Ocean. FEMS Microbiol Ecol 63: 36-45.
Olson RJ, Chisholm SW, Zettler ER, Altabet MA & Dusenberry JA (1990) Spatial and temporal
distributions of prochlorophyte picoplankton in the North Atlantic Ocean. Deep-Sea Res Part aOceano Res Papers 37: 1033-1051.
Parsons TR, Maita Y & Lalli CM (1984) A manual of chemical and biological methods for
seawater analysis. New York: Pergamon Press.
Polovina JJ, Howell EA & Abecassis M (2008) Ocean's least productive waters are expanding.
Geophys Res Lett 35.
Vaulot D, Marie D, Olson RJ & Chisholm SW (1995) Growth of Prochlorococcus, a
photosynthetic prokaryote, in the equatorial Pacific Ocean. Science 268: 1480-1482.
Waterbury JB, Watson SW, Valois FW & Franks DG (1986) Biological and ecological
characterization of the marine unicellular cyanobacterium Synechococcus Photosynthetic
Picoplankton, (Platt T & Li W, eds.), Can Bull Fish Aquat Sci 214: 71-120.
West NJ, Schonhuber WA, Fuller NJ, Amann RI, Rippka R, Post AF & Scanlan DJ (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. Microbiol
147: 1731-1744.
Zubkov MV, Sleigh MA, Burkill PH & Leakey RJG (2000) Picoplankton community structure on
the Atlantic Meridional Transect: a comparison between seasons. Prog Oceano 45: 369-386.
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Zubkov MV, Sleigh MA, Tarran GA, Burkill PH & Leakey RJG (1998) Picoplanktonic community
structure on an Atlantic transect from 50°N to 50°S. Deep Sea Res Part I: Oceano Res Papers 45:
1339-1355.
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3 Trophic interactions - Mixotrophic basis of the Atlantic
Ocean
3.1 Introduction
Prochlorococcus cyanobacteria and the SAR11 group are the most numerous bacterial microbes
in the North Atlantic and South Atlantic subtropical gyres (Section 1), whereas small plastidic
protists, comprising various taxonomic groups (e.g. Prymnesiophyceae and Chrysophyceae)
(Cuvelier, et al., 2008, Lepere, et al., 2009, Jardillier, et al., 2010), are the most abundant among
the eukaryotes, dominating over their aplastidic counterparts (Zubkov, et al., 2007). Current
knowledge about the functioning of these microbe-controlled systems is relatively limited, due
to the difficulty of directly studying microbes in an environment typified by low concentrations
of organic and inorganic macronutrients. Our current understanding of oligotrophic ecosystem
functioning suggests that the roles of different microbial populations are tightly defined. In the
established paradigm (Azam, et al., 1983) for these systems, phytoplankton such as
cyanobacteria and plastidic protists harvest light, fix CO2, and take up inorganic nutrients. These
primary producers fuel the entire system, allowing heterotrophic bacterioplankton to thrive. In
turn, populations of phototrophic and heterotrophic bacterial populations are controlled by
viruses and aplastidic protist predators. Organic matter and inorganic nutrients, released by
these control processes, in addition to cell death and bacterioplankton remineralisation of
dissolved organic matter, sustain heterotrophic bacterioplankton and phytoplankton. However,
more recent observations are at variance with this paradigm. Despite their low phosphate
uptake, plastidic protists are major contributors to CO2 fixation (Section 2, Li, 1994, Jardillier, et
al., 2010). Consequently, the C:P ratio, calculated using CO2 and phosphate uptake rates by
plastidic protists, is unrealistically high, suggesting that osmotrophic uptake of phosphate
cannot satisfy growth requirements. We hypothesize that plastidic protists compensate for the
lack of inorganic nutrients by mixotrophy: They gain energy from sunlight and simultaneously
prey on bacterioplankton to acquire inorganic nutrients, such as phosphate and amino acids.
Here we show that plastidic protists prey on bacterioplankton in the surface mixed layer of both
oligotrophic subtropical gyres and adjoining low-latitude pelagic regions of the Atlantic Ocean
(40°N to 40°S). Owing to their high abundance, plastidic protists prevail over aplastidic protists
in bacterivory. This evidence suggests that mixotrophy is crucial to sustain the functioning of
oligotrophic marine ecosystems.
3.2 Materials and Methods
Sampling
This data, comprising 68 experiments, was collected in the Atlantic Ocean during one AMT
cruise on board the UK RRS James Clark Ross in October–November 2008 and on two AMT
cruises on board the UK RRS James Cook in October–November 2009 and 2010 (Figure 3-1).
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Seawater samples were collected from a depth of 20 m before dawn with 20-L Niskin bottles
mounted on a sampling rosette of a conductivity-temperature-depth profiler (Sea-Bird
Electronics). In 2008 and 2009, experiments were set up within 20 min of sample collection in
the dark at ambient temperature, controlled by a water bath to maintain temperature within
0.5 °C. In 2010, experiments were set up for dark and light measurements in a dark room using
only a dim green light (LEE filter 090; transmission of 20–30% of light at 500–550 nm). Light
incubation experiments were placed in a 6-L water tank illuminated by a warm white lightemitting diode (LED) array (Photon Systems Instruments, Czech Republic). Parallel dark
incubations were done in a similar water tank but were isolated from light. Both tanks were
plumbed into a refrigerated bath (Grant Instruments, UK) to maintain temperature within 0.5 °C
of in situ temperature at the depth of sample collection. The LED array was adjusted to keep
light intensity at 300μmol photons m−2·s−1 inside the incubation bottles.
Figure 3-1 A schematic map of the Atlantic Ocean showing the area sampled in the 2007, 2008, 2009 and
2010 cruises
Cell counting
Bacterioplankton and protist cell concentrations were determined by flow cytometry (Figure
3-2) using FACSort and FACSCalibur instruments (Becton Dickinson). Samples were fixed with
paraformaldehyde (PFA) 1% (wt/vol) final concentration and stained with SYBR Green I DNA dye
(Marie, et al., 1997). A mixture of multifluorescent beads, of diameter 0.5μm and 1.0μm
(Polysciences, USA), was used as an internal standard for fluorescence and flow rates. To
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compare protist population biomass, protist concentrations were multiplied by the
corresponding cell biomass values. To estimate their cell sizes, the three groups of protists were
flow-sorted, sorted cells being deposited on polycarbonate membrane filters with 0.2μm pore
diameter. Filters were mounted onto glass slides and stained excessively with DAPI (final
concentration 1μg ml−1) to reveal cell cytoplasm. Aplast cells were sorted and sized from four
experimental samples on the 2009 cruise and from five experimental samples on the 2010
cruise, which represented all regions studied. Plast-L and Plast-S cells were sorted from five and
four experimental samples on the 2010 cruise, respectively. At least 200 cells were measured at
1,000× magnification of an epifluorescence microscope (Axioscope 2; Zeiss, Germany) to
estimate mean cell diameters. Mean cell diameters of each of the three protist groups were
statistically similar in analyzed samples. The overall mean size of Plast-S cells of 2.0 ± 0.1μm was
significantly lower (t test, P = 0.0002) than the overall mean for Plast-L cells of 3.1 ± 0.3μm,
whereas the overall mean size of Aplast cells of 2.9 ± 0.3μm was statistically similar to the size of
Plast-L cells. To estimate the biomass of the three protist groups, their cell biovolume was
computed assuming that the cells were spheres with a diameter equal to the mean cell size. Cell
biovolume was converted into cell biomass using a specific carbon content of 200fg C·μm−3,
taken as a mean value from Christian and Karl (1994).
Determining rates of protist bacterivory using pulse–chase dual labelling of natural communities
Before an experiment, glass bottles (250mL, Schott; Fisher Scientific, UK) were soaked in 10%
hydrochloric acid and rinsed twice with 50mL of sampled seawater (taken from the same Niskin
bottle as that for the subsequent experiment). Seawater (250mL) from the sample was
subsequently added to each washed glass bottle and spiked with L-[35S]methionine (specific
activity >37TBq mmol-1; Hartmann Analytic, Germany), final concentration 0.25nM or 0.4nM,
and L-[4,5-3H]leucine (specific activity 1.48–2.22TBq mmol-1; Hartmann Analytic, Germany), final
concentration 0.5nM. After 1.5h incubation, nonradioactive methionine and leucine were added
to final concentrations of 0.25μM, or 0.4μM and 0.5μM, respectively, to chase the radioactive
amino acids (Zubkov & Sleigh, 1995, Zubkov & Tarran, 2008). Samples were incubated for 1.5h
to stabilize pulses in bacterioplankton cells before taking subsamples (120mL; fixed with 1%
(w/v) PFA at 3h and 9h) for the measurement of protist tracer uptake rates. After 1h of fixation,
particulate material was collected onto 0.2-μm polycarbonate filters (Nuclepore; Whatman, UK)
to measure the total sample radioactivity.
Flow cytometric cell sorting
For each time point, four different populations (total bacterioplankton (Bpl), Plast-S, Plast-L,
and Aplast protists) were sorted (Figure 3-2). For each population, four to eight replicates of
different cell numbers were sorted (Zubkov & Tarran, 2008). Sorted bacterioplankton cells were
collected onto 0.2μm polycarbonate filters. Sorted protist cells were collected onto 0.8μm
polycarbonate filters to reduce the retention of potentially by-sorted Bpl cells. Filters were
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washed with deionized water to remove contaminating tracer and placed into scintillation vials
to which 5mL of scintillation mixture (GoldStar, Meridian, UK) was added. Subsequently, the
vials were radioassayed for 0.5–2h (depending on sample radioactivity) using an ultra–low-level
liquid scintillation counter (1220 Quantalus; Wallac, UK).
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Figure 3-2. Characteristic flow cytometric signatures of SYBR Green I - DNA stained bacterioplankton (a-b)
and smallest planktonic protists (c-f). The groups were differentiated according to light scattering
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properties (90° or side light scatter, SSC), relative concentration of SYBR Green I stain per particle (green
fluorescence, FL1, 530±30nm), and chlorophyll content (red fluorescence, FL3, >650nm).
Bpl=Bacterioplankton, Plast-S=small, plastidic protists, Plast-L=large, plastidic protists, Aplast=aplastidic
protists
Data Analyses
Using quench curves, the 3H label was deconvoluted from the 35S label, and the amount of each
label was computed as Bq cell−1 by dividing the cumulative 3H or 35S radioactivity by the
corresponding number of sorted cells. Cell bacterivory was calculated according to the following
formula:
Bacterivory = (Prtavg T2 × Bplavg T2 − 1 − Prtavg T1 × Bplavg T1 − 1) × (T2 − T1) − 1 ;
where PrtavgT2 is the average activity of four to eight replicates of one of the protist groups at
the second time point and PrtavgT1 is the same at the first time point; T1 and T2 are the first and
second time points of the experiment (e.g., 3h and 9h, respectively); BplavgT1 and BplavgT2 are
the average activity of four to eight replicates of the bacterioplankton cells at T1 and T2,
respectively. Because of the pulse–chase experimental design, the activity of the Bpl was in
most cases the same at T1 and T2 and a cumulative mean could be used. To verify that the
increase in label between the first and second time points was statistically significant, t tests (P <
0.05) were carried out using SigmaPlot version 11.0 (Systat Software) and Quattro-Pro X4 (Corel)
software. Errors were calculated according to SE propagation procedures. The majority of
experiments (80%) showed a significant difference in protist radioactivity between the two time
points and hence demonstrated bacterivorous activity of the Plast-S, Plast-L, and Aplast cells.
We attribute nonsignificant bacterivory in some experiments to the detection limit of our
method owing to the low radioactivities measured. All estimates of rates of cell bacterivory
were included in calculations of average regional rates. The mean rates were all significantly
higher than zero (t tests, P < 0.05). To calculate regional bacterivory of the protist populations,
bacterivory per cell was multiplied by the corresponding mean concentration of protist cells
ml−1. T-tests were used to compare mean values; SDs, derived from pooled variance, are used to
show variability within regions.
3.3 Results
Protist bacterivory was assessed on three Atlantic Meridional Transect (AMT) research cruises in
October–November 2008, 2009, and 2010 encompassing subtropical oligotrophic gyres of the
Northern and Southern hemispheres and the equatorial convergence area (Fig. 1). Temperate
waters adjoining the Southern gyre were also examined. The results of an earlier study
conducted in 2007 in North Atlantic temperate waters (Zubkov & Tarran, 2008) are included for
comparison. Three populations of the smallest planktonic protists were examined: plastidic
protists (i.e., chloroplast-containing) smaller, 2μm (Plast-S) and larger, 3μm (Plast-L) as well as
aplastidic (without chloroplast) protists, 3μm (Aplast). In the majority of experiments, in all
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regions studied, tracer content of protist cells increased with time during the chase phase in
contrast to stable or slightly decreased tracer content of bacterioplankton cells. The increase
demonstrates bacterivory by the three types of protist cells. Moreover, the influence of light
and dark incubation on bacterivory was studied on the cruise in 2010. No statistically significant
light-induced differences (t-test, P > 0.1) in protist bacterivory were determined. The rates of
cell bacterivory (i.e., the number of bacterioplankton cells consumed per protist cell h−1) by the
Aplast protists were comparable in all regions apart from the Northern temperate (NT) region in
summer (Figure 3-3). The difference between the Southern temperate (ST) and the NT regions
was probably seasonal. Bacterivory by the Plast-L cells was the lowest in the Southern
subtropical gyre (SG) in 2008 and 2010 but was comparable to bacterivory by Aplast cells in the
ST region. Bacterivory by the Plast-S cells was lowest in temperate waters and in the SG in 2010.
It was similar to bacterivory by the Plast-L cells in the NG in 2009 and in the SG in 2008, but
lower in the SG in 2009 and 2010. Rates of bacterivory in the SG varied interannually. Cell
bacterivory by all three types of protists was significantly higher in 2009 compared with 2008
and 2010, whereas the differences between 2008 and 2010 were insignificant (Figure 3-3). On
the other hand, the concentration/biomass of the Plast-S population and the concentration of
bacterioplankton were comparable between the 3 years (Figure 3-4), whereas the
concentration/biomass of Aplast and the Plast-L protists was higher in 2010. Bacterioplankton,
Synechococcus, and Prochlorococcus concentrations in the surface mixed layer of the two gyres
were similar in 2009, whereas Plast-L and Aplast biomass was lower in the SG than in the NG,
and the opposite was true for the Plast-S protists (Figure 3-3). The biomass of Plast-L protists
was highest in all regions, followed by the biomass of Aplast and Plast-S protists (Figure 3-4). The
combined biomass of the two plastidic protist groups made up between 65% and 90% of the
combined biomass of the smallest protists (Figure 3-4). In contrast to cell bacterivory,
population bacterivory (i.e., the total number of bacterioplankton consumed ml−1 h−1 by each
protist population) in the NG and NT regions was not significantly different between the three
protist populations (Figure 3-4). Population bacterivory was significantly higher in more
productive temperate regions followed by the equatorial region, due to higher protist
concentrations (Figure 3-4). In the equatorial waters (EQ), and particularly in the SG in 2009,
bacterivory by the Plast-L population was the highest compared with other populations,
comprising 50% of total protist bacterivory (Figure 3-4). A comparison between the sum of
plastidic populations and the aplastidic population showed a significant difference in population
bacterivory by plastidic and aplastidic protists (t-test, P=0.01). Cumulative bacterivory by
plastidic protists accounted for 60–77% of total protist bacterivory across the Atlantic Ocean.
Furthermore, interannual variability had a minor effect on the domination of bacterivory by
plastidic protists.
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Figure 3-3. A comparison of mean rates of cell bacterivory by the flow-sorted aplastidic (Aplast),
large plastidic (Plast-L) and small plastidic (Plast-S) protists in the five Atlantic regions
(NT=Northern temperate, NG=North subtropical gyre, EQ=equatorial waters, SG=South
subtropical gyre, ST=Southern temperate). The numbers next to the region abbreviations
indicate the year of sampling, and then the numbers in brackets indicate the number of
separate experiments performed in each region. The rates were calculated using 35S-methionine
pulse-chase tracing. Error bars show single standard deviations to indicate the variance of rates
within regions.
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Figure 3-4 A comparison of mean absolute (a) and relative (b) population biomass and mean absolute (c)
and relative (d) population bacterivory of aplastidic (Aplast), large plastidic (Plast-L) and small plastidic
(Plast-S) protists in the five Atlantic regions (NT=Northern temperate, NG=North subtropical gyre,
EQ=equatorial waters, SG=South subtropical gyre, ST=Southern temperate). The numbers next to the
region abbreviations indicate the year of sampling. The numbers in brackets indicate the number of
experiments done in each region. The rates were calculated using 35S-methionine pulse-chase tracing.
Error bars show single standard deviations to indicate the variance of biomass and rates within regions.
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3.4 Discussion
The uniformly higher population rates of bacterivory by plastidic protists compared with
aplastidic protists in the surface mixed layer of the Northern and Southern gyres and the
equatorial region show the large contribution of phytoplankton to harvesting bacterioplankton
in the low-latitude Atlantic Ocean. There are several important implications of this finding. First,
it challenges the long-standing assumption of the total dependence of phytoplankton on
dissolved inorganic nutrients in oligotrophic oceanic waters. Rates of bacterivory by Plast-S cells
were lower in surface waters of the South Atlantic subtropical gyre compared with the North
Atlantic subtropical gyre, which is depleted in phosphate. This suggests that the mixotrophy of
Plast-S cells may be linked to phosphate depletion. However, differences linked to the
seasonality of studies in the gyres (boreal autumn, austral spring) as well as interannual
variability may also play a role here. In contrast, rates of bacterivory by Plast-L protists were
comparable along the whole transect in 2009. Rates of bacterivory by Plast-L cells similar to
those found in 2009 have also been measured in the temperate North Atlantic Ocean (Zubkov &
Tarran, 2008), further supporting the lack of correlation between macronutrient availability and
bacterivory in Plast-L cells. The main temporal variability in Plast-L bacterivory was interannual
in the SG. Both biomass and rates of bacterivory by Plast-L and Aplast populations are broadly
comparable between the phosphate-depleted Northern and relatively phosphate-replete
Southern Atlantic oligotrophic gyres (Figure 3-4). The second implication of our work is related
to the cell metabolism of mixotrophs. Because CO2 fixation as well as predation and respiration
are concomitantly taking place in the same cells, mixotrophy could help to explain the tightness
of biogenic carbon budgets at the community level (Williams, 1998). Tight intercellular coupling
of production and respiration could contribute to the stability of oligotrophic ecosystems in the
absence of seasonal or episodic perturbations (Karl, et al., 2003) such as seasonally accumulated
bioavailable organic matter (Thingstad, et al., 2005), or allochthonous matter transported by
advection (Roussenov, et al., 2006) or deposited from the atmosphere (Dachs, et al., 2005, Calil,
et al., 2011), which enhance growth of opportunistic species and ultimately change the
composition of microbial communities. The third implication concerns the ecological significance
of the smallest plastidic protists in oligotrophic ecosystems. Apart from being key CO2 fixers,
plastidic protists control bacterioplankton abundance, acting as producers of organic matter and
predators at the same time. Such dual control and interdependence of bacterioplankton and
protists could help to explain the constancy of low bacterioplankton concentrations in the
oligotrophic ocean compared with more productive regions (Li, et al., 2004, Zubkov, et al.,
2008). The scarcity of bacterioplankton prey in oligotrophic gyres in turn probably reduces both
propagation of phage infections and growth of specialized predators such as aplastidic protists.
3.5 Conclusion
Oligotrophic subtropical gyres are the largest oceanic ecosystems, covering >40% of the Earth’s
surface. Unicellular Prochlorococcus cyanobacteria and the smallest algae (plastidic protists)
dominate CO2 fixation in these ecosystems, competing for dissolved inorganic nutrients. Here
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we present direct evidence from the surface mixed layer of the subtropical gyres and adjacent
equatorial and temperate regions of the Atlantic Ocean, collected on three Atlantic Meridional
Transect cruises on consecutive years, that bacterioplankton are fed on by plastidic and
aplastidic protists at comparable rates. Rates of bacterivory were similar in the light and dark.
Furthermore, because of their higher abundance, it is the plastidic protists, rather than the
aplastidic forms, that control bacterivory in these waters. In summary, our data shows the
significance and ubiquity of mixotrophy in the survival of the smallest pelagic protists in sunlit
oligotrophic surface waters. This deceptively inefficient lifestyle should reduce nutrient export
and maintain faster nutrient turnover in the surface mixed layer, both of which are essential for
sustainable functioning of oligotrophic ecosystems. Consequently, future food web models
should consider including mixotrophs as a basic ecosystem element.
3.6 Publications
Journal
Hartmann M, Grob C, Tarran GA, Martin AP, Burkill PH, Scanlan DJ, Zubkov MV (2012).
Mixotrophic basis of Atlantic oligotrophic ecosystems. Proc Nat. Acad Sci USA 109: 5756-5760
Hartmann M, Zubkov MV, Scanlan DJ, Lepère C (2013). In situ interactions between
photosynthetic picoeukaryotes and bacterioplankton in the Atlantic Ocean: evidence for
mixotrophy. Environ Microbiol Rep 5: 835-840.
Data series
Hartmann M.; Grob C.; Zubkov M.V. (2014). Influence of light on bacterivory of the smallest
eukaryotes on cruise AMT20 (JC053). British Oceanographic Data Centre - Natural Environment
Research Council, UK. doi:10/sc6.
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Hartmann M, Grob C, Scanlan DJ, Martin AP, Burkill PH & Zubkov MV (2011) Comparison of
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Microbiol 9: 2079-2089.
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Development of global plankton data base and model
system for eco-climate early warning
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Date: 05/05/2017
Issue: 0.1
END OF DOCUMENT
Grob C, Hartmann M, Zubkov MV, Scanlan DJ (2011). Invariable biomass-specific primary
production of taxonomically discrete picoeukaryote groups across the Atlantic Ocean.
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Hill PG, Mary I, Purdie DA, Zubkov MV (2011). Similarity in microbial amino acid uptake in
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