Ecology

Comparison between old and new deciduousness metrics
At first, to check the reliability of the proposed metric, we estimated the deciduousness from the equation proposed by Cuba et al.14 (Eq. 1; referred as ‘old’) and the new metric proposed in this study (Eq. 2; referred as ‘new’) during the extreme and normal rainfall years. The results of dry and moist deciduous samples and 4 pheno-classes revealed an over-estimation and under-estimation of deciduousness with the old-metric, whereas the new metric revealed the accurate relative variability (Fig. 2b,c, Table S1). Table 1 provides the estimated deciduousness values from the old and new metrics for 22 homogeneous sample pixels representing four major vegetation types in the study area (refer Fig. 1 for their spatial locations and Fig. S1 for their annual growth profile). The litter fall information collected from literature revealed a higher litter fall quantity of 10–14.4 Mg Ha−1 year−1 for the moist deciduous forest39,40,41,42 and lower litter fall quantities of 1–8.65 Mg Ha−1 year−1 and 5.63–7.84 Mg Ha−1 year−1 for the dry deciduous forest42,43,44 and the semi-evergreen and evergreen forest44 , respectively. The new metric showed a relatively similar variability in deciduousness to ground observations especially for the moist and dry-deciduous forests than the old metric (Table 1).
Figure 2

Graphical illustration of deciduousness estimation: (a) Theoretical phenology curves from high and low deciduous vegetation and the parameters of deciduousness, (b) Actual RS derived annual growth profiles of moist and dry deciduous vegetation and their deciduousness estimation using the old and new metric, and (c) Annual growth profile of four theoretical pheno-classes for depicting the different magnitude of deciduousness.

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Table 1 Performance of old and new deciduous metric in a normal rainfall year (2011) using 22 samples from different vegetation types (spatial locations of these samples can be seen in Fig. 1).
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Further, the difference between the old and new metric was spatially checked and is shown at the center of Fig. 3, and the actual values are presented in the surrounding in eight different sub-set locations. The difference image denotes the under-estimated (70.76% of forest area) and the over-estimated (29.23% of forest area) deciduousness obtained by the old metric (Fig. 3). The under-estimated area observed was mainly in the moist forested regions of states- Chhattisgarh, Odisha, and Jharkhand states, whereas, the over-estimated area observed was mainly in the dry forested region of states—Madhya Pradesh, Maharashtra, Northern Chhattisgarh and some parts of Jharkhand (Fig. 3). The over- and under-estimations are with respect to the new metric, and not with the real in-situ measurements. However, the new metric is in good agreement with annual growth profiles of different vegetation types, and have positive relation with ground litter fall observations39,40,41,42,43,44.
Figure 3

Difference in the spatial distribution of deciduousness (central figure) and the actual deciduousness (subset boxes) derived from the new and old metric for the year 2011. (These maps were created using ESRI’s ArcMap 10.3—https://desktop.arcgis.com/en/arcmap/, and MS-Office PowerPoint 2007 software).

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The deciduousness derived from these two metrics were also tested for their statistical significance using ANOVA (Table 2). In this test 800 stratified random samples belonging to different deciduous forests of different density classes for dry (2002), normal (2011) and wet (2013) years were used. It was found that the mean deciduousness values from the old metric were similar in the majority of the cases and different rainfall conditions. Hence, it could not be used for understanding rainfall impact on the deciduousness. On the other hand, the new metric performed better than the old metric in terms of its variability under (a) different rainfall conditions (p  More

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Cultivation of novel Nitrospinae representatives
For the initial enrichment of marine NOB, nitrite-containing mineral media were inoculated with coastal surface sediment samples taken in Vancouver, Canada, and Elba, Italy. Within 4 weeks of incubation, nitrite oxidation to nitrate was detected in the cultures and this activity continued to be observed after subsequent replenishment of nitrite and transfers of culture aliquots into fresh medium. Usually, the further purification of NOB from accompanying organisms in the enrichment cultures is hindered by the very slow growth and inability of most NOB (including all cultured Nitrospinae) to grow on solid media. To expedite the purification of Nitrospinae strains from our initial enrichments, which was previously a laborious and lengthy process [24], we developed a method for the physical separation, activity-based identification of NOB, and subcultivation in 96-well microtiter plates. This method uses random, non-fluorescent, single-cell sorting using a fluorescence activated single cell sorting (FACS) instrument paired with a nitrite consumption activity screen (Fig. S1a). Thus, it differs from a previously reported FACS isolation approach for Nitrospira NOB from activated sludge, where the NOB were targeted based on their known cell cluster size and shape, which had been determined by Nitrospira-specific rRNA-targeted FISH analysis before FACS was performed [43]. Our method does not rely on prior knowledge of the identity and morphology of the NOB [43] but is solely based on the detection of nitrite oxidation after sorting. Still, it might allow for a flexible selection of the sorted cell morphologies by adjusting the gating parameters. While the previous method facilitated the isolation of already known NOB, the approach used in our study was designed for the discovery of novel nitrite oxidizers that grow under the given conditions. It may also be suitable for the isolation of other microorganisms that can be efficiently sorted (i.e., grow in suspension or that can be suspended by sonication or other methods) and that perform a specific metabolism of interest, which is detectable by a colorimetric, fluorimetric, or other high-throughput assay. Examples include previously performed, high-throughput enzyme discoveries and the isolation of microalgae [44,45,46]. After sorting cells from the initial Vancouver and Elba enrichment cultures into one 96-well plate each, several wells showed nitrite-oxidizing activity within 3–4 weeks. Cells from three active wells from the Elba enrichment and 4 for the Vancouver enrichment ( >4 were active) were progressively transferred into larger culture volumes. This procedure led to the separate enrichment of two different Nitrospinae, one from each of the initial enrichments, that were identified by 16 S rRNA gene sequencing (Fig. S1b): strain VA (Vancouver) and strain EB (Elba). Interestingly, despite attempted single-cell sorting and various dilution to extinction attempts, an axenic culture could not be established for either of the two obtained Nitrospinae strains. Rather, these cultures represent binary co-cultures each containing one Nitrospinae strain, one alphaproteobacterial strain, and no other detectable microorganisms. Overall, using this single-cell sorting and screening method, we were able to obtain the two binary co-cultures from the environmental samples in ~10 months.
Co-enrichment with Alphaproteobacteria
Pure cultures of the two co-enriched heterotrophic strains were obtained on Marine Broth Agar. Subsequent 16S rRNA gene analysis of these isolates showed that the two Nitrospinae strains had been co-cultured with members of two distinct genera within the Alphaproteobacteria. Strain VA was co-cultured with a bacterium most closely related to a Stappia stellulata strain (NR_113809.1, 99.58% 16 S rRNA gene sequence identity), whereas the EB strain culture contained a Maritimibacter alkaliphilus strain (NR_044015.1, 100% 16 S rRNA gene sequence identity). Both these species have previously been isolated from marine environments and have been described as alkaliphilic chemoorganoheterotrophs that can use a wide variety of simple and complex organic substrates [47, 48]. In our cultures, which were only provided with nitrite for growth, the alphaproteobacterial strains may have lived off simple organic compounds that were excreted by the autotrophic Nitrospinae. Since the NOB could not be grown separately from the heterotrophs, it is tempting to speculate that the Nitrospinae strains also benefitted, for example, from reactive oxygen species (ROS) protection by the heterotrophs. Superoxide dismutase and catalase genes are present in the genomes of both co-cultured alphaproteobacteria, and other isolates of both species are catalase positive [47, 48]. Similar interactions have already been observed in marine autotroph-heterotroph co-cultures, including other nitrifiers [49,50,51].
Phylogeny of the novel Nitrospinae
Closed genomes were reconstructed for both Nitrospinae strains by co-assembling Illumina and Nanopore sequencing data. Since the genome of N. gracilis was nearly completely sequenced but not closed [25] (Table S3), the obtained genomes represent the first closed genomes from Nitrospinae. With a length of >3.9 Mbp, Nitrospinae strain EB has a larger genome than the other cultured Nitrospinae strains sequenced to date (Table S3).
Phylogenetic analysis of the 16S rRNA genes (Fig. S1b) showed that the Nitrospinae strains VA and EB are only distantly related to N. gracilis (93% sequence identity with EB and 91% with VA), N. watsonii (92% sequence identity with EB and 90% with VA), and to each other (90% sequence identity). Since public databases contained only few 16 S rRNA gene sequences closely related to strains VA and EB (Fig. S1b), these strains seem to represent a yet underexplored diversity of Nitrospinae. A phylogenetic tree using conserved concatenated marker proteins from all available Nitrospinae genomes that are >80% complete and 70%, and that these genera are distinct from the genus containing strain VA (Fig. 2). In order to make taxonomic inferences beyond the genus level, the GTDB-TK tool [38] was applied to the Nitrospinae genome dataset. GTDB-TK confirmed that strains VA and EB belonged to the Nitrospinaceae family but, in agreement with the AAI analysis, did not assign them to a genus (Tab. S2). While we showed that nitrite oxidation is spread among phylogenetically distant members within the Nitrospinaceae, no nitrite oxidation phenotype has been observed yet for the Nitrospinae members that do not belong to the Nitrospinaceae family.
Fig. 2: Average amino acid identity (AAI) analysis of the Nitrospinae.

Pairwise AAI values were calculated for the same set of Nitrospinae genomes that was used to reconstruct the phylogenetic tree in Fig. 1. The same tree is used here to annotate the heatmap. The two newly cultured strains are highlighted in red. Clades of the Nitrospinae are indicated as proposed elsewhere [6]. The boxes indicate the genus boundary at 60% identity.

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Taken together, the phylogenetic, ANI, AAI, and GTDB-TK analyses revealed a high genus-level diversity within the Nitrospinae, the vast majority remaining uncultured and poorly characterized. Among all cultured Nitrospinae members, strain VA is most closely related with the uncultured but environmentally abundant clades 1 and 2 [6, 7, 20, 21].
Cell morphology
The morphologies of the new Nitrospinae strains were visualized using SEM, and by 16S rRNA-targeted CARD-FISH using a Nitrospinae-specific oligonucleotide probe (Ntspn759) [21]. Strain VA cells were helically shaped rods (Fig. 3a, d, g), whereas strain EB appeared to be short, slightly curved rods (Fig. 3b, e, h). Interestingly, neither of the two strains displayed the long, slender, rod-shaped morphology that was mostly observed for the previously isolated Nitrospina species, N. gracilis (Fig. 3c, f, i) [23]. The shape of strain VA (Fig. 3a, d, g) rather resembled Nitrospira species [53]. The short, slightly curved rod morphology of strain EB (Fig. 3b, e, h) resembled the compact cell shape reported for aging cultures of N. watsonii [24] and environmental Nitrospinae [5]. While the helical shape of strain VA could be clearly distinguished from the co-cultured Stappia sp. using Nitrospinae-specific FISH (Fig. 3g) and SEM of the Stappia-like isolate (Fig. S3a), assigning a morphology to strain EB was slightly more difficult due a more similar morphotype of the co-cultured Maritimibacter-like bacterium (Figs. 3e and S3b). According to SEM, the isolated Maritimibacter had a coccoid morphology (Fig. S3b) similar to the slightly smaller coccoid cells that were observed in the active co-culture with strain EB (Fig. 3e). Therefore, we assume that the slightly larger, curved rods in the SEM pictures (Fig. 3b, e) were Nitrospinae strain EB cells. However, the previously described morphological variability suggests that the cell shape of Nitrospinae is influenced by the growth stage and environment [23, 24]. Thus, morphology would be of limited use as the sole criterion to differentiate Nitrospinae strains from each other and from other organisms. We propose the name “Candidatus Nitrohelix vancouverensis” VA for strain VA based on its observed morphology and isolation source, and “Candidatus Nitronauta litoralis” EB for strain EB based on its isolation source.
Fig. 3: Scanning electron microscopy (SEM) and CARD-FISH images of the two newly cultured Nitrospinae strains and N. gracilis.

a, d, g “Ca. Nitrohelix vancouverensis” strain VA. b, e, h “Ca. Nitronauta litoralis” strain EB. c, f, i N. gracilis. The short rod in (d) is a cell of the accompanying heterotroph Stappia sp. in the enrichment. e Shows slightly longer and slightly curved rods (strain EB, arrows) and shorter rods, which are cells of the accompanying heterotroph M. alkaliphilus in the enrichment. The scale bars in (a–c) depict 1µm; all other scale bars depict 2µm. g–i The 16S rRNA-targeted probe Ntspn759 was used to detect Nitrospinae cells (magenta), and total nucleic acids were stained with SYBR Green (cyan).

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Nitrite oxidation: activity and kinetics
Both Nitrospinae strains oxidized nitrite stoichiometrically to nitrate (Fig. S4). Exponential growth of “Ca. N. vancouverensis” correlated with the consumption of nitrite (Fig. S4a), whereas “Ca. N. litoralis” did not enter the exponential growth phase during the incubation period (Fig. S4b). During the experiment, the relative abundance of “Ca. N. vancouverensis” compared to the co-cultured Stappia sp. increased pronouncedly from 6 to 75% (Fig. S4a). The relative abundance of “Ca. N. litoralis” compared to the Maritimibacter sp. could not be reliably determined during the incubation experiment (Fig. S4b), but measurements taken after the MR experiments (see below) showed that the relative abundance of “Ca. N. litoralis” ranged from ~56 to 99%. Thus, the quantitative composition of each co-culture appeared to fluctuate and likely depended strongly on the availability of nitrite as the substrate for the NOB strains.
The kinetics of nitrite oxidation of N. gracilis and the two novel Nitrospinae strains were assessed by measuring the nitrite-dependent oxygen consumption in MR experiments. For all three NOB, nitrite oxidation followed Michaelis–Menten kinetics (Fig. S5). The stoichiometry of NO2− and O2 consumption was always close to 1:0.5 (N. gracilis: mean = 1:0.49; s.d. = 0.02; n = 8; “Ca. N. vancouverensis”: mean = 1:0.51; s.d. = 0.01; n = 4; “Ca. N. litoralis”: mean = 1:0.51; s.d. = 0.02; n = 5). This ratio was expected for NOB [54] and indicates that O2 consumption by the co-enriched heterotrophs was only minor during the relatively short MR experiments (maximum 1 h) and did not affect the kinetic analysis of “Ca. N. vancouverensis” and “Ca. N. litoralis”. The apparent half-saturation constant, Km(app), of N. gracilis was determined to be 20.1 µM NO2− (s.d. = 2.1, n = 8) and the maximum reaction rate, Vmax, to be 41.4 µmol NO2− mg protein−1 h−1 (s.d. = 9.4, n = 6), which is highly similar to the previously reported Km(app) and Vmax of the closely related N. watsonii (18.7 ± 2.1 µM NO2− and 36.8 µmol NO2− mg protein−1 h−1) [55]. The Km(app) measured for “Ca. N. litoralis” was 16.2 µM NO2− (s.d. = 1.6, n = 7) and thus resembled the values of N. gracilis and N. watsonii. In contrast, with a Km(app) of 8.7 µM NO2− (s.d. = 2.5, n = 3), “Ca. N. vancouverensis” showed a higher affinity for nitrite that was comparable with non-marine Nitrospira members (Km(app) = 6–9 µM NO2−), which have been the cultured NOB with the highest nitrite affinity known so far [54, 56]. Indeed, strain VA turned out to have the lowest Km(app) of all hitherto analyzed marine NOB in culture (Fig. 4). Interestingly, among all cultured marine NOB, “Ca. N. vancouverensis” is also most closely related to the Nitrospinae clades 1 and 2 that are abundant in oligotrophic waters [20] (see above). However, its Km(app) is still 1–2 orders of magnitude higher than the Km(app) values of nitrite oxidation reported for environmental samples from an OMZ (0.254 ± 0.161 μM NO2−) and South China Sea waters (0.03–0.5 µM) [22, 57]. The very high nitrite affinity observed with these samples might be explained by the presence of uncharacterized nitrite oxidizers, whose nitrite affinity exceeds that of all cultured NOB. However, it remains to be tested whether known NOB can persist under extremely low in situ nitrite concentrations. For example, the half-saturation constant for ammonia oxidizing bacteria spans several orders of magnitude under different growth temperatures [58]. Strongly different substrate affinities have also been observed for Nitrobacter winogradskyi and Escherichia coli under oligotrophic versus copiotrophic growth conditions [59, 60]. Systematic assessments of the kinetic plasticity of NOB under different conditions are still pending, mainly because the production of sufficient biomass of NOB isolates has been a major obstacle for such studies.
Fig. 4: Comparison of the whole-cell apparent half-saturation constants (Km(app)) for nitrite between marine and non-marine NOB.

The Km(app) values measured in this study (highlighted in red) are the mean from all biological replicates (n = 3 for “Ca. Nitrohelix vancouverensis” VA; n = 5 for “Ca. Nitronauta litoralis” EB; n = 8 for N. gracilis). The other Km(app) values were retrieved from previous studies [54, 55, 81].

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General genomic features of cultured Nitrospinae
A pan-genomic analysis of the two novel cultured Nitrospinae strains and N. gracilis showed that these three organisms share a core genome of 1347 proteins, which have at least 50% amino acid sequence identity over 80% of the alignment (Fig. S6). The core genome included universally highly conserved bacterial genes, such as those coding for ribosomal proteins, translational elongation factors and the DNA replication machinery, as well as the genes for the core metabolism of chemolithoautotrophic NOB. Interestingly, among the shared conserved genes we also found highly conserved glutaredoxins, thioredoxin reductases, and peroxidases ( >80% amino acid identity among the respective homologs). Like N. gracilis [25], “Ca. N. vancouverensis” and “Ca. N. litoralis” lack the canonical defense systems of aerobic organisms against oxidative stress, catalase and superoxide dismutase. While the aforementioned core genes could thus be essential for the detoxification of peroxides in all three organisms [61, 62], it remains a mystery how Nitrospinae deal with superoxide radicals [25]. Each of the strains encode a number of unique proteins (Fig. S6), many of which are phage related, corroborating a recently proposed hypothesis predicting extensive phage predation on Nitrospinae [21] (Supplemental Results and Discussion, Fig. S7). Yet, the majority of the variable genome content is still functionally uncharacterized. However, a few genes of the variable genome have known functions and might be important for niche adaptations. In the following sections, we address these genes as well as the shared core metabolism of the three analyzed Nitrospinae.
Nitrite oxidation and respiration
Among the highly conserved proteins are the three known subunits of a periplasmic NXR, NxrABC. Details of the predicted subunit composition and cofactors of the NXR of Nitrospinae, which is closely related to the NXR of Nitrospira, have been described elsewhere [3, 25, 63]. Briefly, all three Nitrospinae strains possess two genomic copies of the substrate-binding subunit NxrA, two or three (only “Ca. N. vancouverensis”) copies of the electron-channeling subunit NxrB (Fig. S8), and several copies of putative NxrC subunits, which may transfer electrons from NXR to a downstream acceptor in the electron transport chain (Table S4). Homologs to all of the different putative NxrC subunits of N. gracilis [25] were also found in “Ca. N. vancouverensis” and “Ca. N. litoralis” (Table S4), but it remains to be determined whether all of these proteins are functionally involved in nitrite oxidation.
The respiratory electron transport chain of NOB is short, as electrons derived from nitrite are directly transferred, via a- or c-type cytochromes, to the terminal oxidase (complex IV) [25, 63, 64]. N. gracilis carries a cbb3-type high affinity heme-copper cyt. c oxidase (HCO) [25], whereas both “Ca. N. vancouverensis” and “Ca. N. litoralis” lack any canonical HCO. However, all three organisms encode highly conserved, putative “bd-like oxidases” [25]. These proteins, which also occur in all Nitrospira genomes, are phylogenetically related with but clearly distinct from the canonical cyt. bd-type quinol oxidases [63]. Interestingly, one variant of the bd-like oxidases from Nitrospira contains all conserved amino acid residues for heme and Cu binding in HCOs [65], indicating that this enzyme could be a novel cyt. c-oxidizing HCO [63]. The bd-like oxidases of N. gracilis, “Ca. N. vancouverensis”, and “Ca. N. litoralis” have most of these conserved residues; however, one of the three histidine ligands of the CuB is replaced with a glutamine, and the histidine ligand of the high-spin heme is replaced with a phenylalanine. Thus, without the cbb3-type oxidase found only in N. gracilis, it remains unclear how the final electron transport step from cyt. c to O2 occurs in “Ca. N. vancouverensis” and “Ca. N. litoralis”. Future biochemical and protein structural research may reveal whether the cyt. bd-like oxidases can catalyze this reaction despite their divergence from bona fide HCOs at two of the predicted cofactor-binding residues, and whether these proteins are capable of proton translocation for proton motive force generation. Corroborating evidence for a function of this enzyme in the context of electron transport stems from its highly conserved genetic synteny within Nitrospinae, Nitrospirae and anammox bacterial genomes. A conserved cluster within these organisms contains a cyt. bd-like oxidase with the aforementioned glutamine and phenylalanine residues, a diheme- and a triheme- cyt. c, and a membrane integral, alternative NxrC subunit (Tab. S4). This putative NxrC might be involved in the electron transfer from NO2− to the terminal oxidase [25, 63, 66].
In addition to the putative cyt. bd-like oxidase discussed above, “Ca. N. litoralis” possesses a canonical cyt. bd-type (quinol) oxidase that is lacking in “Ca. N. vancouverensis” and N. gracilis. Since quinol oxidases cannot accept electrons from the high-potential donor nitrite, we assume that this oxidase receives electrons from quinol during the degradation of intracellular glycogen or during hydrogen oxidation (see below). The cyt. bd-type oxidase may also be involved in oxidative stress defense, as homologous oxidases in other organisms can degrade H2O2 [67] and protect from dioxygen [68]. Taken together, the diverse repertoire of terminal oxidases may be a key feature of Nitrospinae that contributes to the ecological success of this lineage over a broad range of redox conditions in marine ecosystems.
Carbon metabolism and alternative energy metabolisms
Like N. gracilis and other Nitrospinae [6, 25], the novel strains encode all key genes of the reductive tricarboxylic acid (rTCA) cycle for CO2 fixation, including the hallmark enzymes ATP-citrate lyase (ACL), pyruvate:ferredoxin oxidoreductase (POR), and 2-oxogluterate:ferrodoxin oxidoreductase (OGOR) (Table S5). As in N. gracilis, all genes required for the oxidative (oTCA) cycle are also present. All three strains can form glycogen as storage compound, which is degraded via glycolysis and the oTCA cycle. Since N. gracilis lacks pyruvate kinase, the final step of glycolysis may be catalyzed by pyruvate phosphate dikinase (PPDK) in this organism. In contrast, strains VA and EB possess both pyruvate kinase and PPDK, indicating a strict regulatory separation between glycolysis and gluconeogenesis.
Alternative energy metabolisms such as the oxidation of hydrogen, sulfide or organic carbon compounds have been demonstrated in NOB with representatives from the genera Nitrospira, Nitrococcus, Nitrolancea, and Nitrobacter [3, 26,27,28, 69]. Among the three Nitrospinae strains analyzed here, N. gracilis has the largest potential to exploit energy sources other than nitrite: its genome harbors a bidirectional type 3b [NiFe] hydrogenase, which could enable aerobic hydrogen utilization, and a sulfite:cyt. c oxidoreductase [25]. In addition, N. gracilis contains the genes prpBCD for 2-methylisocitrate lyase, 2-methylcitrate synthase, and 2-methylcitrate dehydratase, and might thus be able to catabolically degrade propionate via the 2-methylcitrate pathway (Fig. 5). Of these potential alternative energy metabolisms, “Ca. N. litoralis” shares only the type 3b hydrogenase, whereas “Ca. N. vancouverensis” seems to be an obligate nitrite oxidizer. No genes for the uptake and utilization of urea and cyanate as organic N sources were found in the genomes of the new strains. These genes show a patchy distribution among Nitrospinae [6, 7, 20, 21, 25], suggesting further niche differentiation of these organisms based on the capacity to use organic compounds as sources of reduced N for assimilation.
Fig. 5: Cell cartoon based on the annotation of “Ca. Nitrohelix vancouverensis” VA, “Ca. Nitronauta litoralis” EB and N. gracilis 3/211.

The colored squares indicate the presence or absence of the respective genes, “Ca. Nitrohelix vancouverensis” VA is shown in green, “Ca. Nitronauta litoralis” EB in blue, and N. gracilis 3/211 in purple. The gene annotations are detailed in Table S5. Fd ferredoxin, RE restriction enzymes, TO terminal oxidase.

Full size image

Adaptations to saline environments
The intracellular accumulation of ions or the production of organic osmolytes are two main strategies of microorganisms to cope with the osmotic stress in highly saline, marine environments. Interestingly, the three Nitrospinae strains seem to utilize different osmotic stress defense mechanisms. N. gracilis has the genetic potential to produce glycine betaine, an organic osmolyte that is ubiquitously found in bacteria (Fig. 5) [70]. It also encodes for OpuD, a betaine/carnitine/choline transporter, whereas the genes for glycine betaine synthesis and import are missing in “Ca. N. vancouverensis” and “Ca. N. litoralis”. These two strains harbor the canonical genes ectABC for ectoine biosynthesis (Fig. 5), another widely distributed osmolyte in bacteria [70]. “Ca. N. litoralis” additionally encodes the ectoine hydroxylase EctD and is thus able to form hydroxyectoine. Directly downstream of the (hydroxy-)ectoine synthesis cassette, the two strains further encode an ABC transporter that has similarity to ectoine or amino acid transporters from other organisms and may be utilized for (hydroxy-)ectoine import across the cytoplasmic membrane. Since genes for the synthesis and transport of (hydroxy-)ectoine were also found in “Ca. Nitromaritima” (Nitrospinae clade 1) [6], we assume that usage of (hydroxy-)ectoine is wide-spread among the Nitrospinae. N. gracilis and “Ca. N. litoralis” may also be able to synthesize sucrose as an additional compatible solute (Fig. 5).
Moreover, “Ca. N. vancouverensis” and “Ca. N. litoralis” genomes harbor the gene glsA coding for a glutaminase (Fig. 5), which allows them to deaminate glutamine to glutamate while releasing ammonia. The strains further possess the gltP gene coding for a glutamate/H+ symporter. Both glsA and gltP seem to be lacking in the N. gracilis genome. Since glutamate can play a role in osmoregulation [70], the ability to regulate the intracellular glutamate level via transport or the degradation of glutamine may be one of various adaptations by “Ca. N. vancouverensis” and “Ca. N. litoralis” to rapidly respond to stress caused by fluctuating salinities.
All three Nitrospinae strains have the genomic capacity to synthesize the hopanoids hopan-(22)-ol and hop-22(29)-ene (Fig. 5). Hopanoids are pentacyclic, bacterial lipids that integrate into bacterial membranes, much like cholesterol in eukaryotes [71]. They help regulate membrane fluidity and may be important in highly saline environments [70, 72]. Knock-out studies have shown that cells lacking the ability to make hopanoids are more sensitive to various stresses, such as temperature, pH and osmolarity [72]. Interestingly, a metagenomic study of hopanoid-producing bacteria in the Red Sea revealed Nitrospinae to be among the main organisms harboring squalene hopene cyclase, the key gene for hopanoid production [73]. Other marine NOB (members of Nitrospira, Nitrobacter and Nitrococcus mobilis), and even non-marine Nitrospira and Nitrobacter, also have the squalene hopene cyclase and therefore the genomic potential to produce hopanoids. Insight into the chemical structure of the hopanoids produced by NOB could help to gain insight into early NOB evolution as hopanoids are important lipid biomarkers and commonly used to deduce ancient microbial activity from sediment fossil records [73].
Vitamin B12 auxotrophy
The three Nitrospinae strains lack multiple genes involved in vitamin B12 (cobalamin) synthesis and seem to be auxotrophic for this vitamin, as previously suggested for N. gracilis [74]. To acquire vitamin B12 from the environment, all strains encode the vitamin B12 ABC transporter btuCDF and the outer membrane permease btuB [75, 74]. Alternatively, this transporter may also import cobinamide that may then be converted to vitamin B12 via a salvage pathway that uses the genes yvqK, bluB, and cobS, each encoded in all three Nitrospinae genomes [76]. Hence, the availability of externally supplied vitamin B12 or cobinamide is likely of crucial importance for Nitrospinae in situ and also in lab cultures. The incomplete cobalamin pathway in the already available N. gracilis genome led us to amend the cultivation medium with vitamin B12. Indeed, the addition of vitamin B12, either alone or together with other vitamins, has allowed us to cultivate N. gracilis in a more defined medium that is based on Red Sea salt. Previously, the standard medium for this organism had to be prepared from natural seawater [25]. Furthermore, the addition of vitamin B12 was likely an essential prerequisite for our successful enrichment of novel Nitrospinae after cell sorting. The co-cultured alphaproteobacteria may also provide additional vitamin B12 as they both have the genomic repertoire for its synthesis. In the environment, vitamin B12 could be supplied by different heterotrophic or autotrophic microorganisms, including ammonia-oxidizing thaumarchaeota, which have been shown to produce vitamin B12 [77,78,79] and often co-occur with Nitrospinae [10, 21, 80]. More

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Sampling methods
Ocean sampling at hydrostation S and BATS started in 1954 and 1988, respectively. Hydrostation S began with the pioneering efforts of Hank Stommel (Woods Hole Oceanographic Institution) and colleagues50 at a site approximately 26 km southeast of Bermuda (32° 10′N, 64° 30′W, Fig. 1). The first water–column sampling occurred on the 7th June 1954 from the 61′ R.V. Panularis with more than 1381 cruises conducted up to the present time from the R.V. Panularis II (1967–1983), R.V. Weatherbird (1983–1989); R.V. Weatherbird II (1989–2006) and R.V. Atlantic Explorer (2006–present).
Since the arrival of the R.V. Weatherbird, Hydrostation S has been occupied at near biweekly intervals, with multiple CTD–hydrocasts through the water-column to ~2600 m, and to 4500 m at BATS (more than 450 cruises to the site). Nansen bottles were used for water sampling at first, then 5 L Niskin samplers until October 1988. Thereafter sampling has been conducted with a Seabird 9/11 CTD equipped with 12 L Niskin and Ocean Test Equipment (OTE) samplers.
Water sampling, temperature and CTD measurements
The sampling format has remained substantially consistent for the past 65 years, but with the introduction of CTD–hydrocast sampling in October 1988. From 1954 to 1988, reversing mercury thermometers were used for measurements of temperature until replaced by CTD measurements using a Sea-Bird 9/11 system. Bottle samples before 2012 were taken down to 2600 m at hydrostation S. Since the addition of reliable altimeters to the CTD package, sampling was extended to full ocean depth at both the hydrostation S (~3400 m) and BATS (~4500 m) sites. Numerous sensor configurations have been used on the CTD package (e.g., dual temperature, dual conductivity, dual DO sensors, transmissometer, fluorometer, PAR and altimeter). In contrast, the CTD sampling system has predominately been a Seabird 24-place rosette using 12 L Ocean Test bottles. Before profiling, the CTD is allowed to stabilise at 10 m and once stable, the CTD returns to the surface to start the profile with typical descent rates of 0.5–1.0 m s−1, depending on weather conditions. Water samples are collected on the upcast, whereby the OTE bottles are closed at the target depth after a waiting period of 45 s. The CTD is held at the target depth for another 10 s to allow the SBE35-RT sensor to take an 8 s average. The CTD continues with the upcast at an ascent rate of 0.7–1.0 m s−1. Temperature, conductivity and DO sensors are routinely returned to SeaBird every 6–9 months for routine calibration. The differences between primary and secondary temperature sensors in the deep ocean at BATS ( >3000 m) were 0.002–0.006 °C regardless of time since most recent factory calibration.
Determination of salinity
Salinity samples are typically taken from the OTE bottles at all depths. These samples are collected immediately following DO and CO2 sampling. Samples are taken in 125–250 ml borosilicate glass bottles (Ocean Scientific, UK) that use plastic thimbles to form a better seal. The sample remaining from the previous use is left in the bottles between cruises to prevent salt crystal buildup due to evaporation. When drawing a new sample, the old sample is first discarded over the sampling spigot, and the bottle is rinsed three times with water from the new sample. The bottle is then filled to the shoulder with the sample, and the thimble inserted into the container. The neck of the bottle and the inside of the cap are dried, then the thimble is inserted, and the cap is replaced and firmly tightened. These samples are stored in a temperature-controlled laboratory for later analysis (typically within 1–2 weeks of their collection).
Salinity measurements have been made with a Guildline salinometer at BIOS from 1981 to present (calibrated with IAPSO standard water51) for both BATS and Hydrostation S samples. At present, samples for salinity are analysed on a Guildline Autosal 8400B laboratory salinometer using the manufacturer’s recommended techniques. All readings (10 s average) are taken using the Ocean Scientific interface box and PC software. The salinometer drift during and between successive runs tends to be zero (room temperature carefully controlled and monitored). Bottle salinities are used to calibrate the profiling CTD SBE-04 sensors, and additionally, they are also compared with the downcast CTD profiles to search for possible outliers. Deep-water samples ( >2000 m) are replicated for precision estimates (typically  More