Main

Phosphorus (P) is an important constituent of nucleic acids (DNA, RNA), phospholipids, phosphoproteins and adenosine triphosphate (ATP), which is responsible for the biological transmission of chemical energy2,3,4. Thus, P is essential for all living organisms on Earth. The main form of dissolved inorganic P in the ocean is phosphate, which is an important macronutrient5 and widely used fertilizer that can cause eutrophication and deoxygenation in coastal ecosystems6. One recent study showed that benthic foraminifera from the Peruvian Oxygen Minimum Zone (OMZ) accumulate large amounts of phosphate within their cells1. Their intracellular phosphate concentrations can exceed concentrations in the surrounding seawater by 100–1,000-fold, and their high intracellular phosphate storage is probably involved in the genesis of phosphorites in the Peruvian OMZ1.

Benthic foraminifera are ubiquitous marine protists7. Several foraminiferal species can inhabit extreme O2-depleted environments due to their specific metabolic and morphologic adaptations8. These species might benefit from the current ongoing threat of ocean deoxygenation9. A well-studied and widespread adaptation of foraminifera to O2 depletion is the intracellular storage of nitrate for denitrification, which places them as a key element within the marine nitrogen (N) cycle10,11. Foraminiferal denitrification is partly eukaryotic12,13,14, partly complemented by bacterial symbionts15 and, for some species, is the preferred metabolic pathway over O2 respiration16. Nevertheless, not all foraminifera that inhabit O2-depleted environments can denitrify8 or have other known adaptation mechanisms such as fermentation13,14.

Accumulation of P is also speculated as a common and important foraminiferal adaptation mechanism to O2-depleted environments1. However, the regional distribution and metabolic functions of phosphate storage and subsequent use in foraminifera are poorly understood1. This study aims to analyse the occurrence of foraminiferal phosphate storage from diverse marine environments, and the metabolic functions of the unusually high intracellular phosphate reservoir. Finally, the importance of foraminifera for marine P cycling is discussed.

Intracellular phosphate content was measured in different foraminiferal species from diverse marine environments: an intertidal mudflat at the German Wadden Sea (Friedrichskoog), a seasonal hypoxic fjord basin (Bedford Basin), the seasonally hypoxic Sagami Bay (Japan) and the Mid-Atlantic Ridge (MAR). All analysed benthic foraminifera, except Chilostomella ovoidea from Sagami Bay and some species from the MAR, showed elevated intracellular phosphate levels (Fig. 1, Table 1 and Supplementary Table ST4), indicating the widespread occurrence of foraminiferal phosphate storage. The highest intracellular phosphate concentration was found in the species Ammonia confertitesta from the German Wadden Sea.

Fig. 1: Intracellular phosphate storage in benthic foraminifera from diverse marine environments.
Fig. 1: Intracellular phosphate storage in benthic foraminifera from diverse marine environments.
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a, Species sorted by mean intracellular phosphate concentrations. Column colour indicates the sampling region; n represents the number of biologically independent samples measured for each species. Each sample contained between one and 75 individuals, depending on the size of the species (Supplementary Table ST2). Grey crosses indicate data for individual samples. Error bars are 1 s.d. b, Log–log plot of individual intracellular phosphate content versus mean cell volume of the sample. Each data point represents one sample. Depending on the average body size of the species, each sample contained between one and 75 individuals (Supplementary Table ST2). Black dashed line indicates the correlation found for foraminifera from the Peruvian OMZ1. Anox., anoxic incubation; ox., oxic incubation.

Table 1 Intracellular phosphate storage and mean cell volumes for benthic foraminifera from diverse marine environments
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For assessment of the metabolic function of phosphate storage, intracellular P distribution was mapped in two species from Japan (Ammonia veneta from a cultured strain17 and Bolivina spissa from bathyal Sagami Bay), using energy-dispersive X-ray spectroscopy (EDS) under scanning electron microscopy operated at cryogenic temperature (cryo-SEM). Phosphorus was accumulated, together with calcium (Ca), in cellular structures of size 0.5–2.0 µm (Figs. 2 and 3 and Extended Data Figs. 1 and 2), potential acidocalcisomes based on their elemental compositions, size and appearance. In other organisms, acidocalcisomes are usually also enriched in P and Ca, sometimes containing polyphosphate gels and granules of spherical shape18. The size of acidocalcisomes varies depending on the organism. The typical diameter for protists is 0.4–0.6 µm (ref. 18), but diameters over 1 µm are not uncommon19. Furthermore, P compounds extracted from A. confertitesta were characterized, using 31P nuclear magnetic resonance (NMR) spectroscopy. The data indicate that a large proportion of the extracted phosphate most probably originates from ATP, pyrophosphate and other polyphosphates (Supplementary Note SN1 and Supplementary Figs. SF1 and SF2). The presence of enzymes in polyphosphate metabolism for some benthic foraminifera has previously been shown1. In addition, due to the recent finding of a creatine phosphate metabolism in foraminifera from anoxic Namibian shelf sediments13, the published genomes and transcriptomes of A. confertitesta, A. veneta, Globobulimina pacifica and Reticulomyxa filosa have been screened for homologues of creatine kinase, which was present in all cases (Supplementary Fig. SF4). Creatine phosphate and polyphosphates are both highly energetic, and can be used as an energy source under electron acceptor depletion20,21. Thus, elevated phosphate storage in foraminifera from O2-depleted habitats is probably another adaptation to O2 depletion. In addition, the ubiquitous occurrence of phosphate storage and accumulation of P in organelles that are possibly acidocalcisomes indicates diverse metabolic functions of this intracellular phosphate storage (Supplementary Fig. SF3).

Fig. 2: Elemental maps of B. spissa cross-section.
Fig. 2: Elemental maps of B. spissa cross-section.
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a–d, Cryo-EDS mappings of P and Ca and cryo-SEM images on cross-section of a cryo-fractured specimen of B. spissa. a, P distribution, mapped with cryo-EDS, overlain by the cryo-SEM image of the same region. b–d, Overview of the distribution of P (b), Ca (c) and Mg (d). Arrows indicate structures enriched in both Ca and P, and often in Mg. *, Structures enriched in P and Ca but not in Mg. The cryo-SEM–EDS experiment was repeated on three different specimens of B. spissa. Scale bar, 20 μm.

Fig. 3: Ultrastructure of A. veneta specimens.
Fig. 3: Ultrastructure of A. veneta specimens.
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a, Cryo-SEM image (secondary electron mode) of cross-section of a cryo-fractured specimen of A. veneta. Note the abundant circular structures (denoted by red arrows) that are possibly acidocalcisomes. b, P distribution, mapped with cryo-EDS, is shown in green on a cryo-fractured specimen of A. veneta, overlain by the cryo-SEM image of the same region (SEM image and P distribution are shown individually in Supplementary Fig. SF2). c,d, The circular, slightly P-enriched structures are absent on TEM images of thin sections of A. veneta. Instead, there are abundant empty vesicles visible on the thin sections, which indicates that these structures might have lost their content during fixation and subsequent embedding and polymerization processes. Only one of these vesicles has been found that was not empty (marked with the red arrow in d). This structure was the only structure with measurable P content using coupled TEM–EDS (Extended Data Fig. 4). The cryo-SEM–EDS experiment was repeated on eight different specimens of A. veneta, and the TEM–EDS experiment on three different specimens. Scale bars, 5 μm (a), 2 μm (b,d) and 50 μm (c).

Widespread foraminiferal phosphate storage

These newly acquired data regarding intracellular phosphate storage in benthic foraminifera (Fig. 1 and Table 1) cover a wide range of environments, from shallow marine habitats (for example, the tidal mudflat off Friedrichskoog) to the deep sea (Rainbow Vent Field at MAR). Redox conditions vary from well oxygenated (MAR) to permanently anoxic (Peruvian OMZ centre; Supplementary Table ST1).

Elevated intracellular phosphate content has been found in species from all sampling regions. Only C. ovoidea from Sagami Bay and some species from the MAR did not show elevated intracellular phosphate accumulations. Species from the MAR had very low phosphate storage in general, possibly linked to well-oxygenated environments. Elevated phosphate content at the MAR was found only in Hoeglundina elegans and Rhizammina algaeformis (Table 1). Nevertheless, these two species are relatively large and, normalized to cell volume, intracellular concentrations are still very low (Fig. 1a and Table 1). Concentrations at the Peruvian OMZ, Sagami Bay and the tidal mudflats of Friedrichskoog were comparable, with a slight tendency to higher concentrations at the Peruvian OMZ, where bottom-water O2 concentrations were lowest (Fig. 1 and Supplementary Table ST1). The tendency for higher phosphate storage in more O2-depleted environments supports the premise that elevated intracellular phosphate content is probably an adaptation to O2 depletion.

Specimens of A. confertitesta, having the highest intracellular phosphate concentrations measured, were sampled at the intertidal mudflats of Friedrichskoog. Oxygen penetration depth in intertidal mudflats, which are usually rich in organic matter, can be as shallow as 1 mm (ref. 22). Species of the genus Ammonia cannot denitrify8, but can actively feed and thrive down to a sediment depth of several centimetres23. In addition, to date, no species of the genera Elphidium and Haynesina, which are typically found together with Ammonia spp. in O2-depleted sediment, have been found to denitrify8. This suggests that these taxa must have alternative adaptation mechanism(s) available to cope with O2 depletion. Compared with A. confertitesta, both Elphidium bartletti and Haynesina germanica have moderately high intracellular phosphate concentrations. Whereas kleptoplasty might be a survival strategy under O2 depletion in the case of Elphidium and Haynesina8,24, sequestered chloroplasts in Ammonia spp. are usually dysfunctional25.

Thus, Ammonia spp. must have an alternative survival strategy under anoxic conditions. A. confertitesta is a putative invasive species in shallow European shelf sediments, replacing indigenous Ammonia spp. in these environments, and probably originated from Asia26,27. Thus, it is possible that the high intracellular phosphate storage of A. confertitesta could provide a competition advantage over other autochthonous species in these environments. Bolivina seminuda and Bolivina costata from the Peruvian OMZ showed high phosphate storage, comparable to that of A. confertitesta. These Bolivina species can dominate anoxic and even sulfidic habitats28,29, and are able to denitrify11,16. This suggests that the ability to denitrify and high phosphate storage, as adaptations to O2 depletion, are not mutually exclusive, and provide even further competitive advantages for thriving in environments in which oxygen is low or absent.

Diverse metabolic functions

Newly acquired coupled cryo-SEM–EDS data acquired for A. veneta and B. spissa indicate that phosphate storage is related to additional metabolic functions other than being an adaptation to O2-depleted conditions. Analyses on cells of B. spissa indicate that P is accumulated with Ca, and sometimes with Mg, in solid, granule-like structures (Fig. 2 and Extended Data Fig. 2) of appearance similar to other Ca polyphosphates30,31. In A. veneta, P is accumulated in round structures 0.5–2.0 µm in diameter (Fig. 3). Phosphorus seems to be less locally concentrated in A. veneta than in B. spissa, and not in the form of solid Ca polyphosphates, which indicates a generally lower intracellular phosphate content in A. veneta compared with B. spissa. This is supported by the intracellular phosphate concentrations that have been measured for those species, which were also lower for A. veneta (Fig. 1 and Table 1). These round structures, which are abundant within cells of A. veneta and that have been imaged with cryo-SEM (Fig. 3a,b and Extended Data Fig. 1), are similar in size and shape to acidocalcisomes19, organelles known to accumulate pyrophosphate (diphosphate), and to granules that are enriched in Ca and polyphosphates and other metals such as Mg. Nevertheless, we cannot exclude the possibility that some of these structures are autophagosomes that can also accumulate polyphosphates19. Autophagosomes have a similar size and shape, although they usually show the presence of membranous debris and have a more irregular shape19. Digestive food vacuoles known from foraminifera have a completely different size and shape32 and can most probably be excluded. Putative acidocalcisomes are either absent or empty in transmission electron microscope (TEM) images on thin sections of A. veneta, because abundant and empty round vesicles of the same size are visible (Fig. 3c,d and Extended Data Figs. 3 and 4). During conventional preparation methods of thin sections for TEM imaging, cell fixation involves a dehydration step, decreasing the water content and resulting in the loss of soluble content within the fixed specimen33. In addition, no P accumulations could be found using coupled TEM–EDS on the same thin sections, except in one vesicle that was still filled (Fig. 3d and Extended Data Figs. 3 and 4). When acidocalcisomes are observed with TEM, cryofixation is generally used rather than the traditional thin-section preparation for TEM19. Nevertheless, it has already been recorded that, even when using cryo-TEM, it can be complicated to distinguish acidocalcisomes from other vesicles and vacuoles19. The presence in TEM observations of many empty vesicles of size and shape similar to the round, P-rich structures seen on cryo-SEM–EDS images, gives rise to speculation that many of the empty vesicles observed in A. veneta using TEM are the same structures as the filled ones on cryo-SEM observations, and may possibly be acidocalcisomes.

Acidocalcisomes, which are common in prokaryotes and eukaryotes34 and abundant in many protists19 have never previously been described in foraminifera, despite a long history of ultrastructural analyses of foraminiferal cells32. This is probably on account of the artefacts seen during the non-cryogenic preparation of thin TEM sections described above. Acidocalcisomes can have diverse functions, including the storage of polyphosphate and pyrophosphate and participation in the related energy metabolism, intracellular pH and osmoregulation and Ca storage for the conservation of Ca homeostasis18. If the observed structures are acidocalcisomes, or have a similar function, this would have several implications that could explain why high phosphate storage might provide crucial advantages for species that can accumulate more phosphate than others, including B. costata and A. confertitesta (Fig. 1), as shown below.

  1. (1)

    Rapid osmoregulation. Rapid hydrolysis or synthesis of polyphosphates in acidocalcisomes, which increases or decreases the intracellular electrolyte concentration, has been shown as a reaction to hypo- or hyperosmotic stress in Trypanosoma cruzi, a protist belonging to Excavata35. Both A. confertitesta and B. costata are often found in shallow marine environments that are strongly influenced by tidal cycles26,27,29, and, thus, undergo marked salinity changes, which suggests that this mechanism might be advantageous for rapid osmoregulation.

  2. (2)

    Rapid generation of metabolic energy. Because A. confertitesta and B. costata, as well as other benthic foraminiferal species, are often exposed to anoxia, the storage of energy-rich polyphosphates most probably provides a source of rapidly available energy when these organisms run out of terminal electron acceptors, such as O2 or nitrate. Due to their energy-rich P bonds, polyphosphates are a powerful energy source30, and sulfur bacteria from O2-depleted environments have previously been shown to utilize polyphosphates as an energy source when depleted of of electron acceptors36. Previous comparative genomics analyses have indicated the presence of a polyphosphate metabolism in multiple species of foraminifera1.

  3. (3)

    Intracellular pH regulation with possible relevance for calcification. Most of the foraminifera analysed in this study have calcareous tests. The foraminiferal calcification mechanism involves the elevated concentration of carbonate ions in vacuoles with elevated pH37,38,39. Acidification of acidocalcisomes is driven by vacuolar proton pumps18 and elevates pH in other cell compartments that might be used in the foraminiferal calcification pathway. Thus, if the organelles in foraminifera described above are indeed acidocalcisomes, they might play an important role in the foraminiferal calcification mechanism, with high intracellular phosphate storage probably providing advantages in regard to intracellular pH regulation for calcification. Previous studies show that pH regulation in acidocalcisomes can be crucial in some organisms to adapt to changes in environmental pH40. There are often strong pH gradients in pore waters of tidal flat sediments, within a few millimetres41. Thus, species such as A. confertitesta often undergo rapid pH changes in their microenvironment, and high phosphate storage is probably advantageous as an adaptation to such changes.

Despite the relevance of phosphate storage in acidocalcisomes, the metatranscriptome of foraminifera from a natural anoxic sediment environment off Namibia showed that they encode genes for a creatine phosphate metabolism13. New comparative genomics and transcriptomics analyses on the species A. confertitesta, A. veneta, G. pacifica and R. filosa show that these species also possess and/or express the enzyme creatine kinase and, thus, are able to synthesize and metabolize creatine phosphate (Supplementary Note SN4 and Supplementary Fig. SF4). Creatine phosphate is highly energetic, can rapidly regenerate ATP from ADP and can provide homeostasis of cellular bioenergetics42. Especially in muscle cells, creatine phosphate can rapidly provide energy during periods of elevated activity, or even bursts with high energetic demand43, and is an important contributor at the beginning of anaerobic muscle metabolism44. Thus, it has previously been suggested that creatine phosphate metabolism, observed in foraminifera from anoxic sediments at the Namibian Shelf13, provides energy storage for sudden energetic bursts13, such as feeding by phagocytosis13—that is, vacuolar ingestion of food particles and prey13. Because creatine kinase is encoded in the genome of A. confertitesta (Supplementary Note SN4 and Supplementary Figs. SF4 and SF5), creatine phosphate storage probaby confers on A. confertitesta the possibility of staying active even when exposed to anoxia. Creatine phosphate might even be utilized for energy bursts that are required for the predatory activity of this species that has been observed even under anaerobic conditions several centimetres below the sediment–water interface23. Thus, polyphosphates within acidicalcisomes and creatine phosphate are most probably additional adaptations of certain foraminiferal species to oxygen depletion. Although the metabolic functions of intracellular phosphate storage in foraminifera might be quite diverse (for example, energy supply under O2 depletion, regulation of intracellular pH and electrolyte concentration), it is easy to explain why deep-sea foraminifera from the MAR show lower intracellular phosphate accumulations. Except in some extreme examples, environmental conditions in the well-oxygenated deep sea are very stable, and organisms in this environment rarely experience severe O2 depletion or marked fluctuations in pH or salinity.

Finally, initial exploratory 31P-NMR data on P compounds extracted from A. confertitesta confirm the presence of high levels of pyrophosphate (Fig. 1 and Supplementary Note SN1). In addition, ATP was present in high concentrations and smaller amounts of inorganic polyphosphates were probaby extracted, too, although these are overlain by ATP peaks (Fig. 1 and Supplementary Note SN1). One peak in the 31P-NMR spectrum is close to that of creatine phosphate, but slightly drifted. The 31P-NMR spectrum measured directly on living specimens of A. confertitesta is much more complex and indicates a mixture of more diverse P compounds. Thus, the presence of creatine phosphate in the extracted P compounds should be confirmed in future studies.

Relevance for P cycling and the environment

It is well documented that benthic foraminifera performing denitrification are important for the marine nitrogen cycle, and even outcompete denitrifying bacteria in some O2-depleted environments due to their high abundance8,11,16,45. One recent study showed that the intracellular phosphate concentration in foraminifera from the Peruvian OMZ can be 100–1,000 times higher than in the surrounding pore waters, making them an important and previously overlooked reservoir for benthic inorganic P (ref. 1). That study also indicates that high intracellular phosphate content facilitates the nucleation of phosphorites in this environment1, in a way similar to how certain sulfur bacteria initiate phosphogenesis in regard to their high accumulation of polyphosphates36,46. The parallel enrichment of polyphosphates and Ca in the putative acidocalcisomes (Fig. 2) supports this hypothesis, because phosphorites are phosphatic rocks consisting, to a large degree, of apatite47,48. Rapid hydrolysis of polyphosphates post mortem might initiate apatite nucleation in favourable environments.

A rough extrapolation based on A. confertitesta density in Wadden Sea sediment during a bloom, and on individual intracellular phosphate content, suggests that the phosphate reservoir in A. confertitesta is huge (Supplementary Note SN6). The population density of living A. confertitesta during the bloom at Friedrichskoog was 417 individuals (ind) cm−3 (Supplementary Note SN6), and individual phosphate storage was 413 pmol ind−1. The Wadden Sea has an extension of 11,500 km2, resulting in a total amount of roughly 1,880 t phosphate stored in A. confertitesta within the top 1 cm of sediment. This represents about 5% of the yearly consumption of P-containing fertilizer in Germany in 2021/2022 (Supplementary Note SN6) within only a single foraminiferal species during the snapshot of a bloom, which provides a rough estimate of the importance of the entire group of foraminifera for oceanic P cycling.

A more detailed analysis of benthic foraminiferal phosphate storage in sediments from 135 stations in the Southern North Sea (Supplementary Table ST6) and 35 at the Peruvian Margin28 (Supplementary Table ST7) shows that benthic foraminifera might be an effective buffer for riverine phosphorus runoff in these regions (Fig. 4, Supplementary Note SN8 and Supplementary Fig. SF6). Total foraminiferal phosphate storage in the region of interest of the Southern North Sea (0.0059 ± 0.0014 g m−2; 1 standard error of the mean (s.e.)) is equivalent to roughly 37 days of riverine phosphate runoff in this region (2,583 t yr−1). Foraminiferal assemblages between 10° S and 15° S off Peru store 0.0315 ± 0.0101 g m−2 (1 s.e.), and thus may buffer approximately 21 days of riverine total P runoff in this region. Note that, because data for total riverine phosphate runoff from this region were not available, budgets had to be calculated slightly differently by using, instead, riverine total P runoff (Supplementary Note SN8).

Fig. 4: Distribution of benthic foraminiferal storage.
Fig. 4: Distribution of benthic foraminiferal storage.
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a,b, Maps showing the distribution of benthic foraminiferal phosphate storage in the Southern North Sea (a) and off Peru (b). Benthic foraminiferal phosphate storage was estimated using the composition of living benthic foraminiferal assemblages from 135 stations in the Southern North Sea51,52,53,54,55,56,57 and 35 off Peru1,29 (for details of calculations, see Methods). Maps show the weighted mean. Circles and numbers indicate regional riverine phosphate58 runoff (a) and total P (ref. 59) runoff (b). More detailed budget calculations show that benthic foraminifera might buffer approximately 37 days of riverine phosphate runoff to the Southern North Sea, and roughly 21 days of riverine total P runoff to the Peruvian coast between 10° S and 15° S (Supplementary Note SN8). Maps were generated using Ocean Data View software60.

At the Peruvian OMZ, foraminiferal phosphate storage may be of particular importance. Phosphate from remineralized organic matter adsorbs to iron oxide layers under oxic conditions and is effectively trapped in oxic sediments, whereas it escapes to the water column and is efficiently recycled under markedly O2-depleted conditions49. The high abundance of benthic foraminifera in this region might dampen this phosphate recycling, reduce productivity and thus act as a negative feedback mechanism in the expansion of the OMZ that has been observed since the 1960s50.

Finally, these results indicate that benthic foraminifera can be an important buffer to counteract eutrophication. Denitrifying foraminifera already mitigate eutrophication by reducing reactive nitrate to non-reactive N2 gas. These new findings regarding phosphate storage in foraminifera indicate that these organisms also contribute to de-eutrophication related to their high phosphate accumulations, making them key mediators that probably buffer the effects of coastal eutrophication. Budget calculations of total phosphate stored in benthic foraminifera in the Southern North Sea and the Peruvian OMZ indicate that this phenomenon is also globally important. Nevertheless, reactive P stored in foraminifera is not completely removed from the environment, such as nitrate during denitrification. Thus, intracellular foraminiferal phosphate storage can be seen more as temporary withdrawal from the system—that is, a ‘hidden’ standing stock that is temporarily unavailable for primary production.

Methods

Sediment sampling

For sampling of living benthic foraminifera, samples were taken during various surveys at Sagami Bay (Japan), the Rainbow Vent Field at the MAR, the Bedford Basin (Canada) and the German Wadden Sea (Friedrichskoog). Samples at Sagami Bay were taken during a RV Kaimei cruise in September 2019 and on RV Yokosuka cruises in October 2022 and May 2023. Sediment was retrieved using a multicorer during the former and a push core during the latter at stations in central Sagamy Bay (NSB site; Supplementary Table ST2). Samples from the region around the Rainbow Vent Field were retrieved during RV Meteor cruise no. M176/2 in September 2021, using a multicorer at six stations (Supplementary Table ST2). Samples from the Bedford Basin were retrieved in March 2022 on board the dive vessel EastCom, using a multicorer at three stations (Supplementary Table ST2). Sediments from the intertidal mudflats in Friedrichskoog were retrieved manually at one station in November 2021 and May 2023. The top 1 cm of sediment was scraped off by spoon. Surface sediments from the brackish water salt marsh of Hirakata Bay, Yokohama (Japan) were collected in 2015 (Supplementary Table ST2), and isolated A. veneta strains were maintained in the laboratory.

Preparation of living foraminifera for intracellular phosphate analyses

The top 1 cm of sediment was directly wet sieved over a 125 or 250-µm mesh within a time frame of 2 h following core retrieval, using either filtered regional surface water or nitrate- and phosphate-free artificial seawater (ASW) at ambient salinity prepared using Red Sea Salt. Only certain samples from the Bedford Basin were sieved within 2 days following sampling, using filtered seawater from the Bedford Basin. Sediments retrieved from Friedrichskoog in March 2023 were directly sieved in the field, using surface water.

Living foraminifera specimens for intracellular phosphate analyses were wet picked from the coarse residue (over 250 µm for Friedrichskoog samples and over 125 µm for the remainder). In total, 34 samples were picked that included 15 different benthic foraminifera species. Specimens of A. veneta were obtained from the clonal strain cultured at the Japan Agency for Marine-Earth Science and Technology (JAMSTEC). For a description of culturing methods, see below. One A. veneta sample was prepared from oxic incubation and one from anoxic. Each foraminiferal sample contained between one and 75 living foraminifera specimens depending on the average size of the species (Supplementary Table ST2). Note that the sample of R. algaeformis contained only one large fragment of the organism, which was visibly filled with cytoplasm. All samples were photographed with a stereomicroscopic camera for subsequent determination of foraminiferal biovolume. Afterwards, foraminifera were cleaned and phosphate extracted using the methods described in ref. 1. Specimens were rinsed with phosphate-free ASW, prepared from Red Sea Salt, then transferred to centrifuge tubes with the lowest amount of ASW possible. Next, 3 ml of reverse osmosis water (with conductivity of 0.055 µS cm−1) was added to samples. Within the water, foraminiferal specimens were broken up using a clean pipette tip. A procedural blank underwent the same procedure without foraminiferal specimens, for blank corrections of NO3− analyses (13 procedural blanks in total). All samples were frozen at −20 °C for at least 2 h and subsequently thawed. This procedure was repeated three times. Freeze–thaw injuries damage plasma membranes and increase their permeability61,62. Subsequently, samples were filtered through sterile 0.2-μm cellulose acetate filters.

A slightly adapted protocol was used for the extraction of P compounds for 31P-NMR; these samples were retrieved from Friedrichskoog in March 2023. Two replicates were picked, each containing around 1,000 living specimens of A. confertitesta. Specimens were rinsed with phosphate-free ASW prepared from Red Sea Salt and subsequently transferred to microcentrifuge tubes with the lowest amount of ASW possible. Next, 600 µl of heavy water (D2O) with 1 M KOH was added to samples. Within the water, foraminiferal specimens were broken up using a clean pipette tip. Samples were frozen at −20 °C for at least 2 h and then thawed. This procedure was repeated three times. Subsequently, samples were centrifuged. A further sample was picked that contained roughly 1,500 specimens of A. confertitesta, which were transferred unharmed to a NMR tube containing D2O and with a salinity of 28.

Analyses of extracted intracellular phosphate

Filtered samples were analysed for total dissolved phosphate by segmented flow-injection analysis using a QUAATRO39 (Seal Analytical) autoanalyser, which included a XY2-autosampler unit, at GEOMAR Helmholtz Centre for Ocean Research Kiel (Kiel, Germany). Samples from the MAR were analysed, using the same methods, directly on board during RV Meteor cruise no. M176/2. The system set-up included four channels—for nitrate + nitrite, silicate, nitrite and phosphate—but only the phosphate data were used within this study. The method used for phosphate analysis corresponds to Q-064-05 Rev. 8 (developed by Nederlands Instituut voor Onderzoek der Zee; detection limit 0.004 µmol l−1 and described by QuAAtro Applications).

Biovolumetric determination of living foraminifera

Total foraminiferal cell volume of each species was estimated following previously published methods63. We assumed that internal test volume corresponds to 75% of total test volume and is completely filled with cytoplasm64. Methodology and equations used for precise biovolume estimation in several benthic foraminifera species are given in ref. 65 but, unfortunately, none of the species analysed in our investigation are listed in their study; therefore, the closest geometric shape was used for biovolume estimation (Supplementary Table ST3). In total, the biovolume of 850 foraminiferal specimens was determined.

Note that, for the two shapes related to B. spissa (cone with elliptic base) and C. wuellerstorfi (triaxial ellipsoid), the heights of the specimens would have been required, which were not visible on the images. The following approximations have been used instead: for B. spissa specimens, height was estimated by the previously determined mean height of the species (133 ± 7 μm)16; for C. wuellerstorfi, the average ratio of the shortest diameter on the spiral side to the height was determined (0.424 ± 0.029, 1 s.e.), using published images of the species66 (Supplementary Table ST5).

Determination of living abundance

One sample retrieved from Friedrichskoog in March 2023 was taken to determine the abundance and population density of A. confertitesta. For this sample, we used a square-shaped metal frame with a side length of 10 cm. Within this metal frame, the top 1 cm of sediment was scraped off and collected in a polyethylene bottle. Next, a mixture of ethanol and Rose Bengal (2 g l−1) was added to the bottle until ethanol concentration exceeded 70%. The jar caps were cleaned and applied tightly, the height of the sediment in the jar was marked and the jars stored for at least 14 days at room temperature until further analysis. Subsequently, stained samples were wet sieved over a 125-µm mesh, dried at around 40 °C and the jars filled with water up to the sediment fill mark level. The volume of water representing bulk sediment volume was measured in a graduated cylinder (approximate accuracy ±5 cm3). Subsequently, samples were split using a dry splitter, and specimens of A. confertitesta stained with vital raspberry red were counted under the microscope. Living stained foraminifera were fixed in plummer cells.

Culture of A. veneta

Cultures of A. veneta were from the same strain used by Ishitani et al.17 isolated first in 2015. Specimens were cultured in ASW at a salinity of 35, at 23 °C under 14/10 h light/dark cycles. The specimens used for this experiment were fed, frozen, dead Dunaliella salina (no. NIES-2257). We isolated five specimens with shell diameter 150–300 μm from subculture into 35-mm culture dishes with 5 ml of ASW, with culture for 4 days under both oxic and anoxic conditions. We cultured normally for oxic conditions in an AnaeroPack-Anaero, which can maintain 0% O2 and 15% CO2 for anoxic conditions.

Comparative genomics and metabarcoding

Creatine kinase homologues were identified with the KEGG KAAS tool67 (species: hsa, mmu, rno, dre, dme, cel, ath, sce, ago, cal, spo, ecu, pfa, cho, ehi, eco, nme, hpy, bsu, lla, mge, mtu, syn, aae, mja, ape, mbr, ddi, tet, smin, pti, ehx, gtt, ngr, tva, tbr, spar) as K00933. The following genomes and transcriptomes were screened for comparative genomic analysis:

Subsequently, published transcriptome data of Ammonia spp. were collected from the NCBI database for further identification of this taxon at species level by metabarcoding. Collected raw reads were quality filtered with FASTX-Toolkit 0.0.13 (ref. 69), and those with fewer than 50 bases—or that included ambiguous barcodes and showed poor quality (q-score <20)—were removed.

NMR spectroscopy

All NMR experiments were carried out on a Bruker 600 MHz Avance III HD spectrometer (14.09 T, 600.13 MHz for 1H, and 242.94 MHz for 31P) at 298 K in D2O. One-dimensional 31P{1H} spectra were obtained utilizing a 30° excitation pulse and relaxation delay of 1.0 s. The waltz16 sequence was implemented for proton decoupling. Spectra were acquired at a spectral width of 96,153.84 Hz and 65,536 time domain data points, by recording 1,024 scans for extracted samples and 3,584 for the sample containing live foraminiferal cells31. P chemical shifts were referenced to external phosphoric acid (external measurement). Data were acquired using TopSpin v.3.6.4, and all spectra were processed with Topspin v.4.1.4, applying zero filling and an exponential multiplication of the free induction decay with a line-broadening factor of 1.0 Hz.

Cryofixation for cryo-SEM–EDS

Bolivina spissa specimens were isolated from the topmost 1 cm of sediment directly onboard, immediately following sampling under a stereomicroscope; A. veneta specimens were picked from cultures. Cryofixation followed the protocols of ref. 70. For cryofixation of foraminifera, conductive glue comprising 30 wt% graphite oxide and glycerol was used with probe sonication, following the method described in ref. 71 with solvent modification. Glycerol was applied as a cryoprotectant and viscous dispersant to prevent sinking of foraminifera deep into the glue. The glue was pasted onto an aluminium rivet (diameter 3 mm), each foraminifer specimen was mounted on the glue using an eyelash brush and was then immediately frozen in semifrozen isopentane at −159.8 °C. The rivet was then mounted on an ultramicrotome (Ultracut S equipped with FCS, Leica Microsystems, operated at −130 °C) and the cross-section faced using a diamond knife (Diatome). Faced samples were stored in a container below −160 °C until required for cryo-SEM observation.

Cryo-SEM–EDS

In total, three B. spissa specimens (two from the October 2022 cruise and one from the May 2023 cruise) and eight A. veneta specimens were analysed. Cryo-SEM observation was performed on a Helios G4 UX (Thermo Fisher Scientific) equipped with a cryogenic stage and a cryopreparation chamber (catalogue no. PP3010T, Quorum Technologies). EDS analyses were performed on an Octane Elite Super (C5) (AMETEK), which was attached to the cryo-SEM (software TeamEDS, v.4.6.0052.0238). The sample was mounted on a transfer shuttle in liquid nitrogen, then vacuum transferred to the cryopreparation chamber. Water was sublimed at −80 °C for 8 min to expose the organelle structure, and Cr was then coated by magnetron sputtering at 20 mA for 60 s. Note that sublimation does not melt or sublime glycerol-based glue. We selected Cr because conventional Pt or Au sputtering causes overlap in EDS signals, including P; the K-lines of N and P appear at 0.392 and 2.013 kV, respectively, and the M-lines of Pt and Au at 2.048 and 2.120 keV, respectively. By contrast, the L-line of Cr appears at 0.573 eV, overlapping only with O, and no signals appear between it and its K-line at 5.414 keV. The sputtered sample was transferred to the SEM chamber and maintained below −140 °C; cross-sectional morphology was imaged by secondary electron at 2 kV with 50 V of antibias on the sample, and EDS mapping was performed at 20 kV without antibias.

Spectral treatment aiming to deconvolve signal from noise was performed on EDS elemental maps. Conventional quantitative EDS analyses use correction by atomic number (Z), absorption (A) and X-ray fluorescence (F), called the ZAF method, assuming that the surface is completely flat and that elemental composition, along with depth direction, is homogeneous. However, our cryo-SEM and cryo-EDS maps did not meet these requirements and the apparent EDS maps were correlated with background, so that we could see the similarity between count-per-second (CPS) maps and the EDS maps of low-intensity atoms. Therefore, we tried to suppress the position-dependent background signal72. EDS maps are generated from the number of counts in which X-rays from atom A appear, and count IA is composed of pure signal SA and background BA, where background is mostly due to the bremsstrahlung effect. Because CPS is the sum of signal from all atoms and background, \({\rm{CPS}}={\Sigma }_{i}{S}_{i}+B={\Sigma }_{i}{I}_{i}+{B}_{\bar{i}}\), where i indicates the atom of interest, B is the summation of background intensity for all of the energy and Bi is the summation of background intensity at energy ranges in which characteristic X-ray peaks are absent. The background mostly derives from bremsstrahlung and is thus dependent on beam condition, and here we assume that background shape is the same in any position of EDS maps. The intensity-to-noise ratio (R) of atom i at position (m, n) is defined as equation (1):

$$\begin{array}{c}R({\rm{m}},{\rm{n}})={I}_{i}({\rm{m}},{\rm{n}})/{B}_{i}({\rm{m}},{\rm{n}})={S}_{i}({\rm{m}},{\rm{n}})/{B}_{i}({\rm{m}},{\rm{n}})+1\\ \,\,\,\,=\,{I}_{i}({\rm{m}},{\rm{n}})/({\rm{CPS}}-\sum _{{\rm{i}}}{I}_{i}({\rm{m}},{\rm{n}}),\end{array}$$
(1)

which is related to conventional signal-to-noise ratio Si/Bi. We used R to emphasize localized minor elements hidden under the strong background.

Finally, colocalization of SEM images and the EDS elemental map was performed manually using calcium distribution maps, the main compound of the test in calcitic foraminifera. When needed, we performed EDS map scaling and/or rotation without deformation (that is, warping). For enhanced visualization of elemental distribution on EDS maps, a 16-colour look-up table was applied on EDS maps without grey value modification (Fig. 2c). Then, for all elements (excluding Ca), one-pixel median filtering was performed to smoothe the elemental distribution map and identify enriched ultrastructures regarding the element of interest. Image treatment was performed using the software Fiji73.

TEM–EDS

Foraminifera specimens were fixed with 2.5% glutaraldehyde in filtered ASW for at least 24 h at 4 °C. They were then embedded in 1% aqueous agarose and cut into cubes of roughly 1 mm3. Fixed specimens were embedded in 1% aqueous agarose, decalcified with 0.2% ethylene glycol-bis(2-aminoethylether)-N,N,N’,N’-tetraacetic acid in 0.81 mol l−1 aqueous sucrose solution (pH 7.0) for several days and then rinsed with filtered seawater. For measurement of P in cells with EDS, we did not conduct postfixation with osmium tetroxide, which was the overlapped energy peak position of P. Specimens embedded in agarose were rinsed with filtered ASW, stained with 2% uranyl acetate solution for 2 h at 4 °C, dehydrated in a graded ethanol series and embedded in epoxy resin (Quetol 651).

Ultrathin sections (100 nm) were cut using a diamond knife on an Ultracut S ultramicrotome and then stained with 2% aqueous uranyl acetate and lead staining solution (0.3% lead nitrate and 0.3% lead acetate, Sigma-Aldrich). TEM–EDS imaging was performed on a Tecnai G2 20 (Thermo Fisher Scientific), equipped with a bottom-mounted 2k × 2k Eagle charge-coupled device camera (Thermo Fisher Scientific) and a RTEM-S 61700ME EDS detector, (AMETEK) at an acceleration voltage of 200 kV. Note that the elements used for staining are heavier than Au and do not overlap with P.

Calculation of total phosphate storage in living foraminiferal assemblages

The total dissolved inorganic phosphate pool stored in foraminifera in the sediment column (∑DIPi sed., in mmol m−2) was calculated for locations in the Southern North Sea region around the Wadden Sea (from 2 to 10 °E and from 51 to 55 °N) and for the Peruvian continental margin (from 10 to 15 °S). Assemblage data for the North Sea region include 135 stations from the literature51,52,53,54,55,56,57 and one at Friedrichskoog (from this study). Assemblage data for Peru include 35 stations1,29.

The ∑DIPi sed. for 14 stations off Peru has previously been calculated, and is derived directly from ref. 1 For the remaining stations, ∑DIPi sed. was calculated according to equation (2) using the composition of benthic foraminiferal assemblages and intracellular phosphate content for each species (phosphatei n; Table 1):

$$\sum {{\rm{DIP}}}_{i\,{\rm{sed.}}}=\sum {A}_{n}\times {{\rm{phosphate}}}_{i\,n}\times {10}^{-9},$$
(2)

where An is the abundance (living) of foraminiferal species n (in ind m−2) and phosphatei n is the mean intracellular phosphate content of species n (in pmol ind−1). Individual phosphate storage data for A. veneta were not used for these calculations, because it was the only species for which no measurements were available from environmental samples (from laboratory cultures only). For species with unknown intracellular phosphate storage that share a genus with other species for which intracellular phosphate storage had already been determined, the average individual intracellular phosphate storage for this genus was used. Other species with unknown phosphate storage were excluded from the calculations. Finally, mmol m−2 was converted to g m−2 using the molar mass of phosphate (approximately 95 g mol−1). All assemblage data used for caclulations are summarized in Supplementary Table ST8 (Southern North Sea) and Supplementary Table ST9 (Peru) as downloadable spreadsheets.

Estimation of coastal riverine P runoff

Riverine phosphate runoff for the most important river estuaries of the Southern North Sea (Rhine, Meuse, Noordzeekanaal, Ijsselmeer, Ems, Weser and Elbe) was recorded for the year 2019 from the report of a monitoring programme58. Riverine total P runoff to the Peruvian coast from 10 to 15 °S, was taken from a global modelling study for the year 201559.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.