Nitrifying microorganisms occur across a wide temperature range from 4 to 84 °C and previous studies in geothermal systems revealed their activity under extreme conditions. Archaea were detected to be responsible for the first step of nitrification, but it is still a challenging issue to clarify the identity of heat-tolerant nitrite oxidizers. In a long-term cultivation approach, we inoculated mineral media containing ammonium and nitrite as substrates with biofilms and sediments of two hot springs in Yellowstone National Park (USA). The nitrifying consortia obtained at 70 °C consisted mostly of novel Chloroflexi as revealed by metagenomic sequencing. Among these, two deep-branching novel Chloroflexi were identified as putative nitrite-oxidizing bacteria (NOB) by the presence of nitrite oxidoreductase encoding genes in their genomes. Stoichiometric oxidation of nitrite to nitrate occurred under lithoautotrophic conditions, but was stimulated by organic matter. Both NOB candidates survived long periods of starvation and the more abundant one formed miniaturized cells and was heat resistant. This detection of novel thermophilic NOB exemplifies our still incomplete knowledge of nitrification, and indicates that nitrite oxidation might be an ancient and wide-spread form of energy conservation.
Nitrification, the biological conversion of ammonia to nitrite and nitrate, is a fundamental process of the global nitrogen cycle and must be sustained in engineered and natural environments including those with extreme conditions. Ammonia and nitrite oxidation are accomplished by a number of highly specialized nitrifiers, adapted to a wide temperature spectrum [1, 2]. Described nitrite-oxidizing bacteria (NOB) are scattered among the Alpha-, Beta-, and Gammaproteobacteria [1, 3] or belong to the distinct bacterial phyla Nitrospirae and Nitrospinae [4, 5]. With the isolation of the moderately thermophilic nitrite oxidizer Nitrolancea affiliated with the phylum Chloroflexi , it became obvious that the diversity of NOB is even more complex than previously thought.
Geothermal settings are suitable habitats for a highly active nitrogen cycle and process measurements revealed nitrogen fixation, ammonia oxidation, denitrification and nitrate reduction to ammonium in US hot springs [7, 8]. Chemolithotrophic microorganisms are important primary producers in high-temperature environments  and ammonia oxidation may be a major source for biomass production . The ammonia-oxidizing archaeon (AOA) “Candidatus Nitrosocaldus yellowstonii” with a growth optimum of 72 °C has been isolated from a hot spring in Yellowstone National Park (YNP) . Additional thermophilic AOA from the Nitrosocaldus clade were described recently [12, 13]. Although nitrate formation could be measured in geothermal areas of Iceland and Kamchatka  and calculations of energy yields confirmed that both steps of nitrification are thermodynamically feasible in hot springs , nitrite-oxidizing microorganisms have not yet been identified in these systems. Instead, it was speculated that in geothermal springs nitrite was removed by anaerobic respiration .
Evidence for nitrite oxidation at elevated temperatures was brought by molecular detection and enrichment of Nitrospira, but the temperature optimum of such cultures was in the range of 37–52 °C [15,16,17]. To elucidate the microorganisms responsible for nitrite oxidation in geothermal settings, cultured representatives are valuable to gain a deeper understanding of the physiological and genomic potential of these to date unidentified species. Although the geothermal area in YNP is a “hot spot” for the detection of novel microorganisms [18,19,20], thermophilic NOB with a temperature optimum above 52 °C have not been identified so far. Long-term cultivation and metagenomics listed several bacterial and archaeal candidates for nitrite oxidation in high-temperature bioreactors, but the diverse community impeded identification of single microbes . Just recently, thermophilic, putative nitrite oxidoreductase (NXR)-containing Chloroflexi were enriched along with Nitrosocaldus islandicus . In this study we present first culture-dependent and genome-based insights on microorganisms that drive nitrite oxidation at high temperature.
Materials and methods
Sampling was performed in September 2007 at YNP UWG009 Spring (Lat: 44.4795014, Lon: −110.8518253) and YNP Diadem Spring (Lat: 44.5600075, Lon: −110.8328473) in the Upper and Lower Geyser Basin of YNP. Both non-sulfurous springs are characterized by slightly alkaline to neutral pH values of 8.0 and 7.1 and vent temperatures of 92 °C and 74 °C, respectively (Table S1). UWG009 harbors microbial mats of different colors on the shore (Fig. S1a, b), whereas gray to brownish sediment was sampled from Diadem Spring with a strong acclivity (sampling spots Yell 4–Yell 6; Fig. S1c). The temperatures at the sampling sites were 63 °C and 59 °C, respectively (Table S2).
For enrichment of nitrifying microorganisms, either unbuffered mineral medium for ammonia-oxidizing bacteria (AOB) with 0.5 mM ammonium chloride  or a mineral NOB medium with 0.3 mM, 1 mM, or 3 mM sodium nitrite  was used. Since 2015, the NOB medium was prepared with a modified trace element solution . Half-strength R2A media consisted of 0.25 g yeast extract, 0.25 g proteose peptone, 0.25 g casamino acids, 0.25 g glucose, 0.25 g soluble starch, 0.25 g Na-pyruvate, 0.15 g K2HPO4, 0.025 g MgSO4 × 7H2O, dissolved in 1 L distilled water. The final pH was adjusted to 7.2 with crystalline KH2PO4 and autoclaved for 15 min at 121 °C. R2A plates were prepared by adding 13 g L−1 agarose, nitrite (0.125 mM) and ammonium (1 mM). HWB medium (half-strength) was composed of 0.25 g L−1 peptone, 0.25 g L−1 yeast extract, 0.25 g L−1 meat extract, 0.292 g L−1 sodium chloride, pH 7.3–7.4.
The samples were transported at ambient temperature by courier to the University of Hamburg (Germany). A sample aliquot of 0.5 ml (spring water with biofilm or sediment) was inoculated into 100 ml screw cap Schott bottles, containing 50 ml of mineral medium. Cultures were incubated at 70 °C (±2 °C) and evaporation was compensated for with sterile distilled water. All batch cultures were grown under static conditions. When ammonium was depleted, it was replenished to 400–500 µM with sterile 5 M ammonium chloride solution. Ammonium measurements were done with Quantofix test stripes (Macherey-Nagel, Düren, Germany). The pH value was manually adjusted with 5% NaHCO3 to about 7.4. Cultures were transferred with 1–2% (v/v) inoculum. Since 2011, culturing was continued in 100–300 ml Erlenmeyer flasks at a reduced temperature of 60–65 °C to enhance growth. In Fig. S2, the enrichment procedure is exemplarily shown for culture Yell 2.
In 2015, a follow-up culture of A6 (derived from original culture Yell 2, Fig. S2) was supplemented with nitrite and formate. After 5 months, the nitrifying culture was inoculated into autotrophic NOB medium (containing additionally 0.5 mM ammonium), which was mixed with half-strength R2A medium (initially 1:5 diluted with NOB medium, later 1:10). Subsequently, cultures were fed with nitrite when it was consumed and ammonium (as N-source) when growth ceased.
For electron microscopy, cells were collected, fixed with 2.5% (v/v) glutaraldehyde and 2% (w/v) osmium tetroxide and embedded in a mixture of Spurr and acetone as described . Thin sections were stained with 2% (w/v) uranyl acetate and 2% (w/v) lead citrate. Microscopic examination was carried out with a transmission electron microscope (Zeiss model Leo 906E with a CCD camera model 794). Gamma corrections were applied on pictures of very dark cells. For the visualization of whole cells, 3 µl concentrated biomass was pipetted on EM grids (300 mesh, Stork Veco B.V, The Netherlands) and stained with 2% (w/v) uranyl acetate.
4′−6′-diamidino-2-phenylindole stained cells were observed using a confocal laser scanning microscope LSM 800 equipped with an Airyscan (Zeiss, Jena, Germany) equipped with Plan-Apochromat 63× and 100 × 1.4 oil objectives. Light microscopic images were taken with an AxioScope epifluorescence microscope equipped with a N-Achroplan 100 × 1.25 oil objective and an AxioCam ICc1 1.4-megapixel CCD camera (Zeiss, Jena, Germany).
Fluorescence in situ hybridization
Cells were pelleted at 10 °C and 13,000 × g for 15 min, washed with 0.9% NaCl and fixed with 1:1 (v/v) 96% ethanol:PBS as described previously . To increase permeability, cells were partly treated with Proteinase K (5 mg ml−1) for 30 min at 37 °C and Lysozyme (1 mg ml−1) for 15 min at room temperature. After dehydration in ethanol  cells were hybridized overnight in hybridization buffer containing 20% formamide with the FITC-labeled universal bacterial probe EUB338  and Cy3-labeled probes specific for Ca. Nitrocaldera or Ca. Nitrotheca (Table S3), which were designed as described below. Up to three probes labeled with the same fluorescent dye were used simultaneously to detect Ca. Nitrocaldera to increase signal intensity. Signal specificity was ensured by performing negative control hybridizations using the NON338 probe labeled in Cy3 . Cells were embedded in Citifluor AF1 (Citifluor Ltd, London, UK) prior to microscopic observation at the LSM 800 (Zeiss).
Nitrite and nitrate were determined qualitatively with analytical test stripes (Merck KGaA, Germany) and quantitatively by HPLC via ion pair chromatography on a LiChrospher RP-18 column (125 × 4 mm; Merck) with UV detection in an automated system (Hitachi LaChrom Elite; VWR International GmbH, Darmstadt, Germany). Data acquisition and processing of nitrite and nitrate was performed with the integrated software EZChrom Elite 3.3.2. Ammonium was measured by an ortho-phthalaldehyde fluorescence assay [29, 30]. To test for growth of NOB, nitrite was determined using the Griess–Ilosvay spot test .
DNA for PCR and cloning was isolated using the Ultra-Clean DNA isolation kit (MO BIO Laboratories, Inc. Carlsbad, CA). For Illumina sequencing of culture A5 and A7 DNA was isolated via the RTP® DNA Mini Kit (Stratec molecular, Berlin, Germany) according to the manufacturer’s instruction. DNA of culture A6 was obtained by phenol-chloroform extraction  using TE-sucrose buffer (10 mM Tris, 1 mM Na2EDTA, pH 8.0, 20% (w/v) sucrose) in combination with freeze/thaw cycles (5× liquid nitrogen/65 °C for 5 min). DNA of nitrite-oxidizing cultures was extracted with the PowerSoil® DNA isolation Kit (MO BIO Laboratories, Inc, Carlsbad, CA) according to the manufacturer’s instructions with slight modifications: Before cell disruption via vortexing (step 5) Proteinase K (1 mg ml−1), Lysozyme (4 mg ml−1) and RNase A (1 mg ml−1) were added. The vortexing step was extended to 30 min at 37 °C.
To test for the presence of functional genes of bacterial and archaeal ammonia oxidation, the primer sets amoA-1f/amoA-2r  and Arch-amoAf/Arch-amoAr  were used (Table S3). The presence of NOB belonging to Nitrospira was checked using the 16S rRNA gene-targeted primer set NxrB-169f/NxrB-638r  and of Nitrobacter with the nxrA (encoding NXR subunit A)-targeted primers F1NorA/R1NorA .
Cloning and RFLP analyses
Bacterial 16S rRNA genes were amplified by the bacterial primers 27F and 1492R  (Table S3). The PCR products were ligated into the pGEM-T vector cloning system (Promega, Mannheim, Germany) and transformed into chemically competent Escherichia coli JM109 as described in the manufacturer’s instructions. For partial and near-complete sequencing of clone inserts, the plasmid primers SP6 and T7 were used to reamplify the insert. Restriction digestion was performed with the enzyme HapII. Unique RFLP patterns were identified after separation on a 3% (w/v) agarose gel.
(Semi) specific PCR for thermophilic NOB
Fragments of the nxrA and 16 S rRNA genes of the three putative NOB were amplified using primer sets 1–6 (Tables S3 and S4), which were designed as described below. The following PCR program was used: initial denaturation at 95 °C, 4 min; 36× denaturation at 95 °C, 35 s, annealing at 50 °C, 45 s (primer set 1), elongation at 72 °C, 45 s; final elongation 8 min (primer sets 1, 3, 5) or 12 min (primer sets 2, 4, 6). The annealing temperatures for the other primer sets were 2: 50 °C, 3: 56 °C, 4: 54 °C, 5: 50 °C, and 6: 45 °C.
FISH probe and PCR primer design
16S rRNA-targeted FISH probes and 16S rRNA gene-targeted PCR primers were designed and evaluated using the “design probes” and “match probes” functions of ARB  and the nonredundant SILVA SSURef NR 99 database (releases 123 for probe design and 132 for probe evaluation) . Probe sequences suggested by the probe design tool were checked and refined in the ARB sequence editor against manually curated alignments of all 16S rRNA gene sequences obtained in this study (see below) and a subset of Chloroflexi, including all highly similar sequences present in SILVA SSURef NR 99 release 123 and downloaded from the NCBI nr database, and representative sequences from the more distantly related genera within the phylum. All oligonucleotides were designed to target one of the NOB candidates identified in the enrichment cultures by metagenomic sequencing (see below), and the most closely related sequences present in the dataset (<94% sequence identity and monophyletic clustering in phylogenetic analyses). This rough level of sequence identity was chosen to ensure that the oligonucleotides designed here would also work in case of strain diversity among the enrichment cultures, and to allow future identification of also more distantly related NOB candidates in environmental samples. Probe and primer sensitivity and specificity were queried by matching the newly designed oligonucleotides against the complete SILVA SSURef NR 99 release 132 database allowing for 0 mismatches. Target and nontarget groups were defined based on monophyletic clustering in phylogenetic analyses (see below). All oligonucleotides were ensured to have 100% coverage of the target group, with no perfect-match non-target organism present in the database. Due to the high specificity of the newly designed FISH probes and the lack of pure cultures, no formamide series were performed to determine the optimal formamide concentrations during hybridization. Instead, all probes were employed at 20% formamide in the hybridization buffer (see above).
NxrA-targeted primers were designed manually in the ARB sequence editor. During primer design the main focus was on designing oligonucleotides that specifically distinguish between the NOB candidates present in the enrichments. In case this was not possible, group-specific forward were combined with semi-specific or highly specific reverse primers, which in combination allowed distinguishing between the NOB candidates.
For selected cultures, 10 ng of genomic DNA were used to generate 16S rRNA gene amplicons. For cultures A5, A7, N2a and N2b, amplicons were generated using the primers 341F and 785R , followed by 454 pyrosequencing (300 bp, 3000 reads per sample). For all other cultures the primers 515F and 806R  were used, with subsequent Illumina MiSeq sequencing (2 × 300 bp, 20,000 reads per sample). Operational taxonomic units (OTUs) were obtained from the preprocessed sequences by the qiime (v1.9.1)  de novo OTU picking workflow, summarized and visualized by the qiime summarize_taxa_through_plots.py script. Illumina sequences were processed, classified and summarized by MR DNA and their analysis pipeline (MR DNA, Shallowater, TX, USA) . In short, sequences were joined and depleted of barcodes, followed by removal of sequences < 150 bp or with ambiguous base calls. Sequences were denoised, OTUs generated and chimeras removed. OTUs were defined by clustering at 3% divergence (97% similarity). Final OTUs were taxonomically classified using BLASTn against a curated database derived from NCBI and RDPII (www.ncbi.nlm.nih.gov, http://rdp.cme.msu.edu). 16S rRNA gene amplicon data of the autoclaved culture N7 were obtained from Macrogen Inc. (Seoul, Rep. of Korea) with the bacterial primers 341F and 805R on the Illumina platform (2 × 300 bp, 100,000 reads per sample) with subsequent OTU clustering as described above.
DNA was fragmented using a Bioruptor (Diagenode, Seraing, Belgium). Sequencing libraries were generated using the NEBNext Ultra DNA Library Prep Kit for Illumina (New England Biolabs, Ipswich, USA) as per the manufacturer’s recommendations. Size and quality of the libraries were visualized on a Bioanalyzer High Sensitivity Chip (Agilent Technologies, Santa Clara, USA). Diluted libraries were multiplex sequenced on the Illumina HiSeq 2500 instrument (Illumina, St. Diego, USA) by paired-end sequencing (2 × 100 bp).
Metagenome assembly and binning
Adapter removal, contaminant filtering, and quality trimming of Illumina HiSeq paired-end sequencing reads was performed using BBDUK (BBTOOLS version 37.76) . Terminal base calls with a quality score below Q17 were trimmed. Processed reads with a mean quality score of ≥Q20 and length of ≥70 bp were corrected for sequencing errors using BayesHammer . Corrected reads for all samples were co-assembled using metaSPAdes v3.11.1  with default settings. MetaSPAdes iteratively assembled the metagenome using kmer size of 21, 33, 55, 77, 99, and 127. Trimmed reads were mapped back to the metagenome separately for each sample using Burrows–Wheeler Aligner (BWA 0.7.17) , employing the “mem” algorithm. The sequence mapping files were handled and converted as needed using SAMtools 1.6 . Metagenome binning was performed for contigs greater than 1500 bp using five binning algorithms: BinSanity v0.2.6.1 , COCACOLA , CONCOCT , MaxBin 2.0 2.2.4  and MetaBAT 2 2.12.1 . The five bin sets were supplied to DAS Tool 1.0  for consensus binning to obtain the final optimized bins. The quality of the generated bins was assessed through single-copy marker gene analysis using CheckM 1.0.7 . Subsequently, the binned genomes where annotated using Prokka  against the NCBI RefSeq bacteria_protein database (release 86). Genes of interest where identified by BLAST searches  with selected reference sequences against the nucleotide and protein FASTA output files, or by manual inspection of the Prokka annotation. Functional pathway annotation was performed using the KEGG Automatic Annotation Server .
The absence of additional NXR genes in the unbinned fraction of the metagenome (unbinned contigs ≥1500 bp from low-abundance organisms and contigs <1500 bp) was verified by blasting representative nxrA gene sequences against the complete metagenomic assembly using BLASTN and TBLASTX , and by mapping the corrected reads from all samples against the nxrA gene sequences identified in the binned genomes using BBMap (BBTOOLS version 37.76)  with the minimal alignment similarity cutoff set to 0.7 (parameters minid = 0.70 idfilter = 0.70).
16S rRNA gene sequences present in the genome bins were imported into ARB  in the Silva ARB database SSURef NR 99 release 123 , which was manually updated to contain all highly similar sequences from the NCBI nr database. After automatic alignment (using the Fast align option within the ARB sequence editor) and subsequent manual refinement, a phylogenetic tree was constructed using the PhyML program  implemented in ARB with the GTR nucleotide substitution model, which was identified as best model available in ARB by ModelFinder (http://iqtree.cibiv.univie.ac.at), and 100 nonparametric bootstraps. No conservation filter was applied and only alignment columns without information were filtered out, resulting in 1841 valid alignment positions.
For NxrA analyses, the protein sequences identified in the metagenomic bins were imported and aligned in ARB as described above. Subsequently, phylogenetic trees were calculated using MrBayes  (version 3.2.3, with the parameters aamodelpr = mixed samplefreq = 500 diagnfreq = 5000 nruns = 2 nchains = 4 stoprule = yes stopval = 0.01), and using RAxML  (with the PROTMIX rate distribution and WAG amino acid substitution models and 100 rapid bootstrap runs) and PhyML (with the WAG amino acid substitution model and 100 non-parametric bootstraps) within ARB. The WAG substitution model was identified by MrBayes as the amino acid model with a posterior probability = 1.0. For all NxrA tree calculations a 10% conservation filter was applied, resulting in 1198 valid alignment positions. Partial sequences were omitted from tree calculations.
The phylogenomic tree was reconstructed using UBCG . First, all Chloroflexi (phylum) genomes available in the NCBI RefSeq Assembly database (accession date February 22, 2018) were dereplicated using dREP  (with parameter settings -l 1500000 -comp 90 -con 10 -sa 0.95 -comW 2 -conW 1 -strW 1 -N50W 2 -sizeW 1 --checkM_method taxonomy_wf). Only the best genome assembly was retained for genomes >95% identical. Subsequently, a set of 92 universal bacterial conserved phylogenetic marker proteins was automatically identified, aligned and concatenated in UBCG, followed by tree calculation using RAxML. UBCG also calculated separate trees for each marker protein; the support of each node by these single protein trees is indicated in the final concatenated tree.
Environmental distribution analysis
The integrated microbial next generation sequencing (IMNGS) platform  was used to analyze the environmental distribution of the organisms retrieved via metagenomic sequencing in this study. In short, the 16S rRNA gene sequences extracted from the genome bins were used to query all prokaryotic 16S rRNA amplicon datasets available in the NCBI Sequence Read Archive (SRA), which are integrated into IMNGS as sample-specific sequence databases and OTU-based profiles (accession data: June 16, 2019). This query was performed using 97 and 99% 16S rRNA identity cutoffs. The resulting distribution patterns (expressed in sequence hit counts per amplicon dataset) were summarized to reflect the number of amplicon datasets from a common origin (e.g. hot springs) positive for the respective query sequence. The study origins for studies without automatic classification into one of the main categories within IMNGS were manually curated. Only amplicon datasets with ≥2 query hits were counted as positive. Sequence count summaries were generated in Microsoft Excel using the PivotTable functionalities.
Enrichment of thermophilic NOB with ammonium
Mineral media containing ammonium as sole energy source were used with the initial goal to enrich both, ammonia and nitrite oxidizers, simultaneously. Cultures were inoculated with biofilms and sediments from two hots springs in YNP (UWG009 and Diadem Spring; Fig. S1a–c, Tables S1 and S2). After 6 years of incubation at 65–70 °C and several transfers (Fig. S2) with repeated ammonia consumption (Fig. S3), we obtained a diverse bacterial community in cultures derived from both springs, which mostly consisted of novel members of the phylum Chloroflexi (Fig. 1, Fig. S4, and Table S5). Known AOB were not found and no positive PCR reaction was achieved with the described primer sets for ammonia and nitrite-oxidizing microorganisms (Table S3). The thermophilic AOA Nitrosocaldus yellowstonii  was detected in primary enrichments, but disappeared during further transfers probably due to distinct cultivation conditions. High-throughput metagenomic sequencing revealed candidate genes for the key enzyme of nitrite oxidation (see below), but not for ammonia oxidation. This finding inspired us to initiate nitrite-oxidizing cultures from the ammonia-oxidizing enrichments (Fig. S2) to select for the novel NOB.
Mixotrophic growth of thermophilic NOB
In the beginning of our cultivation attempts, nitrite oxidizers could not reproducibly be grown in standard mineral NOB media. In contrast, oxidation of nitrite to nitrate could be stimulated for unidentified reasons in the cultures that were transferred from AOB medium when ammonium and organic carbon (R2A, 1:5) were added to the mineral NOB medium. Before cells were transferred, the mineral AOB medium had been supplemented with nitrite and formate in order to stimulate growth of mixotrophic NOB . After nitrification was induced, growth of the nitrite-oxidizing consortia was most successful when using a reduced amount of R2A (1:10). At 55 °C and 60 °C, 2.5 mM nitrite was stoichiometrically oxidized to nitrate within 40 days (Fig. 2a), but no nitrite oxidation was observed with highly dilute R2A (1:20) or without R2A. Only little nitrite was consumed using elevated concentrations of organic matter (1:4 or 1:2), which lead to strongly increased growth of heterotrophic bacteria. Tests for optimal growth temperature revealed stoichiometric oxidation of 1 mM nitrite to nitrate in the presence of diluted R2A (1:10) in the range of 50–71 °C with maximum activity at 60 °C (Fig. 2b).
A high-throughput metagenomic sequencing approach included three cultures enriched in mineral AOB medium (A5, A6, and A7) and two cultures grown in mixotrophic NOB medium (N1 and N2a), five cultures which were derived from an initial enrichment inoculated with biofilm sampled in Spring UWG009 (Yell 2; Fig. S2). Co-assembly of the reads from these cultures resulted in a total of 523 contigs (≥1500 bp) with a total length of 16142227 nucleotides (Table S6). The use of a consensus binning approach allowed the retrieval of five distinct metagenome-assembled genomes (MAGs; Table S7). In correlation with 16S rRNA gene amplicon sequencing (Fig. S4) and cloning of the 16S rRNA genes (Table S5), most phylotypes belonged to distinct subgroups, which are (distantly) related to Thermoflexus hugenholtzii  or were identified as members of the genus Thermomicrobium [66, 67]. These thermophilic heterotrophs were originally isolated from sediment of Great Boiling Spring in Nevada (USA) , an alkaline hot spring in YNP  or volcanic soil in Hawaii .
The metagenomes of the nitrite- and ammonia-oxidizing cultures were mostly dominated by a single organism represented by bin_4 (Fig. S5). The 16S rRNA gene sequence of this bacterium, provisionally designated as Candidatus Nitrocaldera robusta, has 86% identity to Thermoflexus hugenholtzii (Fig. 1) and is 99% similar to environmental sequences from Antarctica fumarolic soil . A Thermoflexus-like bacterium (bin_1; 99% 16S rRNA gene identity to T. hugenholtzii) and a Thermomicrobium carboxidum-like bacterium (bin_3; 99% 16S rRNA gene identity to T. carboxidum) were also regularly observed. Similar to bin_4, two additional MAGs (bin_5, named Candidatus Nitrotheca patiens and bin_2, named Candidatus Caldibacter yellowstonii) have no close taxonomically described relatives (Fig. S6). Their closest relatives are T. hugenholtzii (84% 16S rRNA gene identity) and Sphaerobacter thermophilus (85% 16S rRNA gene identity), respectively. Except for Ca. Caldibacter, all phylotypes are highly similar to environmental 16S rRNA gene sequences derived from geothermal settings (Fig. 1).
Genomic identification of putative NOB
Surprisingly, in three of the five MAGs genes encoding the nitrite oxidoreductase complex were detected. This molybdopterin-binding enzyme within the DMSO reductase type II family [69, 70] occurs in two forms: The Nitrobacter-type NXR, including the enzymes of Nitrococcus and Nitrolancea, is characterized by a cytoplasmic orientation of the enzyme complex, while the Nitrospira-type NXR, including the Nitrospina enzyme, is located in the periplasmic space [3, 5, 70]. The NxrA of Nitrobacter, Nitrococcus, and Nitrolancea is related to the alpha subunit of the respiratory membrane-bound nitrate reductase (NarG) of denitrifying bacteria [6, 71], whereas those of Nitrospira and Nitrospina reveal a distinct phylogenetic affiliation with the NxrA of the anaerobic ammonium-oxidizing Brocadiaceae . Recently, an additional distinct type of NXR was described in Nitrotoga, which is distantly related to the enzyme found in Nitrospira and also located in the periplasmic space . In Ca. Nitrocaldera two distinct NXR candidates (Nitrobacter-type and Nitrotoga-type) were found (Fig. 3), with the protein sequence of the first showing 71% identity to the NxrAs of Nitrobacter and Nitrolancea hollandica. The two other MAGs contain only one variant of NXR. Ca. Nitrotheca encodes a Nitrotoga-type, while Thermomicrobium sp. has a Nitrobacter-type NXR with 80% similarity to the enzyme of Ca. Nitrocaldera (Table 1 and Fig. S7a).
In accordance with their affiliation with the Nitrotoga branch (Fig. 3), the second putative NxrA identified in Ca. Nitrocaldera and the enzyme of Ca. Nitrotheca are also predicted to have a periplasmic orientation. They cluster together with molybdopterin oxidoreductases from subsurface metagenomes , which are supposed to originate from autotrophic microorganisms in a thermophilic biofilm (~70 °C) and represent extremophilic “dark matter” . The Nitrotoga-like NxrA-like sequence of Ca. Nitrocaldera is distantly related to a NXR/nitrate reductase of the deeply branching bacterium Candidatus Acetothermum autotrophicum . BLAST analyses identified two partial sequences in the recently described Candidatus Caldiarchaeum subterraneum  as close relative of the NxrA-like protein of Ca. Nitrotheca. These “Aigarchaeota” are a sister lineage to the AOA-containing Thaumarchaeota , but no nitrite-oxidizing potential has been reported to date.
Notably, Ca. Nitrocaldera is the first putative NOB, which contains two different types of NxrA (Table 1 and Fig. 3). It is tempting to speculate that this organism possesses a flexible metabolism with regard to nitrite oxidation, indicating an adaptation to a wide range of nitrite concentrations. Alternatively, only one of the enzymes might be a real NXR, while the second type serves as dedicated nitrate reductase.
Although the Nitrotoga branch of NxrA contains many uncharacterized enzymes also from nitrate-reducing microorganisms, strong support for the nitrite-oxidizing nature of Ca. Nitrotheca was found in an enrichment of Yell 3 (Table S2), which was grown at 55 °C in mixotrophic NOB medium. This culture, consisting of 39% Ca. Nitrotheca, only 5% Ca. Nitrocaldera as second putative NOB and Thermorudis peleae as main heterotroph (Fig. S8a), accumulated 3.4 mM nitrate after the consumption of the same amount of nitrite.
Organization of the NXR operon
In Nitrobacter, the membrane-associated NXR complex consists of the catalytic alpha (NxrA), an electron-transporting beta (NxrB), and a membrane-integral, heme-containing gamma (NxrC) subunit. In addition, the NXR operon encodes a peptidyl-prolyl cis-trans isomerase (NxrX) and a TorD-like chaperone (NxrD) involved in insertion of the molybdopterin cofactor into NxrA. These genes are arranged in a nxrAXBDC order. The operons of the putative Nitrobacter-type NXR in Ca. Nitrocaldera and Thermomicrobium sp. reveal a syntenic genetic organization as found in all other NOB containing this NXR type, with a highly conserved gene cluster furthermore containing a c-type cytochrome and a putative C4-dicarboxylate transporter (Fig. S7a).
Similarly, Ca. Nitrotoga fabula contains genes for the three structural subunits of NXR (NxrABC), as well as a TorD-like chaperone NxrD. Unlike in the Nitrobacter-like enzyme complex, however, the Nitrotoga fabula NxrA contains a N-terminal signal peptide for export into the periplasm via the twin-arginine translocation (TAT) system, and NxrC a signal peptide for translocation via the Sec pathway , as has also been described for Nitrospira defluvii . Furthermore, the NXR gamma subunit does not contain any predicted transmembrane helices, indicating a soluble nature of the enzyme. Contrastingly, while the TAT signal peptide is conserved in the Nitrotoga-type NxrAs of Ca. Nitrocaldera and Ca. Nitrotheca NxrAs, the NxrC subunit of Ca. Nitrocaldera contains one C-terminal transmembrane helix (amino acid positions 304–326) in addition to the N-terminal signal peptide, and the Ca. Nitrotheca NxrC a N-terminal transmembrane helix (amino acid positions 58–80) and no signal peptide. This indicates an anchoring of the NXR complex to the periplasmic face of the cytoplasmic membrane, which might be a necessary adaptation due to the apparent absence of outer membranes in many Chloroflexi . A TorD-like chaperone could not be identified in these genomes (Fig. S7b).
Genomic features of the novel NOB
The total length of the obtained MAGs ranged from 2.4 to 3.6 Mb. The high G + C content of the different genomes (65.2–69.9%; Table S7) is in accordance with known thermophilic Chloroflexi species (64–69%) . The genomes of the two putative NOB Ca. Nitrotheca and Ca. Nitrocaldera are predicted to be near-complete and phylogenetic analyses of concatenated marker protein alignments revealed that these bacteria, along with the nonnitrifying Ca. Caldibacter, form a separate phylogenetic lineage within the phylum Chloroflexi that is distinct from the described species (Fig. S6). Despite their close phylogenetic affiliation, the genome of our Thermoflexus strain shares an average nucleotide identity of 94.8% with Thermoflexus hugenholtzii, which indicates that they form two distinct species .
In Ca. Nitrocaldera, the genetic inventory for CO2 fixation via the Calvin–Benson cycle indicated the potential for autotrophic growth predicted to be ribulose 1,5-bisphosphate carboxylase/oxygenase (Rubisco) with the large subunit (RbcL) having 78% identity to the corresponding enzyme of Synechococcus. Rubisco is also present in Nitrobacter, Nitrococcus, Nitrolancea and Nitrotoga, but not in Nitrospira and Nitrospina, which use the reductive citric acid cycle for carbon fixation [5, 6, 72, 79]. Thus, Ca. Nitrocaldera is a potential chemolithoautotrophic organism, although amendment of organic carbon stimulated nitrite oxidation. It could not be clarified in this study if this effect is due to mixotrophic growth of the novel NOB as described for Nitrobacter  and most other NOB , or if organics mediate the chemical degradation of reactive oxygen species (ROS) against which many nitrifiers are sensitive . Alternatively, organics might have an indirect positive effect by stimulating coexisting heterotrophs, which detoxify ROS or produce metabolites that support growth of the NOB.
In contrast to Ca. Nitrocaldera, Rubisco genes were not identified in the draft genomes of Ca. Nitrotheca or Thermomicrobium sp. Instead, Ca. Nitrotheca encodes all enzymes of the reductive tricarboxylic acid (TCA) cycle and for pyruvate synthesis from acetyl-CoA, with the exception of the key enzyme ATP-citrate lyase. However, a reversibility of citrate synthase was demonstrated recently in Thermosulfidibacter takaii and Desulfurella acetivorans [83, 84], which might also explain the apparent autotrophic potential of Ca. Nitrotheca. Although some reactions of the rTCA cycle are oxygen-sensitive, enzymatic adaptations allow its functioning in aerobic NOB like Nitrospira  as this probably also is the case in nitrite-oxidizing Chloroflexi.
As mentioned above, Ca. Nitrocaldera is related to thermophilic bacteria inhabiting the oligotrophic subsurface of Tramway Ridge, Antarctica (CO2 rich steam fumaroles) . A possible reason for reduced autotrophic growth in our thermophilic cultures is a limitation of carbon dioxide, due to its low solubility at elevated temperatures. However, autotrophic growth was achieved when culture N1 was transferred into mineral NOB medium (culture N7) without R2A, but supplemented with bicarbonate (0.5 mM NaHCO3; Fig. S2). Under these conditions especially the abundance of Ca. Nitrotheca increased, along with Thermoflexus sp. (Fig. S4). In earlier ammonia-oxidizing cultures, amendment of NaHCO3 for pH adjustment might have fulfilled CO2 requirements of the thermophilic NOB, thus allowing lithoautotrophic growth.
Interestingly, all NOB candidates enriched in this study lacked genes for assimilatory nitrite reduction, indicating a dependency on ammonium for growth. The same observation has been made for Nitrolancea hollandica  and thus appears to be a general characteristic of nitrite-oxidizing Chloroflexi and will contribute to their apparent lack of growth in standard NOB medium . A comparison of the main features of Ca. Nitrocaldera, Ca. Nitrotheca and Nitrolancea is shown in Table S8.
Morphological diversity of thermophilic nitrite-oxidizing cultures
Microscopic investigations of nitrite-oxidizing cultures of Yell 2 revealed a strong phenotypic heterogeneity (Fig. S9), but all cultures contained similar morphotypes as seen in the preliminary ammonia-oxidizing enrichments (Fig. S10). Two of the main morphotypes (MT1 and MT3) are very similar in cell shape and ultrastructure to the described species of Thermoflexus  and Thermomicrobium [66, 67], respectively, and MT2 resembles the coccoid cells of Sphaerobacter . Notably, MT4 and MT5 have a unique morphology and ultrastructure (Fig. 4; see supplementary text). They remain the best NOB candidates and their occurrence in nitrite-oxidizing cultures obtained from Yell 2 and Yell 3 was proven by amplicon sequencing and FISH. Probes targeting Ca. Nitrocaldera labeled pleomorphic short rods (Fig. 5a), which gave a strong fluorescent signal at the cell poles (Fig. S11b). Ca. Nitrotheca specific probes labeled ovoid cells and compact short rods in nitrite-oxidizing cultures derived from Yell 2 and Yell 3 (Fig. S12 and S13a).
Putative NOB are heat resistant
During our cultivation approach, the first active nitrite-oxidizing culture N1 was obtained after supplementing the ammonia-oxidizing culture A6 with an aliquot of cell suspension from an ammonia-oxidizing culture inoculated in 2007, which had been “sterilized” by autoclaving (Fig. S2; supplementary text). The goal of this procedure was to possibly add missing trace elements present in the seeding material. Whereas most cells in active cultures containing Ca. Nitrocaldera revealed pleomorphic rods (Fig. S14a), small light-refracting granules were observed in aging cultures (Fig. S14b). This finding indicated the presence of heat-resistant cells and the autoclaving process was repeated with culture N3 (121 °C, 20 min). The autoclaved material was spread on R2A agarose plates (containing nitrite and ammonium) and inoculation of mixotrophic NOB medium with cells from a brownish colony resulted in culture N5. Primarily, Ca. Nitrocaldera was identified by the universal 16S rRNA gene primer set 27F/1492R and by (semi) specific PCR, but after 106 days positive signals for Thermomicrobium sp. were also obtained (Table S4). In subsequent enrichments, they coexisted in mixotrophic NOB medium (cultures N6a + b) and were furthermore accompanied by Geobacillus (Fig. S4). Whereas autoclaving of NOB cultures supplemented with R2A containing glucose selected for the enrichment of Geobacillus, heat treatment of the autotrophic culture N7 proved to be a more appropriate strategy to enrich for the putative NOB. Again, cells of Ca. Nitrocaldera survived, whereas the relative abundance of Ca. Nitrotheca was reduced (Fig. S11a).
Consequently, cells of Ca. Nitrocaldera persisted heat treatment (121 °C) at 1 bar repeatedly, but the mechanism of heat-resistance requires further clarification. One strategy might be the formation of exospores, which has already been observed in the related Chloroflexi Ktedonobacter, Thermosporothrix and Thermogemmatispora . Another survival strategy is the differentiation of vegetative cells into resistant resting structures which are produced under harsh conditions by many non-spore formers. Modified cells of circular shape and reduced size have been shown to be able to persist extreme conditions like wet heat or desiccation [87, 88]. Interestingly, after dormancy with low metabolic activity, Ca. Nitrocaldera depends on the supply of small amounts of organic matter for proliferation and subsequent nitrite oxidation.
Ca. Nitrocaldera appears as main NOB
The molecular, metagenomic, and microscopic results imply that Ca. Nitrocaldera represents the dominant NOB candidate in nitrifying cultures containing ammonium and/or nitrite as energy source (Figs. 5, S4, S5, and Table 1). The abundance of Ca. Nitrotheca declined over time, whereas Ca. Nitrocaldera was maintained during continuous growth in mixotrophic NOB medium (Table S4). Cells retained their nitrite-oxidizing activity with stoichiometric nitrate formation after several transfers (Fig. 2a) and could be enriched to nearly 50% purity when the composition of the mixotrophic medium was modified (Fig. 5). Ca. Nitrocaldera could also be enriched from the second investigated hot spring (Diadem Spring, samples Yell 4 and 5; Table S5) and is supposed to be widely distributed in this thermophilic area.
Environmental distribution of novel chloroflexi
All 16S rRNA amplicon datasets available at NCBI were subsequently screened for the presence of the five novel Chloroflexi species identified by metagenomic sequencing in the ammonia- and nitrite-oxidizing cultures. As expected, sequences closely related to the organisms identified in this study were detected in a range of datasets derived from hot springs and fumarolic sediments (Fig. S15). Also soil and the rhizosphere apparently frequently harbor especially Ca. Nitrotheca and Thermomicrobium-like organisms, and Thermomicrobium furthermore is encountered in many composting systems and activated sludge. Interestingly, this analysis additionally indicated the gut of organisms as diverse as catarina scallops (Argopecten ventricosus), green sea urchins (Lytechinus variegatus), and the house mouse (Mus musculus) as a major habitat of both Ca. Nitrocaldera and Ca. Nitrotheca, but also of Thermoflexus and Ca. Caldibacter, indicating their adaptation to a surprisingly diverse range of ecosystems.
The phylum Chloroflexi is one of the main bacterial groups found in hot springs worldwide  and it also is abundant in various natural and constructed habitats . Our knowledge of the diversity of Chloroflexi has undergone enormous expansion in the last 15 years, revealing diverse physiological features (e.g. photosynthesis, organoheterotrophy or lithoautotrophy) with at least seven subclasses containing both mesophilic and thermophilic representatives .
In this study, two putative nitrite-oxidizing novel Chloroflexi (Ca. Nitrocaldera and Ca. Nitrotheca) were found in nitrifying cultures derived from hot springs in YNP. In contrast, a third NXR-containing bacterium is nearly identical to Thermomicrobium carboxidum , which however may have distinct requirements to consume nitrite compared to Ca. Nitrocaldera. Alternatively, the NXR-like enzyme might function as canonical nitrate reductase in Thermomicrobium sp.
The community of thermophilic NOB and associated heterotrophic bacteria investigated here represents a dynamic system, which is able to adapt to varying culture conditions. They revealed morphological (Figs. S16–S19) as well as physiological flexibility with a proposed potential for autotrophic, mixotrophic and heterotrophic growth. Nevertheless, the complex community consisting of five different Chloroflexi was a very stable consortium adapted to limiting concentrations of nutrients and oxygen, resembling the natural environmental conditions. This community could be maintained for 12 years in mineral AOB or NOB medium.
Both, the putative nitrite oxidizers and the accompanying bacteria belong to the phylum Chloroflexi. Thermoflexus sp., Thermomicrobium sp. and Ca. Caldibacter are assumed to be heterotrophic, but also exist under lithoautotrophic conditions like in initial ammonia-oxidizing cultures. In contrast, the putative NOB Ca. Nitrocaldera and Ca. Nitrotheca appear to have the ability for autotrophic growth, but prefer an ancient chemolithomixotrophic life style . Their abundance increased in incubation strategies similar to culture N1 (Table S4), supplemented once with organic matter and subsequently only supplied with nitrite and ammonium. Ca. Nitrocaldera represents the first heat-resistant nitrite oxidizer, a survival strategy enabling existence in hot springs and volcanic soils. The growth temperature optimum and maximum of 60 °C and 71 °C, respectively, are clearly above those of the thermophilic members of Nitrospira (growth temperature limit 60–65 °C) [15, 16]. A putative thermophilic NOB with nxrAB genes within the Nitrobacter/Nitrolancea cluster was already enriched in co-culture with an ammonia-oxidizing Thaumarchaeon from a hot spring in Iceland , which supports our finding that the phylum Chloroflexi contains several novel NOB.
Filamentous Chloroflexi provide a matrix in which other cells can become stably embedded  and occurrence in multicellular aggregates cause enormous difficulties in isolation procedures. Another reason for the failure of isolation of Ca. Nitrocaldera might be its dependency on a (facultatively) heterotrophic bacterium and complex interactions in the nitrifying consortia are hypothesized. Nitrite was only oxidized in co-culture with Thermomicrobium sp. and/or Thermoflexus sp., which were consistently detected in the nitrifying enrichments since 2007. In some cultures, putative NOB were also accompanied by spore forming members of the genus Geobacillus, which sporadically proliferated when R2A was added and are known to stimulate thermophilic bacteria by CO2 supply . So far, no nitrite-oxidizing pure culture could be obtained.
Dependency on other members of the natural community is a well-known finding in hot springs  and might be due to growth on low-molecular weight organic compounds or other metabolites excreted by others . Bacterial interactions are in the focus of new cultivation concepts  and development of new media and isolation strategies are required. Nevertheless, by the design of novel (semi) specific primer sets, it will now be possible to detect these (and other) new NOB in environmental samples, e.g. from volcanic systems. These reference cultures are of high scientific value to study the diversity and adaptations to low nutrient availability and extreme conditions as well as evolutionary processes, as hot spring conditions are typical to the early environment of the planet earth .
Taxonomic consideration of “Candidatus Nitrocaldera robusta” gen. nov. sp. nov
L. n. nitrum: nitrate, N.L. fem. n. caldera: from the Spanish n. caldera, cauldron; designating a volcanic crater. L. fem. adj. robusta: strong. A robust (autoclaving-resistant) nitrate-forming bacterium from a volcanic caldera. Phylogenetically affiliated with the phylum Chloroflexi.
Pleomorphic rods 0.3 × 0.5–3 µm with an electron-dense cytoplasm, Gram-positive. Lateral vesicle extrusion, produces aggregates, formation of dwarf cells, which are connected by fibers. Aerobic, facultative chemolithoautotroph that oxidize nitrite (0.3–5 mM) to nitrate and use carbon dioxide as carbon source. Growth is stimulated by organic compounds, requires ammonium supplements. Obligately thermophilic with a temperature range of 50–71 °C and an optimum of 60 °C; Heat resistant, survives autoclaving at 121 °C. Prefers neutral pH, but tolerates slightly acidic conditions (pH 5.5). The DNA G + C content is 67 mol%.
Taxonomic consideration of “Candidatus Nitrotheca patiens” gen. nov. sp. nov
L. n. nitrum: nitrate, L.n. theca: capsule, L. adj. patiens: persistent. A perennial, spiny EPS covered putative nitrite-oxidizing bacterium within the phylum Chloroflexi.
Ovoid cells (0.2 × 0.3 µm) extending to rods (up to 1.5 µm) with an electron-dense cytoplasm, Gram-positive. Covered by a regular protein layer and crystal-like extracellular polymeric substances. Aerobic, obligate chemolithoautotroph that oxidizes nitrite to nitrate, requires ammonium supplementation. Obligately thermophilic, grows at 55–70 °C. The DNA G + C content is 69 mol%.
Taxonomic consideration of “Candidatus Caldibacter yellowstonii” gen. nov. sp. nov
L. adj. caldus: hot, N. L. masc. n. bacter: a rod, N. L. masc. adj. yellowstonii: pertaining to the habitat of the type strain, the YNP. Phylogenetically affiliated with the phylum Chloroflexi.
A thermophilic bacterium that oxidizes organic substances at neutral pH and temperatures between 55–70 °C. The DNA G + C content is 70 mol%.
Metagenomic and 16S rRNA gene amplicon sequence data are available in the European Nucleotide Archive (ENA) under accession number PRJEB28556. Requests for bacterial enrichment cultures are subject to the “General Permit Conditions” and cannot be shared by the authors without permission from U.S. National Park Service.
Alawi M, Lipski A, Sanders T, Eva-Maria-Pfeiffer, Spieck E. Cultivation of a novel cold-adapted nitrite oxidizing betaproteobacterium from the Siberian Arctic. ISME J. 2007;1:256–64.
Reigstad LJ, Richter A, Daims H, Urich T, Schwark L, Schleper C. Nitrification in terrestrial hot springs of Iceland and Kamchatka. FEMS Microbiol Ecol. 2008;64:167–74.
Spieck E, Bock E. The lithoautotrophic nitrite-oxidizing bacteria. In: Garrity G, Brenner DJ, Krieg NR, Staley JT, editors. Bergey’s manual of systematic bacteriology. Berlin/Heidelberg, Germany: Springer-Verlag; 2005. p. 149–53.
Ehrich S, Behrens D, Lebedeva E, Ludwig W, Bock E. A new obligately chemolithoautotrophic, nitrite-oxidizing bacterium, Nitrospira moscoviensis sp. nov. and its phylogenetic relationship. Arch Microbiol. 1995;164:16–23.
Lücker S, Nowka B, Rattei T, Spieck E, Daims H. The genome of Nitrospina gracilis illuminates the metabolism and evolution of the major marine nitrite oxidizer. Front Microbiol. 2013;4:3–19.
Sorokin DY, Lücker S, Vejmelkova D, Kostrikina NA, Kleerebezem R, WIC Rijpstra, et al. Nitrification expanded: discovery, physiology and genomics of a nitrite-oxidizing bacterium from the phylum Chloroflexi. ISME J. 2012;6:2245–56.
Hedlund B, Thomas S, Dodsworth J, Zhang C. Life in high-temperature environments. In: Yates M, Nakatsu C, Miller R, Pillai S, editors. Manual of environmental microbiology, 4th ed. Washington, DC., USA: ASM Press; 2016. p. 4.3.4-1–4.3.4-15. https://doi.org/10.1128/9781555818821.ch4.3.4.
Dodsworth J, Hungate B, Hedlund BP. Ammonia oxidation, denitrification and dissimilatory nitrate reduction to ammonium in two US Great Basin hot springs with abundant ammonia-oxidizing archaea. Environ Microbiol. 2011;13:2371–86.
Inskeep WP, Ackerman GG, Taylor WP, Kozubal M, Korf S, Macur RE. On the energetics of chemolithotropy in nonequilibrium systems: case studies of geothermal springs in Yellowstone National Park. Geobiology. 2005;3:297–317.
Dodsworth JA, McDonald AI, Hedlund BP. Calculation of total free energy yield as an alternative approach for predicting the importance of potential chemolithotrophic reactions in geothermal springs. FEMS Microbiol Ecol. 2012;81:446–54.
de la Torre JR, Walker CB, Ingalls AE, Könneke M, Stahl DA. Cultivation of a thermophilic ammonia oxidizing archaeon synthesizing crenarchaeol. Environ Microbiol. 2008;10:810–8.
Abby SS, Melcher M, Kerou M, Krupovic M, Stieglmeier M, Rossel C, et al. Candidatus Nitrosocaldus cavascurensis, an ammonia oxidizing, extremely thermophilic archaeon with a highly mobile genome. Front Microbiol. 2018;9:1–19.
Daebeler A, Herbold CW, Vierheilig J, Sedlacek CJ, Pjevac P, Albertsen M, et al. Cultivation and genomic analysis of ‘Candidatus Nitrosocaldus islandicus,’ an obligately thermophilic, ammonia-oxidizing thaumarchaeon from a hot spring biofilm in Graendalur valley, Iceland. Front Microbiol. 2018;9:1–16.
Becraft ED, Dodsworth JA, Murugapiran SK, Ohlsson JI, Briggs BR, Kanbar J, et al. Single-cell-genomics-facilitated read binning of candidate phylum EM19 genomes from geothermal spring metagenomes. Appl Environ Microbiol. 2016;82:992–1003.
Lebedeva EV, Off S, Zumbrägel S, Kruse M, Shagzhina A, Lücker S, et al. Isolation and characterization of a moderately thermophilic nitrite-oxidizing bacterium from a geothermal spring. FEMS Microbiol Ecol. 2011;75:195–204.
Edwards TA, Calica NA, Huang DA, Manoharan N, Hou W, Huang L, et al. Cultivation and characterization of thermophilic Nitrospira species from geothermal springs in the US Great Basin, China, and Armenia. FEMS Microbiol Ecol. 2013;85:283–92.
Kits KD, Sedlacek CJ, Lebedeva EV, Han P, Bulaev A, Pjevac P, et al. Kinetic analysis of a complete nitrifier reveals an oligotrophic lifestyle. Nature. 2017;549:269–72.
Hugenholtz P, Pitulle C, Hershberger KL, Pace NR. Novel division level bacterial diversity in a Yellowstone hot spring. J Bacteriol. 1998;180:366–76.
Meyer-Dombard DR, Shock EL, Amend JP. Archaeal and bacterial communities in geochemically diverse hot springs of Yellowstone National Park, USA. Geobiology. 2005;3:211–27.
Hedlund BP, Murugapiran SK, Alba TW, Levy A, Dodsworth JA, Goertz GB, et al. Uncultivated thermophiles: current status and spotlight on ‘Aigarchaeota’. Curr Opin Microbiol. 2015;25:136–45.
Kato S, Sakai S, Hirai M, Tasumi E, Nishizawa M, Suzuki K, et al. Long-term cultivation and metagenomics reveal ecophysiology of previously uncultivated Thermophiles involved in biogeochemical nitrogen cycle. Microbes Environ. 2018;33:107–10.
Krümmel A, Harms H. Effect of organic matter on growth and cell yield of ammonia-oxidizing bacteria. Arch Microbiol. 1982;133:50–54.
Spieck E, Lipski A. Cultivation, growth physiology, and chemotaxonomy of nitrite-oxidizing bacteria. In: Klotz MG, editor. Methods in enzymology. 1st ed. Oxford, UK: Academic Press/Elsevier Inc.; 2011. p. 109–30.
Widdel F, Bak F. Gram-negative mesophilic sulfate-reducing bacteria. The Prokaryotes. New York, NY, USA: Springer New York; 1992. p. 3352–78.
Amann R, Ludwig W, Schleifer K. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Rev. 1995;59:143–69.
Manz W, Amann R, Ludwig W, Wagner M, Schleifer K-H. Phylogenetic oligodeoxynucleotide probes for the major subclasses of Proteobacteria: problems and solutions. Syst Appl Microbiol. 1992;15:593–600.
Amann RI, Binder BJ, Olson RJ, Chisholm SW, Devereux R, Stahl DA. Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Appl Environ Microbiol. 1990;56:1919–25.
Wallner G, Amann R, Beisker W. Optimizing fluorescent in situ hybridization with rRNA‐targeted oligonucleotide probes for flow cytometric identification of microorganisms. Cytometry. 1993;14:136–43.
Taylor S, Ninjoor V, Dowd DM, Tappel AL. Cathepsin B2 measurement by sensitive fluorometric ammonia analysis. Anal Biochem. 1974;60:153–62.
Corbin JL. Liquid chromatographic-fluorescence determination of ammonia from nitrogenase reactions: A 2 min assay. Appl Environ Microbiol. 1984;47:1027–30.
Schmidt EL, Belser LW. Autotrophic nitrifying bacteria. In: Weaver RW, Angle JS, Bottomley PJ, editors. Methods of soil analysis. Part 2-microbiological and biochemical properties. Madison, WI, USA: Soil Science Society of America; 1994. p. 159–77.
Sambrook J, Green MR, editors. Extracting DNA from gram-negative bacteria. Molecular cloning: a laboratory manual, 4th ed. New York, NY: Cold Spring Harbor Laboratory Press; 2012 p. 19–20.
Rotthauwe J, Witzel K, Liesack W. The ammonia monooxygenase structural gene amoA as a functional marker: molecular fine-scale analysis of natural ammonia-oxidizing populations. Appl Environ Microbiol. 1997;63:4704–12.
Francis CA, Roberts KJ, Beman JM, Santoro AE, Oakley BB. Ubiquity and diversity of ammonia-oxidizing archaea in water columns and sediments of the ocean. Proc Natl Acad Sci USA. 2005;102:14683–8.
Pester M, Maixner F, Berry D, Rattei T, Koch H, Lücker S, et al. NxrB encoding the beta subunit of nitrite oxidoreductase as functional and phylogenetic marker for nitrite-oxidizing Nitrospira. Environ Microbiol. 2014;16:3055–71.
Poly F, Wertz S, Brothier E, Degrange V. First exploration of Nitrobacter diversity in soils by a PCR cloning-sequencing approach targeting functional gene nxrA. FEMS Microbiol Ecol. 2008;63:132–40.
Lane DJ. 16/23S rRNA sequencing. In: Stackebrandt E, Goodfellow M, editors. Nucleic acid techniques in bacterial systematics. Chichester, UK: John Wiley & Sons; 1991. p. 115–71.
Ludwig W, Strunk O, Westram R, Richter L, Meier H, Yadhukumar A, et al. ARB: a software environment for sequence data. Nucleic Acids Res. 2004;32:1363–71.
Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2013;41:D590–D596.
Herlemann DP, Labrenz M, Jürgens K, Bertilsson S, Waniek JJ, Andersson AF. Transitions in bacterial communities along the 2000 km salinity gradient of the Baltic Sea. ISME J. 2011;5:1571–9.
Caporaso JG, Lauber CL, Walters WA, Berg-Lyons D, Lozupone CA, Turnbaugh PJ, et al. Global patterns of 16S rRNA diversity at a depth of millions of sequences per sample. Proc Natl Acad Sci. 2011;108:4516–22.
Caporaso JG, Kuczynski J, Stombaugh J, Bittinger K, Bushman FD, Costello EK, et al. QIIME allows analysis of high-throughput community sequencing data. Nat Methods. 2010;7:335–6.
Dowd SE, Sun Y, Wolcott RD, Domingo A, Carroll JA. Bacterial tag–encoded FLX amplicon pyrosequencing (bTEFAP) for microbiome studies: Bacterial diversity in the ileum of newly weaned Salmonella-infected pigs. Foodborne Pathog Dis. 2008;5:459–72.
Bushnell B. BBMap short read aligner. Berkeley, CA.: University of California; 2016. http://sourceforge.net/projects/bbmap.
Nikolenko SI, Korobeynikov AI, Alekseyev MA. BayesHammer: Bayesian clustering for error correction in single-cell sequencing. BMC Genomics. 2013;14:S7.
Nurk S, Meleshko D, Korobeynikov A, Pevzner PA. MetaSPAdes: a new versatile metagenomic assembler. Genome Res. 2017;27:824–34.
Li H, Durbin R. Fast and accurate long-read alignment with Burrows-Wheeler transform. Bioinformatics. 2010;26:589–95.
Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, et al. The sequence alignment/map format and SAMtools. Bioinformatics. 2009;25:2078–9.
Graham ED, Heidelberg JF, Tully BJ. BinSanity: unsupervised clustering of environmental microbial assemblies using coverage and affinity propagation. PeerJ. 2017;5:e3035.
Lu YY, Chen T, Fuhrman JA, Sun F, Sahinalp C. COCACOLA: binning metagenomic contigs using sequence COmposition, read CoverAge, CO-alignment and paired-end read LinkAge. Bioinformatics. 2017;33:791–8.
Alneberg J, Bjarnason BS, De Bruijn I, Schirmer M, Quick J, Ijaz UZ, et al. Binning metagenomic contigs by coverage and composition. Nat Methods. 2014;11:1144–6.
Wu YW, Simmons BA, Singer SW. MaxBin 2.0: An automated binning algorithm to recover genomes from multiple metagenomic datasets. Bioinformatics. 2015;32:605–7.
Kang DD, Froula J, Egan R, Wang Z. MetaBAT, an efficient tool for accurately reconstructing single genomes from complex microbial communities. PeerJ. 2015;3:e1165.
Sieber CMK, Probst AJ, Sharrar A, Thomas BC, Hess M, Tringe SG, et al. Recovery of genomes from metagenomes via a dereplication, aggregation and scoring strategy. Nat Microbiol. 2018;3:836–43.
Parks DH, Imelfort M, Skennerton CT, Hugenholtz P, Tyson GW. CheckM: assessing the quality of microbial genomes recovered from isolates, single cells, and metagenomes. Genome Res. 2015;25:1043–55.
Seemann T. Prokka: rapid prokaryotic genome annotation. Bioinformatics. 2014;30:2068–9.
Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ. Basic local alignment search tool. J Mol Biol. 1990;215:403–10.
Moriya Y, Itoh M, Okuda S, Yoshizawa AC, Kanehisa M. KAAS: an automatic genome annotation and pathway reconstruction server. Nucleic Acids Res. 2007;35:W182–W185.
Guindon S, Delsuc F, Dufayard JF, Gascuel O. Estimating maximum likelihood phylogenies with PhyML. Methods Mol Biol. 2009;537:113–37.
Ronquist F, Huelsenbeck JP. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioinformatics. 2003;19:1572–4.
Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics. 2014;30:1312–3.
Na SI, Kim YO, Yoon SH, Ha Smin, Baek I, Chun J. UBCG: Up-to-date bacterial core gene set and pipeline for phylogenomic tree reconstruction. J Microbiol. 2018;56:281–5.
Olm MR, Brown CT, Brooks B, Banfield JF. DRep: a tool for fast and accurate genomic comparisons that enables improved genome recovery from metagenomes through de-replication. ISME J. 2017;11:2864–8.
Lagkouvardos I, Joseph D, Kapfhammer M, Giritli S, Horn M, Haller D, et al. IMNGS: a comprehensive open resource of processed 16S rRNA microbial profiles for ecology and diversity studies. Sci Rep. 2016;6:33721.
Dodsworth JA, Gevorkian J, Despujos F, Cole JK, Murugapiran SK, Ming H, et al. Thermoflexus hugenholtzii gen. nov., sp. nov., a thermophilic, microaerophilic, filamentous bacterium representing a novel class in the Chloroflexi, Thermoflexia classis nov., and description of Thermoflexaceae fam. nov. and Thermoflexales ord. nov. Int J Syst Evol Microbiol. 2014;64:2119–27.
Jackson TJ, Ramaley RF, Meinschein WG. Thermomicrobium, a new genus of extremely thermophilic bacteria. Int J Syst Bacteriol. 1973;23:28–36.
King CE, King GM. Thermomicrobium carboxidum sp. nov., and Thermorudis peleae gen. nov., sp. nov., carbon monoxide-oxidizing bacteria isolated from geothermally heated biofilms. Int J Syst Evol Microbiol. 2014;64:2586–92.
Herbold CW, Lee CK, McDonald IR, Cary SC. Evidence of global-scale aeolian dispersal and endemism in isolated geothermal microbial communities of Antarctica. Nat Commun. 2014;5:1–10.
Meincke M, Bock E, Kastrau D, Kroneck PMH. Nitrite oxidoreductase from Nitrobacter hamburgensis: redox centers and their catalytic role. Arch Microbiol. 1992;158:127–31.
Lücker S, Wagner M, Maixner F, Pelletier E, Koch H, Vacherie B, et al. A Nitrospira metagenome illuminates the physiology and evolution of globally important nitrite-oxidizing bacteria. Proc Natl Acad Sci. 2010;107:13479–84.
Kirstein K, Bock E. Close genetic relationship between Nitrobacter hamburgensis nitrite oxidoreductase and Escherichia coli nitrate reductases. Arch Microbiol. 1993;160:447–53.
Kitzinger K, Koch H, Lücker S, Sedlacek CJ, Herbold C, Schwarz J, et al. Characterization of the first “Candidatus Nitrotoga” isolate reveals metabolic versatility and separate evolution of widespread nitrite-oxidizing bacteria. mBio. 2018;9:1–16.
Anantharaman K, Brown CT, Hug LA, Sharon I, Castelle CJ, Probst AJ, et al. Thousands of microbial genomes shed light on interconnected biogeochemical processes in an aquifer system. Nat Commun. 2016;7:1–11.
Hedlund BP, Dodsworth JA, Murugapiran SK, Rinke C, Woyke T. Impact of single-cell genomics and metagenomics on the emerging view of extremophile “microbial dark matter”. Extremophiles. 2014;18:865–75.
Takami H, Noguchi H, Takaki Y, Uchiyama I, Toyoda A, Nishi S, et al. A deeply branching thermophilic bacterium with an ancient acetyl-CoA pathway dominates a subsurface ecosystem. PLoS ONE. 2012;7.
Nunoura T, Takaki Y, Kakuta J, Nishi S, Sugahara J, Kazama H, et al. Insights into the evolution of Archaea and eukaryotic protein modifier systems revealed by the genome of a novel archaeal group. Nucleic Acids Res. 2011;39:3204–23.
Hugenholtz P, Stackebrandt E. Reclassification of Sphaerobacter thermophilus from the subclass Sphaerobacteridae in the phylum Actinobacteria to the class Thermomicrobia (emended description) in the phylum Chloroflexi (emended description). Int J Syst Evol Microbiol. 2004;54:2049–51.
Rodriguez-R LM, Konstantinidis KT. The enveomics collection: a toolbox for specialized analyses of microbial genomes and metagenomes. 2016.
Daims H, Lücker S, Le Paslier D, Wagner M. Diversity, environmental genomics, and ecophysiology of nitrite-oxidizing bacteria. In: Ward BB, Klotz MG, Arp DJ, editors. Nitrification. Washington, DC, USA: ASM Press; 2011. p. 295–322.
Bock E. Growth of Nitrobacter in the presence of organic matter. Arch Microbiol. 1976;108:305–12.
Lipski A, Spieck E, Makolla A, Altendorf K. Fatty acid profiles of nitrite-oxidizing bacteria reflect their phylogenetic heterogeneity. Syst Appl Microbiol. 2001;24:377–84.
Kim J-G, Park S-J, Sinninghe Damsté JS, Schouten S, Rijpstra WIC, Jung M-Y, et al. Hydrogen peroxide detoxification is a key mechanism for growth of ammonia-oxidizing archaea. Proc Natl Acad Sci. 2016;113:7888–93.
Nunoura T, Chikaraishi Y, Izaki R, Suwa T, Sato T, Harada T, et al. A primordial and reversible TCA cycle in a facultatively chemolithoautotrophic thermophile. Science. 2018;359:559–63.
Mall A, Sobotta J, Huber C, Tschirner C, Kowarschik S, Bačnik K, et al. Reversibility of citrate synthase allows autotrophic growth of a thermophilic bacterium. Science. 2018;359:563–7.
Pati A, la Butti K, Pukall R, Nolan M, del Rio TG, Tice H, et al. Complete genome sequence of Sphaerobacter thermophilus type strain (S 6022 T). Stand Genom Sci. 2010;2:49–56.
Yabe S, Aiba Y, Sakai Y, Hazaka M, Yokota A. A life cycle of branched aerial mycelium- and multiple budding spore-forming bacterium Thermosporothrix hazakensis belonging to the phylum Chloroflexi. J Gen Appl Microbiol. 2010;56:137–41.
Filippidou S, Junier T, Wunderlin T, Kooli WM, Palmieri I, Al-Dourobi A, et al. Adaptive strategies in a poly-extreme environment: differentiation of vegetative cells in Serratia ureilytica and resistance to extreme conditions. Front Microbiol. 2019;10:1–13.
Suzina NE, Mulyukin AL, Kozlova AN, Shorokhova AP, Dmitriev VV, Barinova ES, et al. Ultrastructure of resting cells of some non-spore-forming bacteria. Microbiology. 2004;73:435–47.
Meyer-Dombard DR, Swingley W, Raymond J, Havig J, Shock EL, Summons RE. Hydrothermal ecotones and streamer biofilm communities in the Lower Geyser Basin, Yellowstone National Park. Environ Microbiol. 2011;13:2216–31.
Yamada T, Sekiguchi Y. Cultivation of uncultured chloroflexi subphyla: significance and ecophysiology of formerly uncultured chloroflexi ‘subphylum i’ with natural and biotechnological relevance. Microbes Environ. 2009;24:205–16.
Miller SR, Strong AL, Jones KL, Ungerer MC. Bar-coded pyrosequencing reveals shared bacterial community properties along the temperature gradients of two alkaline hot springs in Yellowstone National Park. Appl Environ Microbiol. 2009;75:4565–72.
Watsuji T, Kato T, Ueda K, Beppu T. CO2 supply induces the growth of Symbiobacterium thermophilum, a syntrophic bacterium. Biosci Biotechnol Biochem. 2006;70:753–6.
Stewart EJ. Growing unculturable bacteria. J Bacteriol. 2012;194:4151–60.
Overmann J, Abt B, Sikorski J. Present and future of culturing bacteria. Annu Rev Microbiol. 2017;71:711–30.
Inskeep W. The YNP metagenome project: environmental parameters responsible for microbial distribution in the Yellowstone geothermal ecosystem. Front Microbiol. 2013;4:1–15.
The authors thank the National Park Service for permission to perform research in Yellowstone National Park (Permit YELL-2007-SCI-5698). Elke Woelken is acknowledged for excellent technical help in electron microscopy and Yvonne Bedarf and Christina Bietz for initial cultivation. We also thank Lia Burkhardt and Kerstin Reumann for sequencing assistance and we are grateful to Ilias Lagkouvardos and Antonios Kioukis for kindly enabling our analyses on the IMNGS platform.
This work was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation grant SP 667/7-1+2) and the Netherlands Organization for Scientific Research (Grants 863.14.019, 016.Vidi.189.050 and SIAM Gravitation Grant 024.002.002).
Conflict of interest
The authors declare that they have no conflict of interest.
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
About this article
Cite this article
Spieck, E., Spohn, M., Wendt, K. et al. Extremophilic nitrite-oxidizing Chloroflexi from Yellowstone hot springs. ISME J 14, 364–379 (2020). https://doi.org/10.1038/s41396-019-0530-9
Applied Microbiology and Biotechnology (2021)