Editor: Wilfrid Mitchell
Triple transcriptional control of the resuscitation promoting factor 2 (rpf2) gene of Corynebacterium glutamicum by the regulators of acetate metabolism RamA and RamB and the cAMP-dependent regulator GlxR
Version of Record online: 18 MAR 2008
© 2008 Federation of European Microbiological Societies Published by Blackwell Publishing Ltd
FEMS Microbiology Letters
Volume 281, Issue 2, pages 190–197, April 2008
How to Cite
Jungwirth, B., Emer, D., Brune, I., Hansmeier, N., Pühler, A., Eikmanns, B. J. and Tauch, A. (2008), Triple transcriptional control of the resuscitation promoting factor 2 (rpf2) gene of Corynebacterium glutamicum by the regulators of acetate metabolism RamA and RamB and the cAMP-dependent regulator GlxR. FEMS Microbiology Letters, 281: 190–197. doi: 10.1111/j.1574-6968.2008.01098.x
- Issue online: 18 MAR 2008
- Version of Record online: 18 MAR 2008
- Received 16 October 2007; accepted 16 January 2008.First published online March 2008.
- Corynebacterium glutamicum;
- resuscitation promoting factor;
- actinobacterial signature protein;
- gene regulation;
- carbohydrate metabolism
The transcriptional regulators RamA, RamB and GlxR were detected to bind to the promoter region of the resuscitation promoting factor 2 (rpf2) gene involved in growth and culturability of Corynebacterium glutamicum. DNA-binding sites were identified by bioinformatic analysis and verified by electrophoretic mobility shift assays with purified hexahistidyl-tagged proteins. Carbon source-dependent deregulation of rpf2 expression was demonstrated in vivo in ramA and ramB mutants and in a C. glutamicum strain overexpressing glxR. The deduced network of regulatory interactions provided insights into the complex regulation pattern of rpf2 expression in C. glutamicum.
The resuscitation promoting factor 2 gene (rpf2) is one of two rpf genes in the genome of Corynebacterium glutamicum (Hartmann et al., 2004). The deduced proteins are characterized by a highly conserved Rpf domain with a length of about 70 amino acids and belong to a protein family that is widely spread in Actinobacteria, such as members of the genera Corynebacterium, Mycobacterium and Streptomyces (Voloshin & Kaprelyants, 2004). The Rpf domain has a lysozyme-like fold (Cohen-Gonsaud et al., 2004) and exhibits peptidoglycan-hydrolyzing activity (Mukamolova et al., 2006; Telkov et al., 2006), which is at the origin of the so-called resuscitation promoting activity of Rpf proteins. First observed in Micrococcus luteus (Mukamolova et al., 1998, 2002a), this function is related to the induction of an actively growing state in temporarily nonculturable cells. Thus Rpf proteins act as intercellular bacterial growth factors in dormant cells, which are found as an alternative resting state in many nonsporulating bacteria. Both rpf genes of C. glutamicum are nonessential, but long-stored cells of a rpf double mutant displayed an extended lag phase after transfer to fresh medium (Hartmann et al., 2004). The Rpf2 protein is located on the cell surface of C. glutamicum and exists in three different glycosylated forms in the supernatant of C. glutamicum cultures. A glycosyltransferase encoded by the C. glutamicum gene pmt is essential for glycosylation of Rpf2 (Mahne et al., 2006).
Owing to their role in resuscitation from dormant cell states, Rpf proteins are highly interesting targets in research on tuberculosis (Mukamolova et al., 2002b). It is estimated by the WHO that about one third of the world population carries a latent tuberculosis infection caused by Mycobacterium tuberculosis cells that persist in the human body after an initial infection. At least some of these cells are in a state of dormancy (Mukamolova et al., 2003) and may play a role in the reactivation of the disease in patients with latent tuberculosis. Insights into the regulation of rpf gene expression might provide an indication of the conditions and signalling events leading to this reactivation. The regulation of the rpfA gene of M. tuberculosis, which is controlled by the transcriptional regulator Rv3676, a cAMP receptor protein (CRP) family member, is the only known example of rpf gene regulation so far (Rickman et al., 2005). Corynebacterium glutamicum can serve as an apathogenic model organism for further investigating the regulation of rpf genes because of its close phylogenetic relationship and its well-investigated transcriptional regulatory network (Brinkrolf et al., 2007).
We present here the identification of three transcriptional regulators, RamA, RamB and GlxR, interacting with the DNA in the promoter region of the C. glutamicum rpf2 gene, which is homologous to the rpfB gene of M. tuberculosis (Ravagnani et al., 2005) and classified as a signature protein for almost all sequenced actinobacterial species (Gao et al., 2006). Furthermore, we report on the regulatory connectivity of the three transcriptional regulators in C. glutamicum.
Materials and methods
Escherichia coli JM109 and BL21 cells were used for expression of recombinant proteins and grown at 37 °C in Luria–Bertani (LB) medium (Sambrook et al., 1989) containing 50 μg mL−1 kanamycin for plasmid selection. Corynebacterium glutamicum strains were cultivated at 30 °C in LB medium or minimal medium CGXII (Keilhauer et al., 1993) with 4% (w/v) glucose or 0.4% (w/v) acetate supplementation.
DNA affinity purification assay
Purification of proteins interacting with DNA fragments was performed as described previously (Gabrielsen et al., 1989). Briefly, the biotinylated DNA fragment R-UF (+300 to −32 relative to the rpf2 start codon) was generated by PCR with the primer pair R-UF_bio (biotin-TAGGAATGAGCCGTCGTT) and R-UF2 (CGGTTGATCCGTGACTTC) and was purified using PCR clean-up columns (Qiagen). The amplified DNA fragments were immobilized with Streptavidin coated magnetic beads (Chemagen) using DNA-binding buffer (50 mM Tris, 0.5 mM EDTA, 1 M NaCl; pH 7.5). The magnetic beads were incubated for 60 min at room temperature with C. glutamicum cell lysates prepared in protein-binding buffer (20 mM Tris, 1 mM EDTA, 10% glycerol, 1 mM dithiothreitol, 100 mM NaCl, 0.05% Triton X100; pH 8.0). Proteins were eluted with elution buffer (20 mM Tris, 1 mM EDTA, 10% glycerol, 1 mM dithiothreitol, 1 M NaCl, 0.05% Triton X100; pH 8.0) after three washing steps with protein binding buffer and were finally separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Protein bands were excised from the gel and subjected to tryptic digestion. Matrix-assisted laser desorption/ionization time-of-flight MS (MALDI-TOF MS) and peptide-mass fingerprinting was performed with an Ultraflex III mass spectrometer (Bruker Daltonics). The mascot software (Perkins et al., 1999) was used to compare the peptide mass patterns obtained with those of all proteins from the theoretical C. glutamicum proteome. The molecular weight search (MOWSE) scoring scheme (Pappin et al., 1993) with a cut-off value of 50 was used for unequivocal identification of proteins.
Protein purification and electrophoretic mobility shift assays (EMSAs)
Recombinant versions of RamA, RamB and GlxR were expressed as hexahistidyl-tagged fusion proteins in E. coli and purified by Ni2+ affinity chromatography as described previously (Gerstmeir et al., 2004; Cramer et al., 2006; Letek et al., 2006). A Cy3-labeled DNA fragment for EMSAs was amplified by PCR with the primer pair R-UF_Cy3 (Cy3-TAGGAATGAGCCGTCGTT) and R-UF2 (see above). For EMSAs with the GlxR protein, either 0.03 pmol of Cy3-labeled PCR product and 40 pmol of GlxR protein or 0.05 pmol of Cy3-labeled 40-bp DNA oligomers and 80 pmol of GlxR protein were incubated for 15 min at room temperature in incubation buffer (137 mM NaCl, 2.7 mM KCl, 10 mM NaHPO4) with the addition of 0.24 mM cAMP where appropriate. Samples were separated on 2% agarose gels that were scanned at a wavelength of 530 nm using a Typhoon scanner (Amersham Pharmacia Biotech). EMSAs with RamA and RamB protein preparations were performed as described before (Cramer & Eikmanns, 2007; Cramer et al., 2007).
Rapid amplification of cDNA ends (RACE)-PCR and reverse transcriptase (RT)-PCR assays
Total RNA from C. glutamicum cells for RACE-PCR and RT-PCR was purified as described previously (Brune et al., 2006). RACE-PCR was performed with the 5′/3′ RACE Kit second generation (Roche Diagnostics) according to the manufacturer's instructions using primers SP1 (CCGGAGGTGATGATGTAA) and SP2 (GCGAACGGTCTCCATATC). RACE-PCR products were sequenced by IIT Biotech GmbH (Bielefeld, Germany). Expression analysis of the rpf2 gene of C. glutamicum was performed by real time RT-PCR assays. One-step RT-PCR assays were carried out with the LightCycler instrument (Roche Diagnostics) in combination with the QuantiTect SYBR green RT-PCR kit (Qiagen) and primers rpf2LC1 (TTGGCAGCTCAGAACGTACA) and rpf2LC2 (AGCTGGAGCTTCTGGATCAT). Product verification was performed by melting curve analysis and conventional gel electrophoresis. Differences in gene expression were determined with the lightcycler software (Roche Diagnostics) by comparing the time of appearance of product in two samples, each measured in duplicate.
Identification of the transcriptional regulators RamA, RamB and GlxR binding to the upstream region of the rpf2 gene
To identify transcriptional regulators potentially involved in the control of the C. glutamicum rpf2 gene, a DNA affinity purification assay was performed to enrich regulatory proteins binding specifically to the promoter region. The 332-bp DNA fragment R-UF from the 5′ region of the rpf2 gene (+300 to −32 relative to the start codon) was generated by PCR and incubated with a cytosolic protein extract harvested from a C. glutamicum culture grown in LB complex medium. A number of proteins binding to the DNA fragment of the rpf2 promoter region were observed after separation by SDS-PAGE (Fig. 1). Analysis by MALDI-TOF MS and peptide-mass fingerprinting led to the identification of eight protein bands representing seven different proteins (Table 1). Among the detected proteins were three transcriptional regulators, all involved in the regulation of corynebacterial carbohydrate metabolism. RamB (cg0444) is a member of the HTH_3 family of transcriptional regulators and known as a repressor of genes of acetate metabolism and the glyoxylate cycle (Gerstmeir et al., 2004). The LuxR family protein RamA (cg2831) is also involved in the regulation of acetate metabolism in C. glutamicum (Cramer et al., 2006) and was furthermore shown to activate the expression of the surface-layer gene cspB (Hansmeier et al., 2006). The third transcriptional regulator apparently interacting with the rpf2 promoter region is GlxR (cg0350), so far described as a repressor of genes of gluconate metabolism (Letek et al., 2006) and the glyoxylate cycle (Kim et al., 2004). As a member of the CRP/FNR family, GlxR binds to its DNA targets in a cAMP-dependent manner (Kim et al., 2004).
|Band||Coding region||Gene name||Protein function||MOWSE score||Peptide coverage (detected fragments)|
|1||cg1354||rho||Transcription termination factor||59||26% (20)|
|2||cg1525||polA||DNA polymerase I, α subunit||162||49% (32)|
|3||cg1998||cglIR||Restriction endonuclease||104||27% (17)|
|4||cg0444||ramB||Transcriptional regulator, HTH_3 family||63||29% (12)|
|5||cg1997||cglIIR||Putative restriction enzyme||90||23% (9)|
|6||cg1997||cglIIR||Putative restriction enzyme||101||34% (11)|
|7||cg2831||ramA||Transcriptional regulator, LuxR family||62||17% (8)|
|8||cg0350||glxR||Transcriptional regulator, CRP/FNR family||174||60% (17)|
Validation of the interaction of RamA with the rpf2 promoter region
To confirm the interaction of RamA with the DNA fragment covering the rpf2 promoter region (Fig. 2), EMSAs were carried out. The 332-bp fragment R-UF was amplified by PCR and used as binding partner in EMSAs. The amplicon was incubated with different amounts of recombinant RamA protein and separated on agarose gels (Fig. 3a). The DNA fragment R-UF was completely retarded with 1.0 μg RamA. Lower amounts of RamA also led to the formation of protein-DNA complexes, but retardation of the DNA was not complete in these samples (Fig. 3a). Bioinformatic analysis using the software reputer (Kurtz & Schleiermacher, 1999) and comparison with known binding motifs revealed two RamA binding sites that are located 112–128 bp upstream of the rpf2 start codon (Fig. 2). The genetic organization of the G-rich sequences resembles known tandem motifs for RamA binding (Cramer et al., 2006).
Real time RT-PCR experiments using the wild-type C. glutamicum ATCC 13032 and the ramA deletion mutant C. glutamicum RG2 (Cramer et al., 2006) supported in vivo the observation that rpf2 gene expression is controlled by RamA. The transcription level of the rpf2 gene was reduced 2.5-fold in the ramA deletion mutant during growth on glucose as the sole carbon source, indicating that RamA is a positive regulator of rpf2 gene expression under this growth condition.
Validation of the interaction of RamB with the rpf2 promoter region
To confirm binding of RamB to the promoter region of the rpf2 gene, EMSAs were performed with the DNA fragment R-UF and different amounts of recombinant RamB protein (Fig. 3b). The formation of DNA-protein complexes was observed after incubation of the R-UF fragment with at least 0.5 μg of RamB protein. The DNA fragment was almost completely retarded using 1.0 μg of RamB protein (Fig. 3b). Bioinformatic analysis using the PoSSuMsearch tool of CoryneRegNet (Baumbach et al., 2006) revealed the presence of a RamB binding site in the rpf2 upstream region, overlapping the −10 region of the rpf2 promoter (Fig. 2).
Comparison of rpf2 gene expression by real time RT-PCR experiments in the ramB deletion mutant C. glutamicum RG1 (Gerstmeir et al., 2004) and the wild-type strain additionally revealed an in vivo influence of the regulatory protein. Whereas no difference in gene expression was observed during growth of both strains on glucose, the expression level of rpf2 was elevated 19.6-fold in RG1 cells when acetate was used as the sole carbon source, indicating a negative regulatory role of RamB in rpf2 expression under this culture condition.
Identification of the GlxR-binding motif in the rpf2 promoter region
The interaction of GlxR with the DNA in the rpf2 promoter region was also confirmed by EMSAs using the DNA fragment R-UF as a binding partner. The R-UF fragment was incubated with 40 pmol of His-tagged GlxR protein in the presence or absence of cAMP and then separated on agarose gels (Fig. 3c). The DNA fragment was clearly retarded after incubation with GlxR in the cAMP-containing buffer, confirming the interaction of the protein with the DNA in the rpf2 promoter region in vitro. Bioinformatic searches with the GlxR consensus sequence TGTGA-N6-TCACA revealed the presence of two putative binding sites for GlxR (GlxR-1 and GlxR-2) in this DNA region (Fig. 2). To validate the interaction of GlxR with these motifs, the Cy3-labeled 40-bp fragments 1037_a (containing GlxR-1) and 1037_b (containing GlxR-2) were synthesized and applied in EMSAs (Fig. 3c). Each fragment contained in the center the putative 16-bp binding motif of GlxR flanked by 12-bp native sequences from the genomic context (Fig. 2). The presence of a DNA band with lower mobility after incubation with GlxR and cAMP proved the interaction of the protein with the 40-bp oligomer 1037_a (Fig. 3c). The cAMP-dependent binding of GlxR to the DNA was clearly demonstrated in vitro as no retarded band was observed for the R-UF and 1037_a samples in the absence of cAMP. No significant interaction was detected with the DNA fragment 1037_b and the 40-bp control fragment 1037_c containing native sequences downstream of the rpf2 start codon (Figs 2 and 3c). The detected binding motif GlxR-1 is apparently responsible for the interaction of GlxR with the rpf2 promoter region.
To get experimental data relating to in vivo effects of GlxR on rpf2 expression, we compared transcription levels in C. glutamicum ATCC 13032 and in a derivative carrying the plasmid pEMCRP (Letek et al., 2006) leading to glxR overexpression. In this strain, rpf2 expression was about eightfold higher than in wild-type cells when acetate was the sole carbon source, whereas no significant difference in expression was observed upon growth on glucose.
Regulatory interactions between the transcriptional regulators RamA, RamB and GlxR
The transcriptional regulation of the rpf2 gene observed in vivo is likely to be the outcome of the inter-relation of all three regulatory proteins binding in the upstream region of this gene (Fig. 2). For a better understanding of the regulatory hierarchy, the connectivity between the transcriptional regulators was investigated. A negative autoregulation of RamA upon growth on glucose-containing minimal medium (Cramer & Eikmanns, 2007) as well as a negative autoregulation of RamB and a carbon source-dependent control of ramB by RamA (Cramer et al., 2007) have been described previously. To complete the reconstruction of regulatory interactions, we searched for putative GlxR-binding sites in the promoter regions of the three regulatory genes by means of bioinformatic methods (Kurtz & Schleiermacher, 1999; Baumbach et al., 2006). Two potential 16-bp binding sites located upstream of the −10 region of the ramB promoter (Fig. 4a) and one binding site overlapping the mapped transcription start site and the deduced −10 promoter region of the glxR gene (Fig. 4b) were detected. To test for GlxR interactions with these motifs, EMSAs were performed using 40-bp fragments as binding partners, consisting of the 16-bp motif flanked on both sides by 12 bp of native genomic sequence. In the presence of cAMP, an interaction between GlxR and the predicted binding site GlxR-2 was demonstrated (Fig. 4a). The position of GlxR-2 between the −10 and −35 promoter regions is indicative of a negative regulation of ramB by GlxR. The second potential binding site within the ramB promoter region (GlxR-1) did not interact with the GlxR protein in vitro (data not shown). Moreover, an interaction of GlxR with the putative binding site in the glxR promoter region was shown by the appearance of retarded DNA-protein complexes on the gel (Fig. 4b), suggesting negative autoregulation of the glxR gene.
In the present study we demonstrate that the expression of the C. glutamicum rpf2 gene is under control of the transcriptional regulators RamA, RamB and GlxR (Fig. 5). RamA positively influenced rpf2 expression upon growth on glucose, whereas RamB repressed gene expression during growth on acetate. This is consistent with the role of rpf genes to control growth (and culturability) of corynebacteria with respect to favorable environmental conditions. Growth rates of C. glutamicum are lower on acetate than on glucose (Wendisch et al., 2000), reflecting the fact that acetate is a suboptimal carbon source for the bacterium. Accordingly, rpf2 expression is enhanced when glucose is used as carbon source, triggering a continuous cell wall expansion as would be required during restart of growth after dormancy.
On the other hand, the in vivo influence of GlxR on rpf2 expression cannot be deduced clearly from the current data. The position of the GlxR binding motif and expression levels upon growth on acetate in cells overexpressing the glxR gene indicated that GlxR exhibited a positive effect on rpf2 expression. Nevertheless, a non-physiological effect caused by the multiple copies of glxR might be considered, as GlxR was shown to bind to the DNA in a cAMP-dependent manner. Because the cytosolic cAMP level is elevated in the presence of glucose and low upon growth on acetate (Kim et al., 2004), it is conceivable that GlxR is unable to function as an activator under acetate conditions. However, in cells carrying multiple copies of glxR, the overexpressed protein might capture intracellular cAMP quantitatively and bind to the DNA even under acetate conditions where the cAMP level is very low (Kim et al., 2004). Alternatively, the negative regulation of ramB by GlxR could also explain the observed up-regulation of rpf2 in glxR multicopy cells upon growth on acetate. Interestingly, the rpfA gene of M. tuberculosis is also positively regulated by the GlxR homologue Rv3676, whose binding site is located about 440 bp upstream of the start codon (Rickman et al., 2005). Further investigations with varying levels of glxR expression might help to uncover this aspect of transcriptional regulation in C. glutamicum.
We also elucidated the regulatory interaction between the three transcriptional regulators and deduced a connectivity diagram, consisting of two interconnected feed-forward loops (FFL) and three negative autoregulations (Fig. 5). Computational and experimental studies revealed some general functions of these regulatory network motifs in bacteria (Alon, 2007). Negative autoregulation in combination with efficient promoters generally speeds up the response time of the cell. Likewise, the incoherent FFL of type 1 generated by RamA and RamB may allow an acceleration of the environmental signal and a rapid response of the regulatory system before final adjustment or switch-off of rpf2 gene expression. On the other hand, the coherent FFL of type 4 assembled by GlxR and RamB may sense signal persistency, thereby filtering noise or fluctuations in the environmental input signal (Alon, 2007). Sensing the persistency of a cAMP signal is reasonable for the corynebacterial cell, because the concentration of this effector molecule of GlxR is apparently related to the status of central metabolism and its continuous presence may indicate optimal conditions for growth. A rapid response to changing environmental conditions may then be triggered by the RamA/RamB system, whose possible effector molecules have not been identified so far (Gerstmeir et al., 2004; Cramer et al., 2006). Consequently, the complex regulation pattern of the rpf2 gene by three interconnected transcriptional regulators reflects the outstanding importance for growth of C. glutamicum to optimize rpf2 gene expression under a variety of fluctuating environmental conditions. In order to better understand the physiological relevance of the described regulatory interactions, in vivo experiments will be required, involving genetically modified C. glutamicum strains with altered regulatory gene dosage.
The authors thank Luis M. Mateos for providing pEMCRP and pETCRP, and Miroslav Pátek for providing pEPR1. B.J. acknowledges the receipt of a grant from the Studienstiftung des Deutschen Volkes.
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