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PLoS One. 2013; 8(12): e82605.
Published online Dec 2, 2013. doi:  10.1371/journal.pone.0082605
PMCID: PMC3846789

Response of the Ubiquitous Pelagic Diatom Thalassiosira weissflogii to Darkness and Anoxia

Adrianna Ianora, Editor

Abstract

Thalassiosira weissflogii, an abundant, nitrate-storing, bloom-forming diatom in the world’s oceans, can use its intracellular nitrate pool for dissimilatory nitrate reduction to ammonium (DNRA) after sudden shifts to darkness and anoxia, most likely as a survival mechanism. T. weissflogii cells that stored 4 mM 15N-nitrate consumed 1.15 (±0.25) fmol NO3- cell-1 h-1 and simultaneously produced 1.57 (±0.21) fmol 15NH4+ cell-1 h-1 during the first 2 hours of dark/anoxic conditions. Ammonium produced from intracellular nitrate was excreted by the cells, indicating a dissimilatory rather than assimilatory pathway. Nitrite and the greenhouse gas nitrous oxide were produced at rates 2-3 orders of magnitude lower than the ammonium production rate. While DNRA activity was restricted to the first few hours of darkness and anoxia, the subsequent degradation of photopigments took weeks to months, supporting the earlier finding that diatoms resume photosynthesis even after extended exposure to darkness and anoxia. Considering the high global abundance of T. weissflogii, its production of ammonium and nitrous oxide might be of ecological importance for oceanic oxygen minimum zones and the atmosphere, respectively.

Introduction

Diatoms are a key group of the eukaryotic phytoplankton of the world’s oceans from polar to tropical latitudes. Pelagic diatoms form massive phytoplankton blooms [1] and may sink to the seafloor in vast abundances [2]. Diatoms are responsible for 40% of the marine primary production, or 20% of the Earth’s primary production [3,4]. Thus, they play a key role in the oceanic C-cycle and their productivity supports large-scale coastal fisheries [5]. Diatoms can also survive for decades buried deep within the dark, O2-depleted sediment layers at the seafloor, where neither photosynthesis nor aerobic respiration is possible [6,7]. The survival mechanism under these non-phototrophic conditions is still poorly understood. Only recently, the dissimilatory use of NO3- by the benthic diatom Amphora coffeaeformis was discovered as a possible survival mechanism in darkness and anoxia [8]. The study revealed that A. coffeaeformis stored NO3- intracellularly and used it for Dissimilatory Nitrate Reduction to Ammonium (DNRA; NO3- NO2- NH4+) after sudden exposure to darkness and anoxia. Briefly, dissimilatory NO3- reduction is an energy-generating pathway where NO3- is taken as electron acceptor instead of O2 in respiratory processes. It preferentially occurs in environments in which O2 is scarce or in which steep O2 gradients exist. In the marine realm, coastal sediments, oceanic Oxygen Minimum Zones (OMZs), and suspended aggregates (“marine snow”) are prominent (micro)environments characterized by O2 shortage (e.g.[9-12]). Besides DNRA, denitrification (NO3- NO2- NO N2O N2) and anammox (oxidation of NH4+ to N2 with NO2- as the electron acceptor) are important dissimilatory NO3- reduction pathways. Dissimilatory NO3- reduction has important implications for the marine N-cycle and is not least due to increasing use of synthetic fertilizers and subsequent pollution of rivers, estuaries, and coastal waters well studied (e.g. 13-17). However, our knowledge is almost exclusively based on prokaryotic studies; research on dissimilatory NO3- reduction by eukaryotes and its quantitative impact on marine N-cycling is still in its infancy. The seminal work on marine eukaryotes that dissimilatorily reduce NO3- was done by Risgaard-Petersen et al. [18]. The authors discovered that the foraminifer Globobulimina pseudospinescens store NO3- in large quantities, and use it for complete denitrification under anoxic conditions. In following studies on diverse benthic foraminifera and a few gromiida from different benthic habitats, denitrification capacity was found for all analyzed species that contained intracellular NO3- [19-21]. In some foraminifera, denitrification is likely carried out by endobionts [22]. The storage of NO3- might be a prerequisite for eukaryotes that can switch between O2 and NO3- respiration, because NO3- can be taken up and stored under favorable, oxic conditions for the usage in habitats that can be temporarily exposed to anoxic conditions.

So far, all marine eukaryotes that have been found to dissimilatorily reduce NO3- originate from benthic habitats in which anoxic conditions are common. This study addresses the response of the pelagic, NO3--storing diatom Thalassiosira weissflogii to darkness and anoxia with respect to dissimilatory NO3- reduction and stability of photopigments. Pelagic diatoms may be exposed to anoxic or hypoxic conditions in algal blooms, if O2 consumption by the community exceeds O2 production, e.g. at night. After the blooms, diatoms might also pass through the anoxic water layers of OMZs [12] and further sink towards the seafloor onto dark/ anoxic sediments [2,23]. The occurrence and viability of Thalassiosira species in marine sediments is indeed well known (e.g. 24-26). We hypothesize that a survival mechanism must exist that is energized by dissimilatory NO3- reduction. To test this hypothesis, we cultured an axenic T. weissflogii strain and followed the consumption of intracellularly stored 15NO3- after a sudden shift to dark/anoxic conditions as well as the production of end products, by-products, and intermediates of denitrification and DNRA. We further investigated the stability of photopigments after exposure to darkness and anoxia as an indicator of the dark survival potential of T. weissflogii.

Materials and Methods

Strain and Cultivation

An axenic strain of the marine pelagic diatom T. weissflogii (CCMP 1336) was obtained from the Provasoli-Guillard National Center for Marine Algae and Microbiota (NCMA; formerly CCMP). The diatoms were cultured in F/2 medium plus silicate [27] prepared with filtered (0.45 μm) and autoclaved North Sea seawater (salinity 35). The cultivation temperature was 15°C, the light:dark cycle was 10:14 h, and the light intensity was 160 μmol photons m−2 s−1. T. weissflogii was frequently checked for possible contaminations with bacteria by careful phase-contrast microscopy and by plating out subsamples of the cultures on nutrient agar plates. Additionally, all T. weissflogii cultures used in the experiments were checked by DAPI staining of cell suspensions immobilized on polycarbonate membrane filters (0.2 μm; Osmonics). A contamination of the diatom strain with prokaryotes was never detected.

Consumption and Production of Inorganic N-compounds in Dark/Anoxic versus Light/Oxic Conditions

The time courses of intracellular NO3- concentrations in T. weissflogii, and NO3- and NH4+ concentrations in the growth medium under dark/anoxic versus light/oxic conditions were followed in a non-labeling experiment and a 15N-stable isotope labeling experiment (see below). For the non-labeling experiment, the cells were washed with sterile NaCl (salinity 35) and centrifuged (10 min at 1000g) three times to remove NO3- from the medium, and transferred into NO3--free artificial seawater. The cell number was determined (see below), and the experiment was started by dividing the culture for (a) the dark/anoxic incubation, and (b) the light/oxic control. For the dark/anoxic incubation, 20 mL of the diatom suspension was transferred into a dark serum bottle (wrapped in aluminum foil), flushed with N2 for 20 min to remove O2, sealed with a gas-tight rubber stopper, and incubated at 15°C. For the light/oxic control, the culture was kept under light/oxic culture conditions (see above). At time intervals of 0, 1, 2, 3, 4, 5, 6, and 7 h, 2 mL diatom suspension each was taken and transferred into a sample tube for centrifugation (10 min at 1000g). To assure anoxia, the dark serum bottle was flushed with N2 after each sampling for 2 min. NO3- and (non-labeled) NH4+ were determined in the cell-free supernatant and the diatom pellet was used for measurements of intracellular NO3-.

NO3- was measured with an NOx analyzer connected to a reaction chamber (CLD 66s plus a Liquid NO Setup; EcoPhysics). In the reaction chamber, acidified VCl3 (0.1 M) reduces NO3- plus NO2- to NO at 90°C, which is then measured by a chemiluminescence detector [28]. If not noted differently, the results of the NO3- plus NO2- analyses are reported as NO3- concentrations throughout, because NO2- concentrations were << NO3- concentrations. For intracellular NO3- measurements, the diatom pellet was directly injected into the reaction chamber where cells burst and release the stored NO3-. Intracellular NO3- concentrations were calculated from the difference of NO3- concentrations in the medium and the cell pellet, the cell numbers in the pellet, and the average cell volume of 1.22 pL [8]. Cell numbers were counted in a Fuchs-Rosenthal counting chamber with phase-contrast microscopy at 400× magnification. The total cell number in the medium was used to calculate the total intracellular NO3- concentration per volume of medium from the cell-specific intracellular NO3- concentration. Ammonium was measured by photometric absorbance determination at λ = 640 nm with a Genesys 10S spectrophotometer (Thermo Scientific; USA) following the sodium-nitroprusside-catalyzed reaction of NH4+ ions with salicylate and hypochlorite [29].

Final Products of Dissimilatory Nitrate Reduction

The time courses of intracellular NO3- consumption and the possible products of dissimilatory NO3- reduction, i.e. NH4+ for DNRA and N2 for complete denitrification, were investigated with a 15N-stable isotope labeling experiment. Prior to the experiment, the (non-labeled) intracellular NO3- pools of T. weissflogii were depleted by a starvation procedure. The cells were separated from the NO3--containing culture medium via gentle centrifugation (10 min at 1000g), transferred into NO3--free artificial seawater [30], and exposed to dark/anoxic conditions for six days. After this pre-incubation, (non-labeled) intracellular NO3- had been completely consumed. For the subsequent storage of intracellular 15N-labeled NO3- (98 atom%; Cambridge Isotope Laboratories), the NO3--starved cells were harvested, re-inoculated into sterile, 15NO3--containing F/2 medium plus silicate in artificial seawater, and cultured under optimal growth conditions for three days (see above). The cells were then washed via gentle centrifugation with sterile NO3--free artificial seawater (salinity 35; 10 min at 1000g) to remove 15NO3- from the medium, and transferred into NO3--free artificial seawater enriched with 200 μM Na-acetate and 25 μM non-labeled NH4+. Thus, the only NO3- source during the 15N-stable isotope labeling experiment was 15NO3- stored intracellularly by the diatoms. The cell density was obtained and the experiment was started by dividing the culture for (a) the dark/anoxic incubation, and (b) the light/oxic control. For the dark/anoxic incubation, ca. 200 mL of the diatom suspension was transferred into a dark bottle (wrapped in aluminum foil) and flushed with He for 30 min to remove O2 and then transferred into 24 replicate 6 mL Labco-exetainers® wrapped in aluminum foil. At time intervals of 1, 2, 3, 4, 5, 6, 8, and 10 h, a He headspace of 3 mL was set in three Labco-exetainers® each, and the diatom cells in the remaining 3 mL were killed with 100 µL ZnCl2 (50%). The Labco-exetainers® were stored upside down at room temperature until measurement of 15N-labeled N2 by gas chromatography-isotope ratio mass spectrometry (GC-IRMS, VG Optima; Isotech). The cell suspension collected during setting the headspace was filled into 15-mL tubes and centrifuged (10 min at 1000g). Part of the cell-free supernatant was used for immediately measuring the extracellular NO3- concentrations, while the pellet was used for intracellular NO3- determination (see above). Further, 1 mL cell-free supernatant was frozen at -20°C until 15NH4+ analysis using the hypobromite assay [31], followed by N2-15N analysis using GC-IRMS. The hypobromite assay actually measures the sum of 15NH4+ and 15N-labeled volatile N compounds such as methyl amines [32]. For the light/oxic control, the culture was kept under light/oxic conditions, and at time intervals of 0, 1, 2, 5, and 10 h, 3 mL cell material each was taken and processed exactly like the material that was obtained during setting the headspace in the dark/anoxic treatment. The sample collected at time point zero was used for both, the dark/anoxic incubation and the light/oxic control.

Intermediates and By-Products of Dissimilatory Nitrate Reduction

The time courses of N2O and NO2- as possible intermediates or by-products of dissimilatory NO3- reduction were measured during the 15N-stable isotope labeling experiment. Nitrous oxide was measured in the headspace of the Labco-exetainers® from the dark/anoxic incubation experiment after N2-15N analysis had been completed (see above; the gas volume removed for N2-15N measurements was taken into account for the subsequent calculation of N2O concentrations), using a GC 7890 (Agilent Technologies) equipped with a CP-PoraPLOT Q column and a 63Ni electron capture detector. Nitrite was determined in the supernatant of the medium from the dark/anoxic and the light/oxic incubation with an NOx analyzer as described for the NO3- determination, except that the reaction chamber contained acidified NaI (2 M) that reduces NO2- to NO at 20°C.

Degradation of Photopigments in Response to Dark/Anoxic Conditions

Chlorophyll a and fucoxanthin were determined in cultures of T. weissflogii that were first exposed to favorable growth conditions (i.e. with light and O2; time 0) and then to dark/anoxic conditions for a time period of 46 weeks. To adjust dark/anoxic conditions, diatom cultures were transferred into gas-tight, dark bottles (wrapped in aluminum foil), flushed with N2 for 30 min and kept at 15°C until sampling. At each time point, 2 mL of the cell suspension was taken in 3 replicates and cell numbers were counted. The samples were freeze-dried for 2 days and 5 mL ice-cold acetone was added for extraction of photopigments. After vigorous mixing and sonication for 5 min, the samples were left over night at -20°C, mixed again, and centrifuged for 5 min at 3000g. The supernatants were filtered (Acrodisc® CR 4 mm, 0.45 µm Versapor®; Gelman Laboratory) and the extracted photopigments were separated by means of HPLC (Waters 2695; U.S.A.) and analyzed by a photodiode array detector (Waters 996; U.S.A.) as described in Stief et al. [26]. In the chromatograms, chlorophyll a and fucoxanthin were identified according to their specific retention time and absorption spectra and the respective peaks were integrated with the Millenium®32 software (Waters, U.S.A.). Calibrations were made with serial dilutions of chlorophyll a and fucoxanthin stock solutions (DHI, Denmark). All procedures were made under dark conditions and using HPLC-grade chemicals.

Results and Discussion

Dissimilatory Nitrate Reduction to Ammonium by T. weissflogii

Our results strongly indicate that the ubiquitous pelagic diatom T. weissflogii is able to perform DNRA, similar to the benthic diatom Amphora coffeaeformis, which was the first phototrophic eukaryote shown to dissimilatorily reduce NO3- under dark/anoxic conditions [8]. Consumption of intracellular NO3- and simultaneous production of NH4+ in response to dark/anoxic vs. light/oxic conditions have been followed in two separate experiments: (a) a non-labeling experiment in which NH4+ was measured photometrically (Figure 1) and (b) a 15N-stable isotope labeling experiment (Figure 2). In both experiments, the rapid consumption of intracellular NO3- and 15NO3- by T. weissflogii was accompanied by the production and release of NH4+ and 15NH4+, respectively, only under dark/anoxic conditions, but not in the presence of light and O2 (Figures 1,,2).2). In the 15N-stable isotope labeling experiment, the initial 15NH4+ concentration was 2 µM because the hypobromite assay actually measures the sum of 15NH4+ and 15N-labeled volatile N compounds such as methyl amines [32]. The concentration of NO3- in the medium, i.e. extracellular NO3-, only decreased under light/oxic conditions, but remained constant after exposure to dark/anoxic conditions (Figure 1). This constant (and not increasing) extracellular NO3- concentration indicates that the intracellular NO3- (expressed in μmol L-1 of growth medium) was indeed consumed by T. weissflogii rather than released from the cells into the medium. Intracellular NO3- was also consumed under light/oxic conditions, even at a higher rate than under dark/anoxic conditions (Tables 1,,2),2), most probably because NO3- was used for assimilation by photosynthetically active diatoms [33-35]. For N-assimilation, NO3- is also reduced to NH4+, but NH4+ is not released from the cells.

Figure 1
Non-labeling experiment.
Figure 2
15N-stable isotope labeling experiment.
Table 1
Cell-specific consumption (neg. values) rates of NO3- by axenic T. weissflogii cultures in response to different experimental conditions for the non-labeling experiment.
Table 2
Cell-specific consumption (neg. values) and production (pos. values) rates of N compounds by axenic T. weissflogii cultures in response to different experimental conditions for the 15N-stable isotope labeling experiment.

In the absence of O2, intracellular NO3- can be used for dissimilation by sulfur bacteria [36-39] and only a few unicellular eukaryotes and fungi (e.g. [8,18,20,40,41]). The ubiquitous diatom T. weissflogii can now be added to the short list of eukaryotes that dissimilatorily reduce NO3-. Notably, T. weissflogii is the first marine pelagic eukaryote shown to have an anaerobic NO3- metabolism, whereas all known eukaryotic NO3- reducers thrive in stratified waters, sediments and soils in which anoxic conditions occur in subsurface layers. So far it is not known, whether DNRA in T. weissflogii is respiratory or fermentative. Briefly, in respiratory DNRA, ATP is generated by an electrochemical proton potential across a cell membrane, at which electrons are transferred from the donor to the acceptor NO3-, and in fermentative DNRA, ATP is generated by substrate-level phosphorylation [16,42-44]. In prokaryotes, the electron donor and acceptor for respiratory DNRA usually originate from an external source and not from cell metabolism, but may be either organic or inorganic, whereas the electron donor in fermentative DNRA is usually organic [45]. So far, the electron donor used by diatoms for DNRA is not known. In our labeling experiment, acetate was added as a potential electron donor. However, it needs to be further investigated, if T. weissflogii can perform DNRA also with intracellularly stored electron donors, like polysaccharides (e.g. chrysolaminarin), and if the external supply of acetate indeed influences the rate of DNRA.

Our experiments revealed that the rate of NO3- consumption after exposure to dark/anoxic conditions depends on the concentration of intracellularly stored NO3-. In the non-labeling experiment, the initial intracellular NO3- concentration was 20 mM, and in the labeling experiment only 4 mM, resulting in a 6 times lower rate of NO3- consumption (Tables 1,,2).2). In the labeling experiment, the production of 15NH4+ (plus N2O and NO2-) by T. weissflogii balanced the consumption of intracellular 15NO3- within the bounds of accuracy (Figure 2; Table 2), whereas in the non-labeling experiment, the net production of NH4+ did not balance the consumption of intracellular NO3- (Figure 1). On average, less than half of the NO3- was found back as NH4+ in the culture medium; further, the NH4+ concentration first increased and then decreased slightly with time (Figure 1). This decrease of the NH4+ concentration in the non-labeling experiment is explained by an uptake of NH4+ by T. weissflogii under dark/anoxic conditions that has also been confirmed in other experiments (data not shown). A dark NH4+ uptake and assimilation, respectively, is generally known for phytoplankton [34,46] and was recently also confirmed by gene expression analysis in Thalassiosira pseudonana [47]. This dark NH4+ uptake is not apparent in the labeling experiment, because the addition of non-labeled NH4+ as background concentration (see Materials and Methods) obscures the putative uptake of 15NH4+. However, under the light/oxic conditions of the labeling experiment, 15NH4+ also decreased after 5 h because the (non-labeled) background NH4+ was completely taken up (data not shown). The labeling approach did not reveal a production of N2-15N by T. weissflogii (Figure 2), which further supports that DNRA and not denitrification is used as a dissimilatory NO3- reduction pathway by T. weissflogii.

Release of Nitrous Oxide and Nitrite during Nitrate Dissimilation

The production of N2O and NO2- in response to dark/anoxic conditions has been followed during the 15N-stable isotope labeling experiment (Figure 3). Both, N2O and NO2- were produced and released from the cells in the same time pattern that has been observed for 15NH4+, and their production apparently mirrors the consumption of intracellular NO3- (Figures 1,,22,,3).3). However, the production rates of N2O and NO2- were about 1000 and 100 times, respectively, lower than the production rate of 15NH4+ (Table 2). Thus, N2O and NO2- are not final products of dissimilatory NO3- reduction, but the congruent time patterns indicate that T. weissflogii releases N2O and NO2- as by-product and intermediate, respectively, of DNRA. In prokaryotes, N2O is a well-known by-product and intermediate of nitrification and denitrification, respectively, and there are some indications that N2O is also released as a by-product of DNRA, which might have been overseen in some organisms [16,48]. In higher plants, N2O is emitted from leaves by plant NO3- assimilation, strictly speaking during photoassimilation of NO2- in the chloroplast [49]. Recently, N2O production was also found in axenic, illuminated cultures of the green algae Chlorella vulgaris [50]. A release of N2O by phototrophic eukaryotes under darkness and anoxia has to our knowledge not been documented so far. Even though the rate of N2O released by T. weissflogii during DNRA might seem low (see above, Table 2), this finding can be of environmental importance because diatoms are highly abundant in the world’s oceans (e.g. [3,51]), hypoxic and anoxic marine environments are spreading [52], and N2O is a particularly strong greenhouse gas [53]. The production of N2O under dark/anoxic conditions has recently also been confirmed for the benthic diatom A. coffeaeformis [8], and it might be worth to screen other benthic and pelagic diatom species for N2O emission under these conditions.

Figure 3
Time courses of extracellular N2O and NO2- concentrations in an axenic T. weissflogii culture in response to dark/anoxic conditions.

The NO2- release during DNRA by T. weissflogii could be due to cell leakage or excretion that is frequently observed in marine phytoplankton, including diatoms [54,55], but has not been linked to a response of phytoplankton to darkness and anoxia so far. The observed NO2- release might be supported by a slightly higher rate of NO3- reduction than NO2- reduction throughout the incubation. Further, there might be a time delay in NO2- reduction to NH4+ because of constitutive expression of the NO3--reductase gene, whereas the (dissimilatory) NO2--reductase gene first needs to be induced by the production of NO2-. T. weissflogii is not able to take up the released NO2- again under dark/anoxic conditions, which is indicated by the observation that the medium NO2- concentration is not decreasing during the incubation (Figure 3). Additionally, intracellular NO2- storage does not occur in T. weissflogii (data not shown), probably because of the toxic effects of NO2- [54].

Slow Degradation of Photopigments in Darkness and Anoxia

To estimate how long T. weissflogii cells retain the ability to operate photosynthesis after exposure dark/anoxic conditions, the fate of the photopigments chlorophyll a and fucoxanthin was followed. Notably, the degradation of the photopigments did not temporally coincide with DNRA by T. weissflogii in response to dark/anoxic conditions. While DNRA activity peaked during the first few hours of dark/anoxic conditions, the major decrease in cellular photopigment contents occurred during the first 3 days (Figures 2,,4).4). After one week of dark/anoxic incubation, the cellular pigment contents had reached a low, but constant level that was maintained for at least 7.5 weeks (Figure 4). These observations are in good agreement with the hypothesis that diatoms use DNRA to enter a resting stage with low metabolic activity, and that T. weissflogii was found to survive at least for 6 weeks after adjusting them to dark/anoxic conditions [8]. Diatoms are known to start photosynthesis and growth very fast after (re)adjusting them to favorable growth conditions, i.e. light and fresh growth medium, even after extended periods of darkness [24,56,57]. To maintain at least low cellular contents of photopigments must be a prerequisite for this. Our experimental design, i.e. that no O2 and prokaryotes were present in the T. weissflogii culture, further led to a decreased rate of degradation, as O2-dependent pigment alteration and grazing-induced cell disruption could not occur [58]. Interestingly, the chloroplasts of T. weissflogii cells showed an autofluorescence even after more than 1 year under dark/anoxic conditions (pictures not shown), which might originate from photopigment degradation products that are still poorly understood [59].

Figure 4
Time courses of the intracellular chlorophyll a and fucoxanthin concentrations in an axenic T. weissflogii culture in response to dark/anoxic conditions.

Ecological and Evolutionary Perspectives

After the benthic diatom A. coffeaeformis was discovered as the first photothrophic eukaryote that dissimilatorily reduces NO3-, it was interesting to ask whether this metabolism also occurs in pelagic diatoms: and indeed, we found T. weissflogii as the so far only marine pelagic eukaryote showing this metabolic trait. The respiration of NO3- by diatoms might be widespread in marine ecosystems and could have so far overseen implications on the marine N-cycle. For benthic foraminifera, Piña-Ochoa et al. [20] calculated a contribution for the removal of fixed N from marine ecosystems that may be equally important to bacterial denitrification in the seafloor. DNRA will not directly remove fixed N, but in anoxic or hypoxic environments, the produced NH4+ can serve as electron donor for anammox that might be especially important in OMZs with high abundances of anammox bacteria [60]. Further research on NO3- respiration by diatoms might also reveal that certain species are capable of other pathways than DNRA, like denitrification as shown for foraminifera [18,20]. Additionally, the exact ambient O2 concentration in the (micro)environment of the diatoms may trigger different dissimilatory NO3- reduction pathways as known from fungi [45].

To date, genes involved in dissimilatory NO3- reduction have not been identified in NO3--respiring diatoms, foraminifera or gromidii. In contrast, several functional genes have been identified in the denitrifying fungus Fusarium oxysporum: a copper-containing NO2- reductase (nirK) and a nitric oxide reductase (P450nor) have been sequenced and characterized [41,61]. Intriguingly, NO3--respiring fungi may use enzymes that are normally involved in assimilatory NO3- reduction in a dissimilatory mode instead [62]. This could also hold true for diatoms. Assimilatory NO3- reductases, multiple transporters for NO3-, and components of a NO3--sensing system have only recently been discovered in diatom genomes [63,64]. First insights into diatom genomes and the ensuing ecophysiological studies revealed a fascinating evolutionary history of diatoms. An unexpected combination of genes by endosymbiotic gene transfer from two secondary endosymbionts to the exosymbiont nucleus, and also horizontal gene transfer led to several additional inclusions from Bacteria and Archaea genomes [63-66]. The diverse assortment of genes results in novel biochemical pathways like the urea cycle [63,65,67-71] that formerly was not known for photosynthetic organisms and congruously makes diatoms for Armbrust et al. [51] to be neither plants nor animals. Further work on diatom genomes could lead to the identification of functional genes involved in dissimilatory NO3- reduction. This would not only convey genetic evidence of dissimilatory NO3- reduction by eukaryotes, but would also provide genetic markers for the cultivation-independent detection of so far unrecognized dissimilatorily NO3- reducing diatoms directly in the environment.

Acknowledgments

We thank the technicians of the Microsensor Group of MPI Bremen for practical assistance and Bo Thamdrup for critically reviewing the manuscript.

Funding Statement

Financial support was provided by the Max Planck Society (Germany), and by a grant from the German Research Foundation awarded to A.K. (KA 3187/2-1). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

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