Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

Depth related adaptations in symbiont bearing benthic foraminifera: New insights from a field experiment on Operculina ammonoides

An Author Correction to this article was published on 19 July 2018

This article has been updated

Abstract

Large benthic foraminifera (LBF) are marine calcifying protists that commonly harbor algae as symbionts. These organisms are major calcium carbonate producers and important contributors to primary production in the photic zones. Light is one of the main known factors limiting their distribution, and species of this group developed specific mechanisms that allow them to occupy different habitats across the light gradient. Operculina ammonoides (Gronovius, 1781) is a planispiral LBF that has two main shell morphotypes, thick involute and flat evolute. Earlier studies suggested morphologic changes with variation in water depth and presumably light. In this study, specimens of the two morphotypes were placed in the laboratory under artificial low light and near the sea floor at depths of 15 m, 30 m, and 45 m in the Gulf of Aqaba-Eilat for 23 days. Differences in growth and symbionts content were evaluated using weight, size, and chlorophyll a. Our results show that O. ammonoides exhibit morphological plasticity when constructing thinner chambers after relocation to low light conditions, and adding more weight per area after relocation to high light conditions. In addition, O. ammonoides exhibited chlorophyll content adaptation to a certain range of light conditions, and evolute specimens that were acclimatized to very low light did not survive relocation to a high light environment, possibly due to photo-oxidative stress.

Introduction

Symbiont-bearing larger benthic foraminifera (LBF) generally live in shallow oligotrophic waters of tropical and sub-tropical seas. They mostly harbor endosymbiotic algae and have complicated internal structures1,2,3. They usually exceed 3 mm3 in volume1, and are an important component of modern low latitudes and ancient oceanic ecosystems. These organisms are major CaCO3 producers, but their calcification is a complex physiological process that is biologically controlled and not well understood. Algal symbiosis is prevalent in many LBF species and hence they are considered as major contributors to primary production in the photic zones of shallow seas4,5,6,7,8. Studies investigating the biology, ecology, depth distribution, morphology and symbiotic relationship with algae in LBF are essential for monitoring the ecological state of tropical ecosystems, for interpretation of their fossil assemblages and for understanding the impact of future climate and oceanographic changes on marine calcifying organisms.

Symbiont-bearing larger benthic foraminifera distribution in shallow marine environments is determined by a set of parameters, of which water depth has a dominant but indirect role5,9,10,11,12,13. Light intensity decreases exponentially with depth because of water absorption and turbidity which are influenced by terrestrial influx and nutrients content8. The characteristics of sea water, dissolved materials and suspended particles affect the wave length of the available light in the benthic habitat. It has been suggested that the type of symbiont associated with specific LBF taxa influences foraminiferal depth distribution14, and that modifications in the shell shapes may be an adaptation for hosting symbionts15,16,17.

Operculina ammonoides (Gronovius, 1781) is a tropical hyaline diatom-bearing LBF, which is found in a wide range of depths5,16,18,19,20. It is closely related to the genus Nummulites (within the same sub-family21), which were widespread throughout the Paleogene (sub) tropics to the extent that they are the principal component of some shallow-water carbonates22. The shell of O. ammonoides is planispiral and exhibits two main morphotypes with various intermediate forms: thick involute morphotype (Fig. 1.1), in which the chamber outer layers cover laterally those of the preceding coil, and flat evolute morphotype (Fig. 1.2), in which the chamber outer layers in a coil do not laterally cover those of the preceding coil. Quite often intermediate semi-involute morphotypes are also found (Fig. 1.3). Previous studies in the Gulf of Aqaba-Eilat described this species as occupying depth interval of 30–150 m, and being very common at water depths greater than 40 m on sandy substrates5,16, with numerical abundance increase of up to 63% in the dead assemblage from 60–90 m23. Few studies indicated that specimens found shallower than 40–60 m are mostly more involute and thicker than specimens from deeper water5,16.

Figure 1
figure1

Variability in the mode of coiling in Operculina ammonoides. (1) Involute. (2) Evolute. (3) Semi-involute (Advolute/Convolute). (A) Side view. (B) Apertural view.

Our own observations show that this species also thrives at depths of 15–30 m at the Northern Gulf where it is found on the sandy and silty sediments and around the sea grass Halophila stipulacea. In this habitat, it appears in very high densities of 5–10 live individuals per cm2, and is a significant component of the dead assemblages20.

Previous studies suggest that O. ammonoides shell changes from involute and semi-involute to evolute with increase in water depth due to lower irradiance and/or current energy5,9,16,19,24. The strong linkage between shell morphology and depth is often assigned to irradiance levels that seem to mainly affect the functionality of the diatom symbionts. However, Oron et al.20 documented variability in the mode of coiling in O. ammonoides population sampled at 27 m water depth after removal of fish farms from the northern Gulf of Aqaba-Eilat in 2008. This was probably an ecophenotypic response which might be typical to this species in impacted, high turbidity environments, but the origin of this phenomenon and its possible linkage to light levels is unknown.

It was previously demonstrated that changes of morphological characters within Operculina and Planoperculina species can be used for depth gradient estimation using regression analyses25. Such methods can be used for high resolution paleodepth, and possibly water turbidity estimation. Studies of living forms can be combined with the rich nummulitids fossil record to better understand growth history and biostratigraphy implications.

Another depth related hypothesis derives from the fact that some groups of foraminifera exhibit sexual dimorphism. This dimorphism involves an asexual generation with a large proloculus (initial chamber) and small test diameter (megalospheric), and a sexual generation with a small proloculus and large test diameter (microspheric). A possible link to dimorphism was suggested based on an increase in proloculus size with depth in O. ammonoides from the Red Sea26. However, this observation could have been the result of the presence of two types of megalospheric forms, as documented in Heterostegina depressa from Hawaii27.

In this study, we combined laboratory and unique in situ experiments to investigate physiological responses of O. ammonoides to depth changes. Our results further develop the perception that the actual depth distribution is a compromise between different habitat requirements, and that the foraminifera shell shape must accommodate the needs of the symbionts when living outside their optimum light range. This study also provides important insights on the functional morphology of large benthic foraminifera, and has a direct implication for interpretations of the paleoecology and taxonomy of fossil nummulitids.

Experimental Design and Methods

Collection of specimens

For involute specimens, sediments were collected by SCUBA diving from soft bottom sites located at the North Beach of the Gulf of Aqaba-Eilat, at 20 m water depth, where O. ammonoides is abundant mostly in its involute and slightly semi-involute forms. The sediments were sieved between 1000 µm and 1500 µm and placed in glass beakers with seawater for collection of climbing individuals.

Evolute specimens were collected from culturing containers in the laboratory containing sediments that were brought by technical Closed-Circuit Rebreathing (CCR) diving from 45 m water depth in the coral reef area off the Inter-University Institute for Marine Sciences in Eilat (IUI). The evolute specimens that were collected for this study were part of a newly formed gamont generation, produced by asexual reproduction of the original population. The laboratory cultures were maintained under low light conditions of ~10 μmol photons m−2 s−1 supplied by a white Light-Emitting Diode (LED) lamp and at 24–25 °C, the average annual temperature in the northern Gulf of Aqaba-Eilat.

Experimental setup

After the collection of live individuals, 110 involute (and slightly semi-involute) and 55 evolute adult specimens were divided into 11 groups of 15 specimens (10 involute, and 5 evolute) and cleaned under the binocular microscope using brushes. All specimens were photographed, and two groups were frozen for later analysis. The nine groups intended for translocation were labeled with the fluorescent probe calcein to indicate the addition of new chambers by cultivating them in filtered seawater with calcein 20 μM for two days. The experimental groups were placed in transparent, perforated (~1 mm) 10 mL plastic tubes. The tubes were attached to small weights and placed near the sea floor at depths of 15 m (three groups), 30 m (three groups) and 45 m (two groups) off the coast of the IUI marine laboratory. One group (“80” m) was placed in the laboratory under low light conditions (~10 μmol photons m−2 s−1) approximating the maximum light levels at ~80 m off the marine laboratory, and was cultured inside the same plastic tubes as the in situ groups in a seawater aquarium in 24–25 °C. The water in the aquarium was changed every 4–5 days with fresh unfiltered sea water collected off the IUI marine laboratory pier. All experimental tubes were cleaned after 11 days, and retrieved after 23 days.

Additional 10 thick involute specimens, that were left after the assembly of the experiment groups, were calcein labeled for 6 days and cultivated in the low light environment of the laboratory for a longer period (43 days) for growth observation purposes.

In addition, 38 involute and 39 evolute dead specimens (empty shells) were collected from the same sediments and culturing containers as the live individuals for establishing a weight to area relationship chart on representative “standard shaped” specimens of both morphotypes.

Temperature and light at the study sites

During the experiment (May–June 2013), HOBO® Pendant Temperature/Light Data loggers were attached to the plastic tubes recording light levels in Lux units (lumen m−2) and temperature. Logging was done at 10 min intervals throughout the experiment. Photosynthetically Active Radiation (PAR), in units of μmol photons m−2 s−1, was calculated using available surface PAR data logged at the meteorological station on the marine laboratory pier (http://www.iui-eilat.ac.il/Research/NMPMeteoData.aspx), combined with typical light attenuation for the month of May in the coral reef area off the IUI marine laboratory28. The maximum daily PAR calculated for depths of 15 m, 30 m and 45 m was 566 μmol photons m−2 s−1, 222 μmol photons m−2 s−1 and 81 μmol photons m−2 s−1 respectively (Fig. 2). In situ temperatures fluctuated between 23.1 °C and 25.5 °C at 15 m, 23.3 °C and 25.1 °C at 30 m and 23 °C and 24.8 °C at 45 m (Fig. 3). The collection site of the involute specimens in the North Beach is characterized by higher turbidity, and the seasonal maximum light levels in 20 m are ~180 μmol photons m−2 s−1, a lower light level than the same depth off the marine laboratory in the south beach (Tamir, unpublished data). However, the involute specimens that were collected there spent ~5 days in the low light environment of the laboratory before the beginning of the experiment.

Figure 2
figure2

Typical daily light profile (a) and maximum light levels (b) during the experiment for the in situ sites at depths of 15 m, 30 m and 45 m, off the IUI Marine Laboratory (May-June 2013). PAR = Photosynthetically Active Radiation.

Figure 3
figure3

Temperature fluctuations throughout the in situ experiment at depths of 15 m, 30 m and 45 m in the experimental sites off the IUI Marine Laboratory (May-June 2013). Color lines are 20 period moving average.

Analytical techniques

Digital and x-ray imaging

Digital images of all specimens were taken before and after the experiment for measurements of shell growth. Normal light photography was done using a Leica stereomicroscope with fitting digital camera. The photographs after the experiment were taken under very low light using high camera sensor sensitivity (ISO) function, to minimally affect the symbionts. Epifluorescent photography was done using a Leica stereomicroscope with a Leica DFC310 FX camera. X-ray computed microtomography on 10 representative specimens was done with Skyscan 1173 Desktop-Micro-CT scanner (Bruker) at the Institute of Paleontology, University of Vienna.

Size/weight measurements and growth calculations

Area measurements were done with Photoshop analysis measurement tools. Area increase was calculated for each individual using the start and end overhead view area.

Weight increase was estimated for each individual using the start calculated weight based on measured area and the weight vs area charts established from empty shells, and the end weight measured after the experiment. Broken specimens were excluded. At the end of the experiment it became clear that the two days cultivation in calcein was not sufficient to label all individuals, so all growth calculations were done using the methods mentioned above.

Chlorophyll measurements

At the end of the experimental period, specimens were frozen in −18 °C and stored in the dark until analyzed for chlorophyll a. Randomly selected specimens from each group were placed in a glass vial containing 2 mL of acetone (90%) in 4 °C in the dark. After 24 h, the extracted chlorophyll was measured using a Chl a Acidification Fluorescent Module in a Turner Designs Trilogy Laboratory Fluorimeter by following EPA Method 445.029. Most measurements were done on a single specimen, to inspect the variability within the population. Specimens used for chlorophyll analysis were later treated in 5% sodium hypochlorite (NaOCl) for 12 h for removing organic matter, and the remaining calcium carbonate was weight to normalize chlorophyll values.

Statistical analysis

Most datasets were found to have non-normal distribution after using Shapiro-Wilk test and histograms observations (Supplementary Table S1), so the Mann-Whitney nonparametric test was used for comparing means. Shapiro-Wilk test and Mann-Whitney test were performed with IBM SPSS Statistics 22 (IBM Corp., Armonk, USA).

Data availability

All data generated or analyzed during this study are included in this article and its Supplementary Information files.

Results

Growth and morphology

The two main morphotypes have a distinctively different weight-to-area relationship (Fig. 4). Namely, envolute specimens are distinctly larger in area per unit of weight (by a factor of ~2). Computed Tomography (CT) scan images of selected specimens of different morphologies from the collection sites all showed megalospheric proloculus with average diameter of 59(±14) µm (Fig. 5), which indicates that the origin of those specimens is by asexual reproduction.

Figure 4
figure4

Area and weight relationships in adult involute and evolute specimens of O. ammonoides. Based on measurements of 38 involute and 39 evolute dead specimens (empty shells) collected from the same sediments and culturing containers as the live individuals used in this experiment.

Figure 5
figure5

Computed Tomography scan images of evolute (a), involute (b) and semi-involute (c) O. ammonoides specimens all showing megalospheric proloculus (large and visible initial chamber). The evolute specimens are from the laboratory and the involute specimens were collected in the field (North Beach 15–20 m).

Growth in both morphotypes showed high variability between individuals. Area increase for involutes was statistically lower in 15 m (p < 0.050, Supplementary Table S2), where average growth per individual was 10%(±5%) compared with 17%(±9%) in the deeper groups (Fig. 6a). All evolute specimens from 15 m were bleached and dead upon retrieval (no pseudopodia activity and no color), and growth in 30 m, 45 m and “80” m was statistically similar (Supplementary Table S2) with an average of 19%(±10%) per individual (Fig. 6b).

Figure 6
figure6

Area increase per individual in “80” m, 45 m, 30 m and 15 m after 23 days of in situ deployment on the sea floor. In both Involute (a) and Evolute (b) specimens, the difference between the 15 m group and the other groups is statistically significant (p < 0.050, Supplementary Table S2). Number of analyzed specimens (n) from left to right for Involute: 10, 20, 30, 30 and Evolute: 5, 10, 15, 15. Box-plot elements: whiskers = most extreme values, middle line = median, upper and lower lines = quartiles, +  = Average. Average area increase in percentage per individual is presented next to the box.

Estimated weight addition for involutes was statistically higher in 15 m and 30 m compared with 45 m and “80” m (p < 0.002, Supplementary Table S3), with an average of 24%(±16%) per individual in the shallower groups and 50%(±23%) per individual in the deeper groups (Fig. 7a). Estimated weight addition for evolutes was statistically similar (Supplementary Table S3) for all surviving groups, with an average of 62%(±27%) per individual (Fig. 7b).

Figure 7
figure7

Estimated weight increase per individual in “80” m, 45 m, 30 m and 15 m after 23 days of in situ deployment on the sea floor. In Involute (a) specimens the difference between the high and intermediate light groups (15 m and 30 m) and the lower light groups (45 and “80” m) is statistically significant (p < 0.002, Supplementary Table S3). In Evolute (b) specimens the 15 m group did not show detectible weight increase, and all other groups are statistically similar (Supplementary Table S3). Number of analyzed specimens (n) from left to right for Involute: 10, 18, 30, 25 and Evolute: 4, 9, 14, 12. Box-plot elements: whiskers = most extreme values, middle line = median, upper and lower lines = quartiles, +  = Average. Average area increase in percentage per individual is presented next to the box.

When comparing area vs weight of the individuals after the experiment with the “standard shape” trendlines from Fig. 4, most individuals from 15 m and 30 m appear under the trendline, meaning they are heavier compared to the “standard shape” population of the same area or diameter (Fig. 8).

Figure 8
figure8

Area vs. weight of all individuals after the experiment and the “standard shape” trendlines from Fig. 4. Most individuals from 15 m and 30 m are heavier compared to the control population of the same area.

Involute specimens that formed more than ~3 new chambers in the deep in situ groups, visibly show thinner chambers which are more consistent in shape and coiling mode with evolute forms (Fig. 9a). The latter observation is even more visible in the additional 10 thick involute specimens that were cultivated in the “80” m conditions for 43 days (Fig. 9b). Abnormal chamber formation was also observed, mostly in evolute specimens, forming chambers with shorter septal distance, shorter chamber height or longer backbend angle (Fig. 9c).

Figure 9
figure9

Growth and morphology changes during the experiment. Involute specimens with “evolute-like” thinner chambers after relocation to 45 m for 23 days (a), and to “80” m (laboratory) for 43 days (b, calcein-labeled), and irregular chamber formation in evolute forms (c). The arrows indicate the beginning of the experiment.

Chlorophyll

In both involute and evolute morphotypes chlorophyll a content in “80” m and 45 m was statistically higher (by a factor of ~3–4) than in 30 m and 15 m (p < 0.002, Supplementary Table S4), excluding the evolute individuals from 15 m that were bleached and dead. Average Chlorophyll a values for involutes were 0.13(±0.03) µg mg−1 in the deeper groups and 0.04(±0.02) µg mg−1 in the shallower groups (Fig. 10a), and average Chlorophyll a values for evolutes was 0.14±0.03 µg mg−1 in the deeper groups and 0.03±0.02 µg mg−1 in the 30 m group (Fig. 10b). Chlorophyll a content negatively correlates to average maximum daily light at the different depths (Fig. 10).

Figure 10
figure10

Chlorophyll a content per mg in “80” m, 45 m, 30 m and 15 m. In both involute (a) and evolute (b) specimens the difference between the low light conditions (“80” m and 45 m) and the intermediate/high light conditions (30 m and 15 m for involute and 30 m for evolute) is statistically significant (p < 0.002, Supplementary Table S4). All evolute specimens at 15 m were bleached and dead upon retrieval. Number of analyzed specimens (n) from left to right for Involute: 10, 8, 8, 8 and Evolute: 6, 8, 8, 8. Box-plot elements: whiskers = most extreme values, middle line = median, upper and lower lines = quartiles, + = Average. PAR = Photosynthetically Active Radiation.

Discussion

Interactions between environmental factors often make it difficult to correlate ecomorphology of LBF with depth distribution and other variables. Haynes15 suggested that shell shape is a compromise between available light, metabolic requirements associated with algal symbiosis and hydrodynamic factors. Many early studies have documented LBF morphology changes with variation in habitat depth, light levels and water motion, and the authors mostly attributed the changes to the association with symbionts. Specifically, for some key species of diatom-bearing LBF (Amphistegina, Heterostegina and Operculina), it has been shown that thicker tests are found in shallow water5,6,19,30,31,32,33 and when cultivated under high light levels32,34. In addition, Amphistegina spp. are thicker in habitats that are exposed to wave action, and produce thicker shells when subjected to water motion during growth in the laboratory32,34,35. Sectioned tests revealed that this feature is a direct result of differences in secondary lamellar thicknesses33,34.

For O. ammonoides specifically, where involute specimens are distinctively heavier compared to evolute specimens of the same size (Fig. 4), the thickening of the test in shallow water appear to be related to changes in the mode of coiling, and the lateral surface of the chamber walls is greater in deeper water5,19,30. It was also concluded that all O. ammonoides ecotypes belong to the same species based on biometric measurements of specimens from different depths, and megalospheric forms were observed among involute and evolute specimens5. Computed Tomography scan images of selected specimens of different morphologies from our collections all showed megalospheric forms (Fig. 5). Therefore, the morphological variability is not related to the sexual or asexual origin of the individuals.

As for growth rates in diatom-bearing LBF, previous work on H. depressa showed optimum growth rates in very low light levels of 300 “daylight neon” Lux (~5 μmol photons m−2 s−1)36,37, and in 600 Lux (~10 μmol photons m−2 s−1)38. Erez39 described a mid-water (25 m) growth and photosynthesis optimum in the diatom-bearing A. lobifera based on in situ experiments, and excluded the simple notion that growth rates decrease with depth as a result of reduction in photosynthetic activity. Furthermore, the 25 m optimum contradicts the reports describing the preferred habitat of A. lobifera (<10 m), implying that the actual depth distribution is a compromise between different habitat requirements, and that the foraminiferal test shape must accommodate the needs of the symbionts when living outside their optimum light range.

In this study, O. ammonoides translocated thick involute specimens showed higher area growth in the lower light conditions of the “80” m, 45 m and 30 m compared to 15 m (Fig. 6a), but higher weight addition in 15 m and 30 m (Fig. 7a). The temperature in the laboratory (24–25 °C) was slightly higher and more stable than in the deep in situ sites (Fig. 3), there was no continuous water motion and the irradiance was constant. However, those parameters did not have a significant effect on survival and growth. Therefore, it is reasonable to assume that the main parameter favoring area growth in this experiment was relatively low light levels of a wide range (~10–222 μmol photons m−2 s−1).

Specimens exhibited morphological plasticity when individuals from 15 m and 30 m became heavier compared to the control population of the same size (Fig. 8), and involute specimens that were relocated to the low light level environments formed new thinner chambers, which were consistent in shape with the evolute forms (Fig. 9a,b). The latter observation agrees with the light-controlled morphology hypothesis described for O. ammonoides, and also indicates that individuals can adapt to new conditions, and balance the thickness and surface area of the test depending on the incoming light.

Another observation we report here is abnormal growth of many of the newly formed chambers during the experiment. The abnormalities include deformed smaller chambers and chambers with shorter septal distance, shorter chamber height or longer chamber backbend angle. The abnormalities are mostly visible in the evolute forms from the in situ groups (Fig. 9c), but appear to some extant in all experiment groups, including the laboratory grown ones (Supplementary Figs S1S9). Previous explanations for deformity in foraminifera include mostly natural environmental stressors and pollution40,41,42 or regeneration after damage43. However, in our experiment, we cannot account for those variables. The common parameter for almost all groups is the relocation and abrupted change in conditions, that may have caused disruption of the cytoskeleton.

Corals that are adapted to low light and were relocated to shallow water usually show high mortality rates44,45,46,47 and/or bleaching48,49. Similarly, LBF have suffered documented bleaching incidents in recent decades. Cases of LBF field populations bleaching are documented in four species of the diatom-bearing Amphistegina (reviewed by Hallock et al.50). Negative response of LBF to high light levels in experiments in the laboratory and in situ was also documented in some cases: A. lobifera, a shallow dwelling (>10 m) species, showed a mid-water (25 m) optimum for growth and photosynthesis39 when cultivated in jars in situ and unable to interact with the substrate (i.e. cryptic behavior). In the laboratory, light levels above A. lobifera optimal range induced oxidative stress and bleaching and caused significant reduction in survivorship51. Amphistegina lessonii, typically a deeper-dwelling species (20–40 m), was photoinhibited in full sunlight. Heterostegina depressa, a diatom-bearing Nummulitidae like O. ammonoides, has reduced growth rates and maximum quantum yields in a very high light regime (with midday peaks at 1200 μmol photons m−2 s−1)52, and showed signs of stress even under illumination of 100 μmol photons m−2 s−1 53. A key difference between the known cases of bleaching in corals and bleaching in LBF is that events of bleaching in corals are mostly correlated with elevated temperatures45, while bleaching in LBF is associated with photoinhibitory stress54,55,56. However, temperature-induced bleaching was reported in three species of diatom-bearing LBF57, and it was shown that more specimens of Amphistegina gibbosa were partially bleached under high light intensities in 32 °C compared with 20 °C and 25 °C55. A current prevailing model for bleaching propose a primary trigger of light-dependent generation of reactive oxygen species by heat-damaged chloroplasts58,59, therefore while bleaching in LBF is linked mostly to light levels, other factors such as temperature, or a combination of factors, may also induce bleaching.

In this study, the evolute specimens that were adapted to low light in the laboratory did not survive the relocation to 15 m (max light levels of 566 μmol photons m−2 s−1). All other groups in this experiment at all depths showed 100% survival. The low-light adapted evolute specimens possibly experienced photo-oxidative stress, as described in corals and in some species of foraminifera. Unlike the relocated evolute specimens, the involute ones could sustain the higher light level at 15 m probably due to their thinker morphology that is more protective of the symbionts since they were collected in the higher (compared to the laboratory) light environment of the North Beach.

Diatom-bearing LBF are a diverse group, and the diversity of diatoms in symbiosis with foraminifera is comparably high. This diversity may be the reason for the wide ecological range of depth and light habitats of this group and their high abundances in those habitats52, and photosynthetic plasticity may be the tribute that allows some species to acclimatize rapidly to different light conditions53. Over the years, O. ammonoides have been suggested to host various species of diatoms belonging to the genera Amphore, Achnanthes, Nitzschia60,61 and Thalassionema62. It has long been known that diatoms can survive at very low light levels and can tolerate relatively high light levels for limited periods of time63. Furthermore, improved growth under low irradiances (<11 μmol photons m−2 s−1) was reported for many diatom species64,65,66, including species that were found in large quantities in O. ammonoides60. The above-mentioned experiments used free living or isolated diatoms, and it should be emphasized here that our study highlights the fact that the hosts test also play a role in controlling and modulating the light received by the endosymbiotic algae. The structure and test shape of the host may have evolved in response to the requirements of the symbionts, and the test shape and thickness balance the surface to volume ratios depending on the incoming light31,33,67. In addition, our study shows that light quantity and quality adaptation mechanism in the holobiont level can be also achieved by increasing the biomass and number of symbionts, or by increasing cellular chlorophyll a content of the symbionts (Fig. 10), as documented in corals and their zooxanthellae68,69,70,71,72,73.

Interestingly, although growth rates did not differ significantly between the low light environments of the laboratory (approximating maximum light at ~80 m), 30 m and 45 m (Fig. 6), the chlorophyll content adaptation seems to have a threshold somewhere between 30 m and 45 m. Both ecotypes show similar chlorophyll content in the “80” m and 45 m, and much lower values in 30 m (Fig. 10). This phenomenon cannot be explained solely by the exponential nature of light attenuation, and might be related to the different components of the light spectrum. The laboratory artificial light, approximating max PAR at ~80 m, was provided by a cold white LED lamp. Cold white LED typically peaks at ~450 nm (blue light), which is similar to the light components reaching the water column below ~30 m in our in situ site, where no light above 600 nm reach 40 m, creating an irradiance field dominated by blue light74. During the last few decades researchers documented specific effects of different light components on the photosynthetic rates and pigment content of various species of marine algae. It is known that the spectral composition of light plays a crucial role in growth rate, photoprotective mechanisms and pigment content in diatoms75. Some species of diatoms grown in blue and blue-green light had distinctively more chlorophyll compared to full spectrum white light of the same intensity.

As for the holobiont level, in the LBF A. gibbosa, higher-energy blue light induced more bleaching than lower-energy white light, but growth rates (in diameter) were higher under blue light. The higher growth rates were explained by the exposure to 20% more useable energy56. Calcification in high light (full spectrum) adapted corals was mostly enhanced after transfer to blue light, but photosynthesis was observed to be significantly less efficient74,76,77. It was suggested that the corals from shallow water are adapted to full light spectra whereas deep corals are adapted to blue light spectra which require more pigments, and that blue light photoreceptors in the coral tissue might be the link between light absorption by the coral host and activation of biological processes that enhance calcification77. Similar mechanisms probably exist in LBF, and chromatic adaptation provide selective advantages by maximizing photosynthetic activity under different spectral conditions.

Finally, it should be noted that light enhanced calcification (LEC) was shown to be independent of symbiont photosynthesis both in planktonic and benthic foraminifera using a photosynthetic inhibitor78,79. More recently it was shown that LEC in corals could proceed almost without photosynthesis of the symbionts under dark blue light77. The implications of these observations for O. ammonoides will need to be evaluated in future work.

Change history

  • 19 July 2018

    A correction to this article has been published and is linked from the HTML and PDF versions of this paper. The error has not been fixed in the paper.

References

  1. 1.

    Ross, C. A. Evolutionary and ecological significance of large calcareous Foraminiferida (Protozoa), Great Barrier Reef (Proceedings of the Second International Coral Reef Symposium Ser. 1, Great Barrier Reef Committee, Brisbane, 1974).

  2. 2.

    Hallock, P. Why are larger foraminifera large? Paleobiology 11, 195–208 (1985).

    Article  Google Scholar 

  3. 3.

    BouDagher-Fadel, M. K. in Evolution and geological significance of larger benthic foraminifera 544 (Elsevier, Amsterdam, 2008).

  4. 4.

    Lee, J. J. & Hallock, P. Algal Symbiosis as the Driving Force in the Evolution of Larger Foraminiferaa. Ann N Y Acad Sci 503, 330–347 (1987).

    ADS  Article  Google Scholar 

  5. 5.

    Reiss, Z. & Hottinger, L. in The Gulf of Aqaba: Ecological micropaleontology. (Springer-Verlag, Berlin; New York, 1984).

  6. 6.

    Hansen, H. J. & Buchardt, B. Depth distribution of Amphistegina in the Gulf of Elat, Israel. Utrecht Micropaleont Bull 15, 205–224 (1977).

    Google Scholar 

  7. 7.

    Langer, M. R., Silk, M. T. & Lipps, J. H. Global ocean carbonate and carbon dioxide production; the role of reef Foraminifera. J Foraminiferal Res 27, 271–277 (1997).

    Article  Google Scholar 

  8. 8.

    Hallock, P. In Modern Foraminifera (ed Sen Gupta, K. B.) 123-124 (Springer, 2002).

  9. 9.

    Hohenegger, J., Yordanova, E., Nakano, Y. & Tatzreiter, F. Habitats of larger foraminifera on the upper reef slope of Sesoko Island, Okinawa, Japan. Mar Micropaleontol 36, 109–168 (1999).

    ADS  Article  Google Scholar 

  10. 10.

    Hohenegger, J., Yordanova, E. & Hatta, A. Remarks on West Pacific Nummulitidae (Foraminifera). J Foraminiferal Res 30, 3–28 (2000).

    Article  Google Scholar 

  11. 11.

    Renema, W. & Troelstra, S. R. Larger foraminifera distribution on a mesotrophic carbonate shelf in SW Sulawesi (Indonesia). Palaeogeogr Palaeoclimatol Palaeoecol 175, 125–146 (2001).

    Article  Google Scholar 

  12. 12.

    Beavington-Penney, S. J. & Racey, A. Ecology of extant nummulitids and other larger benthic foraminifera: applications in palaeoenvironmental analysis. Earth Sci Rev 67, 219–265 (2004).

    ADS  Article  Google Scholar 

  13. 13.

    Renema, W. Habitat selective factors influencing the distribution of larger benthic foraminiferal assemblages over the Kepulauan Seribu. Mar Micropaleontol 68, 286–298 (2008).

    ADS  Article  Google Scholar 

  14. 14.

    Lee, J. J. & Anderson, O. R. In Biology of foraminifera (eds Lee, J. J. & Anderson, O. R.) (Academic Press, London, 1991).

  15. 15.

    Haynes, J. R. Symbiosis, wall structure and habitat in foraminifera. J Foraminiferal Res 16, 40–43 (1965).

    Google Scholar 

  16. 16.

    Hottinger, L. Distribution of larger Peneroplidae, Borelis and Nummulitidae in the Gulf of Elat, Red Sea. Utrecht Micropaleont Bull B 15, 35–110 (1977).

    Google Scholar 

  17. 17.

    Haynes, J. R. In Foraminifera (Palgrave Macmillan, London, 1981).

  18. 18.

    Hottinger, L., Reiss, Z. & Halicz, E. In Recent foraminiferida from the Gulf of Aqaba, Red Sea 179 (Slovenska Akademija Znanosti in Umetnosti, Ljubljana, 1993).

  19. 19.

    Pecheux, M. J. F. Ecomorphology of a recent large foraminifer, Operculina ammonoides. Geobios 28, 529–566 (1995).

    Article  Google Scholar 

  20. 20.

    Oron, S. et al. Benthic foraminiferal response to the removal of aquaculture fish cages in the Gulf of Aqaba-Eilat, Red Sea. Mar Micropaleontol 107, 8–17 (2014).

    ADS  Article  Google Scholar 

  21. 21.

    Holzmann, M. & Hohenegger, J. Molecular data reveal parallel evolution in nummulitid foraminifera. J Foraminiferal Res 33, 277–284 (2003).

    Article  Google Scholar 

  22. 22.

    Jorry, S. J., Hasler, C. & Davaud, E. Hydrodynamic behaviour of Nummulites: implications for depositional models. Facies 52, 221–235 (2006).

    Article  Google Scholar 

  23. 23.

    Perelis-Grossowicz, L., Edelmen-Furstenberg, Y. & Almogi-Labin, A. In Aqaba - Eilat the Improbable Gulf (ed Por, D.) 439–458 (Hebrew University Magnes Press, Jerusalem, 2008).

  24. 24.

    Hohenegger, J. Distribution of living larger foraminifera NW of Sesoko‐Jima, Okinawa, Japan. Mar Eco 15, 291–334 (1994).

    Article  Google Scholar 

  25. 25.

    Yordanova, E. K. & Hohenegger, J. Morphoclines of living operculinid foraminifera based on quantitative characters. Micropaleontology 50, 149–177 (2004).

    Article  Google Scholar 

  26. 26.

    Fermont, W. J. J. Biometrical investigation of the genus Operculina in Recent sediments of the Gulf of Elat. Utrecht micropaleont bull 15, 171–204 (1977).

    Google Scholar 

  27. 27.

    Röttger, R., Irwan, A., Schmaljohann, R. & Franzisket, L. In Endocytobiology (eds Schwemmler, W. & Schenk, H. E. A.) 125–132 (Walter de Gruyter & Co, Berlin, 1980).

  28. 28.

    Dishon, G., Dubinsky, Z., Fine, M. & Iluz, D. Underwater light field patterns in subtropical coastal waters: A case study from the Gulf of Eilat (Aqaba). Isr J Plant Sci 60, 265–275 (2012).

    Article  Google Scholar 

  29. 29.

    Arar, E. J. & Collins, G. B. In Method 445.0: In vitro determination of chlorophyll a and pheophytin a in marine and freshwater algae by fluorescence (United States Environmental Protection Agency, Office of Research and Development, National Exposure Research Laboratory, Washington, DC, 1997).

  30. 30.

    Hottinger, L. & Dreher, D. Differentiation of protoplasm in Nummulitidae (foraminifera) from Elat, Red Sea. Mar Biol 25, 41–61 (1974).

    Article  Google Scholar 

  31. 31.

    Larsen, A. R. Studies of Recent Amphistegina, taxonomy and some ecological aspects. Isr J Earth Sci 25, 1–26 (1976).

    Google Scholar 

  32. 32.

    Hallock, P. Trends in test shape with depth in large, symbiont-bearing foraminifera. J Foraminiferal Res 9, 61–69 (1979).

    Article  Google Scholar 

  33. 33.

    Hallock, P. & Hansen, H. J. Depth adaptation in Amphistegina: change in lamellar thickness. B Geol Soc Denemark 27, 99–104 (1979).

    Google Scholar 

  34. 34.

    Hallock, P., Forward, L. B. & Hansen, H. J. Influence of environment on the test shape of Amphistegina. J Foraminiferal Res 16, 224–231 (1986).

    Article  Google Scholar 

  35. 35.

    ter Kuile, B. & Erez, J. In situ growth rate experiments on the symbiont-bearing foraminifera Amphistegina lobifera and Amphisorus hemprichii. J Foraminiferal Res 14, 262–276 (1984).

    Article  Google Scholar 

  36. 36.

    Röttger, R. & Berger, W. Benthic Foraminifera: morphology and growth in clone cultures of Heterostegina depressa. Mar Biol 15, 89–94 (1972).

    Article  Google Scholar 

  37. 37.

    Röttger, R. Ecological observations of Heterostegina depressa (Foraminifera, Nummulitidae) in the laboratory and in its natural habitat. Maritime Sédiments Spec 1, 75–79 (1976).

    Google Scholar 

  38. 38.

    Williams, D. F., Röttger, R., Schmaljohann, R. & Keigwin, L. Oxygen and carbon isotopic fractionation and algal symbiosis in the benthic foraminiferan Heterostegina depressa. Palaeogeogr, Palaeoclimatol, Palaeoecol 33, 231–251 (1981).

    Article  Google Scholar 

  39. 39.

    Erez, J. Vital effect on stable-isotope composition seen in foraminifera and coral skeletons. Nature 273, 199–202 (1978).

    ADS  Article  CAS  Google Scholar 

  40. 40.

    Almogi-Labin, A., Perelis-Grossovicz, L. & Raab, M. Living Ammonia from a hypersaline inland pool, Dead Sea area, Israel. J Foraminiferal Res 22, 257–266 (1992).

    Article  Google Scholar 

  41. 41.

    Alve, E. Benthic foraminiferal responses to estuarine pollution: a review. J Foraminiferal Res 25, 190–203 (1995).

    Article  Google Scholar 

  42. 42.

    Geslin, E., Debenay, J., Duleba, W. & Bonetti, C. Morphological abnormalities of foraminiferal tests in Brazilian environments: comparison between polluted and non-polluted areas. Mar Micropaleontol 45, 151–168 (2002).

    ADS  Article  Google Scholar 

  43. 43.

    Röttger, R. & Hallock, P. Shape trends in Heterostegina depressa (Protozoa, Foraminiferida). J Foraminiferal Res 12, 197–204 (1982).

    Article  Google Scholar 

  44. 44.

    Dustan, P. Depth-dependent photoadaption by zooxanthellae of the reef coral Montastrea annularis. Mar Biol 68, 253–264 (1982).

    Article  CAS  Google Scholar 

  45. 45.

    Vareschi, E. & Fricke, H. Light responses of a scleractinian coral (Plerogyra sinuosa). Mar Biol 90, 395–402 (1986).

    Article  Google Scholar 

  46. 46.

    Yap, H., Alvarez, R., Custodio, H. & Dizon, R. Physiological and ecological aspects of coral transplantation. J Exp Mar Bio Ecol 229, 69–84 (1998).

    Article  Google Scholar 

  47. 47.

    Iglesias-Prieto, R., Beltran, V., LaJeunesse, T., Reyes-Bonilla, H. & Thome, P. Different algal symbionts explain the vertical distribution of dominant reef corals in the eastern Pacific. Proc R Soc Lond B Biol Sci 271, 1757–1763 (2004).

    Article  CAS  Google Scholar 

  48. 48.

    Baker, A. C. Ecosystems: reef corals bleach to survive change. Nature 411, 765–766 (2001).

    ADS  Article  PubMed  CAS  Google Scholar 

  49. 49.

    Richier, S. et al. Depth-dependant response to light of the reef building coral, Pocillopora verrucosa: Implication of oxidative stress. J Exp Mar Bio Ecol 357, 48–56 (2008).

    Article  CAS  Google Scholar 

  50. 50.

    Hallock, P., Williams, D. E., Fisher, E. M. & Toler, S. K. Bleaching in foraminifera with algal symbionts: implications for reef monitoring and risk asessment. Anuário do Instituto de Geociências 29, 108–128 (2006).

    Google Scholar 

  51. 51.

    Prazeres, M., Uthicke, S. & Pandolfi, J. M. Changing light levels induce photo-oxidative stress and alterations in shell density of Amphistegina lobifera (Foraminifera). Mar Ecol Prog Ser 549, 69–78 (2016).

    ADS  Article  CAS  Google Scholar 

  52. 52.

    Nobes, K., Uthicke, S. & Henderson, R. Is light the limiting factor for the distribution of benthic symbiont bearing foraminifera on the Great Barrier Reef? J Exp Mar Bio Ecol 363, 48–57 (2008).

    Article  Google Scholar 

  53. 53.

    Ziegler, M. & Uthicke, S. Photosynthetic plasticity of endosymbionts in larger benthic coral reef Foraminifera. J Exp Mar Bio Ecol 407, 70–80 (2011).

    Article  Google Scholar 

  54. 54.

    Hallock, P., Talge, H. K., Cockey, E. M. & Muller, R. G. A new disease in reef-dwelling foraminifera; implications for coastal sedimentation. J Foraminiferal Res 25, 280–286 (1995).

    Article  Google Scholar 

  55. 55.

    Talge, H. K. & Hallock, P. Ultrastructural Responses in Field-Bleached and Experimentally Stressed Amphistegina gibbosa (Class Foraminifera). J Eukaryot Microbiol 50, 324–333 (2003).

    Article  PubMed  Google Scholar 

  56. 56.

    Williams, D. E. & Hallock, P. Bleaching in Amphistegina gibbosa d’Orbigny (Class Foraminifera): Observations from laboratory experiments using visible and ultraviolet light. Mar Biol 145, 641–649 (2004).

    Google Scholar 

  57. 57.

    Schmidt, C., Heinz, P., Kucera, M. & Uthicke, S. Temperature-induced stress leads to bleaching in larger benthic foraminifera hosting endosymbiotic diatoms. Limnol Oceanogr 56, 1587–1602 (2011).

    ADS  Article  Google Scholar 

  58. 58.

    Lesser, M., Stochaj, W., Tapley, D. & Shick, J. Bleaching in coral reef anthozoans: effects of irradiance, ultraviolet radiation, and temperature on the activities of protective enzymes against active oxygen. Coral Reefs 8, 225–232 (1990).

    ADS  Article  Google Scholar 

  59. 59.

    Downs, C. A. et al. Oxidative stress and seasonal coral bleaching. Free Radic Biol Med 33, 533–543 (2002).

    Article  PubMed  CAS  Google Scholar 

  60. 60.

    Lee, J. J. et al. Identification and distribution of endosymbiotic diatoms in larger foraminifera. Micropaleontology 35, 353–366 (1989).

    Article  Google Scholar 

  61. 61.

    Lee, J. J. & Correia, M. Endosymbiotic diatoms from previously unsampled habitats. Symbiosis 38, 251–260 (2005).

    CAS  Google Scholar 

  62. 62.

    Holzmann, M., Berney, C. & Hohenegger, J. Molecular identification of diatom endosymbionts in nummulitid Foraminifera. (Symbiosis (Rehovot) Ser. 42, Balaban Publishers, 2006).

  63. 63.

    Richardson, K., Beardall, J. & Raven, J. Adaptation of unicellular algae to irradiance: an analysis of strategies. New Phytol 93, 157–191 (1983).

    Article  Google Scholar 

  64. 64.

    Palmisano, A. C. et al. Shade adapted benthic diatoms beneath Antarctic sea ice. J Phycol 21, 664–667 (1985).

    Article  Google Scholar 

  65. 65.

    Peña, M. Cell growth and nutritive value of the tropical benthic diatom, sp., at varying levels of nutrients and light intensity, and different culture locations. J Appl Phycol 6, 647–655 (2007).

    Article  Google Scholar 

  66. 66.

    Lee, M., Ellis, R. & Lee, J. A comparative study of photoadaptation in four diatoms isolated as endosymbionts from larger foraminifera. Mar Biol 68, 193–197 (1982).

    Article  Google Scholar 

  67. 67.

    Hallock, P. Light dependence in Amphistegina. J Foraminiferal Res 11, 40–46 (1981).

    Article  Google Scholar 

  68. 68.

    McCloskey, L. & Muscatine, L. Production and respiration in the Red Sea coral Stylophora pistillata as a function of depth. Proc R Soc Lond B Biol Sci 222, 215–230 (1984).

    ADS  Article  Google Scholar 

  69. 69.

    Porter, J., Muscatine, L., Dubinsky, Z. & Falkowski, P. Primary production and photoadaptation in light-and shade-adapted colonies of the symbiotic coral. Stylophora pistillata. Proc R Soc Lond B Biol Sci 222, 161–180 (1984).

    ADS  Article  Google Scholar 

  70. 70.

    Kaiser, P., Schlichter, D. & Fricke, H. Influence of light on algal symbionts of the deep water coral Leptoseris fragilis. Mar Biol 117, 45–52 (1993).

    Article  Google Scholar 

  71. 71.

    Masuda, K., Goto, M., Maruyama, T. & Miyachi, S. Adaptation of solitary corals and their zooxanthellae to low light and UV radiation. Mar Biol 117, 685–691 (1993).

    Article  Google Scholar 

  72. 72.

    Leletkin, V., Titlyanov, E. & Dubinsky, Z. Photosynthesis and respiration of the zooxanthellae in hermatypic corals habitated on different depths of the Gulf of Eilat. Photosynthetica 32, 481–490 (1996).

    Google Scholar 

  73. 73.

    Cohen, I. & Dubinsky, Z. Long term photoacclimation responses of the coral Stylophora pistillata to reciprocal deep to shallow transplantation: photosynthesis and calcification. Front Mar Sci 2, 45 (2015).

    Article  Google Scholar 

  74. 74.

    Mass, T. et al. The spectral quality of light is a key driver of photosynthesis and photoadaptation in Stylophora pistillata colonies from different depths in the Red Sea. J Exp Biol 213, 4084–4091 (2010).

    Article  PubMed  CAS  Google Scholar 

  75. 75.

    Kuczynska, P., Jemiola-Rzeminska, M. & Strzalka, K. Photosynthetic pigments in diatoms. Mar Drugs 13, 5847–5881 (2015).

    Article  PubMed  PubMed Central  CAS  Google Scholar 

  76. 76.

    Kühl, M., Cohen, Y., Dalsgaard, T., Jørgensen, B. & Revsbech, N. Microenvironment and photosynthesis of zooxanthellae in scleractinian corals studied with microsensors for O2, pH and light. Mar Ecol Prog Ser 117, 159–172 (1995).

    ADS  Article  Google Scholar 

  77. 77.

    Cohen, I., Dubinsky, Z. & Erez, J. Light Enhanced Calcification in Hermatypic Corals: New Insights from Light Spectral Responses. Front Mar Sci 2, 122 (2016).

    Article  Google Scholar 

  78. 78.

    Erez, J. In Biomineralization and biological metal accumulation (eds Westbroek, P. & de Jong, E. J.) 307–312 (Springer, Renesse, The Netherlands, 1983).

  79. 79.

    ter Kuile, B., Erez, J. & Padan, E. Competition for inorganic carbon between photosynthesis and calcification in the symbiont-bearing foraminifer Amphistegina lobifera. Mar Biol 103, 253–259 (1989).

    Article  Google Scholar 

Download references

Acknowledgements

We wish to express our gratitude to the Inter-University Institute (IUI) of Eilat for providing the facilities to carry out this study. We thank Gal Eyal for accompanying Shai Oron on her CCR diving sessions. The research was supported by the Israel Science Foundation grant No 587/2013. This research is a part of the first author’s PhD dissertation, and was also funded by the Department of Geological and Environmental Sciences at Ben-Gurion University of the Negev and by the Inter-University Institute of Marine Sciences in Eilat (IUI).

Author information

Affiliations

Authors

Contributions

S.O. conducted the experiments and wrote the main manuscript text under the supervision of J.E., S.A. and A.A. J.W. prepared the microtomography scans for Fig. 5. All authors reviewed the manuscript.

Corresponding author

Correspondence to Shai Oron.

Ethics declarations

Competing Interests

The authors declare no competing interests.

Additional information

Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Electronic supplementary material

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Oron, S., Abramovich, S., Almogi-Labin, A. et al. Depth related adaptations in symbiont bearing benthic foraminifera: New insights from a field experiment on Operculina ammonoides. Sci Rep 8, 9560 (2018). https://doi.org/10.1038/s41598-018-27838-8

Download citation

Further reading

Comments

By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.

Search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing