Abstract
Identifying and quantifying nitrogen pools is essential for understanding the nitrogen cycle in aquatic ecosystems. The ubiquitous diatoms represent an overlooked nitrate pool as they can accumulate nitrate intracellularly and utilize it for nitrogen assimilation, dissipation of excess photosynthetic energy, and Dissimilatory Nitrate Reduction to Ammonium (DNRA). Here, we document the global co-occurrence of diatoms and intracellular nitrate in phototrophic microbial communities in freshwater (n = 69), coastal (n = 44), and open marine (n = 4) habitats. Diatom abundance and total intracellular nitrate contents in water columns, sediments, microbial mats, and epilithic biofilms were highly significantly correlated. In contrast, diatom community composition had only a marginal influence on total intracellular nitrate contents. Nitrate concentrations inside diatom cells exceeded ambient nitrate concentrations ∼100–4000-fold. The collective intracellular nitrate pool of the diatom community accounted for <1% of total nitrate in pelagic habitats and 65–95% in benthic habitats. Accordingly, nitrate-storing diatoms are emerging as significant contributors to benthic nitrogen cycling, in particular through Dissimilatory Nitrate Reduction to Ammonium activity under anoxic conditions.
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Introduction
Microorganisms involved in aquatic nitrogen cycling typically exploit freely dissolved inorganic nitrogen species, such as nitrate, nitrite, and ammonium, for the different nitrogen-cycle pathways1. The distribution of freely dissolved nitrogen pools within aquatic ecosystems thus often reflects the distribution and activity of the different nitrogen-cycle pathways and vice versa2,3. Some aquatic microorganisms, however, can accumulate and store nitrate in cell vacuoles at concentrations exceeding ambient concentrations by several orders of magnitude4,5,6. It is commonly believed that intracellular nitrate (ICNO3) is utilized for nitrogen assimilation (N-assimilation) by phototrophic eukaryotes, such as diatoms7,8, and for energy conservation through dissimilatory nitrate reduction by sulfide-oxidizing prokaryotes5,9,10. This paradigm was shattered by the discovery that several groups of marine microbial eukaryotes also utilize ICNO3 for dissimilatory nitrate reduction, i.e., foraminifers11, diatoms12, fungi13, and gromiids14. Diatoms can utilize ICNO3 for Dissimilatory Nitrate Reduction to Ammonium (DNRA) to survive dark and anoxic conditions12.
Since nitrate uptake in excess of immediate nutritional demands (i.e., “luxury uptake”) occurs against a steep concentration gradient, it requires an energy investment5,15. In turn, nitrate storage conveys a competitive advantage in environments where the supply of nitrate is scarce, spatially confined, or temporally variable16. Nitrate uptake and intracellular storage in nitrate-replete subsurface layers allows survival and growth of migrating diatoms in nitrate-depleted surface layers of the ocean17. In benthic habitats with spatially separated electron acceptors (e.g., oxygen and nitrate) and donors (e.g., organic substrates and sulfide), nitrate-storing sulfur bacteria thrive by displaying vertical migration behavior between surface and subsurface layers4,18. Diel vertical migration behavior enabled by ICNO3 storage and DNRA activity was only recently demonstrated for benthic diatoms19.
Diatoms are ubiquitous in shallow freshwater and marine ecosystems where they often dominate phototrophic microbial communities in both pelagic and benthic habitats20,21,22,23. The ability to store nitrate intracellularly has been demonstrated for several diatom isolates and pelagic diatom communities8,12,24,25,26,27, but not in a comprehensive way for the extensive diatom communities in sediments, microbial mats, and biofilms. Thus, the distribution and extent of ICNO3 storage by diatom communities within and across diverse aquatic ecosystems remains unexplored. Knowing the distribution and size of diatom-specific ICNO3 pools can serve as the basis to gauge the influence by diatoms on aquatic nitrogen cycling. Three nitrate-fueled nitrogen-cycle pathways have been described for diatoms so far, N-assimilation7,8, nitrate reduction for dissipating excess photosynthetic energy (NRphot)8,28,29,30, and DNRA under anoxic conditions12,19.
Aside from fueling different nitrogen-cycle pathways, nitrate-storing diatoms may represent a significant local sink of freely dissolved nitrate (FDNO3) in sediments and microbial mats. Efficient ICNO3 storage keeps porewater nitrate concentrations low and thereby increases the diffusive flux of nitrate from adjacent sources (e.g., the water column), which influences nitrate availability for the entire benthic habitat. The local sink of FDNO3 is decoupled in space and time from the ultimate diatom-mediated ICNO3 turnover and thus from the uptake/release of other involved reactive compounds (e.g., ammonium, dissolved inorganic carbon). High diatom abundances may thus lead to misinterpretation of porewater profiles and incorrect rate calculations of nitrogen-cycle pathways in benthic habitats, which should be corrected for by biogeochemical modeling that takes ICNO3 storage into account.
We studied the co-occurrence of diatoms and ICNO3 in phototrophic microbial communities sampled in 117 globally distributed marine and freshwater habitat types (i.e., water column, sediment, microbial mat, and epilithic biofilm). We hypothesized that high diatom abundance coincides with high total ICNO3 contents in space (vertical profiling) and time (seasonal monitoring). The role of diatom community composition for the prevalence of ICNO3 storage was assessed via 18S rRNA gene amplicon sequencing. The rich dataset provided here revealed that habitat type and diatom abundance are key determinants for total ICNO3 contents within aquatic ecosystems, while diatom community composition is of marginal importance. In benthic habitats, most of the total nitrate pool may indeed occur as intracellular nitrate stored by diatoms, suggesting that benthic nitrogen cycling is strongly influenced by diatoms.
Results
Global distribution of intracellular nitrate
A snapshot survey of phototrophic microbial communities was carried out in 117 globally distributed freshwater (69 sites) and marine (48 sites) habitats (Fig. 1, Supplementary Table 1, Supplementary Data 1). Four different habitat types were sampled: water columns (16 sites), sediments (43 sites), microbial mats (22 sites), and epilithic biofilms (36 sites). Open-ocean water column samples are underrepresented in this survey (4 sites). All samples from the snapshot survey were routinely analyzed for photopigments, ICNO3, and freely dissolved nitrate (FDNO3). In two sample subsets, diatom abundance and biovolume (Supplementary Table 2 and Supplementary Data 2), as well as diatom community composition (Supplementary Table 3 and Supplementary Data 3) were determined.
a Global and b Central European distribution of 117 sampling sites (numbered blue circles) at which the occurrence and the determinants of intracellular nitrate were studied. The map was created using the software package ODV 5.6.1 (Schlitzer, Reiner, Ocean Data View, odv.awi.de, 2021). Sampling details are given in Supplementary Table 1 (Supplementary Data 1) and in an interactive map created using Google My Maps (https://bit.ly/2XF9kkQ).
Habitat type
Total ICNO3 contents spanned seven orders of magnitude across all habitat types, from very low levels in water columns to very high levels in epilithic biofilms (Fig. 2 and Supplementary Data 5). ICNO3 contents were significantly correlated with chlorophyll a contents across all habitat types (Supplementary Fig. 1A, R2 = 0.799, p < 0.001, n = 654), suggesting a role of phototrophic microorganisms in ICNO3 storage. ICNO3 contents were even more tightly correlated with fucoxanthin contents across all habitat types (Fig. 2, R2 = 0.864, p < 0.001, n = 654), which is described by the power function ICNO3 = 3.77 ± 1.06 × Fucoxanthin0.753 ± 0.013 (means ± standard error). In turn, fucoxanthin contents were significantly correlated with diatom cell abundance as determined microscopically in a subset of samples (Supplementary Fig. 2, R2 = 0.954, p < 0.001, n = 78). These correlations suggest that diatoms are important for ICNO3 storage in phototrophic microbial communities and that fucoxanthin contents can be used as a proxy for the abundance of nitrate-storing diatoms.
Overall correlation between intracellular nitrate (ICNO3) and fucoxanthin (a diatom marker pigment) across habitat types and climate zones. Total ICNO3 and fucoxanthin contents are expressed per unit volume of the respective habitat. The correlation is described by the power function: ICNO3 (µmol L−1) = 3.77 × Fucoxanthin (µmol L−1)0.75 (R2 = 0.864, p < 0.001, n = 654). Inset shows frequency distribution (in %) of samples in which intracellular nitrate (N) and fucoxanthin (F) were detected (+) or not detected (–).
However, fucoxanthin also occurs in Dinophyceae, Prymnesiophyceae, Chrysophyceae, and Pelagophyceae. Published and newly added data indicate that these microalgal classes store intracellular nitrate at negligible concentrations compared to diatoms (Table 1, Supplementary Table 4, Supplementary Data 4). Additionally, the quantification of several marker pigments in this survey revealed that the relative abundance of the respective microalgal class was not positively correlated with total ICNO3 contents across all habitat types (Supplementary Table 5). Low nitrate storage capacity and lack of correlation between relative abundance and total ICNO3 contents were also found in microalgal classes that do not contain fucoxanthin, i.e., Cryptophyceae, Chlorophyceae, and the prokaryotic cyanobacteria (Table 1, Supplementary Tables 4 and 5). Microalgal classes other than diatoms and cyanobacteria are thus unlikely to contribute substantially to total ICNO3 contents in aquatic habitats.
In contrast, the quantitative relationship between ICNO3 and diatoms is substantiated by finding that ICNO3 and fucoxanthin contents were also correlated within each habitat type (Supplementary Table 6) and that total ICNO3 contents were positively correlated with the fucoxanthin/chlorophyll a ratio (i.e., a proxy for relative diatom abundance) (Supplementary Fig. 1B and Supplementary Table 6). Additionally, most samples (79%) harbored both ICNO3 and fucoxanthin, while samples harboring ICNO3 but lacking fucoxanthin were rare (2%) (Fig. 2).
Freely dissolved nitrate
The influence of FDNO3 concentrations on total ICNO3 contents was ambiguous (Supplementary Fig. 3). Monotonic relationships across habitat types were neither observed for water column nitrate, nor for porewater nitrate (PWNO3) (Supplementary Fig. 3A, B). However, significant, positive correlations were found for sediments and epilithic biofilms (Supplementary Table 6).
Diatom abundance and biovolume
For 6–8 sampling sites within each habitat type (Supplementary Table 2 and Supplementary Data 1), diatom cells were counted and sized by microscopy. Both diatom cell abundance and total diatom biovolume spanned 6–7 orders of magnitude across all habitat types, from very low values in water columns to very high values in mats and biofilms (Fig. 3a and Supplementary Data 6). Total ICNO3 contents were highly significantly correlated with diatom cell abundance and total diatom biovolume across all habitat types (Fig. 3a, b, R2 = 0.823 and R2 = 0.899, respectively, p < 0.001, n = 84) and also within most habitat types (Supplementary Table 6). This confirms that total fucoxanthin contents indeed reflected diatom cell abundance and total diatom biovolume in this survey.
a Diatom abundance and total diatom biovolume, b total ICNO3 contents, and c ICNO3 concentration inside diatom cells in water columns (blue), sediments (brown), microbial mats (green), and epilithic biofilms (cyan). Note different units for ICNO3 in b, c. Percent values in b give the median of the relative contribution of ICNO3 to total nitrate in each habitat type. Multiplication factors in c give the median of the enrichment of ICNO3 concentration inside diatom cells relative to freely dissolved nitrate (FDNO3) concentration in each habitat type. Box plots show median (solid line), 25th and 75th percentiles (box), and 10th and 90th percentiles (whiskers) of nA = 6–8 sampling sites within each habitat type and nB = 3 replicates each.
The relative contribution of ICNO3 to total nitrate in each sample was calculated as ICNO3/(ICNO3 + FDNO3) for water column samples and as ICNO3/(ICNO3 + PWNO3) for sediment, mat, and biofilm samples and expressed in percent. It turned out that ICNO3 accounted for 0.1% of total nitrate in pelagic habitats and 65–95% in benthic habitats (Fig. 3b and Supplementary Data 6). Assuming that the total ICNO3 contents were exclusively stored by diatoms, the average ICNO3 concentration inside diatom cells ranged between 1.6 (diatom mats) and 21.8 mmol L−1 (water column) (Fig. 3c and Supplementary Data 6). Strikingly, the ICNO3 concentrations inside diatom cells exceeded the respective FDNO3 concentrations between 95- and 4082-fold in the different habitat types (Fig. 3c), again assuming that diatoms were the exclusive microorganisms storing ICNO3 in these samples.
Vertical distribution of intracellular nitrate
In 16 vertically profiled aquatic habitats, the total ICNO3 and photopigment contents were generally highest close to the sunlit surface and decreased with depth (Fig. 4, Supplementary Fig. 4, Supplementary Data 7). Fucoxanthin and ICNO3 profiles had matching shapes on a meter (water column), centimeter (sediment), and millimeter scale (microbial mat). This was not the case for phycocyanin, a marker pigment for cyanobacteria, which can be abundant in microbial mats (Fig. 4g and Supplementary Fig. 4). In most sediments and microbial mats, the vertical profiles of ICNO3 and PWNO3 contents had similar shapes with clear surface maxima and very low levels at greater depth (Fig. 4 and Supplementary Fig. 4).
a, c, e, g Total contents of chlorophyll a (Chl a), fucoxanthin (Fuco), and phycocyanin (Phyco) (a cyanobacteria marker pigment) and b, d, f, h freely dissolved nitrate (FDNO3) and intracellular nitrate (ICNO3) in one lake water column, two sediments, and one microbial mat. For sediments and microbial mats, FDNO3 refers to porewater nitrate. For Kongsfjord, no phycocyanin data are available. Note that y-axes have meter, centimeter, and millimeter scales for water column, sediment, and microbial mat, respectively. Means ± standard deviation of n = 3 are shown; for better visibility, vertical sampling points are slightly offset. Vertical profiles for twelve additional sampling sites are shown in Supplementary Fig. 4.
As an extreme case of microbial stratification, thin, golden-brown diatom layers were skimmed off sediment surfaces at nine sampling sites and analyzed separately (Supplementary Table 1 and Supplementary Data 1). Without exception, these “diatom mats” had very high total ICNO3 and fucoxanthin contents. Similarly, epilithic biofilms were detached from the surface of sunlight-exposed stones and pebbles at 36 sampling sites (Supplementary Table 1 and Supplementary Data 1). Epilithic biofilms did not only display the highest average contents of ICNO3 (476 µmol L−1, n = 134) and fucoxanthin (192 µmol L−1, n = 139), but also the single highest values of ICNO3 (3329 µmol L−1) and fucoxanthin (1451 µmol L−1) of all samples analyzed in this survey.
Seasonal variation of intracellular nitrate
In the water column of four seasonally monitored aquatic habitats from the temperate zone, total chlorophyll a, fucoxanthin, and ICNO3 contents showed one or two pronounced maxima during the warm season (Fig. 5 and Supplementary Data 8). ICNO3 maxima coincided to a large extent with fucoxanthin maxima, i.e., with the appearance of diatom blooms. In contrast, total ICNO3 and phycocyanin contents did not co-vary. Coinciding with the emerging spring blooms, FDNO3 concentrations decreased and remained low during the warm season. Nevertheless, total ICNO3 contents remained high, indicating that the low FDNO3 concentrations did not limit nitrate uptake and ICNO3 storage.
a, c, e, g Water column contents of chlorophyll a (Chl a), fucoxanthin (Fuco), and phycocyanin (Phyco) and b, d, f, h freely dissolved nitrate (FDNO3) and intracellular nitrate (ICNO3) in two rivers, one pond, and one coastal marine bay. For better visibility, photopigment and nitrate contents were multiplied by an appropriate factor as indicated in the respective panel. Gray shading marks time periods during which fucoxanthin and ICNO3 maxima appeared. For Århus Bay, no phycocyanin data are available. For Weser, monthly means ± standard deviation of n = 5 were calculated from multiple samplings during 2017–2021.
Diatom community composition
For 7–16 sampling sites within each habitat type (Supplementary Table 3 and Supplementary Data 3), diatoms were identified through 18S rRNA gene sequence analysis. In total, 43 out of 53 samples produced each >5000 sequence reads and were used for further analysis (Supplementary Table 3 and Supplementary Data 3). The ten samples excluded from further analysis produced <5000 sequence reads and corresponded without exception to samples with very low total ICNO3 and fucoxanthin contents and diatom abundance. Based on relative sequence abundance, centric diatoms of the order Thalassiosirales (class Mediophyceae) dominated the communities in most water columns, while pennate diatoms of the order Naviculales (class Bacillariophyceae) dominated the communities in most sediment, mats, and biofilms (Fig. 6). Other abundant diatom orders in benthic habitats included Fragilariales (sediments), Thalassiosirales and Bacillariales (diatom mats), and Achnanthales, Bacillariales, and Cymbellales (epilithic biofilms).
Heatmap of relative 18S rRNA gene sequence abundance at the level of diatom classes and orders for freshwater (F) and marine (M) water column, sediment, diatom mat, and epilithic biofilm samples. Relative sequence abundance scales from white (<0.15%) to dark blue (100%). Sample IDs are given in Supplementary Tab. 1 (Supplementary Data 1).
All habitat types had significantly different diatom communities (ANOSIM: R = 0.274 ± 0.079, p < 0.05), except for sediments and diatom mats, which shared similar diatom communities (ANOSIM: p > 0.05) (Fig. 7 and Supplementary Table 7). There was also a significant difference in diatom community composition between marine and freshwater habitats (ANOSIM: R = 0.53, p < 0.001). Notably, the variation in total ICNO3 contents was only marginally explained by dissimilarities of diatom communities (PERMANOVA: R2 = 0.04, p = 0.011).
NMDS plot of Bray–Curtis dissimilarities from 43 diatom communities with a total of 2157 amplicon sequence variants (ASVs) retrieved from different habitat types.
Discussion
Diatoms as widespread nitrate-storing microorganisms
Our results provide two lines of evidence supporting the hypothesis that diatoms are the major nitrate-storing organisms in phototrophic microbial communities worldwide: (i) the highly significant correlation of total ICNO3 contents with diatom abundance and biovolume (and also with fucoxanthin contents) across all habitat types, and (ii) the spatial and temporal co-occurrence of ICNO3 and fucoxanthin maxima. This fits well with the previously demonstrated high ICNO3 storage capacity of diatom isolates8,12,24,25,26,27. Since fucoxanthin is also found in microalgal classes other than diatoms and since total ICNO3 contents correlated also with chlorophyll a contents, the potential contribution of other microalgal classes to total ICNO3 contents was evaluated. Generally, the contribution of nitrate-storing microorganisms to total ICNO3 contents in the environment increases with the ICNO3 concentration level inside the cell, with cell volume, and with cell abundance. Among microalgae, diatoms store the by far highest ICNO3 concentration levels and possess on average very large cells, which together results in cell-specific ICNO3 contents that are typically two orders of magnitude higher than in any other microalgal class (Table 1). Even large-celled Dinophyceae would have to be ~100 times more abundant than diatoms to contribute the same amount of nitrate to the total ICNO3 contents in a given habitat. However, the peridinin/chlorophyll a ratio (a proxy for the relative abundance of Dinophyceae) was on average >7 times lower in environmental samples than in Dinophyceae isolates (Supplementary Table 5). Additionally, the peridinin/chlorophyll a ratio did not correlate with total ICNO3 contents across all habitat types (Supplementary Table 5). Taken together, it is very unlikely that Dinophyceae contributed substantially to total ICNO3 contents in this survey. The same reasoning applies to Cryptophyceae, Chlorophyceae, cyanobacteria, and the extremely small Pelagophyceae. The pigment-based argumentation is less clear for Prymnesiophyceae and Chrysophyceae as they do not contain unequivocal marker pigments, but their very low ICNO3 storage capacity still disqualifies them as important contributors to total ICNO3 contents. Noteworthy, both fucoxanthin/chlorophyll a and diadinoxanthin/chlorophyll a ratios were nearly the same in environmental samples vs. diatom isolates, which argues for generally high relative diatom abundances in this survey.
Clearly, the most important determinant for the measured total ICNO3 contents is habitat type. The connection between total ICNO3 contents and habitat type must be mediated by the vastly differing diatom abundance and biovolume in phototrophic microbial communities of different aquatic habitats, which culminates in extremely high values of diatom abundance and biovolume and total ICNO3 contents in epilithic biofilms. Additionally, the habitat types differed significantly in diatom community composition, with the exception of sediments and diatom mats. The latter observation is likely explained by the fact that diatom mats had developed on top of sediments and thus diatom communities of the two habitat types may overlap. Different diatom species may possess different ICNO3 storage capacities, as also observed for foraminifers31, which may scale up to differences in total ICNO3 contents in the environment. However, diatom community composition was not a significant determinant of total ICNO3 contents in phototrophic microbial communities. This suggests that the ability to store ICNO3 is widespread in diatoms and/or that species-specific differences in ICNO3 storage capacity are averaged out in the highly diverse diatom communities.
Implications of ICNO3 storage for diatoms
For the individual diatom cell, ICNO3 serves as a buffer for time periods during which the cell is cut off from ambient FDNO3 supply. This may be the case in environments where FDNO3 is spatially constrained as in stratified water columns or benthic habitats17,19, or where FDNO3 concentrations fluctuate over time, such as in tidally flushed sediments32,33. For marine pelagic diatoms, the ICNO3 pool is reported to last for 3–4 days24,34, which is sufficient time for migrating between the nitrate-replete subsurface layer and the nitrate-depleted surface layer in the ocean17. For benthic diatoms, however, the ICNO3 pool is reported to last for ~12–24 h only12,19,33, which is still sufficient time for bridging nitrate-depleted conditions encountered during tidal cycles or migration in sediments or mats with steep concentration gradients19,33. The longer lifetime of the ICNO3 pool in pelagic diatoms is consistent with the higher ICNO3 concentrations inside the cells of pelagic diatoms documented in our survey.
Diatoms can use their ICNO3 pool for N-assimilation24,35. In the size range of diatom cells in our survey (i.e., 550–19450 µm3 per cell), the ICNO3 pool of a single cell corresponded to ~1% of its total nitrogen content as estimated from cell volume36. This low value is consistent with another survey of different marine phytoplankton species24 and implies that diatoms in this size range cannot sustain N-assimilation with their relatively small ICNO3 pool alone, but must rely on intracellular ammonium, urea, and/or amino acids as additional nitrogen sources24,37. In contrast, in the very large pelagic diatom Ethmodiscus sp. (i.e., 5 × 108 µm3 per cell), the ICNO3 pool corresponds to 135% of the particulate organic nitrogen content and can support one cell division within four days34. This discrepancy is explained by the increase in relative vacuole volume with increasing cell size (i.e., from 30–60% for the size range of diatoms in our survey to >90% for Ethmodiscus sp. as estimated from cell volume38).
Diatoms possess an exceptional potential to survive dark and anoxic conditions through physiological and metabolic adaptations, such as DNRA activity, dark fermentation, and resting stage formation12,39,40. Diatom-mediated DNRA activity, in particular, is induced by a change from oxic to anoxic conditions due to, e.g., tidal-induced porewater movement, diatom migration behavior in benthic habitats, or passive sinking of diatom aggregates through an oxygen minimum zone12,18,26. The ICNO3 pool can then be used for energy conservation through anaerobic nitrate respiration until oxic conditions re-establish within 12–24 h12,19,33. In cases where anoxic conditions persist for very long time periods (e.g., after settlement of diatoms onto anoxic sediments), diatoms might use the energy conserved through DNRA activity for the transition to a resting stage, which can be completed within 8–10 h12,41.
Surprisingly, ICNO3 storage was also observed in habitats in which neither FDNO3 limitation, nor anoxic conditions are likely to occur, such as in the water column of nitrate-rich streams. One possible explanation for this phenomenon is that the cytosolic nitrate reductase of diatoms has a relatively low affinity (i.e., a high km value) for nitrate, which requires an intracellular up-concentration to enable high enzymatic conversion rates. Indeed, the km value of assimilatory nitrate reduction is 50–290 µmol L−1 and thus higher than FDNO3 concentrations in many aquatic habitats8. Additionally, in diatoms exposed to rapidly increasing irradiance levels, ICNO3 may be used for NRphot, the dissipation of excess photosynthetic energy via nitrate reduction to nitrite and/or ammonium, which are expelled from the cells8,28,29,30. The NRphot pathway has originally been described for pelagic diatom communities in the oceans, but might also exist in the nitrate-rich streams studied here where diatoms have unlimited access to nitrate, but experience pronounced differences in light levels due to the turbulent water flow. The NRphot pathway may also be of importance in microbial/diatom mats exposed to high irradiation levels, but this remains to be studied. Taken together, nitrate accumulation might be an evolved trait of diatoms that is constitutively expressed despite ample availability of FDNO3.
Implications of ICNO3 storage by diatoms for nitrogen cycling
The total ICNO3 contents measured within a given aquatic habitat can be used to predict the occurrence of specific nitrate-fueled metabolic pathways, if the major nitrate-storing organisms are also known. For example, denitrification and DNRA are likely to occur in oxygen-depleted aquatic habitats in which foraminifers and diatoms, respectively, are abundant11,12. In the phototrophic microbial communities studied here, diatoms were the major nitrate-storing organisms and thus diatom-mediated N-assimilation, NRphot, and DNRA were likely the most important pathways fueled by ICNO3. Utilizing either pathway, diatoms contribute to the retention of fixed nitrogen in the ecosystem in the form of (particulate) organic nitrogen, ammonium, and/or nitrite rather than to the loss of fixed nitrogen to dinitrogen gas as occurs through denitrification and anammox1.
Beyond this qualitative assessment, it is not possible to predict the quantitative extent to which nitrogen cycling is impacted by nitrate-storing diatoms as only the size of the ICNO3 pool was measured but not its turnover. Pure culture studies indicate that the ICNO3 pool of diatoms is turned over on time scales of minutes to hours through NRphot28,29,30 and DNRA12,25. The very few studies on nitrate-storing diatoms in environmental settings report turnover times of the ICNO3 pool on the order of hours to days19,26,33. Ultimately, the rates of nitrogen-cycle pathways fueled by ICNO3 vs. FDNO3 will have to be compared in situ to gauge the contribution by nitrate-storing diatoms to nitrogen cycling, a task that is still pending.
In the pelagic zone of nutrient-rich freshwater and marine ecosystems, total ICNO3 contents are usually in the nanomolar range and contribute <1% to total nitrate, even during diatom blooms42,43,44,this survey. In oligotrophic regions of the open ocean, which are not represented in our dataset, the relative contribution of ICNO3 to total nitrate remains unresolved. However, at nitrate concentrations of ~10 nmol L−1 in the surface water of these regions, only one single cell of Ethmodiscus sp., which stores 11 nmol ICNO3, needs to be present in 1 L seawater to reach a relative contribution of ~50%34. Additionally, NRphot activity by pelagic diatoms may be significant due to turbulent water flow, for example, in coastal upwelling zones of the oceans where diatoms are abundant8,21,28. Since ICNO3 is turned over to and expelled as nitrite and/or ammonium, NRphot activity by pelagic diatoms supplies nitrifying microorganisms with substrates, a link not established in situ so far. High NRphot activity represents also a methodological issue because it is assumed to bias “new production” estimates for the oceans29. DNRA activity by pelagic diatoms is likely precluded by the presence of oxygen in most water columns, but occurs inside anoxic “marine snow” aggregates that display DNRA rates equivalent to 16–28% of denitrification rates26,45. However, even at high aggregate abundance in the water column of oxygen minimum zones, these DNRA rates are low (<1 nmol NH4+ L−1 d−1 (ref. 45) relative to DNRA rates in the ambient water of up to 40 nmol NH4+ L−1 d−1 (ref. 46).
In benthic habitats with abundant diatoms, ICNO3-fueled DNRA rates may indeed rival FDNO3-fueled denitrification rates. In an intertidal sediment, the ICNO3 stored by the diatom-dominated eukaryotic community fueled DNRA activity equivalent to ~10–20% of denitrification activity3,33. In a mat-covered lake sediment, DNRA activity made up 98–99% of total dissimilatory nitrate reduction and was ascribed to the nitrate-storing and migrating diatom Craticula cuspidata19. Notably, this dominance of DNRA coincided with an extremely high relative contribution of ICNO3 to total nitrate of 98%. In a rocky biofilm, high DNRA activity corresponding to 59–69% of total dissimilatory nitrate reduction and was ascribed to anoxic conditions prevailing at night47. While the presence of diatoms in this rocky biofilm was not reported, essentially all epilithic biofilms studied in our survey were dominated by nitrate-storing diatoms and might thus emerge as hotspots of NRphot activity during the day and DNRA activity at night. Notably, these reported relative contributions of ICNO3 turnover by diatoms to benthic nitrogen cycling are in a similar range as the relative contribution of ICNO3 contents to total nitrate contents as established in our survey for benthic habitats (i.e., 65–95%). The reason why diatom-mediated DNRA activity, in particular, might have been overlooked so far pertains to the general neglect of DNRA compared to denitrification and anammox in N-cycle studies3,48 and the common perception that phototrophic communities do not host anaerobic nitrogen cycling because of photosynthetic oxygen production during daytime. However, phototrophic sediments and mats are always underlain by anoxic subsurface layers and even thin phototrophic biofilms may experience anoxia in darkness due to high community respiration rates49,50. Also very large aggregations of pelagic diatoms, such as the centimeter- to meter-scale strands of Melosira arctica underneath sea-ice floes51,52, might emerge as oxygen-depleted microenvironments in which ICNO3 storage and DNRA activity occur. The widespread occurrence of ICNO3 storage by diatoms demonstrated in this survey and the growing number of diatom isolates capable of DNRA strongly suggest that significant DNRA activity will be discovered in many more aquatic ecosystems.
Methods
Sample collection and preparation
Phototrophic microbial communities were sampled across diverse freshwater and marine ecosystems on all continents and in all climate zones (Fig. 1, Supplementary Table 1, Supplementary Data 1). The sampling sites covered four different habitat types: water column, sediment, microbial mat, and epilithic biofilm. Vertical profiles of intracellular nitrate (ICNO3) contents were recorded in the water column, sediment, and microbial mat of 16 sampling sites. Seasonal cycles of ICNO3 contents were monitored in the water column of four sampling sites. Snapshot samples were collected at 117 globally distributed sampling sites.
Water column samples were collected using (i) 2-L polyethylene bottles for surface water samples and (ii) 8-L Niskin bottles for deeper water samples. Water volumes of 0.1–2 L were filtered through pre-combusted 0.3-µm glass fiber filters (GF75, Advantec Inc., U.S.A.) that were stored in 15-mL sample tubes at −20 °C for later analysis of ICNO3 and photopigments. An aliquot of 1 mL filtrate was stored at −20 °C for later analysis of freely dissolved nitrate (FDNO3).
Sediment samples were collected using (i) perspex core liners (3.6 cm internal diameter) to retrieve ~15-cm-long sediment cores or (ii) weighing spoons to scoop the top 0.5 or 1 cm of sediment into 50-mL sample tubes that were 1/3 filled with sediment, 1/3 overlying water (to maintain nitrate availability), and 1/3 air (to maintain oxic conditions). Sediment samples were processed in the laboratory within 1–2 h after collection (sediment cores) or 1–7 days after collection and transport (sediment top layers). Intact sediment cores were sliced at 0.5-1-cm intervals to a total depth of 5–10 cm. Each slice of known wet volume was split into three equal parts and transferred to 15-mL sample tubes for later analysis of nitrate as well as lysozyme- and acetone/ethanol-extracted photopigments. Sediment top layers were vigorously mixed, split into three equal parts, and transferred to 15-mL sample tubes for nitrate and photopigment analyses. The split samples were mildly centrifuged (500 × g for 5 min) to determine the individual wet volume as read from the scale of the sample tube.
Microbial mats were collected as (i) ~10 mm-thick laminated mats that were retrieved using a scalpel and (ii) ~1 mm-thin “diatom mats” that were skimmed off sediment surfaces using a weighing spoon. Laminated mats were originally sampled in the field and maintained in the laboratory under in situ conditions regarding light regime, temperature, and salinity for longer time periods53,54. Mat samples were processed in the laboratory immediately after harvesting them from maintenance tanks (laminated mats) or within 1 h after collection at the sampling site (diatom mats). Laminated mats were sliced using a stainless steel tube (1 cm internal diameter) for coring and a stainless steel rod (1 cm outer diameter) and a spatula for extruding and slicing the mat core, respectively. The thickness of the mat slices was guided by the visible functional layering of each mat. For mats with unclear functional layering, the top 1 cm was treated as one bulk sample. Diatom mat samples were treated in the same way as sediment top layers.
Epilithic biofilms were collected as intact biofilms covering (i) pebbles that were transported in 50-mL sample tubes or (ii) larger stones that were transported in 250-mL plastic containers. Samples were processed in the laboratory within 1–2 h after collection (local sites) or 1–7 days after collection and transport (remote sites). Biofilms were detached from pebbles and larger stones by abrasion and brushing, respectively. All pebbles from one 50-mL sample tube were transferred to a 250-mL polyethylene bottle to which 50 mL of nitrate-free water (NFW) prepared from Red Sea salt (Red Sea Aquatics Ltd., U.K.) was added. For freshwater and marine samples, NFW was prepared at salinities of 0.02 and 35, respectively, and adjusted to pH values of 7.0 and 8.2, respectively. The closed bottle was vigorously shaken for 2 min to enable abrasion of biofilms as the pebbles rubbed against each other. The obtained biofilm suspension was split into three equal parts and transferred into 15-mL sample tubes for ICNO3 and photopigment analyses. The first round of centrifugation at 500 × g for 5 min served to determine the wet volume of the biofilm. The second round of centrifugation at 3000 × g for 5 min served to compact the biofilm pellet so that the clear supernatant could be discarded. Biofilms on larger stones (up to 10 cm in diameter) were thoroughly brushed off using a vegetable brush and with the stone submerged in 100 mL of NFW. The obtained biofilm suspension was processed as described for pebble samples.
Microalgal isolates were screened for their ICNO3 storage capacity in laboratory incubations. Twenty isolates from five microalgal classes for which published ICNO3 data were not available or very sparse and six diatom isolates were studied (Supplementary Table 4 and Supplementary Data 4). The microalgal isolates were grown in F/2 or L1 media in a light:dark cycle of 16:8 h and at 4 or 15 °C. For ICNO3 determination, 20 mL of cell suspension of each isolate was centrifuged at 500 × g for 10 min, the spent growth media potentially still containing nitrate discarded, and the cell pellet resuspended in 20 mL 0.2-µm-filtered NFW amended with 25 µmol L−1 NaNO3. The cell suspension was equally distributed between four transparent 15-mL sample tubes, incubated for a full light:dark cycle of 16:8 h, and harvested at 9 a.m. local time, which was 3 h after the start of the light period. The cell suspensions were thoroughly mixed and a 1-mL subsample was taken and fixed with formaldehyde (1% final concentration) for later cell counting and cell volume determination. The remaining sample volume was used for ICNO3 determination and pigment analysis.
Intracellular nitrate determination
For ICNO3 extraction from water column samples, 5 mL of deionized water was added to the filter in the sample tube before it was snap-frozen in liquid nitrogen. To further promote the destruction of microbial cells on the filter, the tube was transferred to a water bath kept at 80–90 °C until the sample was completely thawed. This freeze-thaw cycle was repeated a second time55. The extracted sample was centrifuged at 3000 × g for 5 min and 1 mL of the supernatant was stored at −20 °C for later analysis.
For ICNO3 and porewater nitrate (PWNO3) extraction from sediment, mat, and biofilm samples, 3 mL of NFW was added to the sample, vigorously mixed, and centrifuged at 3000 × g for 5 min. From the clear supernatant, 1 mL was stored at −20 °C for later PWNO3 analysis. From the remaining sample, ICNO3 was extracted by freeze-thaw cycling.
For ICNO3 extraction from microalgal isolates, the cell suspensions in three replicate tubes were centrifuged at 500 × g for 10 min and 1 mL of the supernatant was taken for FDNO3 analysis. From the remaining sample, ICNO3 was extracted by freeze-thaw cycling. The ICNO3 concentration inside microalgal cells was calculated from the difference between ICNO3 and FDNO3 concentrations, the cell abundance, and the cell volume of the respective microalgal isolate.Nitrate concentrations were measured spectrophotometrically using the protocol by56. This two-step reaction assay combines the Griess reaction for nitrite determination57 and the VCl3 reduction method for NOx (i.e., nitrate plus nitrite) determination58. Since nitrite concentrations were generally negligible, NOx concentrations are reported here as nitrate concentrations.
Total ICNO3 contents of water column samples were calculated from the nitrate concentration in the filter extract corrected for FDNO3 in the water-soaked filter and the volume of filtered water. Total ICNO3 contents of sediment, mat, and biofilm samples were calculated as the difference in nitrate contents before (PWNO3) and after the extraction step (PWNO3 + ICNO3).
The relative contribution of ICNO3 to total nitrate in each sample was calculated as ICNO3/(ICNO3 + FDNO3) for water column samples and as ICNO3/(ICNO3 + PWNO3) for sediment, mat, and biofilm samples and expressed in percent.
Photopigment determination
Photopigments were extracted by adding 5 mL of ice-cold, HPLC-grade acetone:ethanol (1:1) to the sample and keeping it at −20 °C for 24 h during which it was repeatedly shaken by hand. The extracted sample was centrifuged at 3000 × g for 5 min and 1.5 mL of the supernatant was filtered through a 0.45 µm Versapor® membrane (Acrodisc®, Gelman Laboratory) into 2-mL glass vials with a septum cap. Samples were kept in darkness and on ice during extraction and filtration. The extracted pigments were separated and analyzed on a Waters UPLC H-Class system (Waters, Milford, USA) equipped with an Acquity UPLC BEH C18 column (1.7 µm, 2.1 × 150 mm) (Waters, Milford, USA). Solvent gradients, column temperature, and flow rate were as described in ref. 59. In the chromatograms, ten different photopigments were identified according to their specific retention time and absorption spectra and the respective peaks were integrated. Calibrations were made with serially diluted stock solutions of the respective photopigment (DHI, Denmark).
The water-soluble photopigment phycocyanin (a cyanobacteria marker pigment) was extracted by adding 5 mL of lysozyme (15 g L−1 in 65 mM phosphate buffer adjusted to pH 8.2) to the sample and keeping it at 37 °C for 2 h during which it was repeatedly shaken by hand. The lysed sample was centrifuged at 3000 × g for 5 min and 1.5 mL of the clear supernatant was transferred into a 2-mL microcuvette and analyzed spectrophotometrically according to60. The obtained phycocyanin concentrations in g L−1 were recalculated to molar concentrations using the molar mass of phycocyanin of 30 kg mol−1 61.
Diatom abundance and biovolume
For a subset of 6–8 samples from each habitat type and three replicates each, diatom cells were counted and sized by microscopy. Samples were fixed in formaldehyde (1% final concentration) at 4 °C for 24 h. The fixed samples were diluted 100–2000× with 0.2-µm-filtered NFW and vigorously mixed for 2 min. Aliquots of the resulting cell suspension were used for cell counting in a Fuchs-Rosenthal chamber by phase-contrast microscopy12. For each sample and replicate, 140–570 diatom cells were counted, and the cell numbers obtained were recalculated to diatom abundance per unit volume of sample. For biovolume determination, 10–20 light micrographs were taken for each sample and replicate. Length and width of 100–200 individual cells per sample were measured with the image-processing program ImageJ (W.S. Rasband, NIH, Bethesda, U.S.A.). The cell volume of different diatom morphotypes was approximated using equations for different geometric shapes62. The total diatom biovolume per unit volume of sample was calculated from the diatom abundance and the average cell volume determined for each sample.
Diatom community composition
DNA extraction and PCR
For a selected subset of 53 samples representing all habitat types, the diatom community composition was analyzed by 18S rRNA gene sequencing. Water column samples (0.25–1 L) were filtered through 0.22-µm nylon membrane filters (Montamil®, Frisenette ApS, Denmark), from which DNA was extracted using the Allprep DNA/RNA Minikit (Qiagen No. 80204, Germany). The DNA of sediment, mat, and biofilm samples (0.05–5 mL) was extracted using the DNeasy PowerLyzer PowerSoil Kit (Qiagen No. 12855, Germany).
Three rounds of PCR were used to prepare the samples for 18S rRNA gene sequencing; first, the 18S rRNA gene was amplified using the diatom-targeted primer pair D512F/D978R63. The amplification was run in 25 μL reactions containing 2 μL template DNA, 12.5 μL 2X KAPA HiFi Hotstart Readymix (KAPA Biosystems), 0.5 μL bovine serum albumin (10 mg mL−1), 9 μL dH2O and 0.5 μL of each primer (10 pmol μL−1). Cycling conditions were: 95 °C for 3 min, followed by 35 cycles of 95 °C for 45 s, 55 °C for 30 s and 72 °C for 20 s, with a final elongation at 72 °C for 5 min. In the second round of PCR, Illumina adapters were added by using primers carrying the forward and reverse adapter overhang sequences at their 5’ ends. The adapter sequence information is provided in Illumina’s 16S metagenomic sequencing library preparation protocol (Illumina). The reaction conditions consisted of 12.5 μL KAPA HiFi Hotstart Readymix, 2.5 μL of purified PCR product, and 0.5 μL of each primer. Cycling conditions were: 95 °C for 3 min, 10 cycles of 95 °C for 30 s, 55 °C for 30 s, and 72 °C for 30 s, with a final elongation at 72 °C for 5 min. The PCR products from the second PCR were purified with the Agencourt AMPure XP PCR Purification system (Beckman Coulter). Subsequently, indices were added in the third round of PCR using the same conditions as the second PCR, except with only 8 PCR cycles. Products were purified with AMPure XP beads and paired-end sequenced in 2 × 301 cycles on an Illumina MiSeq Desktop sequencer using the MiSeq v3 Reagent Kit according to standard protocols. Samples were sequenced in two separate MiSeq runs.
18S rRNA gene sequence analysis
All analyses were run in R version 4.04. Trimming of adapter and PCR primers were done with cutadapt v0.1.164. The R package “dada2” v. 1.18.065 was used for sequence filtering, denoising, paired-end merging and taxonomic classification. Samples from the two MiSeq runs were filtered and denoised separately to obtain an individual error model for each run with the DADA algorithm. After denoising, the two MiSeq runs were merged for detecting and removing chimeras. Finally, the amplicon sequence variants (ASVs) were taxonomically assigned using the Silva SSU reference database no. 13866. For ASVs with a relative abundance >0.15%, the taxonomic assignment was manually corrected at order and class level according to the taxonomies presented in refs. 67,68, respectively, performing BLAST and Taxonomy Browser searches (NCBI). Four ASVs that were predominantly found in negative PCR controls, but only in low abundance in actual samples, were removed from subsequent analyses.
Statistical analysis
Spearman’s rank correlation coefficients were calculated for monotonic relationships between two variables. To test whether the correlation coefficients were significantly different from zero, two-sided Student’s t-tests with n – 2 degrees of freedom were used. Community analysis of the classified ASVs was done using the R packages phyloseq v1.34.069 and ampvis2 v2.6.5, and all plots were made with ggplot2 v3.3.370. We used vegan v2.5.771 for calculating analysis of similarity (ANOSIM) and permutational multivariate analysis of variance (PERMANOVA) tests.
Reporting summary
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
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Acknowledgements
This study was supported by grants from the German Science Foundation awarded to P.S. (STI 202/6) and A.K. (KA 3187/2-1), by the Max Planck Society (Germany), by the European Union’s Seventh Framework Program for Research and Technological Development including Demonstration Activities awarded to Aarhus Institute of Advanced Studies (Grant agreement No. 609033), and by the European Union’s Horizon 2020 Research and Innovation Program (Grant agreement No. 669947) and the Danish National Research Foundation through the Danish Center for Hadal Research (DNRF145) and the Danish Center for Electromicrobiology (DNRF136). Sample collection was supported by HYDRA Marine Sciences GmbH (Bühl, Germany), PULZ im Rieselfeld (Freiburg, Germany), and MACRORE (European Union’s Horizon 2020 Research and Innovation Program (Grant agreement No. 654182; ENVRIplus)). We would like to thank the following colleagues and friends for their kind and active engagement in collecting samples: Kathrin Graffe, Nicole Ehlers, Tillman Keller, Jutta Drees, Jasper Dammann, Fabian Seredszus, Stefanie Tenten, Georg Supp, Herdís G. R. Steinsdóttir, Mon O. Yee, Hans F. Hansen, Michael W. Hansen, Lorenzo Rovelli, Irina-Elena Antonescu, Irene Olivé, Kelsey I. Miller, Marco Bonaglia, Artur Fink, Miriam Weber, Frank Wenzhöfer, Marit van Erk, and Elisa Merz. Microalgal isolates were kindly provided by Olga Kourtchenko (Gothenburg University Marine Algae Culture Collection, GUMACC), Tina Trautmann, Jan-Hendrik Hehemann, Daniel Doherty, and Hans F. Hansen. The technical support by Erik Laursen, Susanne Nielsen, Anja Niclas, and Gaby Eickert-Groetzschel is acknowledged.
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P.S. designed the study, collected and analyzed samples, analyzed and interpreted data, and conceived and wrote the manuscript. A.K., A.S.C.H.R., R.N.G., and D.B. interpreted data and wrote the manuscript. C.S., M.B.L., and A.G. analyzed samples and data and contributed to manuscript preparation and editing. R.A., M.A., K.A., S.B., J.D., B.F.C., E.G.R., M.H., U.J., A.M., M.M., R.P., T.S.T., A.S., and D.S. collected and processed samples and contributed to manuscript preparation and editing.
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Stief, P., Schauberger, C., Lund, M.B. et al. Intracellular nitrate storage by diatoms can be an important nitrogen pool in freshwater and marine ecosystems. Commun Earth Environ 3, 154 (2022). https://doi.org/10.1038/s43247-022-00485-8
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DOI: https://doi.org/10.1038/s43247-022-00485-8
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