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Extracto de carne---- 0.5 g/L
Peptona de carne---- 10 g/L
Nitrato de potasio---- 1 g/L
Cloruro de sodio ----- 5 g/L
pH final -------------- 7.0 ± 0.2 a 25°C

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Extracto de carne: Aportan vitaminas, minerales, lípidos y proteínas

Peptona de carne: Fuente de aminoácidos

Cloruro de sodio: Mantiene la presión osmótica en un ambiente isotónico

Nitrato de potasio: Sustrato a estudiar (Aceptor final de electrones)

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Siembra por agitación

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37±2ºC durante 24 horas1
35±2°C por18-24 horas²

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Respiración anaerobia (bacterias anaerobias estrictas y facultativas) 

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El aceptor final de electrones en la cadena de trasporte de electrones es el ion NO₃⁻

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How to Inoculate & Interpret a Nitrate Broth Tube - MCCC Microbiology

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Despues del periodo de incubación se realiza la lectura de los resultados agregando los reactivos A, B y C

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Reactivo A: Solución de Acido sulfanílico al 0.8% en acido acético al 5N
Reactivo B: Solución de α-Naftilamina o N,N-dimetil-α-naftilamina al 0.6% en ácido acético 5N

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Dependiendo del tipo de nitrato reductasas que el organismo exprese de su material genético, será la capacidad del macroorganismo de reducir el nitrato

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CNRS, UMR 8261, Université de Paris, Institut de Biologie Physico-Chimique, Paris, France

In oxygen (O ₂) limiting environments, numerous aerobic bacteria have the ability to shift from aerobic to anaerobic respiration to release energy. This process requires alternative electron acceptor to replace O ₂ such as nitrate (NO ₃⁻), which has the next best reduction potential after O ₂. Depending on the organism, nitrate respiration involves different enzymes to convert NO ₃⁻ to ammonium (NH ₄⁺) or dinitrogen (N ₂). The expression of these enzymes is tightly controlled by transcription factors (TFs). More recently, bacterial small regulatory RNAs (sRNAs), which are important regulators of the rapid adaptation of microorganisms to extremely diverse environments, have also been shown to control the expression of genes encoding enzymes or TFs related to nitrate respiration. In turn, these TFs control the synthesis of multiple sRNAs. These results suggest that sRNAs play a central role in the control of these metabolic pathways. Here we review the complex interplay between the transcriptional and the post-transcriptional regulators to efficiently control the respiration on nitrate.

Introduction

Nitrate is an important nutrient for microorganisms and can be used in both assimilatory and dissimilatory pathways. Assimilatory pathways allow the incorporation of nitrogen into the organism's biomass (DNA, proteins.). In contrast, dissimilatory nitrate reduction to ammonium (DNRA) is more a catabolic pathway leading to nitrogen excretion. In some cases, the dissimilatory process can release energy and is therefore termed "nitrate respiration." Several metabolic pathways, such as DNRA, denitrification, and in, some bacteria, anaerobic ammonium oxidation (anammox) (reviewed in Kartal et al., 2012), use the reduction of nitrate or nitrite as the first step in respiration. The final products are NH ₄⁺ for DNRA, and N ₂ for the denitrification and anaerobic anammox pathways ( Figure 1); however, numerous bacteria are only partial denitrifiers since the last enzyme required to convert N ₂O to N ₂ is absent ( Philippot et al., 2011). These diverse respiratory nitrate reduction pathways are not necessarily present simultaneously in a given organism, but can compete for nitrate utilization. Denitrification is probably the best studied nitrate pathway in bacteria and it has been shown that incomplete denitrification can release N ₂O in the atmosphere, a greenhouse gas involved in global warming. Nitrate reduction also plays a key role in bacterial pathogenesis ( Vázquez-Torres and Bäumler, 2016) as well as in the gut colonization by enterobacteria.

Figure 1. Biological pathways of nitrate respiration in bacteria. The major process involved in nitrate transformation to ammonium (NH ₄⁺) is the DNRA pathway (in blue), while those involved in nitrate transformation to N ₂ are denitrification (green), and ammonium oxidation (anammox; red). The main enzymes involved are Nar, a membrane-bound dissimilatory nitrate reductase; Nap, a periplasmic dissimilatory nitrate reductase; NirBD, a nitrite reductase; NrfA and NirK/NirS, nitrite reductases; cNor and qNor, nitric oxide reductases; NosZ, a nitrous oxide reductase; HH, hydrazine hydrolase; HAO, hydrazine oxidoreductase.

Numerous enzymes are involved in these different metabolic pathways, and the coordination of their expression is absolutely required to limit the energetic cost and avoid the accumulation of toxic compounds such as nitric oxide (NO), a free radical which has the propensity to react with other molecules (see below). To correctly regulate the expression of these genes, bacteria use several transcription factors (TFs). One of the most prominent families of transcriptional regulators found to control nitrate respiration is that of the FNR-like proteins.

As it has been shown for other important metabolic pathways, transcriptional regulation is often combined with post-transcriptional control, where non-coding small RNAs (sRNAs) play a key role. Several studies have highlighted the role of sRNAs in nitrogen metabolism and this aspect was reviewed recently ( Prasse and Schmitz, 2018; Muro-Pastor and Hess, 2020). However, recent developments also pointed to an important role for sRNAs in nitrate respiration pathways, which will be addressed in this review.

We will briefly present the diversity of enzymes involved in these metabolic pathways and how the expression of these enzymes is regulated by TFs and sRNAs.

The Diversity of Enzymes Involved in Nitrate Respiration

Nitrate Reductase (Nar, Nap)

For all nitrate respiration pathways mentioned above, the first step consists of the conversion of nitrate to nitrite (NO ₂⁻) by a nitrate reductase. These enzymes use molybdenum as a co-factor. Two types of nitrate reductase have been shown to be involved in nitrate respiration: Nar localized at the membrane and Nap found in the periplasm ( Figure 2). The Nar nitrate reductases are trimeric (NarG/H/I) and encoded by the NarGHJI operon. This operon is widely conserved in bacteria and is found in nitrate-ammonifying (DNRA) bacteria from Escherichia coli to Bacillus subtilis. NarI is a transmembrane protein that receives electrons from the quinone pool and anchors NarG and NarH at the membrane. The electrons are transferred from NarI, a cytochrome b quinol oxidase, to NarG via NarH. NarH and NarG contain Fe-S clusters that receive an electron and transfer it to the active site of NarG ( Figure 2A). The Nar reductase couples nitrate respiration to proton translocation across the membrane, driving ATP generation. NarJ, encoded by the last gene of the operon, is a chaperone protein that plays a role in the maturation and membrane insertion of Nar. Interestingly, E. coli and Salmonella typhimurium possess a duplication of this operon ( narZYWV), which seems to play a greater role in stressed cells rather than anaerobic respiration ( Blasco et al., 1990; Spector et al., 1999). Three nitrate reductases (Nar1, Nar2, and Nar3) have been identified and characterized in Streptomyces coelicolor. Nar1 is induced in spores and competes with aerobic respiration for electrons from menaquinol ( Falke et al., 2019). In contrast, Nar2 and Nar3 are induced by anoxic condition in the mycelium ( Urem et al., 2016; Fischer et al., 2019).

Figure 2. Enzymes involved in dissimilatory nitrate reduction to ammonium (DNRA) in E. coli (A) and in denitrification in P. denitrificans (B). Nar, membrane-bound dissimilatory nitrate reductase; Nap, periplasmic dissimilatory nitrate reductase; NirBD, a sirohaem containing nitrite reductase; NrfA, a cytochrome c nitrite reductase; NirK/NirS, cNor, and qNor, Cu-containing/cd1-type nitric oxide reductases; NosZ, a nitrous oxide reductase; QH2, MKH2, UQH2 : quinone, menaquinone, ubiquinone.

The Nap nitrate reductases are found in the periplasm of many different Gram-negative bacteria (reviewed in Richardson et al., 2007). The NapA subunit, bearing the catalytic active site, is transported across the membrane by the Tat apparatus and requires the NapD chaperone encoded in the same operon ( Grahl et al., 2012). NapA receives electrons from two cytochrome c proteins, the NapC membrane protein and the NapB subunit (review by Richardson et al., 2001) ( Figure 2B). In E. coli, the nap operon also encodes for a quinol-oxidizing system ( napGH) composed of two Fe-S proteins: (1) NapH, a membrane-bound quinol dehydrogenase and (2) NapG, a periplasmic electron transfer adapter protein which can, similarly to NapC, deliver electrons to NapB. However, the exact function of this complex in E. coli is still unclear. The presence of both NapC and NapGH proteins in bacterial genomes is variable. Some bacteria such as Pseudomonas aeruginosa only have the napC gene and others, e.g., ε-proteobacteria such as Campylobacter jejuni or Sulfurimonas denitrificans, only have NapGH ( Sievert et al., 2008; Kern and Simon, 2009; reviewed in Sparacino-Watkins et al., 2014). The nap operon also encodes NapF in E. coli, which is proposed to post-translationally modify NapA prior to its export into the periplasm ( Nilavongse et al., 2006). Interestingly, this protein is absent in its close relative Salmonella. In contrast to the Nar, Nap enzymes are unable themselves to generate a proton motive force. However, Nap can play a role in nitrate respiration when associated with a proton translocating enzyme such as the NADH dehydrogenase NuoA-N ( Moreno-Vivián et al., 1999). In this way, NapA has been shown to support anaerobic respiration in E. coli ( Stewart et al., 2002), and in several bacteria like Bradyrhizobium japonicum ( Bedmar et al., 2005), Rhodobacter sphaeroides ( Liu et al., 1999), Pseudomonas sp. strain G-179 ( Bedzyk et al., 1999), where the Nap enzyme has been shown to be used for the first step of denitrification rather than Nar.

Nitrite Reductase (Nir, Nrf)

The nitrite reductases ensure the reduction of nitrite to NO in denitrifying bacteria (denitrification) and into ammonium in DNRA bacteria, also known as ammonifiers (see below).

In denitrifying bacteria, two types of unrelated nitrite reductases (Nir) have been identified so far (reviewed in Moura and Moura, 2001). One, namely NirK, is an homotrimeric copper-containing enzyme which has been found in both Gram-negative and -positive bacteria ( Suharti and de Vries, 2005). The second, NirS, is a homodimeric cytochrome cd1 nitrite reductase that has been characterized in Gram-negatives ( Figure 2B). Electrons are delivered by the cytochrome bc1 complex. The nirS gene cluster encodes numerous additional proteins (for example the nirXISECFDLGHJN operon in Paracoccus denitrificans) necessary for the correct assembly of the active site haem d1 cofactor of NirS (reviewed in Bali et al., 2014). In contrast to nirS, the nirK gene is generally isolated, but in some cases is in an operon with the nirV gene, whose product may be required for the correct insertion of the copper center. The presence of NirK and NirS has been for a long time proposed to be mutually exclusive. However, recent genomic comparisons have showed that both enzymes can be present in the same organism ( Graf et al., 2014), probably with different roles and regulation, notably in Pseudomonas stutzeri ( Wittorf et al., 2018).

In the nitrate respiratory ammonifiers, such as E. coli and Salmonella enterica, NirK and NirS as well as the nitric oxide reductases (qNor and cNOR, see below) are absent, and the reduction of nitrite is performed by two classes of enzymes:

(1) A cytoplasmic sirohaem-containing nitrite reductase (NirBD), encoded by nirB and nirD ( Figure 2A), with the nirC gene in the same operon probably encoding a nitrite transporter. The nirB and nirD genes are also present in the genome of B. subtilis ( nasDEF). The nasDE genes encode the nitrite reductase subunits and nasF is required for nitrite reductase sirohaem cofactor formation. Interestingly, this enzyme is used for both anaerobic respiration and nitrogen assimilation in B. subtilis ( Ogawa et al., 1995; Nakano et al., 1998).

(2) The periplasmic Nrf enzyme, consisting of a pentahaem cytochrome c nitrite reductase that reduces nitrite by using formate as an electron donor ( Einsle et al., 2002; Figure 2A). NrfA was first identified in E. coli, but is also present in the periplasm of numerous γ-, δ-, and ε-proteobacteria. Similar to the nitrate reductase described previously, electrons are transferred from the membrane quinol dehydrogenase (NrfH or NrfCD) to the cytochrome c reductase NrfA via the small electron transfer protein NrfB (reviewed by Simon, 2002). In E. coli, the nrf operon also encodes three proteins (NrfEFG) dedicated to the attachment of the active site haem group.

Interestingly, NO is also produced by E. coli and S. enterica, despite the fact that they do not possess NirK or NirS enzymes, and is proposed to be a side-product of nitrate respiration. It has been shown that nitrite reduction to NO can be done by the NarG enzyme in Salmonella and represents the major route of NO production in this organism ( Gilberthorpe and Poole, 2008).

Nitric Oxide Reductase (qNOR, cNOR)

The nitric oxide reductases reduce NO into nitrous oxide (N ₂O). They are part of the heme-copper oxidase family but are not able to translocate protons, in contrast to the other enzymes of this family ( Bell et al., 1992; Hino et al., 2010; reviewed in Garcia-Horsman et al., 1994). Two types of NOR have been identified in Gram-negative bacteria: cNOR and qNOR, depending on the electron donor (heme c in cNOR; menaquinol in qNOR). cNOR is composed of two subunits: NorB with the catalytic site receiving electrons from a periplasmic electron donor via the NorC membrane bound cytochrome c ( Figure 2B). cNORs are only found in denitrifying bacteria. Interestingly, a transient interaction between the cNORs and NirS (the cd1 nitrite reductase) has been proposed in P. denitrificans ( Albertsson et al., 2019), and a similar interaction was observed in P. aeruginosa. This interaction is thought to be important for the channeling of the toxic intermediate NO. The NorCB structural subunits are encoded in an operon with other genes ( norEFCBQD). The norQ and norD gene are always linked to norCB, but the norE and norF genes can be encoded separately or even be absent ( Zumft, 2006). The function of these accessory proteins is not well defined. A structural study showed that even if cNOR and qNOR enzymes are not able to pump protons, the catalytic protons can be transferred from bulk water to the buried active site through a specific pathway consisting of a water channel and/or a hydrogen-bond network ( Shiro, 2012).

In contrast to cNOR, the qNOR enzyme is a unique single subunit protein (NorZ) that receives electrons from quinones. The qNOR enzyme has some sequence similarities with both the NorB and the NorC proteins ( Shiro, 2012). The qNOR enzymes have a wider taxonomic distribution and are found in some Archaea and non-denitrifying pathogenic bacteria, where they play a role in NO detoxification ( Hendriks et al., 2000; Sousa et al., 2012).

In the Gram-positive Bacillus azotoformans, a third type of NOR was identified that binds copper (Cu ANOR). This enzyme receives electrons from membrane-bound cytochrome C ₅₅₁ ( Al-Attar and De Vries, 2015) and the large subunit is similar to NorB. The small subunit has no heme c, but a Cu A binding site.

Other non-respiratory enzymes have been identified that can detoxify NO. For example the flavohaemoglobin Hmp which is widespread in bacteria converts NO to NO ₃⁻ ( Vinogradov et al., 2013), and the flavorubredoxin NorV, with its NADH dependent oxidoreductase NorW ( Gomes et al., 2002), reduces NO to NO ⁻. Hmp and NorV are both present in E. coli but their mutation does not impair the reduction of NO, meaning that other NO reduction pathways still need to be characterized ( Vine and Cole, 2011). Since these enzymes are not considered to play a role in respiration, we will not discuss them further in this review.

Nitrous Oxide Reductase (Nos)

Nitrous oxide reductase (Nos) is the last enzyme to complete denitrification by reducing N ₂O into dinitrogen (N ₂). In most denitrifying bacteria, this is a homodimeric copper-enzyme (NosZ) localized in the periplasm ( Figure 2B). However, this enzyme is membrane bound in the Gram-positive bacteria B. azotoformans and Thiobacillus denitrificans ( Hole et al., 1996; Suharti and de Vries, 2005). It was shown in several bacteria, such as P. denitrificans, that electron transfer to Nos is from cytochrome c. NosZ is in general found in an operon with at least 5 other genes ( nosRZDFYL), encoding proteins required for its expression, maturation and maintenance ( Zumft, 2006). This operon sometimes encodes supplementary genes, such as nosX or nosC. In α-, β-, and γ-proteobacteria, the NosR and NosX proteins can bind flavin mononucleotide (FMN) that might be an alternative transport pathway from the quinone pool to NosZ. The NosDFY and L proteins are involved in the maturation of NosZ, and the function of NosC is unknown.

Numerous genomes of ammonifiers also encode an atypical nitrous oxide reductase (cNosZ) ( Sanford et al., 2012; Jones et al., 2013). cNosZ is a variant of the canonical NosZ enzyme, but fused to a cytochrome c domain which transports electrons to the active site ( Simon et al., 2004). Moreover, cNosZ is exported to the periplasm by the Sec secretion pathway and not by the Tat (Twin Arginine translocation) pathway like the classical NosZ enzyme. The genome of Wolinella succinogenes encodes this atypical enzyme in a large operon, nosZBDGC1C2HFYL. The NosG and H are menaquinol dehydrogenases, homologous to the NapGH proteins. The 2 cytochrome c proteins (NosC1 and C2) form a putative electron transport pathway from menaquinol to cNosZ. The nosDFY genes encode a membrane-bound ABC transporter that might be important for the maturation of atypical Nos systems. Finally, NosL could be a chaperone involved in copper metallocenter assembly ( Zumft, 2006).

For a long time, the denitrification and the nitrate ammonification pathways were considered to be mutually exclusive. However, recent genome sequencing data have revealed that some bacteria have all the genes required for both pathways. Shewanella loihica, for example, encodes the NrfA enzyme for the DNRA pathway, and NirK, NorB and NosZ enzymes for the denitrification ( Sanford et al., 2012). Both pathways are functional in this bacterium ( Yoon et al., 2015) and depend on the C/N ratio. Indeed, at low C/N ratio (i.e., electron donor-limiting growth condition) the denitrification pathway is preferred to DNRA. In contrast, at high C/N ratio (i.e., electron-acceptor limiting growth condition), ammonification is dominant ( Yoon et al., 2015). Temperature and pH also influence the nitrate pathways, showing that nitrate respiration is widely connected to environmental conditions.

Some bacteria are also able to release energy by forming N ₂ gas from ammonium and nitrite under anoxic conditions ( Figure 1). This energy pathway (anammox) seems to be limited to the Planctomycetales order. These bacteria have been found in almost all aquatic habitats where oxygen is limiting. This specific pathway requires two type of enzymes, hydrazine hydrolase (HH), which produces hydrazine from ammonium and NO, and hydrazine oxidoreductase (HZO/HAO), which allows the oxidation of hydrazine to N ₂ ( Figure 1). Electrons are thought to be transferred to the quinone pool to ultimately generate a proton motive force.

The numerous enzymes involved in nitrate utilization pathways highlight the need for bacteria to efficiently control their expression at the transcriptional and post-transcriptional levels in response to their specific environments. This control is discussed below, with a focus on the few bacterial species where this has been studied in most detail: E. coli, S. enterica, and B. subtilis for the DNRA pathway, and P. denitrificans and P. aeruginosa for denitrification.

Transcriptional Regulators Controlling the Synthesis of Enzymes Involved in Nitrate Respiration

In bacteria that have the ability to respire on nitrate, this mode of respiration mainly depends on three parameters:

(1) Oxygen limitation

(2) The presence of nitrate (NO ₃⁻) or derived compounds such as nitrite (NO ₂⁻) or nitric oxide (NO) in the medium

(3) An appropriate donor of electrons (i.e., organic carbon compounds).

Several transcriptional regulators play a key role in integrating these signals to modulate the expression of genes related to nitrate respiration. Numerous studies have highlighted the importance of both the FNR-like global TFs controlling several metabolic pathways in order to prepare the cell to anaerobic conditions and the more specific two-component systems (TCSs) that frequently control one metabolic pathway. Most of these regulators can sense different signals like O ₂, NO ₃⁻, NO ₂⁻ or NO (see below). FNR uses an O ₂-labile [4Fe-4S] ²⁺ cluster to sense oxygen. In the absence of O ₂, the cluster is stable and forms a homodimer able to bind its DNA target sequences. In presence of O ₂, the [4Fe-4S] ²⁺ cluster is unstable and the protein cannot dimerize, rendering it inefficient at binding its targets. It has also been demonstrated that the [4Fe-4S] ²⁺ cluster is sensitive to NO in vitro, which also reduces the DNA-binding ability of FNR ( Crack et al., 2013). Depending on the position of the FNR binding site in the promoter region, FNR activates or represses gene expression. In E. coli and B. subtilis, the FNR regulon represents more than 100 genes and its main role is to prepare the cell for anaerobic respiration ( Constantinidou et al., 2006; Reents et al., 2006).

Two-component systems are typically composed of a membrane-bound histidine kinase that senses a specific stimulus and a response regulator that mediates the cellular response by modifying target gene expression. Numerous TCSs, such as NarX-NarL, are involved in nitrate respiration and will be described below. In general, they are more dedicated to nitrate respiration than FNR and their regulon includes less genes than the FNR-like regulon.

This strategy to use both global and more specific transcriptional regulators is similar in numerous bacteria, using either the DNRA or the denitrification pathway to respire on nitrate. This can provide significant advantages that include the integration of several environmental cues, e.g., O ₂ availability via FNR or FNR-like global regulators and the presence of alternative electron acceptors such as nitrate, via the NarX-NarL and NarQ-NarP TCSs. Some variations on this theme exist as well. For example, in B. subtilis and Streptomyces (see below), no transcriptional regulator that directly senses nitrate has been described, i.e., the regulation of the genes involved in the respiration on nitrate is not directly influenced by the presence of nitrate in the environment in this organism.

The different transcriptional regulators, the signals to which they respond, and their targets linked to nitrate respiration are described in the following sections.

Transcriptional Regulation of the DNRA Pathway

A complete review on the conservation of the transcriptional regulators involved in DNRA has been published previously ( Rodionov et al., 2005). Briefly, the global transcriptional regulator FNR sensing O ₂, and the two nitrate-sensing NarQ-NarP and NarX-NarL TCSs have been shown to play a central role in the regulation of anaerobic respiration in E. coli and numerous γ-proteobacteria. In the presence of nitrate, NarX and NarQ phosphorylate NarL and NarP, respectively. Importantly, cross-regulation exists between these two TCSs; NarQ additionally efficiently controls the phosphorylation status of NarL, while NarX can influence NarP much more poorly ( Noriega et al., 2010). Another difference between these two sensors is that NarQ is sensitive to signals other than nitrate, especially nitrite, but not NarX ( Lee et al., 1999). All these transcriptional regulators (FNR, NarXL, and NarQP) are involved in controlling the synthesis of enzymes of the nitrate reduction pathway ( Figure 3). More precisely, they all activate the expression of the nirBDC operon encoding nitrite reductase. The narGHJI operon is activated by NarL and FNR. E. coli and S. typhimurium genomes also encode a second cytoplasmic nitrate reductase ( narZYWV), whose role is still unclear. However, in contrast to the narGHJI operon, these genes are repressed by FNR in S. typhimurium ( Spector et al., 1999). The nap and the nrf genes encoding for the periplasmic nitrate- and nitrite reductase, respectively, are controlled by the NarP response regulator and by FNR ( Figure 3; Jayaraman et al., 1987; Lamberg and Kiley, 2000; Browning et al., 2004). Several other targets of NarX-NarL and NarQ-NarP encode proteins involved in pathways related to nitrate respiration, e.g., NO detoxification ( Filenko et al., 2005) or molybdenum attachment to enzymes such as nitrate reductase ( Hasona et al., 2001). Another important transcriptional regulator involved in nitrate respiration in E. coli is NsrR. This transcriptional repressor has an NO-sensitive [Fe-S] cluster and is not only present in β- and γ-proteobacteria, but also in Streptomyces and Bacillus species ( Rodionov et al., 2005; reviewed in Tucker et al., 2010). Nitrosylation of this factor inhibits its DNA-binding activity, leading to the derepression of the genes controlled by NsrR. In E. coli, NsrR has been shown to regulate 9 operons, including the nitrate- and nitrite reductase operons ( nap and nrf) ( Filenko et al., 2007; Figure 3). In contrast, the nrf operon is not sensitive to the mutation of nsrR in S. typhimurium ( Browning et al., 2010). NsrR was also shown to regulate expression of the Nir and Nor enzymes in denitrifying bacteria such as Neisseria meningitidis ( Heurlier et al., 2008). At last, it should be highlighted that enzymes of the nitrate respiration pathways are most often metalloenzymes that require redox-active cofactors, such as molybdenum, iron or copper (reviewed in Garcia-Horsman et al., 1994, Moreno-Vivián et al., 1999, Sullivan et al., 2013). Interestingly, two regulators of the E. coli nap operon are themselves dependent on such cofactors: ModE, whose activity is induced under anaerobic conditions through molybdenum binding, activates nap expression ( McNicholas and Gunsalus, 2002) while IscR, containing a 2Fe-2S cluster, represses it under aerobic conditions ( Giel et al., 2006). ModE is able to override the control of napF by nitrate when molybdenum is limiting ( McNicholas and Gunsalus, 2002).

Figure 3. Regulation of dissimilatory nitrate reduction to ammonium (DNRA) in E. coli (A) and B. subtilis (B). (A) Nitrate reductase ( nar/nap) and nitrite reductase ( nir/nrfA) genes are co-regulated by FNR and by the two-component system NarQP for the nap, nir, and nrfA operons and NarXL for the nar and nir genes. The nar and nir operons are also repressed by the NsrR TF. narP expression is inhibited by two sRNAs, SdsN, and RprA. Expression of narXL is repressed, probably indirectly, by the DicF sRNA. In contrast, the FNR TF is responsible for fnrS and narS expression. FnrS regulates numerous genes that are dispensable during anaerobic respiration, such as nfsA, encoding an NADPH-dependent nitroreductase. The NarS sRNA is encoded at the 3′ end of the narK mRNA (encoding a transporter of nitrate and nitrite) and represses the expression of nirC, coding for a nitrite transporter. (B) Nitrate reductase gene ( nar) is regulated by FNR. FNR is itself under the control of the ResDE TCS and also controls the expression of nitrite reductase ( nas). NsrR inhibits the expression of the nas operon coding for nitrate and nitrite reductase. ResDE is a transcriptional activator of expression of the RoxS sRNA, which in turn negatively impacts its expression. RoxS regulates numerous genes encoding enzymes of the TCA cycle.

In Mycobacteria, NarL is also responsible for the activation of the narGHJI operon. Although this protein has 74% identity to E. coli NarL, the sensor kinase NarS is not related to NarX. Another TCS, namely DevR/DevS, was identified in Mycobacteria and responds to anaerobic conditions. DevR, the response regulator, also controls the expression of the nar genes and the nitrate transporter NarK2. Interestingly and in contrast to what is known in α- and γ-proteobacteria, the two response regulators DevR and NarL were proposed to form a heterodimer that can efficiently bind the narK2 promoter in Mycobacterium tuberculosis ( Malhotra et al., 2015). Indeed, in vitro binding assays showed that DevR binding on narK2 promoter is stimulated in presence of NarL. More recently, it was shown that the PrrAB TCS, also found in α-proteobacteria (see below), induced the transcription of the DevR response regulator in Mycobacterium smegmatis and is important for ATP generation under aerobic and hypoxic conditions ( Maarsingh et al., 2019).

The NarX-NarL TCS is functionally replaced by the NreABC system in Staphylococci. NreABC has the ability to sense both O ₂ and nitrate, with a higher impact of oxygen compared to nitrate. NreA has a nitrate-binding GAF domain that senses nitrate and NreB, considered as a functional equivalent to FNR, has a [4Fe-4S] ²⁺ cluster that is sensitive to O ₂ ( Müllner et al., 2008). NreB is the sensor histidine kinase and NreC the response regulator of the NreBC TCS. NreB is auto-phosphorylated in anoxic conditions, leading to the phosphorylation of NreC (NreC-P). NreC-P was shown to bind to the GC rich palindromic region of the narGHIJ, nir and narT genes ( Schlag et al., 2008). The NreA protein represents an additional layer of regulation for this TCS. Indeed, in the absence of nitrate, NreA interacts with NreB and inhibits NreB kinase activity ( Niemann et al., 2014; Nilkens et al., 2014).

In B. subtilis, the narGHJI operon is regulated by FNR BS ( Figure 3). In contrast to E. coli FNR, the [4Fe-4S] ²⁺ cluster of FNR BS is at the C-terminus of the protein ( Reents et al., 2006). Interestingly, in this bacterium, nitrate has no direct effect on the narGHJI operon, which seems to be regulated by FNR alone. Indeed, nitrate sensors such as NarXL and NreA have not been identified in B. subtilis. However, on one hand, FNR is transcriptionally regulated by the ResDE TCS that responds to both O ₂ and NO ( Nakano et al., 1996; Nakano and Zhu, 2001; Geng et al., 2007), and on the other, the fnr gene is co-transcribed with the narK gene encoding a nitrite extrusion protein. These results may in part be responsible for the indirect effect of nitrate on nar expression. The NsrR transcriptional repressor also plays a role in nitrate respiration in B. subtilis. The most common target of this regulator in bacteria is the hmp gene, involved in NO detoxification, as observed in B. subtilis ( Kommineni et al., 2012). However, it also regulates numerous other genes, including the nitrate reductase operon ( nasDEF) ( Figure 3; Nakano et al., 2006).

Transcriptional Regulation of the Denitrification Pathway

The regulation of the denitrification pathway has been extensively studied in the γ-proteobacterium P. aeruginosa and in the α-proteobacterium P. denitrificans, and recently reviewed ( Gaimster et al., 2018).

In P. aeruginosa, the denitrification pathway is controlled by the TCS NarXL (sensing NO ₃⁻ and NO ₂⁻), two FNR like-proteins ANR and DNR, and the transcriptional regulator NosR, which is sensitive to NO and O ₂ ( Figure 4). The narK1K2GHJI operon is activated by three transcriptional regulators, the ANR and DNR, and the NarXL TCS. In parallel, it was also shown that NarL represses expression of the nap operon during anaerobic growth ( Van Alst et al., 2009). In the presence of NO, the expression of nirS, norCB and nos genes is activated by DNR ( Arai et al., 1995, 2003). Moreover, ANR also controls the expression of narXL and dnr genes ( Schreiber et al., 2007; Figure 4).

Figure 4. Regulation of denitrification in P. aeruginosa (A) and P. denitrificans (B). (A) Expression of nitrate reductase gene ( nar) is directly co-regulated by the NarXL TCS and the DNR transcriptional regulator. Both regulators (NarXL and DNR) are under the control of the FNR-like regulator ANR. DNR also activates expression of the nitrite reductase gene ( nir). The nitric oxide reductase ( nor) and the nitrous oxide reductase ( nos) genes are repressed by the NosR transcriptional repressor. Transcription of the PaiI sRNA is controlled by the NarXL TCS. (B) Expression of nitrate reductase gene ( nar) is co-regulated by NarR and FnrP. Expression of nitrite reductase ( nir) and nitric oxide reductase ( nor) genes are activated solely by NnrR. Expression of nitrous oxide reductase gene ( nos) is co-regulated by FnrP and NnrR. NirI represses the expression of nir, while NosR repressed that of nos. Moreover, the recently identified DenR (also known as sRNA-29) inhibits nir, nor and nos expression, probably indirectly via a regulator of the GntR type.

In P. denitrificans, three FNR-like proteins are involved in the regulation of denitrification: FnrP and NnrR that sense O ₂ and NO levels, and NarR that senses NO ₃⁻ and NO ₂⁻ ( Lee et al., 2006; Bergaust et al., 2012; Crack et al., 2016; Figure 4). However, in contrast to FnrP, NnrR, and NarR lack the [4Fe-4S] ²⁺ cluster. The NarR TF functionally replaces the NarXL TCS in α-proteobacteria, since no homologs have been identified so far (reviewed in Bueno et al., 2012). The narGHIJ operon is fully induced under anaerobic conditions via FnrP, and by nitrate via the NarR TF ( Wood et al., 2001). The NnrR TF controls the expression of the nir, nor, and nos gene clusters ( Figure 4; Saunders et al., 1999; Van Spanning et al., 1999; Bergaust et al., 2012). Finally, nos gene expression is also regulated by FnrP ( Crack et al., 2016). In addition to these global regulators, two related proteins NirI and NosR, with metal ion-binding or thiol sites (CXXXCP motif), were specifically shown to regulate the expression of the nir and nos genes, respectively ( Saunders et al., 1999). As mentioned above, the detrimental effect of NO is avoided by the interaction between nitric oxide reductase and nitrite reductase in P. denitrificans and P. aeruginosa. However, their co-regulation by NnrR is another way to control NO accumulation.

In the α-proteobacteria B. japonicum, Rhodobacter capsulatus, Sinorhizobium meliloti, and R. sphaeroides, similar redox sensitive TCSs have been identified, namely RegSR, RegBA, ActSR, and PrrBA, respectively. All these TCSs share a high sequence similarity in their C-terminal domain. They are involved in the regulation of different metabolic pathways, including nitrogen fixation, and can functionally replace each other ( Emmerich et al., 2000). The FnrL regulator and the PrrBA TCS are important for photosynthetic gene expression during anaerobic growth of R. sphaeroides. PrrAB was also shown to induce expression of the nitrate reductase NirK, since a PrrA deletion decreases expression of the nirK-lacZ fusion ( Laratta et al., 2002). Another key regulator in R. sphaeroides is the FNR-like protein NnrR, that senses NO, and that is also required for the expression of nirK, as well as that of the norCBQD operon encoding the nitric oxide reductase ( Kwiatkowski and Shapleigh, 1996; Tosques et al., 1996; Laratta et al., 2002). This result shows that the NnrR regulon is indirectly controlled by the PrrAB TCS, since NO (synthetized by nirK) is necessary for NnrR function, and suggests that, in contrast to other denitrifying bacteria, R. sphaeroides uses nitrate respiration to ensure redox balancing during photosynthesis, rather than to support anaerobic growth.

In the ε-proteobacterium W. succinogenes, the nap, nrf, and cnos genes involved in nitrate respiration are upregulated in presence of nitrate, NO or N ₂O but not by nitrite or hydroxylamine ( Kern and Simon, 2016). Three TFs, belonging to the Crp/FNR superfamily, are involved in the upregulation of these genes in specific growth condition, namely NssA, B, and C. Nss factors have an N-terminal effector domain and a C-terminal DNA binding domain. Further study will be necessary to understand how nitrate, NO and N ₂O are sensed by these different TFs. These TFs are homologous to NssR, previously identified in C. jejuni and involved in NO stress resistance ( Elvers et al., 2005). However, in C. jejuni, NssR binding sites have not been identified upstream of the nap and nrf genes ( Elvers et al., 2005).

In S. coelicolor, which possesses three nitrate reductases (Nar1, Nar2, and Nar3), the OsdRK TCS controls the expression of the narGHJI2 operon ( Fischer et al., 2019). Interestingly, Nar1 and Nar3 are not under the control of this TCS, but are induced under anaerobic conditions.

Finally, in the γ-proteobacterium S. loihica PV-4, the global regulator CRP1, which is widely present in bacteria, is necessary to be able to use nitrate in the DNRA pathway, while its paralog CRP2 is required for transcription of the nitrite reductase gene nirK for denitrification ( Liu et al., 2020).

Other transcriptional regulators are involved in the control of genes linked to NO detoxification, such as NorR in E. coli which regulates norV and norW and have previously been reviewed by Spiro (2007). However, we consider these transcriptional regulators as beyond the scope of this review.

In addition to such complex transcriptional control of the respiration processes, it has more recently become evident that an additional layer of post-transcriptional regulation by non-coding RNAs is also crucial in the regulation of cellular respiration. sRNAs can rapidly evolve to adapt to specific conditions (reviewed by Dutcher and Raghavan, 2018) and, not surprisingly, recent discoveries have confirmed a role for sRNAs in nitrate respiration (see also Table 1).

Table 1. sRNAs that could play a role in nitrate respiration.

Small RNAs Involved in Nitrate Respiration

sRNAs Involved in the DNRA Pathway

Small RNAs involved in the respiration on nitrate have been studied in greatest detail so far in the enterobacteria E. coli or S. enterica, where a wealth of data points to the involvement of several sRNAs in the DNRA pathway. As stated above, the NarX-NarL and NarQ-NarP TCSs play key roles in the regulation of the enzymes involved in this process, and a first link between sRNAs and nitrate respiration came from the identification of sRNAs that target these regulators.

SdsN ₁₃₇ is one of three sRNA isoforms and it represses expression of the narP response regulator gene by pairing to the translation initiation region of this mRNA ( Hao et al., 2016). Interestingly, the two other confirmed targets of the SdsN sRNAs are also involved in nitrogen metabolism, as they encode NfsA, a major NADPH-dependent nitroreductase, and HmpA, a nitric oxide dioxygenase. In agreement with the role of NfsA in governing E. coli sensitivity to nitrofuran antibiotics, SdsN was found to protect cells against these compounds ( Hao et al., 2016). The same study showed that the SdsN levels respond to the stress/stationary phase sigma factor RpoS, but also change in response to different nitrogen sources (e.g., ammonium or arginine), in a NarP-dependent manner. It is still unclear, however, whether RpoS and NarP are direct regulators of sdsN gene transcription.

More recently, RprA was identified as a second sRNA that directly controls narP mRNA expression through direct base-pairing with its translation initiation region. Of note, in addition to this canonical regulation, RprA also represses narP by acting at the very 5′ end of the narP mRNA, more than 100 nucleotides upstream of the narP start codon ( Brosse et al., 2021). Even though the precise mechanism for this additional control is unclear, it is most likely to be indirect, suggesting the existence of yet other narP post-transcriptional regulators. Transcription of RprA sRNA is primarily controlled by the Rcs phosphorelay that responds to envelope stress ( Majdalani et al., 2002), and previously reported targets of RprA include the gene encoding the RpoS global stress response protein, and others involved in biofilm formation, conjugation, acid resistance ( Majdalani et al., 2001; Mika et al., 2012; Papenfort et al., 2015; Lalaouna et al., 2018), showing that this sRNA is not just dedicated to the control of the nitrate respiration.

Changing the levels of a regulatory protein may not always lead to an effect on its regulon ( Batchelor and Goulian, 2003). This is especially true in the case of response regulators that most often need to be modified by phosphorylation to be active. At least in the case of narP control by RprA or SdsN ₁₃₇ sRNAs, the overproduction of either sRNA is sufficient to repress the expression of the nap operon, via their effect on narP regulation, suggesting that control by these sRNAs can really impact nitrate respiration ( Brosse et al., 2021).

The narL mRNA, which encodes the response regulator of the NarX-NarL TCS, is also likely subjected to post-transcriptional control by the DicF sRNA. This sRNA accumulates in low oxygen conditions and its gene is present in one copy in the commensal E. coli K-12 genome, but in four copies in the enterohemorrhagic strain 0157:H7 (EHEC) ( Murashko and Lin-Chao, 2017; Melson and Kendall, 2019). DicF regulates the expression of genes important for cell division and metabolism ( Tetart and Bouché, 1992; Balasubramanian et al., 2016) and enhances EHEC virulence ( Melson and Kendall, 2019). narL was among the mRNAs that were differentially expressed in RNAseq analysis performed in the EHEC strain in the presence and the absence of DicF. qPCR data further confirmed that narL mRNA levels increase in the strain deleted for the four dicF genes, and showed that overproduction of any one of the four DicF sRNAs reversed this effect. Whether this is due to a direct interaction between DicF and the narL mRNA, and whether this control is conserved in other bacteria are still open questions.

Finally, there are some indications that the RyhB sRNA could also play a role in the expression of narP and/or narL mRNAs. More precisely, RyhB overproduction significantly repressed the synthesis of a narP-lacZ translational fusion in E. coli ( Hao et al., 2016), while qRT-PCR assays suggested that the two Salmonella RyhB homologs, also called RfrA and RfrB, can activate narL and repress narP expression ( Teixidó and Cortés, 2010). Again, precise mechanisms for these controls remain to be determined, but a convincing pairing can be predicted between RyhB and the narP mRNA, suggesting a direct interaction in this case ( Hao et al., 2016).

As stated above, many enzymes involved in nitrate respiration require metal cofactors such as iron and it is thus interesting that the iron-responsive RyhB sRNA may participate in the regulation of these pathways. This is further exemplified by the regulation of the genes encoding the NirBD nitrite reductase in enterobacteria, since nirB, the first gene of the nirBDC-cysG operon, was predicted as a RyhB target using the CopraRNA pairing prediction algorithm. This interaction was validated by compensatory mutations showing that RyhB represses nirB expression in E. coli ( Wright et al., 2013). Conversely, however, the effects of the two RyhB homologs in Salmonella that have been proposed to activate nirBDC expression upon nitrosative stress may be indirect ( Calderón et al., 2014).

In addition to sRNAs targeting TFs of the nitrate respiration pathway such as the NarP or NarL response regulators, several sRNAs whose transcription depends on these TFs were also identified. A first example is the NarS sRNA whose levels respond to FNR and NarL TFs ( Wang et al., 2020). NarS has been recently identified as an sRNA derived from the 3′ UTR of the narK mRNA, encoding a nitrate and nitrite transporter, through processing by the essential endonuclease RNase E. Transcription of narK is under the direct control of FNR and NarL, which explains how these TFs also control the accumulation of the NarS sRNA. Interestingly, NarS is a Hfq-binding sRNA that represses the expression of the nirC gene, while leaving the expression of the other cistrons of the nirBDC-cysG operon unchanged ( Wang et al., 2020). As NirC is involved in nitrite uptake, it was proposed that the negative control of nirC by NarS could limit nitrite re-import under conditions of narS expression, where nitrite can be exported via NarK or reduced to ammonium.

Other cases of sRNAs whose transcription is under the control of TFs of the nitrate respiration pathway have been reported. As previously mentioned, FNR is a major regulator of gene expression in response to anaerobiosis. It regulates hundreds of genes, including genes whose products are directly related to nitrate respiration such as the nap, nir, and nrf operons in E. coli (see above). The FnrS sRNA was recognized early on as part of the FNR regulon ( Durand and Storz, 2010) and is thus expressed under anaerobic conditions. Of note, fnrS transcription is also controlled by two other major TFs, ArcA (positively) and CRP (negatively), which allows further accumulation of FnrS, under anaerobic conditions and in glucose-based media, respectively ( Durand and Storz, 2010). More than 30 mRNAs are repressed following FnrS pulse-overproduction; they encode proteins involved in diverse functions, such as central metabolism, energy metabolism, aerobic respiration, oxidative stress, that are dispensable under anaerobiosis. FnrS-targets include the NfsA nitroreductase, whose mRNA is also targeted by SdsN, and the CydDC glutathione transporter, important for periplasmic redox control. Interestingly, expression of the cydDC operon is additionally controlled at the transcriptional level by ArcA, NarL, and FNR, with these two latter regulators allowing induction of expression under anaerobic conditions in the presence of alternative electron acceptors, such as nitrate or nitrite ( Cook et al., 1997).

Importantly, the ArcA response regulator not only contributes to FnrS synthesis, but also directly represses the transcription of ArcZ, another Hfq-dependent sRNA that targets multiple mRNAs, including rpoS, eptB, sdaC, tpx, mutS, or tolC ( Papenfort et al., 2009; Mandin and Gottesman, 2010). The ArcB-ArcA TCS is activated under microaerobic conditions ( Alexeeva et al., 2000) and its regulon includes many genes involved in aerobic respiration, thereby partially overlapping the FNR regulon. In addition, and in contrast to FNR, ArcB-ArcA does not directly control expression of genes of the nitrate respiration pathway. However, recent data from RNA-RNA interactome studies strongly support an interaction between the ArcZ sRNA and the napF mRNA and hence suggest an indirect role for ArcA in napF expression through the ArcZ sRNA ( Melamed et al., 2016, 2020).

Transcriptional control of sRNAs by TFs involved in nitrate respiration also is true in Gram-positive bacteria. Transcription of RoxS, the only base-pairing sRNA that is conserved in B. subtilis and Staphylococcus aureus (where it is known as RsaE), is for instance directly controlled by the ResDE TCS (or its homolog SsrAB in S. aureus), allowing RoxS induction in the presence of NO ( Durand et al., 2015). RoxS has many targets, several of which are involved in carbon and energy metabolism, or in amino acid transport. However, it is not yet clear whether this sRNA plays a direct role in nitrate respiration.

sRNAs Involved in the Denitrification Pathway

Although the biology of sRNAs in general has been less studied in denitrifying bacteria, it is already clear that sRNAs are involved in controlling denitrification. A recent transcriptome analysis determined the sRNA composition of P. denitrificans, the model bacterium for the study of denitrification, under different denitrifying conditions: N ₂O-producing or N ₂O-consuming during anaerobiosis, versus aerobic growth without N ₂O ( Gaimster et al., 2016). These conditions were set-up by adding CuSO ₄ to the medium (N ₂O-consuming) or omitting it (N ₂O-producing), as expression of the nitrous oxide reductase NosZ, itself a copper-dependent metalloenzyme required for the last step of the denitrification pathway, depends on copper ( Sullivan et al., 2013). This study allowed the identification of several sRNAs with a potential role in denitrification. Among these, the DenR sRNA (for denitrification repressor, previously known as sRNA-29) was shown later to impact denitrification by repressing the expression of several genes involved in this process, such as nirC, norBC, or nosZ, encoding nitrite reductase, subunits of nitric oxide reductase, and nitrous oxide reductase, respectively. These controls are likely indirect and mediated by DenR promoting the synthesis of a GntR-type transcriptional regulator, whose expression follows that of the sRNA, i.e., is highest in the absence of N ₂O and decreases as N ₂O levels increase ( Gaimster et al., 2019). Of note, this transcriptional regulator was not previously known to control denitrification. As a result, DenR overproduction stalls denitrification at the nitrite reduction step, thereby increasing nitrite levels, while limiting production of NO and N ₂ O.

In P. aeruginosa, another denitrifying bacterium, the 126-nt PaiI sRNA (for P. aeruginosa anaerobically induced RNA I) was identified by RNAseq as the most enriched sRNA under anoxic conditions. Further work showed that PaiI accumulates under anaerobic conditions in the presence of nitrate in an ANR- and NarXL-dependent manner ( Tata et al., 2017). The presence of a close-to-consensus NarL site in the PaiI promoter sequence additionally suggests that NarL directly controls paiI transcription, while the ANR-regulation is most likely indirect via activation of NarXL by ANR ( Figure 3). Interestingly, PaiI-deleted cells display defects in survival after a shift to anoxic conditions in the presence of nitrate and the activity of nitrite reductase is decreased in the absence of the sRNA ( Tata et al., 2017). The molecular basis of these observations is still unclear, but these data nonetheless clearly point to a role for the PaiI sRNA in denitrification.

The PcrZ sRNA of the α-proteobacterium R. sphaeroides is another example of an sRNA responding to regulators of denitrification genes. PcrZ synthesis is induced under low oxygen conditions and this is dependent on the PrrA response regulator and the FnrL TF ( Mank et al., 2012). While the presence of a consensus site for PrrA in the pcrZ promoter strongly suggests a direct regulation, this may not be the case for FnrL control ( Mank et al., 2012). Both PrrA and FnrL are key regulators of R. sphaeroides anaerobiosis gene expression, and their regulons include genes for the photosynthesis apparatus. In turn, the PcrZ sRNA represses expression of several photosynthesis genes through direct pairing with their mRNAs. Interestingly, transcription of these mRNAs is itself activated by PrrA and FnrL; PcrZ thereby participates in an incoherent feed-forward loop to regulate the expression of photosynthesis genes ( Mank et al., 2012). PcrZ does not appear to be directly involved in denitrification at this stage, but is more likely to play a fine-tuning role in the metabolic versatility of R. sphaeroides.

It was also recently proposed that an antisense RNA is produced from the intergenic region between the norQ and norD genes of the norECBQD operon, encoding an NO reductase in the facultative denitrifying bacterium Agrobacterium fabrum ( Lecomte et al., 2020). A possible role for this antisense RNA could be to promote denitrification, since N ₂ O production is reduced more than three-fold after inactivation of this antisense RNA. However, further studies are required to both confirm the existence of this regulatory RNA and address its precise mode of action.

Conclusion

Nitrate respiration is an important metabolic pathway, especially when oxygen is limiting, and is regulated at multiple levels. An important control of the nitrate respiration process is exerted at the transcriptional level allowing the integration of numerous environmental signals to precisely control the expression of enzymes involved in this pathway. More recent studies have shown the additional importance of post-transcriptional regulation: the involvement of sRNAs in nitrate respiration is beginning to be recognized, even though this is so far restricted to only a few model bacterial species. In addition, even in these species, their role is very likely still underestimated. Many mRNAs encoding either enzymes or TFs of the DNRA pathway were for instance significantly enriched upon Hfq co-immunoprecipitation in several studies in enterobacteria, suggesting that they could be subject to post-transcriptional control by base-pairing sRNAs as well. Furthermore, high-throughput analyses relying on the ligation of interacting RNAs following their co-immunoprecipitation with Hfq or ProQ, identified many more putative sRNAs interactions with mRNAs whose products are related to the DNRA pathway (e.g., several nap, nar, nrf, or nir genes) ( Melamed et al., 2016, 2020). In addition, in S. meliloti, Hfq was found to directly interact both upstream and within the fixLJ ORF, encoding an O ₂₋sensing TCS, and to play a role in its expression ( Gao et al., 2016). This control is expected to both directly and indirectly modulate the expression of FixLJ targets, including the nif operon ( nifHDK). Although it is still unknown whether fixLJ regulation by Hfq involves an sRNA or not, this is yet another illustration that the post-transcriptional control of nitrate respiration genes is far from being fully understood.

Finally, it is important to emphasize that, at least in the non-denitrifying bacteria, most studies of gene regulation by sRNAs are performed under aerobic conditions, where expression of genes for nitrate respiration may be much lower than under anaerobic conditions. There is no doubt that additional work under more favorable conditions for nitrate respiration would unravel novel sRNA regulations.

Overall, it thus appears that, as for other important metabolic pathways, nitrate respiration, and more generally nitrogen metabolism, are precisely controlled by multiple regulators acting both at the transcriptional and post-transcriptional levels. In addition to these controls of gene expression, regulation of nitrate respiration could be achieved by modulating the activity of enzymes. Indeed, it was shown that the activity of the E. coli NarGHI nitrate reductase was controlled by the dynamics of the complex, that assembles to the cell poles under anaerobic conditions ( Alberge et al., 2015). As more details of these regulatory mechanisms are revealed, ideally in a large number of bacterial species, it will be interesting to determine the relative contributions of the diverse regulators involved and their respective advantages in relation to the different nitrate respiration pathways and the environmental conditions encountered by bacteria.

Author Contributions

Both authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

Funding

Research in our groups is supported by the CNRS, the "Initiative d'Excellence" program from the French State (Grant "DYNAMO," ANR-11-LABX-0011), the Agence Nationale de la Recherche (ANR grants BaRR-16-CE12-0002-01 and CoNoCo-18-CE12-0025-02 to SD), and the European Research Council (ERC) under the European Union's Horizon 2020 Research and Innovation Programme (Grant Agreement No. 818750 to MG).

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

We are grateful to Mathias Springer and Ciaran Condon for critical reading of the manuscript.

References

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Lee, Y. Y., Shearer, N., and Spiro, S. (2006). Transcription factor NNR from Paracoccus denitrificans is a sensor of both nitric oxide and oxygen: isolation of nnr ∗ alleles encoding effector-independent proteins and evidence for a haem-based sensing mechanism. Microbiology 152, 1461-1470. doi: 10.1099/mic.0.28796-0

Mika, F., Busse, S., Possling, A., Berkholz, J., Tschowri, N., Sommerfeldt, N., et al. (2012). Targeting of csgD by the small regulatory RNA RprA links stationary phase, biofilm formation and cell envelope stress in Escherichia coli. Mol. Microbiol. 84, 51-65. doi: 10.1111/j.1365-2958.2012.08002.x

Murashko, O. N., and Lin-Chao, S. (2017). Escherichia coli responds to environmental changes using enolasic degradosomes and stabilized DicF sRNA to alter cellular morphology. Proc. Natl. Acad. Sci. U. S. A. 114, E8025-E8034. doi: 10.1073/pnas.1703731114 Google Scholar

# data[].attributes.attachment_link.title
Transcriptional and Post-transcriptional Control of the Nitrate Respiration in Bacteria

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In oxygen (O2) limiting environments, numerous aerobic bacteria have the ability to shift from aerobic to anaerobic respiration to release energy. This process requires alternative electron acceptor to replace O2 such as nitrate (NO3-), which has the next best reduction potential after O2.

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Identificación de enterobacterias por la capacidad que tienen para reducir los nitratos a nitrito (Escherichia coli y Klebsiella pneumoniae, son las enterobacterias que producen más carga de enfermedad en los seres humanos)

Esta prueba se emplea en la identificación bioquímica de microorganismos entéricos, no entéricos (Neisseria gonorrhoeae) y de vibrionáceas como V.cholerae y V.parahaemolyticus.

# data[].attributes.attachment_link.title
Combinaciones de bases nutritivas empleadas en el desarrollo del medio caldo nitrato

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Reacción redox

NO₃- + 2H⁺ + 2e⁻ ---> NO₂⁻+H₂O(E₀= 0.43v)
Zn ---> Zn2+ + 2 e- (E₀=0.76 v)
_____________________________________________________________
NO₃⁻+2H⁺+Zn ---> NO₂⁻+H₂O+Zn²⁺

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Si no hay producción de gas y el resultado es negativo

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Existe la posibilidad de que la bacteria tenga la capacidad de reducir el nitrato hasta amoniaco

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Se reporta como:

1. Nitrato positivo: Cambio de color (rosa/rojo) en el medio después de adicionar los reactivos A y B o si después de agregar zinc en polvo no hay cambio de color en el medio

2. Nitrato negativo: No hay cambio de color en el medio despues de agregar los reactivos A y B o si después de agregar zinc en polvo hay un cambio de color en el medio (rosa/rojo)

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Among the human Neisseria and related species, three species - N. mucosa, M. catarrhalis, and K. denitrificans reduce nitrate. The nitrate reduction test is a critical test for differentiating between N. gonorrhoeae and , particularly when strains of K. denitrificans appear to be gram-negative diplococci in stained smears.

Bacterial species may be differentiated on the basis of their ability to reduce nitrate to nitrite or nitrogenous gases. Among the Neisseriaceae of human origin, strains of Neisseria mucosa, Moraxella catarrhalis, and Kingella denitrificans reduce nitrate. Strains of M. catarrhalis and K. denitrificans have been misidentified as N. gonorrhoeae. The nitrate reduction test permits differentiation between these species which are nitrate positive and N. gonorrhoeae (nitrate-negative). The reduction of nitrate may be coupled to anaerobic respiration in some species.

The biochemical pathway involved in nitrate reduction is shown in Figure 1. Nitrate is reduced to nitrite which may then be reduced to nitric oxide, nitrous oxide, or nitrogen (Figure 1).

Figure l. Nitrate reduction pathway.

The nitrate reduction test is based on the detection of nitrite in the medium after incubation with an organism. If present in the medium, nitrite will react with sulfanilic acid (Nitrate reagent A) to form a colorless complex (nitrite-sulfanilic acid). This complex will then yield a red precipitate (prontosil) when nitrate reagent B (alpha-naphthylamine) is then added to the test as shown in Figure 2.

Figure 2. Diagrammatic representation of the detection of nitrite in medium.

A red color will be produced in the medium only when nitrite is present in the medium. Lack of a red color in the medium after the addition of sulfanilic acid and alpha-naphthylamine means only that nitrite is not present in the medium. There may be two explanations for this observation.

The nitrate may not have been reduced; the strain is nitrate-negative.

The nitrate may have been reduced to nitrite which has then been completely reduced to nitric oxide, nitrous oxide, or nitrogen which will not react with the reagents that react with nitrite; the strain is nitrate-positive.

Any test medium that gives a negative result after the addition of the nitrate reagents must be further tested to determine which of the two interpretations is accurate.

A successful nitrate reduction test is dependent on performing the test under the correct conditions.

1. The reaction will occur best if the base medium supports growth of the organism. However, although some Neisseria species do not grow well in broth media, the nitrate reduction test may be performed successfully in a medium that does not support growth by inoculating the medium heavily to provide sufficient preformed enzyme for the reaction to occur.

2. Nitrate reaction occurs only under anaerobic conditions. The nitrate-containing medium is dispensed in tubes to give a low surface area:depth ratio which limits the diffusion of oxygen into the medium, e.g., 5 ml of medium is dispensed in a 13 mm diameter tube. Neisseria and related species use the oxygen in the medium and rapidly produce anaerobic conditions which are ideal for the reduction of nitrate to occur.

The nitrate reduction test is performed in a medium containing 0.2% potassium nitrate. The medium is inoculated heavily with a pure culture of the suspect organism and incubated at 35C to 36.5C for 48 h. in an incubator with or without supplemental carbon dioxide.

Nitrate reduction is detected with the Griess Llosvay reagents, sulfanilic acid and alpha-naphthylamine. Sulfanilic acid (Nitrate reagent A) is added to the incubation mixture and forms a complex (nitrite-sulfanilic acid) with any nitrite present in the medium. When alpha-naphthylamine (Nitrate reagent B) is added to the incubated medium, a red precipitate (prontosil) will be formed with any nitrite-sulfanilic acid complex present in the medium.

An organism may be reported as nitrate-positive if a red color develops in the medium after Nitrate reagents A and B are added to the medium, indicating that the organism has reduced nitrate to nitrite.

The absence of a red color after the addition of both reagents does not automatically mean that the organism is unable to reduce nitrate. Strains may have reduced the nitrate to nitrite, and then reduced the nitrite completely to nitrogenous gases which are not detected when Nitrate reagents A and B are added to the medium. If the medium does not change color after the addition of sulfanilic acid and alpha-naphthylamine, a small amount ("knife point") of zinc dust is added to the incubated medium. The zinc dust will catalyze the reduction of nitrate to nitrite chemically. Thus, if the nitrate has not been reduced by the organisms, i.e., they are nitrate-negative, it will be reduced by the zinc dust and a red color will develop in the incubated medium within 15 min. If no color develops in the incubated medium after the addition of zinc dust, the organisms have not only reduced nitrate to nitrite, but have reduced nitrite to nitrogenous gases; these organisms are also nitrate-positive.

Although nitrate medium is supplied with inverted Durham tubes to detect gas production, gas production is not recorded for Neisseria species. Although some species may reduce nitrate beyond nitrite to nitrogenous gases, gas may not accumulate in the tube. The accumulation of gas is dependent on the rate at which it is produced. When gas is produced very slowly, it may dissolve in the medium and not accumulate in the Durham tube.

Optimum specimen: A pure culture of a suspect gram negative, oxidase-positive diplococcus ( Neisseria spp. or M. catarrhalis) on chocolate agar incubated in a carbon dioxide-enriched atmosphere at 35C to 36.5C for 18 to 24 h.

Unacceptable specimen: Cultures of isolates on chocolate agar incubated in a carbon dioxide- enriched atmosphere at 35C to 36.5C for more than 24 h.
Compromising factors affecting test results:

Test medium must be inoculated heavily enough to permit reaction to occur with preformed enzymes. Insufficient inoculum may not permit organisms to use up oxygen to produce anaerobic conditions in which nitrate reduction can occur.

Too much zinc dust added to the incubated tube may result in very rapid reduction of nitrate beyond nitrite to nitrogenous gases so that nitrite is not detected.

Stability of specimen: Detection of nitrate reduction for Neisseria and related species is dependent on the presence of preformed enzymes.

Tests should only be performed with inoculum harvested from 24 h cultures.

Nitrate medium should be inoculated within 30 min. of removing culture from the incubator; prolonged exposure of the culture at room temperature may result in diminished enzyme activity.

Medium: Nitrate broth (Heart infusion broth containing 0.2% potassium nitrate)

Heart infusion broth (Difco), 25.0 g
Potassium nitrate, 2.0 g
Distilled water, 1000.0 ml

1. Dissolve the ingredients in distilled water; adjust solution to pH 7.0.

2. Dispense 5 ml aliquots of the broth into 16 mm x 100-mm tubes with gas inserts (Durham tubes, 6-mm x 50-mm).

3. Autoclave for 15 min at 121C.

Store medium at 4C to 10C (refrigerated) until used. Prewarm the medium to room temperature before inoculation.

Reagents: Sulfanilic acid solution (Nitrate reagent A): 0.8% in 5N acetic acid
Chemical Name: 4-aminobenzene sulfonic acid
Store Nitrate reagent A at 15C to 30C (room temperature) for up to 3 months, in dark. Reagents may be stored in dark brown glass containers; bottles may be wrapped in aluminum foil to ensure darkness.

Alpha-Naphthylamine solution (Nitrate reagent B): 0.6% in 5N acetic acid
Chemical Name: N,N-dimethyl-1 naphthylamine
Store Nitrate reagent B at 2C to 8C (refrigerated) for up to 3 months, in dark. Reagents may be stored in dark brown glass containers; bottles may be wrapped in aluminum foil to ensure darkness.

Zinc powder, Reagent grade: Store at room temperature (15C to 30C)

Warning: Acetic acid is corrosive. Contact with skin may cause blisters and burns. In case of contact, flush eyes and skin immediately with plenty of water (for at least 15 min.)

QC strains:

Nitrate reductase-positive control: Kingella denitrificans, CDC 10,236

Nitrate reductase-negative control: Neisseria gonorrhoeae, ATCC 43069

QC strains are stored at -70C in a solution of tryptic soy broth containing 20% glycerol. Reactions of control strains should be confirmed at the time frozen stocks are prepared. QC strains may be stored at -70C for up to 2 years.
Procedure:
QC strains are tested in the same manner as clinical isolates. QC strains should be subcultured at least once after the initial culture from the frozen specimen before the test is performed. Clinical isolates may be subcultured from selective medium or purified subcultures. Ensure that cultures are pure.

1. Thaw vials of control strains stored at -70C.

2. Streak onto plates of chocolate agar or supplemented GC agar for isolation. Incubate at 35C to 36.5C in a carbon dioxide-enriched atmosphere for 18 to 24 h.

3. With a sterile swab, prepare a heavy suspension of well-isolated colonies from a pure culture of the isolate incubated on chocolate medium at 35C to 36.5 C in a carbon dioxide-enriched atmosphere for 18 to 24 h. Inoculate test medium to give heavy turbidity.

Note: Strains of N. gonorrhoeae and some other Neisseria spp. may not grow in this medium. Thus, the reaction may be dependent on preformed enzyme.

4. Incubate inoculated media and an uninoculated medium control medium tube at 35C to 36.5 C in a carbon dioxide-enriched atmosphere for 48 h.

5. After 48 h. incubation add, with Pasteur pipettes, 5 drops of Reagent #A, followed by 5 drops of Reagent #B to each tube. Shake the tube well to mix reagents with medium.

Examine the suspension for a pink-red color which should develop within a few minutes if the medium is still warm. The reaction may take a little longer if the medium is cold when the reagents are added.
If the suspension turns pink-red before the addition of Zn powder, the recation is positive and the test is completed. Do not perform step 4.
Reactions observed with the uninoculated medium control, and nitrate-negative, and nitrate-positive isolates are illustrated in Figure 3, Figure 4, and Figure 5, respectively.

6. If the suspension is colorless after the addition of reagents A and B, add a small amount (4 to 5 mg; "sharp knife point") of zinc dust to the medium. Shake the tube vigorously and allow it to stand at room temperature for 10-15 min.

If the medium remains colorless after the addition of Zn powder, the test result is positive.
If the medium turns pink after the addition of Zn powder, the result is negative.

7. Read and record results.

Figure 3. Reaction Observed with Uninoculated Nitrate Medium.

Figure 4. Reaction Observed with Nitrate-Negative Species.

Figure 5. Reactions Observed with Nitrate-Positive Species.

Quality Control Schedule:

A nitrate reductase QC test is performed each day clinical isolates are tested.

The nitrate reduction test may give false-negative or false positive results if the medium is not produced accurately or the test is not performed accurately. The reaction in this test is dependent upon a number of factors.

Failure to detect pink color in the uninoculated medium control tube after the addition of Zn dust may be due (1) to the medium not containing nitrate or (2) to the addition of too much zinc dust which has catalyzed the reduction of nitrate beyond nitrite to nitrogenous gases. The simplest solution is to obtain more nitrate medium ensuring that nitrate has been added to the base medium. Alternatively, inoculate the medium with a positive control strain, but test for a reaction after a shorter incubation time; strains of N. mucosa will produce a positive nitrite reaction after a few hours incubation. If the medium is confirmed to contain nitrate, repeat the test until you have determined the correct amount of zinc dust to add. It is critical to learn how much zinc dust to add to the test. Addition of too much zinc dust can result in a false-positive result.

If a pink color is detected in the uninoculated medium control after nitrate reagents A and B have been added to the medium, the medium is contaminated with nitrite. The only solution is to obtain a new batch of medium that is not contaminated with nitrite.

In medium containing nitrate, the failure of the positive control strain, Kingella denitrificans, to give a positive reaction would occur only if the strain is not K. denitrificans. Recheck the identity of the positive control strain. Select a new culture of the control strain and repeat the test. Similarly, if a positive nitrate reductase test is obtained with the negative control strain, N. gonorrhoeae, either the negative control strain is not N. gonorrhoeae or the culture is contaminated with a nitrate-positive organism. Recheck the purity and identity of gonococcal reference strain. Repeat test with a pure culture of a confirmed culture of N. gonorrhoeae.

The nitrate reduction reaction indicates the ability of organisms to reduce nitrate, a reaction that occurs only under anaerobic conditions; the reaction will not occur if the organisms receive a continuous supply of oxygen. Thus, the reaction may not occur in still cultures (particularly of slow-growing species) in which the medium is distributed in shallow layers that permit oxygen to diffuse into the medium. A test to determine if oxygen is present in the medium may be made by adding a drop of oxidase reagent to the medium. If the medium turns purple, the medium contains oxygen and the nitrate reduction reaction may not occur. If the medium remains colorless, the medium does not contain oxygen and the nitrate reduction test may occur. It has been noted that N. gonorrhoeae cells use up the oxygen rapidly if sufficient cells are inoculated into the medium. If oxidase reagent is added after approximately 1 to 2 hr incubation, the medium will remain clear. Because the oxidase reagent kills the gonococci present in the medium, the medium will gradually turn purple, starting at the top of the tube, as the oxygen diffuses into the medium. If the medium is dispensed in tubes of different dimensions from those suggested above, ensure that the surface area-to-depth ratio is at least equal to or smaller than those suggested above. If the diameter of the tube into which the medium is dispensed is larger than that described above, use a larger volume of medium to maintain the same surface area-to-depth ratio.

The nitrate reduction reaction may not occur if the medium in which the test is performed does not permit normal growth of the organism. However, the test may be performed in a medium that does not support growth of the organisms if the inoculum is sufficiently dense that preformed enzymes can exhaust the existing oxygen supply and reduce the nitrate at a faster rate than that at which the oxygen diffuses into the medium.

Note: To check that the oxygen has been removed from the medium, add 2 to 3 drops of oxidase reagent to a duplicate of the inoculated medium. If the oxygen has been adequately removed from the medium, the oxidase reagent will not turn purple immediately. If the medium contains dissolved oxygen, the oxidase reagent will turn purple. Note also that the nitrate reduction test may be performed in medium to which the oxidase reagent has been added.

When Reagent A is added to the test medium, nitrite produced as a result of the reduction of nitrate will form a complex with the sulfanilic acid that produces a red precipitate with the alpha-naphthylamine in Reagent B. The presence of a red color in the test medium indicates that nitrite is present as a result of the reduction of nitrate. However, the absence of a red color after the addition of Reagents A and B does not necessarily mean that the nitrate was not reduced. The failure to develop a red color may mean (1) that the nitrate was not reduced, or (2) that the nitrite, produced as a result of the reduction of nitrate, has itself been reduced to nitrogenous gases. To determine whether the nitrite has been reduced, place a small quantity of zinc dust in the incubation mixture if it is colorless after the addition of Reagents A and B. The zinc dust catalyzes the reduction of nitrate to nitrite; a red color should develop in the medium which still contains unreduced nitrate. It is important, however, not to add too much zinc dust; excess zinc dust will catalyze the reduction of nitrite produced from that nitrate, resulting in a colorless medium and the incorrect interpretation of the test as positive (a false-positive result).

A positive nitrate reductase test is obtained with the negative control strain, N. gonorrhoeae, after the addition of the zinc dust indicates that the nitrate has been reduced beyond nitrite, probably due to the addition of too much zinc dust to the test. Repeat the test making sure to add very little zinc dust. The pink color, indicating that the organism has not reduced nitrate, may take 10 to 15 min. to develop. Do not add more zinc dust! Wait for the color to develop. If no color has developed in 30 min., interpret the test as positive.

If the test is performed properly and quality control strains give appropriate results, there should be no limitations to this test. Care must be taken to ensure that all components of the test are performed properly .
No identification of genus or species can be made on the basis of the nitrate reduction test alone.
Isolates may be reported as Nitrate-positive if nitrite (pink color) is detected in the inoculated medium after the addition of reagents A and B or if no color is detected in the medium after the addition of zinc dust.

Isolates may be reported as Nitrate-negative if nitrite is not detected (no color change) after the addition of reagents A and B, or if a pink color develops after the addition of zinc dust to the inoculated medium.

Knapp JS, Clark VL. Anaerobic growth of Neisseria gonorrhoeae coupled to nitrite reduction. Infect Immun 1984;46:176-181.

Skerman VBD. 1967. p.218 - 220. A guide to the identification of the genera of bacteria. The Williams & Wilkins Co., Baltimore, MD.

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Among the human Neisseria and related species, three species - N. mucosa, M. catarrhalis, and K. denitrificans reduce nitrate. The nitrate reduction test is a critical test for differentiating between N. gonorrhoeae and , particularly when strains of K. denitrificans appear to be gram-negative diplococci in stained smears.