Several foraminifera are deposit feeders that consume organic
detritus (dead particulate organic material with entrained bacteria).
However, the role of such foraminifera in the benthic food web remains
understudied. Foraminifera feeding on methanotrophic bacteria, which are
13C-depleted, may cause negative cytoplasmic and/or calcitic δ13C values. To test whether the foraminiferal diet includes
methanotrophs, we performed a short-term (20 h) feeding experiment with
Nonionellina labradorica from an active Arctic methane-emission site (Storfjordrenna, Barents Sea)
using the marine methanotroph Methyloprofundus sedimenti and analysed N. labradorica cytology via transmission
electron microscopy (TEM). We hypothesised that M. sedimenti would be visible post-experiment in degradation vacuoles, as evidenced by their ultrastructure.
Sediment grains (mostly clay) occurred inside one or several degradation
vacuoles in all foraminifers. In 24 % of the specimens from the feeding
experiment degradation vacuoles also contained bacteria, although none could
be confirmed to be the offered M. sedimenti. Observations of the apertural area after
20 h incubation revealed three putative methanotrophs, close to clay
particles, based on bacterial ultrastructural characteristics. Furthermore,
we noted the absence of bacterial endobionts in all examined N. labradorica but confirmed
the presence of kleptoplasts, which were often partially degraded. In sum,
we suggest that M. sedimenti can be consumed via untargeted grazing in seeps and that N. labradorica
can be generally classified as a deposit feeder at this Arctic site.
Introduction
In methane seep sites, the upward migration of methane affects the
pore-water chemistry of near-surface sediments, where benthic foraminifera
live (e.g. Dessandier et al., 2019). Extremely light
isotopic signals of δ13C have been measured in seep-associated
foraminiferal calcite tests (Wefer et al., 1994; Rathburn et al., 2003;
Hill et al., 2004b; Panieri et al., 2014). Studies specifically looking at
living (rose bengal stained) foraminiferal tests support the hypothesis that
the carbon isotopic composition is strongly influenced by the porewater dissolved inorganic carbon (DIC)
(McCorkle et al., 1990). Interspecific δ13C differences
between species with similar depth indicate sometimes taxon-specific
“vital” effects (McCorkle et al., 1990). Those “vital” effects describe
the biology of the different species, which could reflect different feeding
patterns. It has been suggested that Nonionella auris is an indicator of methane release and
possibly ingests 13C-depleted methane-oxidising bacteria
(Wefer et al., 1994). Recently, Melonis barleeanus (Williamson, 1858)
collected from an active methane seep site was found to be closely
associated with putative methanotrophs (Bernhard and Panieri,
2018), providing impetus to examine feeding habits of foraminifera living in
or around methane seeps.
Methanotrophs produce the biomarker diplopterol, which has an extremely
light δ13C signature (-60 ‰)
(Hinrichs et al., 2003). Our hypothesis is that if foraminifera
ingest methanotrophs, δ13C values of foraminiferal cytoplasm
should be altered by their diet. Experiments using a high-pressure culturing
system revealed the difficulty of measuring the sensitive relationship between
methane exposure and the foraminifera Cibicides wuellerstorfi. However, it was shown in one
experiment using entire cores that a methane source was reflected in δ13C of foraminiferal calcite (Wollenburg et al., 2015). It is
also not yet conclusive if diet can influence foraminiferal calcite as new
calcite did not form during experiments (Mojtahid et
al., 2011).
Another hypothesis to explain extremely light δ13C values
recorded in benthic foraminiferal calcite is that foraminifera assimilate
carbon as 13C-depleted methane-derived DIC, which would lead to
extremely light δ13C values. The possibility that
13C-depleted DIC from the pore water can be assimilated by foraminifera
is currently debated. Some studies suggest it is not possible
(Herguera et al., 2014), while others assert the feasibility
that foraminifera calcify close to seeps (Rathburn et al., 2003; Hill et
al., 2004a; Panieri et al., 2014). The problem lies in the calcite tests
and the difficulty of assessing the time of death of these protists in the
sediment. Several studies found that the lightest isotopic δ13C
values were measured in tests coated with methane-derived authigenic carbonate
(MDAC) overgrowth, which happens after the death of the foraminifer
(Torres et al., 2010; Panieri et al., 2014; Consolaro et al., 2015;
Panieri et al., 2017; Schneider et al., 2017). However, light δ13C values remain in many tests after MDACs are removed
(Panieri et al., 2014) and have been measured also in primary
calcite, without MDACs, from tests in methane-rich environments
(e.g. Mackensen, 2008; Dessandier et al., 2019). These observations again
point to the role of food influencing the cytoplasmic δ13C.
Foraminifera play an important role in the carbon cycle on the deep seafloor
(Nomaki et al., 2005) where feeding behaviour and food
preference vary with species (Nomaki et al., 2006). Selected
species of deep-sea benthic foraminifera have been shown to feed selectively
on 13C-labelled algae from sedimentary organic matter but unselectively
on 13C-labelled bacteria of the strain Vibrio (Nomaki et al.,
2006). A study from the seafloor around Adriatic seeps suggested that
δ13C of foraminiferal cytoplasm could be influenced by feeding
on the sulfur-oxidising bacterium Beggiatoa, whose abundance was also positively
correlated with foraminiferal densities (Panieri, 2006). Benthic foraminifera have a variety of different trophic strategies and consume many kinds of foods. There are deposit feeders, exclusively herbivorous or carnivorous species, and suspension feeders ingesting dissolved organic matter (DOM) (reviewed in Lipps, 1983). Deposit feeders are omnivorous,
gathering fine-grained sediment (e.g. clay) and associated bacteria,
organic detritus (dead particulate organic material), and, if present, diatom
cells using their pseudopodia. Based on the ultrastructure of the diet found
in vacuoles several species of foraminifera from different habitats have
already been classified to be deposit feeders (Goldstein and
Corliss, 1994).
Here we investigate if Nonionellina labradorica would feed in a short-term feeding experiment on the
marine methanotroph Methyloprofundus sedimenti and compare its ultrastructure on experimental specimens
and field specimens. Nonionellina labradorica is an abundant species in the North Atlantic
(Cedhagen, 1991) and occurs together with N. digitata in Svalbard fjord
sediments (Hald and Korsun, 1997; Shetye et al., 2011; Fossile et al.,
2020). In addition to its wide distribution, it is an especially interesting
experimental species for feeding studies because it hosts kleptoplasts,
i.e. sequestered chloroplasts, of diatom origin inside its cytoplasm
(Cedhagen, 1991; Jauffrais et al., 2019b). Nonionellina labradorica's aperture shows a specific
ornamentation, possibly a morphological adaptation to this “predatory”
mode of life for obtaining the kleptoplasts (Bernhard and Bowser, 1999).
Denitrification has been speculated for N. labradorica (reviewed in Charrieau et al.,
2019) because the foraminiferal genus Nonionella can denitrify, which was
demonstrated on two species (Risgaard-Petersen et al., 2006; Choquel et
al., 2021) but not yet on N. labradorica. Our study analysed contents of the degradation
vacuoles of this species from an active methane-emitting site in the Arctic
(Storfjordrenna, Barents Sea) before and after a feeding experiment.
Materials and methodsSite description and sampling living foraminifera
The sampling site was located approx. 50 km south of Svalbard at 382 m water
depth at the mouth of Storfjordrenna (Serov et al., 2017). The
site is characterised by several large gas hydrate pingos (GHPs), which
actively vent methane over an area of 2.5 km2. Our samples were taken
at GHP3, which is referred to as an underwater gas-hydrate-bearing mound
(Hong et al., 2017, 2018). GHP3 is a ∼ 500 m
diameter and 10 m tall mound that actively vents methane (Fig. 1). Marine
sediment samples were collected during CAGE cruise 18-05 supported by the
research vessel Kronprins Haakon in October 2018 and sampled by the remotely operated
vehicle (ROV) Ægir. A blade corer (BLC18; surface dimensions
27×19 cm,
Fig. 1c) was used to retrieve marine sediment in the vicinity of bacterial
mats (GPS 76∘6′23.7′′ N, 15∘58′1.7′′ E). Once on board the
blade corer was immediately sampled to retrieve living (cytoplasm
containing) foraminifera using a small aquarium hose targeting the first centimetre
(∼ 0–1 cm). The sediment was collected in petri dishes and wet
sieved to a size range of 250–500 µm. The species N. labradorica, which was abundant
in that layer, was subsequently used for a feeding experiment described in
detail below.
Description of the sampling site Gas Hydrate Pingo 3 (GHP3), a
gas-hydrate-bearing mound, located in Storfjordrenna, Barents Sea. (a) Map
illustrating Svalbard archipelago and the sampling site, approx. 50 km
offshore. (b) Map of sampling site GHP3; active gas bubble release is marked
on the top of the underwater mount, the yellow star indicates location of push
corer PUC2 (geochemical analyses), and the black square indicates location of BLC18
(sediment source for experiment). (c) Underwater image of retrieval of BLC18
taken by ROV camera illustrating the colouration of sediment with the
sea floor visible in background.
Geochemistry of the study site
For geochemical analysis of the study site a push corer (PUC2; henceforth
referred to as geochemistry core) was taken to obtain measurements of
δ13CDIC and sulfate because the blade corer (BLC18) did not
allow those measurements. PUC2 was taken in close vicinity to BLC18,
∼ 5 m apart (see Fig. S1 in the Supplement). Pore-water samples were taken from
PUC2 using rhizons that were inserted through pre-drilled holes in the core
tube at intervals of 1 cm (Table S1). Acid-washed 20 mL syringes were
attached to the rhizons for pore-water collection. Depending on the amount
of pore water collected, the samples were split for δ13CDIC and sulfate measurements. To the samples, 10 µL of
saturated HgCl2 (aqueous) was added to stop microbial activity and
stored in cold conditions (5 ∘C). A Thermo Scientific GasBench II
coupled to a Thermo Scientific MAT 253 isotope-ratio mass spectrometer (IRMS) at the Stable Isotope Laboratory
(SIL) at CAGE, UiT The Arctic University of Norway, was used to determine δ13CDIC of the
pore water. Anhydrous phosphoric acid was added to small glass vials (volume
4.5 mL) that were closed and flushed with helium 5.0 gas before the
pore-water sub-sample was measured. A porewater sub-sample (volume 0.5 mL)
was then added through the septa with a syringe needle, followed by
equilibration for 24 h at 24 ∘C to liberate the CO2 gas.
Three solid calcite standards with a range of +2 to -49 ‰ were used for normalisation to δ13C Vienna Pee Dee Belemnite (VPDB).
Correction of measured δ13C by -0.1 ‰ was
done to account for fractionation between gas and aqueous in sample
vials. Instrument precision for δ13C on a MAT253 IRMS was ±0.1 ‰ (SD). Sulfate was measured with a Metrohm ion
chromatography instrument equipped with column Metrosep A Supp 4 and eluted
with 1.8 mmol L-1 Na2CO3+1.7 mmol L-1 NaHCO3 at the
University of Bergen.
Culturing of the marine methanotroph M. sedimenti
Methyloprofundus sedimenti PKF-14 had been previously isolated from a water-column sample collected at
Prins Karls Forland, Svalbard, in the laboratory at UiT in Tromsø.
Methyloprofundus sedimenti were cultured in 10 mL batches of a 35:65 mix of 1/10 nitrate mineral salt
medium (NMS) and sterile filtered seawater using 125 mL
Wheaton® serum bottles with butyl septa and
aluminium crimp caps (Teknolab®). Methane was
injected to give a headspace of 20 % methane in air, and the bottles were
incubated without shaking at 15 ∘C in darkness. Purity of the
cultures and cell integrity was verified by microscopy and by absence of
growth on agar plates with a general medium for heterotrophic bacteria
(tryptone, yeast extract, glucose, and agar).
Experimental set-up
On the ship, N. labradorica (Fig. 2a, b) specimens showing dark greenish-brown cytoplasm
were picked using sable artist brushes under a stereomicroscope immediately
after wet sieving the sediment using natural seawater delivered from the
ship pump. Living specimens had a partly inorganic covering surrounding the
test, which was gently removed using fine artist brushes. Those so-called
cysts are nothing unusual with many foraminiferan taxa (Heinz et al.,
2005).
Exemplary illustration of Nonionellina labradorica, utilised in
this study. (a) Reflected light microscopy image from a specimen directly
after sampling; the white arrowhead indicates aperture location. (b) Scanning
electron image from a specimen before molecular analysis was performed; the
white arrowhead indicates aperture location. (c) Transmission electron
microscopy image of a culture of Methyloprofundus sedimenti, the marine
methanotroph used in the feeding experiment. The characteristic features for
methanotroph identification are the typical type I intracytoplasmic membranes
(ICMs). Furthermore, other internal structures visible are storage granules
(SGs) and a gram-negative cell wall (GNCW).
Our specimens were subsequently rinsed twice in filtered artificial seawater
to remove any sediment before placing them into the experimental petri
dishes. Care was taken that those were minimally exposed to light during
preparation of the experiment as kleptoplasts are known to be highly light
sensitive in this foraminifer (Jauffrais et al., 2019b).
The experiment with M. sedimenti was conducted for a total duration of 20 h to resemble
previous experiments on N. labradorica using transmission electron microscopy (TEM) and
nanometre-scale secondary ion mass spectrometry isotopic imaging
(NanoSIMS) (Jauffrais et al., 2019b), and it included two more
time points at 4 and 8 h. A short pre-experimental phase (2–4 h) was
included before the start of the feeding experiment to allow specimens to
acclimate. During the pre-experimental phase specimens were not fed, and they
resided in the petri dishes to adjust to the experimental conditions. The
feeding experiment consisted of several small petri dishes (3.5 cm
∅, 3 mL) each containing five N. labradorica in artificial seawater (ASW) at ambient salinity 35 (Red
Sea Salt). Petri dishes were sealed with Parafilm® and covered with aluminium foil and placed inside the incubator in
complete darkness. Temperature inside the chamber was maintained at
2–3 ∘C, which is within the range of the site's bottom-water
temperature (-1.8–4.6 ∘C) (Hong et al., 2017).
The feeding of M. sedimenti was performed once at the beginning of the experiment by
adding 100 µL of culture to 3 mL of artificial seawater to produce a
final concentration of ∼1×106 bacteria mL-1 in each
petri dish. Previously conducted feeding studies were used as guides:
Muller and Lee (1969) used 1×l04 bacteria mL-1
seawater and Mojtahid et al. (2011) used 4×108 bacteria mL-1 seawater.
Five foraminifera, which served as initial/field specimens (Table 1), were
fixed without M. sedimenti incubation. The respective petri dishes were incubated for 4,
8, and 20 h to determine if incubation duration influenced response of the
foraminifera to the methanotroph. One petri dish containing five
foraminifera, which were un-fed and fixed at 20 h, served as a negative
“control”. After the end of the respective incubation times, each
foraminifer was picked with a sterilised fine artist brush, which was
cleaned in 70 % ethanol between each specimen, and placed individually
into a fixative solution (4 % glutaraldehyde and 2 % paraformaldehyde
dissolved in ASW).
Summary of TEM observations of Nonionellina labradorica comparing field specimens and
experimental specimens. Field specimens (initials) were not fed, and a
non-fed control was not preserved after a 20 h incubation. The only putative
methanotrophs were observed and imaged in specimens from the 20 h
incubation. Bacteria of unknown origin were described as rod-shaped cells in
the degradation vacuoles.
Duration ofFood providedSampleCytoplasm: degradation Aperture region:experiment(yes (x)/no)IDvacuole contents (putative) methanotrophs(h per field sample)BacteriaClay/inorganicsField samples (initials)noE1noxnonoE3noxnonoE5noxnonoE6noxno4xE25noxnoxE27xxnoxE28noxnoxE29noxno8xE14xxnoxE15noxnoxE16noxnoxE17noxno20xE36xx1xxE37xxnoxE38noxnoxE39nox2xControl (20)noE44noxnoTransmission electron microscopy (TEM) preparation
Samples of N. labradorica preserved in fixative solution were transported to the
University of Angers, where they were prepared for ultrastructural analysis
using established protocols (Lekieffre et al., 2018). Four
embedded foraminiferal cells per treatment were sectioned using an
ultramicrotome (Leica® Ultracut S) equipped with
a diamond knife (Diatome®, ultra 45∘).
Grids were stained using UranyLess® EM Stain
(EMS, USA). Ultra-thin sections (70 nm) were observed with a JEOL JEM-1400
TEM at the SCIAM facility, University of Angers.
To document the ultrastructure of M. sedimenti, a sub-sample of the culture used for
experiments was imaged with TEM (Fig. 2c). To do so, an exponentially
growing culture was collected, centrifuged, pre-fixed with 2.5 % (w/v)
glutaraldehyde in growth medium overnight, washed in PBS (phosphate buffered
saline), and then post-fixed with 1 % (w/v) aqueous osmium tetroxide for 1.5 h at room temperature. After dehydration in an ethanol series, the
samples were embedded in an EPON-equivalent (SERVA) epoxy resin. Ultra-thin
sections were cut on a Leica EM UC6 ultramicrotome and stained with 3 %
(w/v) aqueous uranyl acetate followed by staining with lead citrate
(Reynolds, 1963) at 20 ∘C for 4–5 min. The samples were examined
with a JEOL JEM-1010 transmission electron microscope at an accelerating
voltage of 80 kV with a Morada camera system at the Advanced Microscopy Core
Facility (AMCF), Faculty of Health Science, UiT The Arctic University of
Norway.
Foraminifera ultrastructural observation and image processing
Four specimens per experimental time point (initials, 4, 8, and 20 h) plus one
un-fed (control) specimen were examined with the TEM. From each specimen, a
minimum of 50 TEM images were taken, including images detailing the
degradation vacuoles (approx. 5–27 images per specimen). Before the
ultrastructure was examined in detail, an overview images was created of
each section to illustrate the number of chambers and size of the specimen.
Images were blended together using Photoshop CS5 (see Fig. 4–5a).
Thereafter, the ultrastructure was examined at different parts of the cell:
(a) in the interior to document vitality, (b) on degradation vacuoles to
determine their contents, and (c) at the exterior to survey for microbes
entrained in remnant “reticulopodial trunk” material. All images made
during the observations of the TEM sections are deposited at Zenodo (10.5281/zenodo.6941739; Schmidt et al., 2022).
Molecular genetics and morphology
DNA metabarcoding and morphological documentation were performed on 13
specimens of N. labradorica. Briefly, live specimens were dried on micropalaeontological
slides and transported in a small container, cooled with ice-pads, to the
University of Angers. All specimens were imaged for morphological analysis
using a scanning electron microscope (SEM; EVOLS10, ZEISS, Fig. S1) followed
by total DNA being individually extracted in dissolved organic carbon (DOC) buffer (Pawlowski, 2000).
To amplify foraminiferal DNA, a hot start polymerase chain reaction (PCR; 2 min at 95 ∘C)
was performed in a volume of 25 µL with 40 cycles of 30 s at
95 ∘C, 30 s at 50 ∘C, and 2 min at 72 ∘C,
followed by 10 min at 72 ∘C for final extension. Primers s14F3 and
sB were used for the first PCR and 30 cycles at an annealing temperature of
52 ∘C (other parameters unchanged) for the nested PCR with primers
s14F1 and J2 (Pawlowski, 2000; Darling et al., 2016). Positive
amplifications were sequenced directly with the Sanger method at Eurofins
Genomics (Cologne, Germany). For taxonomic identification, DNA sequences
were compared first with BLAST (basic local alignment search tool)
(Altschul et al., 1997) and then within an alignment
comprising other nonionids implemented in SeaView (Gouy et al.,
2010) and corrected manually.
ResultsSample description and geochemistry of the study site
The visual observation of the sediments within the blade corer BLC18
immediately after sampling (Fig. 1c) indicated that the sediment appeared
light grey to yellowish in the upper part until approx. 13 cm and dark brown
from approx. 13 cm to the bottom. The sulfate measured in the pore water of the
geochemistry core (PUC2) declined from ∼ 2750 ppm at the
sediment–water interface to ∼ 706 ppm at approximately 13 cm
(see Fig. S1, Table S1). A decline in sulfate concentration indicates that
the anaerobic oxidation of methane (AOM) occurred at approx. 13 cm depth. The
sulfate–methane transition zone (SMTZ) characterised by a DIC value of -32 ‰ at approx. 13 cm sediment depth can be considered
shallow based on the global average (Egger et al., 2018).
Ultrastructure of methanotroph culture used in the feeding experiment
Transmission electron microscopy was performed on culture aliquots to allow
morphological comparison to previously published work
(Tavormina et al., 2015). Methyloprofundus sedimenti strain PKF-14 cells are coccoid
to slightly elongated in shape and are characterised by typical type I stacked
intracytoplasmic membranes (ICMs) (Fig. 2c). It has storage granules (SGs) and
a gram-negative cell wall (GNCW), which are not uniquely characteristic of
methanotrophs (Fig. 2c). Additionally, 16S rRNA gene sequencing was
performed (data not shown) to confirm it to be similar to the published
M. sedimenti (Tavormina et al., 2015).
Foraminiferal ultrastructure from an Arctic seep environmentGeneral ultrastructure
All 17 specimens examined for ultrastructure were considered living at the
time of observation (Fig. 3) as the mitochondria had characteristic double
membranes and occasionally visible cristae (Nomaki et al., 2016).
Cytoplasm exhibited several vacuoles and kleptoplasts concentrated in the
youngest chambers (Fig. 3a), and, in some specimens, a nucleus with nucleoli
was visible (Fig. 3b). Kleptoplasts were numerous throughout the cytoplasm
and occurred in the form of a single chloroplast (Fig. 3a–b) or as double
chloroplasts (Fig. S2a–d). Not all kleptoplasts were intact; some showed
peripheral degradation of the membranes indicated by an increasing number of
white areas between pyrenoid, lamella, and thylakoids (Fig. S2a–d). The
mitochondria occurred often in small clusters of two to five throughout the
cytoplasm and were oval, round, or kidney-shaped in cross section (Fig. 3e–f). Peroxisomes in N. labradorica occurred mostly as pairs (Fig. 3c) or small clusters
of three to four spherical organelles (Fig. S3a). Sometimes, but not always,
peroxisomes were associated with endoplasmic reticulum (Fig. S3b) but could
also occur alone. Golgi apparatus (Fig. 3d) had intact membranes, often
occurring near mitochondria.
Transmission electron micrographs showing cellular ultrastructure
of N. labradorica. (a) Cytoplasm showing parts of two chambers, with nucleus
with nucleoli, vacuoles, and several kleptoplasts, (b) nuclear envelope,
nucleoli, and kleptoplasts, (c) peroxisomes and electron opaque bodies, (d)
Golgi, and (e–f) mitochondria. V = vacuole, c = kleptoplast, nu = nucleoli,
n = nucleus p = peroxisome, eo=electron opaque body, m = mitochondrion,
fv = fibrillar vesicle, li = lipid droplet. Scales: (a) 2 µm, (b) 1 µm, (c–f) 200 nm.
Ultrastructure of aperture-associated bacteria
In total, three putative methanotrophs were identified in the vicinity of
two specimens (sample E39, Fig. 4; E37, Fig. 5). These microbes were
identified adjacent to reticulopodial remains (Fig. 4b). As an aid for
identification of M. sedimenti we used the characteristics shown in the literature
(Tavormina et al., 2015) and our own TEM observation
obtained from M. sedimenti culture (Fig. 2c). As noted, M. sedimenti is characterised by a typical
type I intracytoplasmic stacked membrane (ISM). Other characteristics, which
are not specific for methanotrophs, included storage granules (SGs) and a
typical gram-negative cell wall (GNCW) (Fig. 2c). On specimen E39 from the
20 h treatment, we found the methanotroph exhibiting the clearest internal
structure, having both typical type I intracytoplasmic stacked membranes
(ISM) and SGs (Fig. 4c).
Transmission electron micrographs of N. labradorica from 20 h
treatment (sample E39). (a) Stitched cross section of TEM images showing
location of methanotroph at the aperture region (black rectangle is the
location of image shown in panel b). (b) Location of two putative
methanotrophs next to sediment particles and putative reticulopodial remains
(black rectangle is location of image shown in panel c). (c) Close-up of two
putative methanotrophs revealing detailed features for identification, such
as typical type I stacked intracytoplasmic membranes (ICMs), and other
characteristics, such as storage granules (SGs) and gram-negative cell wall
(GNCW). Scale bars: (a) 100 µm, (b) 1 µm, (c) 200 nm.
Transmission electron micrographs of N. labradorica from 20 h
treatment (sample E37). (a) Stitched cross section of TEM images showing
location of putative methanotroph (black rectangle) at the aperture region.
(b) Location of the putative methanotroph next to sediment particles and putative reticulopodial remains. (c) Close-up of putative
methanotroph showing several storage granules (SGs) throughout its cell, which is surrounded by a gram-negative cell wall (GNCW). Scale bars: (a) 100 µm, (b) 0.5 µm, (c) 200 nm.
Contents of degradation vacuoles
Digestive vacuoles and food vacuoles are often summarised as degradation
vacuoles in the literature (Lekieffre et al., 2018), and this
makes sense for our study as well. A degradation vacuole is a vacuole where
enzymatic activities degrade contents, often making them unidentifiable
(Bé et al., 1982; Hemleben et al., 1989).
Sediment particles were present in many degradation vacuoles. The sediment
grains were easy to recognise in the TEM image as angular grains inside the
vacuoles, next to organic debris, which can have many different shapes. Each
specimen had at least one degradation vacuole and mostly several, which were
filled with sediment particles (Table 1). If a sediment particle was
visible, the vacuole was defined as a degradation vacuole (dv), and if it
was not and empty, then it was defined as a standard vacuole (v) (Fig. 6).
The observed entrained sediment particles were platelets, likely clay from
the seafloor, and hence show that the vacuole must contain foreign objects,
around which degradation processes have started. A total of 4 of 17 specimens
examined (23 %) had one or more bacteria of various sizes inside their
degradation vacuoles next to sediment particles (Fig. 6c, f).
TEM micrographs of N. labradorica showing degradation vacuoles
containing miscellaneous items, including bacteria (b), inorganics (clay
platelets), and unidentifiable remains after 4 h incubation, which are shown
enlarged on the left side of the image in a zoomed-in window (a, b; specimens E27,
E28, respectively), after 8 h incubation (c, d; specimen E14), and after 20 h
incubation (e, f; specimens E36, E37, respectively). v = vacuole,
dv = degradation vacuole, c = kleptoplast, p = peroxisome,
m = mitochondrion, li = lipid, g = Golgi. Scales: (a, c–f) 1 µm, (b)
2 µm.
Foraminiferal genetics
A total of 6 of 13 specimens analysed for genetics were positively amplified and
sequenced (Fig. S4). The sequences are deposited in GenBank under the
accession numbers MN514777 to MN514782. When comparing them via BLAST, they
were between 98.6 % and 99.6 % identical to published sequences
belonging to foraminifera identified as the morphospecies N. labradorica from the
Skagerrak, Svalbard, and the White Sea (Holzmann and Pawlowski, 2017;
Jauffrais et al., 2019b). Sequences were also included in an alignment
comprising other nonionids implemented in SeaView (not shown) and corrected
manually to check the BLAST search. This step confirmed the BLAST
identification.
DiscussionSampling site and geochemistry
The sampling site of blade corer BLC18 was in close proximity
(∼ 50 m) to an active methane vent releasing methane bubbles
at the gas hydrate pingo (GHP3) (Serov et al., 2017). At such
sites with high methane fluxes, the SMTZ (sulfate–methane transition zone)
is shallow as sulfate in the sediment is readily consumed in the first tens
of centimetres (Barnes and Goldberg, 1976; Iversen and Jørgensen,
1993) by sulfate-reducing bacteria (SRB) (reviewed in
Carrier et al., 2020). Geochemical analysis of PUC2 revealed an SMTZ at approx.
13 cm, which is rather shallow (Egger et al., 2018) as it can
also be several metres deep in other sites (reviewed in
Panieri et al., 2017). Similar geochemical characteristics can be considered
at the sampling location of living specimens (BLC18) given the close
proximity of the two locations. The geochemical data at PUC2 allow us
conclude that the site where living foraminifera were collected can be
classified as an active methane emission site.
Possible association with putative methanotrophs
The possible association of Nonioninella labradorica with methanotrophs was documented via presence
of two putative methanotrophs, based on microbial ultrastructure
(Tavormina et al., 2015). The documentation of this
possible association with putative methanotrophs likely is due to the
feeding experiment. However, there is a small possibility that the
associated methanotrophs were field remains. Another benthic foraminifer,
Melonis barleeanus, has been noted to have clumps of putative methanotrophs at the apertural
opening of field-collected specimens (Bernhard and Panieri,
2018). However, the non-selective deposit-feeding behaviour of N. labradorica, which we
describe for this species for the first time, shows that methanotrophs may
be ingested via untargeted grazing.
Degradation vacuoles show a large number of sediment particles and few
bacteria
Our results of the feeding experiment show that 23 % of the examined N. labradorica
specimens contained bacteria inside their degradation vacuoles. That is not
a large proportion compared to the presence of sediment particles, which
occurred in 100 % of the examined foraminifers. From this result, however,
we infer that N. labradorica at this site is a deposit feeder, feeding on organic detritus
and associated bacteria. The bacteria observed in the degradation vacuoles
resembled those from other deep-sea foraminifera (Globobulimina pacifica and Uvigerina peregrina) and the
shallow-dwelling genus Ammonia (Goldstein and Corliss, 1994). Salt-marsh
foraminifera also feed on bacteria and detritus, as observed in TEM studies
(Frail-Gauthier et al., 2019). Scavenging on bacteria
has also been observed by other foraminifera from intertidal environments
such as Ammonia tepida or Haynesina germanica (Pascal et al., 2008) and is a logical
consequence of detritus feeding. Certain foraminifera have been shown to
selectively ingest algae/bacteria according to strain (Lee et al.,
1966; Lee and Muller, 1973). From laboratory cultures we know that several
foraminifera cultures require bacteria to reproduce, as antibiotics
inhibited reproduction (Muller and Lee, 1969). Future
studies will need to employ additional molecular tools to determine the food
contents inside the cytoplasm (e.g. Salonen et al., 2019). For
example, a recent study used metabarcoding to assess the contribution of
eukaryotic operational taxonomic units (OTUs) associated with intertidal foraminifera, revealing that
Ammonia sp. T6 preys on metazoans, whereas Elphidium sp. S5 and Haynesina sp. S16 were more likely to
ingest diatoms (Chronopoulou et al., 2019).
General ultrastructure of N. labradorica collected in a seep environment
Our observations also included the intact nature of all major organelle
types of this species as this was essential to conclude vitality after the
experiment (Nomaki et al., 2016). Mitochondria and kleptoplasts were
generally homogeneously distributed throughout the cytoplasm confirming
previous observations of six N. labradorica from the Gullmar Fjord (Lekieffre et al.,
2018; Jauffrais et al., 2019b). If mitochondria are concentrated
predominately under pore plugs, it can be an indicator that the electron
acceptor oxygen is scarce in their environment as the pores are the direct
connection from the cell to the environment. This has been observed in
several other studies in which mitochondria were accumulated under pores in N. stella
(Leutenegger and Hansen, 1979) and Bolivina pacifica (Bernhard et
al., 2010).
Even though our study did not focus on kleptoplasts, we could observe that
kleptoplasts were occasionally degraded, which could have happened (a) during sampling, (b) due to exposure to microscope lights, or (c) due to the
age and condition of kleptoplasts inside the host. Kleptoplasts in N. labradorica have
been studied in detail describing their diatom origin (Cedhagen,
1991), sensitivity to light, and missing photosynthetic functionality
(Jauffrais et al., 2019b).
Conclusions
Based on the content of degradation vacuoles, we conclude that N. labradorica from our
study site, an active methane emitting site in the Barents Sea, is a
deposit feeder. It ingests large amounts of sediment particles, together with
bacteria. For two specimens of the feeding experiment, putative methanotrophs
were observed near the N. labradorica aperture, suggesting ingestion of M. sedimenti via “untargeted
grazing”. Further studies are needed on feeding strategies of other
palaeo-oceanographically relevant foraminifera to detangle the relationship
between δ13C of foraminiferal calcite, their cytoplasm, and their
dietary composition.
Data availability
Datasets containing TEM images are downloadable at Zenodo (10.5281/zenodo.6941739, Schmidt et al., 2022). Molecular sequence data are deposited at GenBank
under the accession numbers MN514777 to MN514782.
Sample availability
Samples are available upon request, and TEM thin sections are archived at the University of Angers.
The supplement related to this article is available online at: https://doi.org/10.5194/bg-19-3897-2022-supplement.
Author contributions
GP, EG, and CS designed the project and experiment; CS and EG collected samples; CS performed experiments; CS and HR prepared samples; CS, JMB, EG, and CL conducted TEM observations and interpretations; MS conducted molecular genetics; CS, GP, JMB, and EG wrote the paper; CL, MMS, MS, and HR provided critical review and comments on the manuscript; MMS, MS, and CL contributed reagents, materials, and analysis tools.
Competing interests
The contact author has declared that none of the authors has any competing interests.
Disclaimer
Publisher’s note: Copernicus Publications remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Acknowledgements
We thank the captains, crew members, and scientists of R/V Kronprins Haakon and ROV Ægir Team, Anne-Grethe Hestnes for culturing the methanotroph, Florence Manero, Romain Mallet, and Rodolphe Perrot (SCIAM microscopy facility, Univ Angers) for their TEM and SEM expertise, Sunil Vadakkepuliyambatta for assistance with mapping (Fig. 1), and Sophie Quinchard (LPG) for molecular analyses. Joan M. Bernhard was partially supported by US NSF 1634469, WHOI's Investment in Science Program, and the Région Pays de la Loire through the FRESCO Project.
Financial support
This research has been supported by the French scientific programme MOPGA (Make our Planet Great Again) managed by the National Research Agency; the Norwegian Research Council through the Centre for Arctic Gas Hydrate, Environment and Climate (project number 223259); NORCRUST (project number 255250); and by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) – 444059848.
The publication of this article was funded by the Open Access Fund of the Leibniz Association.
Review statement
This paper was edited by Aninda Mazumdar and reviewed by Jutta Wollenburg and two anonymous referees.
ReferencesAltschul, S. F., Madden, T. L., Schäffer, A. A., Zhang, J., Zhang, Z.,
Miller, W., and Lipman, D. J.: Gapped BLAST and PSI-BLAST: a new generation
of protein database search programs, Nucl. Acid. Res., 25, 3389–3402,
10.1093/nar/25.17.3389, 1997.Barnes, R. O. and Goldberg, E. D.: Methane production and consumption in
anoxic marine sediments, Geology, 4, 297–300, 10.1130/0091-7613(1976)4<297:MPACIA>2.0.CO;2, 1976.Bé, A. W. H., Spero, H. J., and Anderson, O. R.: Effects of symbiont
elimination and reinfection on the life processes of the planktonic
foraminifer Globigerinoides sacculifer, Mar. Biol., 70, 73–86,
10.1007/BF00397298, 1982.Bernhard, J. M. and Bowser, S. S.: Benthic foraminifera of dysoxic
sediments: chloroplast sequestration and functional morphology, Earth-Sci.
Rev., 46, 149–165, 10.1016/s0012-8252(99)00017-3, 1999.Bernhard, J. M. and Panieri, G.: Keystone Arctic paleoceanographic proxy
association with putative methanotrophic bacteria, Sci. Rep.-Uk, 8, 10610,
10.1038/s41598-018-28871-3, 2018.Bernhard, J. M., Goldstein, S. T., and Bowser, S. S.: An ectobiont-bearing
foraminiferan, Bolivina pacifica, that inhabits microxic pore waters:
cell-biological and paleoceanographic insights, Environ. Microbiol.,
12, 2107–2119, 10.1111/j.1462-2920.2009.02073.x, 2010.
Carrier, V., Svenning, M. M., Gründger, F., Niemann, H., Dessandier,
P.-A., Panieri, G., and Kalenitchenko, D.: The Impact of Methane on
Microbial Communities at Marine Arctic Gas Hydrate Bearing Sediment,
Front. Microbiol., 24, 11, 10.3389/fmicb.2020.01932, 2020.Cedhagen, T.: Retention of chloroplasts and bathymetric distribution in the
sublittoral foraminiferan Nonionellina labradorica, Ophelia, 33, 17–30,
10.1080/00785326.1991.10429739, 1991.Charrieau, L. M., Ljung, K., Schenk, F., Daewel, U., Kritzberg, E., and Filipsson, H. L.: Rapid environmental responses to climate-induced hydrographic changes in the Baltic Sea entrance, Biogeosciences, 16, 3835–3852, 10.5194/bg-16-3835-2019, 2019.Choquel, C., Geslin, E., Metzger, E., Filipsson, H. L., Risgaard-Petersen, N., Launeau, P., Giraud, M., Jauffrais, T., Jesus, B., and Mouret, A.: Denitrification by benthic foraminifera and their contribution to N-loss from a fjord environment, Biogeosciences, 18, 327–341, 10.5194/bg-18-327-2021, 2021.Chronopoulou, P.-M., Salonen, I., Bird, C., Reichart, G.-J., and Koho, K.
A.: Metabarcoding insights into the trophic behavior and identity of
intertidal benthic foraminifera, Front. Microbiol., 10, 1169,
10.3389/fmicb.2019.01169, 2019.Consolaro, C., Rasmussen, T. L., Panieri, G., Mienert, J., Bünz, S., and Sztybor, K.: Carbon isotope (Δ13C) excursions suggest times of major methane release during the last 14 kyr in Fram Strait, the deep-water gateway to the Arctic, Clim. Past, 11, 669–685, 10.5194/cp-11-669-2015, 2015.Darling, K. F., Schweizer, M., Knudsen, K. L., Evans, K. M., Bird, C.,
Roberts, A., Filipsson, H. L., Kim, J.-H., Gudmundsson, G., Wade, C. M.,
Sayer, M. D. J., and Austin, W. E. N.: The genetic diversity, phylogeography
and morphology of Elphidiidae (Foraminifera) in the Northeast Atlantic, Mar.
Micropaleontol., 129, 1–23, 10.1016/j.marmicro.2016.09.001, 2016.Dessandier, P.-A., Borrelli, C., Kalenitchenko, D., and Panieri, G.: Benthic
Foraminifera in Arctic Methane Hydrate Bearing Sediments, Front.
Mar. Sci., 6, 765, 10.3389/fmars.2019.00765,
2019.Egger, M., Riedinger, N., Mogollón, J. M., and Jørgensen, B. B.:
Global diffusive fluxes of methane in marine sediments, Nat. Geosci.,
11, 421–425, 10.1038/s41561-018-0122-8, 2018.Fossile, E., Nardelli, M. P., Jouini, A., Lansard, B., Pusceddu, A., Moccia, D., Michel, E., Péron, O., Howa, H., and Mojtahid, M.: Benthic foraminifera as tracers of brine production in the Storfjorden “sea ice factory”, Biogeosciences, 17, 1933–1953, 10.5194/bg-17-1933-2020, 2020.Frail-Gauthier, J. L., Mudie, P. J., Simpson, A. G. B., and Scott, D. B.:
Mesocosm and Microcosm Experiments On the Feeding of Temperate Salt Marsh
Foraminifera, J. Foraminifer. Res., 49, 259–274, 10.2113/gsjfr.49.3.259, 2019.Goldstein, S. T. and Corliss, B. H.: Deposit feeding in selected deep-sea
and shallow-water benthic foraminifera, Deep Sea Res. Pt. I, 41, 229–241, 10.1016/0967-0637(94)90001-9, 1994.
Gouy, M., Guindon, S., and Gascuel, O.: SeaView version 4: a multiplatform
graphical user interface for sequence alignment and phylogenetic tree
building, Mol. Biol. Evol., 27, 221–224, 10.1093/molbev/msp259, 2010.Hald, M. and Korsun, S.: Distribution of modern benthic foraminifera from
fjords of Svalbard, European Arctic, J. Foramin. Res.,
27, 101–122, 10.2113/gsjfr.27.2.101, 1997.
Heinz, P., Geslin, E., and Hemleben, C.: Laboratory observations of benthic
foraminiferal cysts, Mar. Biol. Res., 1, 149–159, 2005.Hemleben, C., Spindler, M., and Anderson, O. R.: Modern planktonic foraminifera , Springer, Berlin, 363 pp., 10.1007/978-1-4612-3544-6, 1989.Herguera, J. C., Paull, C. K., Perez, E., Ussler Iii, W., and Peltzer, E.:
Limits to the sensitivity of living benthic foraminifera to pore water
carbon isotope anomalies in methane vent environments, Paleoceanography, 29,
273–289, 10.1002/2013PA002457, 2014.Hinrichs, K.-U., Hmelo, L. R., and Sylva, S. P.: Molecular fossil record of
elevated methane levels in late Pleistocene coastal waters, Science, 299,
1214–1217, 10.1126/science.1079601, 2003.Hill, R., Schreiber, U., Gademann, R., Larkum, A. W. D., Kuhl, M., and
Ralph, P. J.: Spatial heterogeneity of photosynthesis and the effect of
temperature-induced bleaching conditions in three species of corals, Mar. Biol., 144, 633–640, 10.1007/s00227-003-1226-1, 2004a.Hill, T. M., Kennett, J. P., and Valentine, D. L.: Isotopic evidence for the
incorporation of methane-derived carbon into foraminifera from modern
methane seeps, Hydrate Ridge, Northeast Pacific, Geochim. Cosmochim.
Acta, 68, 4619–4627, 10.1016/j.gca.2004.07.012,
2004b.Holzmann, M. and Pawlowski, J.: An updated classification of rotaliid
foraminifera based on ribosomal DNA phylogeny, Mar. Micropaleontol., 132,
18–34, 10.1016/j.marmicro.2017.04.002, 2017.Hong, W.-L., Torres, M. E., Carroll, J., Crémière, A., Panieri, G.,
Yao, H., and Serov, P.: Seepage from an arctic shallow marine gas hydrate
reservoir is insensitive to momentary ocean warming, Nat. Commun., 8, 15745,
10.1038/ncomms15745, 2017.Hong, W. L., Torres, M. E., Portnov, A., Waage, M., Haley, B., and Lepland,
A.: Variations in gas and water pulses at an Arctic seep: fluid sources and
methane transport, Geophys. Res. Lett., 45, 4153–4162, 10.1029/2018GL077309, 2018.Iversen, N. and Jørgensen, B. B.: Diffusion coefficients of sulfate and
methane in marine sediments: Influence of porosity, Geochim.
Cosmochim. Acta, 57, 571–578,
10.1016/0016-7037(93)90368-7, 1993.Jauffrais, T., LeKieffre, C., Schweizer, M., Geslin, E., Metzger, E.,
Bernhard, J. M., Jesus, B., Filipsson, H. L., Maire, O., and Meibom, A.:
Kleptoplastidic benthic foraminifera from aphotic habitats: insights into
assimilation of inorganic C, N and S studied with sub-cellular resolution,
Environ. Microbiol., 21, 125–141, 10.1111/1462-2920.14433, 2019b.
Lee, J. J. and Muller, W. A.: Trophic dynamics and niches of salt marsh
foraminifera, Am. Zool., 13, 215–223, 1973.
Lee, J. J., McEnery, M., Pierce, S., Freudenthal, H., and Muller, W.: Tracer
experiments in feeding littoral foraminifera, J. Protozool.,
13, 659–670, 1966.LeKieffre, C., Bernhard, J. M., Mabilleau, G., Filipsson, H. L., Meibom, A.,
and Geslin, E.: An overview of cellular ultrastructure in benthic
foraminifera: New observations of rotalid species in the context of existing
literature, Mar. Micropaleontol., 138, 12–32, 10.1016/j.marmicro.2017.10.005, 2018.Leutenegger, S. and Hansen, H. J.: Ultrastructural and radiotracer studies
of pore function in foraminifera, Mar. Biol., 54, 11–16,
10.1007/BF00387046, 1979.Lipps, J. H.: Biotic Interactions in Benthic Foraminifera, in: Biotic
Interactions in Recent and Fossil Benthic Communities, edited by: Tevesz, M.
J. S. and McCall, P. L., Springer US, Boston, MA, 331–376,
10.1007/978-1-4757-0740-3_8, 1983.Mackensen, A.: On the use of benthic foraminiferal δ13C in
palaeoceanography: constraints from primary proxy relationships, Geological
Society, London, Special Publications, 303, 121–133, 10.1144/SP303.9, 2008.McCorkle, D., Keigwin, L., Corliss, B. H., and Emerson, S. R.: The influence of microhabitats on the carbon isotopic composition of deep-sea benthic foraminifera, Paleoceanography, 5, 161–185, 10.1029/PA005i002p00161, 1990.Mojtahid, M., Zubkov, M. V., Hartmann, M., and Gooday, A. J.: Grazing of
intertidal benthic foraminifera on bacteria: Assessment using pulse-chase
radiotracing, J. Exp. Mar. Biol. Ecol., 399, 25–34,
10.1016/j.jembe.2011.01.011, 2011.Muller, W. A. and Lee, J. J.: Apparent Indispensability of Bacteria in
Foraminiferan Nutrition, J. Protozool., 16, 471–478,
10.1111/j.1550-7408.1969.tb02303.x, 1969.Nomaki, H., Heinz, P., Nakatsuka, T., Shimanaga, M., and Kitazato, H.:
Species-specific ingestion of organic carbon by deep-sea benthic
foraminifera and meiobenthos: In situ tracer experiments, Limnol. Oceanogr.,
50, 134–146, 10.4319/lo.2005.50.1.0134, 2005.Nomaki, H., Heinz, P., Nakatsuka, T., Shimanaga, M., Ohkouchi, N., Ogawa, N.
O., Kogure, K., Ikemoto, E., and Kitazato, H.: Different ingestion patterns
of C-13-labeled bacteria and algae by deep-sea benthic foraminifera, Mar.
Ecol. Prog. Ser., 310, 95–108, 10.3354/meps310095, 2006.Nomaki, H., Bernhard, J. M., Ishida, A., Tsuchiya, M., Uematsu, K., Tame,
A., Kitahashi, T., Takahata, N., Sano, Y., and Toyofuku, T.: Intracellular
Isotope Localization in Ammonia sp. (Foraminifera) of Oxygen-Depleted
Environments: Results of Nitrate and Sulfate Labeling Experiments, Front. Microbiol., 7, 163, 10.3389/fmicb.2016.00163,
2016.Panieri, G.: Foraminiferal response to an active methane seep environment: A
case study from the Adriatic Sea, Mar. Micropaleontol., 61, 116–130,
10.1016/j.marmicro.2006.05.008, 2006.Panieri, G., James, R. H., Camerlenghi, A., Westbrook, G. K., Consolaro, C.,
Cacho, I., Cesari, V., and Cervera, C. S.: Record of methane emissions from
the West Svalbard continental margin during the last 23.500yrs revealed by
δ13C of benthic foraminifera, Global Planet. Change, 122,
151–160, 10.1016/j.gloplacha.2014.08.014, 2014.Panieri, G., Lepland, A., Whitehouse, M. J., Wirth, R., Raanes, M. P.,
James, R. H., Graves, C. A., Crémière, A., and Schneider, A.:
Diagenetic Mg-calcite overgrowths on foraminiferal tests in the vicinity of
methane seeps, Earth Planet. Sci. Lett., 458, 203–212, 10.1016/j.epsl.2016.10.024, 2017.Pascal, P.-Y., Dupuy, C., Richard, P., and Niquil, N.: Bacterivory in the
common foraminifer Ammonia tepida: Isotope tracer experiment and the
controlling factors, J. Exp. Mar. Biol. Ecol., 359, 55–61, 10.1016/j.jembe.2008.02.018, 2008.
Pawlowski, J.: Introduction to the molecular systematics of foraminifera,
Micropaleontology, 46, 1–12, 2000.Rathburn, A. E., Pérez, M. E., Martin, J. B., Day, S. A., Mahn, C.,
Gieskes, J., Ziebis, W., Williams, D., and Bahls, A.: Relationships between
the distribution and stable isotopic composition of living benthic
foraminifera and cold methane seep biogeochemistry in Monterey Bay,
California, Geochem. Geophy. Geosy., 4, 1220, 10.1029/2003GC000595, 2003.Reynolds, E. S.: The use of lead citrate at high pH as an electron-opague stain in electron microscopy, J. Cell Biol., 17, 208–212, 10.1083/jcb.17.1.208, 1963.Risgaard-Petersen, N., Langezaal, A. M., Ingvardsen, S., Schmid, M. C.,
Jetten, M. S. M., Op den Camp, H. J. M., Derksen, J. W. M., Piña-Ochoa,
E., Eriksson, S. P., Peter Nielsen, L., Peter Revsbech, N., Cedhagen, T.,
and van der Zwaan, G. J.: Evidence for complete denitrification in a benthic
foraminifer, Nature, 443, 93, 10.1038/nature05070, 2006.Salonen, I. S., Chronopoulou, P.-M., Bird, C., Reichart, G.-J., and Koho, K.
A.: Enrichment of intracellular sulphur cycle–associated bacteria in
intertidal benthic foraminifera revealed by 16S and aprA gene analysis, Sci.
Rep.-Uk, 9, 1–12, 10.1038/s41598-019-48166-5,
2019.Schmidt, C., Geslin, E., Bernhard, J. M., LeKieffre, C., Svenning, M. M., Roberge, H., Schweizer, M., and Panieri, G.: Dataset of publication: Deposit-feeding of Nonionellina labradorica (foraminifera) from an Arctic methane seep site and possible association with a methanotroph, Zenodo [data set], 10.5281/zenodo.6941739, 2022.Schneider, A., Crémière, A., Panieri, G., Lepland, A., and Knies,
J.: Diagenetic alteration of benthic foraminifera from a methane seep site
on Vestnesa Ridge (NW Svalbard), Deep Sea Res. Pt. I, 123, 22–34, 10.1016/j.dsr.2017.03.001, 2017.Serov, P., Vadakkepuliyambatta, S., Mienert, J., Patton, H., Portnov, A.,
Silyakova, A., Panieri, G., Carroll, M. L., Carroll, J., Andreassen, K., and
Hubbard, A.: Postglacial response of Arctic Ocean gas hydrates to climatic
amelioration, P. Natl. Acad. Sci., 114,
6215–6220, 10.1073/pnas.1619288114, 2017.Shetye, S., Mohan, R., Shukla, S. K., Maruthadu, S., and Ravindra, R.:
Variability of Nonionellina labradorica Dawson in Surface Sediments from
Kongsfjorden, West Spitsbergen, Acta Geologica Sinica – English Edition, 85,
549–558, 10.1111/j.1755-6724.2011.00450.x,
2011.Tavormina, P. L., Hatzenpichler, R., McGlynn, S., Chadwick, G., Dawson, K.
S., Connon, S. A., and Orphan, V. J.: Methyloprofundus sedimenti gen. nov.,
sp. nov., an obligate methanotroph from ocean sediment belonging to the
`deep sea-1'clade of marine methanotrophs, Int. J. Syst. Evol. Microbiol.,
65, 251–259, 10.1099/ijs.0.062927-0, 2015.Torres, M. E., Martin, R. A., Klinkhammer, G. P., and Nesbitt, E. A.: Post
depositional alteration of foraminiferal shells in cold seep settings: New
insights from flow-through time-resolved analyses of biogenic and inorganic
seep carbonates, Earth Planet. Sci. Lett., 299, 10–22, 10.1016/j.epsl.2010.07.048, 2010.Wefer, G., Heinze, P. M., and Berger, W. H.: Clues to ancient methane
release, Nature, 369, 282, 10.1038/369282a0,
1994.
Wollenburg, J. E., Raitzsch, M., and Tiedemann, R.: Novel high-pressure
culture experiments on deep-sea benthic foraminifera – Evidence for methane
seepage-related δ13C of Cibicides wuellerstorfi, Mar.
Micropaleontol., 117, 47–64, 2015.