SGM Journals
PHYSIOLOGY AND BIOCHEMISTRY

l-Glutamine as a nitrogen source for Corynebacterium glutamicum: derepression of the AmtR regulon and implications for nitrogen sensing

Nadine Rehm, Tobias Georgi, Eva Hiery, Ursula Degner, Alfred Schmiedl, Andreas Burkovski, Michael Bott

  • 1Lehrstuhl für Mikrobiologie, Friedrich-Alexander-Universität Erlangen-Nürnberg, D-91058 Erlangen, Germany
  • 2Institut für Biotechnologie 1, Forschungszentrum Jülich, D-52425 Jülich, Germany
  • 3Lehrstuhl für Biochemie, Friedrich-Alexander-Universität Erlangen-Nürnberg, D-91058 Erlangen, Germany
  • Correspondence
    Michael Bott
    m.bott{at}fz-juelich.de

    • Microbiology 2010; 156(10):3180–3193 · https://doi.org/10.1099/mic.0.040667-0

    View at publisher PubMed

    Abstract

    Corynebacterium glutamicum, a Gram-positive soil bacterium employed in the industrial production of various amino acids, is able to use a number of different nitrogen sources, such as ammonium, urea or creatinine. This study shows that l-glutamine serves as an excellent nitrogen source for C. glutamicum and allows similar growth rates in glucose minimal medium to those in ammonium. A transcriptome comparison revealed that the nitrogen starvation response was elicited when glutamine served as the sole nitrogen source, meaning that the target genes of the global nitrogen regulator AmtR were derepressed. Subsequent growth experiments with a variety of mutants defective in nitrogen metabolism showed that glutamate synthase is crucial for glutamine utilization, while a putative glutaminase is dispensable under the experimental conditions used. The gltBD operon encoding the glutamate synthase is a member of the AmtR regulon. The observation that the nitrogen starvation response was elicited at high intracellular l-glutamine levels has implications for nitrogen sensing. In contrast with other Gram-positive and Gram-negative bacteria such as Bacillus subtilis, Salmonella enterica serovar Typhimurium and Klebsiella pneumoniae, a drop in glutamine concentration obviously does not serve as a nitrogen starvation signal in C. glutamicum.

    • ↵†These authors contributed equally to this work.

    • Microarray data and experimental details have been deposited in the Gene Expression Omnibus (GEO) database with the accession number GSE19779.

    Edited by: H.-M. Fischer

    INTRODUCTION

    Corynebacterium glutamicum is a Gram-positive soil bacterium which was isolated by Kinoshita and co-workers in a screening programme for l-glutamate-producing bacteria (Kinoshita et al., 2004; Udaka, 1960) and subsequently used for the industrial production of amino acids. Today, large amounts of l-glutamate (more than 1 524 000 tonnes per year) and l-lysine (more than 812 800 tonnes per year) are produced by using C. glutamicum strains (Hermann, 2003; Leuchtenberger et al., 2005). The genome sequence of this species became available in 2003 (Ikeda & Nakagawa, 2003; Kalinowski et al., 2003) and provided the basis for global analyses such as transcriptomics (Hüser et al., 2003; Polen & Wendisch, 2004; Wendisch, 2003) or proteomics (Bendt et al., 2003; Schaffer et al., 2001) and to establish C. glutamicum as a model organism for high-GC Gram-positive bacteria.

    A focus of our previous studies was a detailed understanding of nitrogen metabolism and its control in C. glutamicum. Ammonium assimilation involves three key reactions (Fig. 1⇓): the reductive amination of 2-oxoglutarate to l-glutamate by glutamate dehydrogenase (GDH), the ATP-dependent conversion of l-glutamate and ammonia to l-glutamine by glutamine synthetase (GS), and the conversion of l-glutamine and 2-oxoglutarate to two molecules of l-glutamate by glutamate : 2-oxoglutarate aminotransferase (also known as glutamate synthase; GOGAT). Inspection of the genome revealed another putative enzyme that might be of importance for the conversion of l-glutamine to l-glutamate, namely glutaminase (GlsK). However, the function of this protein has not yet been analysed experimentally.

    Figure image not available in archive
    Fig. 1.

    Overview on enzymes, genes and reactions relevant for nitrogen assimilation in C. glutamicum. GDH, glutamate dehydrogenase; GOGAT, glutamate : 2-oxoglutarate aminotransferase (also glutamate synthase); GlsK, putative glutaminase; GS, glutamine synthetase.

    The regulation of nitrogen metabolism in C. glutamicum has been intensively studied. Expression of nitrogen-controlled genes is regulated by the TetR-family protein AmtR (Jakoby et al., 2000), which blocks transcription of various genes during growth in nitrogen-rich medium. At least 35 genes are directly controlled by the AmtR repressor (Beckers et al., 2005; Buchinger et al., 2009) and these encode transporters and enzymes for ammonium assimilation (amtA, amtB, glnA, gltBD, dapA), creatinine (codA, crnT) and urea (urtABCDE, ureABCEFGD) metabolism, a number of biochemically uncharacterized enzymes and transport systems, as well as signal transduction proteins (glnD, glnK). In contrast with most TetR-type regulators, the dissociation of AmtR from its target promoters is not triggered by the binding of a low-molecular mass ligand, but by complex formation with the PII-type signal transduction protein GlnK (Beckers et al., 2005). This interaction requires that the trimeric GlnK protein is adenylylated at tyrosine residue 51. Adenylylation is catalysed by the adenylyltransferase GlnD in response to nitrogen deprivation, but it is not yet clear how this is sensed by the enzyme. Global analyses on nitrogen metabolism focused on either AmtR (Buchinger et al., 2009) or the cell's response to ammonium limitation or starvation (Silberbach et al., 2005a, b).

    The work presented here analyses the use of glutamine as a nitrogen source for C. glutamicum. Glutamine is not only a key metabolite for nitrogen assimilation and transferase reactions but also a marker metabolite for nitrogen supply in different Gram-negative and Gram-positive bacteria such as Salmonella enterica serovar Typhimurium (Ikeda et al., 1996), Klebsiella pneumoniae (Schmitz, 2000) and Bacillus subtilis (Hu et al., 1999). In enterobacteria, the NtrBC two-component signal transduction system activates transcription of σ54-dependent promoters in response to nitrogen limitation (Weiss et al., 2002). Phosphorylation of the response regulator NtrC by the NtrB kinase is controlled by the nitrogen regulatory protein PII (GlnB) mainly depending on the cellular glutamine concentration (via uridylylation/deuridylylation of PII by the uridylyltransferase GlnD). In B. subtilis, GlnR, TnrA and GltC (Schreier et al., 1989; Wray et al., 1996, 1998; Picossi et al., 2007) co-operate to regulate nitrogen metabolism (for review, see Fisher, 1999; Sonenshein, 2007; Amon et al., 2010). GlnR is active during growth with excess nitrogen, where it represses the expression of glnRA, ureABC and tnrA (for review, see Sonenshein, 2007; Amon et al., 2010), whereas TnrA is active during nitrogen-limited growth, where it activates and represses the expression of genes involved in the transport and metabolism of nitrogen compounds. The glnA gene encodes GS, which is feedback-inhibited by glutamine. The activities of TnrA and GlnR are controlled through direct protein–protein interactions by feedback-inhibited GS (FBI-GS), which is only present under nitrogen excess. FBI-GS activates the DNA-binding activity of GlnR through a transient association where FBI-GS acts as a chaperone that stabilizes GlnR–DNA complexes (Fisher & Wray, 2008, 2009). In contrast, TnrA is inactivated by FBI-GS through the formation of a stable complex which effectively blocks the DNA-binding ability of TnrA (Wray & Fisher, 2008).

    The results presented in this paper indicate that glutamine is an excellent nitrogen source for C. glutamicum and that its metabolism is dependent on GOGAT. Furthermore, the data suggest that it is unlikely that glutamine serves as an indicator of the cellular nitrogen status.

    METHODS

    Bacterial strains, media and growth conditions.

    All strains and plasmids used in this work are listed in Table 1⇓. The C. glutamicum type strain ATCC 13032 (Abe et al., 1967) was used as wild-type. For growth experiments and DNA microarray experiments, cells from a fresh LB agar plate (Sambrook et al., 2001) were inoculated into 70 ml LB medium (Sambrook et al., 2001) in a 500 ml baffled shake flask and this first preculture was incubated overnight at 30 °C and 120 r.p.m. For the second preculture and the main culture, a modified CGXII medium (Keilhauer et al., 1993) was used which contained 70 mM glucose as a carbon source and either 70 mM l-glutamine or 150 mM ammonium sulfate as a sole nitrogen source. The medium was supplemented with 0.03 g 3,4-dihydroxybenzoate l−1 as iron chelator and 0.2 mg biotin l−1. Before inoculation of the second preculture to OD600 0.1 and of the main cultures to OD600 0.5 (or 1.0 for growth experiments), cells were washed with culture medium without any carbon or nitrogen source. The second preculture was incubated overnight and the main cultures were incubated for the indicated times at 30 °C and 120 r.p.m. For the transcriptome comparison, the main cultures were grown to OD600 4.1±0.2 before cells were harvested and used for RNA preparation. For all cloning purposes, Escherichia coli DH5αMCR (Grant et al., 1990) was used as host and cells were cultivated aerobically at 37 °C.

    Table 1.

    Strains and plasmids used in this study

    Global gene expression analysis.

    Preparation of RNA and synthesis of fluorescently labelled cDNA were carried out as described previously (Lange et al., 2003; Möker et al., 2004). Custom-made whole-genome DNA microarrays for C. glutamicum ATCC 13032 printed with 70-mer oligonucleotides were obtained from Operon (Cologne) and are based on the genome sequence with GenBank accession no. NC_006958 (Kalinowski et al., 2003). Hybridization and stringent washing of the microarrays were performed according to the supplier's instructions. Hybridization was carried out for 16–18 h at 42 °C using a MAUI hybridization system (BioMicro Systems). After washing, the microarrays were dried by centrifugation (5 min, 1600 g) and fluorescence was determined at 532 nm (Cy3-dUTP) and 635 nm (Cy5-dUTP) with 10 μm resolution using an Axon GenePix 6000 laser scanner (Axon Instruments). Quantitative image analysis was carried out using GenePix image analysis software and results were saved as a GPR file (GenePix Pro 6.0, Axon Instruments). For data normalization, GPR files were processed using the BioConductor/R-packages limma and marray (). Processed and normalized data as well as experimental details [these conformed to the MIAME guidelines (Brazma et al., 2001)] were stored in the in-house microarray database for further analysis (Polen & Wendisch, 2004) and in the Gene Expression Omnibus (GEO) repository under the accession number GSE19779.

    Using DNA microarray technology, the mRNA concentrations of C. glutamicum wild-type grown with glutamine as a sole nitrogen source were compared with those of wild-type cells grown with ammonium sulfate as a sole nitrogen source. As outlined above, the strains were cultivated in CGXII minimal medium with 70 mM glucose as carbon source and either 70 mM l-glutamine or 150 mM ammonium sulfate as sole nitrogen source. RNA used for the synthesis of labelled cDNA was prepared from cells in the exponential growth phase at OD600 4.1±0.2. The comparison was performed three times, starting from independent cultures. To filter for differentially expressed genes and reliable signal detection in each of the three comparisons, the following quality filter was applied: (i) flags ≤0 (GenePix Pro 6.0), (ii) signal/noise ≥3 for Cy5 (F635Median/B635Median, GenePix Pro 6.0) or Cy3 (F532Median/B532Median, GenePix Pro 6.0), (iii) ≥twofold change on average in the comparison of wild-type glutamine/wild-type ammonium sulfate, (iv) significant change (P<0.05) by Student's t-test.

    RNA hybridization.

    RNA probes for the analysis of transcription of individual genes were generated by PCR and subsequent labelling with DIG RNA-labelling mix and T7 RNA polymerase (Roche Diagnostics). The oligonucleotides used for the gdh probe were gdh-for (5′-CCCGGGTGACATCGGAGTTGGTGG) and gdh-rev (5′-GGGCCCTAATACGACTCACTATAGGGGAGCTTAGCCACGTCAAC), for the glnA probe, glnA-for (5′- CCAAACCGTTGACGTGCG) and glnA-rev (5′- GGGCCCTAATACGACTCACTATAGGGGGATCCGGTGATTGGGATAC), for the gltB probe, gltB-for (5′-CCACTGGTGTGCTGAAGGTGATGTCC) and gltB-rev (5′-GGGCCCTAATACGACTCACTATAGGGGAAACGCTTC), and for the gltD probe, gltD-for (5′-GGGCCCCCGCGAAGGCTGGGTGCAACC) and gltD-rev (5′-GGGCCCTAATACGACTCACTATAGGGGTGAGGTAAT). The underlined sequences represent the T7 promoter sequence. All oligonucleotides were used at a final concentration of 0.5 μM. Chromosomal DNA as template and the following PCR programme were used: 3 min at 94 °C, 30 cycles of 15 s at 94 °C followed by 15 s at 60 °C and 1 min at 72 °C, then 10 min at 72 °C followed by cooling to 4 °C. The RNA isolated from cells was spotted directly onto nylon membranes using a Schleicher & Schuell Minifold I dot blotter. Hybridization of DIG-labelled RNA probes was detected with Kodak X-OMAT X-ray films using alkaline phosphatase-conjugated anti-DIG Fab fragments and CSPD as a light-emitting substrate as recommended by the supplier (Roche Diagnostics).

    General molecular biology techniques.

    For plasmid isolation, transformation and cloning, standard techniques were used (Ausubel et al., 1987; Sambrook et al., 2001). E. coli strain DH5αMCR was used as the cloning host. Competent C. glutamicum cells were prepared according to the method described by van der Rest et al. (1999). Chromosomal C. glutamicum DNA was isolated as described by Eikmanns et al. (1994). DNA sequence analyses were carried out using an ABI sequencer and the freely available software Chromas Lite.

    Construction of deletion mutants.

    Unmarked deletions were introduced into the genome of C. glutamicum according to the protocol described by Schäfer et al. (1994) and Niebisch & Bott (2001). All deletions were verified by PCR and the flanking regions in the genome were sequenced as a control (data not shown). For construction of the glsK deletion plasmid, 0.7 kb DNA fragments covering the upstream and the downstream region of glsK were amplified and fused by overlap–extension PCR using chromosomal DNA from ATCC 13032 as a template. The oligonucleotide primers were designed in such a way that an XbaI restriction site (flanking the upstream fragment) or an SphI restriction site (flanking the downstream fragment) were introduced (5′-CGCTCTAGAGCGAATCCACCGAAATCGTCTTCATCT-3′, 5′-GCCCATAAAGCTACCGGAGAATCAAGACTAGTGAT-3′, 5′-ATCACTAGTCTTGATTCTCCGGTAGCTTTATGGGC-3′, 5′-CGCGCATGCTTTAAGCTGCGAGGCCTTCTTCGAGTTT-3′, restriction sites shown in bold). The resulting 1.4 kb DNA fragment was ligated to XbaI/SphI-restricted vector pK19mobsacB, leading to plasmid pK19mobsacB-ΔglsK. This plasmid was then applied to introduce a chromosomal deletion of glsK as described previously (Schäfer et al., 1994) into strains ATCC 13032, LNΔgdh and LNΔgltBD leading to strains NR-2, NR-3 and NR-4, respectively. In order to generate a chromosomal gdh deletion in the C. glutamicum mutant strains LNΔgltBD and NR-4, plasmid pK18Δgdh was used (Müller et al., 2006).

    Construction of a gltBD overexpression plasmid.

    A plasmid for overexpression of gltBD, designated pEKEx-gltBD, was generated as follows. First, a 2.7 kb fragment of the 5′-region of gltB (gltB′) was amplified by PCR using chromosomal DNA as template and the oligonucleotides 5′-GGGCCCGGATCCATGAAACCACAAGGACTCTAC-3′ and 5′-GGGCCCGGTACCGGCTTCAGCAGAAATCGAGCC-3′. The gltB′ fragment was ligated into pEKEx2 using the BamHI and Acc65I sites introduced by the oligonucleotides (shown in bold), leading to the plasmid pEKEx-gltB′. The second fragment of 3.5 kb, harbouring the 3′-region of gltB and the entire gltD gene (gltBD′′), was amplified by PCR using chromosomal DNA as template and the oligonucleotides 5′-GGGCCCCGGACCGCAAGCCAATTTCGGTG-3′ and 5′-GGGCCCGGTACCCTAGACAGCCAGCGGCACGTC-3′. The gltBD′′ fragment was cloned into pEKEx-gltB′ using the RsrII restriction site, which naturally occurs in the downstream region of gltB, and the Acc65I restriction site, introduced by PCR (shown in bold). The resulting plasmid, pEKEx-gltBD, was sequenced as a control.

    Enzyme activity measurements and protein determination.

    GS activity was determined by using the protocol described by Shapiro & Stadtman (1970). The protein content of cell extracts was determined using a Bradford assay (Roth).

    Western blot analyses.

    For the preparation of C. glutamicum cell extracts, 2 ml aliquots were transferred from the culture to tubes containing 300 mg glass beads (0.25 mm, Sigma) and immediately frozen in liquid nitrogen. Samples thawed on ice were disrupted using a FastPrep FP120 instrument (Q-BIOgene). Subsequently, glass beads and cell debris were removed by centrifugation. Protein concentrations were determined as described above. SDS-PAGE was carried out using 10 % acrylamide gels and a Tricine buffer system as described by Schägger & von Jagow (1987). After electrophoresis, the gel-separated proteins were transferred onto a PVDF membrane by electroblotting and incubated with a rabbit polyclonal antiserum against C. glutamicum GS. Antibody binding was visualized by appropriate secondary antibodies coupled to alkaline phosphatase and the BCIP/NBT alkaline phosphatase substrate (Sigma-Aldrich).

    Measurement of intracellular l-glutamine concentrations.

    Cells were grown as described above in minimal medium and 2–4 ml culture was harvested in the exponential growth phase by centrifugation (2 min, 8000 g, 25 °C). Cell pellets were washed once with 4 ml PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, adjusted with HCl to pH 7.4). Metabolites were extracted from the cell pellets by adding 4 ml 80 % (v/v) ethanol, previously heated to 80 °C, to the cell pellets. The extracts were then incubated under shaking for 30 min at 80 °C. After centrifugation, the supernatant containing the extracted metabolites was kept and the disrupted cells were again treated with ethanol as described above. The second supernatant was mixed with the first and an appropriate amount was dried using a vacuum centrifuge. Metabolites were dissolved in double distilled water (1 ml) and the solution was filtered through a 10 kDa exclusion membrane in order to remove proteins and other small cell particles. The samples were then used for metabolite quantification by liquid chromatography coupled to mass spectrometry. The LC-MS system used consisted of a liquid chromatograph DIONEX ICS-3000 equipped with an Autosampler AS-50, a column ACCLAIM-OA 5 μm, 4×250 mm, and a precolumn ACCLAIM-OA 5 μm, 4.3×10 mm (all from Dionex). The chromatograph was operated by the software Chromatography MS Link 2.0.0.2315 (DC/MS-Link 2.0; Dionex) which is a plug-in module of the MS software analyst 1.4.2 (Applied Biosystems). MDS SCIEX TurboV (MDS Analytical Technologies) was used as an ion source. Samples were kept at 8 °C in the autosampler before injection. The system was equilibrated with buffer (2.5 mM ammonium acetate, pH 3.5). A flow rate of 0.25 ml min−1 and a column temperature of 30 °C were used. The run time of each sample took about 50 min. A gradient of 50 mM ammonium acetate, rising from 5 to 30 % (v/v), then decreasing to 5 % (v/v) again, was used, followed by increasing the acetonitrile content from 0 to 90 % (v/v) to elute hydrophobic amino acids and wash the column. Glutamine eluted at a retention time of 9.8 min. Analysis was carried out by a mass spectrometer 3200 QTRAP (Applied Biosystems) using a multiple reaction monitoring mode (MRM) with a dwell time of 50 ms per MRM. The detection parameters for glutamine were as follows: ion-source-specific parameters CUR, 10 p.s.i.; IS, 5500 V; GS1 , 40 p.s.i.; GS2 ,  45 p.s.i.; TEM, 500 °C. The compound-specific parameters in MRM mode were as follows: MRM transition, 147 to 130; DP, 23.5 V; EP, 5.5 V; CEP, 14 V; CE, 14 V; CXP, 2.5 V. The data obtained were analysed by the analyst 1.4.2 software. Internal glutamine concentrations of C. glutamicum cells were then calculated from the data using the following correlations: OD600=1 corresponds to 0.3 mg cell dry weight ml−1 and 1 mg cell dry weight corresponds to a cytoplasmic volume of 1.6 μl (Botzenhardt et al., 2004).

    RESULTS

    Growth of C. glutamicum on glutamine as the carbon and/or nitrogen source

    In order to determine the growth parameters of C. glutamicum with l-glutamine as the sole nitrogen and/or carbon source, wild-type cells were cultivated in CGXII minimal medium containing different carbon and nitrogen sources. As shown in Fig. 2⇓, C. glutamicum wild-type ATCC 13032 was able to grow with 70 mM l-glutamine as the sole carbon and nitrogen source to a final OD600 ∼10±1.2, but with a very slow growth rate (μ) between 0.02 and 0.03 h−1. The additional presence of 150 mM ammonium sulfate had no influence on glutamine-dependent growth. In contrast, cells cultivated with 70 mM glucose and 70 mM l-glutamine as the sole nitrogen source showed a similar growth rate (μ=0.34±0.01 h−1) to cells grown with 70 mM glucose and 150 mM ammonium sulfate as the sole nitrogen source (μ=0.33±0.01 h−1). The final OD600 was 28.7±3.8 for growth on glucose and l-glutamine and 18.4±0.9 for growth on glucose and ammonium, indicating that l-glutamine is incorporated into biomass to the same extent as during growth on l-glutamine as the sole carbon and nitrogen source. In summary, these data show that l-glutamine is metabolized slowly as a carbon source but serves as an excellent nitrogen source, allowing the same growth rates as allowed by ammonium.

    Figure image not available in archive
    Fig. 2.

    Growth of C. glutamicum wild-type in carbon- and nitrogen-free CGXII medium supplemented with 70 mM glucose and 150 mM ammonium sulfate (▪), 70 mM glucose and 70 mM l-glutamine (□), 70 mM glutamine and 150 mM ammonium sulfate (▴), or 70 mM glutamine (▵). Representative results of at least three independent experiments are shown.

    Comparison of global gene expression profiles of C. glutamicum grown on l-glutamine or ammonium as a sole nitrogen source using DNA microarrays

    To analyse how l-glutamine influences global gene expression when used as a sole nitrogen source instead of ammonium, DNA microarray analyses were performed. For this purpose, RNA was isolated from exponentially growing cells (OD600 4.1±0.2) cultivated in CGXII medium containing glucose as the carbon source and either l-glutamine (70 mM) or ammonium sulfate (150 mM) as the nitrogen source. As shown above, the growth rates of C. glutamicum in these two media are very similar and therefore growth-rate-dependent effects on transcription were expected to be negligible. In fact, transcription of genes whose expression is typically growth-rate-dependent, such as the atpABCDEFGH operon or ribosomal genes, was not altered. Overall, 65 genes showed significantly (P<0.05) changed mRNA levels by a factor of two or more (average of three biological replicates). These genes can be subdivided into three groups (Table 2⇓).

    Table 2.

    Genes whose mean mRNA ratio was altered ≥twofold when C. glutamicum was grown in glucose minimal medium with l-glutamine instead of ammonium as the sole nitrogen source

    The ratios represent mean values from three DNA microarray experiments starting from independent cultures.

    Group A comprises 26 genes which belong to the nitrogen starvation stimulon of C. glutamicum and are regulated by the transcriptional master regulator of nitrogen metabolism, AmtR (Jakoby et al., 2000). All of these genes showed increased transcript abundance on l-glutamine as the nitrogen source (Table 2⇑). Among these genes were glnD and glnK, encoding key proteins for nitrogen signal transduction (Meier-Wagner et al., 2001), which showed 21- and 23-fold increased mRNA levels, respectively. The glnA gene, which codes for GS, was 10-fold upregulated. The genes of the gltBD operon, encoding the large and the small subunit of GOGAT (Beckers et al., 2001), showed 93- and 231-fold increased mRNA levels, respectively, and the genes for ammonium uptake, amtA and amtB, displayed 1826- and 141-fold increased mRNA levels, respectively. Furthermore, expression of several operons for uptake und utilization of alternative nitrogen sources was upregulated in cells growing with l-glutamine as a sole nitrogen source. These were the creatinine uptake and utilization genes cg0103 and cg0104 (664- and 39-fold increased mRNA levels, respectively) (Bendt et al., 2004), the urease operon ureABCEFGD (Nolden et al., 2000), whose genes were at least 18-fold upregulated, the urea uptake system urtABCDE (Beckers et al., 2004), whose expression was at least 28-fold increased, and soxA (143-fold) and ocd (750-fold), which are supposed to be involved in the degradation of sarcosine and ornithine and are found in an operon together with amtA (Jakoby et al., 1999). The gene cg1295 showed a 14-fold increased mRNA level and is the only gene of this group whose function is presently unknown. Together, 24 known AmtR-regulated genes in C. glutamicum were strongly induced by at least a factor of 9.5 when cultivated with l-glutamine instead of ammonium as the sole nitrogen source. The five remaining genes of the AmtR regulon observed here either were not differentially expressed by a factor of at least two (cg1646, gluD, gdh and mez) or, in the case of cg1296, showed a strongly increased mRNA level, but exhibited too much variation to meet the P-value cut-off. The data clearly show that growth of C. glutamicum with l-glutamine as the sole nitrogen source causes derepression of the AmtR-regulated genes (Beckers et al., 2005; Jakoby et al., 2000).

    Group B (Table 2⇑) comprises 14 genes, which were previously shown to be repressed under excess iron conditions by DtxR, the master regulator of iron homeostasis in C. glutamicum (Brune et al., 2006; Wennerhold & Bott, 2006). The mRNA level of these genes was lower by a factor of between two and four in cells grown with l-glutamine as the nitrogen source compared with cells grown with ammonium. Twelve of the 14 genes are involved in binding or uptake of iron: cg0589, cg0590, cg0591, cg0770, cg0926 and cg0927 encode components of siderophore ABC transporters; cg0922, cg0924, cg1418 and cg3404 code for secreted siderophore-binding lipoproteins; cg0466 encodes a haem transport-associated protein; and cg0921 encodes a cytoplasmic siderophore-interacting protein (Frunzke & Bott, 2008; Wennerhold & Bott, 2006). Besides the genes involved in iron uptake, the mRNA level of the ripA gene was decreased by a factor of three. The AraC-type regulator RipA represses a set of genes encoding prominent iron proteins, such as aconitase or succinate dehydrogenase, under iron starvation and in this way reduces the iron demand of the cell (Wennerhold et al., 2005). The function of the protein encoded by cg0160, whose mRNA level was also twofold lower during growth on l-glutamine as the nitrogen source, is unknown. The above results indicate that the iron starvation response is turned on earlier during growth with ammonium than during growth with l-glutamine as a nitrogen source.

    Recently, evidence was provided that the iron starvation response is induced by acidic pH (Follmann et al., 2009). Therefore, we followed the pH during growth of C. glutamicum with glutamine or ammonium as the nitrogen source. Indeed, a difference in the pH profiles was observed. With glutamine, a slight increase in the pH from 7.0 to 7.4 was observed within 33 h of cultivation, whereas with ammonium, the pH dropped from 7.0 to 6.1 in the same period. At the time when the cells were harvested for RNA isolation (∼5 h after the start of cultivation), there was a pH difference of about 0.10–0.15 between the cultures (data not shown), which might at least partially be responsible for the differences in the expression of the DtxR target genes.

    Group C (Table 2⇑) comprises a functionally diverse set of 25 genes whose mRNA levels were changed at least twofold and which do not belong to the AmtR or DtxR regulons. It includes genes involved in carbon metabolism, transport, transcriptional regulation and other functions. Three genes involved in the utilization of ethanol showed three- to fourfold lower mRNA levels with l-glutamine as the nitrogen source, i.e. those encoding alcohol dehydrogenase (adhA), acetaldehyde dehydrogenase (cg3096) and isocitrate lyase (aceA). On the other hand, the mRNA level of the gluconeogenic pck gene encoding phosphoenolpyruvate carboxykinase was twofold increased in cells grown with l-glutamine. For most of the genes summarized in group C, an obvious relationship to the utilization of l-glutamine as a nitrogen source is missing (see also Discussion).

    Growth of mutant strains with ammonium as the nitrogen source

    The microarray data suggest that one or several AmtR-controlled genes have a role in the utilization of glutamine as the nitrogen source. This assumption was investigated further by mutant analyses. C. glutamicum has two primary pathways for the synthesis of l-glutamate and l-glutamine (Fig. 1⇑), which are the major nitrogen donors for biosynthetic reactions (Jakoby et al., 1997; Shiio & Ozaki, 1970; Tesch et al., 1998). GDH assimilates ammonium by the reductive amination of 2-oxoglutarate to l-glutamate. Alternatively, ammonium is assimilated via the GS/GOGAT pathway. GS uses ammonium to amidate l-glutamate under consumption of ATP to form l-glutamine. The amide group is then transferred reductively to 2-oxoglutarate by GOGAT, resulting in the net conversion of 2-oxoglutarate to l-glutamate. l-Glutamine might be metabolized by a presently uncharacterized glutaminase, encoded by the C. glutamicum glsK gene (cg2728), to l-glutamate and ammonia. In order to characterize the function of these different enzymes in glutamine metabolism, a series of mutant strains was constructed (details in Table 1⇑) and tested first for growth on ammonium as the nitrogen source.

    During growth in CGXII minimal medium with glucose as the carbon source and ammonium sulfate as the nitrogen source, the wild-type had a doubling time of 115±4 min. Growth was unaltered in the ΔglsK mutant NR-2 (112±6 min), the ΔgltBD mutant LNΔgltBD (113±7 min) and the ΔgltBDΔglsK double mutant NR-4 (111±6 min). In contrast, gdh deletions significantly affected growth with ammonium. The Δgdh mutant LNΔgdh showed a doubling time of 193±23 min, which was essentially unaltered in combination with a glsK mutation in the ΔgdhΔglsK double deletion strain NR-3 (175±16 min). In combination with a gltBD deletion, the negative impact of a gdh deletion on growth was even more pronounced. For the ΔgltBDΔgdh mutant NR-5, a doubling time of 229±39 min was observed; for the ΔgltBDΔgdhΔglsK triple mutant NR-6 this was 286±48 min. The results are summarized in Table 3⇓.

    Table 3.

    Growth parameters of C. glutamicum strains cultivated with different nitrogen and carbon sources

    td, Doubling time; values are mean±sd. –, Not determined.

    As shown recently (Müller et al., 2006), a gdh mutation elicits a partially deregulated nitrogen starvation response in C. glutamicum, as shown by partial adenylylation of GlnK and the increased transcription of glnA and gltBD. Ammonium is then assimilated exclusively via the GS/GOGAT pathway under an extra consumption of 1 mol ATP per mol ammonium assimilated, which might be the reason for the impaired growth of strain LNΔgdh.

    The ΔgltBDΔgdh double mutant showed a strongly decreased growth rate. As this strain can still assimilate ammonium via GS, we assume that glutamate synthesis is the main factor that contributes to decreased growth. In the absence of GDH and GOGAT, the synthesis of glutamate can be achieved by transamination of 2-oxoglutarate using another amino acid as the amino donor. However, this route requires an alternative ammonium assimilation reaction. One possibility for such a reaction is aspartate ammonia lyase (aspartase), which catalyses the reversible conversion of fumarate and ammonia to aspartate. This enzyme is used in industry for the production of aspartate. Aspartate can then serve as an amino donor for the transamination of 2-oxoglutarate to glutamate, presumably by the transaminase AspT (Marienhagen et al., 2005). C. glutamicum possesses an aspartate ammonia lyase encoded by cg1697 and thus could make use of it for ammonia assimilation in a ΔgltBDΔgdh double mutant. Another reaction involved in ammonia assimilation is the one catalysed by diaminopimelate dehydrogenase. This enzyme is involved in l-lysine biosynthesis and catalyses the reductive amination of piperideine-2,6-dicarboxylate to dl-diaminopimelate, which is subsequently decarboxylated to l-lysine. However, in contrast with l-aspartate it is unknown whether dl-diaminopimelate can serve as an amino donor in a transaminase reaction. Furthermore, there are many genes of unknown function present in the genome of C. glutamicum, which might provide a further route to glutamate synthesis together with one of the different transaminase reactions, which have only been partially characterized to date (Marienhagen et al., 2005; Marienhagen & Eggeling, 2008).

    The triggering of the nitrogen starvation response in the different strains was verified on the level of gene expression, GS content and GS activity (Fig. 3a–c⇓). In short, glnA and gltBD expression, GS protein level and GS activity were increased in all strains carrying a gdh deletion, confirming the results obtained by Müller et al. (2006) which showed that a partial nitrogen starvation response is elicited by a lack of GDH.

    Figure image not available in archive
    Fig. 3.

    Expression analyses, GS protein levels and GS activities of mutant strains grown with ammonium (a–c) or glutamine (d–f) as the nitrogen source. (a, d) RNA hybridization using probes against gdh, glnA, gltB and gltD. Total RNA was prepared from the wild-type (bar 1) and from mutant strains NR-2 (ΔglsK; bar 2), LNΔgdh (Δgdh; 3), NR-3 (ΔgdhΔglsK; 4), LNΔgltBD (ΔgltBD; 5), NR-4 (ΔgltBDΔglsK; 6), NR-5 (ΔgltBDΔgdh; 7) and NR-6 (ΔgltBDΔgdhΔglsK; 8). Two micrograms total RNA per sample was spotted. (b, e) GS protein levels. Cell extracts (10 μg protein per lane) were separated by using SDS-PAGE and were visualized by Western blotting with anti-GS antiserum. (c, f) Specific GS activity. The average activity of three independent measurements is shown for each strain. Error bars, sd.

    Growth of mutant strains with l-glutamine as the nitrogen source

    As shown by the transcriptome analyses presented above, growth with l-glutamine as the sole nitrogen source elicited the nitrogen starvation response in C. glutamicum wild-type. The molecular basis of this phenomenon was investigated by growth experiments with the mutants described above (Table 3⇑). With a doubling time of 130±8 min, growth of the wild-type with l-glutamine was almost as fast as with ammonium. Also, the ΔglsK mutant NR-2 (126±2 min), the Δgdh mutant LNΔgdh (125±12 min) and the ΔgdhΔglsK double deletion strain NR-3 (127±13 min) reached wild-type growth rates. In contrast, growth of all gltBD mutants was significantly impaired. The ΔgltBD mutant LNΔgltBD had a doubling time of 303±51 min, the ΔgltBDΔglsK double mutant NR-4 of 376±122 min, the ΔgltBDΔgdh strain NR-5 showed a doubling time of 468±8 min, and for the ΔgltBDΔgdhΔglsK triple mutant NR-6 a doubling time of 474±58 min was observed. These results indicate that GOGAT is the main glutamine-metabolizing enzyme in C. glutamicum and provide an explanation for the derepression of the AmtR-regulated genes, since the GOGAT-encoding gltBD operon is under strict AmtR control in C. glutamicum (Beckers et al., 2001).

    The triggering of the nitrogen starvation response in the different mutants was tested again using RNA hybridization experiments, Western blotting and GS activity measurements (Fig. 3d–f⇑). As expected from the DNA microarray data, the mRNA levels of glnA, gltB, and gltD were high, as were the GS protein levels. GS activity was high in most strains [2–3 U (mg protein)−1] but was significantly lower in strains NR-4 (ΔgltBDΔglsK) and NR-6 (ΔgdhΔgltBDΔglsK). Since the GS protein level was also high in these strains, the reduced activity seems to be due to activity regulation by adenylylation via the adenylyltransferase GlnE (Nolden et al., 2001a).

    Effect of plasmid-encoded gltBD

    In light of the idea that GOGAT is primarily responsible for glutamine metabolism, it should be possible to rescue a chromosomal gltBD deletion by plasmid-encoded gltBD. For this complementation assay, growth of the wild-type and the ΔgltBD mutant carrying either pEKEx2 or pEKEx2-gltBD was tested in the presence of 1 mM IPTG. In fact, growth of the ΔgltBD mutant in CGXII medium with glucose as the carbon source and l-glutamine as the nitrogen source was improved when it carried the gltBD expression plasmid, but it did not reach wild-type levels (Fig. 4⇓). Doubling times observed were 108 min for the wild-type, 230 min for strain LNΔgltBD, 248 min for strain LNΔgltBD carrying the control plasmid pEKEx2 and 164 min for strain LNΔgltBD carrying pEKEx-gltBD. This supports the proposed function of GOGAT in l-glutamine metabolism. The reason for the observed incomplete complementation of strain LNΔgltBD by the multi-copy plasmid pEKEx-gltBD remains unclear. Transcription of plasmid-borne and chromosomally expressed gltB and gltD was tested by RNA hybridization experiments and similar transcript levels were found (data not shown).

    Figure image not available in archive
    Fig. 4.

    Growth complementation of C. glutamicum LNΔgltBD. The C. glutamicum wild-type (▪) and strains LNΔgltBD (□), LNΔgltBD/pEKEx2 (▴) and LNΔgltBD/pEKEx-gltBD (▵) were grown in nitrogen-free CGXII glucose minimal medium supplemented with 70 mM l-glutamine and 1 mM IPTG. Representative results of five independent experiments are shown.

    Glutamine as a sensor molecule for nitrogen control

    As described above, GOGAT is important for efficient utilization of glutamine as a nitrogen source. Since the corresponding gltBD operon is under strict nitrogen regulation, derepression of transcription is a prerequisite for fast growth with glutamine as a sole nitrogen source. Previous work showed that growth on glutamine as a sole carbon and nitrogen source results in relatively high intracellular glutamine levels in the range of 100 mM (Niebisch et al., 2006), which indicates that C. glutamicum does not sense nitrogen starvation as a drop in glutamine concentration. This assumption was further investigated here.

    When l-glutamine was measured in the wild-type grown with ammonium as the nitrogen source, a cytoplasmic glutamine concentration of 15±1 mM was determined, which is in the range of previously published data (Müller et al., 2006; Nolden et al., 2001b). About twofold higher concentrations were found when the cells were grown with 70 mM l-glutamine as the nitrogen source with a cytoplasmic l-glutamine concentration of 31±7 mM for the wild-type. These data clearly indicate that a drop in the cytoplasmic glutamine concentration does not serve as a signal for nitrogen starvation in C. glutamicum. When both ammonium and glutamine were used as nitrogen sources, an intracellular concentration of 35±5 mM l-glutamine was determined, indicating that the presence of ammonium did not inhibit glutamine uptake. It was previously suggested that ammonium might be a sensor metabolite for the nitrogen status of the cell (Müller et al., 2006; Nolden et al., 2001b). In fact, when an ammonium pulse was given to glutamine-grown cells with derepressed AmtR target genes, gltB expression ceased within minutes in these cells (Fig. 5⇓), making it even more likely that ammonium is an effector molecule.

    Figure image not available in archive
    Fig. 5.

    Suppression of the nitrogen starvation response by ammonia as tested by RNA hybridization using a gltB probe. Two micrograms total RNA per sample was spotted. C. glutamicum wild-type was grown in minimal medium with 70 mM glucose, 150 mM ammonium sulfate and 70 mM l-glutamine (a), or with 70 mM glucose and 70 mM glutamine (b), and samples for RNA isolation were taken in the early exponential growth phase at OD600 ∼4. A pulse of 150 mM ammonium sulfate was given to culture (b) and samples for RNA isolation were taken from this immediately after ammonium addition (c), or after 5 (d), 15 (e), 30 (f), 60 (g) or 120 (h) min.

    DISCUSSION

    In this work, we showed that l-glutamine serves as a good nitrogen source for C. glutamicum and allows similar growth rates to ammonium. DNA microarray analysis revealed that the target genes of the master regulator of nitrogen metabolism, AmtR, are derepressed in cells cultivated with l-glutamine as a sole nitrogen source (Table 2⇑). As discussed in more detail below, this response is due to the requirement of GOGAT for glutamine utilization, whose structural genes gltBD are repressed by AmtR. Besides the members of the AmtR regulon, more than a dozen genes of the DtxR regulon showed a lower mRNA level in glutamine-grown cells compared with ammonium-grown cells (Table 2⇑). DtxR directly senses the intracellular concentration of Fe2+ and functions as a master regulator of iron homeostasis in C. glutamicum: it represses or activates more than 60 genes under iron-sufficient conditions (Brune et al., 2006; Frunzke & Bott, 2008; Wennerhold & Bott, 2006). Under iron limitation, bound Fe2+ dissociates from DtxR and the protein dissociates from DNA, causing derepression or deactivation of its target genes. The majority of the repressed genes encode iron uptake systems, but some also encode transcriptional regulators, such as RipA, which is responsible for reducing the cellular iron demand under iron limitation (Wennerhold et al., 2005). A possible explanation for the higher mRNA level of DtxR target genes in ammonium-grown cells could be the lower pH (0.1–0.15 pH units) of these cultures compared with the glutamine-grown cultures at the time when the cells were harvested for RNA isolation. It was recently shown that the iron starvation response is induced by acidic pH (Follmann et al., 2009).

    Besides the targets of AmtR and DtxR, 25 other genes of highly diverse functions also showed more than twofold-altered mRNA levels in glutamine-grown cells (Table 2⇑, group C). For most of these genes, such as cg1832, cg1833, cg3138, cg3139 and cg3140, we do not have an explanation for their differential expression. The 0.3- to 0.4-fold decreased expression of the genes encoding alcohol dehydrogenase (adhA), aldehyde dehydrogenase (cg3096) and isocitrate lyase (aceA) in glutamine-grown cells might be due to an altered activity of the transcriptional regulator RamA, which is known to positively regulate these genes (Arndt & Eikmanns, 2007; Auchter et al., 2009; Cramer et al., 2006). The gene pyrG (cg1606) presumably encodes CTP synthetase (EC 6.3.4.2), an enzyme catalysing the ATP-dependent amination of UTP to CTP with either ammonia or l-glutamine as the source of nitrogen. Its sevenfold increased expression in glutamine-grown cells might indicate that its activity with glutamine is lower than with ammonia. The pyrG gene could be a novel member of the AmtR regulon, as its expression was also threefold increased in a ΔamtR mutant compared with the wild-type (Buchinger et al., 2009).

    Analysis of mutant strains lacking the genes for a putative GlsK (glsK), GDH (gdh) or GOGAT (gltBD), either alone or in combination, revealed that the latter enzyme is crucial for glutamine metabolism as its absence leads to an almost threefold increase in the doubling time. The fact that the gltBD operon is part of the AmtR regulon (Beckers et al., 2001, 2005; Schulz et al., 2001) provides an explanation for the triggering of the nitrogen starvation response. GDH and GlsK are not relevant for glutamine utilization as long as GOGAT is present, as the doubling times of the Δgdh and ΔglsK mutants were similar to the wild-type. In the absence of GOGAT, however, both GlsK and GDH appear to contribute to glutamine metabolism, as indicated by the significantly prolonged doubling times of the ΔgltBDΔglsK and ΔgltBDΔgdh double mutants compared with the ΔgltBD single mutant (Table 3⇑). In order to better understand the role of the GlsK protein, a biochemical characterization of its properties will be required. In other bacteria such as Rhizobium etli (Durán et al., 1995, 1996), GlsK was shown to have a significant role in the metabolism of l-glutamine, especially when glutamine is provided as a carbon source.

    A prerequisite for the utilization of l-glutamine is its transport into the cell. C. glutamicum was previously reported to possess a secondary Na+-dependent glutamine uptake system which uses both the membrane potential and the sodium ion gradient as driving forces (Siewe et al., 1995). Glutamine uptake showed Michaelis–Menten kinetics, with a Km of 36 μM and a Vmax of 12.5 nmol min−1 (mg dry weight)−1 at pH 7. The Km value for the co-transported sodium ion was reported to be 1.4 mM. The gene(s) encoding this glutamine uptake system has not been identified yet. In the transcriptome comparison performed in our work, none of the genes with an increased mRNA level during growth with l-glutamine as the nitrogen source was annotated as a putative transporter for this or a related amino acid. This might suggest that expression of the gene encoding the secondary l-glutamine transporter is constitutive and not induced by glutamine. In E. coli, an ABC transport system encoded by the glnHPQ operon is responsible for glutamine uptake (Nohno et al., 1986). C. glutamicum contains a gene that has been annotated as glnH (cg3045) and may encode a glutamine-binding lipoprotein. However, glnH is not associated with genes characteristic for ABC transporters, but with a gene encoding a membrane protein of unknown function (glnX, cg3044) and the serine/threonine protein kinase gene pknG (Niebisch et al., 2006). Interestingly, mutants lacking glnX, glnH or pknG showed a growth defect on agar plates containing l-glutamine as the sole carbon and nitrogen source. Evidence was provided that this phenotype is not caused by a defect in glutamine uptake, but by an inhibition of the 2-oxoglutarate dehydrogenase complex (Bott, 2007; Niebisch et al., 2006).

    Growth on glutamine obviously led to a change in the concentration of a metabolite indicating the cellular nitrogen status. In enterobacteria, nitrogen supply in the cells is perceived by measuring two metabolites of the central nitrogen cycle. The NtrBC two-component signal transduction system activates transcription of σ54-dependent promoters in response to nitrogen limitation (Weiss et al., 2002). Phosphorylation of the response regulator NtrC by the NtrB kinase is controlled by the nitrogen regulatory protein PII (GlnB) depending on the cellular glutamine concentration (via uridylylation/deuridylylation of PII by the uridylyltransferase GlnD; Zhang et al., 2010) and on the cellular 2-oxoglutarate concentration as well as the adenylate energy charge (Jiang & Ninfa, 2009a, b). At low glutamine and high 2-oxoglutarate concentrations, the activation of the phosphatase activity of NtrB by PII is inhibited and the inhibition of the autophosphorylation activity of NtrB by PII is relieved. As a consequence, NtrC is phosphorylated and NtrBC-regulated genes are expressed.

    In C. glutamicum, high 2-oxoglutarate concentrations favour modification of the PII-type protein GlnK and transcription of AmtR-regulated genes (Müller et al., 2006). However, these results obtained with a gdh deletion strain also suggest that there is a second marker metabolite, since only a partially deregulated nitrogen control was observed, as indicated by a partially, not fully, adenylylated GlnK protein. Due to its central position in nitrogen metabolism and the situation in enterobacteria and B. subtilis, l-glutamine would be a possible candidate for such a marker metabolite. However, as the AmtR regulon was derepressed in l-glutamine-grown cells although the intracellular l-glutamine concentration was higher than in ammonium-grown cells, l-glutamine is obviously not involved in nitrogen sensing in C. glutamicum.

    The data obtained here are in accordance with previous suggestions, which favour ammonium instead of glutamine as the sensor of the cellular nitrogen supply (Müller et al., 2006; Nolden et al., 2001b). Further work is necessary to strengthen this model.

    Acknowledgments

    The authors wish to thank Lars Nolden (Cologne) for providing strain LNΔgltBD, Julia Frunzke (Jülich) for testing the pH profiles and Tino Polen (Jülich) for deposition of the microarray data in the GEO database. This work was supported by the Bundesministerium für Bildung und Forschung (BMBF) within the GenoMik-Plus programme.

    References

    1. Abe, S., Takayama, K. & Kinoshita, S. (1967). Taxonomical studies on glutamic acid producing bacteria. J Gen Appl Microbiol 13, 279–301.
    2. Amon, J., Titgemeyer, F. & Burkovski, A. (2010). Common patterns – unique features: nitrogen metabolism and regulation in Gram-positive bacteria. FEMS Microbiol Rev 34, 588–605.
    3. Arndt, A. & Eikmanns, B. J. (2007). The alcohol dehydrogenase gene adhA in Corynebacterium glutamicum is subject to carbon catabolite repression. J Bacteriol 189, 7408–7416.
    4. Auchter, M., Arndt, A. & Eikmanns, B. J. (2009). Dual transcriptional control of the acetaldehyde dehydrogenase gene ald of Corynebacterium glutamicum by RamA and RamB. J Biotechnol 140, 84–91.
    5. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A. & Struhl, K. (editors) (1987). Current Protocols in Molecular Biology. New York: Wiley.
    6. Beckers, G., Nolden, L. & Burkovski, A. (2001). Glutamate synthase of Corynebacterium glutamicum is not essential for glutamate synthesis and is regulated by the nitrogen status. Microbiology 147, 2961–2970.
    7. Beckers, G., Bendt, A. K., Krämer, R. & Burkovski, A. (2004). Molecular identification of the urea uptake system and transcriptional analysis of urea transporter- and urease-encoding genes in Corynebacterium glutamicum. J Bacteriol 186, 7645–7652.
    8. Beckers, G., Strösser, J., Hildebrandt, U., Kalinowski, J., Farwick, M., Krämer, R. & Burkovski, A. (2005). Regulation of AmtR-controlled gene expression in Corynebacterium glutamicum: mechanism and characterization of the AmtR regulon. Mol Microbiol 58, 580–595.
    9. Bendt, A. K., Burkovski, A., Schaffer, S., Bott, M., Farwick, M. & Hermann, T. (2003). Towards a phosphoproteome map of Corynebacterium glutamicum. Proteomics 3, 1637–1646.
    10. Bendt, A. K., Beckers, G., Silberbach, M., Wittmann, A. & Burkovski, A. (2004). Utilization of creatinine as an alternative nitrogen source in Corynebacterium glutamicum. Arch Microbiol 181, 443–450.
    11. Bott, M. (2007). Offering surprises: TCA cycle regulation in Corynebacterium glutamicum. Trends Microbiol 15, 417–425.
    12. Botzenhardt, J., Morbach, S. & Krämer, R. (2004). Activity regulation of the betaine transporter BetP of Corynebacterium glutamicum in response to osmotic compensation. Biochim Biophys Acta 1667, 229–240.
    13. Brazma, A., Hingamp, P., Quackenbush, J., Sherlock, G., Spellman, P., Stoeckert, C., Aach, J., Ansorge, W., Ball, C. A. & other authors (2001). Minimum information about a microarray experiment (MIAME) –toward standards for microarray data. Nat Genet 29, 365–371.
    14. Brune, I., Werner, H., Hüser, A. T., Kalinowski, J., Pühler, A. & Tauch, A. (2006). The DtxR protein acting as dual transcriptional regulator directs a global regulatory network involved in iron metabolism of Corynebacterium glutamicum. BMC Genomics 7, 21.
    15. Buchinger, S., Strösser, J., Rehm, N., Hänßler, E., Hans, S., Bathe, B., Schomburg, D., Krämer, R. & Burkovski, A. (2009). A combination of metabolome and transcriptome analyses reveals new targets of the Corynebacterium glutamicum nitrogen regulator AmtR. J Biotechnol 140, 68–74.
    16. Cramer, A., Gerstmeir, R., Schaffer, S., Bott, M. & Eikmanns, B. J. (2006). Identification of RamA, a novel LuxR-type transcriptional regulator of genes involved in acetate metabolism of Corynebacterium glutamicum. J Bacteriol 188, 2554–2567.
    17. Durán, S., Pont, G. D., Huertazepeda, A. & Calderón, J. (1995). The role of glutaminase in Rhizobium etli – studies with a new mutant. Microbiology 141, 2883–2889.
    18. Durán, S., Sanchez Linares, L., Huerta Saquero, A., DuPont, G., Huerta Zepeda, A. & Calderón, J. (1996). Identification of two glutaminases in Rhizobium etli. Biochem Genet 34, 453–465.
    19. Eikmanns, B. J., Kleinertz, E., Liebl, W. & Sahm, H. (1991). A family of Corynebacterium glutamicum/Escherichia coli shuttle vectors for cloning, controlled gene expression, and promoter probing. Gene 102, 93–98.
    20. Eikmanns, B. J., Thum-Schmitz, N., Eggeling, L., Lüdtke, K. U. & Sahm, H. (1994). Nucleotide sequence, expression and transcriptional analysis of the Corynebacterium glutamicum gltA gene encoding citrate synthase. Microbiology 140, 1817–1828.
    21. Fisher, S. H. (1999). Regulation of nitrogen metabolism in Bacillus subtilis: vive la difference! Mol Microbiol 32, 223–232.
    22. Fisher, S. H. & Wray, L. V. (2008). Bacillus subtilis glutamine synthetase regulates its own synthesis by acting as a chaperone to stabilize GlnR–DNA complexes. Proc Natl Acad Sci U S A 105, 1014–1019.
    23. Fisher, S. H. & Wray, L. V. (2009). Novel trans-acting Bacillus subtilis glnA mutations that derepress glnRA expression. J Bacteriol 191, 2485–2492.
    24. Follmann, M., Ochrombel, I., Krämer, R., Trötschel, C., Poetsch, A., Rückert, C., Hüser, A., Persicke, M., Seiferling, D. & other authors (2009). Functional genomics of pH homeostasis in Corynebacterium glutamicum revealed novel links between pH response, oxidative stress, iron homeostasis and methionine synthesis. BMC Genomics 10, 621.
    25. Frunzke, J. & Bott, M. (2008). Regulation of iron homeostasis in Corynebacterium glutamicum. In Corynebacteria: Genomics and Molecular Biology, pp. 241–266. Edited by Burkovski, A.. Norfolk, UK: Caister Academic Press.
    26. Grant, S. G. N., Jessee, J., Bloom, F. R. & Hanahan, D. (1990). Differential plasmid rescue from transgenic mouse DNAs into Escherichia coli methylation-restriction mutants. Proc Natl Acad Sci U S A 87, 4645–4649.
    27. Hermann, T. (2003). Industrial production of amino acids by coryneform bacteria. J Biotechnol 104, 155–172.
    28. Hu, P., Leighton, T., Ishkhanova, G. & Kustu, S. (1999). Sensing of nitrogen limitation by Bacillus subtilis: comparison to enteric bacteria. J Bacteriol 181, 5042–5050.
    29. Hüser, A. T., Becker, A., Brune, I., Dondrup, M., Kalinowski, J., Plassmeier, J., Pühler, A., Wiegrabe, I. & Tauch, A. (2003). Development of a Corynebacterium glutamicum DNA microarray and validation by genome-wide expression profiling during growth with propionate as carbon source. J Biotechnol 106, 269–286.
    30. Ikeda, M. & Nakagawa, S. (2003). The Corynebacterium glutamicum genome: features and impacts on biotechnological processes. Appl Microbiol Biotechnol 62, 99–109.
    31. Ikeda, T. P., Shauger, A. E. & Kustu, S. (1996). Salmonella typhimurium apparently perceives external nitrogen limitation as internal glutamine limitation. J Mol Biol 259, 589–607.
    32. Jakoby, M., Tesch, M., Sahm, H., Krämer, R. & Burkovski, A. (1997). Isolation of the Corynebacterium glutamicum glnA gene encoding glutamine synthetase I. FEMS Microbiol Lett 154, 81–88.
    33. Jakoby, M., Krämer, R. & Burkovski, A. (1999). Nitrogen regulation in Corynebacterium glutamicum: isolation of genes involved and biochemical characterization of corresponding proteins. FEMS Microbiol Lett 173, 303–310.
    34. Jakoby, M., Nolden, L., Meier-Wagner, J., Krämer, R. & Burkovski, A. (2000). AmtR, a global repressor in the nitrogen regulation system of Corynebacterium glutamicum. Mol Microbiol 37, 964–977.
    35. Jiang, P. & Ninfa, A. J. (2009a). α-Ketoglutarate controls the ability of the Escherichia coli PII signal transduction protein to regulate the activities of NRII (NtrB) but does not control the binding of PII to NRII. Biochemistry 48, 11514–11521.
    36. Jiang, P. & Ninfa, A. J. (2009b). Sensation and signaling of α-ketoglutarate and adenylylate energy charge by the Escherichia coli PII signal transduction protein require cooperation of the three ligand-binding sites within the PII trimer. Biochemistry 48, 11522–11531.
    37. Kalinowski, J., Bathe, B., Bartels, D., Bischoff, N., Bott, M., Burkovski, A., Dusch, N., Eggeling, L., Eikmanns, B. J. & other authors (2003). The complete Corynebacterium glutamicum ATCC 13032 genome sequence and its impact on the production of L-aspartate-derived amino acids and vitamins. J Biotechnol 104, 5–25.
    38. Keilhauer, C., Eggeling, L. & Sahm, H. (1993). Isoleucine synthesis in Corynebacterium glutamicum: molecular analysis of the ilvB-ilvN-ilvC operon. J Bacteriol 175, 5595–5603.
    39. Kinoshita, S., Udaka, S. & Shimono, M. (2004). Studies on amino acid fermentation. Part I. Production of L-glutamic acid by various microorganisms. J Gen Appl Microbiol 50, 331–343.
    40. Lange, C., Rittmann, D., Wendisch, V. F., Bott, M. & Sahm, H. (2003). Global expression profiling and physiological characterization of Corynebacterium glutamicum grown in the presence of l-valine. Appl Environ Microbiol 69, 2521–2532.
    41. Leuchtenberger, W., Huthmacher, K. & Drauz, K. (2005). Biotechnological production of amino acids and derivatives: current status and prospects. Appl Microbiol Biotechnol 69, 1–8.
    42. Marienhagen, J. & Eggeling, L. (2008). Metabolic function of Corynebacterium glutamicum aminotransferases AlaT and AvtA and impact on l-valine production. Appl Environ Microbiol 74, 7457–7462.
    43. Marienhagen, J., Kennerknecht, N., Sahm, H. & Eggeling, L. (2005). Functional analysis of all aminotransferase proteins inferred from the genome sequence of Corynebacterium glutamicum. J Bacteriol 187, 7639–7646.
    44. Meier-Wagner, J., Nolden, L., Jakoby, M., Siewe, R., Krämer, R. & Burkovski, A. (2001). Multiplicity of ammonium uptake systems in Corynebacterium glutamicum: role of Amt and AmtB. Microbiology 147, 135–143.
    45. Möker, N., Brocker, M., Schaffer, S., Krämer, R., Morbach, S. & Bott, M. (2004). Deletion of the genes encoding the MtrA–MtrB two-component system of Corynebacterium glutamicum has a strong influence on cell morphology, antibiotics susceptibility and expression of genes involved in osmoprotection. Mol Microbiol 54, 420–438.
    46. Müller, T., Strösser, J., Buchinger, S., Nolden, L., Wirtz, A., Krämer, R. & Burkovski, A. (2006). Mutation-induced metabolite pool alterations in Corynebacterium glutamicum: towards the identification of nitrogen control signals. J Biotechnol 126, 440–453.
    47. Niebisch, A. & Bott, M. (2001). Molecular analysis of the cytochrome bc1–aa3 branch of the Corynebacterium glutamicum respiratory chain containing an unusual diheme cytochrome c1. Arch Microbiol 175, 282–294.
    48. Niebisch, A., Kabus, A., Schultz, C., Weil, B. & Bott, M. (2006). Corynebacterial protein kinase G controls 2-oxoglutarate dehydrogenase activity via the phosphorylation status of the OdhI protein. J Biol Chem 281, 12300–12307.
    49. Nohno, T., Saito, T. & Hong, J. (1986). Cloning and complete nucleotide sequence of the Escherichia coli glutamine permease operon (glnHPQ). Mol Gen Genet 205, 260–269.
    50. Nolden, L., Beckers, G., Möckel, B., Pfefferle, W., Nampoothiri, K. M., Krämer, R. & Burkovski, A. (2000). Urease of Corynebacterium glutamicum: organization of corresponding genes and investigation of activity. FEMS Microbiol Lett 189, 305–310.
    51. Nolden, L., Farwick, M., Krämer, R. & Burkovski, A. (2001a). Glutamine synthetases of Corynebacterium glutamicum: transcriptional control and regulation of activity. FEMS Microbiol Lett 201, 91–98.
    52. Nolden, L., Ngouoto-Nkili, C. E., Bendt, A. K., Krämer, R. & Burkovski, A. (2001b). Sensing nitrogen limitation in Corynebacterium glutamicum: the role of glnK and glnD. Mol Microbiol 42, 1281–1295.
    53. Picossi, S., Belitsky, B. R. & Sonenshein, A. L. (2007). Molecular mechanism of the regulation of Bacillus subtilis gltAB expression by GltC. J Mol Biol 365, 1298–1313.
    54. Polen, T. & Wendisch, V. F. (2004). Genomewide expression analysis in amino acid-producing bacteria using DNA microarrays. Appl Biochem Biotechnol 118, 215–232.
    55. Sambrook, J., MacCallum, P. & Russell, D. (2001). Molecular Cloning. A Laboratory Manual, 3rd edn. Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press.
    56. Schäfer, A., Tauch, A., Jäger, W., Kalinowski, J., Thierbach, G. & Pühler, A. (1994). Small mobilizable multipurpose cloning vectors derived from the Escherichia coli plasmids pK18 and pK19 – selection of defined deletions in the chromosome of Corynebacterium glutamicum. Gene 145, 69–73.
    57. Schaffer, S., Weil, B., Nguyen, V. D., Dongmann, G., Günther, K., Nickolaus, M., Hermann, T. & Bott, M. (2001). A high-resolution reference map for cytoplasmic and membrane-associated proteins of Corynebacterium glutamicum. Electrophoresis 22, 4404–4422.
    58. Schägger, H. & von Jagow, G. (1987). Tricine-sodium dodecyl sulfate-polyacrylamide gel electrophoresis for the separation of proteins in the range from 1 to 100 kDa. Anal Biochem 166, 368–379.
    59. Schmitz, R. A. (2000). Internal glutamine and glutamate pools in Klebsiella pneumoniae grown under different conditions of nitrogen availability. Curr Microbiol 41, 357–362.
    60. Schreier, H. J., Brown, S. W., Hirschi, K. D., Nomellini, J. F. & Sonenshein, A. L. (1989). Regulation of Bacillus subtilis glutamine synthetase gene expression by the product of the glnR Gene. J Mol Biol 210, 51–63.
    61. Schulz, A. A., Collett, H. J. & Reid, S. J. (2001). Nitrogen and carbon regulation of glutamine synthetase and glutamate synthase in Corynebacterium glutamicum ATCC 13032. FEMS Microbiol Lett 205, 361–367.
    62. Shapiro, B. M. & Stadtman, E. R. (1970). Glutamine synthetase (Escherichia coli). Methods Enzymol 17A, 910–922.
    63. Shiio, I. & Ozaki, H. (1970). Regulation of nicotinamide adenine dinucleotide phosphate-specific glutamate dehydrogenase from Brevibacterium flavum, a glutamate-producing bacterium. J Biochem 68, 633–647.
    64. Siewe, R. M., Weil, B. & Krämer, R. (1995). Glutamine uptake by a sodium-dependent secondary transport system in Corynebacterium glutamicum. Arch Microbiol 164, 98–103.
    65. Silberbach, M., Hüser, A., Kalinowski, J., Pühler, A., Walter, B., Krämer, R. & Burkovski, A. (2005a). DNA microarray analysis of the nitrogen starvation response of Corynebacterium glutamicum. J Biotechnol 119, 357–367.
    66. Silberbach, M., Schäfer, M., Hüser, A. T., Kalinowski, J., Pühler, A., Krämer, R. & Burkovski, A. (2005b). Adaptation of Corynebacterium glutamicum to ammonium limitation: a global analysis using transcriptome and proteome techniques. Appl Environ Microbiol 71, 2391–2402.
    67. Sonenshein, A. L. (2007). Control of key metabolic intersections in Bacillus subtilis. Nat Rev Microbiol 5, 917–927.
    68. Tesch, M., Eikmanns, B. J., de Graaf, A. A. & Sahm, H. (1998). Ammonia assimilation in Corynebacterium glutamicum and a glutamate dehydrogenase-deficient mutant. Biotechnol Lett 20, 953–957.
    69. Udaka, S. (1960). Screening method for microorganisms accumulating metabolites and its use in the isolation of Micrococcus glutamicus. J Bacteriol 79, 754–755.
    70. van der Rest, M. E., Lange, C. & Molenaar, D. (1999). A heat shock following electroporation induces highly efficient transformation of Corynebacterium glutamicum with xenogeneic plasmid DNA. Appl Microbiol Biotechnol 52, 541–545.
    71. Weiss, V., Kramer, G., Dunnebier, T. & Flotho, A. (2002). Mechanism of regulation of the bifunctional histidine kinase NtrB in Escherichia coli. J Mol Microbiol Biotechnol 4, 229–233.
    72. Wendisch, V. F. (2003). Genome-wide expression analysis in Corynebacterium glutamicum using DNA microarrays. J Biotechnol 104, 273–285.
    73. Wennerhold, J. & Bott, M. (2006). The DtxR regulon of Corynebacterium glutamicum. J Bacteriol 188, 2907–2918.
    74. Wennerhold, J., Krug, A. & Bott, M. (2005). The AraC-type regulator RipA represses aconitase and other iron proteins from Corynebacterium under iron limitation and is itself repressed by DtxR. J Biol Chem 280, 40500–40508.
    75. Wray, L. V. & Fisher, S. H. (2008). Bacillus subtilis GlnR contains an autoinhibitory C-terminal domain required for the interaction with glutamine synthetase. Mol Microbiol 68, 277–285.
    76. Wray, L. V., Ferson, A. E., Rohrer, K. & Fisher, S. H. (1996). TnrA, a transcription factor required for global nitrogen regulation in Bacillus subtilis. Proc Natl Acad Sci U S A 93, 8841–8845.
    77. Wray, L. V., Zalieckas, J. M., Ferson, A. E. & Fisher, S. H. (1998). Mutational analysis of the TnrA-binding sites in the Bacillus subtilis nrgAB and gabP promoter regions. J Bacteriol 180, 2943–2949.
    78. Zhang, Y., Pohlmann, E. L., Serate, J., Conrad, M. C. & Roberts, G. P. (2010). Mutagenesis and functional characterization of the four domains of GlnD, a bifunctional nitrogen sensor protein. J Bacteriol 192, 2711–2721.