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Environmental Microbiology (2015) 17(9), 3379–3390

doi:10.1111/1462-2920.12813

Evidence for synergistic control of glutamate biosynthesis by glutamate dehydrogenases and glutamate in Bacillus subtilis

Lorena Stannek,1 Martin J. Thiele,1 Till Ischebeck,2 Katrin Gunka,1 Elke Hammer,3 Uwe Völker3 and Fabian M. Commichau1* 1 Department of General Microbiology, Institute of Microbiology and Genetics, Georg-August-University Göttingen, Grisebachstr. 8, Göttingen D-37077, Germany. 2 Department for Plant Biochemistry, Albrecht-von-Haller-Institute for Plant Sciences, Georg-August-University Göttingen, Grisebachstr. 8, Göttingen D-37077, Germany. 3 Interfaculty Institute for Genetics and Functional Genomics, University Medicine Greifswald, Friedrich-Ludwig-Jahnstr. 15a, Greifswald D-17475, Germany. Summary In the Gram-positive bacterium, Bacillus subtilis glutamate is synthesized by the glutamine synthetase and the glutamate synthase (GOGAT). During growth with carbon sources that exert carbon catabolite repression, the rocG glutamate dehydrogenase (GDH) gene is repressed and the transcription factor GltC activates the expression of the GOGAT encoding gltAB genes. In the presence of amino acids of the glutamate family, the GDH RocG is synthesized and the enzyme prevents GltC from binding to DNA. The dual control of glutamate biosynthesis allows the efficient utilization of the available nutrients. Here we provide genetic and biochemical evidence that, like RocG, also the paralogous GDH GudB can inhibit the transcription factor GltC, thereby controlling glutamate biosynthesis. Contradictory previous observations show that high level of GDH activity does not result in permanent inhibition of GltC. By controlling the intracellular levels of glutamate through feeding with exogenous arginine, we observed that the GDH-dependent control of GltC and thus expression of the gltAB genes inversely Received 22 December, 2014; revised 4 February, 2015; accepted 11 February, 2015. *For correspondence. E-mail [email protected]; Tel. (+49) 551 393 3796; Fax (+49) 551 393 3808.

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd

correlates with the glutamate pool. These results suggest that the B. subtilis GDHs RocG and GudB in fact act as glutamate sensors. In conclusion, the GDH-mediated control of glutamate biosynthesis seems to depend on the intracellular glutamate concentration. Introduction The major amino group donor for nitrogen-containing building blocks is glutamate, which can be synthesized de novo via two metabolic pathways (Wohlheuter et al., 1973; Magasanik, 2003; Gunka and Commichau, 2012). In both pathways, 2-oxoglutarate (2-OG) serves as the carbon backbone that is derived from the tricarboxylic acid cycle. Therefore, glutamate biosynthesis is a key metabolic intersection that links carbon to nitrogen metabolism (Commichau et al., 2006; Sonenshein, 2007). In the longer glutamate biosynthetic pathway, which involves the ATP-dependent glutamine synthetase (GS) and the glutamate synthase (GOGAT), 2-OG and ammonium are converted to glutamate. In the shorter pathway, the NADPH2-dependent glutamate dehydrogenase (GDH) directly converts 2-OG and ammonium to glutamate. The latter pathway seems to be clearly more profitable to the cell because it does not consume ATP. In bacteria like the Gram-negative organism Escherichia coli, the GDH can only synthesize glutamate if sufficient ammonium is available because the enzyme has a much lower affinity for ammonium than the GS (Helling, 1994; Reitzer, 2003). In contrast to the situation in E. coli, in the Gram-positive model bacterium Bacillus subtilis, glutamate is exclusively synthesized by the combined action of the GS and the GOGAT (Fig. 1; Belitsky and Sonenshein, 1998; Commichau and Stülke, 2008; Commichau et al., 2008). In B. subtilis, the hexameric GDH RocG has a very low affinity for ammonium and the enzyme is strictly devoted to glutamate catabolism (Fig. 1; Commichau and Stülke, 2008; Commichau et al., 2008; Gunka et al., 2010). In B. subtilis, glutamate biosynthesis is controlled by the availability of carbon and nitrogen sources (for a recent review Gunka and Commichau, 2012). In the presence of the preferred carbon source glucose, which exerts

3380 L. Stannek et al. Fig. 1. Schematic illustration of de novo glutamate biosynthesis and Roc pathway for utilization of arginine and related amino acids. PtsG, glucose permease; GS, glutamine synthetase; GOGAT, glutamate synthase; RocG and GudB, glutamate dehydrogenases; GltC, transcriptional regulator of the GOGAT-encoding gltAB genes; RocA and PutC, Δ1-pyrroline-5-carboxylate dehydrogenases; RocD, ornithine aminotransferase; RocF, arginase; RocC and RocE, arginine permeases; 2-OG, 2-oxoglutarate; TCA, tricarboxylic acid.

carbon catabolite repression (CCR), the DNA-binding transcription factor GltC activates transcription of the gltAB genes and the encoded GOGAT synthesizes glutamate (Bohannon and Sonenshein, 1989; Wacker et al., 2003; Picossi et al., 2007). At the same time CCR prevents transcription of the rocG gene and the catabolically active GDH RocG is not synthesized (Belitsky and Sonenshein, 2004; Belitsky et al., 2004). The carbon source-dependent induction and repression of the gltAB and rocG genes, respectively, ensure that the cell synthesizes sufficient glutamate, which is needed to achieve high growth rates if a good carbon source is available. In contrast, in the absence of CCR-mediating carbon sources like succinate, the gltAB genes are not expressed (Belitsky and Sonenshein, 2004; Commichau et al., 2007a). The gltAB genes are also not expressed if the bacteria grow with nitrogen sources like arginine, which induces the expression of the arginine degradation pathway, including the GDH-encoding rocG gene (Fig. 1; Gardan et al., 1997; Belitsky and Sonenshein, 1998). The underlying mechanism of how the expression of the gltAB genes is regulated by signals derived from carbon and nitrogen metabolism has been enigmatic for a long time. Recently, we found that the GDH RocG directly binds to the transcription factor GltC, and prevents it from activating expression of the gltAB genes (Commichau et al., 2007b; Herzberg et al., 2007). The observation that the GDH RocG negatively controls the activity of GltC and thus expression of the gltAB genes is in line with the observation that the gltAB genes are constitutively expressed in a rocG mutant (Belitsky and Sonenshein, 2004; Belitsky et al., 2004; Commichau et al., 2007a). The GDH RocG is a trigger enzyme that is active in metabolism and in controlling gene expression (Commichau and Stülke, 2008). This sophisticated control of glutamate biosynthesis by RocG ensures that the glutamatesynthesizing GOGAT is only formed if there is no exogenous source of glutamate like arginine available and if

the demand for the important amino group donor glutamate is high. However, there are some discrepancies to be resolved because constitutive expression of the rocG gene does not result in a permanent inhibition of the transcription factor GltC (see below). In addition to the rocG gene, the genomes of B. subtilis laboratory strains such as 168 contain the cryptic gudBCR gene, encoding the enzymatically inactive GDH GudBCR (Belitsky and Sonenshein, 1998; Zeigler et al., 2008). The GDH GudBCR is non-functional because the gudBCR gene contains a perfect 18 bp-long direct repeat (DR) and the resulting duplication of three amino acids in a conserved region of the encoded protein renders it enzymatically inactive (Belitsky and Sonenshein, 1998). Although the gudBCR gene is constitutively transcribed, the encoded GudBCR protein is subject to rapid proteolytic degradation, especially under conditions of glucose starvation (Gerth et al., 2008; Gunka and Commichau, 2012; Gunka et al., 2012). It is interesting to note that B. subtilis can afford to inherit the gudBCR gene and to permanently synthesize a non-functional enzyme. The maintenance of the cryptic gudBCR gene might allow the bacteria to efficiently cope with a certain supply of nutrients. Indeed, it can be advantageous for B. subtilis to only synthesize the GDH RocG if the supply with exogenous glutamate is low. By contrast, synthesis of two functional GDHs can be advantageous for the bacteria because high-level GDH activity allows the utilization of glutamate as the single carbon and nitrogen source (Belitsky and Sonenshein, 1998; Gunka et al., 2013; Stannek et al., 2014). Strains synthesizing the two GDHs RocG and GudB can easily be isolated on glutamate-containing minimal medium. Interestingly, the spontaneously emerging mutants harbour the mutated gudB allele, lacking one repeat unit of the 18 bp-long DR that is present in the gudBCR gene (Belitsky and Sonenshein, 1998; Gunka et al., 2013). The encoded GDH GudB is enzymatically active and not subject to rapid proteolytic degradation (Gunka and Commichau, 2012; Gunka et al., 2012; 2013). Suppressor mutants that

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

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Fig. 2. Western blot analysis to estimate the GDH levels in strains expressing either rocG or gudB, or both GDHs. The B. subtilis strains GP28 (rocG::Tn10 ΔgudB), GP342 (rocG gudBCR), GP32 (rocG::Tn10 gudB) and GP804 (rocG gudB) were grown in CSE minimal medium in the absence (−) and in the presence of 0.5% (w/v) glucose (Glc) and arginine (Arg) respectively. Cells were harvested at an OD600 between 0.5 and 0.8. After electrophoresis in a 12.5% sodium dodecyl sulfate polyacrylamide (SDS-PAA) gel and transfer onto a polyvinylidene difluoride (PVDF) membrane, RocG and GudB were detected using a rabbit polyclonal antibodies raised against RocG. Fifteen microgram of total cell protein per lane were applied. n.d. = not determined.

have spontaneously mutated the cryptic gudBCR do also rapidly occur in the background of a B. subtilis rocG mutant strain during growth on rich medium (Belitsky and Sonenshein, 1998; Gunka et al., 2013). Under these growth conditions, a B. subtilis strain lacking GDH activity has a severe growth defect that is probably due to the accumulation of unknown toxic intermediates of the arginine, ornithine and citrulline (Roc) degradation pathway (Fig. 1) and because of de-regulation of glutamate biosynthesis (Commichau et al., 2007a). However, under laboratory growth conditions, one GDH is sufficient for maintaining glutamate homeostasis (Gunka and Commichau, 2012). Previous studies suggest that the active GudB variant can compensate for the loss of RocG. This work provides genetic and biochemical evidence that, like RocG, the active GDH GudB directly binds to the transcription factor GltC, thereby controlling glutamate biosynthesis in B. subtilis. Moreover, we found that the GDHs interact with each other in vivo. We also shed light on the surprising previous observation that high-level GDH activity does not result in permanent inhibition of GltC but rather in high-level activity of the transcription factor. By metabolome analyses of a strain that synthesizes constant amounts of RocG, we show that the GDHdependent control of GltC activity directly correlates with the intracellular amounts of glutamate. These results suggest that the GDH acts as a glutamate sensor and that the GDH-mediated control of glutamate biosynthesis depends on the intracellular glutamate pool.

Results GudB is a trigger enzyme that is active in metabolism and in controlling glutamate biosynthesis To study the role of GudB in the control of glutamate biosynthesis, we assessed the activity of a translational gltA-lacZ fusion in strain GP32 (rocG::Tn10 gudB gltAlacZ) that synthesizes only the GDH GudB. The wild-type

strain GP342 (rocG gudBCR gltA-lacZ) that only synthesizes RocG and the GDH-negative strain GP28 (rocG::Tn10 ΔgudB gltA-lacZ) served as controls. All strains were grown in CSE minimal medium containing succinate (S) and glutamate (E) as sources of carbon and nitrogen respectively. We also performed Western blot analyses to monitor the GDH level (Fig. 2). As expected and reported previously (Belitsky and Sonenshein, 2004; Commichau et al., 2007a), when cultivated in CSE medium, the gltA promoter was highly active in strain GP28 that does not synthesize a GDH (Table 1; Fig. 2). By contrast, the gltA promoter showed only basal activity in the RocG-synthesizing wild-type strain GP342 and in strain GP32 that synthesizes GudB instead of RocG (Fig. 2). Thus, like RocG, the enzymatically active GDH GudB seems to be able to control glutamate biosynthesis. Next, we analysed the effect of the carbon and nitrogen sources glucose and arginine, respectively, on the expression of the gltAB genes in the wild-type strain GP342, in the GDH-deficient strain GP28 and in strain GP32 that synthesizes only GudB. As expected, the lack of RocG, being the negative effector of GltC, results in constitutive Table 1. Analysis of gltA expression. β-Galactosidase activitya Strain

Relevant genotype

GP28 GP342 GP32 GP804

rocG::Tn10 ΔgudB rocG gudBCR rocG::Tn10 gudB rocG gudB

309 8 12 36

+ Glc

+ Arg

+ Glc + Arg

335 216 160 160

n.g. 6 12 2

262 12 6 4

a. Bacteria were grown in C minimal medium. Succinate (Suc), Glucose (Glc), glutamate (Glt) and arginine (Arg) were added to final concentrations of 0.5% (w/v; Suc, Glc and Arg) or 0.8% (w/v; Glt). β-Galactosidase activities are given as units per milligram of protein. Cells were harvested at an OD600 between 0.5 and 0.8. Experiments were carried out at least threefold. The maximum deviation of the series of representative data shown here was less than 30%. n.g. = No growth.

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

3382 L. Stannek et al. expression of the gltAB genes in strain GP28 (Fig. 2; Table 1). While in the wild-type strain GP342 expression of the gltAB genes was induced by glucose, the addition of arginine alone or when added together with glucose resulted in strong expression and repression of the rocG and gltAB genes respectively (Table 1, Fig. 2). These results are in perfect agreement with previous observations, suggesting that high amounts of RocG cause inhibition of GltC (Commichau et al., 2007b). Surprisingly, expression of the gltAB genes in strain GP32 was also induced by glucose despite the presence of significant amounts of the GDH GudB (Table 1, Fig. 2). This might indicate that GudB requires an unknown cofactor to prevent binding of GltC to the gltA promoter. It has been suggested previously that this cofactor might be derived from the Roc pathway (see below; Belitsky and Sonenshein, 2004; Gunka et al., 2010). Indeed, the addition of arginine alone or in combination with glucose completely abolished expression of the gltAB genes in strain GP32. In conclusion, the enzymatically active GDH GudB can fully compensate for the loss of RocG. GudB directly interacts with the transcription activator GltC As shown above, the expression of the gltAB genes is completely restored in a strain expressing the

enzymatically active GDH GudB (see Table 1). This observation suggests that like RocG also GudB controls glutamate biosynthesis by a direct protein–protein interaction with the DNA-binding transcription factor GltC. To test this hypothesis, we performed a Strep–protein interaction experiment (SPINE) that has been used previously to monitor the interaction between RocG and GltC in vivo (Commichau et al., 2007b; Herzberg et al., 2007). For this purpose, the C-terminally Strep-tagged GltC protein (bait) was expressed from a multi-copy plasmid pGP916 in the rocG mutant strain GP32-pGP916 (rocG::Tn10 gudB gltC-Strep) harbouring the gudB allele, which encodes the functional GDH GudB. The strain GP27-pGP916 (rocG ΔgudB gltC-Strep) expressing the rocG gene served as the positive control. Moreover, isogenic strains carrying the empty vector pBQ200 served as negative controls. All strains were grown in CSE-Glc-Arg minimal medium, containing succinate (Suc) and glucose (Glc) as sources of carbon, and glutamate (Glt) and arginine (Arg) as sources of nitrogen. For the detection of protein–protein interactions, we performed SPINE using formaldehyde as a cross-linker. The samples of the SPINE were analysed by SDS-PAGE and Western blotting. As expected, no protein was purified from the crude extracts obtained from the strains that harbour the empty vector (Fig. 3A). By contrast, GltC together with RocG was purified from crude extracts of strain GP27-pGP916 (rocG ΔgudB gltC-Strep),

Fig. 3. In vivo cross-linking experiment to study the interaction between the transcription factor GltC and the GDHs RocG and GudB. A. Protein complexes were isolated from the B. subtilis strains GP27-pGP916 (rocG ΔgudB gltC-Strep) and GP32-pGP916 (rocG::Tn10 gudB gltC-Strep) synthesizing RocG and GudB, respectively, together with the C-terminally Strep-tagged GltC protein. As indicated in the figure, RocG and in the elution fractions of strains GP27-pGP916 and GP32-pGP916, respectively, were identified by mass spectrometry (lower and upper bands; Table S3). Strains GP32 and GP27 carrying the empty vector pBQ200 (no gltC) served as negative controls. All strains were grown in CSE-Glc-Arg minimal medium containing 0.5% glucose and 0.5% arginine. (A) Washing (W) and elution (E) fractions from each purification (15 μl) were loaded onto the sodium dodecyl sulfate polyacrylamide (SDS-PAA) gel and the proteins were visualized by silver staining. M, protein standard. B. Western blot analysis to verify the presence of GltC and of the GDHs RocG and GudB in the elution fractions that were obtained from crude extracts of strains GP32-pGP916 and GP27-pGP916 using GltC and RocG polyclonal antibodies.

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

Control of glutamate biosynthesis in Bacillus subtilis which confirmed the previous observation that RocG interacts with GltC (Commichau et al., 2007b; Herzberg et al., 2007). Moreover, when GltC-Strep served as the bait protein in strain GP32-pGP916 (rocG::Tn10 gudB gltC-Strep) synthesizing the functional GDH GudB, two proteins appeared in the elution fraction, suggesting that like RocG also GudB forms a complex with GltC. Western blot analysis revealed that these proteins were indeed GudB and GltC (Fig. 3B). To ultimately verify the identity of the proteins in the elution fraction, the bands corresponding to RocG (lower band, 46 kDa) and GudB (higher band, 47 kDa) of the silver-stained SDS-PAGE were cut out and analysed by mass spectrometry. The mass spectrometric analyses revealed that RocG and GudB were present in the elution fractions of strains GP27-pGP916 (rocG ΔgudB gltC-Strep) and GP32-pGP916 (rocG::Tn10 gudB gltC-Strep) respectively (Table S3). Taken together, the results unequivocally indicate that both GDHs RocG and GudB can physically interact with the transcription factor GltC. Interactions between the GDHs in vivo As shown above, both GDHs may directly bind to GltC and control the DNA-binding activity of the transcription factor (see Fig. 3A and B). Since we used strains that either expressed rocG or gudB, we were curious whether RocG and GudB may heterohexamerize in vivo. To address his question, we performed a SPINE using the strain GP442-pBP179 (rocG Strep-gudB) that

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synthesizes the enzymatically active GDHs RocG and Strep-GudB. For this purpose, the bacteria were grown in CSE-Glc-Arg minimal medium and the protein–protein interactions were fixed in the growing cells by formaldehyde cross-linking (see above). The bacteria were collected by centrifugation, disrupted and the protein complexes were isolated from cell-free crude extracts by affinity purification. Aliquots of the final washing and elution steps were subjected to SDS-PAGE, and the proteins were visualized by silver staining. As shown in Fig. 4A, two proteins were visible on the gel, which were most likely the Strep-GudB bait protein (48 kDa) and the GDH RocG (46 kDa). Mass spectrometric analysis of the silver-stained bands revealed that RocG (identified by 16 and 46 in the upper and lower bands respectively) was indeed co-purified with the Strep-tagged GudB protein (identified by 40 and 28 peptides in the upper and lower bands respectively) (Table S4). To confirm the interaction by a complementary method, we used the bacterial adenylate cyclase two-hybrid (B2H) system (Karimova et al., 1998). We also tested whether the enzymatically inactive GudBCR protein is able to interact with the active GDHs GudB and RocG. As expected, the B2H analysis revealed that all GDH variants interact with each other and that GudB interacts with RocG (Fig. 4B). Moreover, we observed that enzymatically inactive GudBCR protein, which is subject to rapid proteolytic degradation in B. subtilis interacts with its active counterpart. In contrast to this, no interaction was observed between inactive GudBCR protein and RocG (Fig. 4B). Therefore, it seems

Fig. 4. Interactions among the B. subtilis glutamate dehydrogenases. A. In vivo cross-linking experiment to study the interaction between the GDHs RocG and GudB. Protein complexes were isolated from the B. subtilis strains BP442-pBP179 (rocG ΔgudB Strep-gudB) synthesizing RocG together with the N-terminally Strep-tagged GudB protein. The RocG and GudB proteins in the upper and lower bands of the elution fraction were identified by mass spectrometry (Table S4). The strain was grown in CSE-Glc-Arg minimal medium containing 0.5% glucose and 0.5% arginine. Forty microlitre elution fraction were loaded onto the sodium dodecyl sulfate polyacrylamide (SDS-PAA) gel and the proteins were visualized by silver staining. M, protein standard. B. B2H analysis to study the interactions among the active GDHs RocG and GudB, and the inactive GudBCR protein. The rocG, gudB and gudBCR genes were cloned in the plasmids pUT18, pUT18C, p25-N and pKT25. Plasmids pUT18 and pUT18C allow the expression of the GDHs fused either to the N- or the C-terminus of the T18 domain of the B. pertussis adenylate cyclase respectively. Plasmid p25-N and pKT25 allow the expression of the GDHs fused to the N- or the C-terminus of the T25 domain of the adenylate cyclase. The E. coli transformants were incubated for 24 h at 28°C.

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

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unlikely that the GudBCR protein interferes with the control of GltC activity by RocG in vivo. The active GDHs RocG and GudB share 74% sequence identity, which might allow a strong interaction among the active GDHs. Since the inactive GDH GudBCR seems to be completely misfolded (Stannek et al., 2015), one could imagine that less residues in the inactive GudBCR protein needed for the interaction with RocG are accessible. Taken together, it seems highly likely that the enzymatically active GDHs RocG and GudB can form homo- or heterohexameric complexes, and each of the complexes is probably able to control the DNA-binding activity of the transcription factor GltC (see Discussion). The effect of GDH levels on glutamate biosynthesis It is interesting to note that as long as the rocG gene is intact, the cryptic gudBCR allele is stably inherited in the domesticated B. subtilis strain 168 (Belitsky and Sonenshein, 1998; Zeigler et al., 2008; Gunka and Commichau, 2012; Gunka et al., 2012). Recently, we have shown that the presence of only one GDH-encoding gene (rocG) in the genome of the laboratory strain provides the bacteria with a selective growth advantage when exogenous glutamate is scarce (Gunka and Commichau, 2012; Gunka et al., 2012; Stannek et al., 2014). However, it is unclear if the growth advantage is simply due to the reduced glutamate-degrading enzyme activity or if high GDH levels disturb regulation of glutamate biosynthesis. To test whether the presence of two enzymatically active GDHs affects expression of the gltAB genes, we monitored the activity of the gltA promoter in strain GP804 (rocG gudB gltA-lacZ). We also analysed the cellular GDH levels by Western blotting (Fig. 2). As shown in Table 1, in comparison to strains GP342 and GP32 synthesizing RocG and GudB, respectively, expression of the gltAB genes was threefold to fourfold enhanced in strain GP804 during growth in CSE medium. Thus, increased GDH levels (see Fig. 2) indeed disturb regulation of glutamate biosynthesis. This finding is in good agreement with previous observations showing that GltC is not inhibited when RocG, the negative effector of GltC, is overproduced (Belitsky and Sonenshein, 1998; Gunka et al., 2010). By contrast, no difference in glucose- and arginine-dependent regulation of glutamate biosynthesis was observed between the strain GP804 and the strains GP342 and GP32 that synthesize only one GDH. In all strains, the gltAB genes were highly expressed in the presence of glucose but repressed with arginine alone or when added in combination with glucose (Table 1). In conclusion, the presence of only one functional GDH-encoding gene in the laboratory strain 168 seems to be advantageous for the cells because it prevents the formation of a futile cycle: the simultaneous

degradation of glutamate and deregulation of the gltAB genes that in turn results in glutamate biosynthesis (Fig. 1). Synergistic control of GltC activity by GDH and glutamate Previously, it has been reported that overexpression of the rocG gene resulted in high-level expression of the gltAB genes (Belitsky and Sonenshein, 2004; Gunka et al., 2010). Moreover, in the present study, we show that expression of the gltAB genes was enhanced in a strain that synthesized RocG together with the enzymatically active GDH GudB (Table 1). Surprisingly, the addition of arginine rescued the inhibitory effect of RocG and GudB on gltAB expression. This observation suggests that the GDHs degrade a Roc pathway-derived cofactor that is needed for the GDH-dependent regulation of glutamate biosynthesis in B. subtilis (Fig. 1). To narrow down the cofactor, we monitored the activity of the gltA promoter and simultaneously determined the cellular amounts of the intermediates of the Roc pathway in strain BP220-pBP529 (rocG::Tn10 ΔgudB ΔgltAB gltA-lacZ rocG) that was grown with various amounts of arginine. This strain is unable to synthesize glutamate de novo and constitutively expresses the rocG gene from plasmid pGP529 (see Fig. S1). Therefore, we expected that the GDH-dependent control of GltC activity by intermediates of the Roc pathway should strictly depend on exogenously available arginine. Indeed, with low amounts of exogenous arginine, the gltA promoter in strain BP220-pBP529 was highly active but the activity gradually decreased with increasing amounts of arginine (Fig. 5A). By contrast, the gltA promoter and thus GltC was constitutively active in the control strain BP220pBQ200 (rocG::Tn10 ΔgudB ΔgltAB gltA-lacZ) carrying the empty plasmid (Fig. 5A). Next, using gas chromatography coupled with mass spectrometry (GC-MS), we determined the intracellular levels of the intermediates of the Roc pathway in strain BP220-pBP529 that was grown with various amounts of arginine (see Experimental procedures). As shown in Fig. 5B, by increasing the amounts of arginine from 0.01% to 0.05%, the cellular glutamate levels also increased. By contrast, arginine and ornithine only accumulated in the cells when 0.5% arginine was added to the medium. The fact that in the range between 0.01% and 0.05% of exogenous arginine, only the cellular glutamate levels inversely correlated with the activity of the gltA promoter (r = −0.96; P-value of < 0.01) strongly suggests that RocG and glutamate synergistically inhibit the DNAbinding activity of GltC (Fig. 5A and B). In conclusion, RocG and GudB seem to act as sensors for glutamate, and depending on the intracellular concentration of the central metabolite, the GDHs adjust glutamate biosynthesis accordingly (see Discussion).

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

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Fig. 5. Inverse correlation between GltC activity and intracellular glutamate levels. A. Activity of the gltA promoter in strain BP220-pBP529 (rocG::Tn10 ΔgudB ΔgltAB rocG gltA-lacZ) constitutively expressing rocG and in the control strain BP220-pBQ200 (rocG::Tn10 ΔgudB ΔgltAB gltA-lacZ) carrying the empty plasmid. The bacteria were grown in CSE-Glc minimal medium supplemented with the indicated amounts of arginine. The β-galactosidase assay was carried out at least threefold. Representative results from one series are shown. B. Metabolome analysis to the intracellular levels of the intermediates of the Roc pathway in strain BP220-pBP529 (rocG::Tn10 ΔgudB ΔgltAB gltA-lacZ) that was grown in CSE-Glc minimal medium supplemented with the indicated amounts of arginine. The mean values from five biological replicates are shown. The maximum deviation was < 30%. The Pearson correlation coefficients (r) were calculated to evaluate the correlations between the metabolites of the arginine degradation pathway and the activity of the gltA promoter.

Discussion In the present study, we show that the enzymatically active GDH GudB can control glutamate biosynthesis in B. subtilis through a direct protein–protein interaction with GltC, the transcriptional activator of the gltAB genes. Thus, like RocG, also the ‘decryptified’ GDH GudB lacking the duplication of three amino acids in its active centre is a bifunctional enzyme. It belongs to the class of so-called trigger enzymes, which are active in metabolism and in controlling gene expression (Commichau and Stülke, 2008). It has been previously suggested that RocG and GudB constitute the major and minor GDHs, respectively, and that the GDHs allow B. subtilis to utilize amino acids of the glutamate family (Belitsky and Sonenshein, 1998). In contrast to the rocG gene, whose expression is tightly controlled by signals derived from carbon and nitrogen metabolism, the gudB gene is constitutively transcribed (Belitsky and Sonenshein, 2004; Belitsky et al., 2004; Gunka and Commichau, 2012; Gunka et al., 2012). Except for the domesticated B. subtilis strains 160, 166 and 168 that harbour the cryptic gudBCR allele, the lessdomesticated strains express the gudB allele, encoding the functional GDH GudB (Zeigler et al., 2008). Thus, the genetic makeup of the common laboratory strain 168 is rather unnatural and does not reflect the situation in the non-domesticated B. subtilis strains and their derivatives. Therefore, we suggest that GudB and RocG constitute the

major and minor GDHs, of which the latter enzyme is only required by the bacteria when amino acids of the glutamate family (e.g. arginine) are available in the environment in high amounts (see Fig. 1). The present study also revealed that the enzymatically active GDHs RocG and GudB interact with each other using a B2H system and by performing SPINE. These observations suggest that the B. subtilis GDHs are able to form heterohexamers that can, similar to the GDH homohexamers, regulate the DNA-binding activity of GltC. However, it is yet unclear whether heterohexamerization is of in vivo relevance in a strain encoding two functional GDHs. It is interesting to note that the genome of B. subtilis encodes two GDHs that can fully replace each other: GudB as well as RocG are both active in degradation of glutamate and in controlling the DNA-binding activity of the transcription factor GltC (Commichau et al., 2007b). Intra-species homologues like the B. subtilis GDHs can be acquired by gene duplication or by horizontal gene transfer. Several lines of evidence support the hypothesis that the gudB and rocG genes have rather been emerged through gene duplication than through horizontal gene transfer. Both GDHs are bifunctional enzymes having the same interaction partners in the cell and the biochemical properties of the enzymes are almost identical (Gunka et al., 2010). Moreover, the structural similarity of the GDHs, which is certainly due to presence of 74% of

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

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Fig. 6. Model for the control of GltC activity by glutamate dehydrogenase (GDH) and glutamate. Enzymatically idle GDH complexes indicated by black circles do not inhibit the DNA-binding activity of the transcription factor GltC. Catabolically active GDH complexes indicated by black rectangles inhibit GltC.

identical amino acid in the proteins, allow them to form heteromultimers. It is commonly well accepted that gene duplication contributes to the emergence of novel genetic information because both paralogues might acquire a new function by spontaneous mutagenesis and fixation of the beneficial mutation by purifying selection (Ohno, 1970; Gevers et al., 2004; Conant and Wolfe, 2008). Indeed, adaptive specialization after gene duplication is important for the diversification of transcription factors and for the development of enzymes with altered substrate specificities (Teichmann and Babu, 2004; Zhang et al., 2004; Peréz et al., 2014). Alternatively, gene duplication and the resulting increase in gene dosage enable bacteria to cope with stressful conditions like high temperatures (Riehle et al., 2001). It has also been suggested that duplicated genes are rapidly lost by the accumulation of deleterious mutations (Lynch and Conery, 2000; Lynch, 2007). Thus, if the duplicated genes are retained in the genome over evolutionary timescales either changes in one or both copies must occur for subfunctionalization of the encoded proteins, or regulation of the genes must have diverged (Gu et al., 2005). Obviously, the latter is true for the GDH-encoding genes in B. subtilis. As mentioned above, while the gudB gene is constitutively transcribed, rocG expression strongly depends on the availability of amino acids such as arginine. It is likely to assume that the redundancy of the GDH-encoding genes and their differential regulation provide the bacteria with a selective growth advantage under specific growth conditions. Indeed, high-level GDH activity is required for the efficient utilization of amino acids of the glutamate family (e.g. arginine) and the simultaneous inhibition of glutamate biosynthesis, which is not required under these growth conditions (see Fig. 1). The present study also revealed that high-level GDH activity results in high-level expression of the gltAB genes

(Table 1; Belitsky and Sonenshein, 2004; Gunka et al., 2010). These observations seem to be somewhat contradictory because one would expect that overproduction of the negative effector of GltC, which can be either RocG or GudB, or probably both GDHs in a complex, must cause permanent inhibition of the transcription factor. The deregulation of glutamate biosynthesis in bacteria endowed with high level of GDH activity might indicate that the GDHs need a co-factor for being able to control the DNA-binding activity of GltC. The fact that the addition of arginine rescued the inhibitory effect of RocG and GudB on gltAB expression indeed suggests that an intermediate derived from the Roc pathway and the GDHs synergistically control GltC activity (Table 1). The metabolome analysis in the present study revealed a strong inverse correlation between activity of GltC and the intracellular glutamate pool. As shown in Fig. 5, despite the presence of the GDH RocG, the transcription factor GltC was highly active when the intracellular amount of glutamate was low. In contrast, the DNAbinding activity of GltC was severely reduced when the cellular amount of glutamate was high. These observations strongly suggest that glutamate is the cofactor of the GDHs allowing them to control GltC activity. There is also genetic evidence supporting this hypothesis (Belitsky and Sonenshein, 2004). It has been shown that GltC is no longer controlled by arginine in a rocA putC mutant strain lacking the enzymes that are needed for the conversion of Δ1-pyrroline-5-carboxylate to glutamate (see Fig. 1). Thus, the GDHs seem to act as glutamate sensors, which, when a certain intracellular glutamate concentration has been reached, prevent glutamate biosynthesis by inhibiting the DNA-binding activity of GltC (Fig. 6). This elegant regulatory circuit enables B. subtilis to tightly adapt glutamate biosynthesis in order to keep the intracellular glutamate pool constant during growth in different nutritional environments.

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

Control of glutamate biosynthesis in Bacillus subtilis The molecular details of the regulatory interaction between the GDHs and the transcription factor GltC should be elaborated in future studies. Co-crystallization experiments or the determination the interaction surface between the enzymes and the transcription factor via chemical cross-linking in combination with mass spectrometry may be valid methodologies.

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(2 μg ml and 25 μg ml respectively). In B. subtilis, amylase activity was detected after growth on plates containing nutrient broth (7.5 g l−1), 17 g Bacto agar l−1 (Becton Dickinson) and 5 g hydrolysed starch l−1 (Carl Roth). Starch degradation was detected by sublimating iodine onto the plates. −1

−1

Construction of B. subtilis mutant strains Experimental procedures Chemicals, media and DNA manipulation Oligonucleotides used in this study that are listed in Table S1 and were purchased from Sigma-Aldrich (Taufkirchen, Germany). Chromosomal DNA was isolated from B. subtilis using the DNeasy Blood & Tissue Kit (Qiagen, Hilden, Germany). Plasmid DNA was isolated using the Nucleospin Extract Kit (Macherey-Nagel, Düren, Germany). DNA fragments that were generated by the polymerase chain reaction (PCR) were purified using the PCR Purification Kit (Qiagen). Phusion DNA polymerase, restriction enzymes and T4 DNA ligase were purchased from Thermo Scientific (Schwerte, Germany) and used according to the manufacturer’s instructions. Miscellaneous chemicals and media were purchased from Sigma-Aldrich, Carl Roth (Karlsruhe, Germany) and Becton-Dickinson (Heidelberg, Germany). Plasmids were sequenced by the SeqLab Sequence Laboratories (Göttingen, Germany).

Bacterial strains and growth conditions All B. subtilis strains used in this work are listed in Table S2. B. subtilis was grown in sporulation (SP) medium, Luria-Bertani (LB) medium or in C minimal medium supplemented with tryptophan (at 50 mg l−1) (Commichau et al., 2007b). CSE medium is C minimal medium supplemented with sodium succinate (6 g l−1) and potassium glutamate (8 g l−1). C-Glc is C minimal medium supplemented with glucose (5 g l−1), and CS is supplemented with sodium succinate (6 g l−1) (Commichau et al., 2007b). Additional sources of carbon and nitrogen were added as indicated. E. coli was grown in LB medium and transformants were selected on plates containing ampicillin (100 μg ml−1). LB, SP and C minimal medium plates were prepared by the addition of 17 g Bacto agar l−1 (BectonDickinson) to LB, SP or CS medium respectively.

DNA manipulation, transformation and phenotypic analysis E. coli DH5α was used for cloning experiments and transformation of E. coli was performed as described previously (Sambrook et al., 1989). E. coli transformants were selected on LB plates containing ampicillin (100 μg ml−1). B. subtilis was transformed with plasmid or chromosomal DNA according to the two-step protocol described previously (Kunst and Rapoport, 1995). Transformants were selected on SP plates containing kanamycin (10 μg ml−1), chloramphenicol (5 μg ml−1), spectinomycin (150 μg ml−1), tetracycline (10 μg ml−1) or erythromycin plus lincomycin

Deletion of the gudB and gltAB genes in the strains BP442 and GP807, respectively, was achieved by transformation with long-flanking homology (LFH) PCR products as described previously (Wach, 1996). The LFH PCR products were constructed using oligonucleotides (see Table S2) to amplify DNA fragments flanking the target genes and intervening kanamycin (aphA3) and tetracycline (tet) antibiotic resistance cassettes from plasmids pDG780 and pDG1514 respectively (Guérout-Fleury et al., 1995).

β-Galactosidase assay Quantitative studies of lacZ expression in B. subtilis were performed as follows: cells were grown at 37°C and 220 r.p.m. in 10–12 ml of C minimal medium supplemented with different carbon and nitrogen sources as indicated. The medium was supplemented in 100 ml shake flasks and the cells were grown without additional aeration. Cells were harvested at an optical density OD600 of 0.6 to 0.8. Specific β-galactosidase activities were determined with cell extracts obtained by lysozyme treatment as described previously (Kunst and Rapoport, 1995). One unit of β-galactosidase is defined as the amount of enzyme which produces 1 nmol of o-nitrophenol per min at 28°C. The BioRad dye-binding assay was used to determine the protein concentrations.

Western blotting For Western blot analyses, the cells were grown as described for the β-galactosidase assay. The proteins were separated by 12.5% SDS-PAGE and transferred onto polyvinylidene difluoride membranes (Bio-Rad, Munich, Germany) by electroblotting. A rabbit anti-RocG (1:15,000) antibody that was shown to recognize RocG as well as GudB served as the primary antibody (Commichau et al., 2007a,b). The antibody was visualized using anti-rabbit immunoglobulin G-alkaline phosphatase secondary antibodies (Promega, Mannheim, Germany) and the CDP-Star detection system (Roche Diagnostics, Basel, Switzerland) as described previously (Commichau et al., 2007b).

Detection of protein–protein interactions The isolation of protein complexes from B. subtilis cells was performed using the SPINE technology (Herzberg et al., 2007). Briefly, the cells were grown at 37°C and 220 r.p.m. in 2.5 l shake flasks supplemented with 500 ml CSE-Glc-Arg medium without additional aeration. To facilitate cross-linking of transiently interaction proteins, the cells were treated for

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

3388 L. Stannek et al. 20 min with the membrane-permeable cross-linker formaldehyde (0.6%, w/v) (Herzberg et al., 2007). The cells were harvested by centrifugation and disrupted using a French Press (18 000 psi, 138 000 kilopascals; Spectronic Instruments, G. Heinemann, Neuss, Germany). The Strep-tagged proteins together with their potential interaction partners were purified from cell-free crude extracts using the StrepTactin : Strep-tag purification system (IBA, Göttingen, Germany) and desthiobiotin (2.5 mM) as eluent. Selected fractions of the SPINE were analysed by silver staining as described previously (Nesterenko et al., 1994), and the proteins were detected and identified by Western blotting and mass spectrometry respectively.

Protein identification by mass spectrometry Silver nitrate-stained gel slices were destained by incubation in 30 mM K3[Fe(CN)6], 100 mM Na2S2O3 until colourless and washed three times in water before processing gel slices as previously described (Thiele et al., 2007). Briefly, gel pieces were washed twice with 200 μl of 20 mM NH4HCO3, 50% (v/v) acetonitrile (ACN) for 30 min at 37°C and dried by adding 200 μl of ACN two times for 15 min. Trypsin solution (10 ng μl−1 trypsin in 20 mM ammonium bicarbonate) was added until gel pieces stopped swelling, and digestion was allowed to proceed for 16–18 h at 37°C. Peptides were extracted from gel pieces by incubation in an ultrasonic bath for 30 min in 40 μl of 0.1% (v/v) acetic acid followed by a second extraction with 40 μl of 50% ACN in 0.05% acetic acid. The supernatants containing peptides were collected, combined, can depleted by evaporation and transferred into microvials for mass spectrometric analysis. Peptides were separated by a nonlinear water–ACN gradient in 0.1% acetic acid on a nanoAcquity UPLC reverse phase column (BEH130, C18, 100 μm × 100 mm, Waters Corporation, Milford, MA, USA) with a nano-UPLC system (Waters) coupled on line with a LTQ (linear trap quadrupole) Orbitrap mass spectrometer (Thermo Electron, Bremen, Germany) operated in data-dependent MS/MS mode. Proteins were identified by searching all MS/MS spectra against a B. subtilis protein database (4254 entries; extracted from SubtiList using SEQUEST version 2.7 rel. 11) (Sorcerer built 4.04, Sage-N Research, Milpitas, CA, USA) (see Tables S3 and S4). Initial mass tolerance for peptide identification on mass spectrometry (MS) and MS/MS peaks were 10 ppm and 1 Da respectively. Up to two missing tryptic cleavages were allowed. Methionine oxidation (+15.99492 Da) and propionamide modification on cysteine (+71.037109 Da) were set as variable modifications. Protein identification results were evaluated by determination of probability for peptide and protein assignments provided by PeptideProphet and ProteinProphet (Institute for Systems Biology, Seattle, WA, USA) incorporated in the Scaffold software package release 4.3.2 (Proteome Software, Portland, OR, USA). Proteins were identified by at least two peptides with minimal peptide scores of XCorr = 2.2 at z = 2 and XCorr = 2.5 at z = 3 and a peptide probability >95% reflecting protein probability of >95%.

Bacterial two-hybrid analysis Primary protein–protein interactions were identified by bacterial two-hybrid analyses (Karimova et al., 1998). The bac-

terial two-hybrid system is based on the interaction-mediated reconstitution of the Bordetella pertussis adenylate cyclase (CyaA) activity in E. coli. Functional complementation between the T18 and T25 fragments of the CyaA protein as a consequence of the interaction between two proteins results in the synthesis of cAMP, which is monitored by measuring the β-galactosidase activity of the cAMPcatabolite activator protein-dependent promoter of the E. coli lac operon. The plasmids p25-N (Claessen et al., 2008) and pKT25 allow the expression of proteins fused to the C- and N-terminus, respectively, of the T25 fragment of the CyaA protein. Plasmids pUT18 and pUT18C allow the expression of proteins fused to the C- and N-terminus, respectively, of the T18 fragment of CyaA (Karimova et al., 1998). The plasmids constructed for the bacterial two-hybrid assay (Table S2) were used for co-transformation of the E. coli strain BTH101 (Δcya), and the protein–protein interactions were then analysed by plating the cells on LB agar plates containing 100 μg ml−1 ampicillin, 50 μg ml−1 kanamycin, 80 μg ml−1 5-bromo-4-chloro-3-indolyl β-D-galactopyranoside (X-Gal, dissolved in N,N-dimethylformamide to a final concentration of 80 mg ml−1) and 0.5 mM isopropyl β-Dthiogalactopyranoside. The plates were incubated for a maximum of 72 h at 30°C.

Analysis of metabolite pools For the metabolite measurements by GC-MS analysis, the bacteria were grown over night in 100 ml shake flasks containing 7 ml CSE-Glc minimal medium supplemented with different arginine concentrations (0.01%, 0.015%, 0.025%, 0.05% and 0.5%) at 37°C and 220 r.p.m. Next day, the cultures were used to inoculate 250 baffled shake flasks containing 60 ml CSE-Glc medium supplemented with the respective arginine concentrations to an OD600 of 0.1. The cells were grown at 37°C without additional aeration. At an OD600 between approximately 0.4 and 0.5, 20 ml of each culture were directly harvested via filtration sampling as described previously (Meyer et al., 2013). In this study, the bacteria were removed from the filters by adding 4 ml extraction solution (cooled 60% ethanol) containing the internal standard (0.0125 mg L-glutamic acid 15N ml−1). Cell disruption was performed by a freeze/thaw cycle as described previously (Meyer et al., 2013). Centrifugation was carried out for 10 min at 4°C and 8000 r.p.m. to remove the filter and the cell fragments. After combining the supernatants of the first and second extraction, the samples were dried under nitrogen stream. The polar fraction was extracted and derivatized with 30 μl methoxyamine hydrochloride and 60 μl N-methyl-N(trimethylsilyl) trifluoroacetamide as previously described (Bellaire et al., 2014) to transform the metabolites into their methoxyimino (MEOX) and trimethylsilyl (TMS) derivatives. The samples were analysed on an Agilent 5973 Network mass selective detector connected to a Agilent 6890 gas chromatograph equipped with a capillary HP5-MS column (30 m × 0.25 mm; 0.25 μm coating thickness; J&W Scientific, Agilent Waldbronn, Germany). Helium was used as carrier gas (1 ml min−1). The inlet temperature was set to 230°C and the temperature gradient applied was 50°C for 2 min, 50–330°C at 5 K min−1 330°C for 2 min. Electron energy of 70 eV, an ion source temperature of 230°C, and a transfer

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390

Control of glutamate biosynthesis in Bacillus subtilis line temperature of 330°C was used. Spectra were recorded in the range of 71–600. For quantification, the ions with the following mass-to-charge ratio were used: 2-OG (1 MEOX, 2 TMS), 198 Da e−1; glutamic acid (3 TMS), 348 glutamic acid15 N (3 TMS), 349; glutamine (3 TMS), 156 Da e−1; ornithine (4 TMS), 142 Da e−1 and arginine [-NH3] (3 TMS), 256 Da e−1. For absolute quantification, glutamate was quantified versus a L-glutamic acid-15N internal standard. All other compounds were quantified against the standard using a calibration curve performed with pure substances based on a total of 15 measurements for each substance.

Acknowledgements The students Dagmar Akkermann, Advait Jagirdir, Dmitrij Rekhter and Lara Schmitz of the lab course ‘Advanced practical training in Microbiology 2013’ of the Göttingen University are acknowledged for the help with some experiments. We are grateful to Ivo Feußner for providing his analytical facility for the determination of metabolite pools. We are grateful to Sabine Lentes and Sabine Freitag for technical assistance. We wish to thank Jörg Stülke for helpful discussions. Maxwell Bain and Bernard Göbel are acknowledged for critical reading of the manuscript. This work was supported by the DFG (Grant No. CO 1139/1-1), the Fonds der Chemischen Industrie (http://www.vci.de/fonds), the Göttingen Centre for Molecular Biology (GZMB) and the Max-BuchnerForschungsstiftung (http://www.dechema.de/mbf.html; MBFSt-Kennziffer 3381).

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Supporting information Additional Supporting Information may be found in the online version of this article at the publisher’s web-site: Fig. S1. Western blot analysis showing constitutive expression of RocG. Table S1. Oligonucleotides used in this study. Table S2. Bacterial strains and plasmids used in this study. Table S3. Number of spectral counts as a measure of abundance of the proteins identified by mass spectrometry in the gel slices (see Fig. 3A). Table S4. Number of spectral counts as a measure of abundance of the proteins identified by mass spectrometry in the gel slices (see Fig. 4A).

© 2015 Society for Applied Microbiology and John Wiley & Sons Ltd, Environmental Microbiology, 17, 3379–3390