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Environmental Microbiology (2014) 16(10), 3263–3274

doi:10.1111/1462-2920.12546

The nitrate-sensing NasST system regulates nitrous oxide reductase and periplasmic nitrate reductase in Bradyrhizobium japonicum

Cristina Sánchez,1† Manabu Itakura,1† Takashi Okubo,1 Takashi Matsumoto,2 Hirofumi Yoshikawa,2,3 Aina Gotoh,4 Masafumi Hidaka,4 Takafumi Uchida4 and Kiwamu Minamisawa1* 1 Graduate School of Life Sciences, Tohoku University, 2-1-1 Katahira, Aoba-ku, Sendai, 980-8577, Japan. 2 Department of Bioscience and 3Genome Research Center, NODAI Research Institute, Tokyo University of Agriculture, 1-1-1 Sakuragaoka, Setagaya-ku, Tokyo, 156-8502, Japan. 4 Graduate School of Agricultural Science, Tohoku University, 1-1 Amamiya-machi, Tsutsumidori, Aoba-ku, Sendai, 981-8555, Japan. Summary The soybean endosymbiont Bradyrhizobium japonicum is able to scavenge the greenhouse gas N2O through the N2O reductase (Nos). In previous research, N2O emission from soybean rhizosphere was mitigated by B. japonicum Nos++ strains (mutants with increased Nos activity). Here, we report the mechanism underlying the Nos++ phenotype. Comparative analysis of Nos++ mutant genomes showed that mutation of bll4572 resulted in Nos++ phenotype. bll4572 encodes NasS, the nitrate (NO3−)-sensor of the two-component NasST regulatory system. Transcriptional analyses of nosZ (encoding Nos) and other genes from the denitrification process in nasS and nasST mutants showed that, in the absence of NO3−, nasS mutation induces nosZ and nap (periplasmic nitrate reductase) via nasT. NO3− addition dissociated the NasS-NasT complex in vitro, suggesting the release of the activator NasT. Disruption of nasT led to a marked decrease in nosZ and nap transcription in cells incubated in the presence of NO3−. Thus, although NasST is known to regulate the NO3−mediated response of NO3− assimilation genes in

Received 29 April, 2014; accepted 13 June, 2014. *For correspondence. E-mail [email protected]; Tel. (+81) 22 217 5684; Fax (+81) 22 217 5684. †C.S. and M.I. contributed equally to this work.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd

bacteria, our results show that NasST regulates the NO3−-mediated response of nosZ and napE genes, from the dissimilatory denitrification pathway, in B. japonicum. Introduction Nitrous oxide (N2O) is a powerful greenhouse gas with an atmospheric lifetime of 114 years. Its global warming potential is 300 times that of the equivalent amount of carbon dioxide (CO2) (Thomson et al., 2012). N2O is also the major ozone-depleting substance in the stratosphere (Ravishankara et al., 2009). Soils can act as both a source of N2O and a sink, but on a global scale, the source activity largely dominates the sink activity, accounting for 70% of the atmospheric loading of N2O (Hénault et al., 2012). N2O is produced in the soil during several microbial processes, but mainly through denitrification, as an intermediate product, and by nitrification, as a by-product of ammonium oxidation (Baggs, 2011). Reduction of N2O to N2 by N2O reductase (Nos) is currently the only known pathway for the removal of N2O from soils and water (Thomson et al., 2012). Nos is encoded by nosZ and is typically found in denitrifying bacteria that reduce N2O to N2. Recently, it has been proposed that abundant groups of non-denitrifying N2O reducers with atypical Nos are potential contributors to N2O reduction in a broad range of habitats (Sanford et al., 2012). Soybean is a globally important leguminous crop. Soybean hosts endosymbiotic N2-fixing soil bacteria (rhizobia) that can both produce and reduce N2O through denitrification. Denitrification, the dissimilatory reduction of NO3– (nitrate) to N2, in the soybean endosymbiont Bradyrhizobium japonicum USDA110 requires four enzymes: periplasmic nitrate reductase (Nap), nitrite reductase (NirK), nitric oxide reductase (Nor) and Nos. The enzymes and accessory functions are encoded by the napEDABC, nirK, norCBQD and nosRZDYFLX gene clusters respectively (Bedmar et al., 2005). As in many other denitrifiers, full expression of the denitrification genes in strain USDA110 requires both oxygen limitation and the presence of an N oxide. Microaerobic induction of nap, nirK, nor, and nos depends on the FixLJ-FixK2

3264 C. Sánchez et al. regulatory cascade (Velasco et al., 2004; Bueno et al., 2012). FixLJ is a two-component regulatory system consisting of the heme-based sensor kinase FixL and the FixJ response regulator. FixK2 belongs to the bacterial family of FNR/CRP-type transcriptional regulators and activates nap, nirK, and nnrR (Bueno et al., 2012 and references therein). The FNR-like activator NnrR has been proposed to activate nor genes in response to an N oxide (Bueno et al., 2012 and references therein). The low oxygenresponsive NifA protein, which activates the transcription of essential N2-fixation genes, is also involved in the expression of nap, nirK and nor (Bueno et al., 2012 and references therein). In soybean rhizosphere, a large proportion of N2O is emitted during degradation of the root nodules, so N2O emissions often peak before or after the harvest period (Yang and Cai, 2005; Inaba et al., 2009; 2012). Sameshima-Saito and colleagues showed that soybean roots nodulated with B. japonicum Nos+ strains (+, that carry nos genes) could scavenge N2O in the soil, and thereby lessen N2O emission from soybean fields to the atmosphere (Sameshima-Saito et al., 2006). This offered a promising perspective for the mitigation of N2O emission, especially in soils that are inefficient in reducing N2O, as in the case when native strains are nosZ− dominant and lack nosZ (Sameshima-Saito et al., 2006; Hénault et al., 2012). In a previous study, we generated Nos++ strains (mutants with increased Nos activity) by using a proofreading-deficient mutant of B. japonicum USDA110 combined with selection pressure for N2O respiration (Itakura et al., 2008). Inoculation of soybeans with the Nos++ strain 5M09 mitigated N2O emission in both pot and field experiments (Itakura et al., 2013). We have also shown that in strain 5M09, transcription of nosZ, nosR and nosD (genes from the N2O reduction cluster), and napA (encoding the catalytic subunit of the periplasmic nitrate reductase), was higher than in USDA110 (Sánchez et al., 2013). The induced transcription of nos and nap genes did not depend on the FixLJ-FixK2-NnrR and RegSR-NifA regulatory cascades (Sánchez et al., 2013). In this paper, we describe the causal gene for the Nos++ phenotype and the underlying mechanism. Results Genome determination for the B. japonicum Nos++ strains In a previous study, Nos++ strains 5M09, 5M14 and 20M19 were isolated from the same population (population M), and exhibited significantly higher Nos activities than the wild-type USDA110 (Itakura et al., 2008). In the present study, we isolated four new Nos++ strains from two different populations: strains G2, G4 and G9 from population G, and strain K2 from population K. Similar to the refer-

ence strain 5M09, Nos activity values in free-living cells of the strains isolated from populations G and K were about four times that in strain USDA110 (Table 1). To identify the causal gene or genes responsible for the Nos++ phenotype, the genomes of M strains (5M09, 5M14 and 20M19) and G strains (G2, G4 and G9) were sequenced using the Illumina Genome Analyser II and compared. In all cases, the covered genomic bases were more than 99.3% (Supporting Information Table S1). Nucleotide substitutions (SNP, single nucleotide polymorphism) were the major mutations (94–100% of the total), but deletions and insertions were also detected (Supporting Information Table S2). Among the nucleotide substitutions, transversions were most common (Supporting Information Table S2). This result is in line with that reported in other studies of mutator-mediated mutagenesis (Shiwa et al., 2012). When eight target genes that were commonly mutated in strains 5M09, 5M19 and 20M19 were sequenced by the Sanger method, the results confirmed the validity of SNP detection by the Illumina technology (Supporting Information Fig. S1). Strains from population M carried 52–66 mutations (Fig. 1A). 5M09, 5M14 and 20M19 shared 24 mutations (Supporting Information Fig. S2). Strains from population G carried 24–50 mutations (Fig. 1A). G2, G4 and G9 only shared four mutations (Supporting Information Fig. S2). The only mutated gene shared between the strains from populations M and G was bll4572; this gene was mutated in all six strains (Fig. 1A and B; Supporting Information Fig. S2). In M strains, substitution of 281C to A created a premature stop codon (298 of 388 residues are missing), whereas in G strains substitution of 302T to G led to an amino acid change (101leucine to arginine, L101R) (Fig. 1B, Supporting Information Fig. S3A). The Nos++ strain K2 was not included in the Illumina genome analysis, but its bll4572 sequence revealed a single nucleotide deletion (701G) that caused a translational frameshift (the

Table 1. N2O reductase (Nos) activity in Bradyrhizobium japonicum free-living and symbiotic cells (bacteroids). Nos activity (nmol N2O h–1 mg protein–1) Strain

Free-living

Bacteroids

USDA110 5M09 G2 G4 G9 K2 nasS (bll4572) nasT (bll4573) nasST

89 ± 35 397 ± 102b 334 ± 43b 399 ± 74b 384 ± 55b 344 ± 34b 429 ± 127b 147 ± 64a 184 ± 63a

225 ± 39a 316 ± 23b NM NM NM NM 345 ± 6b NM NM

a

Data are means ± standard deviations. Values labelled with the same letter do not differ significantly (Tukey’s honestly significant difference test, P < 0.05, n = 3–12). NM, not measured.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

NasS mutation induces N2O reductase activity

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Fig. 1. Identification of bll4572 (nasS) as the causal gene responsible for the Nos++ phenotype. A. Mutation distribution on the chromosomes of Nos++ strains isolated from population M (5M09, 5M14 and 20M19) and population G (G2, G4 and G9). The numbers in parentheses indicate the number of mutations in each strain; see also Supporting Information Fig. S2. B. Mutations in bll4572, at the gene and protein levels, in the different Nos++ strains.

last 54 residues are not conserved) (Fig. 1B, Supporting Information Fig. S3A). These results suggested that the mutation in bll4572 is involved in the enhancement of Nos activity in the Nos++ mutants.

Mutational analysis of bll4572 To examine whether bll4572 is responsible for the enhancement of Nos activity, a B. japonicum USDA110 derivative mutant was constructed by replacing the wildtype allele with that in Nos++ strain 5M09 (Fig. 1B). Both the 5M09 and bll4572 mutant cells exhibited similar Nos activity, at about 4.8 and 4.5 times, respectively, the value in wild-type cells (Table 1). Anaerobic growth of 5M09 and bll4572 mutant cells with N2O as the sole electron acceptor was significantly higher than that of wild-type cells, reaching an OD660 of 0.54 after 7 days compared with 0.38 for the wild-type (Fig. 2A). These results suggest that the mutation in bll4572 enhances the Nos activity in B. japonicum. To examine the symbiotic phenotype of the bll4572 mutant, we performed a plant inoculation test. Symbiotic cells of the bll4572 mutant and strain 5M09 exhibited similar values of Nos activity, which were significantly higher than the value in symbiotic cells of the wild-type (Table 1). This suggests that bll4572 is involved in the N2O reduction process also under symbiotic conditions. Nitrogen fixation-dependent growth of soybean plants inoculated with the bll4572 mutant was similar to that of the wild-type (Supporting Information Fig. S4A). In addition,

plants inoculated with the wild-type or the bll4572 mutant exhibited similar values of plant dry weight, nodule number and nodule fresh weight (Supporting Information Fig. S4B–D). These results indicate that bll4572 does not adversely affect the development and functioning of nodules or plant growth under the conditions tested.

bll4572 and bll4573 are homologues to nasST bll4572 encodes a 388-amino acids protein that is predicted to be a putative NO3− transport protein (Kaneko et al., 2002). bll4572 product is homologous to NrtA (32% identity), which is the periplasmic component of an ABCtype uptake system for NO3− in Synechocystis sp. PCC 6803 (Koropatkin et al., 2006). bll4572 product conserves seven of the eight NO3−-binding residues of NrtA (Supporting Information Fig. S3B), suggesting that it might be an NO3− binding protein. However, it lacks the N-terminal periplasmic leader sequence and other residues that are present in NrtA and that are required for periplasmic export and membrane tethering (Supporting Information Fig. S3B), suggesting that it may be a soluble cytoplasmic protein. bll4573 gene (adjacent to the bll4572) encodes a 196amino acids protein that is predicted to be a twocomponent response regulator (Kaneko et al., 2002). bll4573 product shares 41–49% sequence identity with NasT, an RNA-binding antiterminator involved in the transcriptional regulation of NO3−/NO2− (nitrite) assimilation genes in Azotobacter vinelandii DJ (Gutiérrez et al., 1995;

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

3266 C. Sánchez et al. Fig. S5): an N-terminal CheY-like receiver domain (REC domain, amino acids 8–122), found in the response regulators of two-component regulatory systems (Shu and Zhulin, 2002); and a C-terminal RNA-binding domain (an ANTAR domain, amino acids 128 to 189) that is conserved in several antiterminators (Morth et al., 2004). These observations suggest that bll4572 and bll4573 encode the NO3–/NO2– responsive two-component system NasS (Bll4572)-NasT (Bll4573), respectively, where NasS is a NO3−/NO2− sensor and NasT is a transcription antiterminator (Romeo et al., 2012; Wang et al., 2012; Luque-Almagro et al., 2013). Bradyrhizobium japonicum USDA110 NasS shares 38–40% sequence identity with the NasS proteins of A. vinelandii, P. aeruginosa and P. denitrificans (Supporting Information Fig. S3B). Typically, nasST genes are clustered together with other genes involved in NO3– assimilation, such as genes that encode NO3– transporters, assimilatory nitrate reductase and assimilatory nitrite reductase (Luque-Almagro et al., 2011; Fig. 3). Bradyrhizobium japonicum USDA110 nasST genes are located in a putative NO2− assimilation gene cluster that contains a ferredoxin nitrite reductase (bll4571) and a flavoprotein α-subunit of a probable sulfite reductase (bll4570) (Kaneko et al., 2002; Fig. 3). Regulation of nos and nap genes by the NasST system in the absence of nitrate Fig. 2. Growth of Bradyrhizobium japonicum USDA110 and the 5M09, nasS (bll4572), nasT (bll4573) and nasST mutant strains on HMM medium at 30°C with reciprocal shaking at 300 r.p.m. under (A) anaerobic conditions with 20% N2O (v/v) as the electron acceptor and (B) anaerobic conditions with 20 mM KNO3 as the electron acceptor. Growth was measured by recording the optical density (660 nm) of the cultures on a daily basis (n = 4).

Wang et al., 2012), Pseudomonas aeruginosa PAO1 (Romeo et al., 2012) and Paracoccus denitrificans PD1222 (Luque-Almagro et al., 2013) (Supporting Information Fig. S5). bll4573 product contains two domains that are also present in other NasT proteins (Gutiérrez et al., 1995; Romeo et al., 2012; Wang et al., 2012; Luque-Almagro et al., 2013; Supporting Information

In A. vinelandii and P. denitrificans, NasS and NasT proteins act as repressor and activator, respectively, of the expression of genes for the NO3−/NO2− assimilatory pathway (generally, nas genes) (Gutiérrez et al., 1995; Luque-Almagro et al., 2013). Recent work in P. denitrificans has demonstrated that NasS and NasT form a stable complex that is sensitive to the presence of NO3−/NO2− (Luque-Almagro et al., 2013). Binding of NO3−/ NO2− by the sensor NasS triggers the release of NasT, which binds the leader sequence upstream of the nas operon (Luque-Almagro et al., 2013), thereby preventing hairpin formation and allowing complete transcription of the nas operon (Romeo et al., 2012; Wang et al., 2012). This agrees with the observation that in the nasS mutant of B. japonicum, which carries a truncated NasS protein

Fig. 3. Gene alignments of nasS and nasT and genes that encode nitrate or nitrite assimilation in Bradyrhizobium japonicum USDA110, Paracoccus denitrificans PD1222, Azotobacter vinelandii DJ and Pseudomonas aeruginosa PAO1. Open reading frames that encode the following putative products are coloured as follows: nitrate sensor (nasS) and antiterminator (nasT), black; NO3−/ NO2− transporters, diagonal grey lines; nitrate reductase, light grey; nitrite reductase, dark grey.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

NasS mutation induces N2O reductase activity (Fig. 1B, Supporting Information Fig. S3A), Nos activity was induced (Table 1). Thus, we were interested in testing whether transcription of nosZ depends on nasS and nasT. β-Galactosidase activity by a PnosZ-lacZ transcriptional fusion was examined in nasS−, nasT− and nasS−T− genetic backgrounds. Since the direction of transcription in B. japonicum USDA110 is from nasT to nasS (Fig. 3), we constructed nasT and nasST in frame deletion mutants in order to avoid a polar effect on nasS. Before the β-galactosidase assay, cells were incubated under aerobic or anaerobic conditions for 24 h in the absence of NO3−. Under aerobic and anaerobic conditions, the β-galactosidase activity in a PnosZ-lacZ transcriptional fusion was significantly higher in the nasS and 5M09 strains than in the wild-type (Table 2). This confirms that nasS is a repressor of the transcription of nosZ in B. japonicum. The nasST double mutant exhibited aerobic and anaerobic values of β-galactosidase activity not significantly different from those in the wild type (Table 2). This result suggests that the enhancement of nosZ transcription induced by the nasS mutation depends on the nasT gene. Indeed, Nos activity in free-living cells of the nasST double mutant decreased (relative to that in the nasS mutant) to around the wild-type value (Table 1). Furthermore, anaerobic growth of the nasST double mutant with N2O as the electron acceptor was much lower than that of the nasS mutant, reaching an OD660 of 0.19, compared with 0.54 for the nasS strain after 7 days growth (Fig. 2A). To examine whether the transcription of nap genes also depends on NasST, we analysed the β-galactosidase activity by a PnapE-lacZ transcriptional fusion in nasS−, nasT− and nasS−T− genetic backgrounds. The β-galactosidase activity in a PnapE-lacZ fusion was

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higher in the nasS and 5M09 strains than in USDA110 under aerobic and anaerobic conditions (Table 2). The induction of napE expression depended on the nasT product since β-galactosidase activity in the nasST mutant was not significantly different from that in USDA110 (Table 2). The β-galactosidase activities by the PnirK-lacZ and PnorC-lacZ fusions in the nasS and nasST mutants did not differ significantly from that in USDA110 (Table 2), in line with the absence of nirK and nor induction in 5M09 reported earlier (Sánchez et al., 2013). This suggests that the regulation of transcription of denitrification genes by NasST is specific to the nos and nap genes, and occurs under aerobic and anaerobic conditions. Regulation of nos and nap genes by NasST system in the presence of nitrate To test whether B. japonicum NasST forms a complex, which is sensitive to the presence of NO3−, His6-tag-fused NasS and GST-fused NasT were independently produced in E. coli and purified (Fig. 4A). To establish the composition of the NasS-NasT complex of B. japonicum, a mixture of purified NasS and NasT was subjected to sizeexclusion chromatography in an analysis similar to that for P. denitrificans (Luque-Almagro et al., 2013). The fractions corresponding to elution volumes of 67–79 ml were collected (fraction no. 1–6), and subjected to SDS-PAGE analysis (Fig. 4B). In the absence of NO3−, NasS and NasT were eluted at 68 ml as a single peak. SDS-PAGE analysis revealed equivalent amounts of NasS and NasT in the first three fractions (1–3), indicating that NasS and NasT form a complex in the absence of NO3−. In the presence of 1 mM NO3−, the absorbance peak was shifted to 78 ml. SDS-PAGE analysis of the eluted protein

Table 2. β-Galactosidase activity mediated by nos, nor, nirK and nap promoters in B. japonicum cells incubated under aerobic (+O2) or anaerobic (−O2) conditions in the absence (–KNO3) or in the presence (+KNO3) of 10 mM KNO3. β-Galactosidase activity (Miller units) PnosZ-lacZ

–KNO3

+KNO3

Strain

+O2

USDA110 5M09 nasS nasT nasST USDA110 5M09 nasS nasT nasST

23 ± 4 66 ± 8b 86 ± 19b 27 ± 10a 27 ± 9a 79 ± 7a 86 ± 7a 98 ± 5b 31 ± 3c 32 ± 2c

PnorC-lacZ +O2

–O2 a

55 ± 10 143 ± 26b 161 ± 17b 49 ± 18a 51 ± 19a 559 ± 65a 543 ± 61a 691 ± 166a 185 ± 25b 188 ± 36b a

PnirK-lacZ +O2

–O2

10 ± 3 10 ± 4ac 9 ± 3a 19 ± 3b 17 ± 2bc 10 ± 1a 10 ± 1a 9 ± 1a 12 ± 2a 10 ± 1a a

47 ± 16 43 ± 8a 40 ± 8a 61 ± 20a 63 ± 24a 659 ± 116a 636 ± 178a 691 ± 183a 546 ± 182a 611 ± 180a a

PnapE-lacZ +O2

–O2

31 ± 4 31 ± 8a 31 ± 9a 39 ± 1a 41 ± 2a 24 ± 3a 21 ± 5a 21 ± 7a 29 ± 1a 31 ± 5a a

140 ± 42 131 ± 36a 142 ± 28a 171 ± 17a 170 ± 23a 361 ± 46a 359 ± 75a 423 ± 112b 481 ± 38b 554 ± 70b a

–O2

41 ± 14 88 ± 20b 101 ± 24b 31 ± 11a 32 ± 10a 76 ± 7a 86 ± 2b 83 ± 2ab 34 ± 1c 38 ± 3c a

124 ± 4a 252 ± 48b 230 ± 8b 77 ± 21a 77 ± 17a 516 ± 125a 419 ± 151a 590 ± 95a 172 ± 39b 150 ± 40b

Data are means ± standard deviations. Values labelled with the same letter do not differ significantly (Tukey’s honestly significant difference test, P < 0.05, n = 4). Shading indicates differential regulation compared with the wild-type USDA110.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

3268 C. Sánchez et al. Fig. 4. Characteristics of recombinant NasS and NasT proteins. A. His6-tag-fused NasS (42 kDa) and GST-fused NasT (49 kDa) were independently expressed in E. coli and purified using affinity column chromatography, and protein purity was assessed by SDS-PAGE using Coomassie Brilliant Blue staining. B. NO3−-dependent dissociation of purified NasS-NasT was analysed by means of size-exclusion chromatography. Representative chromatograms for protein elution recorded in the absence (solid line) and presence (dotted line) of 1 mM NO3− are shown. SDS-PAGE analysis of the protein content is shown below the chromatogram. Protein bands were visualized by means of Coomassie Brilliant Blue staining. mAU, milli-absorbance units.

revealed that peak of NasS elution shifted to the last three fractions (4–6), indicating that NasS is a monomer in the presence of NO3−. These results were similar to those for NasS-NasT from P. denitrificans, which formed a complex in the absence of NO3− and dissociated in the presence of NO3− (Luque-Almagro et al., 2013). Therefore, we conclude that NasS-NasT of B. japonicum is also a regulatory complex that dissociates upon sensing NO3−. We also examined the β-galactosidase activity in the PnosZ-lacZ and PnapE-lacZ transcriptional fusions in the presence of 10 mM KNO3. We observed no differences between the wild-type USDA110 and the nasS mutant under aerobic and anaerobic conditions (Table 2). This suggests that NO3− is counteracting the NasS-mediated repression of nos and nap transcription in the wild-type (Fig. 5). In addition, the nasT mutant clearly exhibited lower aerobic and anaerobic βgalactosidase activities than the wild-type (Table 2), suggesting that NasT is required for the induction of nos and nap by NO3−. Unlike nos and nap, mutation of nasT had no effect on nirK and nor transcription in the presence of NO3− (Table 2). To test whether transcriptional activity of napE correlates to Nap activity, we analysed methyl viologen (MV)dependent nitrate reductase activity in cells incubated under anaerobic conditions in the presence of 10 mM KNO3. Wild-type and nasS mutant cells exhibited similar values of Nap activity (700 ± 119 and 769 ± 189 nmol NO2− h–1 mg protein–1 respectively). However, Nap activity in nasT and nasST mutant cells decreased drastically (to 11.7 ± 7 and 12 ± 4 nmol NO2− h–1 mg protein–1 respectively). Since MV does not easily traverse the cytoplasmic membrane (Jones et al., 1976), it is likely that nitrate reductase activity resulted exclusively from Nap. Thus, these results suggest that the NasST system is a key

player for regulation of the periplasmic NO3− reduction in B. japonicum. However, anaerobic growth of nasT and nasST mutant cells with 20 mM KNO3 did not differ significantly from that in the wild-type and nasS mutant strain (Fig. 2).

Fig. 5. Model for regulation of nap and nos genes by NasST system in B. japonicum USDA110 and nasS and Nos++ mutant strains. In the absence of nitrate [NO3− (−)], NasS and NasT form a stable complex, which is dissociated only in the case of nasS mutation (mutated NasS is indicated by a circle with dotted line). In the presence of nitrate [NO3− (+)], binding of NO3− by the sensor NasS triggers release of the antiterminator NasT. Regulation of nap, nir, nor and nos genes by FixLJ-FixK2-NnrR and RegSR-NifA cascades is also shown (adapted from Velasco et al., 2004; Bueno et al., 2012). Dotted arrows indicate that the regulatory mechanism is not fully understood. Therefore, the regulation of NasT target genes may be direct or indirect.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

NasS mutation induces N2O reductase activity Discussion The concentration of N2O has risen during the past century and a half, mainly due to agriculture and the increased use of industrial fertilizers (Hénault et al., 2012; Thomson et al., 2012). Since the 1997 Kyoto Protocol, many of the non-biological emissions of N2O have been systematically lowered, but N2O emissions from agriculture are essentially unchanged (Thomson et al., 2012). Temporal and spatial hot spots of N2O fluxes represent a large part of these emissions. This is the case for soybean fields in the late growth period, during degradation of the root nodules (Inaba et al., 2009; 2012; Hénault et al., 2012). While N2O generation is due to a diversity of microbial enzymes, N2O consumption appears to be due exclusively to Nos (Baggs, 2011; Thomson et al., 2012). Comparative genome analysis identified a gene involved in the Nos++ phenotype of strains isolated by using a proofreading-deficient mutant of B. japonicum USDA110 combined with selection pressure for N2O respiration (Itakura et al., 2008). When we compared the genomes of the Nos++ strains from populations M and G, we found that they shared 24 and 4 mutations respectively (Supporting Information Fig. S2). However, only one candidate gene, nasS, was shared between the two populations. Since a B. japonicum USDA110 derivative mutant strain carrying the nasS allele from reference strain 5M09 exhibited the Nos++ phenotype, a mutation in the nasS gene appears to be the mechanism responsible for the enhancement of Nos activity in Nos++ strains. In addition, nasS bacteroids exhibited the Nos++ phenotype and normal N2-fixing phenotype, suggesting that NasS also regulates the transcription of nos genes under symbiotic conditions but is not involved in N2-fixation. Three different mutations in nasS led to the induction of Nos activity in the Nos++ strains in the absence of NO3−. In M strains, the production of a truncated NasS, likely unable to interact with NasT (Fig. 5, NO3− [−]), may allow the transcription of nosZ by NasT antitermination, and thus the enhancement of Nos activity. In strain K2, the production of a NasS protein, in which the last 54 residues are not conserved, probably led to an inability to interact with NasT (Fig. 5, NO3− [−]). In the case of strains from population G, the replacement of 101leucine by arginine in NasS led to protein misfolding and aggregation (data not shown). Thus, it is reasonable to hypothesize that in G strains, NasS-L101R is not correctly folded and that NasT exists in the free form that can activate nosZ expression (Fig. 5, NO3− [−]). The NasST system is typically involved in the NO3−responsive regulation of genes in the NO3−/NO2− assimilatory pathway (generally, nas genes), and it appears to be widely distributed in Gram-negative bacteria, including

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symbionts, pathogens and denitrifiers, which can assimilate NO3−/NO2− in the absence of other N sources (Luque-Almagro et al., 2011). nasST genes typically cluster within or in close proximity to nas loci (Luque-Almagro et al., 2011). In B. japonicum, the genes that encode the putative assimilatory nitrate and nitrite reductases are located at distinct loci (Kaneko et al., 2002; Sánchez et al., 2011). nasST cluster together with bll4571, which encodes the putative assimilatory nitrite reductase (Kaneko et al., 2002). This supports a role of nasST in the regulation of the NO3−/NO2− assimilation genes of B. japonicum. In B. japonicum, nas genes have not been characterized, but this bacterium is known to be able to assimilate and denitrify 15NO3− simultaneously to 15 NH4+ and 15N2 under anaerobiosis (Vairinhos et al., 1989), suggesting that NO3− assimilation can occur when required. nasST genes can also be located at different loci from the NO3−/NO2− assimilation gene cluster. Thus, a possible role as regulators of other metabolic pathways has also been suggested (Luque-Almagro et al., 2011). In B. japonicum, maximal induction of denitrification genes (nap, nirK, nor and nos) requires oxygen limitation and the presence of NO3− or a derived N oxide (Velasco et al., 2004; Bueno et al., 2012). The response to low oxygen conditions is mediated by the FixLJ-FixK2 regulatory cascade, and the N oxide-mediated induction of nor genes has been suggested to depend on NnrR (Velasco et al., 2004; Bueno et al., 2012; Fig. 5). Here, we show that, in the presence of NO3−, the NasS-NasT complex of B. japonicum was dissociated (Fig. 4), and the nasT mutant had 70% lower transcription of nos and nap genes than the wild-type under both aerobic and anaerobic conditions (Table 2). These observations suggest that NasST is involved in NO3−-mediated induction of nos and nap (Fig. 5, NO3− [+]). Nos and Nap enzymes are typically involved in denitrification, the dissimilatory NO3− reduction to N2 concomitant to energy conservation. In B. japonicum USDA110, complete denitrification of NO3− to N2 requires the Nap, NirK, Nor and Nos enzymes. However, nirK and nor transcription was not affected by the nasST mutation. This suggests an alternative physiological role, independent of denitrification, for the Nap and Nos enzymes in response to NasST regulation. It has been reported that the reduction of NO3− and N2O by Nap and Nos is not coupled obligatorily to denitrification (Zumft, 1997, and references therein). Our results indicate that NO3− might counteract the NasS-mediated inhibition of nap and nos genes by allowing the dissociation of the antiterminator NasT from the NasS-NasT complex (Fig. 5, NO3− [+]). This is the same mechanism described for regulation of nas genes by NasST in A. vinelandii and P. denitrificans (Gutiérrez et al., 1995; Wang et al., 2012; Luque-Almagro et al.,

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

3270 C. Sánchez et al. 2013). Several studies have provided evidence that NasT binds to the leader sequence in mRNA upstream of the nas operon, thereby preventing hairpin formation and allowing complete transcription of the nas genes (Romeo et al., 2012; Wang et al., 2012; Luque-Almagro et al., 2013). We carried out RNA secondary structure analysis using Mfold (Zuker, 2003) and the WebGeSTer Database (Mitra et al., 2011), but we could not find similar hairpin structures associated with transcription antitermination upstream of nosR or nosZ and napE, suggesting that the regulation of these genes by NasST might be indirect, through an intermediate regulator. Nap has been proposed as the enzyme responsible for anaerobic growth of B. japonicum under NO3−-respiring conditions by reducing NO3− to NO2−, which can be further reduced by the denitrification reactions to N2 (Delgado et al., 2003). Yet we found that even when Nap activity decreased by 98% in nasT mutant cells incubated anaerobically with NO3−, the growth under the same conditions did not differ significantly from that in the wild-type (Fig. 2B). This suggests that anaerobic growth under NO3−-respiring conditions is being supported by another nitrate reductase, whose activity is localized in the cytoplasm. One possibility might be the membrane-bound respiratory nitrate reductase (Nar), although there is evidence that B. japonicum USDA110 lacks the Nar system (Delgado et al., 2003). A second possibility might be the assimilatory nitrate reductase enzyme. This enzyme may be present in B. japonicum USDA110, with blr2809 putatively encoding the large catalytic subunit, NasA (Sánchez et al., 2011). However, if NasST is regulating assimilatory nitrate reductase in B. japonicum, as has been described for several bacteria (Luque-Almagro et al., 2011), this activity would also be absent in a nasT mutant. This remains to be established. Our use of proofreading-deficient mutants to obtain B. japonicum Nos++ strains followed by comparative genome analysis identified a new and unexpected system for the regulation of periplasmic NO3− reduction and N2O reduction. Although the biological significance of regulation of Nap and Nos by NasST remains to be established, our findings raise many questions, whose answers will improve the understanding of N transformation in the environment, including the mechanisms by which N2O is generated and reduced. An interesting issue is whether NasST system regulates nos genes in other denitrifying bacteria, such P. denitrificans, various pseudomonads, the alfalfa endosymbiont Sinorhizobium meliloti (Galibert et al., 2001; Torres et al., 2011) and the photosynthetic rhizobia Bradyrhizobium BTAi1 that nodulates Aeschynomene plants (Giraud et al., 2007). Emission of N2O from soybean rhizosphere was significantly decreased by inoculation with B. japonicum Nos++ strains (Itakura et al., 2013), suggesting that NO3− concentration

in the rhizosphere was not high enough to induce NasSNasT dissociation in B. japonicum wild type (Fig. 5, NO3− [−]). This shows that the enhancement of N2O reductase activity by rhizobial nasS mutation is a promising strategy to decrease the emission of this greenhouse gas from legume fields into the atmosphere. Experimental procedures Bacterial strains and growth conditions Bacterial strains and plasmids are listed in Supporting Information Table S3. Bradyrhizobium japonicum cells were cultured at 30°C in HM salt medium (Cole and Elkan, 1973) supplemented with 0.1% arabinose and 0.025% (w/v) yeast extract (Difco). HM medium was further supplemented by adding trace metals (HMM medium) for the denitrification assays (Sameshima-Saito et al., 2006). Escherichia coli cells were grown at 37°C in Luria-Bertani medium (Miller, 1972). Antibiotics were added to the media at the following concentrations: for B. japonicum, 100 μg tetracycline (Tc) ml−1, 100 μg kanamycin (Km) ml−1 and 100 μg polymyxin B ml−1; for E. coli, 50 μg Tc ml−1, 50 μg Km ml−1 and 50 μg ampicillin ml−1.

Plant inoculation Surface-sterilized soybean seeds (Glycine max ‘Enrei’) were germinated in sterile vermiculite for 2 days at 25°C and then transplanted to a Leonard jar pot (Leonard, 1943), which contained sterile vermiculite and nitrogen-free nutrient solution (Okazaki et al., 2004). The seeds were then inoculated with Bradyrhizobium japonicum at 1 × 107 cells per seed. Plants were then grown in a phytotron (Koito Industries) for 37 days at 25°C/20°C with a 16 h light/8 h dark photoperiod (270 μmol s−1 m−2). Shoots (separated from roots at the cotyledonary node) were dried to a constant weight at 80°C and dry weight was measured. Nodule number and nodule fresh weight were determined per plant.

Isolation of B. japonicum populations with higher N2O activity The B. japonicum Nos++ strains were isolated by introducing a plasmid containing a mutated dnaQ gene (pKQ2) and then performing enrichment culture under selection pressure for N2O respiration, as previously described (Itakura et al., 2008). Introduction of the mutator plasmid pKQ2 into B. japonicum USDA110 cells was carried out using triparental mating, as described previously (Itakura et al., 2008). Three independent populations (M, G and K) were generated from three independent triparental matings. Selection for N2O respiration, plant inoculation, single-colony isolation via plant passage and polymerase chain reaction (PCR) verification of plasmid elimination were also performed, as described previously (Itakura et al., 2008).

Genome sequence analysis Determination of genome sequences was performed using an Illumina Genome Analyser II (Illumina). Briefly, 3 μg of

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

NasS mutation induces N2O reductase activity genomic DNA was fragmented to an average size of 500 bp on a Covaris S2 system (Covaris). Sequencing libraries were generated from the fragments using a Multiplexing Sample Preparation Oligonucleotide Kit (Illumina) and sequenced as paired-end 100 bp reads according to the manufacturer’s protocols. The read data were deposited in the DDBJ Sequence Read Archive under accession number DRA001174 (http://www.ddbj.nig.ac.jp/). Sequencing reads were mapped to the genome of B. japonicum USDA110 (accession no. NC_ 004463) using the BWA software (Ver. 0.5.1) with default parameter values (Li and Durbin, 2009). To call mutations, we used the SAMTOOLs software (Ver. 0.1.9) (Li et al., 2009) and applied the following additional criteria. There must be at least five aligned reads, and the proportion of reads that called a variant must be more than 90%.

Sanger dideoxy sequencing Sanger DNA sequencing was carried out on an ABI 3130xl Genetic Analyzer (Applied Biosystems) using BigDye terminator chemistry (Ver. 3.1) (Applied Biosystems) according to the manufacturer’s instructions. The PCR products were run in a sequencing reaction with the following thermocycling conditions: 25 cycles of 96°C for 10 s, 50°C for 5 s and 60°C for 4 min.

Plasmid and strain construction The nasS mutant of B. japonicum USDA110 was constructed by replacing wild-type bll4572 with the mutant allele from 5M09. First, a 1777 bp HindIII-PstI fragment containing the bll4572 region from B. japonicum 5M09 was cloned into the pUC19 vector (New England Biolabs), yielding the plasmid pMS0101. A 1785 bp HindIII-XbaI fragment from pMS0101 was then cloned into the Kmr-suicide vector pK19mobsacB (Schäfer et al., 1994) to yield the plasmid pMS0102. To construct nasT in frame deletion mutants (nasT and nasST) of B. japonicum USDA110, DNA fragments corresponding to the upstream and downstream regions of bll4573 were individually cloned as 604 bp HindIII-BamHI and 629 bp BamHI-XbaI fragments into the pBluescriptSK(+) vector (Stratagene), yielding the plasmids pMS0301 and pMS0302 respectively. The 604 bp HindIII-BamHI fragment from pMS0301 was then cloned into pMS0302, yielding the plasmid pMS0303. Finally, a 1.2 kb HindIII-XbaI fragment (containing an in frame deletion of 281 bp in bll4573) from pMS0303 was cloned into the Kmr-suicide vector pK18mobsacB (Schäfer et al., 1994) to yield the plasmid pMS0304. Plasmid pMS0102 or pMS0304 was transferred by conjugation from E. coli DH5α to B. japonicum USDA110 for markerless mutant construction. Triparental matings were conducted using pRK2013 as a helper plasmid. Kmr transconjugants were selected and grown in the presence of 10% sucrose to force loss of the vector-encoded sacB gene. The resulting colonies were checked for Km sensitivity. The desired mutations in nasS and nasT were confirmed by sequence analysis and PCR respectively. To obtain the nasST double mutant, plasmid pMS0304 was transferred by conjugation from E. coli DH5α to B. japonicum nasS for markerless mutant construction, as described above. The desired mutation in nasT was confirmed by PCR.

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For the β-galactosidase assays, chromosomally integrated transcriptional lacZ fusions with the napE, nirK, norC and nosZ promoters were used. The plasmids pBG0614 (Robles et al., 2006), pRJ2498 (Mesa et al., 2003), pRJ2499 (Mesa et al., 2003) and pNOSLZch (Sánchez et al., 2013) were integrated in B. japonicum USDA110, and the nasS, nasT and nasST mutant strains (see Supporting Information Table S3). Acquisition of Tcr indicated that the suicide plasmid containing the fusion had been integrated into the chromosome after a single recombination event, which generated a tandem duplication. Integration of the fusion plasmid into the chromosome was tested by PCR.

Analytical methods For the MV-dependent nitrate reductase assay, B. japonicum cells were grown aerobically in 15 ml of HM medium, collected by centrifugation (8000× g, 10 min at 4°C), and washed twice. They were then adjusted to an optical density at 660 nm of about 0.2 by the addition of HMM medium supplemented with 10 mM KNO3. Twenty millilitre of the cell preparations were incubated in 80 ml tubes. The tubes were sealed and the gas phase was replaced with N2 for the anaerobic incubation for 24 h at 30°C with reciprocal shaking at 300 r.p.m. MV-dependent nitrate reductase activity was measured as described previously (Sánchez et al., 2010). For the N2O reductase assay, B. japonicum cells were grown aerobically in 15 ml of HM medium, collected by centrifugation (8000× g, 10 min at 4°C), washed twice and finally re-suspended in 5 ml of HMM medium. Bacteroids were prepared as described previously (Itakura et al., 2013). The N2O reductase assay was performed as described previously (Itakura et al., 2013). Protein concentrations were estimated using the Bio-Rad assay (Bio-Rad) with a standard curve of varying bovine serum albumin concentrations. For the β-galactosidase assay, aerobically grown cells were collected and washed twice. They were then adjusted to an optical density (660 nm) of about 0.2 by the addition of HMM medium. Five millilitre of the cell preparations were incubated in 35 ml tubes aerobically or anaerobically in the absence or in the presence of 10 mM KNO3 for 24 h at 30°C with reciprocal shaking at 300 r.p.m. For the anaerobic incubation, the tubes were sealed and the gas phase was replaced with N2. Final optical density (660 nm) was about 0.5 for aerobic incubations (in the absence and in the presence of 10 mM KNO3), and 0.2 and 0.25 for anaerobic incubations in the absence and in the presence of 10 mM KNO3 respectively. β-Galactosidase activity was determined with permeabilized cells as described previously (Miller, 1972).

Cloning, expression and purification of NasS and NasT Genes encoding NasS (bll4572) and NasT (bll4573) were amplified by PCR, using KOD plus neo DNA polymerase (Toyobo) according to manufacturer’s instructions, from a pUC-based clone library (brp12240) using the primer pairs 5′-ATGACCGGACCGCTTC-3′ and 5′-GGCCTTCCAGCGA CC-3′ for bll4572, and 5′-GTTCTGTTCCAGGGGCCCAT GAGCGCCGAGCAGTCG-3′ and 5′-GATGCGGCCGCTC GAGTCATTTCAGCATCTCCGACGC-3′ for bll4573. The

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

3272 C. Sánchez et al. amplified genes were inserted into expression vectors amplified by PCR using the In-Fusion HD Cloning Kit (Takara Bio). pET28a was amplified by the primer pair 5′-GGTCGCT GGAAGGCCCACCACCACCACCACCACTGAGATCCGGC TGCTAAC-3′ and 5′-GAAGCGGTCCGGTCATGGTATATCT CCTTC-3′, and pGEX-4T was amplified by the primer pair 5′-TGACTCGAGCGGCCGCATCGTG-3′ and 5′-GGGCCC CTGGAACAGAACTTCC-3′. nasS was inserted into pET28a (Novagen) to add a His6-tag at the C-terminus of the recombinant protein, and nasT was cloned into pGEX-4T (GE Healthcare) to add a glutathione-S-transferase at the N-terminus. The resultant plasmids for the expression of NasS and NasT (pET28a-NasS and pGEX-NasT) were transformed into E. coli BL21 CodonPlus (Stratagene) and E. coli BL21 (Novagen) respectively. The transformants were grown at 37°C in 200 ml of 2 × YT medium until an absorbance of 0.6 at 600 nm. Gene expression was induced by 0.1 mM isopropyl β-D-thiogalactopyranoside and continued at 15°C for 20 h. The cells were harvested by centrifugation at 8000× g for 5 min at 4°C, then were suspended in buffer A (50 mM Tris-HCl, pH 8.0, 300 mM NaCl). NasS was purified by immobilized metal affinity column chromatography. Soluble cell extract was loaded onto a Ni-NTA agarose column (Qiagen). The column was then washed with buffer A containing 10 mM imidazole to elute the unbound protein. Bound protein was eluted with a buffer A containing 500 mM imidazole. NasT was purified using a Glutathione HiCap column (Qiagen) according to the manufacturer’s instructions. Fractions containing purified NasS or NasT were desalted using Bio-Gel P-6 Desalting Cartridge (Bio-Rad), followed by buffer exchange with 10 mM HEPES (pH 8.0).

Site-directed mutagenesis of nasS Site-directed mutagenesis of NasS-L101R was carried out using the QuikChange kit (Agilent). The pET28a plasmid carrying the nasS gene was used as the template for PCR using the primer pair 5′-GACGCTTCATGCCGCGCGAA TGGAGGAGATCGAC-3′ and 5′-GTCGATCTCCTCCATT CGCGCGGCATGAAGCGTC-3′ to produce the plasmid pNasS-L101R.

Size-exclusion chromatography Analytical size-exclusion chromatography was performed on a 120 ml HiLoad 16/600 Superdex 200 pg column (GE Healthcare) equilibrated with 10 mM HEPES (pH 8.0) containing 150 mM NaCl. The column was loaded with 1 ml of purified protein (8 nmol each of NasS and NasT). The absorbance at 280 nm was monitored.

Bioinformatics Amino acid sequences were obtained from the UniProt database (http://www.uniprot.org/) using the following accession numbers: B. japonicum USDA110 NasS, Q89LH2; Paracoccus denitrificans PD1222 NasS, A1BAH4; Azotobacter vinelandii DJ NasS, Q44530; Pseudomonas aeruginosa PAO1 NasS, Q9I2V6; Synechocystis sp. PCC6803 NrtA, P73452; B. japonicum USDA110 NasT,

Q89LH1; P. denitrificans PD1222 NasT, A1BAH5; A. vinelandii DJ NasT, Q44531; and P. aeruginosa PAO1 NasT, Q9I2V7. Sequences were aligned using the ClustalΩ algorithm (Sievers et al., 2011) at http://www.uniprot.org/align/. Domain searching was performed using ScanProsite (de Castro et al., 2006) at http://prosite.expasy.org/scanprosite/.

Acknowledgements We thank S. Mesa and M.J. Delgado (EEZ-CSIC, Granada, Spain) for kindly providing plasmids pBG0614, pRJ2498 and pRJ2499. We thank H. Mitsui (Graduate School of Life Sciences, Tohoku University, Sendai, Japan), M.J. Delgado and R. Kugimiya (Neo-Morgan Laboratory, Tokyo, Japan) for useful discussions regarding the preparation of this manuscript. This work was supported financially by BRAIN grant from the Ministry of Agriculture, Forestry, and Fisheries of Japan, and by Grant-in-Aid for Scientific Research (A) 23248052 and 26252065 from the Ministry of Education, Culture, Sports, Science, and Technology of Japan to K.M.; C.S. was supported by a Japan Society for the Promotion of Science fellowship.

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Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site:

Fig. S1. Sanger sequencing analysis of eight genes commonly mutated in Bradyrhizobium japonicum USDA110 and strains 5M09, 5M19 and 20M19. Mutated nucleotides are highlighted in red. Fig. S2. Summary of mutations detected by genome comparison analysis of the Bradyrhizobium japonicum Nos++ strains. Fig. S3. Alignments of amino acid sequences encoded by bll4572 (nasS). A. Amino acid sequences encoded by bll4572 in Bradyrhizobium japonicum USDA110 and Nos++ strains 5M09, G2 and K2. Non-conserved amino acids are highlighted in red. B. Amino acid sequences of NrtA from Synechocystis sp. PCC6803 (S_NrtA), NasS from B. japonicum USDA110 (Bj_Nas), Paracoccus denitrificans PD1222 (Pd_NasS), Azotobacter vinelandii DJ (Av_Nas) and Pseudomonas aeruginosa PAO1 (Pa_NasS). Nitrate-binding residues in NrtA and NasS are highlighted in black. In NrtA, the TAT signal peptide is highlighted in grey. In (A) and (B), identical and similar residues are indicated by asterisks (*) and colons (:) respectively. Fig. S4. Symbiotic phenotype of Bradyrhizobium japonicum USDA110, and of the 5M09, G2 and nasS (bll4572) mutant strains 37 days after inoculation: (A) nitrogen fixationdependent growth of soybean plants; (B) plant dry weight; (C) nodule number; (D) nodule fresh weight. Non, non-inoculated control. Values labelled with the same letter do not differ significantly (Tukey’s honestly significant difference test, P < 0.05, n = 6). Fig. S5. Sequence alignments of NasT from Bradyrhizobium japonicum USDA110 (Bj_NasT), Paracoccus denitrificans PD1222 (Pd_NasT), Azotobacter vinelandii DJ (Av_NasT) and Pseudomonas aeruginosa PAO1 (Pa_NasT). The response regulatory domain (REC) is highlighted in black, and the RNA-binding domain (ANTAR) is highlighted in grey. Identical and similar residues are indicated by asterisks (*) and colons (:) respectively. Table S1. Results of the sequencing analysis of the Nos++ strains using the Illumina Genome Analyser II. Table S2. Mutational events in the Nos++ strains. Table S3. Bacterial strains and plasmids used in this study.

© 2014 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 16, 3263–3274

The nitrate-sensing NasST system regulates nitrous oxide reductase and periplasmic nitrate reductase in Bradyrhizobium japonicum.

The soybean endosymbiont Bradyrhizobium japonicum is able to scavenge the greenhouse gas N2O through the N2O reductase (Nos). In previous research, N2...
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