The rate and fate of N2 and C fixation by marine diatom-diazotroph symbioses

Abundances of N₂ fixing symbioses in the WTNA

To date, the various marine symbiotic diatoms are notoriously understudied, and hence our understanding of their abundances and distribution patterns is limited [7]. In general, these symbiotic populations are capable of forming expansive blooms, but largely co-occur at low densities in tropical and subtropical waters with a few rare reports in temperate waters [26,27,28,29, 39,40,41,42]. The Rhizosolenia-Richelia symbioses have been more commonly reported in the North Pacific gyre [26, 27, 31], and the western tropical North Atlantic (WTNA) near the Amazon and Orinoco River plumes is an area where widespread blooms of the H. hauckii-Richelia symbioses are consistently recorded [28, 29, 42,43,44,45,46,47].

In the summer of 2010, bloom densities (10⁵−10⁶ cells L⁻¹) of the H. hauckii-Richelia symbioses were encountered at multiple stations with mesohaline (30–35 PSU) surface salinities (Supplementary Table 1). The R. clevei-Richelia symbioses were less abundant (2–30 cells L⁻¹). Similar densities of H. hauckii-Richelia have been reported in the WTNA during spring (April–May) and summer seasons (June–July) (28–29; 46). In fall 2011, less dense symbiotic populations (0–50 cells L⁻¹) were observed, and the dominant symbioses was the larger cell diameter (30–50 µm) H. membranaceus associated with Richelia. Previous observations of H. membranaeus-Richelia in this region are limited and reported as total cells (i.e., 12-218 cells) and highest numbers recorded in Aug–Sept in waters near the Bahama Islands [43]. On the other hand, Rhizosolenia-Richelia are even less reported in the WTNA, and most studies by quantitative PCR assays based on the nifH gene (for nitrogenase enzyme for N₂ fixation) of the symbiont (44; 46–7). Unlike qPCR which cannot resolve if the populations are symbiotic or active for N₂ fixation, the densities and activity reported here represent quantitative counts and measures of activity for symbiotic Rhizosolenia.

The WTNA is largely influenced by both riverine and atmospheric dust deposition (e.g., Saharan dust) [48], including the silica necessary for the host diatom frustules, and trace metals (e.g., iron) necessary for photosynthesis by both partners and the nitrogenase enzyme (for N₂ fixation) of the symbiont. We observed similar hydrographic conditions (i.e., low to immeasurable concentrations of dissolved N, sufficient concentrations of dissolved inorganic P and silicates, and variable surface salinities; 22; 28–29; 40–47) as reported earlier that favor high densities of H. hauckii-Richelia blooms. Unfortunately our data is too sparse to determine if these conditions are in fact priming and favoring the observed blooms of the H.hauckii-Richelia symbioses in summer 2010, and to a lesser extent in the Fall 2011.

A biometric relationship between C and N activity and host biovolume

The diatom-Richelia symbioses are considered highly host specific [10, 11], however, the driver of the specificity between partners remains unknown. We initially hypothesized that host selectivity could be related to the N₂ fixation capacity of the symbiont. Moreover, it would be expected that the larger H. membranaceus and R. clevei hosts which are ~2–2.5 and 3.5–5 times, respectively, larger in cell dimensions than the H. hauckii cells would have higher N requirements (Supplementary Table 2). In fact, recently it was reported that the filament length of Richelia is positively correlated with the diameter of their respective hosts [22]. Thus, to determine if there is also a size dependent relationship between activity and cell biovolume, the enrichment of both ¹⁵N and ¹³C measured by SIMS was plotted as a function of symbiotic cell biovolume.

Given the long incubation times (12 h) and previous work [32] that show fixation and transfer of reduced N to the host is rapid (i.e., within 30 min), we expected most if not all of the reduced N, or enrichment of ¹⁵N, to be transferred to the host diatom during the experiment (Fig. 1). Therefore, we measured and report the enrichment for the whole symbiotic cell, rather than the enrichment in the individual partners (Supplementary Table 2; Fig. 2). The enrichment of both ¹³C/¹²C and ¹⁵N/¹⁴N was significantly higher in the larger H. membranaceus-Richelia cells (atom % ¹³C: 1.5628–2.0500; atom % ¹⁵N: 0.8645–1.0200) than the enrichment measured in the smaller H. hauckii-Richelia cells (atom % ¹³C: 1.0700–1.3078; atom % ¹⁵N: 0.3642–0.7925) (Fig. 2) (¹³C, Mann–Whitney p = 0.009; ¹⁵N, Mann–Whitney p < 0.001). In addition, the higher enrichment also corresponded to higher rates of N₂ fixation (e.g., 12.6–28.4 fmol N cell⁻¹ h⁻¹ for H. membranaceus-Richelia compared to 0.1-4.29 fmol N cell⁻¹ h⁻¹ for H. hauckii-Richelia; Supplementary Table 2) and C fixation (219–685 fmol C cell⁻¹ h⁻¹ compared to 2.74–59.8 fmol C cell⁻¹ h⁻¹, respectively; Supplementary Table 2).

Similar findings of higher ¹⁵N enrichment and N₂ fixation rates were also reported by Martínez-Pérez et al. [5], when comparing the larger N₂ fixing UCYN-A2-haptophyte symbioses to the smaller UCYN-A1-haptophyte symbiotic cells. Here, however, pooling all the SIMS measurements and biovolume estimates for the three different symbioses, we present a robust relationship between enrichment (atom % ¹³C or atom % ¹⁵N) and biovolume for both C and N (Fig. 2). It suggests that the larger size demands a higher activity for both N and C, which would be expected from allometric theory. Here, allometry refers to the relationship that biological processes scale by body-size [49].

Light dependence of N₂ fixation by diatom-Richelia symbioses

In order to determine if metabolic activity for the various symbioses is light dependent, the stable isotope incubation experiments were set up under simulated light conditions of the water column (i.e., 0.1−100% incidence surface light). Activity rates are reported for both bulk (see below) and single cells.

Although the data is more limited for the two larger symbiotic diatoms: H. membranaceus and R.clevei-Richelia symbioses, both follow the expected trend for phototrophic organisms to have higher C fixation in cells collected nearer to the surface and incubated under higher light intensities [50]. For example, the H. membranaceus-Richelia symbioses collected nearer to the surface had higher ¹³C enrichment and rates of C fixation than those collected from a deeper depth (25 m). However, ¹⁵N enrichment and corresponding N₂ fixation rates for the H. membranaceus-Richelia symbioses were similar regardless of their depth of collection. The two R. clevei–Richelia symbioses measured high and comparable enrichment of ¹³C and to a lesser extent ¹⁵N (atom % ¹³C = 1.2053 and 1.2346; atom % ¹⁵N = 0.7543 and 0.4480) to that measured in the H. membranaceus-Richelia symbioses (Supplementary Table 2).

Given the higher number of measurements for the H. hauckii-Richelia symbioses at multiple light levels, we plotted the individual rates of N₂ and C fixation as a function of photosynthetically active radiation (PAR) measured at the time of collection (Fig. 3). A hyperbolic tangent model [33] fit well for the N₂ fixation rate (adjusted R² = 0.56). This indicates that N₂ fixation followed a light dependence and saturation kinetics of maximum N₂ fixation (N₂fix_max) and enrichment (¹⁵N_max) at the highest irradiances. Unexpectedly, the light response curve for C fixation could not be fit with the same or any saturation model. In fact, the ¹³C enrichment and corresponding estimated C fixation rates were highly variable (Supplementary Table 2). N₂ fixation in heterocystous cyanobacteria is fueled by C fixation and thus one expects parallel trends in activity for C and N₂ fixation [51]. We hypothesize that the fixed C of both partners is partitioned into growth and respiration to fuel other metabolic activities (i.e., N₂ fixation), and resulted in highly variable enrichment patterns. The estimated N- and C- based growth rates (see below) indicated that the cells were also growing at different rates and additionally could influence the observed variation. Further experimentation would be required to test these hypotheses and both require shorter incubations and multiple time points than presented here.

Similar lab experiments for determining the light dependency on N₂ fixation have been reported in the facultative Calothrix SC01 strain that forms symbioses with Chaetoceros diatoms, in a few older lab studies of R. clevei-Richelia and a recent report on H. hauckii-Richelia enrichment cultures [21, 33, 52, 53]. In these earlier works, the same light dependent N₂ fixation is shown. The strong light dependent activity shown here, however, was somewhat surprising given that measures were derived from field populations, unlike these other experiments which were done in controlled laboratory settings and additionally Calothrix SC01 was growing asymbiotic (21; 52–53). Furthermore, our measurements are pooled from three different stations with experiments that span approximately four weeks and two stations ~700 km apart (Supplementary Fig. 2). The wild populations studied here were likely different subpopulations and/or at different stages of their growth cycle, and still the response in their N₂ activity fit well to the curve. In fact, N and C-based growth rates (Supplementary Table 2) identified significant differences in estimated growth rates between the cells collected at the three stations (C-based growth rate: Kruskal–Wallis, p = 0.09; N-based growth rate: Kruskal–Wallis, p = 0.004). These growth rates were calculated from the SIMS derived cellular ¹³C/¹⁵N enrichment and initial N/C content based on biovolume (see Supplementary Materials). Moreover, both N-based and C-based growth was depth dependent, where cells nearer to the surface (stations 2 and 19) had higher estimated growth rates (Supplementary Table 2; Kruskal–Wallis, p = 0.002 and p = 0.008, respectively). Hence, genetically distinct symbiotic strains have a similarly robust N₂ fixing activity response to light despite varying growth states.

Increased N₂ fixation by symbiotic cyanobacteria is largely under the host control in many terrestrial symbioses (17; 54). A similar scenario was suggested in the early work of applying NanoSIMS to field collected symbiotic diatoms because higher rates of N₂ fixation were reported in the symbiotic populations compared to asymbiotic ones [32]. However, in terrestrial symbiotic examples, the number of symbionts per host is dramatically different. Typically, one to two Richelia filaments associate with a Hemiaulus spp. host, whereas in plants, a symbiotic chamber houses 100’s to 1000’s of symbionts [17, 54]. Moreover, in most terrestrial examples, the number of heterocysts per symbiont filament increases after establishment [17, 18], however in Richelia (and Calothrix), the heterocysts are single and terminal. Thus, Richelia sustains a high N₂ fixing capacity by maintaining short filaments and a high heterocyst to vegetative cell ratio [23]. It is plausible and hypothesized that Richelia acquires energy from their hosts, e.g., in the form of C substrates. This would be a particularly attractive strategy for the internal Richelia symbionts of Hemiaulus spp. because they reside in close proximity to the host photosynthetic machinery [22].

Reduced C and N₂ fixation when host is inhibited

To better understand the potential control and role of C fixation mediated by the host diatoms, ¹⁵N₂ and ¹³C-bicarbonate incubation experiments were treated with cycloheximide, a eukaryotic cytosolic protein translation inhibitor. Here, the enrichment of ¹⁵N and ¹³C were visualized and measured by SIMS in both the symbiont and the host separately. DCMU (3-(3,4-dichlorophenyl)−1, 1-dimethylurea) and chloramphenicol are other common inhibitors, however, both have been shown to inhibit several cyanobacteria strains, including heterocystous types [55]. Cycloheximide is not inhibitory to cyanobacteria and often applied as a cultivation strategy to avoid enrichment of eukaryotes [56]. Earlier lab studies on diatoms (and other eukaryotic algae, i.e., dinoflagellates) have shown that at similar concentrations (0.1–10 μg/ml) of cycloheximide, as used here, can reduce key metabolic processes, including photosynthesis and energy generation [57,58,59]. Hence, cycloheximide seemed to be an attractive and appropriate inhibitor for shutting down the diatom host photosynthesis, because the cyanobacterial symbiont would not be influenced.

The enrichment of both ¹⁵N/¹⁴N and ¹³C/¹²C was significantly reduced in the inhibited cells (Fig. 4; Supplementary Table 3; T test; p < 0.03). The decreased enrichment was measured in the host cells and only the heterocysts of Richelia filaments compared to the respective control cells. In fact, in the inhibited cells, the enrichment for ¹⁵N/¹⁴N was clearly localized to the symbiont filament, whereas ¹³C/¹²C was reduced overall without an apparent localization (Fig. 4B). The average decrease in ¹³C/¹²C and ¹⁵N/¹⁴N enrichment was one to two times, or 16–17% and 36–46% reduced in the hosts and heterocysts of inhibited cells, respectively. The reduction in enrichment resulted in a 66–78% and 76% reduction in estimated rates of C and N₂ fixation, respectively, when the cells were inhibited. These results are congruent with studies on terrestrial-based symbioses, where the CO₂ (and NH₄⁺) assimilation rates are measurably depressed in symbiotic cyanobacteria (17–18; 54). Increased N₂ fixation is expected when the host is present, as it is the basis of the partnership and has been previously reported [32]. Moreover, it could be that the increased C fixation by the host leads to C:N ratio favoring increased N₂ fixation. The latter hypothesis remains to be tested.

The reduction in both C and N₂ fixation in inhibited cells indicates that the host is required, and likely controls the symbiont’s metabolism. In heterocystous cyanobacteria, there is a coordinated effort between vegetative cells and heterocysts to exchange energy, organic C and fixed N. Reduced C is rapidly transported to the heterocysts from vegetative cells, and N₂ fixed in heterocysts is exported to vegetative cells [60,61,62]. The ¹³C/¹²C of vegetative cells was similar between inhibited and control cells (t test, p = 0.08); hence C fixation by Richelia persisted in the absence of the host.

The ¹³C/¹²C in terminal heterocysts of untreated cells was, however, significantly higher than the inhibited cells indicating that the host must also contribute to the increased ¹³C/¹²C enrichment measured in the heterocysts. One plausible scenario could be that reduced C substrates derived from the host photosynthesis are transferred to the Richelia vegetative cells and subsequently transported to the heterocysts. Encoded in the Richelia RintHH01 genome is an invertase (InvB), an enzyme used to irreversibly split sugar [23]. In Anabaena sp. 7120, a related heterocystous cyanobacteria, InvB is heterocyst specific and sucrose functions as a reduced C substrate for heterocysts that is transported from vegetative cells [63, 64]. Unfortunately, our incubation period was too long to clearly resolve the transfer of C from the host to symbiont. Shorter time incubations akin to a pulse-chase experiments would be desirable for estimating the gradual transfer, or perhaps a dual label incubation experiment of ¹⁴C- and ¹³C-labels combined with NanoSIMS to trace the fate of labeled carbon [65].

Using the quantitative measures by SIMS in the control cells, we estimate that 8–22% of assimilated ¹³C in the symbiont is derived from the host, and 78–91% of the host ¹⁵N is derived from the symbiont. Our estimates are consistent with a recent cellular model of Hemiaulus-Richelia that reported 25% of fixed C in the symbiont is derived from the host, and 82% of fixed N in the host is from the symbiont [30]. In the cellular model, however, input values were largely derived from literature values of biovolume, biovolume to C relationships, and assumed stoichiometry [30]. The variation we measured here in the host derived C (8–22%) could be attributed to our long incubation times and thus “bottle effects” [66]. It is also likely that some fixed C was respired in order to fuel N₂ fixation. Moreover, we have little knowledge of the life history of the cells prior to collection, hence we cannot discount that the population of cells were in varying growth states and contributed to the observed variation. In fact, the estimated growth (both N- and C- based) was higher in the control cells compared to the inhibited cells (p = 0.009). The latter is expected because activity was largely inhibited in the treated cells. However, in both the control and inhibited cells, the symbiont (vegetative cells) had higher estimated growth than the respective host cell (Supplementary Table 3; p = 0.02). In plant-cyanobacteria symbioses, slower growth is reported in the symbionts [18]; hence our results are unexpected and suggest uncoordinated growth cycles between the partners.

Impact of symbiotic diatoms on bulk N₂ and C fixation

Studies on rate and fate of N₂ and C fixation by the symbiotic diatoms are fewer compared to investigations on other N₂ fixers i.e., Trichodesmium, unicellular diazotrophs. In fact, searching the open world data archive PANGEA for datasets reported for the latter groups, and Richelia finds 1564, 126, and 35 datasets, respectively. Thus, the data presented here and motivation for the work was to contribute new rate information for these biogeochemical relevant yet under-reported populations.

To date, there are a few estimates of N₂ fixation by the Hemiaulus-Richelia populations from the same WTNA region. These earlier studies attribute a large source of new N to the photic zone derived from these symbiotic diatom populations [28, 29, 67, 68]. Moreover, this new N contributes significantly to C export in the region [29, 67, 68] and likewise in other regions where diatom-Richelia symbioses persist (i.e., N. Pacific: 26–27; 31). However, the previous measurements of N₂ fixation in the WTNA were indirect because acetylene reduction assays were used and applied to cell concentrates (i.e., plankton “slurries”). Thus, the individual cell activity cannot be distinguished. Combining the single cell measurements by SIMS with cell abundances by microscopy (Supplementary Table 1, 2) we could more accurately estimate the fraction of N₂-fixation attributable to the symbiotic populations.

It is important to note that although our measures are on individual cells, the populations were incubated in whole water bottle experiments. Hence, we cannot discount that some enrichment in the individual cells is derived from the transfer of reduced substrates from co-occurring populations of other N₂ fixers. In 2010, however, incubations were performed at stations almost entirely dominated by H.hauckii-Richelia cells (>10⁴ cells L⁻¹), whereas in 2011, biomass in general was low, and observations of other N₂ fixers i.e., Trichodesmium spp., Crocosphaera watsonii, were rare in general and in particular at the stations assayed for N₂ and C fixation measurements.

In 2010, we estimate that between 12–53% of the bulk N₂ fixation was accounted for by the H. hauckii-Richelia populations. Although the symbiotic cell densities at depth (11–21 m) of station 2 were slightly higher than the populations in the surface (2–4 m), the surface H. hauckii-Richelia cells had higher individual N₂ fixation rates (0.86–3.58 fmol N cell⁻¹ h⁻¹) and therefore made a larger contribution (53%) to the total fixed N₂. Meanwhile, cell densities at station 25 (10⁵ cells L⁻¹) were similar to station 2, however rates of N₂ fixation (0.10–0.35 fmol N cell⁻¹ h⁻¹) were reduced, and therefore the estimated contribution (12–17%) to bulk N₂ fixation was lower at station 25. The latter findings are directly relevant to models that use abundance estimates (including mRNA abundances of nifH genes for nitrogenase), rather than in situ activity to estimate new N contributing to N budgets [e.g., 69, 70].

The contribution to total C fixation by the symbioses was estimated to be far less than their N contribution. For example, H. hauckii-Richelia accounts for only 2–5% of bulk C fixation at stations 2 and 25, and is comparatively less than the contribution of 12–53% to bulk N₂ fixation. Observations of the H. hauckii-Richelia populations during the bloom reported that one or both partners possessed variable cell integrity [46]. For example, long diatom chains (8–12 symbiotic cells per chain and >50 symbiotic cells in a chain) were reported at station 2 with fully intact symbiotic Richelia filaments (2–3 vegetative cells and terminal heterocyst), and at station 25 chains were short (1–2 symbiotic cells) and associated with short Richelia filaments (only terminal heterocyst). Moreover, the symbiotic H. hauckii hosts possessed poor chloroplast auto-fluorescence at station 25 [46]. Given that the cells selected for NanoSIMS were largely single cells, rather than chains, we suspect that these cells were in a less than optimal cell state, which was also reflected in the low ¹³C/¹²C enrichment ratios and low estimated C-based growth rates (0.30–57 div d⁻¹). These are particularly reduced compared to the growth rates recently reported for enrichment cultures of H. hauckii-Richelia (0.74–93 div d^−1§) (Supplementary Table 2) [33].

In 2011, higher cellular N₂ fixation rates (15.4–27.2 fmols N cell⁻¹ h⁻¹) were measured for the large cell diameter H. membranaceus-Richelia, symbioses. Despite high rates of fixation, cell abundances were low (4–19 cells L⁻¹), and resulted in a low overall contribution of the symbiotic diatoms to the whole water N₂ (>1%) and C-fixation (>0.01%). The estimated C-based growth rates for H. membranaceus were high (1.9–3.5 div d⁻¹), whereas estimated N-based growth rates (0.3–4 div d⁻¹) were lower than previously published (33; 52–53). Hence the populations in 2011 were likely in a pre-bloom condition given the low cell densities.

Estimating symbiotically derived reduced N to surface ocean

To date, determining the fate of the newly fixed N from these highly active but fragile symbiotic populations has been difficult. Thus, we attempted to estimate the excess N fixed and potentially available for release to the surround by using the numerous single cell-specific rates of N₂ fixation determined by SIMS on the Hemiaulus spp.-Richelia symbioses (Supplementary Materials). Because the populations form chains during blooms and additionally sink, we calculated the size-dependent sinking rates for both single cells and chains (>50 cells). Initially we hypothesized that sinking rates of the symbiotic associations would be more rapid than the N excretion rates, such that most newly fixed N would contribute less to the upper water column (sunlit).

The sinking velocities were plotted (Fig. 5) as a function of cell radius at a range (min, max) of densities and included two different form resistances (∅ = 0.3 and 1.5). As expected, the combination of form resistance and density has a large impact on the sinking velocity. For example, a H. hauckii cell of similar radius (10 μm) and density (3300 kg m⁻³) but higher form resistance (0.3 vs. 1.5) sinks twice as fast at the lower form resistance (Fig. 5). This points to chain formation (e.g., increased form resistance) as a potential ecological adaptation to reduce sinking rates. Recently, colony formation was identified as an important phenotypic trait that could be traced back ancestrally amongst both free-living and symbiotic diatoms that presumably functions for maintaining buoyancy and enhancing light capture [22].

The concentration of fixed N surrounding a H. hauckii and H. membranaceus cell were modeled (Supplementary Materials; Supplementary Table 4; Fig. 6). First, the cellular N requirement (Q_N, mol N cell⁻¹) for a cell of known volume, V, as per the allometric formulation of Menden-Deuer and Lessard [71] is calculated by the following.

Volume calculations assume a cylindrical shape; whereas exudation assumes an equivalent spherical volume. Then, using published growth rates of 0.4 d⁻¹ and 0.68 d⁻¹ for the symbioses [52, 53], N uptake rate (V_N) necessary to sustain the Q_N was determined. N loss was assumed to be a constant fraction (f) of the V_N; this fraction was assumed to be 7.5% and 11% for H. hauckii and H. membranaceus, respectively, or the estimated excess N which was fixed given the assumed growth rate [31]. The excretion rate (E_N) of the individual cells was then calculated as

The concentration of fixed N surrounding the cell (C_r) was iteratively calculated by the following:

The concentric radius (r_x) as per Kiørboe [72] uses a diffusivity of N assumed to be 1.860 × 10⁻⁵ cm² sec⁻¹ and the background concentration of N (C_i) is assumed to be negligible. Figure 5 presents the results for the two symbioses: H. membranaceus and H. hauckii at the two growth rates and as chains or singlets. Mean sinking rates for cells with a high form resistance (e.g., chains) are <10 m d⁻¹ (Fig. 5). Simplified exudation calculations assuming no motility suggest that the phycosphere surrounding individual cells in these chains would range from ~3–70 nmol N L⁻¹ at the cell surface and ~1–25 nmol N L⁻¹ at a distance of twice the equivalent spherical radius (Fig. 6). Sinking would deform these plumes and increase flux away from the cellular boundary layer so these are considered maximal near-cell concentrations of exudate. More complex mechanistic models of diffusive fluid dynamics (e.g., x, y, z) would be needed to simulate deformation of exudate plumes as a function of sinking speeds and the flow field (laminar or turbulent) [73,74,75].

In oligotrophic regimes such as WTNA, where N concentrations in the surface mixed layer are often below detection, the symbiotically derived reduced N serves as an important source of new N to the surface ocean. Based on our calculations, we would predict the phycosphere surrounding these symbioses to be N-rich, and given sinking speeds <10 m d⁻¹, these exudates retain the availability of limiting nutrients in the surface waters and fuel regional primary production rather than contributing to the mesopelagic nutrient inventory.

Conclusions

Despite widespread distributions and occasional large-scale blooms, diatom-Richelia symbioses have been largely understudied. Our understanding of their metabolic activities and partner interactions has been limited to N₂ fixation and the exchange of fixed N, respectively. Here, we report both the N₂ and C fixation activity of single cells from three different diatom-Richelia symbiotic populations collected and assayed in the wild. A robust biometric relationship was identified where larger symbiotic cells have higher activity. N₂ fixation by one symbiotic population appears to be light dependent, whereas unexpectedly C fixation is highly variable and independent of light. Inhibitor experiments designed to shut down the host photosynthesis and communication resulted in depressed C and N₂ fixation activity suggesting that the hosts are the primary C fixing partner and likely control their symbionts N₂ fixing activity. Single cell rates and estimated sinking velocities were combined with a simplified model to predict that most of the fixed N is released in the upper water column by the symbiotic diatoms. In summary, the observations reported here for both abundance and in situ activity contributes to an improved understanding of symbiotic diatom distribution, ecology, and contribution to N/C cycling.

Source: Ecology - nature.com