# Species-specific mechanisms of benthic foraminifera in response to shell dissolution

Maxime Daviray, Milla Lesut, Emmanuelle Geslin, Edouard Metzger, Inge van

Dijk, Julia Courtial

► To cite this version:

Maxime Daviray, Milla Lesut, Emmanuelle Geslin, Edouard Metzger, Inge van Dijk, et al. Species-specific mechanisms of benthic foraminifera in response to shell dissolution. Marine Micropaleontology, 2025, 200, pp.102490. 10.1016/j.marmicro.2025.102490. hal-05187663

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# Species-specific mechanisms of benthic foraminifera in response to shell dissolution

Maxime Daviray*, Milla Lesut, Emmanuelle Geslin, Edouard Metzger, Inge van Dijk, Julia Courtial

Univ Angers, Nantes Université, Le Mans Université, CNRS, Laboratoire de Planétologie et Géosciences, LPG UMR 6112, 49000 Angers, France

## ARTICLE INFO

**Keywords:**

Haynesina germanica

Ammonia confertitesta

acidification

kleptoplast

behavioural traits

## ABSTRACT

Ammonia confertitesta and Haynesina germanica are two common estuarine benthic foraminifera subject to sediment acidification. Nevertheless, mechanisms involved in their response to acidification are still poorly understood. Since *H. germanica* is kleptoplastic and photosynthetically active, unlike *A. confertitesta*, these species were cultured in controlled experiments to determine whether these mechanisms could mitigate acidification-induced shell dissolution. Both living and dead specimens were incubated at two pH (8.0 and 6.8) and two light conditions (0 and 24 µmol photon m⁻²·s⁻¹) for 18 days. For each species, respiration and photosynthesis rates were calculated based on oxygen measurements. At the end of incubation, foraminiferal viability was assessed with CellTracker Green™ biomarker, and each test was categorised according to a dissolution scale (DS) using SEM. For both species, in acidic conditions, the tests of dead specimens were significantly more dissolved than the tests of living specimens, suggesting active mechanisms providing tolerance to acidification. For the living specimens, no significant difference in the DS distribution was observed between the two species at both conditions, suggesting that kleptoplast photosynthetic activity in *H. germanica* does not provide additional resistance to acidification. Until at least day 12, respiration data revealed a different biological activity for the two species, and we observed distinct behaviours (e.g., encystment and pseudopod emission). These suggest each species exhibits species-specific responses to cope with acidification. On day 18, respiration rates and binocular observations showed low biological activity, suggesting dormancy or death. Further investigation is required to identify the cellular mechanisms involved to counter acidification stress.

## 1. Introduction

Carbonate system parameters in estuarine ecosystems undergo large variations on daily (i.e., the ebb and flow tide) to seasonally (i.e., winter flooding/summery low river discharge) time scales, due to both natural and anthropogenic factors (i.e., temperature, salinity, pH, organic matter inputs). Therefore, these transitional environments are subjected to a wide range of acidification processes leading to carbonate dissolution in sediments, such as low pH/low alkalinity freshwater discharges (Cai et al., 2021), organic and inorganic environmental pollution (Andersson et al., 2005; Wallace et al., 2014), cable bacteria activity (Nielsen et al., 2010) or even local oxygen release and organic acid exudation from seagrass rhizosphere (Brodersen et al., 2018; Long et al., 2008). These processes pose significant threats to calcifying organisms, such as benthic foraminifera.

Benthic foraminifera are present in high abundances in estuarine

environments which provide a wide diversity of habitats (Debenay et al., 2000, 2006; Horton and Murray, 2007). Despite their adaptation to transitional environments, estuarine species with a calcareous shell, or so-called test, can be affected by sediment acidification processes. Numerous studies report *in situ* observations of advanced dissolution of the calcareous tests of living foraminifera, at times even revealing the inner organic lining (Alve and Nagy, 1986; Buzas-Stephens and Buzas, 2005; Cesbron et al., 2016; Charrieau et al., 2018a; Daviray et al., 2024; Fouet et al., 2024; Haynert et al., 2012; Polovodova and Schonfeld, 2008; Schönfeld and Mendes, 2022). Species-dependent foraminiferal responses to acidification have been highlighted in previous laboratory cultures of large benthic photosymbiont-bearing foraminifera (e.g., Charrieau et al., 2022; Dämmer et al., 2023; Fujita et al., 2011; Kuroyanagi et al., 2021; Prazeres et al., 2015; Reymond et al., 2013; Schmidt et al., 2014; Sinutok et al., 2011) or small benthic foraminifera (Allison et al., 2010; Charrieau et al., 2018b; Haynert et al., 2011, 2014;

<footnote>* Corresponding author.
E-mail address: maxime.daviray@outlook.com (M. Daviray).</footnote>
Keul et al., 2013). This feature has also been documented in taphonomical experiments (Haynert et al., 2014; Nguyen et al., 2009).

In this study, we focus on *Ammonia confertitesta* and *Haynesina germanica* which are common species on European intertidal areas (Debenay et al., 2000; Fouet et al., 2022; Francescangeli et al., 2020; Murray and Alve, 2000; Sen Gupta, 1999). These two hyaline species have a trochospiral and planispiral perforated carbonate test, respectively. They have different food sources and feeding strategies: *Ammonia* is an omnivorous genus and feeds from organic detritus to microalgae and meiofauna (Chronopoulou et al., 2019; Dupuy et al., 2010; Haynert et al., 2020; Mojtahid et al., 2011; Pascal et al., 2008; Schweizer et al., 2022; Wukovits et al., 2018), whilst *H. germanica* feeds mainly on large diatoms (Austin et al., 2005; Chronopoulou et al., 2019; Jesus et al., 2021; Schweizer et al., 2022; Ward et al., 2003). However, unlike *Ammonia*, *H. germanica* is a kleptoplastic foraminifera (Cesbron et al., 2017; Jauffrais et al., 2016, 2017; Knight and Mantoura, 1985; Lopez, 1979) that retains photosynthetically active chloroplasts, known as “kleptoplasts”, from the diatoms it feeds on (Pillet et al., 2011). These kleptoplasts can remain photosynthetically active for up to 21 days in the laboratory after field sampling (Jauffrais et al., 2017), allowing *H. germanica* a mixotroph strategy. The kleptoplasty observed in foraminifera is thought to constitute an additional carbon supply and could also be involved in the uptake of inorganic nitrogen or sulphate (Grzymski et al., 2002; Jauffrais et al., 2019a, 2019b; LeKieffre et al., 2018). The photosynthetic activity involves the local consumption of dissolved CO₂ that can induce an increase in pH in the microenvironment of the specimen (Glas et al., 2012; Jørgensen et al., 1985; Rink et al., 1998; Zerveas et al., 2021).

Previous observations showed a dominance of *H. germanica* and lower number of *Ammonia* spp. in the first centimetres of the sediment under acidic conditions (Daviray et al., 2024; Fouet et al., 2022; Francescangeli et al., 2020). Living *H. germanica* specimens with slightly dissolved tests were also observed in a monospecific community under acidic conditions (Daviray, 2024). The mechanisms involved in the response to acidification are still poorly understood. Based on the field observations, we hypothesised species-specific responses of *H. germanica* and *A. confertitesta* to acidification-induced shell dissolution. In this study, we investigated the potential role of test structure (general shell morphology and thickness) and active mechanisms (metabolic and behavioural) by incubating both living and dead foraminifera in acidified and non-acidified conditions. This allows us to test if living foraminifera can actively resist shell dissolution, and if photosynthetic activity of kleptoplasts provides better tolerance for *H. germanica* compared to kleptoplast-barren *Ammonia* spp..

## 2. Material & Methods

### 2.1. Foraminiferal Sampling

The first millimetres of sediment were sampled at low tide on 6 March 2024 from the Bourgneuf Bay mudflat (French Atlantic coast; 47.013° N, 2.019° W). Seawater (salinity: 26.5; pH: 8.09) was taken from a channel near the mudflat. In the laboratory, seawater was pre-filtered using Fisherbrand® 300-µm filters (Fisher Scientific, Illkirch) and microfiltered on 0.45-µm GF/C membranes (Whatman, Maidstone, UK). A small amount of seawater (~1 L) was filtered through a Minisart® regenerated cellulose syringe 0.2-µm filter (Fisher Scientific, Illkirch). The sediment was kept overnight in the laboratory at room temperature.

The >150-µm fraction of sediment was sieved with 0.2-µm microfiltered seawater. Living specimens (i.e., cytoplasm coloured and evenly distributed except in the last chambers) and dead specimens (i.e., empty tests) with glassy and transparent tests of *Ammonia* spp. and *H. germanica* were collected with a brush under a binocular magnifier (Leica MZ 12.5). The specimens collected therefore showed no mark of dissolution. Specimens with such marks (e.g. whitish test, cracks,

missing chambers) were therefore excluded. Two successive baths of 0.2-µm microfiltered seawater were used to limit bacterial and microalgal contamination. A total of 360 living specimens per species, 360 dead *H. germanica* specimens and 300 dead *Ammonia* spp. were selected. In Bourgneuf Bay mudflat, *Ammonia* spp. included two pseudocryptic species belonging to the *Ammonia tepida* morphogroup (Cushman, 1926): *A. confertitesta* Zheng, 1978, and *A. aberdoveyensis* Haynes, 1973. They are difficult to distinguish with a binocular (Richirt et al., 2021) and were identified at a species level at the end of the experiment, based on the specimens from control conditions (n = 82), with a DEBEN Hitachi T4000 scanning electron microscope (SEM). These observations revealed a ~95 % dominance of *A. confertitesta*. Based on this result, our knowledge (pers. comm. Emmanuelle Geslin and Julia Courtial,) and two published theses which focussed on this specific site (Choquel, 2021; Richirt, 2020), we will assume that the *Ammonia* spp. specimens belong to *A. confertitesta* for the remainder of this study.

### 2.2. Experimental Set-up

The incubations started two days after the field sampling and were carried out for 18 days in a temperature-controlled room at 16°C (porewater temperature on site). Four transparent plastic aquaria (5.2 L) were filled with 4 L of 0.45-µm filtered seawater (Figure 1). Each aquarium was fitted with a clamp cover to keep the medium airtight and a 2-cm diameter opening in the lid for the probes, which can be closed with a rubber stopper. A magnetic stirrer as used to oxygenate and homogenise the system. Two different lighting settings were set up to investigate the potential role of the photosynthetic activity of kleptoplast foraminifera in resistance to acidification: two aquaria were subjected to permanent darkness guaranteed by enclosing the culture set-up in an opaque black bag. The other two were exposed to a 6h/18h light/dark cycle under a cold-light fluorescent lamp (Lumix day light, L30W/865, Osram). This photoperiod assumes that first sediment millimetres in turbid environments as intertidal mudflats are exposed to light during diurnal low tides. Light intensity was below the alteration value for the photosynthetic metabolism of *H. germanica* (70 µmol photons.m⁻².s⁻¹; Jauffrais et al., 2016), and it was slightly above the constant half-saturation (between 13 and 21 µmol photons.m⁻².s⁻¹; Jauffrais et al., 2016) around 24 µmol photons.m⁻².s⁻¹ (MQS-B quantum sensor and ULM-500 photometer, Walz).

In each light setting, one aquarium was acidified by bubbling with CO₂. The CO₂ bubbling system consisted of a vacuum flask containing 1 L of demineralised water and 60 g of sodium bicarbonate (NaHCO₃). This solution was bubbled with ambient air to release the dissolved CO₂ into the aquarium to acidify the seawater to the target pH≈6.8. This target pH was chosen to mimic porewater microhabitat acidity as it is a low value often observed *in situ* in the first millimetres of sediment in transitional environments (Alve and Nagy, 1986; Cesbron et al., 2016; Fouet et al., 2024; Schönfeld and Mendes, 2022). The second aquarium was unaltered, with a pH between 7.7-8.0. Temperature, salinity and pH were measured daily with a multimeter (Multi 3620 IDS set C, WTW) with TetraCon 925 and Sentix 940 probes (WTW). The pH sensor was calibrated using three NBS standard solutions (4.0, 7.0, 9.2). During the experiment, if an increase in pH was observed in the acidic aquarium, the CO₂ bubbling system was reactivated to achieve the target value of pH≈6.8. Living and dead specimens of *A. confertitesta* and *H. germanica* were distributed in 40-mm borosilicate glass VWR® crystallisers, filled with 0.2-µm microfiltered seawater and closed with a net (100-µm mesh) to prevent specimens from escaping. No sediment was added to better control and stabilise the seawater carbonate chemistry. The dissolution of carbonate particles in the sediment could disrupt DIC and alkalinity in the relatively limited water volume used in this experiment, as discussed in Bernhard et al. (2021). Three crystallisers each species containing thirty live specimens, three crystallisers containing thirty dead *H. germanica* and three crystallisers containing twenty-five dead *A. confertitesta* are placed in each aquarium corresponding to a specific
<figure><img src="image_3.png" /><figcaption>**Figure 1.** Schematic illustration of the experimental set-up. Two pH conditions (control~8.0 and acidified~6.8) were tested under two lighting settings (A: light with 6/18h light/dark cycle and B: darkness) resulting in a total of four 5-L aquaria. The CO₂ bubbling system consisted of bubbling ambient air into a vacuum flask containing a solution of sodium bicarbonate to release the dissolved CO₂ into the aquarium to be acidified. A magnetic stirrer as used to oxygenate and homogenise the system. Three batches of 30 living specimens per species, three batches of 30 dead H. germanica (1) and three batches of 25 dead A. confertitesta (2) were incubated in each aquarium.</figcaption></figure>

light and pH condition. Because the three crystallisers were together in the same aquarium, they are pseudo-replicates and not true replicates.

## 2.3. Seawater carbonate chemistry

Every two days, 10 mL of seawater was sampled in triplicate from each aquarium to analyse total alkalinity (TA) using an automatic titration system (Titrino 785 Metrohm) with a HCl standard solution of 0.1 M (Gran, 1952). Here we assumed that TA consisted mainly of carbonate species. To compensate for the seawater sampling, 0.45-µm filtered seawater was added occasionally to keep the volume in the experiment as constant as possible. Physio-chemical parameters of each experimental condition are summarized in Table 1. The pH (NBS scale), TA, water temperature, and salinity were used to calculate dissolved inorganic carbon (DIC) and calcium carbonate saturation state ($\Omega_{Ca}$) using the CO2Sys v3.0 software (Lewis et al., 1998; Pierrot et al., 2021) with equilibrium constants of Cai and Wang (1998) for estuarine environments. For each aquarium, all these parameters remained stable with very slight variations over time. Total alkalinity and DIC were higher in acidified conditions compared to controlled conditions (around 2.5 vs

<caption>**Table 1**</caption>

Statement of carbonate parameters for the incubation duration (18 days). Values indicate mean ± standard deviation (n = 10). Dissolved Inorganic Carbon (DIC) and calcite saturation state ($\Omega_{Ca}$) were calculated from temperature, salinity, pH and total alkalinity (TA) using the CO2Sys software.

<table><thead><tr><th rowspan="2">Lighting setting<br>pH condition</th><th colspan="2">Light/dark cycle</th><th colspan="2">Dark</th></tr><tr><th>Controlled<br>(pH≈8.0)</th><th>Acidified<br>(pH≈6.8)</th><th>Controlled<br>(pH≈8.0)</th><th>Acidified<br>(pH≈6.8)</th></tr></thead><tbody><tr><td>T (°C)</td><td>17.4 ± 0.5</td><td>17.1 ± 1.1</td><td>17.3 ± 0.7</td><td>17.2 ± 0.7</td></tr><tr><td>Salinity</td><td>27.1 ± 0.1</td><td>27.1 ± 0.1</td><td>27.1 ± 0.1</td><td>27.1 ± 0.1</td></tr><tr><td>pH</td><td>7.97 ± 0.13</td><td>6.81 ± 0.05</td><td>7.89 ± 0.11</td><td>6.80 ± 0.04</td></tr><tr><td>TA (mM)</td><td>2.33 ± 0.03</td><td>2.49 ± 0.09</td><td>2.31 ± 0.07</td><td>2.51 ± 0.10</td></tr><tr><td>DIC (mM)</td><td>2.21 ± 0.06</td><td>2.95 ± 0.13</td><td>2.22 ± 0.04</td><td>2.98 ± 0.14</td></tr><tr><td>$\Omega_{Ca}$</td><td>2.59 ± 0.62</td><td>0.20 ± 0.02</td><td>2.16 ± 0.50</td><td>0.20 ± 0.02</td></tr></tbody></table>

2.3 mM for alkalinity and 2.9 vs 2.2 mM for DIC respectively). Calcite saturation state was undersaturated in pH≈6.8 conditions compared to controlled conditions (0.2 vs ≥ 2.0 respectively).

## 2.4. Oxygen consumption and production measurements

As the foraminifera had been manipulated on day 1, the first oxygen measurements were done after a day/night of stabilisation on day 2. Then, oxygen measurements were performed on day 12 and 18 based on observations of the dissolution stage of the living foraminifera (see section 2.5 below). On day 2, three sub-sets of five living specimens of both species were randomly selected from the control condition (pH≈8) under a light/dark cycle, considered as their initial reference state. Oxygen production/consumption measurements were conducted on these specimens to assess their physiological state at the start of the experiment. On days 12 and 18, one group of five specimens of *A. confertitesta* and *H. germanica* from each pH and lighting setting, was randomly collected to measure O₂ production and consumption. For *H. germanica* in the control condition with light/dark cycle, a second group of five specimens was studied to confirm any reduction of the photosynthetic activity. Foraminiferal consumption rates were calculated from specimens/overlying water oxygen gradients (Cesbron et al. (2017) adapted from Høgslund et al. (2008)), using a 50-µm diameter Clark-type microelectrode (OXI50 - Unisense; Revsbech, 1989) mounted on a motorised micromanipulator (Unisense).

In short, measurements were carried out on groups of five foraminifera in a glass microtube with a 1-mm internal diameter, filled with 0.2-µm microfiltered seawater maintained in an aquarium of demineralised water at stable temperature (16°C). O₂ measurement started at 200 µm above the foraminifera until 1200 µm with a 100-µm steps. Each O₂ profiling was repeated between 5 and 25 times in light (700 µmol photons.m⁻².s⁻¹) and then in darkness. Five parallel O₂ gradients were selected based on the best R² (above 0.995). The O₂ flux was determined using the Fick's law and then multiplied by the exchange surface area to obtain the O₂ rate of the foraminifera pool. After the O₂ measurements, the specimens were photographed under a stereomicroscope and
reintroduced into their respective experimental condition. Using pho-
tographs with FIJI (Schindelin et al., 2012) the diameter of the fora-
minifera was measured and their volume calculated, estimating that the
test has a half-sphere shape (Geslin et al., 2011). The biovolume (cell
volume $V_B$) was considered as 75 % of the test volume according to Eq.
1:

$$
V_B = 0.75 * \frac{1}{2} \left[ \frac{4}{3} * \pi * \left( \frac{d_1 + d_2}{4} \right)^3 \right] \quad (1)
$$

with $d_1$ and $d_2$, the diameters along the length and width of the foraminifera. $O_2$ consumption and production rates $R(O_2)$ were related to the biovolume of the five foraminiferal specimens according to Eq. 2:

$$
R(O_2) = -D_0 \frac{dC}{dz} * \frac{S}{n * V_B} \qquad (2)
$$

with $D_0$, the diffusion coefficient of dioxygen, $\frac{dC}{dz}$, the oxygen gradient diffusion, S, the glass microtube section, n, the number of specimens in the microtube, and $V_B$, the biovolume. The net photosynthesis rate *NPR* was estimated subtracting the $O_2$ consumption rate in light on from the rate in light off as Eq. 3:

$$
\mathrm{NPR} = R(\mathrm{O}_2)_{\text{light on}} - R(\mathrm{O}_2)_{\text{light off}} \quad (3)
$$

2.5. Foraminiferal observations and evaluation of test dissolution

Observations with the binocular microscope were carried out on
living and dead foraminifera from each condition and for both species on
day 2, 12 and 18 to assess their state. Purely observations of the fora-
minifera from one crystalliser were carried out for each condition on day
7 using an optical inverted microscope (Axiovert 25 HBO 50/AC, Zeiss
equipped with a Nikon D750 camera). The aim was to quickly assess the
overall state of the foraminifera before repeating a mid-term oxygen
measurement. At the end of the 18 days of incubation, all recovered
specimens were labelled with fluorescent CellTracker Green™
biomarker (Invitrogen; 1 mg.L⁻¹; Bernhard, 2000) overnight at room
temperature, and then fixed in ethanol (98 %). An epifluorescent ste-
reomicroscope equipped with epifluorescence optics (Olympus SZX-16)
was used to distinguish surviving foraminifera (fluorescent labelled) and
to calculate survival rates, defined here as the ratio of fluorescent
specimens to the total number of recovered specimens in each crystal-
liser (pseudo-replicate) at the end of the incubation.

All foraminiferal tests were observed using a DEBEN Hitachi T4000
scanning electron microscope (SEM) (non-metallised samples; 15kV; wd
= 7.7 mm; partial vacuum = -60 Pa). Their dissolution stage (DS) was
classified with a dissolution scale based on Daviray et al. (2024). The
established scale for Ammonia spp. Starts from dissolution stage 0 (DS-0,
"intact test") to dissolution stage 5 (DS-5, "fully dissolved test" revealing
the inner organic lining). Since no DS-5 was observed for H. germanica
by Daviray et al. (2024), the scale goes from DS-0 to dissolution stage 4
(DS-4, "star-shape"). In this study, we added a sixth stage for
H. germanica (DS-5, "fully dissolved test") for specimens which could not
be collected due to their excessive fragility.

Unrecovered specimens at the end of the experiment were catego-
rized differently for the groups “dead foraminifera” and “living speci-
mens”. Dead foraminifera that were missing at the end of the
incubations in the acidified conditions were considered as fully dis-
solved. These specimens are noted as “lost” and attributed to DS-5 group
for statistics. Unrecovered foraminifera from the “living specimens”
group, are excluded from further analysis and not used for calculating
survival rates as we cannot identify the cause of their disappearance.
Loss could be due to dissolution, or movement of the specimens out of
the container, or handling errors despite the efforts to check and clean
the nets carefully during the harvesting. Resulting DS-groups from
different conditions and between species were compared statistically

using the R software. As the data did not follow a normal distribution
and some of them were ordinal, a Kruskal-Wallis test (p-value < 0.05,
with a Bonferroni correction) followed by a Fisher LSD (Least Significant
Difference) were applied to determine which groups were significantly
different from the others.

3. Results

3.1. Foraminiferal survival rate and test recovery

At the end of the experiment, regardless of the species, fewer spec-
imens were recovered under acidified conditions compared to the con-
trol for the incubated living specimens (Table 2). Additionally, the
missing A. *confertitesta* specimens were greater than those of
*H. germanica* for each condition, with the highest loss for both species
occurring in the dark acidified condition. From the specimens recovered,
the survival rate at the end of the experiment was over 78 % for all
conditions, with higher survival rates in the acidified conditions
compared to the controls for both species. On species level, the survival
rates were consistently higher for *H. germanica* than for *A. confertitesta*
across all conditions, particularly in darkness, where *H. germanica*
exhibited an average survival rate approximately 7.5 % higher than that
of *A. confertitesta* (Table 2).

3.2. Observations throughout the incubation

On day 7 of the incubation, observations revealed a transparent film at the bottom of the crystallisers, which may correspond to the development of a bacterial biofilm. This biofilm was more developed in aquariums exposed to the light/dark cycle, particularly in the acidified condition. In the acidified light/dark cycle aquaria, living specimens of both species were observed agglomerating particles of this biofilm around their cells, a process known as "encystment" (Heinz et al., 2005). A relatively higher number of *A. confertitesta* individuals were encysted compared to *H. germanica*. Photographs of observations performed on day 12 revealed encystment of few specimens of *A. confertitesta* and *H. germanica*, and emissions of pseudopodia for *A. confertitesta* (Supplementary Figure 1). Encysted specimens were no longer observed on day 18.

Microscope observations showed that the living and dead specimens
in the control conditions were in the same dissolution state on day 7 as at
the beginning of the experiment for both species: a mix of DS-0 and DS-1.
On the contrary in the acidified conditions, most of the living specimens

**Table 2**

Number of living specimens per species recovered at the end of the incubation and survival rates for Ammonia confertitesta and Haynesina germanica in relation to light and pH conditions. Values indicate means ± standard deviation between the three pseudo-replicates incubated under the same condition. The number of specimens recovered corresponds to those present in a crystallizer at the end of incubation, given that thirty foraminifera were introduced per crystallizer at the start of the experiment. Survival rate was calculated on recovered specimens.

<table>
  <thead>
    <tr>
      <th rowspan="2">Lighting setting<br>pH condition</th>
      <th colspan="2">Light/dark cycle</th>
      <th colspan="2">Dark</th>
    </tr>
    <tr>
      <th>Controlled<br>(pH≈8.0)</th>
      <th>Acidified<br>(pH≈6.8)</th>
      <th>Controlled<br>(pH≈8.0)</th>
      <th>Acidified<br>(pH≈6.8)</th>
    </tr>
  </thead>
  <tbody>
    <tr>
      <td><strong>A. confertitesta</strong><br>Recovered<br>individuals</td>
      <td>27 ± 3</td>
      <td>23 ± 6</td>
      <td>28 ± 2</td>
      <td>20 ± 5</td>
    </tr>
    <tr>
      <td>Survival rate<br>(%)</td>
      <td>83 ± 13</td>
      <td>95 ± 1</td>
      <td>79 ± 1</td>
      <td>87 ± 17</td>
    </tr>
    <tr>
      <td><strong>H. germanica</strong><br>Recovered<br>individuals</td>
      <td>30 ± 1</td>
      <td>26 ± 3</td>
      <td>29 ± 2</td>
      <td>25 ± 1</td>
    </tr>
    <tr>
      <td>Survival rate<br>(%)</td>
      <td>88 ± 2</td>
      <td>98 ± 2</td>
      <td>86 ± 3</td>
      <td>95 ± 5</td>
    </tr>
  </tbody>
</table>
showed a whitish test, and some of them seemed to be dead as soon as
day 7 (i.e., empty and white test). Furthermore, in the acidified darkness
condition, the cytoplasm of living *A. confertitesta* seemed more stained
and their tests less fragile than living *H. germanica*. Taking a look at the
tests of initial dead specimens, the *A. confertitesta* shells appeared fragile
and brittle, with in general the last chamber dissolved, whereas the
shells of *H. germanica* were highly damaged and seemed to have almost
reached stage 5 (DS-5).

### 3.3. Test dissolution stages

The proportions of dissolution stages (DS) of living and dead foraminifera in experimental conditions are presented in Figure 2, and median DS specimens per conditions and per species are presented in Figure 3. The dissolution of the tests was significantly greater in the acidified conditions compared to the control conditions, regardless of species, lighting setting, or initial viability (living vs dead specimens). When comparing living and dead foraminifera in acidified conditions, the general dissolution stage of dead foraminifera was much higher compared to the living specimens.

#### 3.3.1. Dissolution of Living Specimens

In the control conditions, recovered living specimens of both species

showed no statistic difference in dissolution, except for the
*A. confertitesta* specimens in darkness (Figure 2). Almost no dissolution
was observed (median DS: DS-0) in most conditions, whilst
*A. confertitesta* incubated in darkness showed slight dissolution (median
DS: DS-1). However, because of the discrepancy in the control, it was not
possible to establish a causal effect for *A. confertitesta* incubated in the
two lighting settings nor comparing the two species in the dark. In
acidified conditions, over 60 % of living *H. germanica* showed a mod-
erate dissolution stage (DS-3) in both lighting settings, and more than 15
% an advanced dissolution stage (DS-4 or DS-5), whilst over 50 % of
living *A. confertitesta* showed a slight dissolution stage (DS-0, DS-1 and
DS-2). Despite those observations, no significant difference was
observed between the two species.

#### 3.3.2. Dissolution of dead foraminifera tests

In the control conditions, the dead *H. germanica* were significantly less dissolved than *A. confertitesta*, related to their stage at the initial collection, preventing a statistical comparison of the effect of pH between the two species (Figure 2). Lost specimens in the dead groups are considered to be fully dissolved. In acidified conditions, the dead loss at the end of incubations was over 45 % for *A. confertitesta* and over 70 % for *H. germanica* (Figure 2). Loss for dead *A. confertitesta* was about 30 % lower than for *H. germanica* in acidified conditions. Dead foraminifera

<figure><img src="image_4.png" /><figcaption>**Figure 2.** Dissolution stages after 18 days of incubation for A) living and B) dead specimens of *A. confertitesta* and *H. germanica* for each pH and lighting settings. The "lost" stage corresponds to missing dead specimens, which are included in the total count (n) of individuals. For statistical analysis, lost specimens are included in the DS-5 group. Lower-case letters denote statistically different groups from the Kruskal-Wallis test (p < 0.05).</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Figure 3.** Ammonia confertitesta and Haynesina germanica specimens after 18 days of incubation in different experimental conditions. From left to right: white light microscopy, epifluorescence microscopy after CTG labelling, and SEM images. The specimens presented here have a dissolution stage (DS) corresponding to the median degree observed for each condition. A white cross indicates no image was acquired</figcaption></figure>

were significatly (p < 0.05) more dissolved in acidified conditions compared to control conditions for each species and lighting settings. In acidified conditions, all the dead specimens were fully dissolved (DS-5), with a lot of missing organic linings (i.e., total disappearance).

### 3.4. Foraminiferal oxygen consumption and production

The oxygen consumption and production rates of *H. germanica* and *A. confertitesta* for experimental conditions on day 2, 12 and 18 are presented in Figure 4. On day 2, the O₂ rate of both species was measured on specimens from the control condition with light/dark cycle. The O₂ consumption of *A. confertitesta* specimens did not differ significantly whether the light was switched on (O₂ rate = 2.31 ± 0.31 pmol O₂·s⁻¹·mm⁻³) or off (O₂ rate = 2.29 ± 0.77 pmol O₂·s⁻¹·mm⁻³). When the light was off, *A. confertitesta* specimens exhibited respiration rates one to three times higher (2.29 ± 0.77 pmol O₂·s⁻¹·mm⁻³) than *H. germanica* (0.91 ± 0.10 pmol O₂·s⁻¹·mm⁻³). For *H. germanica*, the difference of O₂ consumption between the light-off (positive values) and the light-on conditions (negative values) indicated photosynthetic

activity and a net oxygen production. At the beginning of the experiment, the kleptoplasts of *H. germanica* showed a calculated net photosynthesis of 1.54 ± 0.3 pmol O₂·s⁻¹·mm⁻³ (Table 3).

On days 12 and 18, *A. confertitesta* O₂ consumption was higher in acidified condition compared to control condition for both light conditions (Figure 4; Supplementary Table 1). On day 12 for *H. germanica*, specimens from the control condition subjected to the light/dark cycle exhibited sufficient O₂ production to compensate for respiration (net photosynthesis rate of 2.20 ± 0.14 and 5.23 ± 0.27 pmol O₂·s⁻¹·mm⁻³ for the replicate 1 and replicate 2 respectively; Supplementary Table 1). For the *H. germanica* specimens incubated in the darkness conditions, the net photosynthesis rate was in the same order of magnitude (2.86 ± 0.23 pmol O₂·s⁻¹·mm⁻³ in the control condition, and 2.08 ± 0.32 pmol·s⁻¹·mm⁻³ in the acidified condition) without compensation for respiration. In the acidified conditions, photosynthetic activity was lower in the light/dark cycle (0.98 ± 0.18 pmol·s⁻¹·mm⁻³). *Haynesina germanica* specimens incubated in the dark exhibited respiration rates two to three times higher than those incubated in light/dark cycle. On day 18, the net photosynthesis rate of *H. germanica* remained insignificant in all
<figure><img src="image_6.png" /><figcaption>**Figure 4.** Oxygen consumption/production of *A. confertitesta* and *H. germanica* per experimental condition on day 2, 12 and 18 at 16°C. Yellow plots represent the “light on” measurements, and the grey plots represent “light off” measurements. Each bar indicates the average of 5 oxygen profiles. Day 2 oxygen measurements were conducted on three pools of five foraminifera, the error bars represent standard deviation, and the lower-case letters represent the groups from the Kruskal-Wallis test (p < 0.05). On days 12 and 18, oxygen measurements were performed on one pool of five foraminifera, and the error bars represent the technical variability. Two pools of *H. germanica* were used in “light on” condition. Black crosses indicate no data acquisition.</figcaption></figure>

conditions (0.30 ± 0.05 pmol.s⁻¹.mm⁻³), except for control condition in light/dark cycle where the net photosynthesis rate was twice higher (0.73 pmol O₂.s⁻¹.mm⁻³) without compensate for respiration anymore.

# 4. Discussion

## 4.1. Responses of dead and living specimens to acidification

At the end of the experiment, all the empty tests of dead foraminifera incubated in the acidified conditions were severely corroded or fully dissolved in 18 days (DS-5 or lost; Figure 2). The binocular observations on day 7 suggested an early dissolution of dead *H. germanica* tests compared to *A. confertitesta* ones. In the same culture condition, the number of *H. germanica* tests recovered was lower than that of *A. confertitesta* at the end of incubation (≤ 25 % and > 50 % respectively). Since lost tests of dead foraminifera are considered fully dissolved, this suggests a greater sensitivity of *H. germanica* tests to acidification. *H. germanica* has a test wall about 48 % thinner than *Ammonia tepida* for the same number of calcite layers (Cisneros-Lazaro et al., 2022), and the cogwheels, structural blocks of calcite that build up

the foraminiferal test, are smaller for *H. germanica* compared to *A. confertitesta* (van Dijk et al., 2020). Since the suture areas of cogwheels can be potentially more susceptible to alteration (Cisneros-Lazaro et al., 2022), *H. germanica* specimens have a relatively larger surface area potentially susceptible to dissolution. This specific structural difference of tests could therefore explain the discrepancy between the living foraminiferal community and the thanatocoenosis observed *in situ* in sediment submitted to dissolution process (Alve and Nagy, 1986; Daviray et al., 2024; Murray and Alve, 1999). Our observations suggest that the tests of *H. germanica* may have a lower potential to be preserved in the sediment record than these of *A. confertitesta* in acidic conditions. This is consistent with field observations from the Trieux estuary in France (Daviray, 2024) where the living communities were dominated by *H. germanica* while the dead assemblages contain more *A. confertitesta* tests than *H. germanica* ones.

In contrast to the dead specimens, about 70 % of living specimens of both species that are recovered from acidified conditions were only slightly dissolved (from DS-0 to DS-2; Figure 2). These results demonstrate that these two species can maintain shell integrity at low pH conditions. Numerous experiments from literature show varying results
<caption>**Table 3**</caption>

Net photosynthesis rates calculated based on the average O₂ consumption and production rates of *H. germanica* for each experimental condition at 16°C. Values indicate the mean net photosynthesis rate calculated ± standard deviation. These values come from calculations based on O₂ gradient measurements carried out at least five times on the same pool of foraminifera.

<table><thead><tr><th rowspan="2">Lighting setting</th><th colspan="2">Light/dark cycle</th><th colspan="2">Dark</th></tr><tr><th>Control (pH≈8.0)<br>1.04 ±<br>0.02</th><th>Acidified (pH≈6.8)</th><th>Control (pH≈8.0)</th><th>Acidified (pH≈6.8)</th></tr></thead><tbody><tr><td>pH condition</td><td></td><td></td><td></td><td></td></tr><tr><td>Day 2<br>(pmol O₂·s⁻¹·mm⁻³)</td><td>1.74 ±<br>0.39<br>1.80 ±<br>0.03<br>2.20 ±</td><td>/</td><td>/</td><td>/</td></tr><tr><td>Day 12<br>(pmol O₂·s⁻¹·mm⁻³)</td><td>0.14<br>5.23 ±<br>0.27<br>0.53 ±</td><td>0.98 ± 0.18</td><td>2.86 ± 0.23</td><td>2.08 ± 0.32</td></tr><tr><td>Day 18<br>(pmol O₂·s⁻¹·mm⁻³)</td><td>0.12<br>0.93 ±<br>0.09</td><td>0.34 ± 0.16</td><td>0.30 ± 0.05</td><td>0.18 ± 0.08</td></tr></tbody></table>

for test dissolution of various foraminiferal species incubated in conditions of (near) undersaturation with calcite ($\Omega_{Ca} \ge 1$; Charrieau et al., 2022; Guamán-Guevara et al., 2019; Haynert et al., 2011, 2014; McIntyre-Wressnig et al., 2013, 2014; Mojtahid et al., 2023; Prazeres et al., 2015; Vogel and Uthicke, 2012). After one and a half months of incubation at $\Omega_{Ca} = 0.5$, Dissard et al. (2010) did not observe any trace of dissolution in *A. tepida* and even new chambers calcified. In contrast, Charrieau et al. (2018b) showed the test dissolution of *Elphidium crispum* occurring from day 9 when exposed to low pH/calcite undersaturated seawater (pH ~ 7.2, $\Omega_{Ca} \sim 0.6$), and day 17 for *Ammonia* sp.. These observations seem to indicate that responses to acidification vary between species. However, the differences in the experimental set-ups and incubation conditions between these experiments, as well as the qualitative nature of dissolution scales (Charrieau et al., 2018b; Daviray et al., 2024; Schönfeld and Mendes, 2022) cause important limitations when comparing studies. Quantification of test dissolution by conducting 3D analyses by µCT-scan (Choquel et al., 2023; François et al., 2022; Kuroyanagi et al., 2021) would greatly improve the analysis accuracy to allow mass balance approaches. Nevertheless, although in our study difference in the dissolution state was observed between living *A. confertitesta* and living *H. germanica* in this study, the different responses of living and dead specimens presented here is extremely evident. It suggests that living foraminifera have active mechanisms to tolerate acidification-induced shell dissolution.

Counter intuitively, we observed that the survival rates based on CTG incubations were higher in acidic conditions compared to control conditions for both species studied (Table 2). We know that dormancy might cause a minor underestimation of survival rates for symbiont-bearing *Amphistegina gibbose* (Ross and Hallock, 2018). This would suggest that in our study, dormancy occurred in the foraminifera under control conditions rather than in those exposed to acidic conditions, which appears unlikely. Furthermore, as more than 80% of the specimens were labelled, we suggest that even in a state of activity slowdown, about 15 hours incubation in the presence of CTG enables the active cytoplasm to be labelled. We hypothesise that, in acidic conditions, foraminifera may rapidly enter a phase of activity reduction in response to stress. In contrast, specimens in control conditions continue to exhibit normal activity. Then, in the long term, the absence of food may probably result in a higher mortality rate among these active specimens than among those that have already reduced their activity over a period of several days. High survival rates in acidified conditions have been previously observed in experimental studies for small coastal benthic species like *Ammonia aomoriensis* (Haynert et al., 2014), *A. confertitesta*,

*Bulimina marginata*, *Cassidulina laevigata* (Mojtahid et al., 2023), *A. tepida* (Charrieau et al., 2018b), *Bolivina argentea* and *Bulimina marginata* (McIntyre-Wressnig et al., 2014) and endosymbiontial large coastal benthic species (Charrieau et al., 2022; McIntyre-Wressnig et al., 2013; Prazeres et al., 2015; Vogel and Uthicke, 2012). A limited number of studies showed a clear mortality in acidified conditions compared to the control, like for *Elphidium williamsoni* (pH = 7.3; Guamán-Guevara et al., 2019) or *A. aomoriensis* (pH = 7.2; Haynert et al., 2011). The pH conditions in all these previous published experiments were higher (pH ≥ 7.2) than those tested here, over much longer incubation periods (in general over a month) and feeding. Nevertheless, these results support our observations and show the ability of foraminifera to survive low pH conditions. Still, success in survival to exposure to low pH conditions seems to be species-specific. The mechanisms behind these observed differences in species resistance remains enigmatic. In our study, we explored whether the stress caused by acidification had a different impact on the general biological activity of these two species by studying their respiration rates.

Respiration rates on day 2 revealed that *H. germanica* exhibited lower respiration in the dark compared to *A. confertitesta* (Figure 4), suggesting a difference in vital activity. The respirations rates obtained for these two species were of the same magnitude as in the literature (between 0.53 and 3.30 pmol O₂·s⁻¹·mm⁻³ for *A. tepida*, and between 0.43 and 2.65 pmol O₂·s⁻¹·mm⁻³ for *H. germanica*, Supplementary Table 3; Cesbron et al., 2016, 2017; Deldicq et al., 2020; Geslin et al., 2011, and references inside; Jauffrais et al., 2016). In our case the biovolume of *A. confertitesta* was lower than in Geslin et al. (2011) (Supplementary Table 3), however, the relationship between biovolume and O₂ rate highlighted by these authors is not sufficient to explain our observations. The higher respiration measurements in acidified conditions than control conditions for *A. confertitesta* on days 12 and 18 suggest a stress generated by low pH on the contrary of *H. germanica*. Haynesina germanica showed higher respiration rate in the darkness compared to light/dark conditions, and much more in the dark acidified condition. This difference of respiration was not further observed on day 18. It could suggest that *H. germanica* was much more active in the dark and stressed by low pH. The photosynthetic activity of *H. germanica* will be discussed later. In the next paragraph we will explore what strategies and mechanisms enable living foraminifera to protect both the test and the cell from acidic stress.

## 4.2. Foraminiferal mechanisms to tolerate acidification

### 4.2.1. Foraminiferal behavioural skills

#### 4.2.1.1. Wearing armour (encystment or calcification).
Foraminifera can use their environment to respond to stress. During the incubations, specimens of *A. confertitesta* were often observed encysted with particles agglomerated by their pseudopodia net (Supplementary Figure 1). This observation was less frequent for *H. germanica*. This process has been commonly observed in previous experimental studies (Diz et al., 2012; Dupuy et al., 2010; Heinz et al., 2005; Maire et al., 2016). These cysts may create microenvironments where the chemical (carbonate concentration, calcite saturation state, pH) and food conditions can differ significantly from those of the culture seawater (Diz et al., 2012). Encystment could therefore be a behavioural adaptation in response to acidification and starvation. This is supported by the observations on *A. aomoriensis*, which demonstrated a greater sensitivity to acidification when specimens were directly exposed to acidified seawater compared to incubation in their natural sediment (Haynert et al., 2014). The presence of sediment therefore seems to reduce environmental stress by offering potential chemical microhabitats to support growth and development of benthic foraminifera. In our experiment, the development of a bacterial biofilm in the light/dark cycle conditions facilitated encystment of living specimens and could have contributed to the
acidification tolerance. This would explain the higher dissolution stages observed in the living *A. confertitesta* in the permanent darkness compared to the light/dark condition (Figure 2), where the biofilm was denser, and the cysts of agglomerated particles were relatively thicker.

Another hypothesis could be that living foraminifera are able to maintain their shell integrity by calcifying new layers of calcite during chamber addition events throughout incubation. Numerous studies have shown that hyaline foraminifera precipitate a thin calcite layer on the entire shell when chambers are added (Erez, 2003; Geerken et al., 2022; Jonkers et al., 2016; Nehrke et al., 2013; van Dijk et al., 2019). Calcification is also a period of high intracellular pH control for foraminifera (de Nooijer et al., 2009). It lasts from 4 to 7 hours (Geerken et al., 2022; Glas et al., 2012; Toyofuku et al., 2017) and may somehow buffer the low pH of the microenvironment around the shell. However, the outer shell of the foraminifer remains exposed to a low pH under acidic conditions for the rest of the time. However, chamber formation for Ammonia is strongly linked to feeding (Ernst et al., 2006; Goldstein and Moodley, 1993; Nomaki et al., 2008), while Haynesina germanica needs living diatoms (LeKieffre et al., 2018; Pillet et al., 2011; Schweizer et al., 2022). Since no food was offered to the foraminifera in our set-up, regular chamber formation events are unlikely to have occurred. Unfortunately, our experimental set-up does not allow the verification of the presence or absence of chamber addition event, so we cannot further explore this hypothesis.

### 4.2.1.2. Networking (pseudopodial net for motility or trophic strategy).
Foraminiferal pseudopodia are also involved to motility (Geslin et al., 2004; Goldstein et al., 1995; Moodley et al., 1998; Travis and Bowser, 1991). Motion strategies have already been identified in *A. tepida* and *H. germanica* (Deldicq et al., 2020; Guilhermic et al., 2023; Maire et al., 2016), and could contribute to stress tolerance by swerving local changes in sediment. These studies show no significant difference in motion between these two species. In our experiment, on day 12, *A. confertitesta* showed more extensive pseudopodial nets compared to *H. germanica* (Supplementary Figure 1). The fact that *H. germanica* was less active overall compared to *A. confertitesta* could be due to a specific functional trait. The emission of pseudopods is also associated with feeding. As *A. confertitesta* is heterotrophic (Chronopoulou et al., 2019; Haynert et al., 2020; Mojtahid et al., 2011; Pascal et al., 2008; Schweizer et al., 2022; Wukovits et al., 2018), it could be feeding on the biofilm. According to this hypothesis, the greater oxygen consumption during the light/dark cycle in acidic condition observed on day 12 (Figure 4), when the biofilm is the most developed, would be due to greater activity linked to consumption of the biofilm in a stressful environment (low pH and starvation). Under the same conditions, this was not observed for *H. germanica*, which has a mixotrophic strategy (photosynthetically active using kleptoplasts and feeding mainly on diatoms; Austin et al., 2005; Chronopoulou et al., 2019; Schweizer et al., 2022; Ward et al., 2003). *Haynesina germanica* therefore probably prefers to photosynthesise rather than explore its environment in search of food. Therefore, higher oxygen consumption rates in darkness suggest a more heterotrophic activity for *H. germanica* (Chronopoulou et al., 2019; Mojtahid et al., 2011; Pascal et al., 2008; Schweizer et al., 2022). No pseudopodial emissions were observed anymore on day 18 for *A. confertitesta*, which could be explained by a disturbance in the biological activity as observed after some time in previous experimental studies (Le Cadre et al., 2003; Stouff et al., 1999; Travis and Bowser, 1991) like dormancy (LeKieffre et al., 2017).

### 4.2.1.3. Waiting out the storm (reducing metabolism).
For *A. confertitesta*, we observed a progressive decrease in respiration throughout the incubation period, indicating a reduction of activity (Figure 4). It could be a sign of a state of dormancy, a common stress adaptation strategy in benthic foraminifera (LeKieffre et al., 2017; Ross and Hallock, 2016). This strategy protects the cell until environmental conditions become

favourable again and can, therefore, serve as a solution to temporary environmental changes. Higher survival rate in acidified conditions can suggest a slowing down of the cell, likely due to the absence of food. However, this passive strategy does not appear to be suitable for maintaining a test during an acidification event, as living benthic foraminifera specimens with fully dissolved tests have been observed in *situ* (Alve and Nagy, 1986; Cesbron et al., 2016; Charrieau et al., 2018a; Daviray et al., 2024). It has been shown that foraminifera can reconstruct their test after dissolution when environmental conditions become favourable (Bentov et al., 2009; Charrieau et al., 2022; Haynert et al., 2012; Kurtarkar et al., 2011; Le Cadre et al., 2003). This strategy may depend on the species because not all species have an inner organic lining that is thick or resistant enough to remain “naked” in the environment (Cohen and Guber, 1968; Tyszka et al., 2021). Our experiments did not reveal such difference, whilst most of the living specimens fully dissolved observed in *situ* belonged to the genus *Ammonia*. Incubated *Ammonia* spp. has shown thickening of the organic lining under anoxic conditions (Koho et al., 2018), or under high heavy metal concentrations (Le Cadre and Debenay, 2006). This was neither investigated previously nor observed in our experiment for *H. germanica*.

### 4.2.2. What about kleptoplast photosynthetic activity?

One of the biggest differences between *A. confertitesta* and *H. germanica* that we can further investigate in our study, is the ability of *H. germanica* to perform photosynthesis through kleptoplasty. The difference of respiration between both species is evident at day 2 and day 12 whilst at day 18, it is not so clear potentially due to potential dormancy state (Figure 4). The *H. germanica* kleptoplasts were still photosynthetically active on day 12 of experiments, and the estimates of the net photosynthesis ratio were of the same order of magnitude (low) as in the literature (Supplementary Table 3; Cesbron et al., 2017; Jauffrais et al., 2016; Lopez, 1979). We might expect that acidification, by increasing dissolved CO₂ concentration would enhance RuBisCO activity and positively impact photosynthetic activity. This contrasts with our observations: the net photosynthetic rate calculated on day 12 was not higher in acidified conditions compared to the control, and the photosynthetic activity was almost lost in acidified light/dark cycle. On day 18, net photosynthesis was negligible under all conditions indicating inactivity or even loss of the kleptoplasts, except for the control under light/dark cycle, where fluorescent images demonstrated the presence of kleptoplasts (Figure 3). Moreover, *H. germanica* respiration was higher in the dark acidified conditions (Figure 4), suggesting a combined stress effect of acidity and darkness on the biological activity of *H. germanica*. Jauffrais et al. (2016) demonstrated that kleptoplasts remained active for 11 to 21 days in permanent darkness, and for 9 to 12 days under low light conditions (25 µmol.photons·m⁻²·s⁻¹). In our study, kleptoplasts remained active until at least the 12th day with a similar light intensity for the light/dark cycle (~24 µmol photons·m⁻²·s⁻¹). However, this photosynthetic activity over a period of at least 12 days does not seem to have contributed to a significant advantage against the dissolution of *H. germanica* compared with the *A. confertitesta*. The progressive loss of photosynthetic activity by *H. germanica* in acidified conditions could be explained by the digestion of their kleptoplasts under stress and starving conditions (Jauffrais et al., 2016, 2019a). The disruption of photoprotection for the diatom kleptoplasts, such the xanthophyll cycle1 mechanism (Jauffrais et al., 2017; Lavaud et al., 2007), or the accumulation of detrimental reactive oxygen species like in diatoms (Kan et al., 2023; Waring et al., 2010), might be responsible for such loss.

Finally, considering the progressive loss of kleptoplast photosynthetic activity and the absence of a significant difference in the dissolution stage between the light/dark cycle and permanent darkness in acidified conditions at the end of the experiment (Figure 2), the photosynthetic activity of kleptoplasts does not appear to be involved in resistance to acidification-induced shell dissolution during long-term exposure to seawater with low pH. Previous studies have measured
the pH rise in the photosymbiotic foraminiferal microenvironment, to estimate the extent to which this could compensate for ocean acidification induced decreases in pH (Glas et al., 2012; Jørgensen et al., 1985; Köhler-Rink and Kühl, 2000, 2001; Rink et al., 1998). For example, in planktonic foraminifera, *Orbulina universa* showed a photosynthesis rate of 67.8 pmol O₂·s⁻¹·mm⁻³ (standardised at 16°C) at saturating irradiances inducing a 0.5 pH increase above ambient seawater of test surface (Rink et al., 1998), and *Globigerinoides sacculifer* a net photosynthesis about 47.6 pmol O₂·s⁻¹·mm⁻³ (standardised at 16°C) inducing a 0.4 pH raise around the test surface (Jørgensen et al., 1985). Large benthic foraminifera, *Marginopora vertebralis*, *Amphistegina lobifera* and *Amphisorus hemprichii*, showed a photosynthesis rate standardised at 16°C of 6.5, 2.1 and 1.9 pmol O₂·s⁻¹·mm⁻³ and an associated pH raise of 0.3, 0.1 and 0.1 respectively (Köhler-Rink and Kühl, 2000, 2001). Here, *H. germanica* net photosynthesis was about 2.0 pmol O₂·s⁻¹·mm⁻³ (700 µmol photons·m⁻²·s⁻¹) being 24 to 34 times lower than photosymbiotic planktonic foraminifera, and in the same order of magnitude as photosymbiotic large benthic foraminifera. A pH increase in the foraminiferal microenvironment of *H. germanica* test due to photosynthesis in these experimental conditions (low irradiance about 24 µmol photons·m⁻²·s⁻¹) could be negligible, which might explain why we do not observe a higher tolerance of *H. germanica* compared to *A. confertitesta*. A pH monitoring experiment close to the test by exposing *H. germanica* to darkness and saturating irradiance for kleptoplasts could confirm whether *H. germanica* can significantly modify the pH of its microenvironment or not.

To sum up, from day 2 to day 12, the effect of the parameters on living specimens of both species are quite visible through encystment, pseudopodial net emission, or O₂ consumption. For *A. confertitesta*, in controlled conditions, O₂ consumption rates decrease probably due to a lack of food, whilst in acidified conditions O₂ consumption rates increase, either because of acid stress or as a result of biofilm grazing in starving condition. Encystment and pseudopodial emission are both behavioural responses to these hypotheses. For *H. germanica*, when it is mixotrophic (in light/dark cycle conditions), test dissolution appears stronger than in *A. confertitesta*, mostly due to the specific test structure. In addition, acidification harms the kleptoplasts resulting in a reduction in the net photosynthesis rate. When *H. germanica* is heterotrophic (in darkness), O₂ consumption is the most elevated in acidified condition, likely as a result of acid stress. On day 18 (end of incubation), O₂ consumption rates were lower than on day 12, suggesting dormancy. In summary, both species showed similar degrees of dissolution after incubation in acidified conditions regardless of light exposure conditions. We have identified multiple mechanisms that may partly explain the tolerance of estuarine foraminifera to acidification. However, the effect of acidification on the cellular metabolism of foraminifera and the (species-) specific metabolic pathways involved in resistance to acidification remain to be identified. To this end, it could be interesting to carry out comparative transcriptome analyses of foraminifera incubated under control and acidification conditions.

## 5. Conclusion

By comparing the dissolution of foraminifera tests of living and dead specimens incubated in an acidification experiment, our study demonstrates that living benthic foraminifera have active mechanism(s) to tolerate environment acidification. We explored various mechanisms for *A. confertitesta* and *H. germanica* to counter acidification-induced shell dissolution including activity of kleptoplasty and the differences in test morphology. The ability and success to mitigate or even negate dissolution are species-specific and efficient for short-term protection of the

test. In general, the tests of *H. germanica* specimens appear more sensitive to acidification, likely due to the thinner test thickness compared to *A. confertitesta* and (the lack of) certain behaviour skills (e.g., particle encystment). The photosynthetic activity of the kleptoplasts of *H. germanica* does not seem to be a strategy in long-term tolerance to acidification. To summarise, our study highlights that resistance mechanisms are species-specific and should therefore be further investigated to understand the impact of acidification process on benthic ecosystem functioning and the test preservation in fossil record.

## CRediT authorship contribution statement

**Maxime Daviray:** Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization. **Milla Lesut:** Writing – review & editing, Writing – original draft, Visualization, Resources, Investigation, Formal analysis, Data curation. **Emmanuelle Geslin:** Writing – review & editing, Visualization, Validation, Supervision, Funding acquisition, Conceptualization. **Edouard Metzger:** Writing – review & editing, Visualization, Validation, Supervision, Funding acquisition, Conceptualization. **Inge van Dijk:** Writing – review & editing, Visualization, Validation, Supervision, Funding acquisition, Conceptualization. **Julia Courtial:** Writing – review & editing, Writing – original draft, Visualization, Validation, Supervision, Resources, Project administration, Methodology, Investigation, Funding acquisition, Formal analysis, Data curation, Conceptualization.

## Author statement

We declare that this manuscript is original, has not been published before and is not currently being considered for publication elsewhere.

We confirm that the manuscript has been read and approved by all named authors and that there are no other persons who satisfied the criteria for authorship but are not listed. We further confirm that the order of authors listed in the manuscript has been approved by all of us.

We understand that the Corresponding Author is the contact for the Editorial process. He is responsible for communicating with the other authors about progress, submissions of revisions and final approval of proofs.

## Funding

The authors received funding from the OSUNA (internal program, project AKA), and from the University of Angers, France.

## Declaration of competing interest

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:

Julia Courtial reports financial support was provided by Nantes Atlantic Observatory of the Sciences of the Universe. If there are other authors, they declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

## Acknowledgements

We thank Chistine BARRAS, Meryem MOJTAHID, and Aubin THIBAULT DE CHANVALON for their advice on the experimental design and improvement of the set-up.
## Appendix A. Appendix. Taxonomy of the foraminiferal species cited in the manuscript

<table><thead><tr><th>Ecology</th><th>Species name</th></tr></thead><tbody><tr><td rowspan="13">Benthic</td><td>Ammonia aberdoveyensis Haynes, 1973</td></tr><tr><td>Ammonia confertitesta Zheng, 1978</td></tr><tr><td>Ammonia tepida morphogroup (Cushman, 1926)</td></tr><tr><td>Ammonia beccarii (Linnaeus, 1758) as Ammonia aomoriensis (Asano, 1951) †</td></tr><tr><td>Amphisorus hemprichii Ehrenberg, 1839</td></tr><tr><td>Amphistegina gibbosa d'Orbigny, 1839</td></tr><tr><td>Amphistegina lobifera Larsen, 1976</td></tr><tr><td>Bolivina argentea Cushman, 1926</td></tr><tr><td>Bulimina marginata d'Orbigny, 1926</td></tr><tr><td>Cassidulina laevigata d'Orbigny, 1826</td></tr><tr><td>Elphidium crispum (Linnaeus, 1758)</td></tr><tr><td>Elphidium williamsoni Haynes, 1973</td></tr><tr><td>Haynesina germanica (Ehrenberg, 1840)</td></tr><tr><td>Marginopora vertebralis Quoy &amp; Gaimard, 1830</td></tr><tr><td>Quinqueloculina seminula (Linnaeus, 1758)</td></tr><tr><td>Rosalina globularis d'Orbigny, 1826</td></tr></tbody></table>

Planktonic *Trilobatus sacculifer* as *Globigerinoides sacculifer* (Brady, 1877)

*Orbulina universa* d'Orbigny, 1839

## Appendix B. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.marmicro.2025.102490.

### Data availability

All the data are published within this paper and in the Supplement. The raw data used to make the figures are available on request.

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