# Direct pathway of incorporating dietary nitrogen in shell-bound matrix of the planktic foraminifera Trilobatus sacculifer

Wei-Ning Fang, Oscar Branson, Er-Wen Yang, Wen-Hui Chen, Ren-Yi Cai-Li, Howard J Spero, Jennifer Fehrenbacher, Lael Vetter, Charlotte Lekieffre, Haojia Ren

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Wei-Ning Fang, Oscar Branson, Er-Wen Yang, Wen-Hui Chen, Ren-Yi Cai-Li, et al. Direct pathway of incorporating dietary nitrogen in shell-bound matrix of the planktic foraminifera Trilobatus sacculifer. Earth and Planetary Science Letters, 2025, 654, pp.119231. 10.1016/j.epsl.2025.119231. hal-05454657

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# Direct pathway of incorporating dietary nitrogen in shell-bound matrix of the planktic foraminifera *Trilobatus sacculifer*

Wei-Ning Fang<sup>a,b,*</sup> <sup>●</sup>, Oscar Branson<sup>a</sup>, Er-Wen Yang<sup>b</sup>, Wen-Hui Chen<sup>b,●</sup>, Ren-Yi Cai-Li<sup>b,●</sup>, Howard J. Spero<sup>c</sup>, Jennifer Fehrenbacher<sup>d,●</sup>, Lael Vetter<sup>e</sup>, Charlotte LeKieffre<sup>f</sup>, Haojia Ren<sup>b,*</sup> <sup>●</sup>

## ARTICLE INFO

Editor: Tristan J. Horner

**Keywords:**

Foraminifera-bound δ¹⁵N
Foraminifera culturing
Trophic effects
Primary organic sheet
Nitrogen metabolism
calcification

## ABSTRACT

The stable isotopes of organic nitrogen (N) preserved within the fossil tests of foraminifera have been used to reconstruct past changes in surface ocean nitrogen cycling processes. Modern observations show temporal and spatial covariations of δ¹⁵N between planktic foraminifera and particulate organic matter in the surface ocean, suggesting that heterotrophic foraminifera record the N isotopic compositions of their diet. However, little is known about the underlying mechanisms of N translocation from diet into foraminiferal biomass (i.e., intrashell protoplasm) and finally embedded within their mineralized shells. We investigate the pathways of N uptake from diet into foraminiferal calcite tests by feeding the planktic, dinoflagellate-bearing *Trilobatus sacculifer* with two strains of brine shrimp (*Artemia*) with different ¹⁵N/¹⁴N isotopic compositions, and monitoring the δ¹⁵N evolution of both biomass and shell-bound matrix. The two feeding groups show comparable results, that δ¹⁵N of both the biomass and shell-bound matrix evolve towards their diet sources without expressing trophic enrichment in δ¹⁵N. However, we observe that the δ¹⁵N of the biomass and shell-bound matrix exhibit distinct mixing behaviors. Biomass δ¹⁵N is well modelled as a mixture between the original biomass and newly metabolised dietary intake, suggesting *T. sacculifer* has a closed N system with minimum N leakage. Meanwhile, shell-bound δ¹⁵N quickly approaches the δ¹⁵N of the diet, and is offset from the biomass δ¹⁵N. This indicates a direct pathway of N incorporation from the diet into shell-bound organics during calcification, without significant exchange with the overall biomass pool.

## 1. Introduction

The isotopic composition of nitrogen (N) contained in proteins embedded in the calcite tests of planktic foraminifera, known as 'foraminifera-bound or FB-δ¹⁵N', provides an invaluable archive of marine paleo-nutrient conditions (e.g., Martínez-García et al., 2014; Ren et al., 2009, 2017; T. Wang et al., 2018) (δ¹⁵N = {[(¹⁵N/¹⁴N)sample/(¹⁵N/¹⁴N)N₂ in air] - 1}x1000 (‰ vs. air)). In oligotrophic surface water of low-latitude oceans, FB-δ¹⁵N is found to correspond to the δ¹⁵N of subsurface nitrate (NO₃), the predominant source of new N provided to the nitrate-limited euphotic zone through vertical mixing (Ren et al.,

2012; Smart et al., 2018). In these regions, FB-δ¹⁵N is used to track thermocline nitrate δ¹⁵N (δ¹⁵N-NO₃) changes, which is in turn influenced by N input and loss processes regulating the thermocline and global ocean NO₃ inventory (e.g., Ren et al., 2009, 2017; T. Wang et al., 2018). In nutrient-rich regions such as the Southern Ocean surface, FB-δ¹⁵N is observed to track changes in the surface δ¹⁵N-NO₃ (Smart et al., 2020). The surface δ¹⁵N-NO₃ rises as NO₃ concentration is lowered by biological uptake since phytoplankton preferentially incorporate lighter ¹⁴N (Granger et al., 2004; Wada and Hattori, 1978). In these regions, FB-δ¹⁵N is applied to reconstruct changes in NO₃ consumption in the euphotic zone, as determined by nutrient supply from depth and

<footnote><sup>a</sup> Department of Earth Sciences, University of Cambridge, Downing St CB2 3EQ, UK</footnote>

<footnote><sup>b</sup> Department of Geosciences, National Taiwan University, Taipei, Taiwan</footnote>

<footnote><sup>c</sup> Department of Earth and Planetary Sciences, University of California, Davis, CA 95616, USA</footnote>

<footnote><sup>d</sup> College of Earth, Ocean, and Atmospheric Sciences, Oregon State University, Corvallis, OR 97331, USA</footnote>

<footnote><sup>e</sup> Department of Geosciences, University of Arizona, Tucson, AZ 85721 USA</footnote>

<footnote><sup>f</sup> University Grenoble Alpes, CNRS, CEA, INRAe, IRIG-LPCV, Grenoble, France</footnote>

<footnote>* Corresponding authors.
E-mail addresses: wnf21@cam.ac.uk (W.-N. Fang), abbyren@ntu.edu.tw (H. Ren).</footnote>
biological uptake at the surface (Martínez-García et al., 2014; Ren et al., 2015; Straub et al., 2013).

Foraminifera-bound N serves as a more reliable proxy archive than bulk sedimentary N because the organics are tightly bound within the calcite crystal structure of the test (Branson et al., 2016). This binding prevents δ¹⁵N alterations by biological degradation (Altabet, 1988), diagenetic alteration (Altabet and Francois, 1994), and mixing with terrestrial N (Schubert and Calvert, 2001). As such, FB-δ¹⁵N has been increasingly used to reconstruct past changes in marine N sources, extending as far back as to the early Cenozoic (Kast et al., 2019; X. T. Wang et al., 2022). Despite the expanded use of the FB-δ¹⁵N proxy in palaeoceanographic studies, the pathways and processes of incorporating environmental information into the shell-bound matrix are still poorly understood. The exchange of N between foraminifera and the environment, the internal N cycling processes within foraminifera, as well as the formation of shell-bound organics all have the potential to influence the final δ¹⁵N recorded in foraminiferal shells (Fig. 1).

Foraminifera-bound δ¹⁵N is tightly coupled with environmental δ¹⁵N-NO₃⁻. However, planktic foraminifera are heterotrophic zooplankton that do not directly uptake inorganic NO₃⁻ from the water column. Although many planktic species have photosynthetic symbionts that are capable of assimilating NO₃⁻ from surroundings, this intake pathway is likely very limited given the low nutrient concentration in the mixed layer where most planktic foraminifera thrive. It has therefore been assumed that FB-δ¹⁵N inherits its isotopic signal from the foraminifera diet, and reflects food-web N from the lower trophic levels. The planktic foraminifera food source spans a wide range of suspended particulate organic N (SPN), including detritus, bacteria, phytoplankton, and other zooplankton (Anderson et al., 1979; Spindler et al., 1984). The

influence from diet is particularly evident in recent time-series studies that find seasonal and spatial covariances between FB-δ¹⁵N and δ¹⁵N-SPN in nutrient-depleted and nutrient-rich waters (Smart et al., 2018, 2020). Foraminifera may also assimilate ammonium (NH₄⁺) directly from their environment. The ability to assimilate NH₄⁺ has been demonstrated in several benthic (Bird et al., 2020; Jauffrais et al., 2019; Lekieffre et al., 2018) as well as planktic foraminifera (Bird et al., 2020; Lekieffre et al., 2020), regardless of whether they are kleptoplastic/symbiotic.

Endosymbionts may play an important role in internal N cycling process. Ammonium and perhaps other forms of organic N are produced during protein metabolism (Bidigare, 1983). The lighter ¹⁴N isotope is preferentially incorporated into the NH₄⁺ pool, the excretion of which to the environment leads to isotopic enrichment of the consumers' biomass. The expression of this isotopic elevation is referred to as trophic enrichment, and is typically between 2 to 4‰ for each trophic level (Checkley and Miller, 1989; Minagawa and Wada, 1984). Endosymbionts can reabsorb excreted NH₄⁺ (LeKieffre et al., 2020) and translocate the photosynthetic by-product back to its host (LeKieffre et al., 2018). This process may reduce bulk ¹⁵N-enrichment in foraminiferal δ¹⁵N relative to diet by reducing or even eliminating N loss to the environment (Uhle et al., 1997, 1999). In support of this hypothesis, recent studies show that foraminifera species hosting dinoflagellate symbionts typically have lower δ¹⁵N in both biomass (i.e., foraminifera's protoplasm) and shell-bound matrix relative to species without algal symbionts or hosting different symbionts collected from the same environment (Cai-Li et al., 2024, submitted; Ren et al., 2012; Smart et al., 2018)

The process of N translocation from source to biomass and to organic matrix, along with the associated isotopic fractionation, also remains

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Nitrogen (N) pathways in planktic foraminifer-symbiont system (adapted from Uhle et al., 1999), subdivided into the internal (includes foraminiferal shell and spine microenvironment, shown as the lighter oval area) and the external (beyond the spine halo, shown as the darker peripheral area) cycling regions. A foraminifer stores protein-N primarily inside the chamber space although a small amount of protein (< 0.5 %) is incorporated into the calcite chamber walls. The shell-bound N is found in the Primary Organic Sheet (POS) and subsequent Organic Layers (OL), which are composed of biomineralizing template molecules that are laid down before chamber calcification events. In this study we focus on dietary N incorporation into the dinoflagellate-bearing *T. sacculifer*. The pathways being investigated via our culture experiments are marked in solid lines (on the right side of the figure), while the others in dash lines.</figcaption></figure>
unclear. Within the foraminifera, most of the N is in its protoplasm biomass. Only a very small fraction of N is 'trapped' within the calcite test. This shell-bound N is thought to be primarily in proteins and polypeptides (Robbins and Brew, 1990; Robbins and Donachy, 1991; Stathoplos and Tuross, 1994) within the Primary Organic Sheet (POS, also referred to as the Primary Organic Membrane, POM in early studies; see clarification in De Nooijer et al., 2014; Erez, 2003), which is laid down as a template structure at the onset of mineralization (Bé and Hemleben, 1970; Spero, 1988). Additional calcite and organic N are added as the shell wall thickens, possibly associated with secondary organic sheets (Erez, 2003; Reiss, 1957). As calcite is deposited, the organics are 'locked' within the test, recording the transient δ¹⁵N of the shell-bound matrix at the time of test formation. This is distinct from biomass δ¹⁵N, which may evolve throughout the life of the foraminifera with additional dietary intake. The shell-bound and biomass N may thus have different integration time during the growth of a single foraminifer. Modern observations of planktic species indicate an overall strong δ¹⁵N similarity between biomass and shell-bound N when comparing the average values across time and multiple species, but the δ¹⁵N relationship seems to vary among species, with relatively large variability among individual specimens (Ren et al., 2012; Smart et al., 2018, 2020). This variability may result from the differences in the processes of translocating biomass N into the calcite shell, and/or the different residence times of the two N pools (biomass vs. shell-bound N) within a foraminifer.

To confidently interpret FB-δ¹⁵N as a proxy for marine N cycling, we must understand the mechanisms of N uptake into the shell, and potential biases that might arise from them. Previous attempts focus on calibration from field observations; we present geochemical data from

culture experiments to identify the pathway(s) of N uptake from diet to shell-bound matrix. To achieve this, we track the evolution of δ¹⁵N in foraminiferal biomass and shell-bound N in response to controlled diet with a known and distinct isotopic composition. We selected the planktic, spinose, dinoflagellate-bearing *Trilobatus sacculifer* as our model species. It is the most commonly measured species in FB-δ¹⁵N applications, and is shown to have similar δ¹⁵N values to ambient environmental NO₃⁻ sources (Ren et al., 2012; Smart et al., 2018). These culture experiments allow us to quantify trophic effects in *T. sacculifer* during dietary uptake, trace the N incorporation from diet source into foraminiferal biomass and ultimately their calcifying matrix, and constrain the metabolism rate of N within a foraminiferal cell. Understanding these processes will address significant knowledge gaps in our mechanistic understanding of N incorporation, improving our ability to reconstruct nutrient dynamics in ancient oceans from FB-δ¹⁵N.

## 2. Materials and methods

### 2.1. Location

Sample collection and foraminifera culturing took place at the Green Island Marine Research Station, Taiwan (22°40′N, 121°29′E). Green Island is situated along the southeastern coast of Taiwan in the mainstream of the Kuroshio current (Fig. 2). The current transports warm (surface temperature ≈ 28 °C) oligotrophic (surface [NO₃⁻] ≈ 0 µM) seawater that originates from the North Equatorial Current. Tropical zooplankton communities are prevalent, with planktic foraminifera species such as *Globigerinoides ruber*, *Trilobatus sacculifer*, *Globigerinella siphonifera*, and *Globigerinella calida*, among which *G. ruber* is the most

<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Green Island in the oligotrophic western Pacific Ocean is situated in the northward flowing Kuroshio current. Our sampling site (red dot) is on the north side of the island where water depth is > 200 m. After collection, foraminifera were transported back and cultured at the Green Island Marine Research Station (GIMRS, white triangle). The mean annual surface NO₃⁻ concentration is from *World Ocean Atlas* 2018 (Garcia et al., 2019).</figcaption></figure>
abundant (Taylor et al., 2018). Sea surface temperatures (SST) vary seasonally from 26 °C in winter to 29 °C in summer. The mixed layer depth (MLD) ranges from 40 to 65 m in winter to 15 m in summer, averaging 25 to 30 m annually (CTD data from Ocean Data Bank, National Science and Technology Council, https://www.odb.ntu.edu.tw/). During our sampling periods, the SST was observed to be approximately 27 to 28 °C, and the MLD ranged from 30 to 50 m.

## 2.2. Sample collection

In May 2019 and September 2023, individual spinose planktic foraminifera, plankton tow materials, and seawater were collected north of Green Island in open water (depth > 200 m) (Fig. 2 & 3). SCUBA divers hand collected live *T. sacculifer* specimens for lab culture from 1 to 5 m depths following standard protocols (Huber et al., 1996). Bulk plankton material was obtained using an open plankton net (202 µm mesh) from surface to 25 m depths. For culture experiments, large volumes of surface seawater were collected with a 20 L acid-cleaned Nalgene bottle, filtered (0.2 to 0.22 µm, hydrophilic polyethersulfone or polycarbonate membrane filter), and used within two days. Unprocessed samples were frozen at -20 °C for later N composition and δ¹⁵N analyses.

## 2.3. Culture experiments

Upon return to the research station, foraminifera were identified at the species level and measured for size under an inverted microscope, then transferred individually into clean 120 ml Wheaton jars containing filtered seawater. Individuals were observed daily with a hand lens, and their ontogenesis (chamber formation events) recorded. Temperature was maintained at ~25 °C (±5 °C) by air-conditioners in 2019, and at 27 °C (±0.5 °C) by recirculating water baths in 2023. Aquarium lights (VIPARSPECTRA, 165 W LED) provided high-intensity illumination (400–1000 µmol photons/m²/s, PPFD) to maximize symbiont photosynthesis (Jørgensen et al., 1985; Spero and Parker, 1985), and on a 12 h:12 h light/dark cycle to mimic the day/night cycle.

In 2019, individual *T. sacculifer* were randomly separated into two

groups, each fed with a different strain of commercially available *Arte mia* nauplii with distinctive biomass δ¹⁵N: the Great Salt Lake strain from Ocean Star International (GSL; biomass δ¹⁵N = 10.5 ± 0.3‰ 1 SD; N content per nauplii = 7.0 ± 1.6 nmol N 1 SD; number of measure ments (n) = 4) and the Tibetan strain from Aquamaster (Tibetan; biomass δ¹⁵N = 4.8 ± 0.1‰ 1 SD; N content per nauplii = 12.7 ± 2.1 nmol N 1 SD; n = 7) (Spero et al., 1993; details in Section 2.4.2 and 3.2.1.) (Fig. 3). The Tibetan strain of *Arte mia* nauplii has roughly double the biomass N content of the GSL strain. Thus, the GSL group was fed one nauplius daily, while the Tibetan group was fed one every other day, ensuring comparable N ingestion rates under identical conditions. Both groups exhibited similar growth patterns (Fig. 4 & Table S1). Culture ended when the foraminifera either underwent gametogenesis or died before gamete release. About 2 % of specimens were terminated while still alive at the field season's end. Pre-gametogenesis specimens retained most of their protoplasm and had about twice the N content of post-gametogenesis specimens (Section 3.2.3). We did not differentiate individuals by end status but assessed potential isotopic changes due to biomass N loss from gametogenesis (see below). Cultured shells and remaining protoplasm were dried down on 10-hole microfossil slides and preserved in desiccator prior to analyses.

Individual chambers of cultured *T. sacculifer* in 2019 were amputated from the shell using a scalpel under a dissecting microscope (Cang and Shackleton, 1990; Spero and Lea, 1993). Chamber number in the shell whorl was determined according to their size and relative position, and classified into three types: 'wild' grown chambers already present when the specimen was collected in the field (Ch0); the first new chamber after feeding experiment started (Ch1), and subsequent chambers formed during the feeding experiment (Ch2, which included the second, third and occasionally the fourth chamber that formed in culture). The three chamber types were pooled separately in 12 ml pre-combusted glass vials with a clean brush (Table S2), with each type containing pooled samples from 79 to 153 chambers. The remaining protoplasm and shell-bound matrix sequestered within the calcite were analyzed separately for N content and δ¹⁵N following protocols in Section 2.4.1.

To constrain the evolution of biomass δ¹⁵N under controlled diet and the biomass changes associated with gametogenesis, additional

<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Experiment procedures. Samples were divided into two categories: materials collected in field and foraminifera undergoing culture. Unless specified otherwise, samples were collected in May 2019. In the culture, *T. sacculifer* were fed with two strains of *Arte mia* (GSL: the Great Salt Lake strain and *Tibetan*: the Tibetan strain, details in text) with different biomass δ¹⁵N values, referred to as diet δ¹⁵N in the figure. The sample types that were analyzed for δ¹⁵N are indicated in grey rectangles.</figcaption></figure>
<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Distribution of the cultured *T. sacculifer* in 2019, plotted against their (a) initial size, (b) survival time in culture, (c) number of new chambers formed and (d) end status. No significant difference is observed between the two feeding groups. For the GSL group, each foraminifer is fed one *Artemia* every day. For the Tibetan group, each foraminifer is fed one *Artemia* every other day.</figcaption></figure>

*T. sacculifer* specimens cultured in 2023 were collected for biomass analyses (Fig. 3). The specimens were fed with one GSL *Artemia* nauplii per day in the morning. After 2, 4, 6, and 8 days, two specimens were killed instantly at -80 °C to preserve the fresh biomass. This was conducted 6 to 24 h after feeding, when no undigested *Artemia* was visible in the foraminiferal spines. After day eight, feeding was stopped to allow natural gametogenesis. By comparing biomass values prior and after gametogenesis, we could constrain the loss of biomass and its δ¹⁵N changes associated with gametogenesis.

## 2.4. Analytical methods

### 2.4.1. Foraminifera specimens from tow materials and culture experiments

For tow-collected/cultured specimens, we measured the δ¹⁵N of bulk/remnant protoplasm and organic N within calcite chamber walls with the persulfate-denitrifier method (Knapp et al., 2005; Ren et al., 2009). This involved persulfate oxidation of the organic N into NO₃⁻ before and after acid digestion of the calcite shell, followed by denitrifier method for δ¹⁵N analyses. We used 5 or 6 ml of persulfate oxidation reagent (POR; 1 g recrystallized K₂S₂O₈ + 1 g NaOH in 100 ml Milli-Q water) to oxidize protoplasm prior to acid digestion. We then transferred 4 ml of the supernatant containing NO₃⁻ converted from protoplasm N to new pre-combusted glass vials. The remaining calcite free of protoplasm were rinsed with Milli-Q water 3 times, dried at 50 °C, and weighed (0.5 to 5 mg) into pre-combusted 4 ml vials, where they were dissolved in 2–3 N HCl to release the shell-bound organic N. The shell-bound N is subsequently oxidized with 1 ml POR (0.3 g recrystallized K₂S₂O₈ + 0.5 to 0.7 g NaOH in 100 ml Milli-Q water). International

standard USGS 40 (δ¹⁵N = -4.5‰ vs. air) and a laboratory standard made of a mixture of 6-aminocaproic acid and glycine (A + G5, δ¹⁵N = 5.4‰ vs. air) were oxidized alongside the shell samples and POR blanks. All samples were pH adjusted to between 4 and 7, and analyzed for concentration with the chemiluminescence method (Braman and Hendrix, 1989) and δ¹⁵N with denitrifer method (Knapp et al., 2005; Ren et al., 2009). The POR recipe differs for each type of analyses to ensure complete oxidation and to minimize blank contribution.

Given the low N content in shell-bound matrix, the total N for chamber-specific shell-bound δ¹⁵N even after pooling 79 to 153 individual chambers is small, only allowing single measurement on each chamber type. We take the following steps to ensure the precision of the measurements and constrain uncertainties. After accounting for material loss during cleaning, the total sample ranges from 0.58 to 3.93 mg of calcite for each chamber type. This then yields a total N content from 2.7 to 25.3 nmol. Considering our blank size of 0.24 nmol of N, the blank-to-total N content for all the measurements are below 10% (ranging from 1 to 9%). The N content for the shell-bound material ranges from 4.0 to 7.5 nmol N/mg of cleaned calcite, which is comparable to reported N content in fossil foraminifera (Ren et al., 2017), and well within the range of observed N content in fresh foraminifera collected in plankton tows (Smart et al., 2018).

The two organic standards are used to constrain the uncertainties. Both standards are treated the same way as the samples, with comparable range of blank-to-total N ratio (1 to 8%). We apply the same blank correction method using the isotope mass balance equation for all references and samples. After blank correction, the average δ¹⁵N value of these reference material is 0.01‰ lower than their nominal δ¹⁵N value,
with 1 SD of ±0.18‰. We thus report an analytical error of ±0.2‰ for the chamber-specific FB-δ¹⁵N. The raw data, references, blanks, and blank correction are all provided as a supplementary file for further evaluation.

### 2.4.2. *Artemia*

Foraminifera ingest only the soft biomass of *Artemia*, discarding the outer chitin carapaces (Uhle et al., 1997). To separate the two, 50 individuals' tissues were dissolved in 1 ml of 10 % NaOH at 70 °C for 30 min (Perga, 2010), then centrifuged to settle the carapaces. The upper 50 µl liquid containing dissolved biomass was pipetted to 4-ml pre-combusted glass vials. The carapaces were rinsed with Milli-Q water at least 3 times before transfer to pre-combusted glass vials, dried at 50 °C, and treated with 40 µl of 3 N HCl to dissolve any carbonate (Perga, 2010). The *Artemia* biomass liquid and chitin precipitates were oxidized separately with 2 ml of POR (0.3 g recrystallized K₂S₂O₈ + 0.4 g NaOH in 100 ml Milli-Q water), followed by pH adjustment and analyses of concentration and isotope as with the foraminifera samples.

### 2.4.3. *Mass spectrometry methods*

The N₂O converted by denitrifiers was measured by gas chromatography connected to an isotope ratio mass spectrometry (IRMS; Thermo 253 PLUS, connected to Thermo Conflo IV open split interface) (Sigman et al., 2001; Weigand et al., 2016) in the Department of Geosciences, National Taiwan University. Bacterial blanks and two NO₃ isotopic standards, IAEA-N3 (δ¹⁵N = 4.7‰ vs. air) and USGS34 (δ¹⁵N = -1.8‰ vs. air), were run with the samples to monitor the bacterial conversion and the stability of the mass spectrometer. The precision for denitrifier method is ±0.08‰ (1 SD) at 2 nmol of N constrained by the two nitrate standards, and the precision generally improves at higher N level (Fig. S1). The δ¹⁵N of oxidized samples were corrected for POR blanks by isotope balance equations. The δ¹⁵N precision for the persulfate-denitrifier method is within ±0.2‰ (1 SD), based on analytical errors of the organic standards processed along with the samples. We report 1 SD for all the samples for which we can make duplicated measurements. For chamber-specific FB-δ¹⁵N, we provided the details to derive the uncertainty above. The raw data, references, blanks, and blank correction for all the δ¹⁵N measurements in this study are provided as a supplementary file for further evaluation.

## 3. Results

### 3.1. *Field samples*

Along with *T. sacculifer*, two other spinose planktic species, *Globigerinoides ruber* and *Globigerinella siphonifera*, were analyzed from 2019 tow collections. Both biomass δ¹⁵N and FB-δ¹⁵N were measured on *T. sacculifer* and *G. ruber*, while only biomass δ¹⁵N was measured for *G. siphonifera* due to insufficient samples. The average biomass δ¹⁵N values were 1.6‰ (±0.2‰ 1 SD; *n* = 13) for *G. ruber*, 2.1‰ (±0.6‰ 1 SD; *n* = 23) for *T. sacculifer*, and 3.0‰ (±0.4‰ 1 SD; *n* = 5) for *G. siphonifera*. The FB-δ¹⁵N values were 1.4‰ (±0.1‰ 1 SD; *n* = 2) and 2.2‰ (±0.7‰ 1 SD; *n* = 3) for *G. ruber* and *T. sacculifer*, respectively. The similar δ¹⁵N values between biomass and shell-bound N are consistent with previous findings in the Atlantic Ocean (Smart et al., 2018). The species difference also aligns with surface sediment and plankton net studies (Ren et al., 2012; Smart et al., 2018). The biomass δ¹⁵N of tow-collected *T. sacculifer* from 2023 matched 2019 values at 2.2‰ (±0.4‰ 1 SD; *n* = 9) within uncertainty. Biomass N content per *T. sacculifer* was 9.6 nmol N (±4.3 nmol 1 SD; *n* = 23) in 2019, and 7.3 nmol N (±2.3 nmol 1 SD; *n* = 9) in 2023. These measurements serve as control points for cultured specimens in both years.

### 3.2. *T. sacculifer* feeding experiments

#### 3.2.1. *Artemia*

The GSL and Tibetan strains of *Artemia* have biomass δ¹⁵N values of 10.5‰ (±0.3‰ 1 SD; *n* = 4) and 4.8‰ (±0.1‰ 1 SD; *n* = 7), respectively. The δ¹⁵N values of their chitinous carapace were at 10.4‰ (±0.5‰ 1 SD; *n* = 3) and 4.6‰ (±0.7‰ 1 SD; *n* = 8), similar to their respective biomass. For N content, the GSL strain has 7.0 nmol N (±1.6 nmol N 1 SD; *n* = 4) in biomass and 0.7 nmol N (±0.1 nmol N 1 SD; *n* = 3) in carapace per nauplius, while the Tibetan strain has 12.7 nmol N (±2.1 nmol N 1 SD; *n* = 7) in biomass and 0.9 nmol N (±0.5 nmol N 1 SD; *n* = 8) in carapace per nauplius. The Tibetan strain biomass N content is approximately twice that of the GSL strain. No significant fractionation between biomass and carapace δ¹⁵N was observed, contrary to Uhle et al. (1999), who reported a 7.3‰ enrichment in biomass δ¹⁵N. Since *Artemia* sheds its exoskeleton as it grows, it is possible that the periodical molting leads to carapace differentiation (Criel and Macrae, 2002) which may cause biomass-chitin δ¹⁵N fractionation through aging. However, since the nauplii measured here and used in our feeding experiments were only 1-day old, it may be too short to generate a substantial biomass-chitin fractionation. As the carapace is discarded by foraminifera after feeding (Uhle et al., 1997), the *Artemia* δ¹⁵N hereafter only refers to its biomass δ¹⁵N.

#### 3.2.2. *Biomass N evolution during feeding experiments*

Biomass includes the total organic material from the foraminifera host and its intracellular symbionts. Assuming that an average *T. sacculifer* specimen has on the order of 1000 dinoflagellate symbionts (Hönisch et al., 2021; Spero and Parker, 1985) with 10 to 50 picogram N per cell as reported in related species (Menden-Deuer and Lessard, 2000; Nielsen, 1996), symbionts would account for 0.7 to 3.6 nmol N in the bulk biomass. This is a non-negligible fraction of the total biomass. This study analyzes the total biomass without differentiating these pools, though potential isotopic differences between them warrant further investigation.

During the 2023 field season, each *T. sacculifer* was fed one GLS *Artemia* nauplii daily, with two specimens archived every other day to track biomass δ¹⁵N and N content evolution. The tow-collected *T. sacculifer* from the same season are considered our baseline values (δ¹⁵N = 2.2 ± 0.4‰ 1 SD; N content = 7.3 ± 2.3 nmol N/indiv. 1 SD; *n* = 9). The cultured *T. sacculifer*'s biomass δ¹⁵N increased from the baseline to 8.3‰ (*n* = 1), 9.6‰ (±0.4‰ 1 SD; *n* = 2), 10.0‰ (±0.1‰ 1 SD; *n* = 2), and finally 10.5‰ (±0.1‰ 1 SD; *n* = 2) after consuming 2, 4, 6, and 8 *Artemia* nauplii over the 8-day culture (Fig. 5 & Table S3). Foraminiferal biomass steadily increases by 8.2 nmol N (±0.6 nmol N 1 SD) per *Artemia* consumed. No significant increase in biomass N content was observed after consuming two *Artemia* compared to tow-collected *T. sacculifer*, likely due to variability in N content and size between tow-collected and cultured specimens.

#### 3.2.3. *Biomass N changes associated with gametogenesis*

We also investigate biomass changes associated with gametogenesis by comparing specimens before and after gamete release, after feeding the same number of *Artemia* in 2023. Before gametogenesis (*n* = 2), specimens had a biomass N content of 55.7 nmol N (±3.7 nmol N 1 SD) and a δ¹⁵N value of 10.5‰ (±0.1‰ 1 SD). After gamete release (*n* = 1), which corresponds to a partial emptying of protoplasm from the foraminifera shell, the biomass N content decreases to 33.7 nmol N. However, the δ¹⁵N value remained the same (10.5‰) (Fig. 5 & Table S3). These results suggest that *T. sacculifer* loses ~40% of its biomass during gametogenesis without altering the δ¹⁵N of residual protoplasm.

#### 3.2.4. *Shell-bound N evolution during feeding experiments*

In May 2019, in total 298 *T. sacculifer* individuals were collected and successfully cultured. The ontogenetic history of the two feeding groups is summarized in Fig. 4 and Table S1. On average, individuals lived ~6
<figure><img src="image_8.png" /><figcaption>Fig. 5. Bulk biomass N evolution of *T. sacculifer* feeding on the GSL strain *Artemia* in 2023. As foraminifera grow and digest their *Artemia* prey, the biomass N content increases and its δ¹⁵N approaches the diet δ¹⁵N. Gametogenic ('Gam.', red cross) individuals lose ~40 % of their biomass without altering the δ¹⁵N of remaining protoplasm. No increase in biomass N content is observed when comparing foraminifera recently fed with two *Artemias* to their tow-collected baseline (grey dot). The two stages are thus connected in dashed lines as their initial N content might not be comparable, probably due to a smaller averaged size of the foraminifera selected into the experiment. Data are provided in Table S3.</figcaption></figure>

days and developed a new chamber every ~2.5 days. No systematic differences were observed between feeding groups, indicating similar physiological conditions for both.

The δ¹⁵N of cultured *T. sacculifer* diverges significantly from that of tow-collected specimens, showing elevated δ¹⁵N in both shell-bound N and remnant protoplasm. This reflects a N-mixture of the baseline δ¹⁵N from tow-collected *T. sacculifer* and δ¹⁵N from their *Artemia* diet (Fig. 6). For both feeding groups, FB-δ¹⁵N in chambers from growth stages 'Ch0-Ch1-Ch2' increased towards their diet δ¹⁵N: '7.3-7.9-10.2‰' for the GSL and '4.1-4.9-5.1‰' for the Tibetan group, indicating incorporation of dietary N into shell-bound N. Each chamber type's FB-δ¹⁵N represents a single measurement. For the Tibetan group, FB-δ¹⁵N of Ch2 is slightly more elevated than the dietary δ¹⁵N with marginal significance (p-value = 0.09, two-sample t-test performed using the ttest_ind_from_stats function from the sciPy.stats module, SciPy v1.14.1). In contrast, the δ¹⁵N of the remnant protoplasm remained constant across stages, at 8.5‰ (±0.1‰ 1 SD; n = 3) for the GSL and 4.5‰ (±0.2‰ 1 SD; n = 3) for the Tibetan group.

# 4. Discussion

## 4.1. *T. sacculifer* biomass expresses no trophic effect

Trophic enrichment of δ¹⁵N is ubiquitous across marine food web. Isotopic fractionation occurs during the production of NH₄⁺ through deamination of proteins and amino acids, when lighter ¹⁴N is preferentially incorporated into NH₄⁺ (Bada et al., 1989; Macko et al., 1986). Loss of this isotopically depleted NH₄⁺ will then increase the δ¹⁵N of a consumer's biomass. The trophic effect is thus defined as the N isotopic difference between the consumer and its diet, typically observed to be +2 to 4‰ for each trophic level (Checkley and Miller, 1989; Minagawa and Wada, 1984). However, during the biomass tracing experiments in 2023, *T. sacculifer* expressed negligible trophic effect: the foraminiferal biomass δ¹⁵N evolves towards, but never exceeds the diet δ¹⁵N (Fig. 5& Table S3).

It is possible to lower trophic enrichment if the primary N waste is dissolved organic nitrogen (DON) instead of NH₄⁺. For example, it has been reported that changes in food quality could have an effect on the trophic enrichment (Miller and Roman, 2008). However, this is unlikely to eliminate any fractionation during the metabolism. In addition, we report a constant growth rate (8.2 ± 0.6 nmol N/Artemia consumed 1 SD) in the *T. sacculifer* biomass N content comparable to the N content of each *Artemia* (7.0 ± 1.6 nmol N/indiv. 1 SD; n = 4). This would suggest that *T. sacculifer* is a closed system with no N leakage. In support of this, we found that both the NH₄⁺ and DON concentrations in the culture vessels remained relatively constant during the culture experiments in 2019 (Fig. S2), detailed in a separate manuscript (Cai-Li et al., 2024, submitted).

These observations require an efficient internal N recycling mechanism to minimize N loss. *Trilobatus sacculifer* is one of the common planktic foraminifera species with dinoflagellate symbionts, dominating the tropical and subtropical regions, where density stratification of the water column restricts vertical mixing of deep nutrients into the photic zone. In such oligotrophic environments, direct uptake of nutrient N from the environment by these dinoflagellate symbionts is likely very limited given the low concentrations of major nutrients in the surface ocean. It is more likely that these algae utilize dissolved inorganic N derived from foraminiferal host excretion, thereby creating an efficient nutrient recycling system (LeKieffre et al., 2020) (Fig. 1). This recycling system is crucial for their success in nutrient-limited environments and explains their lower FB-δ¹⁵N relative to other surface-dwelling spinose species with non-dinoflagellate symbiont (e.g. *Globigerinella siphonifera*) or symbiont-barren species (e.g. *Hastigerina pelagica* or *Globigerina bulloides*) (Ren et al., 2012; Smart et al., 2018).

In summary, the biomass tracing experiment demonstrates that *T. sacculifer* expresses a negligible trophic effect during our culture experiments and can be treated as a closed system with little to no N waste release to the environment. With single dietary input, the biomass δ¹⁵N at any given stage can be calculated by the following mass balance model:

$$ \text{Biomass } \delta^{15}\text{N} = \text{Biomass } \delta^{15}\text{N}_{\text{Baseline}} * (1 - f_{\text{Diet}}) + \delta^{15}\text{N}_{\text{Diet}} * f_{\text{Diet}} \quad (1) $$

where the biomass δ¹⁵N is calculated from the baseline values measured from the tow-collected *T. sacculifer* and *Artemia* diet of either strain, depending on their respective δ¹⁵N and relative contributions of each source (f) to the total N content.

To further validate this idea, we revisited the remnant protoplasm δ¹⁵N measured in 2019. Since our biomass tracing experiment showed negligible isotopic changes associated with gametogenesis despite partial biomass loss (Section 3.2.3), we can assume the remnant biomass δ¹⁵N represent their final biomass δ¹⁵N regardless of the specimens' end status. Both feeding groups showed homogeneous δ¹⁵N signal: 8.5‰ (±0.1‰ 1 SD; n = 3) for the GSL and 4.5‰ (±0.2‰ 1 SD; n = 3) for the Tibetan group, irrespective of chamber stage (Ch0, Ch1, or Ch2) (Fig. 6).
<figure><img src="image_9.png" /><figcaption>**Fig. 6.** Foraminiferal $\delta^{15}$N of cultured *T. sacculifer* and their tow-collected baseline (bars showing $\pm$1 SD) in 2019. Left: shell-bound N, displayed along with the chamber types in growth order on the x-axis (Ch0: the wild chambers, Ch1: the first chamber that grows under culture, Ch2: the second and the following chambers that grows under culture). Bars showing analytical errors of organic standards instead of repeated measurements due to sample scarcity. Right: biomass, here referring to either the final remnant protoplasm after dead/gametogenesis for the cultured specimens, or the bulk biomass for the tow-collected ones as collected alive (see text). Diet $\delta^{15}$N (*Artemia* biomass only, shaded areas showing $\pm$1 SD) shown as dashed lines in both panels. Signals of shell-bound N and remnant protoplasm are both elevated towards the diet sources under culture.</figcaption></figure>

This consistency is expected since the remnant biomass $\delta^{15}$N reflects the cell's final composition, not at the time of chamber formation (unlike shell-bound N, Section 4.2). Using Eq. (1), we can also calculate a theoretical final biomass $\delta^{15}$N from the baseline biomass values in 2019 (tow-collected *T. sacculifer*: $\delta^{15}$N = 2.1 ± 0.6‰ 1 SD; N content = 9.6 ± 4.3 nmol N/indiv. 1 SD; n = 23) and gross intake of the designated diet. The result with propagated errors is 9.0‰ (±0.7‰ 1 SD) for the GSL and 4.3‰ (±0.2‰ 1 SD) for the Tibetan group, which is within ±5 percent (%) difference from the measurements. The error bars for the modelled biomass $\delta^{15}$N derived from uncertainties in both the $\delta^{15}$N and N content of tow-collected *T. sacculifer* biomass and *Artemia* diet. Calculations were performed individually for each specimen and then averaged. These results give us confidence that the evolution of biomass $\delta^{15}$N can be back-calculated according to the amount of diet digested at each stage of chamber production (Table S4). Using this closed-system model, we reconstructed the biomass $\delta^{15}$N values as each specimen developed a chamber during culture experiment in 2019. The values are then averaged across specimens and compared to the observed evolution of FB-$\delta^{15}$N changes (Fig. 7).

## 4.2. From dietary N to shell-bound N

In the 2019 experiment, we observe that FB-$\delta^{15}$N values approached the diet $\delta^{15}$N as foraminifera grew by adding chambers. The shell-bound N captures the $\delta^{15}$N signal of organic matter incorporated during chamber formation, and is thus able to record the changes through each development stage. This matches the expected patterns from foraminiferal biomass $\delta^{15}$N changes as new N is added during the experiment.

However, the FB-$\delta^{15}$N is consistently higher than the modelled biomass $\delta^{15}$N at any given ontogenetic stage (Fig. 7a&b).

The FB-$\delta^{15}$N of Ch0 (existing chambers upon collection) is elevated compared to the baseline value of FB-$\delta^{15}$N measured from tow-collected specimens. We attribute this offset to thickening of chamber walls associated with subsequent chamber additions. While shell-bound organic material is considered mainly in the POS within new chambers, additional Organic Layers (OL) may be added with secondary calcite in lamellar perforate foraminifera (Bé and Hemleben, 1970; Sadekov et al., 2005), or fragmentarily throughout the test structure (Branson et al., 2016) (Fig. 1). Consequently, Ch0 $\delta^{15}$N is likely determined by both the original chamber N and the new N addition in the feeding experiment when subsequent chambers formed. The $\delta^{15}$N of Ch0 from both the GSL and the Tibetan groups suggests that secondary calcite with additional OL can significantly contribute N (>60 %) to the entire chamber (Fig. 7c&d). By the same token, all the $\delta^{15}$N of chambers we analyzed is expected to express mixed signals contributed by various calcite layers formed successively. Nevertheless, the thickening effect should be less and less significant in the younger Ch1 and Ch2, but the $\delta^{15}$N difference between shell-bound matrix and biomass persists, thus requiring other explanations. This isotopic difference contrasts with field observations, which show an overall similarity between the two, indicating that it cannot be explained by isotopic fractionation during N incorporation into the shell.

Alternatively, the offset between shell-bound and biomass $\delta^{15}$N can result from different N uptake pathways into these two N pools. Specifically, newly metabolized dietary N is directly incorporated into the shell-bound matrix without passing through the foraminiferal bulk
<figure><img src="image_10.png" /><figcaption>**Fig. 7.** Evolution of measured FB-δ¹⁵N (see Fig. 6) versus modelled biomass δ¹⁵N through chamber development. (a) & (b) The modelled biomass δ¹⁵N is calculated based on a closed N model with a sole input from diet intake (Eq. (1)), assuming an initial biomass δ¹⁵N same as the tow-collected specimens. End-stage biomass δ¹⁵N of the two feeding groups (see Fig. 6) are also plotted in the panels to compared with their theoretical results based on an averaged known total dietary intake (see text). The modelled final biomass δ¹⁵N can be slightly lower than that at the 3rd chamber formation. This is because the end-stage is an average of all specimens, but not all specimens formed three chambers before being archived. Notice the modelled biomass δ¹⁵N falls behind the FB-δ¹⁵N at any given stage. Calculated from these isotopic values, the (c) & (d) shell-bound N and (e) & (f) biomass N show the relative contributions between baseline N and dietary N through time (Eq. (1)). Data are provided in Table S4. For measured shell-bound δ¹⁵N in (a) and (b), the error bar is derived from organic standards (1 SD), same as in Fig. 6. The error bar for tow-collected specimens is the scaled 1 SD of individual tow measurements. The error bars for modeled biomass δ¹⁵N accounts for the variabilities in the δ¹⁵N and N content in tow-collected foraminifera, *Artemia* diet, and the number of total *Artemia* uptake for each specimen.</figcaption></figure>

biomass (Fig. 8). In this model, the biomass N is a mixture of existing biomass and newly ingested dietary N, with δ¹⁵N reflecting the relative contribution of each endmember (Eq. (1)). Shell-bound N is the instantaneous product of the newly incorporated dietary N. The rapid development and the persistency of the isotopic difference between biomass and shell-bound N in culture experiments is thus explained by a switch in the diet δ¹⁵N.

It has been proposed that organelles called fibrillar bodies in planktic foraminifera are responsible for synthesizing proteins/polysaccharides and potentially serving as N reservoirs for biomineralization (LeKieffre et al., 2020; Spero, 1988). During calcification, the stored biomineralization proteins in these fibrillar bodies are released and transported to

the site of calcification, supplying protein-N to the shell-bound matrix. Building on this hypothesis, we propose that the fibrillar bodies accumulate N directly from the diet source, before the dietary N mixes with the rest of the biomass (Fig. 8). A key feature of this model is that the fibrillar bodies are not homogenous, but accumulate N continuously, inheriting the isotopic composition of the main new N source at the time they assemble the matrix proteins (i.e., dietary N, in our experiments). This allows each fibrillar organelle to have a distinct isotopic composition. Evidence from LeKieffre et al. (2020) shows that after exposure to spiked ¹⁵NH₄⁺, most but not all fibrillar bodies are enriched in ¹⁵N. Specifically, they observe two fibrillar bodies in close proximity with very different isotopic compositions. One is ¹⁵N-enriched while the
<figure><img src="image_11.png" /></figure>

**Fig. 8.** The model of dietary N incorporation into a foraminifer. After digested in vacuoles, the dietary N is released to the association probably in form of $NH_4^+$. It then blends into original biomass without leakage and form a ‘mixed’ biomass at any given time. Meanwhile, a parallel path exists from dietary N into shell-bound N. Organelle fibrillar bodies (‘f’; f1 represents the original fibrillar bodies, and f2 represents the dietary N enriched fibrillar bodies) are proposed as reservoirs of shell-bound matrix (LeKieffre et al., 2020; Spero, 1988). The fibrillar bodies directly receive metabolised dietary N from vacuoles and are sequentially formed in a foraminiferal cell (see evidence in Fig. 9). As a result, the bodies are not necessarily homogenous and can bear distinct isotopic compositions at the time they are produced. In our culture experiments, a mixture of fibrillar proteins from before (in white, containing baseline N possibly derived from diet acquired in wild) and after (in black, containing metabolised dietary N in the lab) the culture is embedded into shell-bound N. However, the baseline N signal should be progressively diluted in newly formed chambers as fibrillar bodies are replenished with dietary N provide in lab. The sequential formation model is supported by a NanoSIMS image from LeKieffre et al., 2020 (cropped from their Fig. 2, upper left corner of the 18 h $^{15}$NH$_4^+$ section), where there are coexisting non-enriched and $^{15}$N-enriched fibrillar proteins formed before/after the spike was added.

other is at ‘background’ levels (see their cropped NanoSIMS image in Fig. 8). This suggests that the fibrillar body lacking $^{15}$N-enriched signals accumulated those proteins before the spike was added, supporting their ability to aggregate proteins over time and retain the $^{15}$N/$^{14}$N isotopic composition at the time of their formation.

The uptake of dietary N by fibrillar bodies is demonstrated by the translocation of $^{15}$N into these organelles 20 h after being fed with $^{15}$N-labelled *Artemia* (LeKieffre, unpublished data, 2024; Fig. 9). Notably, the $^{15}$N-enriched fibrillar bodies were not reported in LeKieffre et al. (2020) by the end of their 8 h incubation (see their Fig. 4, upper right).

<figure><img src="image_12.png" /></figure>

**Fig. 9.** TEM and NanoSIMS images of cytoplasm from an *Orbulina universa* specimen, 20 h after being fed with $^{15}$N-labelled *Artemia*. (a) & (d) TEM, (c) & (f) NanoSIMS, and (b) & (e) TEM/NanoSIMS overlay. These unpublished images were part of the LeKieffre et al., 2020 study using living foraminifera collected off Santa Catalina Island, California, USA in 2015. The images show the fibrillar bodies (‘f’), thought to produce organic matrix proteins, and dinoflagellate symbionts (‘d’) that dominate the initial uptake and incorporation of $^{15}$N-enriched $NH_4^+$ from digested prey. This supports our hypothesis that dietary N is translocated directly into the organelle that produces the matrix proteins which are sequestered in the foraminifera chamber wall. Because digestion metabolizes 100 % of the vacuole prey N, there is no isotopic fractionation between prey and $NH_4^+$ so the matrix proteins in the chamber wall have similar isotopic values to the prey $\delta^{15}$N.
This discrepancy might be due to two possibilities: it might take more
than 8 h for dietary N to be digested, released to the protoplasm, and
translocated into fibrillar body proteins. Alternatively, the TEM sections
in LeKieffre et al. (2020) may have missed recently filled ¹⁵N-enriched
fibrillar bodies, as these organelles are more numerous in the latest
formed chambers (Lee et al., 1965; Spero, 1988). While Fig. 9 does not
distinguish between our two scenarios, it does demonstrate that dietary
N is translocated into fibrillar body proteins after digestion and N
release. This supports our model that dietary N is stored in fibrillar
bodies before being incorporated into shell-bound matrix, although the
rate of the process worth further investigation.

Our model also allows us to estimate the residence time of filled
fibrillar bodies in a foraminifer cell. Using the modelled biomass δ¹⁵N
and the dietary δ¹⁵N as two end members (Eq. (1)), we can estimate that
the shell ‘Ch0-Ch1-Ch2’ contains approximately ‘61–68–96 %’ and
‘72–100–100 %’ of N derived from experimental diet in the GSL and the
Tibetan groups respectively (Fig. 7c&d). Since it takes 2.5 days on
average to grow a new chamber under the culture condition (Table S1),
the two groups imply a 5- and 2.5-day period, respectively, to
completely turn over fibrillar bodies. Given the larger isotopic difference
between baseline biomass and the dietary N, the GSL group should
provide a more accurate estimate. We thus speculate that as the internal
N reservoir for shell-bound matrix, fibrillar bodies have a residence time
of approximately 5 days based on our observations. This time may vary
with changes in the feeding rate, i.e. higher feeding rate (greater than 7
nmol N/day in our experiment) may reduce the turnover time. The shell-
bound N is more readily influenced by dietary uptake in the Tibetan
group, which may result from its higher N uptake per feeding event. The
GSL group is fed one Artemia every day, so the feeding rate is more
continuous. The Tibetan group is fed one Artemia every other day. We
adopt this feeding strategy to ensure that both groups have the same
overall N uptake from diet, given the different N content in the two
Artemia strains. However, it is possible that this feeding rate difference
may create some discrepancies in the estimate of the residence time of
the fibrillar bodies. In particular, the old N material in the fibrillar
bodies may be drained more completely in the Tibetan group before
being fed with new N, and the larger uptake of N during each feeding
could replenish the fibrillar bodies more quickly, both of which may
ensure the fibrillar bodies in the Tibetan group to be more quickly
replaced with new materials.

The distinct paths of dietary N incorporation into biomass and shell-
bound reservoirs may partly explain the variabilities between the
biomass and FB-δ¹⁵N in field observations (Smart et al., 2018, 2020). As
shell chambers increase in size through ontogeny, the shell calcite and
shell-bound matrix might thus be biased towards the last larger/more
massive chambers. On the individual level, FB-δ¹⁵N might thus reflect
the most recent dietary N, which would likely involve larger
zooplankton, while biomass δ¹⁵N represents integrated N consumption
over the foraminifer's life. However, these variabilities would be
smoothed away averaged across multiple individuals, so the biomass
and FB-δ¹⁵N are observed to show close similarity in the modern ocean
when averaged among different measurements and plankton tows from
the same sampling location/site. Exceptions may exist in some cases,
and are discussed in the next section.

4.3. *From culture experiments to field observations*

In natural environments, dietary N of planktic foraminifera likely
derives from a fraction of the surface suspended particulate nitrogen
(SPN) pool, dominated by living organisms such as phytoplankton,
various zooplankton, and bacteria (Anderson et al., 1979; Spindler et al.,
1984). Observations from both nitrate-depleted and nitrate-rich regions
report strong correlation among FB-δ¹⁵N, δ¹⁵N-SPN, and δ¹⁵N of sinking
particles in plankton tows and sediment traps (Ren et al., 2012; Smart
et al., 2018, 2020). Our culture experiments confirm this, by showing
the tight δ¹⁵N coupling between diet and shell-bound N, and together

they argue that FB-δ¹⁵N is primarily reflective of dietary N sources.

Observations from the modern ocean also show elevated δ¹⁵N in
foraminifera biomass and shell-bound matrix relative to δ¹⁵N-SPN, even
in dinoflagellate-bearing species (Ren et al., 2012; Smart et al., 2018,
2020). This δ¹⁵N elevation is 1.2‰ in the Southern Ocean (Orbulina
universa), and 2.0‰ in the Sargasso Sea (averaged among all
dinoflagellate-bearing species). This elevation is lower than the typical
δ¹⁵N difference between heterotrophs and their diet (Checkley and
Miller, 1989; Minagawa and Wada, 1984), but is still significant
comparing to the negligible trophic effect in our experiments. It is likely
that the dinoflagellate-bearing planktic foraminifera consume a fraction
of the SPN with higher δ¹⁵N relative to bulk SPN. In the subtropical
Atlantic, eukaryotic phytoplankton is found to have similar δ¹⁵N to the
underlying thermocline NO₃, and higher than the prokaryotic phyto-
plankton (Fawcett et al., 2011). If the dinoflagellate-bearing species
primarily consume eukaryotic phytoplankton in the surface ocean, we
may still expect strong δ¹⁵N correlation between SPN and foraminifera
in the absence of any trophic effect.

The turnover time of SPN in the surface ocean ranges from days to months (Eppley et al., 1983), and δ¹⁵N-SPN is determined by the δ¹⁵N of new N sources, NO₃ uptake and export, as well as NH₄⁺ recycling in the euphotic layer (Fawcett et al., 2014; Lourey et al., 2003). The biomass δ¹⁵N of dinoflagellate-bearing planktic foraminifera reflects integrated N consumption over its 2 to 4 week lifespan (e.g., Bé et al., 1977; Erez et al., 1991), while its FB-δ¹⁵N could be weighted towards the dietary intake at the end of the foraminifera's life cycle. If the δ¹⁵N-SPN in the surface ocean remains constant over the life cycle of the planktic foraminifera, both biomass and FB-δ¹⁵N are expected to be similar and vary with surface δ¹⁵N-SPN.

In the oligotrophic regions, both the concentrations and δ¹⁵N values of the SPN remain relatively constant through the year (Altabet, 1988), so the biomass- vs. FB-δ¹⁵N difference should be small in these environments. However, significant spatial and temporal changes in δ¹⁵N-NO₃ and δ¹⁵N-SPN at surface ocean are observed in the upwelling regions (DiFiore et al., 2006; Rafter et al., 2013). According to Rayleigh fractionation kinetics, the isotopic composition of the residual surface NO₃ pool (and thus of its SPN products) increases exponentially with NO₃ drawdown (Altabet, 2006; DiFiore et al., 2006). Hence, δ¹⁵N-NO₃ changes are most apparent at the end of production periods or at the edge of upwelling region. Foraminifera produced during these periods or at these regions may develop isotopic difference, with FB-δ¹⁵N values rising above the biomass δ¹⁵N. For instance, the FB-δ¹⁵N in *Globorotalia truncatulinoides* is consistently higher compared to its biomass δ¹⁵N from the same tows in the Southern Ocean, which is attributed to different integration time and dietary shift during its seasonal migration to shallow waters (Smart et al., 2020). Our unpublished data from the Central Equatorial Pacific also show a clear biomass- vs. FB-δ¹⁵N difference in *T. sacculifer* (n = 6 for biomass; n = 2 for shell-bound N analysis) and *Pullentiatina obliquiloculata* (n = 3 for biomass; n = 1 for shell-bound N analysis) measured from the same net tows conducted in September 2010 (Cruise KH10-4). The FB-δ¹⁵N is 3.5‰ and 4.0‰ higher than the biomass δ¹⁵N for the two species, respectively (Fig. 10). The equatorial Pacific is characterized by strong seasonal upwelling, which generally peaks in boreal summer and fall due to the strong cross-equatorial trade winds (Chang, 1994, 1996; Li and Philander, 1996). During periods with strong upwelling, thermocline NO₃ is brought into the euphotic layer, where preferential biological uptake of the ¹⁴N isotope enriches the δ¹⁵N of the surface NO₃ over time (Rafter and Sigman, 2016). The δ¹⁵N composition of foraminifera diet in surface ocean is thus likely to increase during the life cycle of these foraminifera, causing the FB-δ¹⁵N values to be more elevated compared to their biomass. In this perspective, FB-δ¹⁵N in a sedimentary record from the upwelling regions may be biased towards higher δ¹⁵N values during the more productive seasons. However, as sedimentary archive integrates foraminifera produced during different seasons, any seasonality-induced bias only affects paleoceanographic reconstructions if this seasonality
<figure><img src="image_13.png" /><figcaption>**Fig. 10.** Unpublished tow-collected foraminiferal biomass and FB-δ¹⁵N from the Central Equatorial Pacific (5°30′N, 164°12′W and 7°06′N, 166°30′W; Cruise KH10-4).</figcaption></figure>

has changed over time.

We acknowledge that the chamber-specific FB-δ¹⁵N data presented in this study are based on single measurements derived from one planktic foraminifera species. Therefore, our findings may not fully represent the general behavior of all foraminifera. In natural environments, additional factors, such as differences in life span and dwelling depth, may further contribute to species-specific biomass- vs. FB-δ¹⁵N patterns. Despite these potential limitations, the dietary influence inferred from pooled analysis of over 100 individuals reveals isotopic shifts in the experiment that are well beyond the uncertainty of analytical errors. Our study suggests that changes in dietary uptake could be a plausible explanation for the field observations.

## 5. Conclusions

Previous studies in modern water columns and surface sediments show: (1) Similar δ¹⁵N values for biomass and shell-bound N in planktic foraminifera; (2) Strong temporal and spatial δ¹⁵N correlation between foraminifera and SPN in the surface ocean; (3) Dinoflagellate-bearing species have lower δ¹⁵N than non-dinoflagellate bearing planktic foraminifera species.

Our culture experiments provide a mechanistic explanation for these observations. The lack of trophic enrichment in *T. sacculifer* result from minimizing N waste, likely through active internal N recycling processes between the host and their dinoflagellate symbionts. Comparison with other planktic foraminifera without symbionts or with other types of symbionts is needed to validate this. Our findings help to simplify interpretations of the FB-δ¹⁵N of dinoflagellate-bearing planktic foraminifera.

We identify a direct pathway for incorporating newly metabolized dietary N into shell-building proteins, without mixing with the bulk biomass. This may indicate a prioritization for shell construction in planktic foraminifera. The dietary N signal is instantaneously locked within the shell matrix without isotopic fractionation, permitting the application of FB-δ¹⁵N to reconstruct past changes in nutrient sources and cycling in the surface ocean.

## CRediT authorship contribution statement

**Wei-Ning Fang:** Writing – original draft, Visualization, Investigation, Formal analysis, Conceptualization. **Oscar Branson:** Writing – review & editing, Supervision, Investigation, Funding acquisition. **Er-**

**Wen Yang:** Investigation. **Wen-Hui Chen:** Investigation. **Ren-Yi Cai-Li:** Investigation. **Howard J. Spero:** Writing – review & editing, Investigation. **Jennifer Fehrenbacher:** Writing – review & editing, Investigation. **Lael Vetter:** Writing – review & editing, Investigation. **Charlotte LeKieffre:** Writing – review & editing, Investigation. **Haojia Ren:** Writing – original draft, Supervision, Funding acquisition, Conceptualization.

## Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgments

This study is supported by MOST 107-2636-M-002-002, MOST 111-2116-M-002-032-MY3, NSTC 112CC-4 to H. Ren, and Leverhulme Trust RL-2022-05 and a Royal Society Exchange Grant IEC\R3\223073 to O. Branson. We thank M. Kelsey Lane, Theresa Fritz-Endres, Wen-Hsuan Liao, William Dorrien-Smith, Yurika Ujiie, Yoshiyuki Ishitani, Laurie Charrieau, Pratyusha Madhnure, Sophie Nuber, Heng-Chun Kuo, Madison East and Alice Ball for their assistance during field sampling and culture experiments in 2019 and 2023; Yi-Chi Chen, Hailing Ren, Yu-Chen Liu, Ya-Hsien Lan for their help with subsequent sample analyses. We appreciate the support and hospitality from staff at the Marine Science Center at Green Island Marine Research Station, especially the help from Ming-Jay Ho and Hsiang-Yi Chen.

## Supplementary materials

Supplementary material associated with this article can be found, in the online version, at doi:10.1016/j.epsl.2025.119231.

## Data availability

The data presented in this paper are available in the supplementary files.

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