# Innovative TEM-coupled approaches to study foraminiferal cells

Hidetaka Nomaki, Charlotte Lekieffre, Stéphane Escrig, Anders Meibom, Shinsuke Yagyu, Elizabeth Richardson, Takuya Matsuzaki, Masafumi Murayama, Emmanuelle Geslin, Joan Bernhard

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## To cite this version:

Hidetaka Nomaki, Charlotte Lekieffre, Stéphane Escrig, Anders Meibom, Shinsuke Yagyu, et al. "Innovative TEM-coupled approaches to study foraminiferal cells." Marine Micropaleontology, 2018, 138, pp.90-104. 10.1016/j.marmicro.2017.10.002. hal-02872784

HAL Id: hal-02872784

https://univ-angers.hal.science/hal-02872784v1

Submitted on 13 Jun 2025

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Innovative TEM-coupled approaches to study foraminiferal cells

Hidetaka Nomakiª, Charlotte LeKieffreᵇ, Stéphane Escrigᵇ, Anders Meibomᵇ,ᶜ, Shinsuke

Yagyuᵈ, Elizabeth A. Richardsonᶜ, Takuya Matsuzakiᵈ, Masafumi Murayamaᵈ,

Emmanuelle Geslinf, Joan M. Bernhardg

ªDepartment of Biogeochemistry, Japan Agency for Marine-Earth Science and Technology,
2-15 Natsushima-cho, Yokosuka, Kanagawa, 237-0061, Japan

ᵇLaboratory for Biological Geochemistry, School of Architecture, Civil and Environmental
Engineering (ENAC), Ecole Polytechnique Fédérale de Lausanne (EPFL), 1015 Lausanne,
Switzerland

ᶜCenter for Advanced Surface Analysis, Institute of Earth Sciences, University of Lausanne,
Switzerland

ᵈCenter for Advanced Marine Core Research, Kochi University, B200, Monobe,
Nankoku-city, Kochi, 783-8502, Japan

ᵉGeorgia Electron Microscopy, University of Georgia, Athens, GA 30602, USA

fLPG-BIAF, University of Angers, UMR CNRS 6112, 49095 Angers Cedex, France

g Department of Geology and Geophysics, Woods Hole Oceanographic Institution, Woods
Hole, MA 02543, USA

* Corresponding author: nomakih@jamstec.go.jp

Key words: Ultrastructure, NanoSIMS, micro-X-ray CT, correlative microscopy, isotope
mapping, microhabitat
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Abstract

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Transmission electron microscope (TEM) observation has revealed much about the basic cell biology of foraminifera. Yet, there remains much we do not know about foraminiferal cytology and physiology, especially for smaller benthic foraminifera, which inhabit a wide range of habitats. Recently, some TEM-coupled approaches have been developed to study correlative foraminiferal ecology and physiology in detail: Fluorescently Labeled Embedded Core (FLEC)-TEM for observing foraminiferal life-position together with their cytoplasmic ultrastructure, micro-X-ray computed tomography (CT)-TEM for observing and reconstructing foraminiferal cytoplasm in three dimensions (3D), and TEM-Nanometer-scale secondary ion mass spectrometry (NanoSIMS) for mapping of elemental and isotopic compositions at sub-micrometer resolutions with known ultrastructure.
In this contribution, we review and illustrate these recent advances of TEM-coupled methods.
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# 1. Introduction

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Benthic foraminifera are unicellular eukaryotes that are abundant constituents of the marine seafloor. Their habitats are diverse and, thus, as a group they encounter many different environmental conditions such as temperature, salinity, dissolved oxygen concentrations, water depths, trophic conditions, and redox conditions (reviewed in Murray 2006). It is expected that foraminifera have physiological adaptations to these various environments. Studies of their cytoplasmic structure are therefore keys to understanding how these microbial eukaryotes adapt to those different environmental conditions. Ultrastructural observation using a transmission electron microscope (TEM) has successfully been applied to foraminifera since the 1960s (Hedley et al. 1967; Dahlgren 1967a, b). The relationships between foraminiferal cellular ultrastructure and environments have been studied in a few cases (e.g., Leutenegger and Hansen 1979; Bernhard and Bowser 2008; Bernhard et al. 2006a, 2010a; 2010b). For instance, *Virgulinella fragilis*, which can thrive in micro-oxic and hydrogen sulfide-enriched environments, is known to support a double symbiosis with kleptoplasts and putative δ-proteobacteria (Bernhard 2003; Tsuchiya et al. 2015). Although ultrastructural observations have cast light on many different aspects of foraminiferal metabolic diversity, TEM observation alone is not enough to determine the functions of all organelles. For example, incubation experiments are needed to establish the function of putative “symbionts” such as photosynthetic kleptoplasts (e.g., Jauffrais et al. 2016, 2017).

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Also, recent advances on fine-scale analyses of porewater chemistry in sediments indicate both vertical and horizontal gradients and heterogeneity of constituents such as oxygen (Glud et al. 2005), nitrate (Glud et al. 2009; Metzger et al., 2016), dissolved reactive phosphorus, dissolved iron (Thibault de Chanvalon et al. 2015), and probably other compounds as well, at sub-millimeter length scales. Many benthic foraminifera are infaunal, particularly in mesotrophic to eutrophic sediments where they can utilize organic matter even in deep sediments (e.g., >3 cm below the sediment-water interface; Jorissen et al. 1995). The micro-environments of infaunal foraminifera are, thus, highly diverse and heterogeneous, and can change temporally (Glud et al. 2005). Although foraminifera are able to migrate within sediments (Alve and Bernhard 1995; Moodley et al. 1998; Gross 2000; Heinz et al. 2001; Geslin et al., 2004; Nomaki et al. 2005; Koho et al. 2011), the ultrastructural characteristics observed with the TEM are likely to reflect the micro-environments that cannot be characterized with bulk-sediment analysis.

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Recently, innovative methods using the TEM and coupled recent microanalytical technologies have been developed and applied to foraminiferal studies in efforts to overcome these challenges. The Fluorescently Labeled Embedded Core (FLEC) method is an excellent approach to reveal foraminiferal life position in sediments. The fluorescent tags are used to label metabolically active cells in sediments and their positions are maintained in the
sediments with glutaraldehyde fixation and subsequent resin embedding. Because FLEC also
labels additional organisms like ciliates and prokaryotes, it is also possible to observe
interactions between foraminifera and these organisms together with foraminiferal
ultrastructure observed with the TEM of the same sliced plane.

The complete sequential thin-section analysis of a whole organism or an
array-tomography technique can be applied to study small-sized specimens (Yabuki et al.
2012, 2013; Micheva and Smith 2007). Because foraminifera are relatively large cells
(hundreds of µm), however, the complete ultrastructure of an entire foraminifera is difficult to
reconstruct from TEM observations. One solution to the foraminiferal size issue is
micro-X-ray computed tomography (CT), which also provides information on internal
structures. Micro-X-ray CT has been used to assess attributes regarding foraminiferal tests
(shells), such as biometry (Speijer et al. 2008; Briguglio et al. 2011, 2013; Hohenegger and
Briguglio 2012; Ferrández Cañadell et al. 2014) and test-wall dissolution by means of X-ray
attenuation (Johnstone et al. 2010; Iwasaki et al. 2015). Nomaki et al. (2015) applied X-ray
CT to foraminiferal cells prepared for TEM (i.e., resin embedded; stained with heavy metals).
Because micro-X-ray CT is nondestructive, observations of foraminiferal cytoplasmic
structure with this technique prior to sectioning for TEM allow us to determine the precise
positioning of sections that have particular cytoplasmic structures, such as extended
reticulopodia, the cell periphery, chamber plugs, vacuoles, or multiple nuclei. The X-ray CT
can also determine the surface area and volume of foraminiferal cytoplasm and selected
structures, thus, permitting quantitative analyses of foraminiferal biovolume and growth
pattern through chamber addition, as well as allowing an estimation of volumes and surface
areas of vacuoles, which contain soluble compounds (Khalifa et al. 2016).

The combination of TEM and Nanometer-scale secondary ion mass spectrometry (NanoSIMS) can spatially investigate the elemental and isotopic compositions of organisms at sub-micrometer resolution (e.g., Finzi-Hart *et al.* 2009; Pernice *et al.* 2012; Carpenter *et al.* 2013; Hoppe *et al.*, 2013; Kopp *et al.* 2015). In pulse-chase experiments with isotopically labeled substrates, it is therefore possible to quantitatively evaluate where within a cell, such as specific organelle types or endobionts, a certain molecule or its metabolic derivates were located at a given point in time (Nomaki *et al.* 2016; LeKieffre *et al.* 2017).

Here, we present three advanced TEM-coupled technologies applied to foraminiferal
research; FLEC-TEM; micro-X-ray CT-TEM, and TEM-NanoSIMS. All these TEM-related
methods provide valuable information on foraminiferal biology and ecology. Examples of
these correlative technologies are presented along with summaries of each method, notes
regarding advantages and disadvantages, and possible future applications.

**2. Materials and Procedures**
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## 2.1. FLEC-TEM

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Specimens were prepared using the FLEC-TEM method described by Bernhard and Richardson (2014). Foraminifera-bearing sediment cores were collected between July 23-25, 2007 from an area off Monterey, California (USA) with active hydrocarbon seepage, at approximately 36°44.7′N 122°16.6′W (~1003 m water depth; Bernhard et al. 2010b), using pushcores manipulated by the ROV *Jason*. The viability indicator CellTracker Green CMFDA (Thermo Fisher Scientific; Bernhard *et al.*, 2006b) was injected into the pushcores and allowed to incubate *in situ* for approximately 1 day (Bernhard *et al.* 2009). For this study, two pushcores were utilized. One core, designated as the seep core, was collected adjacent to a clam bed (Bernhard *et al.*, 2010b) while the other, designated the non-seep core, was collected >25 m from visible active seepage. Aboard the surface vessel RV *Atlantis*, each core was subsampled with 1.5-cm diameter syringe cores, which were preserved in ~3% glutaraldehyde / 0.1 M cacodyllic acid (pH 7.2). At WHOI (USA), syringe cores were further processed for FLEC as described in Bernhard and Bowser (1996) and Bernhard *et al.* (2003). Polymerized FLEC cores were sectioned perpendicular to the sediment-water interface using a Beuhler Low Speed Saw for rocks. Resulting sections were ~0.5mm in thickness. Fluorescent foraminifera were observed using an epifluorescence-equipped dissecting microscope (Leica FLIII with fluorescein / Green Fluorescent Protein filters; 490-nm bandpass excitation; 520-nm bandpass emission), which was used to note foraminiferal location in sediment slices along with their depth below the sediment-water interface. Select foraminifera were further imaged with an Olympus Fluoview Laser Scanning Confocal Microscope (LSCM) and then excised from their FLEC sediment slice using a sharp razor blade. Foraminifera were identified using morphology, test texture (mineralized vs. agglutinated), and knowledge of the local foraminiferal assemblage. In many cases, identification to species was not possible.

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Blocks were trimmed as appropriate for TEM (Bernhard and Richardson, 2014), and sectioned with Diatome diamond knives into 1-µm thick sections for light microscopy and 70-nm thin sections for transmission electron microscopy. Thick sections were stained with toluidine blue and imaged with a Zeiss Axiovert 40C Tissue Culture microscope. Thin sections were post-stained with a solution of uranyl acetate (4%) in ethanol and a modified version of Sato's lead citrate (Hanaichi et al., 1986). Transmission electron microscopy observations were made using a Zeiss 10CA TEM at the Marine Biological Laboratory (Woods Hole, MA, USA) or a JEOL JEM1011 TEM at the Georgia Electron Microscopy facility, University of Georgia (Athens, GA, USA).

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## 2.2. Micro-X-ray CT

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### 2.2.1. Foraminiferal sampling and fixation
The intertidal foraminifera *Ammonia* sp. (phylotype T6; Holzmann 2000, Hayward et al. 2004) was collected from the Nojima tidal flat (35°19.4N 139°38.1E; water depth, ~0 m) of Tokyo Bay on March 21, 2014. The deep-sea foraminiferal samples were collected from central Sagami Bay, Japan (35°01.3N 139°22.1E; water depth 1430 m), during a cruise of *R/V Kaimei* KM16-01 in April 2016. At the Nojima tidal flat, the top ~1 cm of sediment was collected into several 4 L buckets for foraminiferal fixation in the laboratory. For Sagami Bay, sediment samples collected with a push corer were immediately sliced onboard into 0.5- or 1-cm sections to a depth of 6 cm. The sediment samples were fixed in 10% glutaraldehyde in cacodylate buffer to a final concentration of 4.0% glutaraldehyde. In the laboratory, specimens that had an aggregation of sediment particles around their aperture and/or whose test was mostly filled with cytoplasm were isolated from the >125-µm size fraction using a binocular stereomicroscope.

### 2.2.2. Post-fixation, staining and embedding into resin

Sample post-fixation using osmium tetroxide, staining with uranyl acetate, and subsequent embedding into resin block followed the protocol for micro-X-ray and TEM sample preparation described previously (Nomaki et al. 2015). To yield high-contrast specimens for X-ray CT analysis, specimens were post-fixed and stained with osmium tetroxide and uranyl acetate. Osmium tetroxide is a conventional post-fixation agent that preserves lipids, and also some amino acids, peptides, and proteins to a lesser extent (Hayat 2000). Uranyl acetate stains nucleic acids and, to a lesser extent, proteins (Hayat 2000).

### 2.2.3. Micro-X-ray CT imaging

The resin-embedded Os- and U-stained foraminiferal specimens were imaged using a model Xradia 410 Versa microfocus X-ray CT system (Zeiss). An X-ray filter was attached to the X-ray source to reduce the X-ray energy to yield better resolution. The X-ray source setting was 40 kV and 200 µA for *Bolivina spissa*, and the exposure time for each projection was 10 seconds. A total of 1601 projections were performed during 360° rotation and the resolution of the obtained image was 1.07 µm/pixel. For *Ammonia* sp. (phylotype T6), different settings were applied: 60 kV and 130-µA X-ray source with 5 second projection for a total of 801 projections per 360° rotation. The short projection time and lower numbers of projections for *Ammonia* sp. (phylotype T6) resulted in lower quality images compared to those of *B. spissa*. The obtained data were reconstructed using the manufacturer-supplied software and Osirix MD 6.5 (Pixmeo) software.

## 2.3. TEM-NanoSIMS

TEM-NanoSIMS analyses of benthic foraminifera was conducted independently in
two different laboratories. Although the principal methods for our two sets of correlative TEM-NanoSIMS analyses were similar, there are several minor differences between Nomaki et al. (2016) and LeKieffre et al. (in press), as summarized in Tables 1 and 2. Below we describe similarities and differences in methodology.

### 2.3.1. Incubation with stable-isotope tracer(s)

Benthic foraminifera collected from Nojima tidal flat (Japan), Bourgneuf Bay (France), or Gullmar Fjord (Sweden) (Table 1, Nomaki et al. 2016, LeKieffre et al. in press and unpublished data) were incubated for different durations with different isotope tracers (as noted below). *Ammonia* sp. (phylotype T6) collected from the Nojima tidal flat were incubated with sediment either in dysoxic or anoxic conditions. Foraminifera were isolated from Bourgneuf Bay sediments (*Ammonia* sp. phylotype T6 and *Haynesina germanica*) and Gullmar Fjord sediments (*Nonionellina labradorica*) and incubated without sediments in artificial seawater (ASW, RedSea salt, salinity: 35). All foraminiferal specimens were incubated with additions of ¹³C, ¹⁵N, or ³⁴S tracer(s) (Table 1, Nomaki et al. 2016; LeKieffre et al. in press and unpublished data).

### 2.3.2. Fixation and embedding into resin

After isotopic incubations, the Nojima flat sediments including benthic foraminifera were fixed in 10% glutaraldehyde in cacodylate buffer to a final concentration of 4.0% glutaraldehyde. Specimens whose tests (except for the ultimate one or two chambers) were filled with bright yellow cytoplasm typical of this *Ammonia* sp. (phylotype T6) were selected for analysis. Foraminifera from Bourgneuf Bay and Gullmar Fjord were, at the end of their incubation in isotope-spiked ASW, immediately fixed according to the chemical fixation protocol described in LeKieffre et al. (this issue). These glutaraldehyde-fixed foraminiferal specimens were either embedded in agarose (Nomaki et al. 2016) or directly decalcified (LeKieffre et al., in press), and then all specimens were post-fixed using osmium tetroxide. The Nojima specimens were additionally stained with uranyl-acetate "en-bloc" for 2 h followed by rinsing with distilled water. All samples were subsequently embedded in resin (Nomaki et al. 2016, LeKieffre et al. this issue, Koho et al. this issue).

### 2.3.3. Sectioning, TEM imaging and coating

Embedded specimens were sectioned into ultra-thin (60-nm to 70-nm thick) or semi-thin (200-nm thick) sections. Sections were transferred to copper grids and stained with 2% aqueous uranyl acetate only (LeKieffre et al. in press) or 2% aqueous uranyl acetate followed by 0.3% lead nitrate and 0.3% lead acetate solution (Nomaki et al. 2016). In Nomaki et al. (2016), specimens' ultra-thin sections were used for detailed observations of organelle membranes to check cell integrity (e.g., intact mitochondrial cristae and
double-membranes); semi-thin sections were used for subsequent TEM-NanoSIMS analyses. In the case of LeKieffre et al. (in press) materials, the same ultra-thin sections were used for both TEM and NanoSIMS. Once the target area for NanoSIMS analyses was selected, that targeted region was observed sequentially and a composite of TEM images was reconstructed. TEM-imaged sections intended for NanoSIMS analysis were coated with 20 nm of osmium (Nomaki et al. 2016) or 10 nm of gold (LeKieffre et al. in press) to prevent samples from electron charging during NanoSIMS analysis.

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### 2.3.4. NanoSIMS measurements

Grids containing metal-coated sections were placed into the sample holder of a nanometer-scale secondary ion mass spectrometer (NanoSIMS NS50 or NanoSIMS 50L). The settings of the NanoSIMS analyses are summarized in Table 2.

During analysis, secondary ions were extracted with a primary $Cs^+$ beam. Different settings were used for the $Cs^+$ beam, resulting in a slightly different primary beam spot size on the two instruments (Table 2). A multi-collector system was used to simultaneously measure five to seven different secondary ions (Nomaki et al. 2016; LeKieffre et al. in press). Areas of 8 × 8-µm to 30 × 30-µm were imaged with either 128 × 128 or 256 × 256 pixels rastered images. Sequential layers of images (up to 50) were accumulated to produce one image for each given area. The data quantification was performed after a drift correction of all layers, using the software ImageJ or L'IMAGE developed by Dr. Larry Nittler, Carnegie Institution of Washington, USA.

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### 2.3.5. Data analysis

Correlation between NanoSIMS and TEM images was done by aligning the $^{12}$C$^{14}$N⁻ ion image with the corresponding ultrastructural features in a TEM image. Values of $^{15}$N and $^{34}$S enrichments were obtained from the $^{12}$C$^{15}$N⁻/$(^{12}C^{14}N⁻ + ^{12}C^{15}N⁻)$ or $^{12}$C$^{15}$N⁻/$^{12}$C$^{14}$N⁻ and $^{34}$S⁻/$(^{32}S⁻ + ^{34}S⁻)$ or $^{34}$S⁻/$^{32}S⁻$ ratios respectively, and presented as ‰ enrichment over the natural isotope ratio measured in unlabeled samples prepared and analyzed identically. These ratios were calculated for selected regions of interest, using the program Look@NanoSIMS (Polerecky et al., 2012). This allows comparison of different cell compartments within the cytoplasm, or between different time points or incubation conditions (e.g., pH, light, salinity, temperature, oxygen, presence/absence of a certain nutrient). It is noted that comparisons of isotopic ratios between samples from experiments using different sample preparation methods should be treated with caution as differences in sample preparation can lead to different isotopic ratios (Musat et al., 2014). For the data presented here, such effects did not significantly impact data interpretations.

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## 3. Results and Discussions
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## 3.1. FLEC-TEM

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### 3.1.1. Results

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Here, two foraminifera were designated for fluorescently labeled embedded core (FLEC)-TEM analyses, one individual from each ROV-collected core. In the non-seep core, the specimen was located at a sediment depth of 1.7 cm below the sediment-water interface. The upper edge of the core section shown in Fig. 1A was not the sediment-water interface. This specimen was tentatively identified as a *Buliminella tenuata* due to knowledge of the area's foraminiferal community (Bernhard et al., 2001; Bernhard et al., 2010b); it appeared to have intact cytoplasmic vacuoles in LSCM images (Fig. 1B). The test of this specimen was calcite, as evidenced by its appearance in reflected light (Fig. 1A) and in semi-thin section (Fig. 1C). The ultimate chamber of this individual contained little cytoplasm (Fig. 1B, C), which is to be expected given this is where reticulopods are deployed and retracted through the aperture (e.g., Alexander and Banner, 1984). This specimen also appeared to have a distinct layer of fluorescent material attached to the test (shell) exterior, as noted in the LSCM micrograph (Fig. 1B). In the sediments, the specimen was slightly inclined from horizontal, with the apertural end oriented deepest (Fig. 1B, C). What appears to be an open pore space is adjacent to the specimen's aperture (Fig. 1B, C).

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In TEM micrographs of *B. tenuata* from the non-seep core, large vacuoles were visible (Fig. 2A). It is notable that osmication is not included in the FLEC-TEM preparation procedure. Thus, the membranes in FLEC-TEM specimens do not appear as distinctly as those membranes observed in specimens that were conventionally prepared for TEM (i.e., with osmication). What appeared to be degradation vacuoles (i.e., food vacuoles; see LeKieffre et al., this issue) were relatively common (Fig. 2A, B), containing items appearing to be prokaryotes as well as unidentifiable materials. Mitochondria were identified by their shape and size (Fig. 2B-D); in some cases cristae and the double membrane were discernable (Fig. 2C), but not clearly. The mitochondria appeared to occur at the cell periphery, as well as far from the periphery, well within the cell. Electron-opaque bodies, sensu LeKieffre et al. (this issue) (also known as "electron dense bodies"; Nomaki et al., 2016), were also observed (Fig. 2C-E). Golgi were also noted (Fig. 2 B-D), as were peroxisomes with endoplasmic reticulum (Fig. 2E). The crystalized core (catalase) of peroxisomes (Fig. 2E inset; Bernhard and Bowser, 2008) was not as distinctive in these preparations compared to those subject to conventional TEM processing with osmication. There was no evidence of prokaryotic endobionts (i.e., endosymbionts) in this foraminiferan. Examination of the area immediately adjacent to what would have been the test exterior revealed that the fluorescent material observed in the LSCM image (Fig. 1B) was not prokaryotic (i.e., ectobionts) but was most likely foraminiferal reticulopods (Fig. 2F, G). Because we did not fix the FLEC cores to preserve reticulopods (Travis and Bowser 1986, Bernhard et al., 2013), well-preserved
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reticulopodial networks were not expected, but such structures (Fig. 2F, G) are not inconsistent with reticulopodial cytoplasm.

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The foraminifer from the seep core was agglutinated, evidenced by the brownish color of the test (Fig. 3A), especially when compared to a calcareous (white) foraminifer in the same image. This agglutinated seep specimen was clearly biserial but we were not confident in identifying this specimen to a genus or species. The specimen aperture was located at a sediment depth of 1.7 mm below the sediment-water interface (Fig. 3A, B), near a juvenile clam. Microbes (eukaryotes and/or prokaryotes) also occurred in these sediments (Fig. 3B), but no additional foraminifera visible in reflected light (Fig. 3A) were living (i.e., fluorescent; Fig. 3B). The biserial seep specimen was oriented with its aperture facing downward, at a ~45° angle from horizontal (Fig. 3A, B). As in the *B. tenuata*, the ultimate (youngest) chamber of this agglutinated individual was less fluorescent, indicating presence of less cytoplasm than in middle chambers (Fig. 3B, C). Mineral grains in the test of the specimen can be discerned in some places (Fig. 3C).

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The LSCM image of the biserial agglutinant shows it had visible cytoplasmic vacuoles, as well as one large nucleus (Fig. 3C). Compared to the exterior surface of the non-seep specimen's test, this seep specimen lacked an obvious layer of fluorescent material on its test exterior. However, it is clear that short filamentous prokaryotes occurred in sediments in the vicinity of this seep specimen (Fig. 3C). What appears to be either a feeding bolus or reticulopodial trunk is located in the apertural region (Fig. 3C). Because of this specimen's large size, only the older portion of its test was excised and subject to ultrastructural analysis.

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The TEM micrographs of the agglutinated seep specimen show the presence of large cytoplasmic vacuoles (Fig. 4), most of which lacked preserved membranes. Degradation vacuoles were present (Fig. 4A, B) but not as common as in the non-seep *B. tenuata* specimen. In the space between the organic lining and the interior surface of the test, prokaryotes were sometimes observed (Fig. 4B). There was no evidence of endosymbionts in the endoplasm, although some vacuoles appeared to contain one or two items that appeared to be prokaryotes (Fig. 4A); the identity and role of these microbes is not certain but the reader is directed to the contribution in this issue on prokaryotic associates (e.g., symbionts, parasites; Bernhard et al., this issue). Smaller vacuoles (Fig. 4C, D) appeared to have preserved membranes. Given the slight contrast due to lack of osmication (see above), mitochondria were identified on the basis of size and shape (Fig. 4C, D). In some mitochondria (Fig. 4C, D), a double membrane was clearly noted. The mitochondria did not appear to occur exclusively at the cell periphery. Peroxisomes were noted occasionally in characteristic distributions with endoplasmic reticulum at vacuole peripheries (Fig. 4D; Bernhard and Bowser, 2008); Golgi was also noted (not shown).

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344 3.1.2 FLEC-TEM Methodological notes

345 Correlative epifluorescence and TEM methods are not novel (e.g., Rieder and Bowser, 1987). The FLEC-TEM approach, which was recently described (Bernhard and Richardson, 2014), is novel in that it allows examination of cellular attributes in specimens that are 346 located within sediments, where they lived at the time of fixation. The ability to observe the 347 life position of foraminifera, or any other meiofauna, is advantageous to ecological studies 348 but the method does have disadvantages, some of which are discussed in Bernhard and 349 Richardson (2014). Briefly, the disadvantages of FLEC-TEM are: (1) Sectioning of 350 specimens among sediments will expose the diamond knife to heavy use, likely causing 351 damage and the need for resharpening. A novel method that merges use of a focused ion 352 beam (FIB) with the use of a scanning electron microscope (SEM) can produce TEM-quality 353 ultrastructural information in cells (Narayan and Subramaniam, 2015), although this 354 FIB-SEM method has not been performed on foraminifera, to our knowledge. (2) Also, as 355 noted, the lack of osmication prevents preservation and detection of some membranes. 356 Apparent lack of membranes in FLEC-TEM specimens does not necessarily mean the 357 membranes did not exist at the time of fixation. (3) Another disadvantage involves fixation 358 rate. Typically, when materials are preserved in glutaraldehyde, the fixation process is quick 359 because specimens are immersed directly into the fixative solution. With FLEC fixation, 360 however, the fixative must percolate into the sediments contained in the syringe core. 361 Depending on the pore-water content and sediment porosity, as well as specimen location 362 (e.g., deep infaunal vs. epifaunal), fixation of FLEC samples may be considerably slower 363 than immediate fixation of isolated individuals. The slower speed of fixation could impact 364 cellular preservation in unknown ways. (4) Foraminiferal identification at the species or 365 genus level hinges on the plane of section; confident identification can be difficult to 366 impossible. Depending on the study objective(s), such constraints may not be an issue. (5) 367 For some studies, it is imperative to distinguish living from dead specimens (e.g., Bernhard et 368 al., 2010b). Our experience with FLEC-TEM to date indicates that viability in specimens 369 prepared with this method is inconclusive because, for example, mitochondrial status (cristae 370 and double membrane) is not clearly discernable. While significantly degraded cytoplasm is 371 easily detected, any conclusions asserting that a specimen analyzed with FLEC-TEM was 372 living at the time of preservation should be made with due caution. 373

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376 3.1.3 Methodological advantages: Merging ecology and physiology

377 Advantages of the FLEC-TEM method include the ability to pair fine-scale 378 distribution with cellular attributes, as described in Bernhard and Richardson (2014) and 379 discussed further below. Because foraminifera move relatively slowly, we expect that the 380 position of individual foraminifera observed in FLEC sections is where the individual lived, 381 unless it was entrained down the core barrel at the edge of the core. Thus, we can distinguish
between surface-living (epifaunal) and deep-dwelling (infaunal) specimens, and we can
observe the fitness of each of those cells. In addition to determining position in sediments and
orientation of the aperture, FLEC also allows observation about the microhabitat of the
individual foraminifer. Thus, the sediment texture, fabric, and pore space can be assessed, as
can attributes regarding the neighboring microbial community. If neighbors have distinct
morphologies, we can determine if particular types of organisms consistently associate
together (Bernhard et al. 2003).

On the cellular scale, FLEC-TEM allows insights regarding adaptations and diet,
which can be particularly insightful when considering the fine-scale location of any given
individual within sediments, especially in chemoclines where steep geochemical gradients
exist. Furthermore, FLEC-TEM can determine if individuals collected from oxygen-depleted
environments have specialized features, such as prokaryotic associates, putative symbionts,
numerous peroxisomes, or peripheral / pore-plug associated mitochondria.

3.1.4 Observations on the paired ecology and physiology of these seep and non-seep
foraminifers

Due to the discernable double membrane and cristae noted in some mitochondria,
relatively intact peroxisomes, endoplasmic reticulum and Golgi, as well as abundant
degradation vacuoles, and presence of reticulopodial remains on the test exterior (Fig. 2), we
assert that the non-seep *Buliminella tenuata* specimen was active (i.e., living) at the time of
fixation. Because this specimen was located at a sediment depth of 1.7 cm, the pore-waters
bathing this specimen at the time of collection likely were depleted in oxygen (e.g., Bernhard,
1992). There was no evidence of the foraminifer being directly associated with any burrows
or tubes, although one tube was present in FLEC sections from the other side of the 1.5-cm
diameter FLEC core. Burrows and tubes can oxidize pore waters at depth (e.g., Thibault de
Chanvalon et al., 2015). We note that this non-seep *B. tenuata* had mitochondria both at the
cell periphery as well as further inside the cell. Foraminifera from low-oxygen habitats often
concentrate mitochondria at the cell periphery (e.g., Bernhard et al., 2010a; Leutenegger and
Hansen, 1979; see also LeKieffre et al., this issue) but can also have mitochondria throughout
the cell. Additional evidence that the specimen was living in oxygen-depleted pore waters is
the presence of characteristic peroxisome-endoplasmic reticulum complexes that are often
noted in specimens from low-oxygen habitats (Bernhard and Bowser, 2008; see also
LeKieffre et al., this issue). The specimen lacked evidence of endosymbionts, which is
consistent with the non-seep habitat.

The results presented here on the agglutinated seep foraminifer are valuable because,
to our knowledge, observations of the cell ultrastructure and organelle attributes of
multi-chambered deep-sea agglutinated foraminifera are not published. This lack of published
micrographs likely is because such robust agglutinated tests are extremely challenging to
420 section.

421 As with the non-seep *B. tenuata*, assessing viability in the seep-associated biserial
422 agglutinated foraminiferan is difficult. Mitochondria with double membranes and partially
423 discernable cristae were present, as were additional organelles (e.g., peroxisomes,
424 endoplasmic reticulum, small vacuoles with membranes, Golgi), suggesting the specimen
425 was living or recently living. The association of the peroxisomes with the periphery of large
426 vacuoles adds support that this agglutinated foraminifer was living in the seep habitat because,
427 as noted, many foraminifera living in steep chemoclines have such associations (Bernhard
428 and Bowser, 2008; Bernhard et al., 2010b; Martin et al., 2010). The presence of a bolus or
429 reticulopodial trunk in the apertural region (Fig. 3C) provides additional evidence that the
430 agglutinated specimen was active at the time of fixation. The relative dearth of degradation
431 vacuoles suggests the specimen may not have been particularly active at the time of fixation,
432 although it appears that the foraminifer was ingesting prokaryotes (Fig. 4A). It is possible that
433 more degradation vacuoles appeared in the younger part of the specimen that was not
434 examined with TEM.

435 The seep-associated agglutinated foraminiferan did not appear to be migrating toward
436 the sediment-water interface because its aperture was oriented downward (Fig. 3). Generally,
437 foraminifera move in the direction that their aperture points (e.g., Heinz 1999; H. Nomaki,
438 pers. obs.). Thus, if we assume the specimen was living at the time of fixation, it was not
439 adverse to the localized seep chemistry. Unfortunately, we lack porewater chemical analyses
440 on this specific FLEC core, but the pushcore was collected adjacent to an active
441 chemosynthetic mussel bed. Also, we note presence of some filamentous prokaryotes (Fig.
442 3C) in the vicinity of the foraminiferan; these filaments could be the sulfide oxidizing
443 *Beggiatoa*, which is commonly known to occur in seeps (e.g., Lloyd et al. 2010).
444

445 3.2. Micro-X-ray CT-TEM
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446 3.2.1. Results

447 The X-ray computed tomography (CT) imaging using the Os- and U-stained
448 foraminifera embedded in resin block showed three-dimensional (3D) structures of
449 foraminiferal cytoplasm (Figs. 5 and 6, Supplementary movies 1, 2, and 3). Using different
450 contrast settings in the software, we were able to observe both external morphology (e.g.,
451 Supplementary movie 2) and internal structures larger than several micrometers (e.g.,
452 Supplementary movie 3). In a specimen of *Ammonia* sp. (phylotype T6), we observed that the
453 cytoplasm of the ultimate 3 to 5 chambers were less dense in comparison to that of the other
454 chambers based on the differences in X-ray attenuation (Fig. 5A, C, Supplementary movie 1),
455 probably reflecting more vacuoles (Nomaki et al. 2014) and less lipid droplets (Koho et al.
456 this issue), together with less staining of the cytosol (data not shown). Many chambers in the
457 last whorl had relatively large vacuoles (~50 µm in diameter) close to the axis of rotation (Fig.
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5C). A chamber marked with a white arrowhead in Fig. 5A showed a cytoplasmic depression; this void between the calcite test and cytoplasm is sometimes observed for certain specimens and might be an artifact due to the shrinkage of the cytoplasm during the fixation process. In a specimen of *Bolivina spissa*, the last 2 chambers were “frothy” and not all the chamber space was occupied with cytoplasm (Fig. 6A, B, Supplementary movie 2). The proloculus cytoplasm also looked less dense than other chambers’ cytoplasm (Fig. 6C). Cross sectional views showed many vacuoles of 10- to 30-µm diameter throughout all chambers (Fig. 6D, E). Another obvious structure that was observed with micro-X-ray CT is plugs, which had high electron density due to uranium staining (Fig. 6D, Supplementary movie 3; Nomaki et al. 2015). Plugs are hypothesized to provide a means to partition portions of the cell not only among different chambers but also into different volumes, which, hypothetically, perform different intracellular biochemical reactions (Hottinger 2000; Ferrández Cañadell 2002). Sometimes, plugs were observed in each chamber of an entire foraminiferal specimen (Nomaki et al. 2015). In some specimens, sediment particles around the aperture and trunks of reticulopods extending from the aperture were also observed (not shown).

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### 3.2.2. Advantages and applications

The advantage of the micro-X-ray CT is to provide an overall view of cytoplasmic conditions, allowing one to target characteristic areas or organelles (e.g., vacuoles, plugs, nucleus). These observations greatly help us to determine the positions of TEM sections. Also, the whole-specimen perspective helps to evaluate the representativeness of the observed ultra-thin section(s) in the observed specimen. Pre-observation of the resin-embedded specimens using micro-X-ray CT before sectioning for TEM observation may become common when micro-X-day CT scanning becomes more affordable and faster.

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Another advantage of micro-X-ray CT is the ability to quantify the volume or surface area of the region of interest (Nomaki et al. 2015). The volume of cytoplasm in each chamber allows, e.g., calculation of the biovolume, growth pattern through chamber addition, and assessment of fitness. The total cell volume and volume of each vacuole also provides information to estimate the total chemical contents or concentrations in vacuoles. The cytoplasm-occupancy ratio seems to differ greatly between chambers and between species (Figs. 5 and 6, Nomaki et al. 2015), reflecting the volume of vacuoles that might be affected by their environment (LeKieffre et al., this issue).

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### 3.2.3. Spatial resolution limitations

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The spatial resolution of the micro-X-ray CT continues to improve; the current spatial resolution of the commercial micro-X-ray CT, e.g. Xradia 410 Versa (ZEISS, this study) or SkyScan 1272 (Bruker; Nomaki et al. 2015), is as low as 0.35 µm. Furthermore, because the differences in X-ray attenuation between metal-labeled cytoplasm and resin are not large as
between calcite (test) and air (e.g., Iwasaki et al. 2015), identification of small organelles like
mitochondria and peroxisomes remains difficult. Certain studies using full-field or scanning
X-ray microscopes and scanning transmission X-ray microscopes reported 50- or 60-nm
resolution of internal fungal features (Larabell and Le Gros 2004; Le Gros et al. 2005),
indicating there is potential to observe small organelles in foraminifera. Such high-resolution
images, however, can be achieved only for some limited target areas, not for whole
foraminiferal cells. High-resolution 3D observations of limited areas are also achieved using
a focused ion beam scanning electron microscope (FIB-SEM; e.g., Lucas et al. 2014)
technology at a voxel size of 5 to 10 nm with a maximal sample size of 50 × 30 × 30 µm in
volume, which remains a small fraction of foraminiferal cell size.

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### 3.3. TEM-NanoSIMS

#### 3.3.1. Elemental and isotope mapping and quantification

Nanometer-scale secondary ion mass spectrometry (NanoSIMS) analyses on a target
area of ultra-thin sections or semi-thin sections (Fig. 7A) previously observed with a TEM
(Fig. 7B, C) provide qualitative elemental concentration data and stable isotope ratios for
imaged and identifiable ultrastructural features (Fig. 7D, E). NanoSIMS <sup>12</sup>C<sup>14</sup>N- ion images
contain a wealth of ultrastructural information that enables precise correlation with TEM
images (Fig. 7D, E).

An example of isotopic ratio analysis using Look@NanoSIMS showed quantitative
information on isotopic compositions of each region of interest (ROI) (Fig. 8). In this
example, the foraminifer was fed with <sup>13</sup>C-enriched diatoms (d), which were observed with
their silica frustules in the foraminiferal endoplasm (Fig. 8A). The ROIs based on the TEM
image can be correlated with obtained NanoSIMS images (Fig. 8B, C, D), including isotopic
ratios (Fig. 8E), and the quantitative data can be summarized into a table (Fig. 8F).

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#### 3.3.2. Examples of 5 different isotope labels

Elemental ratios provide qualitative indications of relative C and N contents in
measured areas corresponding to different organelles or ultrastructural features. For example,
regions of lipid droplets exhibited lower N concentrations relative to other organelles and the
cytosol (Fig. 7D, E). This may reflect low concentrations of N in lipid droplets (and also in
the embedding resin), which have been thought to store lipids as a carbon source (LeKieffre
et al. 2017, submitted), although the actual contents of “lipid droplets” are not well
understood. On the other hand, certain organelles like mitochondria exhibited higher N
concentrations than those of the cytosol (Fig. 7D, E; see clearer image in Fig. 2 of Nomaki et
al. 2016).

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The stable isotopic compositions of C, N, and S, which are calculated with the ratios
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of ¹³C¹²C⁻/¹²C₂⁻, ¹²C¹⁵N⁻/(¹²C¹⁴N⁻ + ¹²C¹⁵N⁻) or ¹²C¹⁵N⁻/¹²C¹⁴N⁻, and ³⁴S⁻/(³²S⁻ + ³⁴S⁻) or ³⁴S⁻/³²S⁻, respectively, show the localization of the added stable isotope labels (Fig. 9). For example, the addition of ¹³C-labeled diatoms revealed the incorporation of food materials (Fig. 9A). While most ¹³C-labeling was observed within the organic material contained in the diatom frustules, one spot outside a diatom frustule exhibited high ¹³C-concentrations (lower left of Fig. 9A), suggesting the C in this region was derived from a digested diatom.

The addition of ¹³C-enriched inorganic C (H¹³CO₃⁻) provides information on autotrophic/anaplerotic C fixation into biomass (Fig. 9B). In the example provided in Fig. 9B, foraminifera were supplied with H¹³CO₃⁻ to investigate the autotrophic metabolism of kleptoplastic benthic foraminifera. Because soluble ¹³C-compounds are generally lost from the cell during TEM sample preparation (see section 3.2.4.), the isotope enrichments measured with the NanoSIMS represent ‘structural’ organic molecules derived from H¹³CO₃⁻. This data showed the ¹³C-assimilated by photosynthetic symbiont and its subsequent transfer and re-processing within the foraminiferal cell.

The assimilation of inorganic sources of ¹⁵N (e.g., ammonium or nitrate) can provide an important assessment of nitrogen assimilation and translocation within foraminiferal cells (Fig. 9C, D). Both ammonium and nitrate are important N sources for amino acid synthesis in autotrophic organisms, including foraminiferal symbionts or organelles like kleptoplasts. Here, assimilation of ¹⁵N originating from ¹⁵NH₄⁺ was observed as small spots of less than 1 µm in diameter, corresponding to electron-opaque bodies (Fig. 9C). Similar assimilation of ¹⁵N into electron opaque bodies (called “electron dense bodies” in Nomaki et al. 2016) was also observed in the specimens incubated with ¹⁵NO₃⁻ (Fig. 9D). Nitrogen isotopic compositions of amino acids in *Ammonia* sp. (phylotype T6) incubated with ¹⁵NO₃⁻ indicated that those ¹⁵N labels existed as amino acids probably originated from endobiotic prokaryote (Nomaki et al. 2014; reported as *Ammonia beccarii*). Enrichments in ¹⁵N were obvious in electron-opaque bodies of the *Ammonia* sp. (phylotype T6) incubated in anoxia (Fig. 9D), but were also obvious in mitochondria, cytoplasm, and food vacuoles in dysoxia-incubated specimens, suggesting metabolism varied under different oxygen concentrations (Nomaki et al. 2016). Organic lining and lipid droplet regions were modestly labeled with ¹⁵N even after 14 days, indicating slower turnover than for other organic matter.

Using ³⁴SO₄²⁻ as an isotope tracer in *Ammonia* sp. (phylotype T6) allowed visualization of ³⁴S localization in the cytoplasm (Fig. 9E), likely resulting from metabolism related to sulfur assimilation and/or sulfate reduction by symbionts. Enrichments in ³⁴S were detected in various opaque bodies, but enrichment was also detected in the cytoplasm near vacuoles or as remains inside vacuoles (Fig. 9E). As observed for ¹⁵N, ³⁴S assimilation was more pronounced in the specimens incubated with dysoxia versus those incubated in anoxia.
### 3.3.3 Methodological advantages and applications

The main advantage of correlated TEM-NanoSIMS observations is the small scale imaging of elements and isotopes. Indeed, this method allows the sub-cellular localization of isotopically labeled compounds, and gives the possibility to quantify isotopic ratios at the scale of organelles (micron to sub-micron scales). Such knowledge has wide applications to studies of foraminiferal biology and physiological processes such as symbiosis, kleptoplasty, trophic metabolism or denitrification. The use of labeled key elements combined with an appropriate incubation time(s) (see below) can also be used to study foraminiferal metabolism under different laboratory-controlled conditions, such as oxic vs. dysoxic conditions or light/dark incubations. For instance, symbiosis between ecto-/endobionts or algal symbionts and foraminifera, widely reported in the literature (e.g., Lee and Hallock 1987; Lee and Anderson 1991; Bernhard et al. 2000, 2006a, 2010a), are obvious candidates for correlated TEM-NanoSIMS study, because little is known yet about the role of the symbionts or the metabolite transfer between the symbionts and the host cell. For example, phylogenetic studies of *Virgulinella fragilis* endobionts suggest they are putative sulfate reducing δ-proteobacteria (Tsuchiya et al. 2015). The close phylogenetic relationship between the *V. fragilis* endobionts and sulfate reducing δ-proteobacteria suggests these endobionts use organic matter from the host for carbon oxidation. In another example, a study using a radio-isotope tracer in foraminifera containing different algal symbionts suggested that metabolites exchanged between host foraminifera and algal symbionts differed between diatom-retaining foraminifera and dinoflagellate-retaining foraminifera (Kremer et al. 1980). However the Kremer et al. study performed a bulk cell analysis of radio-isotope label transfer, without any spatial information inside the foraminifera. The use of TEM-NanoSIMS with stable isotope labeling could identify the transfer of target elements from specific symbionts to foraminifera or *vice versa* at a sub-micron scale.

Time-series sampling in incubation experiments is important for elucidating the actual pathways of the label into foraminiferal ultrastructure or symbiotic partners. For instance, the use of ¹³C-labeled diatoms as a foraminiferal food source (Figs. 8 and 9A) in a longer incubation reveals the ¹³C-assimilation into the foraminiferal biomass. The extent of labeling differs between ultrastructural features (i.e., heterogeneity), showing localization in specialized features such as lipid droplets (LeKieffre et al., 2017). Further, prolonged incubations should result in the ¹³C-label being respired into ¹³CO₂ and the ¹³C-signal in the cytoplasm decreasing over time. The use of NanoSIMS to trace ¹³C- or ¹⁵N-labeled organic matter with appropriate time series sampling, thus has great advantages in metabolic and stoichiometric studies of foraminifera, because one could estimate not only food selectivity but also turnover rates of each organelle type.

### 3.3.4. Limitations and challenges
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NanoSIMS analysis cannot discriminate which organic molecules were isotopically labeled. Thus, NanoSIMS imaging cannot determine the exact nature of the compounds transferred, but isotope-label localization with ultrastructural observations can provide some guidance. Nevertheless, employing a suite of multiple approaches, such as compound specific isotope analysis of metabolites, genomic and transcriptome analyses, organelle purification with their chemical analyses, in combination with TEM-NanoSIMS, will provide a clearer comprehension of foraminiferal cell metabolism.

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Isotopically labeled compounds originally introduced into experiment treatments are not necessarily stable. As incubations proceed over time, labeled substrates can be transformed into other compounds and be available to all entrained organisms in the experimental conditions. Thus the observed isotope-enriched parts in the foraminifera do not necessarily indicate the assimilation of the original labeled compounds, but could indicate the assimilation of other compounds metabolically derived from the original substrates. For example, ammonium or nitrate can be quickly metabolized into ammonium/nitrite/nitrate via microbial processes in the seawater or in sediments (Francis et al. 2007; Libes 2009). Thus the isotope enrichments observed in a long-term experiment requires careful interpretations. Short-term experiments, on the other hand, may be insufficient to label materials for TEM-NanoSIMS analyses that have lower sensitivity than conventional bulk isotope analyses such as EA/IRMS.

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Another limitation of the current TEM-NanoSIMS method is that the content of vacuoles and associated soluble compounds in the cytosol (e.g., free amino acids, inorganic carbon, or ions like nitrate and ammonium) are lost during sample preparation. This is unfortunate because these compounds are very important for assessing foraminiferal denitrification, calcification, assimilation and dissimilation processes. Cryofixation of foraminiferal cells (Goldstein and Richardson 2002, this issue) is a potential way to examine those soluble components in the foraminiferal cytosol. Cryofixation is a fixation protocol that preserves a sample with high-pressure freezing at extremely low temperature (Moor 1987). Recently, elemental mapping of the foraminiferal cytoplasm, vesicles and vacuoles using cryofixation reported presence of vacuoles containing calcium in seawater in a cryo-fixed *Amphistegina lobifera* (Khalifa et al, 2016). This is a good example that cryofixation maintains soluble compounds in vacuoles that may be lost during conventional sample preparation protocols. NanoSIMS analyses of cryopreserved specimens should provide more complete information on elemental and isotopic localizations in foraminiferal cells.

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From a logistical and resource perspective, TEM-NanoSIMS is a time-consuming method due to sample preparation steps, TEM observation, and isotope mapping using NanoSIMS. Additionally, the instrumentation is costly and can be difficult to access. These facts result in generally low numbers of analyzed TEM-NanoSIMS specimens per experimental condition, hampering generalization of results.

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## 4. Conclusion

The three TEM-related techniques provide a novel perspectives into foraminiferal ecology and biology on different spatial and temporal scales. FLEC-TEM provides sub-millimeter-scale information on foraminiferal life positions and microhabitats in terms of living depth and surrounding milieu including sediment fabric, porosity and meio- and microfaunal communities, all related to foraminiferal ultrastructure and fitness. Micro-X-ray CT-TEM provides 3D views of foraminiferal cells to assess cytoplasmic features such as vacuolization extent to provide volumetric and areal information useful to biomass/biovolume assessments. TEM-NanoSIMS provides micron to nanometer scale information on the distributions of elemental and isotopic compositions in foraminifera along with ultrastructural information, yielding important information about foraminiferal metabolism at the organelle to endobiont scale. Although each of these techniques has limitations, the advantage of correlation with TEM imaging provides different ways to link cell ultrastructures with metabolic processes, ultimately better defining foraminiferal roles in biogeochemical cycling and microbiome ecosystem functioning.

## 5. Acknowledgments

We thank Ms Kaya Oda (JAMSTEC) for the preparation of the Japanese TEM samples; Katsuyuki Uematsu and Akihiro Tame (Marine Works, Japan) for TEM observations; Akizumi Ishida, Yuji Sano, Naoto Takahata (The University of Tokyo) for NanoSIMS analyses; Chon Chen (JAMSTEC) for help with the onboard sampling of the sediment samples from the bathyal Sagami Bay during the KM16-01 cruise. The electron microscopy platform at the University of Lausanne (Switzerland) is thanked for access and technical assistance. JMB thanks her collaborators Tony Rathburn and Jon Martin on the original seep foraminifera project, Sam Bowser and Sue Goldstein for insightful conversations, Kasia Hammar and Louie Kerr for TEM assistance at the Marine Biological Laboratory (Woods Hole, MA, USA), and the Georgia Electron Microscopy facility at UGA. The comments raised from Prof. Howard Spero and an anonymous reviewer greatly improved our manuscript. This contribution was edited by Prof. Richard Jordan. This work was financially supported by the Grant-in-Aid for Scientific Research from the Ministry of Education, Culture, Sports, Science and Technology, Japan (Scientific Research (C) grant number 17K05697 to HN) and the Swiss National Science Foundation (grant no. 200021_149333). JMB's contributions were funded by US NSF grants OCE-0551001 and OCE-1634469, the WHOI Robert W. Morse Chair for Excellence in Oceanography, and The Investment in Science Fund at WHOI. The micro-X-ray CT imaging was performed under the cooperative research program of Center for Advanced Marine Core Research (CMCR), Kochi
686 University (accept No. 17A021).

687

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753 Dahlgren, L., 1967b. On the nuclear distribution of RNA and DNA and on the ultrastructure 754 of nuclei and adjacent cytoplasm of the foraminifers *Hippocrepinella alba* 755 Heron-Allen and Earland *Globobulimina turgida* (Bailey). Zool. Bidrag, Uppsala Bd 756 37, 113-138.

757 Ferrández Cañadell, C., 2002. Multicellular-like compartmentalization of cytoplast in fossil 758 larger foraminifera. Lethaia 35, 121-130 doi: 10.1111/j.1502-3931.2002.tb00073.x

759 Ferrández Cañadell, C., Briguglio, A., Hohenegger, J., Woger, J., 2014. Test fusion in adult 760 foraminifera: A review with new observations of an Early Eocene *Nummulites*
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Figure legends

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**Figure 1 (view in color).** Light micrographs of *Buliminella tenuata* in living position in non-seep sediments. A. Reflected light image showing excised portion of FLEC section that includes the foraminifer (white arrowhead). The black marking (white ^) is a permanent marker to guide in locating the specimen during excision. Note that the top of the sediment in the image is not the sediment-seawater interface (see text). B. LSCM micrograph showing the fluorescent specimen in sediments. Note the vacuolated nature of the cytoplasm, relatively empty ultimate chamber and veil of fluorescent material on the test exterior (arrows). sg1 and sg2 mark particular sediment grains that are also visible in C. C. Transmitted light micrograph of a thick section stained with toluidine blue, showing the specimen after sectioning. Note the holes where the sediment grains (sg1, sg2) occurred. t = test. * = open pore space adjacent to aperture. Scales: A = 500 µm; B, C = 100 µm.

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**Figure 2.** Transmission electron micrographs of *B. tenuata* collected from non-seep sediments. A. Overview of chamber periphery showing cytoplasm with large vacuoles (v) and degradation vacuoles (dv). po = pores; t = location of former test; white arrowhead = prokaryote in degradation vacuole. B. Internal edge of chamber showing mitochondria (m) and degradation vacuoles. g = Golgi. C. Periphery of chamber, tangentially through pores, showing mitochondria associated with pores and Golgi. ol= organic lining; eo= electron-opaque body; black arrow = mitochondrial double membrane. Inset: cross section through pore showing preservation of fibrous material. D. Chamber periphery showing mitochondria and Golgi. E. Peroxisomes (p) complexed with endoplasmic reticulum (er), showing darker core (inset). F,G. Images taken outside the foraminiferan test, which existed in the light-colored spaces. Areas of interest are the blebs (*), which likely were reticulopods that produced the fluorescence on the test exterior in Fig. 1B. Scales: A = 2 µm; B,G = 1 µm; C-F = 0.5 µm.

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**Figure 3 (view in color).** Light micrographs of an agglutinated biserial foraminifer in living position in seep sediments. A, B. Paired images showing specimens in life position in sediments preserved with FLEC. A. Double exposure micrograph of epifluorescence and reflected light showing foraminifer in life position 1.3 mm below the sediment-water interface and close to a juvenile bivalve (+). White and black arrow = cross section of a calcareous foraminifer. B. Epifluorescence micrograph revealing microbes (fluorescent items including prokaryotes and/or eukaryotes; some are labeled with white arrowheads) shown in this image. Note the calcareous foraminifer noted in A is not fluorescent (i.e., dead). C. LSCM micrograph showing the vacuolated nature of the cytoplasm, nucleus (black *), relatively empty ultimate
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(youngest) chamber (#), feeding bolus or reticulopodial trunk near the aperture (white arrow), and mineral grains in test (white *). Note the fine filaments in the adjacent sediments (white arrowheads). t = test. Scales: B = 1 mm (A = same magnification as B); C = 200 µm.

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**Figure 4.** Transmission Electron Micrographs of a biserial agglutinated foraminifer collected from seep sediments. A. Overview, including portion of cell periphery with organic lining (ol), showing cytoplasm with large empty vacuoles (v) and some appearing to include 1-2 prokaryotes (black arrows) and digestive vacuoles (dv). Fractured mineral grains from test (t) are evident; empty (white) space is space formerly occupied by sediment grains (i.e., foraminiferal test). B. Overview showing prokaryotes (arrows) occurring in the lumen between organic lining and interior surface of test. C. Area showing what are interpreted to be mitochondria (m), along with vacuoles and organic lining. Some mitochondria have faintly visible double membranes (white arrowhead). D. More mitochondria with faint double membranes and cristae (white arrowheads), along with peroxisomes (p) and endoplasmic reticulum (er). Scales: A,B = 2 µm; C = 1 µm; D = 0.5 µm.

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**Figure 5 (view in color).** Micro-X-ray CT slice images of *Ammonia* sp. (phylotype T6) incubated under dysoxic condition. A. Reconstructed micro-X-ray CT image of the *Ammonia* sp. White arrowhead shows a depressed cytoplasm in the chamber. B. Cross-sectional view of the CT image indicated as b1'-b2' to b1"-b" in A. C. A cross-sectional view from the spiral side. D. A cross-sectional view of the CT image indicated as d1'-d2' to d1"-d2" in A. In the panels B, C, and D, white indicates more X-ray absorbance, meaning higher electron density. Note that this specimen was sectioned for TEM observation before carrying out the micro-X-ray CT imaging. Scale bars = 100 µm.

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**Figure 6 (view in color).** Micro-X-ray images of *Bolivina spissa* collected from the bathyal Sagami Bay. A. Reconstructed micro-X-ray CT image of the *Bolivina spissa*. B. A view from the aperture side indicated as the arrow b in A. C. A view from the proloculus side indicated as the arrow c in A. D. A cross-sectional view of the *Bolivina spissa* indicated as the plane d in A. Many vacuoles (v) and a part of plug (pl) are visible. E. A cross-sectional view of the *Bolivina spissa* indicated as the plane e in A. In panels D and E, white indicates more X-ray absorbance, meaning higher electron density. Scale bars = 100 µm.

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**Figure 7 (view in color).** Examples of TEM-NanoSIMS analysis on *Ammonia* sp. (phylotype T6) incubated under dysoxic conditions. A. Osmium coated semi-thin sections of *Ammonia* sp. (phylotype T6) on a copper grid. Note that the TEM observations (B, C, D, and E) were carried out before the osmium coating. B. Low-magnification TEM image of the whole specimen. The square marked with c indicate the area where montage images of TEM were

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made. C. Montage images of TEM which were observed with ×3,500 magnification. The squares marked with d and e indicate the area where NanoSIMS analysis were performed. D and E. Correlative TEM images and NanoSIMS images on nitrogen mapping ($^{12}C^{14}N^-$ count per each pixel) and the ratio of $^{15}N/^{14}N$ ($^{12}C^{15}N^-$ count / $^{12}C^{14}N^-$ count per each pixel). m: mitochondria, li: lipid droplet, v: vacuole. D: 15 µm x 15 µm. E: 12 µm x 12 µm.

**Figure 8 (view in color).** Isotopic signal quantification on NanoSIMS images with the software Look@NanoSIMS (Polerecky et al., 2012). The example illustrated in this figure is a quantification of the $\delta^{13}C$ in the endoplasm of a species of *Ammonia* sp. (phylotype T6) which was fed with $^{13}C$-enriched diatoms. A: TEM micrograph of the endoplasm of an *Ammonia* sp. (phylotype T6) B: Corresponding $^{12}C^{14}N$ NanoSIMS image. C: Overlay of the TEM and $^{12}C^{14}N$ NanoSIMS images following the alignment procedure with the software Look@NanoSIMS; blue: TEM, red: $^{12}C^{14}N$ image. D: Drawing of regions on interest (ROIs) on the overlay image (here the ROIs are diatoms). The ROIs are numbered order from left to right. E: Application of the ROI template on the aligned $\delta^{13}C$ NanoSIMS image. F: Results compiled in a table by the software, each line corresponds to one ROI: identification (#i), positions x and y of the ROI central point (Xi and Yi), $\delta^{13}C$ enrichment value (MEANi, here in ‰), Poisson error (Poiss_Ei, theoretical precision of the mean), size: diameter in µm (SIZESi) and in pixels (PIXELSi), and classification (here "d" for diatom). d: diatom. Scale bars: 5 µm.

**Figure 9 (view in color).** TEM micrographs (left) and corresponding NanoSIMS images (right) of the endoplasm of different foraminiferal species incubated with different sources of isotopes. A. *Ammonia* sp. (phylotype T6) fed with a $^{13}C$-enriched diatom biofilm. The $^{13}C$ signal could be traced in the diatoms ingested inside the foraminiferal endoplasm (for more details see LeKieffre et al. in press). B. *Haynesina germanica* incubated with $H^{13}CO_3^-$ as a source of inorganic carbon. C. *Nonionellina labradorica* incubated with $^{15}NH_4^+$ as a source of inorganic nitrogen. D. *Ammonia* sp. (phylotype T6) incubated with $^{15}NO_3^-$ as a source of inorganic nitrogen in anoxia. E. *Ammonia* sp. (phylotype T6) incubated with $^{34}SO_4^{3-}$ as a source of inorganic sulfur in dysoxia. White arrowheads indicate the labeled isotope localization in the foraminiferal cytoplasm and the black arrowheads indicate the corresponding region in the TEM images. d: diatom, n: nucleus, v: vacuole, c: chloroplast, m: mitochondria, ob: electron opaque body, eb: elongated body, dv: degradation vacuole.

**Supplementary movie 1.** Reconstructed 3-D micro-X-ray CT image of *Ammonia* sp. (phylotype T6), obtained with the Xradia 410 Versa microfocus X-ray CT system. Note that this specimen was sectioned for TEM observation before conducting micro-X-ray CT imaging. The ultimate 3 to 4 chambers look frothy (less densely packed) in comparison to
other chambers. Among the ultimate 3 to 4 chambers, the outer portions of the chambers appear denser than the inner portions (closest to the axis of rotation), reflecting what appears to be more vacuoles nearer the aperture of this species. Particles showing high X-ray attenuation are (mineral) particles originally attached on the umbilical part outside the calcite test.

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**Supplementary movie 2.** Reconstructed 3-D micro-X-ray CT image of *Bolivina spissa*, obtained with the Xradia 410 Versa microfocus X-ray CT system, showing external surface of the cytosol, together with large vacuoles.

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**Supplementary movie 3.** Reconstructed 3-D micro-X-ray CT image of the same *Bolivina spissa* specimen in Supplementary movie 2, but the obtained image was processed to visualize the inner structure. In the cross section, plugs separating cytoplasm between chambers, shown in red (high densities), were clearly observed through the entire cross section. Note that the color contrast was altered from Supplementary movie 2 to better illustrate the plugs.

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A

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B

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C

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<figure><img src="image_20.png" /><figcaption>Figure 5</figcaption></figure>
<figure><img src="image_21.png" /><figcaption>Figure 6</figcaption></figure>
<figure><img src="image_22.png" /><figcaption>Figure 7</figcaption></figure>
<figure><img src="image_23.png" /><figcaption>Figure 8</figcaption></figure>

F: Results example

<table><thead><tr><th>#i</th><th>Xi</th><th>Yi</th><th>MEANi</th><th>Poiss_Ei</th><th>SIZEi</th><th>PIXELSi</th><th>LWRatio</th><th>Class</th></tr></thead><tbody><tr><td>1</td><td>19.7</td><td>102.57</td><td>5.24E+02</td><td>2.33E+00</td><td>5.53</td><td>1750</td><td>2.37 d</td><td></td></tr><tr><td>2</td><td>20.12</td><td>187.37</td><td>6.15E+02</td><td>3.25E+00</td><td>4.39</td><td>1101</td><td>1.48 d</td><td></td></tr><tr><td>3</td><td>36.55</td><td>153.26</td><td>2.94E+02</td><td>1.78E+00</td><td>4.27</td><td>1042</td><td>1.45 d</td><td></td></tr><tr><td>4</td><td>52.36</td><td>98.96</td><td>5.25E+01</td><td>5.15E-01</td><td>2.93</td><td>490</td><td>1.09 d</td><td></td></tr><tr><td>5</td><td>69.63</td><td>125.67</td><td>5.39E+01</td><td>4.90E-01</td><td>3.13</td><td>561</td><td>1.17 d</td><td></td></tr><tr><td>6</td><td>87.33</td><td>103.72</td><td>8.72E+01</td><td>7.83E-01</td><td>3.03</td><td>526</td><td>1.38 d</td><td></td></tr><tr><td>7</td><td>88.91</td><td>169.81</td><td>8.89E+01</td><td>4.26E-01</td><td>5.74</td><td>1886</td><td>3.22 d</td><td></td></tr><tr><td>8</td><td>95.39</td><td>205.81</td><td>3.59E+02</td><td>3.47E+00</td><td>2.52</td><td>363</td><td>1.25 d</td><td></td></tr><tr><td>9</td><td>109.7</td><td>131.24</td><td>4.41E+02</td><td>2.32E+00</td><td>4.47</td><td>1143</td><td>1.35 d</td><td></td></tr><tr><td>10</td><td>127.26</td><td>190.31</td><td>8.05E+01</td><td>6.46E-01</td><td>3.39</td><td>657</td><td>1.16 d</td><td></td></tr><tr><td>11</td><td>139.52</td><td>87.55</td><td>8.24E+01</td><td>5.03E-01</td><td>4.45</td><td>1135</td><td>1.83 d</td><td></td></tr><tr><td>12</td><td>151.32</td><td>172.55</td><td>1.39E+02</td><td>1.04E+00</td><td>3.47</td><td>689</td><td>1.93 d</td><td></td></tr><tr><td>13</td><td>165.31</td><td>229.7</td><td>5.58E+02</td><td>4.02E+00</td><td>3.13</td><td>561</td><td>1.58 d</td><td></td></tr><tr><td>14</td><td>178.91</td><td>166.39</td><td>1.34E+02</td><td>6.65E-01</td><td>5.21</td><td>1553</td><td>2.27 d</td><td></td></tr></tbody></table>
<figure><img src="image_24.png" /><figcaption>Figure 9</figcaption></figure>
<table><caption>Table 1. Incubation periods and stable isotope labels used for TEM-NanoSIMS analyses.</caption><thead><tr><td>Species</td><td>Incubation period</td><td>Added isotope label</td></tr></thead><tbody><tr><td>Ammonia sp. phytotype T6<sup>1</sup></td><td>24 h</td><td><sup>13</sup>C-enriched diatom biofilm</td></tr><tr><td>Haynesina germanica<sup>1</sup></td><td>16 h</td><td>NaH<sup>13</sup>CO<sub>3</sub></td></tr><tr><td>Nonionellina labradorica<sup>1</sup></td><td>7 h</td><td><sup>15</sup>NH<sub>4</sub>Cl</td></tr><tr><td>Ammonia sp. phylotype T6<sup>2</sup></td><td>14 d</td><td>NaH<sup>13</sup>CO<sub>3</sub>, Na<sup>15</sup>NO<sub>3</sub>, Na<sub>2</sub><sup>34</sup>SO<sub>4</sub></td></tr></tbody></table>

<footnote n="1">¹ LeKieffre et al. (2017 and unpublished data)</footnote>

<footnote n="2">² Nomaki et al. (2016)</footnote>
<table><caption>Table 2. NanoSIMS measurement settings of Nomaki et al. (2016) and LeKieffre et al. (2017 and unpublished data).</caption><thead><tr><th></th><th><sup>13</sup>C/<sup>12</sup>C, <sup>15</sup>N/<sup>14</sup>N analyses<sup>1</sup></th><th><sup>15</sup>N/<sup>14</sup>N analyses<sup>2</sup></th><th><sup>34</sup>S/<sup>32</sup>S analysis<sup>2</sup></th></tr></thead><tbody><tr><td>Measurement area</td><td>15×15 or 30×30 µm<br>(256×256 pixels)</td><td>Typically 8×8 to 15×15 µm<br>(256×256 pixels)</td><td>Typically 8×8 to 16×16 µm<br>(128×128 or 256×256 pixels)</td></tr><tr><td>Primary beam</td><td>about 2 pA Cs<sup>+</sup> beam</td><td>about 1 pA Cs<sup>+</sup> beam</td><td>about 5 pA Cs<sup>+</sup> beam</td></tr><tr><td>Primary beam focus size</td><td>120-150 nm</td><td>&lt; 600 nm</td><td>&lt; 1000 nm</td></tr><tr><td>Primary beam dwell-time</td><td>5 milliseconds/pixel</td><td>&lt; 1 millisecond/pixel</td><td>&lt; 1 millisecond/pixel</td></tr><tr><td>Mass resolution</td><td>8000-10000 at 10% peak height</td><td>&gt; 8000 at 10% peak height</td><td>&gt; 8000 at 10% peak height</td></tr><tr><td>Numbers of sequential image at 1 site</td><td>6 to 10</td><td>Typically 50</td><td>Typically 20</td></tr></tbody></table>

<footnote n="1">¹ LeKieffre et al. (2017 and unpublished data)</footnote>

<footnote n="2">² Nomaki et al. (2016)</footnote>