Characterization and Use of Catabolite-Repressed Promoters from Gluconate Genes in Corynebacterium glutamicum
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细菌学杂志 2006年第1期
area de Microbiologia, Facultad de Biologia, Universidad de Leon, 24071 Leon, Spain,1 area de Microbiologia, Facultad de Medicina, Universidad de Oviedo, 33006 Oviedo, Spain2
ABSTRACT
The genes involved in gluconate catabolism (gntP and gntK) in Corynebacterium glutamicum are scattered in the chromosome, and no regulatory genes are apparently associated with them, in contrast with the organization of the gnt operon in Escherichia coli and Bacillus subtilis. In C. glutamicum, gntP and gntK are essential genes when gluconate is the only carbon and energy source. Both genes contain upstream regulatory regions consisting of a typical promoter and a hypothetical cyclic AMP (cAMP) receptor protein (CRP) binding region but lack the expected consensus operator region for binding of the GntR repressor protein. Expression analysis by Northern blotting showed monocistronic transcripts for both genes. The expression of gntP and gntK is not induced by gluconate, and the gnt genes are subject to catabolite repression by sugars, such as glucose, fructose, and sucrose, as was detected by quantitative reverse transcription-PCR (qRT-PCR). Specific analysis of the DNA promoter sequences (PgntK and PgntP) was performed using bifunctional promoter probe vectors containing mel (involved in melanin production) or egfp2 (encoding a green fluorescent protein derivative) as the reporter gene. Using this approach, we obtained results parallel to those from qRT-PCR. An applied example of in vivo gene expression modulation of the divIVA gene in C. glutamicum is shown, corroborating the possible use of the gnt promoters to control gene expression. glxR (which encodes GlxR, the hypothetical CRP protein) was subcloned from the C. glutamicum chromosomal DNA and overexpressed in corynebacteria; we found that the level of gnt expression was slightly decreased compared to that of the control strains. The purified GlxR protein was used in gel shift mobility assays, and a specific interaction of GlxR with sequences present on PgntP and PgntK fragments was detected only in the presence of cAMP.
INTRODUCTION
Corynebacterium glutamicum is a nonpathogenic gram-positive soil bacterium of considerable interest to biotechnological industries for its role in amino acid and nucleotide production (46); in addition, C. glutamicum presents an emerging role as a nonpathogenic model for the study of related bacteria with clinical relevance, such as pathogenic corynebacteria (23) or mycobacteria (16). The genera Corynebacterium and Mycobacterium are included in the suborder Corynebacterineae, which belongs to the mycolata, a broad and diverse group of mycolic acid-containing actinomycetes (52). C. glutamicum is able to grow on a variety of carbohydrates and organic acids (such as acetate, lactate, and gluconate) used as single and combined sources of carbon and energy; gluconate is an important carbon source for many microorganisms in their natural environments and is required by Escherichia coli to colonize the mouse large intestine (71). In general, only two specific enzymes are required for gluconate catabolism; extracellular gluconate is incorporated into bacterial cells by a specific gluconate permease (GntP) and phosphorylated to 6-phosphogluconate by a gluconate kinase (GntK) (31). Thus, 6-phosphogluconate can be channeled either through the pentose phosphate pathway or through the Entner-Doudoroff pathway, which is present only in certain microorganisms (11, 59).
In E. coli, GntI is considered to be the main system for gluconate metabolism which comprises (i) an operon with three components, gntR-gntK-gntU, coding for a repressor protein, gluconate kinase, and gluconate permease, respectively (32) and (ii) two additional genes for gluconate permeases (gntP and gntT) scattered throughout the E. coli chromosome (39, 60). In several Bacillus species, gnt genes are clustered and form the gntRKPZ operon, encoding the repressor, gluconate kinase, gluconate permease, and 6-phosphogluconate dehydrogenase, respectively (19, 63, 77). Both systems are transcriptionally regulated, i.e., induced by gluconate and repressed by glucose (9, 58, 60). In E. coli, another subsidiary gluconate system (GntII) has been previously described (30), but it is now considered the catabolic pathway for L-idonate where D-gluconate is an intermediary (4); the idn idonate operon (the former GntII) is repressed by a regulatory protein (IdnR) and globally repressed by the carbon source (5).
Carbon catabolite repression (CCR) is an environment-sensing mechanism used by bacteria for establishing priorities in carbon metabolism. In gram-negative bacteria, the foremost component of transcriptional control is cyclic AMP (cAMP) receptor protein (CRP), which acts as a global transcriptional activator in the presence of cAMP (8). The CCR mechanism in low-G+C-content gram-positive bacteria is different and is not based on the presence of a typical CRP protein able to bind or respond to cAMP (65). In this group of bacteria, three elements are crucial to CCR (68): CcpA (carbon catabolite protein), HPr (transcriptional regulator), and CRE (catabolite-responsive element). The location of the CRE site within a given promoter appears to determine whether CcpA functions as an activator when CRE is located upstream of the promoter or as a repressor when CRE is overlapping or downstream of the promoter (48).
Many global and specific transcriptional regulators involved in gene expression have been described for bacteria. In C. glutamicum, only the following six transcriptional regulators have been deeply analyzed during the last couple of years: the negative transcriptional regulators McbR (64), RamB (24), AcnR (41), and HspR (18); the transcriptional activator ClgR (17, 18); and GlxR, which is involved in modulating expression of aceB (codes for malate synthase). GlxR shares common features with the CRP from E. coli, such as a 27% identity in amino acid sequence and the presence of a cAMP-binding motif where cAMP is the modulating agent involved in gene expression; heterologous glxR can complement E. coli CRP mutants (37). In most cases, inactivation of the aforementioned regulators leads to a different expression pattern of the target gene, except in the case of GlxR, where glxR mutants have not been obtained. In addition, several C. glutamicum genes are transcriptionally regulated by a dual mechanism: GlxR and RamB regulate aceB (24, 37), and the aconitase gene acn is regulated by AcnR plus at least one more system (41).
Gene-modulated systems are powerful tools for studying gene function and for the validation of drug targets in bacteria (14). In E. coli and Bacillus, there are useful inducible and repressible systems based on Plac, Ptac, or phage promoters (15) to assess the effect of the expression or depletion of cloned gene products; it is also possible to modulate the expression of the associated genes using regulated promoters based on the arabinose operon from E. coli (26, 50) or the xylose operon from Bacillus subtilis (38). Unfortunately, there are not efficient systems available to modulate gene expression in corynebacteria, and systems based on Plac (lactose) or Pbad (arabinose) from E. coli or Pxyl (xylose) from Bacillus do not work proficiently in corynebacteria, due mainly to the inability to metabolize those carbon sources (6, 62). Owing to the huge biotechnological importance of C. glutamicum, its genome has been sequenced by different groups and is now available (29, 34). Our interest in gluconate metabolism in C. glutamicum arose from results obtained when we were studying the nonspecific integration of suicide plasmids in the gntP gene of C. glutamicum (47) and continues presently because the regulatory properties of the gnt promoters may be a way to control expression of cell division genes in corynebacteria (61, 62).
In this work, we analyzed the genes involved in gluconate metabolism with special emphasis on their promoters. Herein, we provide evidence of the absence of a gntR-like gene in the genome of C. glutamicum and suggest that the expression of the gntK and gntP genes is governed by a global regulatory mechanism responsive to the carbon source. In addition, an applied example based on divIVA gene expression using the modulated gnt promoters is shown.
MATERIALS AND METHODS
Bacterial strains, plasmids, and culture conditions. The bacterial strains and plasmids used in this study are listed in Table 1. E. coli strains were grown on Luria-Bertani complex medium (27) or VB minimal medium (74) at 37°C. Corynebacterial strains were grown at 30°C in Trypticase soy broth from Oxoid (TSB complex medium without glucose), TSA (TSB supplemented with 2% agar), and minimal medium (MM) for corynebacteria lacking a carbon source (35) and supplemented with either 1% glucose (MMG), 1% gluconate (MMGn), 1% acetate (MMA), 1% fructose (MMF), or 1% sucrose (MMS). Where specified, TSA, TSB, or MM was supplemented with different quantities (up to 4%) of sugars and/or organic acids. When required, the following supplements were added to the culture media: kanamycin (50 μg/ml for E. coli; 25 μg/ml for corynebacteria), ampicillin (100 μg/ml), and chloramphenicol (100 μg/ml for E. coli; 10 μg/ml for corynebacteria). For melanin production in corynebacteria, TSA was supplemented with tyrosine (8.2 mM) and cupric sulfate (2 mM) added after sterilization. All cultures were grown in Erlenmeyer flasks at 30°C (for Corynebacterium strains) or 37°C (for E. coli) at 250 rpm. Corynebacterial cells were harvested at an optical density at 600 nm (OD600) of 1.5 for conjugation assays and for RNA isolation.
DNA/RNA manipulations and analyses. Plasmid DNA isolation, restriction endonuclease digestion, agarose gel electrophoresis, Southern blot analysis, and molecular cloning were performed by using standard procedures (36). PCR products in this work were generated by Pfu (for expression) and Taq DNA polymerase (Stratagene) using the C. glutamicum ATCC 13032 genome as the template and primers listed in Table 2. E. coli cells were transformed by the method of Hanahan (27). Mobilization of plasmids from E. coli to coryneform strains was accomplished as described previously (47).
DNA labeling and hybridization for Southern blotting was performed using a digoxigenin (DIG) High Prime kit from Roche (Mannheim, Germany) according to the manufacturer's specifications. DNA labeling and detection for gel-retarding analysis was performed using a second-generation DIG gel shift kit from Roche (Mannheim, Germany). PCR-amplified DNA was always sequenced when it was used for expression.
RNA for Northern blotting, reverse transcription-PCR (RT-PCR), quantitative RT-PCR (qRT-PCR), and rapid amplification of cDNA ends (RACE) analysis was isolated from C. glutamicum strains grown in TSB or in MM containing different metabolizable carbon sources (acetate, gluconate, glucose, fructose, and sucrose) using an RNeasy commercial kit (QIAGEN); for Northern blots, 15-μg portions of total cellular RNA were denatured by incubation for 10 min at 65°C in formaldehyde-MOPS (morpholinepropanesulfonic acid) gel-loading buffer, electrophoresed through a 1.5% agarose gel containing formaldehyde-MOPS buffer, and then transferred to nylon membranes (Hybond; Amersham). Filters were hybridized with a 600-bp PstI-BglII internal fragment of the gntP gene (Fig. 1A) or with the whole gntK gene (740 bp) (Table 2; Fig. 1B) obtained by PCR amplification using gnt3/gnt4 probes. Probes were labeled by nick translation (Amersham) using [-32P]dCTP (6,000 Ci/mmol; Amersham) and purified by using a Magic-Clean system (Promega). Filter hybridization and film development were accomplished with conventional protocols (36).
For RT-PCR, 1 μg of the isolated RNA was used as the template to generate single-strand cDNA from the gntP and gntK genes with avian myeloblastosis virus reverse transcriptase (first-strand cDNA synthesis kit for RT-PCR; Roche) and primers gnt8 and gnt10, respectively (Table 2; Fig. 1A and B). Primers were designed using the Primer Express software v2.0 (Applied Biosystems) and checked for PCR amplification using C. glutamicum genomic DNA as the template. RNA samples were also tested for contamination with genomic DNA.
For qRT-PCR, 1/20 of each RT-PCR product was used as the template for DNA amplification, using specific primer pairs for each gene (gnt7/gnt8 for gntP and gnt9/gnt10 for gntK [Table 2; Fig. 1A and B]), 12.5 μl of master mix reagent (Applied Biosystems), and up to 25 μl of water. Reactions were performed with an ABI Prism 7000 sequence detection system (Applied Biosystems), and the results were processed using specific software (ABI Prism 7000 SDS software). The results obtained were referred to as the CT (cycle threshold) values. In all cases, the oligonucleotides used in qRT-PCR were designed to have similar melting temperatures (59°C) and to amplify DNA fragments of similar lengths (around 50 nucleotides).
For identification of the transcriptional start site of gnt genes, a 5'-to-3' RACE kit from Roche (Mannheim, Germany) was used. For the assay, 3 μg of RNA was used as the template for specific gntK cDNA production using primer gnt11, and the PCR was performed using the primer pair gnt12/poliT (the latter from the kit); for gntP, cDNA was generated using primer gnt13 and the primer pair gnt14/poliT for the further PCR (Table 2; Fig. 1A and B). The purified PCR products were isolated and cloned into pGEM-TEasy vector, yielding recombinant plasmids pGEM-TK and pGEM-TP (Table 1); these were further sequenced (five of them), and their corresponding transcriptional start sites were determined.
Comparisons of DNA or protein sequences were carried out online (http://www.ebi.ac.uk) using FASTA (57), BLAST (3), and CLUSTAL W (10). Phylogenetic analyses were performed using MEGA3 (molecular evolutionary genetics analysis software) (42) with the neighbor-joining method; this information is shown in the supplemental material.
Construction of recombinant plasmids. gntP was identified in a 4.1-kb sequenced fragment (EMBL databank accession number, AJ296014) obtained from a genomic library of C. glutamicum ATCC 13032 (28) by use of an internal fragment of the gntP gene as a probe (47); the 4.1-kb BamHI fragment was subcloned in pUC18, yielding plasmid pULMV00 (Table 1). A 2.7-kb BamHI-KpnI internal fragment of pULMV00 containing the gntP gene was subcloned into pUC18, yielding the recombinant plasmid pULMV05 (Fig. 1A; Table 1).
gntK was amplified from the C. glutamicum chromosome with gnt3/4 primers, which contained restriction sites for BamHI and HindIII at their 5' ends (Table 2; Fig. 1B). The amplified 740-bp DNA fragment was isolated, digested, and ligated to the BamHI-HindIII-digested pBluescript SK, yielding plasmid pKSK11 (Fig. 1B).
Internal fragments of gntP (600 bp) and gntK (200 bp) were obtained from plasmids pULMV05 (PstI plus BglII digested) and pKSK11 (HaeIII digested and purified the 200-bp fragment), respectively, and ligated to the mobilizable plasmid pK18mob, yielding plasmids pKP1 for gntP and pKMM3 for gntK (Table 1; Fig. 1A and B). Both plasmids are mobilizable and were used to obtain the kanamycin-resistant mutant strains C. glutamicum GNTP and C. glutamicum GNTK (Table 1).
To complement C. glutamicum GNTP, the whole gntP gene was isolated as a 2.7-kb BamHI-KpnI (blunt-ended) fragment from plasmid pULMV05 and ligated to pECM2 (BamHI-SmaI digested) (Table 1), yielding plasmid pECP (Table 1; Fig. 1A). To complement C. glutamicum GNTK, gntK was isolated from plasmid pKSK11 as a 740-bp BamHI-HindIII fragment (the latter target was blunt ended) and cloned into pECM2 (BamHI-SmaI digested), making plasmid pECK (Table 1; Fig. 1B); pECP and pECK were mobilized from E. coli to C. glutamicum, and transconjugants were selected by chloramphenicol resistance.
A promoter probe vector to quantify promoter strength in C. glutamicum was constructed using the egfp2 gene (69) as the reporter as follows. An EcoRI-NdeI fragment containing the xylanase promoter (PxysA) from pXEGFP2 (Table 1) was replaced by the 0.9-kb kan promoter (Pkan) from Tn5 present in plasmid pXHis1-NPro (1), yielding plasmid pKEGFP2 (Table 1). A BglII fragment from pKEGFP2 containing the Pkan-egfp2 gene was subcloned into the bifunctional mobilizable plasmid pEM2, yielding plasmid pEGFP (Fig. 1C). Any promoter with EcoRI-NdeI ends can be introduced into pEGFP by replacement of the EcoRI-NdeI fragment (Pkan). Plasmid pEGNC (Table 1), a Pkan-less pEGFP derivative plasmid obtained by EcoRI-NdeI digestion and blunted by Klenow treatment and ligation was always used as a negative control for fluorescence. C. glutamicum strains containing pEGFP derivatives were grown in liquid media (OD600 = 0.5); samples were then loaded into 96-well microtiter plates, and their fluorescence levels were measured on a Synergy HT multidetection microplate reader fluorimeter (Bio-Tek Instruments, Inc.).
To detect the presence of functional promoters immediately upstream from the gntP and gntK genes, DNA fragments were amplified by PCR using genomic DNA from C. glutamicum and the primer pairs gnt1/gnt2 for PgntP and gnt5/gnt6 for PgntK (Table 2; Fig. 1A and B). The PgntP-amplified fragment (270 bp) was BamHI-NdeI digested and ligated to the E. coli promoter probe vector pJMFA24 (Table 1), yielding plasmid pJMF-MP (Fig. 1A). Similarly, the PgntK-amplified fragment (250 bp) was EcoRI-NdeI digested and ligated to pJMFA24, affording plasmid pJMF-MK (Fig. 1B). PgntP and PgntK were isolated from plasmids pJMF-MP and pJMF-MK, respectively, by EcoRI-NdeI digestion, and DNA fragments were further subcloned into the bifunctional E. coli-C. glutamicum promoter probe plasmids pEMel-1 and pEGFP (Table 1), generating plasmids pEMel-P, pEMel-K, pEGFP-MP, and pEGFP-MK (Table 1; Fig. 1C).
For the cloning of the glxR gene from C. glutamicum, a 684-bp fragment corresponding to the full open reading frame (ORF) was PCR amplified using the primer pair crp3/crp4 (Table 2), digested with NdeI, and subcloned into the NdeI-digested plasmids pNPro and pET-28a (Table 1), yielding plasmids pNProCRP and pETCRP, respectively (see below). pNProCRP was BglII digested, and the 1.5-kb fragment containing the glxR gene under the kanamycin promoter (Pkan-glxR) was subcloned into the bifunctional mobilizable plasmid pEM2 (BamHI digested; Fig. 1C), making plasmid pEMCRP (Table 1), which was further transferred to C. glutamicum.
An internal fragment from glxR (300 bp) was PCR amplified using the primer pair crp1/crp2 (Table 2), and the purified fragment was digested with EcoRI and further subcloned into pK18mob, resulting in plasmid pKCRP (Table 1). Plasmid pKCRP was transferred from E. coli S17-1 to C. glutamicum by conjugation to disrupt the chromosomal copy of glxR.
The presence of a single NdeI site in plasmids pEGFP-MP and pEGFP-MK allowed us to study the expression of any C. glutamicum gene as a translational fusion with EGFP2 (as an NdeI-ended fragment) under the control of PgntP or PgntK. To isolate the divIVA gene as an NdeI fragment, a 1.1-kb PCR product corresponding to the full ORF without a stop codon was obtained from the chromosomal DNA of C. glutamicum ATCC 13032 using the primer pair div1/div2 (Table 2), digested with NdeI, and cloned into the NdeI-digested plasmids pEGFP-MP and pEGFP-MK (Table 1; Fig. 1C), yielding plasmids pEPDG and pEKDG (Table 1), which contained the divIVA-egfp2 translational fusion under the control of PgntP and PgntK, respectively.
Purification of His-tagged GlxR protein in E. coli. Plasmid pETCRP carries the coding sequence of GlxR with an amino-terminal six-His tag label. Purification of the His-tagged protein from IPTG (isopropyl- D-thiogalactopyranoside)-induced (1 mM) E. coli JM109 (DE3)(pETCRP) cells grown at 37°C in LB medium supplemented with 100 μg/ml ampicillin was accomplished according to the standard procedures of the manufacturer (Novagen). When indicated, the six-histidine tag was removed by treatment with thrombin (Novagen) after purification.
EMSA. Three DNA probes obtained from C. glutamicum were used for the electrophoretic mobility gel shift assays (EMSA); two of the probes correspond to the 270-bp PgntP and 250-bp PgntK fragments, and the third was a 250-bp fragment (ParsB1) corresponding to the promoter region of the arsenic detoxification operon from C. glutamicum (53), which was used as a negative control due to the absence of hypothetical CRP-binding sequences. Probes were nonradioactively labeled by use of a DIG High Prime kit from Roche (Mannheim, Germany), and samples were further diluted and dot blot analyzed to optimize the quantity of DNA used in the assay. The binding reaction mixture contained the labeled DNA fragments (around 10 ng), the purified GlxR protein (from 0.1 to 5 μg), 10 mM phosphate buffer, 137 mM NaCl, 2.7 mM KCl, 0.05 μg of poly(dI-dC), and cAMP (0.05 to 0.6 mM) when indicated; 10 μl of the reaction mixture was incubated at 25°C for 30 min and further analyzed on 6% sodium dodecyl sulfate-polyacrylamide (SDS-PAGE) gels in the presence or absence of cAMP. After electrophoresis, gels were blotted to nylon membrane (Hybond) using an electroblotting device (Bio-Rad). For detection, the chemiluminescent substrate CSPD (Amersham) was used and further exposed with X-ray films. Specific binding was confirmed by adding to the reaction mix the same unlabeled DNA probe (100 to 1,000 times).
Microscopic techniques. Fluorescence microscopy of C. glutamicum cells was performed according to protocols described previously (61).
RESULTS
Molecular analysis of the gnt genes from C. glutamicum ATCC 13032. Several years ago, we described the characterization of a transconjugant (C. glutamicum TRA-8) obtained by stable integration of a suicide conjugative plasmid in the gntP gene from C. glutamicum (47). This gene (EMBL databank accession number, AJ296014) was cloned in plasmid pULMV05 (Fig. 1A), and sequence analysis revealed the presence of a 463-amino-acid ORF with significant homology to bacterial gluconate permeases. Once the complete genome sequence of C. glutamicum (NC003450) became available (29), a second gene involved in gluconate metabolism (gntK) was amplified from the C. glutamicum chromosome by PCR; the expected 740-bp PCR DNA fragment was obtained and cloned into pBluescript SK, affording plasmid pKSK11 (Fig. 1B). In both cases, the sequences surrounding gntP and gntK were analyzed in order to identify any other gene that might be involved in gluconate catabolism (especially for the gntR gene, encoding the gluconate repressor), although the search was unsuccessful.
Examination of the DNA sequences located upstream from gntP and gntK revealed the presence of several putative –10 and –35 RNA polymerase-binding sites for C. glutamicum, and therefore it was necessary to characterize the transcription start points (TSPs) for both genes. PCR-amplified fragments were subcloned into pGEM-TEasy by use of a 5'-to-3' RACE kit, and the resulting plasmids, pGEM-TP and pGEM-TK, were sequenced, leading to the identification of TSPs for gntP and gntK (Fig. 1A and B); in both cases, the TSP was a cytosine. The C. glutamicum gntP and gntK –10 boxes are much more conserved than the –35 boxes, compared with the consensus –10 and –35 hexamers described for corynebacteria (TAnaaT and ttGcca, respectively [lowercase letters indicate less conserved nucleotides]) (55, 56); in both cases, the –10 and –35 boxes were separated by 18 nucleotides (Fig. 1A and B). In addition, two putative CRP-binding sites (CBS) were located upstream from gntP (5'-TGTGG-N6-TCTCA-3') and gntK (5'-TGTGA-N6-ACACC-3') (Fig. 1A and B). Both boxes matched the consensus CRP-binding site box (5'-TGTGA-N6-ACACT-3') described previously by de Crombrugghe et al. (12). No putative consensus binding sequences for GntR (5'-ATGTTA-N4-TAACAT-3') or IdnR (5'-ATGTTA-N4-TAACGT-3') from E. coli (5, 60) or for GntR interaction (the dyad symmetry sequence ATACTTGTA) from B. subtilis (22) were found upstream from the gntP or gntK genes. The absence of consensus GntR binding sites upstream from the gnt genes and the failure to find a gntR homolog on the chromosome of C. glutamicum might suggest a nonconventional mechanism for regulation of the gnt genes in this bacterium.
Gene disruption and complementation of the gnt genes. To confirm the participation of gntP and gntK in gluconate catabolism in C. glutamicum, we performed gene disruption experiments using conjugative suicide plasmids containing internal fragments of both gnt genes. Suicide plasmids pKP1 and pKMM3 (Fig. 1A and B) were transferred by conjugation from E. coli to C. glutamicum RES167, and in both cases kanamycin-resistant transconjugants were obtained, suggesting that the disruption of both genes is not lethal. As expected, the transconjugants were able to grow in MMG but not in MMGn (containing gluconate instead of glucose). Transconjugants disrupted either in gntP or in gntK were named C. glutamicum GNTP and C. glutamicum GNTK, respectively. These mutants were able to grow in MMGn when plasmids containing the whole gntP or gntK genes were introduced by conjugation into C. glutamicum GNTP or C. glutamicum GNTK (pECP or pECK, respectively; Fig. 1A and B). Therefore, it is reasonable to point out that the products of the gntP and gntK genes seem to be required for gluconate catabolism in C. glutamicum.
Expression analysis of the gnt genes. Northern analyses were done using RNA obtained from C. glutamicum RES167 grown in minimal media containing 1% acetate (MMA), 1% gluconate (MMGn), or 1% glucose (MMG) or in TSB. The probes used for Northern blotting were the internal PstI-BglII fragment from gntP (600 bp) and the whole gntK gene (740 bp) isolated from pKSK11 (Fig. 1A and B). Figure 2 shows (i) that gntP and gntK are expressed as monocistronic transcripts of 1.5 and 0.6 kb, respectively; (ii) that equivalent expression was observed in samples from MMA and MMGn; and (iii) that little expression (MMG) or no apparent expression (TSB) for either gnt gene indicates the importance of the carbon source in gnt gene expression.
To precisely quantify the expression levels of gntP and gntK under different culture conditions, qRT-PCR was performed with mRNA isolated from C. glutamicum grown in MM supplemented with different carbon sources and concentrations (acetate, gluconate, glucose, fructose, and sucrose). mRNAs from different cultures were used to obtain specific cDNA for gntP and gntK, using primers gnt8 and gnt10, respectively, and further amplified using the primer pairs gnt7/gnt8 and gnt9/gnt10, respectively (Fig. 1A and B). Table 3 summarizes the qRT-PCR data as follows. (i) The expression level of the gntK gene is always higher than that of gntP in the same culture conditions (indicated by lower CT values). (ii) The estimated numbers of specific mRNA molecules for gntK or gntP in cultures containing gluconate or acetate (at 1% or 4%) were equivalent, suggesting a noninducible gnt gene expression in presence of gluconate. (iii) For different nutritional conditions, the scale of expression of gnt genes was as follows (from high to low): acetate/gluconate, glucose, fructose, and sucrose. (iv) The expression of the gnt genes was dependent not only on the catabolized carbon source but also on its concentration in the culture media. (v) The expression levels of the gnt genes in media supplemented with gluconate plus glucose or gluconate plus fructose were slightly lower than the expression levels in glucose or fructose medium lacking gluconate.
Homologous and heterologous analysis of the promoter activity from the gnt genes. Promoter regions of gntP (PgntP; 270 bp) and gntK (PgntK; 250 bp) were PCR amplified and cloned into the E. coli promoter probe vector pJMFA24, which contains the promoterless kan gene as the reporter, yielding plasmids pJMF-MP and pJMF-MK (Fig. 1A and B). E. coli strains containing plasmids pJMF-MP and pJMF-MK were resistant to up to 50 μg/ml of kanamycin. Therefore, the PgntP and PgntK promoters from C. glutamicum are recognized by the E. coli RNA polymerase, like many corynebacterial promoters (55, 56). PgntP and PgntK were also cloned into the bifunctional promoter probe vectors pEMel-1 and pEGFP (Table 1; Fig. 1C) to confirm their promoter activities in corynebacteria. C. glutamicum transconjugants containing plasmids pEMel-P and pEMel-K (Table 1) were grown separately in TSA medium supplemented with different carbon sources and with tyrosine and cupper sulfate for the identification of a dark color in thestrain when the mel operon is expressed. As can be seen in Fig.3A, similar expression levels of mel under the control of PgntP or PgntK were phenotypically observed in the presence of the organic acids gluconate, acetate, and lactate (the latter not shown), regardless of the concentrations used. The strongest repressor effect was caused by sucrose, followed in order of strength of effect by fructose, ribose, glucose, and mannose. In the presence of two carbon sources (combining gluconate, fructose, and glucose), the repressor effect seems to be stronger than that in the presence of individual sugars; alone, these results support those obtained by qRT-PCR analysis but include additional sugars (ribose and mannose). Minor differences were seen, but they could be attributed to the fact that the growth media used in the experiments differed (MM for qRT-PCR and TSA for mel expression).
To monitor the promoter activity of PgntP and PgntK in the presence of different carbon sources using an alternate method, pEGFP derivative plasmids pEGFP-MP and pEGFP-MK were used (Fig. 1C). C. glutamicum(pEGFP-MP) and C. glutamicum(pEGFP-MK) were grown in liquid MM or TSB in the presence of different carbon sources at various concentrations. Plasmid pEGFP, which contains a strong and constitutive promoter (Pkan) not repressed by these carbon sources, was used as a control; the fluorescence value obtained from C. glutamicum(pEGFP) cultured on MM or TSB was set at 100%, and the other data were normalized to this value. The results in Fig. 3B indicate the following. (i) The level of fluorescence from C. glutamicum(pEGFP-MK) (PgntK) was always higher than that from C. glutamicum(pEGFP-MP) (PgntP), independent of the carbon source or growth medium (MM or TSB). (ii) The repressor effect was always 50 to 60% higher in complex medium (TSB) than in defined medium (MM). (iii) Plasmids containing PgntK and PgntP were repressed to the same extent by mannose, glucose, ribose, fructose, and sucrose, with the last exerting the greatest catabolic repression. (iv) The repressor effect was dependent on the concentration of the carbon source (increasing at higher concentrations). (v) The repression effect seems to increase slightly when gluconate and an additional repressor sugar are present in the media.
PgntP or PgntK can be used to modulate the expression of essential cell division genes. The divIVA gene from Brevibacterium lactofermentum (presently known as C. glutamicum [44]) is located downstream from the cell division cluster (dcw), and its product (DivIVA) has a possible function at the growing cell poles of C. glutamicum (61). Overexpression of the translational fusion divIVA-egfp2 (from plasmid pEAG2) in C. glutamicum led to an altered morphology: the cells were rounded, larger, and swollen and tended to grow in a polar fashion, with the DivIVA-EGFP2 product located mainly at the cell poles. It has been suggested that DivIVA participates in the maintenance of cell morphology in C. glutamicum (61).
We constructed plasmids pEPDG and pEKDG (Table 1), in which PgntP and PgntK controlled the expression of the divIVA-egfp2 translational fusion. The plasmids were transferred by conjugation to C. glutamicum ATCC 13032 for comparison with our previous results (61). Transconjugants containing either plasmid were grown in TSB or MM containing different sugars, and the expression levels relative tothe morphological changes were compared to that of divIVA-egfp2 under its own promoter (plasmid pEAG2; Table 1). C. glutamicum(pEAG2) displayed the previously described altered morphology (Fig. 4) independently of the medium used for growth (TSB or MM). However, when C. glutamicum(pEPDG) and C. glutamicum(pEKDG) were grown in TSB, their morphologies were similar to that of the wild type, suggesting low levels of expression of DivIVA-EGFP2 under the control of PgntP or PgntK (Fig. 4). This is supported by the low level of apical fluorescence detected from C. glutamicum(pEPDG) or C. glutamicum(pEKDG).
C. glutamicum(pEPDG) and C. glutamicum(pEKDG) were also grown in MM in the presence of gluconate (1%) or fructose (4%) to control the expression of divIVA-egfp2 under the control of PgntP or PgntK. As can be observed in Fig. 4, larger and swollen cells were observed in the presence of gluconate, but the cell size and the fluorescence level were not affected as much by the presence of fructose. It can also be observed that the fluorescence of the DivIVA-EGFP2 fusion under PgntK was stronger than that under PgntP, confirming our results regarding promoter strength (see above).
Cloning and expression of the glxR gene. As indicated above, no typical binding domains for GntR interaction were found in the promoter region of gntP or gntK; however, consensus sequences corresponding to the CBS described for catabolite repression in gram-negative bacteria were found upstream from gntP and gntK. We then performed a search for a hypothetical crp gene on the genome sequences of C. glutamicum, C. efficiens, and C. diphtheriae (GenBank accession numbers BX927147, BA4035, and BX248353, respectively). We were able to identify a sequence encoding a common 227-amino-acid CRP protein, previously described as the glyoxylate regulator GlxR (37). This protein showed significant homology with putative transcriptional regulators from Mycobacterium tuberculosis and Streptomyces coelicolor and with CRP regulators from E. coli and Vibrio cholerae. To test the possible participation of the GlxR protein on the regulation of the gnt genes, the C. glutamicum crp1 gene (also called the glxR gene herein) was PCR amplified and subcloned into pNPro (pNProCRP), and the BglII cassette was cloned into the bifunctional E. coli-corynebacteria expression vector pEM2 (Fig. 1C), yielding the plasmid pEMCRP, which contains the glxR gene under the kan promoter (Pkan-glxR). With the glxR gene present on a multicopy plasmid, C. glutamicum(pEMCRP) showed delayed growth, in agreement with the results previously published by Kim et al. (37). This strain was grown in MMG (1%) or MMGn (1%), and the expression of gntP and gntK was evaluated by qRT-PCR; the transcription levels of the gnt genes in C. glutamicum(pEMCRP) were slightly lower than those of C. glutamicum(pEM2) and C. glutamicum without plasmid, which were used as CT control values (Table 3).
The suicide plasmid pKCRP, which contains a 300-bp internal fragment from glxR (Table 1), was repeatedly transferred from E. coli to C. glutamicum by conjugation, but kanamycin-resistant transconjugants were never obtained; this result might indicate that glxR is essential for the viability of C. glutamicum, as was described previously (37).
Purification of the GlxR protein and binding analysis to the gnt promoter regions. A His-tagged GlxR protein was purified from E. coli(pETCRP) and analyzed by SDS-PAGE (Fig. 5A); its molecular mass (27 kDa) coincides with the expected size deduced from the gene sequence.
To determine if purified protein (His-tagged GlxR) binds to the gnt promoter regions, PgntP and PgntK were used as probes in gel shift mobility assays. We used the promoter region of the arsenic detoxification operon ParsB1 (53) as the negative control. Starting from the protocol described by Kim et al. (37), we optimized the reaction conditions for each probe and obtained gel-retarding bands only for probes PgntP and PgntK. The presence of retarding bands was directly dependent on the presence of cAMP in the reaction mixture (Fig. 5B); bands were detectable when the cAMP values were between 0.05 and 0.6 mM (and above). No retarding bands were observed in the absence of cAMP or His-tagged GlxR (Fig. 5B). Treating His-tagged GlxR with thrombin to remove the His tag had no effect on binding (data not shown). When the ParsB1 probe was used, no retardation was observed (data not shown). In addition, the binding specificity of GlxR to PgntP or PgntK was confirmed by adding increasing amounts of unlabeled probe (Fig. 5B). No differences were found on the levels of retarding bands from the gel shift assay when cAMP was present or absent in the SDS-PAGE and/or in the electrophoresis buffer.
DISCUSSION
C. glutamicum can metabolize numerous carbon sources, including glucose, fructose, sucrose, ribose, mannose, lactate, acetate, and gluconate. In general, gluconate is incorporated into bacterial cells by a gluconate permease (GntP) and is then converted to 6-phosphogluconate by a gluconate kinase (GntK); 6-phosphogluconate is channeled through the pentose phosphate pathway and through the Entner-Doudoroff pathway, if present (59). Owing to the presence of edd and the absence of eda, which encode the Entner-Doudoroff enzymes 6-phospho-gluconate dehydratase and 2-keto-3-deoxy-6-phosphogluconate-aldolase, respectively, in the genome of C. glutamicum (as it was deduced from the database), it may be assumed that gluconate/6-phosphogluconate is metabolized through the pentose phosphate pathway. To our knowledge, no studies specifically addressing gluconate metabolism in corynebacteria have been conducted, except for those regarding the use of gluconate as a secondary carbon source in nongrowing cells of C. glutamicum (7, 43).
Microorganisms able to metabolize gluconate can contain a typical gnt operon which includes genes for the repressor protein (gntR), for gluconate permease (gntP or the equivalent), and for gluconate kinase (gntK). This organization is present in the E. coli GntI system (72) and in B. subtilis (20), Pseudomonas aeruginosa (accession number, NC002516), Staphylococcus aureus (NC002952), Streptomyces coelicolor A3 (2) (NC003888), and Propionibacterium acnes (NC006085), as shown in Fig. 6. A gnt operon structure lacking gntR is present on the genomes of Neisseria meningitidis (NC003116) and Corynebacterium diphtheriae (NC002935). The most frequent situation is the presence of one or two genes of the gnt system scattered on the chromosome, as occurs in Clostridium perfringens (NC003366), Enterococcus faecalis (NC004668), Haemophilus influenzae (NC007146), Lactobacillus acidophilus (NC006814), Corynebacterium efficiens, and C. glutamicum (Fig. 6).
A second hypothetical gluconate kinase gene (gntK2) was located in a recent sequence analysis of the C. glutamicum and C. efficiens genomes (Fig. 6); comparative analysis of the ORFs showed that the gntK2 gene is larger than gntK (494 and 156 amino acids, respectively) and also showed certain similarities to the gntK gene from Bacillus (513 amino acids); moreover, phylogenetic analysis of bacterial GntK performed with the MEGA3 program displays different associations for gntK and gntK2 (see Fig. S1 in the supplemental material); the former is well positioned close to the actinobacteria, whereas gntK2 (absent in C. diphtheriae) is related to the low-G+C-content gram-positive bacteria. In addition, the evidence obtained from this work suggests that gntK2, if functional, is not involved in gluconate catabolism in C. glutamicum, since C. glutamicum GNTK (a mutant with gntK disrupted) was unable to grow in MM containing gluconate. A similar analysis was performed with gntP (see Fig. S2 in the supplemental material) and gntR (see Fig. S3 in the supplemental material) and showed a high level of conservation (likeness) in the sequences of both genes among the analyzed bacteria (even though in this analysis no specific gntR was found in C. glutamicum or within the Corynebacterineae group).
Northern blotting analysis showed that gntP and gntK are transcribed in C. glutamicum as single monocistronic units of 1.5 and 0.6 kb, respectively. The expression of the gntK and gntP genes is not induced by gluconate, since equivalent levels of expression were obtained when acetate was the only carbon source present in the media (Fig. 2; Table 3). An important repressor effect was observed in the presence of several assimilable carbon sources, such as glucose, fructose, and sucrose (Table 3). The repression caused by sucrose and fructose is the strongest and highest when the sugar concentration is increased (Table 3). In all cases, the basal level of expression of the gntK gene was higher than that in gntP (Fig. 2; Table 3). The repressor effect of glucose or fructose did not revert when gluconate was added to the culture; on the contrary, a slight increase of gnt gene repression was continuously observed by qRT-PCR (Table 3) and by mel (Fig. 3A) and gfp (Fig. 3B) expression analysis. A slight repressor effect was also observed when cells were grown in MMGn containing 4% gluconate rather than 1% (Table 3 and Fig. 3B).
Parallel gene expression results were obtained when C. glutamicum strains containing bifunctional promoter probe vectors with mel and egfp2 under the control of gnt promoters (PgntP and PgntK) were assayed; in addition to the results described above, we observed repression of the reporter geneswhen ribose or mannose was present (Fig. 3A and B). Strains containing plasmids with the fully expressed mel operon are easily visualized by its typical dark color. The assay was performed in complex medium (TSA) (Fig. 3A), since phenotypical differences when cells were grown in MM were not appreciated due to the high level of expression of the mel operon in these culture conditions. In addition, to evaluate the fluorescence levels from cultures containing pEGFP derivatives grown at different conditions, we applied a fast and confident method in which the promoter fusion results were correlated to those obtained by qRT-PCR without the tedious process of mRNA isolation. The data differences seen in these two procedures could be due to the presence of several copies of the promoter probe vector in the cells, which contrasts with the quantification by qRT-PCR of single-copy gene expression.
Several putative regulators with partial similarity to CcpA were found in Streptomyces (73) and Corynebacterium (54). However, C. glutamicum does not have genetic information for HPr kinase/phosphatase, the central enzyme of CCR in low-GC-content gram-positive bacteria (54); therefore, due to the presence of typical CBS upstream from gntP and gntK, we decided to look for CRP-type regulators. We searched the C. glutamicum genome for a putative crp gene using the crp homologues of Mycobacterium and Streptomyces and phylogenetic analysis (see Fig. S4 in the supplementary material). We found that the crp1 gene (CG0350; Fig. 6) was exactly the same as glxR as described by Kim et al. (37) for the strain C. glutamicum ATCC 13059. These authors mentioned the heterologous complementation of E. coli CRP mutants by GlxR protein and suggested a possible interaction of GlxR with other promoters (37). glxR seems to be essential for cell viability, in agreement with the results previously mentioned (37) and in contrast to all transcriptional regulators described so far for C. glutamicum, where the corresponding encoded genes were not essential (24, 41, 64). We indicate the presence (based on database gene alignment) of the crp and ccpA genes located on the chromosome of the above-mentioned bacteria (Fig. 6).
Due to the presence of typical CBS upstream from gntP and gntK, we looked for such sites around the promoter region of C. glutamicum genes shown to be regulated by carbon sources, such as aceA and aceB (37), acn (41), gluA (40), and cspB (70). The aceA and aceB genes coding for isocitrate lyase and malate synthase, respectively, are transcriptionally regulated by the RamB repressor; the effect of RamB was evaluated in media containing glucose (24). In addition, GlxR has been described as a carbon catabolite transcriptional regulator involved in aceB gene expression (37), although an equivalent mechanism is mentioned for aceA. The expression of acn, which codes for aconitase, is modulated by the transcriptional repressor AcnR (41). However, in acnR mutants of C. glutamicum, an increase in aconitase activity (1.5- to 2.3-fold) in cultures grown on acetate, citrate, or propionate compared to the activity of cultures grown on glucose was observed. Therefore, an additional carbon-dependent regulatory mechanism has been suggested (41). gluA is the first gene from the gluABCD operon involved in glutamate uptake in C. glutamicum (40); a repressor effect on glutamate uptake was established when cells were grown in the presence of different carbon sources (such as glucose or fructose) and compared to cells grown on glutamate as the sole carbon source (54). In the case of cspB (coding for the S-layer protein PS2), the catabolite repressor effect was measured by analyzing the amount of PS2 present in cultures grown in the presence of lactose or glucose or by measuring the galactosidase activity from C. glutamicum strains containing the cspB promoter fused to lacZ (PcspB-lacZ) grown in media containing lactate or glucose. Both methods showed a repressive effect of glucose (70). The CBS consensus sequence, TGTGA-N6-ACACT, matched with the analyzed carbon catabolite-regulated promoters from C. glutamicum (Fig. 7). The physical position of CBS on PgntP and PgntK could be a determining factor for the modulation of gene expression, as with CRE sequences (48). In the case of PgntP, the hypothetical CBS is partially included in the –35 box, and therefore its inherent promoter activity could be lower and could strengthen the repressor effect of the sugars (Fig. 1A; Table 3). The contrary may be true for PgntK, where the CBS is located upstream of the –35 box (Fig. 1B; Table 3). Therefore, based on our results for the expression of the gnt genes (Fig. 2 and Table 3) and the amount of EGFP2 produced from gnt promoters (Fig. 3B), we can conclude that gntP is expressed in C. glutamicum at lower levels and is more sensitive to catabolite repression than gntK.
EMSA using purified GlxR from C. glutamicum and PgntP and PgntK showed a specific DNA-GlxR interaction; therefore, the gnt genes would be subject to carbon catabolite regulation, with the binding of GlxR to the promoters mediated by the intermediate regulator, cAMP. In C. glutamicum, high levels of cAMP are present when cells are grown on glucose medium, while low levels are present when cells are grown on acetate (37, 45). The repressor effect observed in C. glutamicum grown in media containing sucrose or fructose was high compared with that seen with other sugars (ribose, glucose, or mannose) or organic acids (acetate, lactate, or gluconate); this could be due to the specific or unspecific mechanism of sugar uptake (25, 49) and/or to the generation of certain glycolytic intermediates. These intermediates could be involved directly or indirectly with cAMP production in catabolite repression. We also observed a higher repressor effect when C. glutamicum was grown in complex medium (TSB) than when it was grown in defined medium (MM) (Fig. 2; Fig. 3A and B). In B. subtilis, the inositol dehydrogenase gene (idn) is repressed by the presence of carbohydrates (51); analogous results, described by Deutscher et al. (13) for a B. subtilis mutant, showed that the inositol dehydrogenase activity was almost completely repressed when the strain was grown in rich medium but only partially repressed when the grown was in MM (23-fold and 4-fold, respectively). We do not know the possible mechanism involved in the increased repressor effect of TSA/TSB medium, but equivalent results were obtained when Luria-Bertani complex medium (lacking glucose) was used. Perhaps the presence of very little sugar in the complex medium would be responsible for a certain repressor effect, but it is more probable that additional unknown factors are involved.
In this work, we describe the fact that gntP and gntK, which are involved in gluconate metabolism in C. glutamicum, are subject to carbon catabolite regulation mediated by cAMP andGlxR; expression of the glxR gene does not seem to be thelimiting factor, as its overexpression [in C. glutamicum(pEMCRP)] decreased the expression of the gnt genes slightly (Table 3). However, the presence of additional regulatory mechanisms involved in gnt gene expression is possible. The promoters of the gluconate system in corynebacteria display a weak to strong repression as a function of the media (complex or minimal) and of the presence of certain sugars (mannose, glucose, ribose, fructose, and sucrose). The effect of catabolic repression is higher when the repressor agents, sugars, are present in large quantities.
The availability of regulated promoters in corynebacteria will allow us to control the expression of essential genes, particularly those studied in basic and medical research. An example of the regulation of an essential gene from C. glutamicum which is involved in cell division is reported in this work (Fig. 4); the overexpression of divIVA from C. glutamicum leads to an aberrant cell morphology. We were able to modify the cell morphology from large and irregular to normal by use of vectors containing the divIVA-egfp2 gene fusion product under the control of the gnt promoters and specific nutritional conditions.
ACKNOWLEDGMENTS
We thank Brett Geissler for the critical revision of the manuscript.
M. Letek, N. Valbuena, E. Ordoez, and A. Ramos were recipients of fellowships from the Junta de Castilla y Leon (E.O. and A.R.) and Ministerio de Educacion y Ciencia (M.L. and N.V.). This work was supported by grants from the Junta de Castilla y Leon (LE 24/01), Diputacion de Leon (2003FMC), Universidad de Leon (ULE-2001-03), and Ministerio de Ciencia y Tecnologia (BIO 2002-03223).
Supplemental material for this article may be found at http://jb.asm.org/.
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ABSTRACT
The genes involved in gluconate catabolism (gntP and gntK) in Corynebacterium glutamicum are scattered in the chromosome, and no regulatory genes are apparently associated with them, in contrast with the organization of the gnt operon in Escherichia coli and Bacillus subtilis. In C. glutamicum, gntP and gntK are essential genes when gluconate is the only carbon and energy source. Both genes contain upstream regulatory regions consisting of a typical promoter and a hypothetical cyclic AMP (cAMP) receptor protein (CRP) binding region but lack the expected consensus operator region for binding of the GntR repressor protein. Expression analysis by Northern blotting showed monocistronic transcripts for both genes. The expression of gntP and gntK is not induced by gluconate, and the gnt genes are subject to catabolite repression by sugars, such as glucose, fructose, and sucrose, as was detected by quantitative reverse transcription-PCR (qRT-PCR). Specific analysis of the DNA promoter sequences (PgntK and PgntP) was performed using bifunctional promoter probe vectors containing mel (involved in melanin production) or egfp2 (encoding a green fluorescent protein derivative) as the reporter gene. Using this approach, we obtained results parallel to those from qRT-PCR. An applied example of in vivo gene expression modulation of the divIVA gene in C. glutamicum is shown, corroborating the possible use of the gnt promoters to control gene expression. glxR (which encodes GlxR, the hypothetical CRP protein) was subcloned from the C. glutamicum chromosomal DNA and overexpressed in corynebacteria; we found that the level of gnt expression was slightly decreased compared to that of the control strains. The purified GlxR protein was used in gel shift mobility assays, and a specific interaction of GlxR with sequences present on PgntP and PgntK fragments was detected only in the presence of cAMP.
INTRODUCTION
Corynebacterium glutamicum is a nonpathogenic gram-positive soil bacterium of considerable interest to biotechnological industries for its role in amino acid and nucleotide production (46); in addition, C. glutamicum presents an emerging role as a nonpathogenic model for the study of related bacteria with clinical relevance, such as pathogenic corynebacteria (23) or mycobacteria (16). The genera Corynebacterium and Mycobacterium are included in the suborder Corynebacterineae, which belongs to the mycolata, a broad and diverse group of mycolic acid-containing actinomycetes (52). C. glutamicum is able to grow on a variety of carbohydrates and organic acids (such as acetate, lactate, and gluconate) used as single and combined sources of carbon and energy; gluconate is an important carbon source for many microorganisms in their natural environments and is required by Escherichia coli to colonize the mouse large intestine (71). In general, only two specific enzymes are required for gluconate catabolism; extracellular gluconate is incorporated into bacterial cells by a specific gluconate permease (GntP) and phosphorylated to 6-phosphogluconate by a gluconate kinase (GntK) (31). Thus, 6-phosphogluconate can be channeled either through the pentose phosphate pathway or through the Entner-Doudoroff pathway, which is present only in certain microorganisms (11, 59).
In E. coli, GntI is considered to be the main system for gluconate metabolism which comprises (i) an operon with three components, gntR-gntK-gntU, coding for a repressor protein, gluconate kinase, and gluconate permease, respectively (32) and (ii) two additional genes for gluconate permeases (gntP and gntT) scattered throughout the E. coli chromosome (39, 60). In several Bacillus species, gnt genes are clustered and form the gntRKPZ operon, encoding the repressor, gluconate kinase, gluconate permease, and 6-phosphogluconate dehydrogenase, respectively (19, 63, 77). Both systems are transcriptionally regulated, i.e., induced by gluconate and repressed by glucose (9, 58, 60). In E. coli, another subsidiary gluconate system (GntII) has been previously described (30), but it is now considered the catabolic pathway for L-idonate where D-gluconate is an intermediary (4); the idn idonate operon (the former GntII) is repressed by a regulatory protein (IdnR) and globally repressed by the carbon source (5).
Carbon catabolite repression (CCR) is an environment-sensing mechanism used by bacteria for establishing priorities in carbon metabolism. In gram-negative bacteria, the foremost component of transcriptional control is cyclic AMP (cAMP) receptor protein (CRP), which acts as a global transcriptional activator in the presence of cAMP (8). The CCR mechanism in low-G+C-content gram-positive bacteria is different and is not based on the presence of a typical CRP protein able to bind or respond to cAMP (65). In this group of bacteria, three elements are crucial to CCR (68): CcpA (carbon catabolite protein), HPr (transcriptional regulator), and CRE (catabolite-responsive element). The location of the CRE site within a given promoter appears to determine whether CcpA functions as an activator when CRE is located upstream of the promoter or as a repressor when CRE is overlapping or downstream of the promoter (48).
Many global and specific transcriptional regulators involved in gene expression have been described for bacteria. In C. glutamicum, only the following six transcriptional regulators have been deeply analyzed during the last couple of years: the negative transcriptional regulators McbR (64), RamB (24), AcnR (41), and HspR (18); the transcriptional activator ClgR (17, 18); and GlxR, which is involved in modulating expression of aceB (codes for malate synthase). GlxR shares common features with the CRP from E. coli, such as a 27% identity in amino acid sequence and the presence of a cAMP-binding motif where cAMP is the modulating agent involved in gene expression; heterologous glxR can complement E. coli CRP mutants (37). In most cases, inactivation of the aforementioned regulators leads to a different expression pattern of the target gene, except in the case of GlxR, where glxR mutants have not been obtained. In addition, several C. glutamicum genes are transcriptionally regulated by a dual mechanism: GlxR and RamB regulate aceB (24, 37), and the aconitase gene acn is regulated by AcnR plus at least one more system (41).
Gene-modulated systems are powerful tools for studying gene function and for the validation of drug targets in bacteria (14). In E. coli and Bacillus, there are useful inducible and repressible systems based on Plac, Ptac, or phage promoters (15) to assess the effect of the expression or depletion of cloned gene products; it is also possible to modulate the expression of the associated genes using regulated promoters based on the arabinose operon from E. coli (26, 50) or the xylose operon from Bacillus subtilis (38). Unfortunately, there are not efficient systems available to modulate gene expression in corynebacteria, and systems based on Plac (lactose) or Pbad (arabinose) from E. coli or Pxyl (xylose) from Bacillus do not work proficiently in corynebacteria, due mainly to the inability to metabolize those carbon sources (6, 62). Owing to the huge biotechnological importance of C. glutamicum, its genome has been sequenced by different groups and is now available (29, 34). Our interest in gluconate metabolism in C. glutamicum arose from results obtained when we were studying the nonspecific integration of suicide plasmids in the gntP gene of C. glutamicum (47) and continues presently because the regulatory properties of the gnt promoters may be a way to control expression of cell division genes in corynebacteria (61, 62).
In this work, we analyzed the genes involved in gluconate metabolism with special emphasis on their promoters. Herein, we provide evidence of the absence of a gntR-like gene in the genome of C. glutamicum and suggest that the expression of the gntK and gntP genes is governed by a global regulatory mechanism responsive to the carbon source. In addition, an applied example based on divIVA gene expression using the modulated gnt promoters is shown.
MATERIALS AND METHODS
Bacterial strains, plasmids, and culture conditions. The bacterial strains and plasmids used in this study are listed in Table 1. E. coli strains were grown on Luria-Bertani complex medium (27) or VB minimal medium (74) at 37°C. Corynebacterial strains were grown at 30°C in Trypticase soy broth from Oxoid (TSB complex medium without glucose), TSA (TSB supplemented with 2% agar), and minimal medium (MM) for corynebacteria lacking a carbon source (35) and supplemented with either 1% glucose (MMG), 1% gluconate (MMGn), 1% acetate (MMA), 1% fructose (MMF), or 1% sucrose (MMS). Where specified, TSA, TSB, or MM was supplemented with different quantities (up to 4%) of sugars and/or organic acids. When required, the following supplements were added to the culture media: kanamycin (50 μg/ml for E. coli; 25 μg/ml for corynebacteria), ampicillin (100 μg/ml), and chloramphenicol (100 μg/ml for E. coli; 10 μg/ml for corynebacteria). For melanin production in corynebacteria, TSA was supplemented with tyrosine (8.2 mM) and cupric sulfate (2 mM) added after sterilization. All cultures were grown in Erlenmeyer flasks at 30°C (for Corynebacterium strains) or 37°C (for E. coli) at 250 rpm. Corynebacterial cells were harvested at an optical density at 600 nm (OD600) of 1.5 for conjugation assays and for RNA isolation.
DNA/RNA manipulations and analyses. Plasmid DNA isolation, restriction endonuclease digestion, agarose gel electrophoresis, Southern blot analysis, and molecular cloning were performed by using standard procedures (36). PCR products in this work were generated by Pfu (for expression) and Taq DNA polymerase (Stratagene) using the C. glutamicum ATCC 13032 genome as the template and primers listed in Table 2. E. coli cells were transformed by the method of Hanahan (27). Mobilization of plasmids from E. coli to coryneform strains was accomplished as described previously (47).
DNA labeling and hybridization for Southern blotting was performed using a digoxigenin (DIG) High Prime kit from Roche (Mannheim, Germany) according to the manufacturer's specifications. DNA labeling and detection for gel-retarding analysis was performed using a second-generation DIG gel shift kit from Roche (Mannheim, Germany). PCR-amplified DNA was always sequenced when it was used for expression.
RNA for Northern blotting, reverse transcription-PCR (RT-PCR), quantitative RT-PCR (qRT-PCR), and rapid amplification of cDNA ends (RACE) analysis was isolated from C. glutamicum strains grown in TSB or in MM containing different metabolizable carbon sources (acetate, gluconate, glucose, fructose, and sucrose) using an RNeasy commercial kit (QIAGEN); for Northern blots, 15-μg portions of total cellular RNA were denatured by incubation for 10 min at 65°C in formaldehyde-MOPS (morpholinepropanesulfonic acid) gel-loading buffer, electrophoresed through a 1.5% agarose gel containing formaldehyde-MOPS buffer, and then transferred to nylon membranes (Hybond; Amersham). Filters were hybridized with a 600-bp PstI-BglII internal fragment of the gntP gene (Fig. 1A) or with the whole gntK gene (740 bp) (Table 2; Fig. 1B) obtained by PCR amplification using gnt3/gnt4 probes. Probes were labeled by nick translation (Amersham) using [-32P]dCTP (6,000 Ci/mmol; Amersham) and purified by using a Magic-Clean system (Promega). Filter hybridization and film development were accomplished with conventional protocols (36).
For RT-PCR, 1 μg of the isolated RNA was used as the template to generate single-strand cDNA from the gntP and gntK genes with avian myeloblastosis virus reverse transcriptase (first-strand cDNA synthesis kit for RT-PCR; Roche) and primers gnt8 and gnt10, respectively (Table 2; Fig. 1A and B). Primers were designed using the Primer Express software v2.0 (Applied Biosystems) and checked for PCR amplification using C. glutamicum genomic DNA as the template. RNA samples were also tested for contamination with genomic DNA.
For qRT-PCR, 1/20 of each RT-PCR product was used as the template for DNA amplification, using specific primer pairs for each gene (gnt7/gnt8 for gntP and gnt9/gnt10 for gntK [Table 2; Fig. 1A and B]), 12.5 μl of master mix reagent (Applied Biosystems), and up to 25 μl of water. Reactions were performed with an ABI Prism 7000 sequence detection system (Applied Biosystems), and the results were processed using specific software (ABI Prism 7000 SDS software). The results obtained were referred to as the CT (cycle threshold) values. In all cases, the oligonucleotides used in qRT-PCR were designed to have similar melting temperatures (59°C) and to amplify DNA fragments of similar lengths (around 50 nucleotides).
For identification of the transcriptional start site of gnt genes, a 5'-to-3' RACE kit from Roche (Mannheim, Germany) was used. For the assay, 3 μg of RNA was used as the template for specific gntK cDNA production using primer gnt11, and the PCR was performed using the primer pair gnt12/poliT (the latter from the kit); for gntP, cDNA was generated using primer gnt13 and the primer pair gnt14/poliT for the further PCR (Table 2; Fig. 1A and B). The purified PCR products were isolated and cloned into pGEM-TEasy vector, yielding recombinant plasmids pGEM-TK and pGEM-TP (Table 1); these were further sequenced (five of them), and their corresponding transcriptional start sites were determined.
Comparisons of DNA or protein sequences were carried out online (http://www.ebi.ac.uk) using FASTA (57), BLAST (3), and CLUSTAL W (10). Phylogenetic analyses were performed using MEGA3 (molecular evolutionary genetics analysis software) (42) with the neighbor-joining method; this information is shown in the supplemental material.
Construction of recombinant plasmids. gntP was identified in a 4.1-kb sequenced fragment (EMBL databank accession number, AJ296014) obtained from a genomic library of C. glutamicum ATCC 13032 (28) by use of an internal fragment of the gntP gene as a probe (47); the 4.1-kb BamHI fragment was subcloned in pUC18, yielding plasmid pULMV00 (Table 1). A 2.7-kb BamHI-KpnI internal fragment of pULMV00 containing the gntP gene was subcloned into pUC18, yielding the recombinant plasmid pULMV05 (Fig. 1A; Table 1).
gntK was amplified from the C. glutamicum chromosome with gnt3/4 primers, which contained restriction sites for BamHI and HindIII at their 5' ends (Table 2; Fig. 1B). The amplified 740-bp DNA fragment was isolated, digested, and ligated to the BamHI-HindIII-digested pBluescript SK, yielding plasmid pKSK11 (Fig. 1B).
Internal fragments of gntP (600 bp) and gntK (200 bp) were obtained from plasmids pULMV05 (PstI plus BglII digested) and pKSK11 (HaeIII digested and purified the 200-bp fragment), respectively, and ligated to the mobilizable plasmid pK18mob, yielding plasmids pKP1 for gntP and pKMM3 for gntK (Table 1; Fig. 1A and B). Both plasmids are mobilizable and were used to obtain the kanamycin-resistant mutant strains C. glutamicum GNTP and C. glutamicum GNTK (Table 1).
To complement C. glutamicum GNTP, the whole gntP gene was isolated as a 2.7-kb BamHI-KpnI (blunt-ended) fragment from plasmid pULMV05 and ligated to pECM2 (BamHI-SmaI digested) (Table 1), yielding plasmid pECP (Table 1; Fig. 1A). To complement C. glutamicum GNTK, gntK was isolated from plasmid pKSK11 as a 740-bp BamHI-HindIII fragment (the latter target was blunt ended) and cloned into pECM2 (BamHI-SmaI digested), making plasmid pECK (Table 1; Fig. 1B); pECP and pECK were mobilized from E. coli to C. glutamicum, and transconjugants were selected by chloramphenicol resistance.
A promoter probe vector to quantify promoter strength in C. glutamicum was constructed using the egfp2 gene (69) as the reporter as follows. An EcoRI-NdeI fragment containing the xylanase promoter (PxysA) from pXEGFP2 (Table 1) was replaced by the 0.9-kb kan promoter (Pkan) from Tn5 present in plasmid pXHis1-NPro (1), yielding plasmid pKEGFP2 (Table 1). A BglII fragment from pKEGFP2 containing the Pkan-egfp2 gene was subcloned into the bifunctional mobilizable plasmid pEM2, yielding plasmid pEGFP (Fig. 1C). Any promoter with EcoRI-NdeI ends can be introduced into pEGFP by replacement of the EcoRI-NdeI fragment (Pkan). Plasmid pEGNC (Table 1), a Pkan-less pEGFP derivative plasmid obtained by EcoRI-NdeI digestion and blunted by Klenow treatment and ligation was always used as a negative control for fluorescence. C. glutamicum strains containing pEGFP derivatives were grown in liquid media (OD600 = 0.5); samples were then loaded into 96-well microtiter plates, and their fluorescence levels were measured on a Synergy HT multidetection microplate reader fluorimeter (Bio-Tek Instruments, Inc.).
To detect the presence of functional promoters immediately upstream from the gntP and gntK genes, DNA fragments were amplified by PCR using genomic DNA from C. glutamicum and the primer pairs gnt1/gnt2 for PgntP and gnt5/gnt6 for PgntK (Table 2; Fig. 1A and B). The PgntP-amplified fragment (270 bp) was BamHI-NdeI digested and ligated to the E. coli promoter probe vector pJMFA24 (Table 1), yielding plasmid pJMF-MP (Fig. 1A). Similarly, the PgntK-amplified fragment (250 bp) was EcoRI-NdeI digested and ligated to pJMFA24, affording plasmid pJMF-MK (Fig. 1B). PgntP and PgntK were isolated from plasmids pJMF-MP and pJMF-MK, respectively, by EcoRI-NdeI digestion, and DNA fragments were further subcloned into the bifunctional E. coli-C. glutamicum promoter probe plasmids pEMel-1 and pEGFP (Table 1), generating plasmids pEMel-P, pEMel-K, pEGFP-MP, and pEGFP-MK (Table 1; Fig. 1C).
For the cloning of the glxR gene from C. glutamicum, a 684-bp fragment corresponding to the full open reading frame (ORF) was PCR amplified using the primer pair crp3/crp4 (Table 2), digested with NdeI, and subcloned into the NdeI-digested plasmids pNPro and pET-28a (Table 1), yielding plasmids pNProCRP and pETCRP, respectively (see below). pNProCRP was BglII digested, and the 1.5-kb fragment containing the glxR gene under the kanamycin promoter (Pkan-glxR) was subcloned into the bifunctional mobilizable plasmid pEM2 (BamHI digested; Fig. 1C), making plasmid pEMCRP (Table 1), which was further transferred to C. glutamicum.
An internal fragment from glxR (300 bp) was PCR amplified using the primer pair crp1/crp2 (Table 2), and the purified fragment was digested with EcoRI and further subcloned into pK18mob, resulting in plasmid pKCRP (Table 1). Plasmid pKCRP was transferred from E. coli S17-1 to C. glutamicum by conjugation to disrupt the chromosomal copy of glxR.
The presence of a single NdeI site in plasmids pEGFP-MP and pEGFP-MK allowed us to study the expression of any C. glutamicum gene as a translational fusion with EGFP2 (as an NdeI-ended fragment) under the control of PgntP or PgntK. To isolate the divIVA gene as an NdeI fragment, a 1.1-kb PCR product corresponding to the full ORF without a stop codon was obtained from the chromosomal DNA of C. glutamicum ATCC 13032 using the primer pair div1/div2 (Table 2), digested with NdeI, and cloned into the NdeI-digested plasmids pEGFP-MP and pEGFP-MK (Table 1; Fig. 1C), yielding plasmids pEPDG and pEKDG (Table 1), which contained the divIVA-egfp2 translational fusion under the control of PgntP and PgntK, respectively.
Purification of His-tagged GlxR protein in E. coli. Plasmid pETCRP carries the coding sequence of GlxR with an amino-terminal six-His tag label. Purification of the His-tagged protein from IPTG (isopropyl- D-thiogalactopyranoside)-induced (1 mM) E. coli JM109 (DE3)(pETCRP) cells grown at 37°C in LB medium supplemented with 100 μg/ml ampicillin was accomplished according to the standard procedures of the manufacturer (Novagen). When indicated, the six-histidine tag was removed by treatment with thrombin (Novagen) after purification.
EMSA. Three DNA probes obtained from C. glutamicum were used for the electrophoretic mobility gel shift assays (EMSA); two of the probes correspond to the 270-bp PgntP and 250-bp PgntK fragments, and the third was a 250-bp fragment (ParsB1) corresponding to the promoter region of the arsenic detoxification operon from C. glutamicum (53), which was used as a negative control due to the absence of hypothetical CRP-binding sequences. Probes were nonradioactively labeled by use of a DIG High Prime kit from Roche (Mannheim, Germany), and samples were further diluted and dot blot analyzed to optimize the quantity of DNA used in the assay. The binding reaction mixture contained the labeled DNA fragments (around 10 ng), the purified GlxR protein (from 0.1 to 5 μg), 10 mM phosphate buffer, 137 mM NaCl, 2.7 mM KCl, 0.05 μg of poly(dI-dC), and cAMP (0.05 to 0.6 mM) when indicated; 10 μl of the reaction mixture was incubated at 25°C for 30 min and further analyzed on 6% sodium dodecyl sulfate-polyacrylamide (SDS-PAGE) gels in the presence or absence of cAMP. After electrophoresis, gels were blotted to nylon membrane (Hybond) using an electroblotting device (Bio-Rad). For detection, the chemiluminescent substrate CSPD (Amersham) was used and further exposed with X-ray films. Specific binding was confirmed by adding to the reaction mix the same unlabeled DNA probe (100 to 1,000 times).
Microscopic techniques. Fluorescence microscopy of C. glutamicum cells was performed according to protocols described previously (61).
RESULTS
Molecular analysis of the gnt genes from C. glutamicum ATCC 13032. Several years ago, we described the characterization of a transconjugant (C. glutamicum TRA-8) obtained by stable integration of a suicide conjugative plasmid in the gntP gene from C. glutamicum (47). This gene (EMBL databank accession number, AJ296014) was cloned in plasmid pULMV05 (Fig. 1A), and sequence analysis revealed the presence of a 463-amino-acid ORF with significant homology to bacterial gluconate permeases. Once the complete genome sequence of C. glutamicum (NC003450) became available (29), a second gene involved in gluconate metabolism (gntK) was amplified from the C. glutamicum chromosome by PCR; the expected 740-bp PCR DNA fragment was obtained and cloned into pBluescript SK, affording plasmid pKSK11 (Fig. 1B). In both cases, the sequences surrounding gntP and gntK were analyzed in order to identify any other gene that might be involved in gluconate catabolism (especially for the gntR gene, encoding the gluconate repressor), although the search was unsuccessful.
Examination of the DNA sequences located upstream from gntP and gntK revealed the presence of several putative –10 and –35 RNA polymerase-binding sites for C. glutamicum, and therefore it was necessary to characterize the transcription start points (TSPs) for both genes. PCR-amplified fragments were subcloned into pGEM-TEasy by use of a 5'-to-3' RACE kit, and the resulting plasmids, pGEM-TP and pGEM-TK, were sequenced, leading to the identification of TSPs for gntP and gntK (Fig. 1A and B); in both cases, the TSP was a cytosine. The C. glutamicum gntP and gntK –10 boxes are much more conserved than the –35 boxes, compared with the consensus –10 and –35 hexamers described for corynebacteria (TAnaaT and ttGcca, respectively [lowercase letters indicate less conserved nucleotides]) (55, 56); in both cases, the –10 and –35 boxes were separated by 18 nucleotides (Fig. 1A and B). In addition, two putative CRP-binding sites (CBS) were located upstream from gntP (5'-TGTGG-N6-TCTCA-3') and gntK (5'-TGTGA-N6-ACACC-3') (Fig. 1A and B). Both boxes matched the consensus CRP-binding site box (5'-TGTGA-N6-ACACT-3') described previously by de Crombrugghe et al. (12). No putative consensus binding sequences for GntR (5'-ATGTTA-N4-TAACAT-3') or IdnR (5'-ATGTTA-N4-TAACGT-3') from E. coli (5, 60) or for GntR interaction (the dyad symmetry sequence ATACTTGTA) from B. subtilis (22) were found upstream from the gntP or gntK genes. The absence of consensus GntR binding sites upstream from the gnt genes and the failure to find a gntR homolog on the chromosome of C. glutamicum might suggest a nonconventional mechanism for regulation of the gnt genes in this bacterium.
Gene disruption and complementation of the gnt genes. To confirm the participation of gntP and gntK in gluconate catabolism in C. glutamicum, we performed gene disruption experiments using conjugative suicide plasmids containing internal fragments of both gnt genes. Suicide plasmids pKP1 and pKMM3 (Fig. 1A and B) were transferred by conjugation from E. coli to C. glutamicum RES167, and in both cases kanamycin-resistant transconjugants were obtained, suggesting that the disruption of both genes is not lethal. As expected, the transconjugants were able to grow in MMG but not in MMGn (containing gluconate instead of glucose). Transconjugants disrupted either in gntP or in gntK were named C. glutamicum GNTP and C. glutamicum GNTK, respectively. These mutants were able to grow in MMGn when plasmids containing the whole gntP or gntK genes were introduced by conjugation into C. glutamicum GNTP or C. glutamicum GNTK (pECP or pECK, respectively; Fig. 1A and B). Therefore, it is reasonable to point out that the products of the gntP and gntK genes seem to be required for gluconate catabolism in C. glutamicum.
Expression analysis of the gnt genes. Northern analyses were done using RNA obtained from C. glutamicum RES167 grown in minimal media containing 1% acetate (MMA), 1% gluconate (MMGn), or 1% glucose (MMG) or in TSB. The probes used for Northern blotting were the internal PstI-BglII fragment from gntP (600 bp) and the whole gntK gene (740 bp) isolated from pKSK11 (Fig. 1A and B). Figure 2 shows (i) that gntP and gntK are expressed as monocistronic transcripts of 1.5 and 0.6 kb, respectively; (ii) that equivalent expression was observed in samples from MMA and MMGn; and (iii) that little expression (MMG) or no apparent expression (TSB) for either gnt gene indicates the importance of the carbon source in gnt gene expression.
To precisely quantify the expression levels of gntP and gntK under different culture conditions, qRT-PCR was performed with mRNA isolated from C. glutamicum grown in MM supplemented with different carbon sources and concentrations (acetate, gluconate, glucose, fructose, and sucrose). mRNAs from different cultures were used to obtain specific cDNA for gntP and gntK, using primers gnt8 and gnt10, respectively, and further amplified using the primer pairs gnt7/gnt8 and gnt9/gnt10, respectively (Fig. 1A and B). Table 3 summarizes the qRT-PCR data as follows. (i) The expression level of the gntK gene is always higher than that of gntP in the same culture conditions (indicated by lower CT values). (ii) The estimated numbers of specific mRNA molecules for gntK or gntP in cultures containing gluconate or acetate (at 1% or 4%) were equivalent, suggesting a noninducible gnt gene expression in presence of gluconate. (iii) For different nutritional conditions, the scale of expression of gnt genes was as follows (from high to low): acetate/gluconate, glucose, fructose, and sucrose. (iv) The expression of the gnt genes was dependent not only on the catabolized carbon source but also on its concentration in the culture media. (v) The expression levels of the gnt genes in media supplemented with gluconate plus glucose or gluconate plus fructose were slightly lower than the expression levels in glucose or fructose medium lacking gluconate.
Homologous and heterologous analysis of the promoter activity from the gnt genes. Promoter regions of gntP (PgntP; 270 bp) and gntK (PgntK; 250 bp) were PCR amplified and cloned into the E. coli promoter probe vector pJMFA24, which contains the promoterless kan gene as the reporter, yielding plasmids pJMF-MP and pJMF-MK (Fig. 1A and B). E. coli strains containing plasmids pJMF-MP and pJMF-MK were resistant to up to 50 μg/ml of kanamycin. Therefore, the PgntP and PgntK promoters from C. glutamicum are recognized by the E. coli RNA polymerase, like many corynebacterial promoters (55, 56). PgntP and PgntK were also cloned into the bifunctional promoter probe vectors pEMel-1 and pEGFP (Table 1; Fig. 1C) to confirm their promoter activities in corynebacteria. C. glutamicum transconjugants containing plasmids pEMel-P and pEMel-K (Table 1) were grown separately in TSA medium supplemented with different carbon sources and with tyrosine and cupper sulfate for the identification of a dark color in thestrain when the mel operon is expressed. As can be seen in Fig.3A, similar expression levels of mel under the control of PgntP or PgntK were phenotypically observed in the presence of the organic acids gluconate, acetate, and lactate (the latter not shown), regardless of the concentrations used. The strongest repressor effect was caused by sucrose, followed in order of strength of effect by fructose, ribose, glucose, and mannose. In the presence of two carbon sources (combining gluconate, fructose, and glucose), the repressor effect seems to be stronger than that in the presence of individual sugars; alone, these results support those obtained by qRT-PCR analysis but include additional sugars (ribose and mannose). Minor differences were seen, but they could be attributed to the fact that the growth media used in the experiments differed (MM for qRT-PCR and TSA for mel expression).
To monitor the promoter activity of PgntP and PgntK in the presence of different carbon sources using an alternate method, pEGFP derivative plasmids pEGFP-MP and pEGFP-MK were used (Fig. 1C). C. glutamicum(pEGFP-MP) and C. glutamicum(pEGFP-MK) were grown in liquid MM or TSB in the presence of different carbon sources at various concentrations. Plasmid pEGFP, which contains a strong and constitutive promoter (Pkan) not repressed by these carbon sources, was used as a control; the fluorescence value obtained from C. glutamicum(pEGFP) cultured on MM or TSB was set at 100%, and the other data were normalized to this value. The results in Fig. 3B indicate the following. (i) The level of fluorescence from C. glutamicum(pEGFP-MK) (PgntK) was always higher than that from C. glutamicum(pEGFP-MP) (PgntP), independent of the carbon source or growth medium (MM or TSB). (ii) The repressor effect was always 50 to 60% higher in complex medium (TSB) than in defined medium (MM). (iii) Plasmids containing PgntK and PgntP were repressed to the same extent by mannose, glucose, ribose, fructose, and sucrose, with the last exerting the greatest catabolic repression. (iv) The repressor effect was dependent on the concentration of the carbon source (increasing at higher concentrations). (v) The repression effect seems to increase slightly when gluconate and an additional repressor sugar are present in the media.
PgntP or PgntK can be used to modulate the expression of essential cell division genes. The divIVA gene from Brevibacterium lactofermentum (presently known as C. glutamicum [44]) is located downstream from the cell division cluster (dcw), and its product (DivIVA) has a possible function at the growing cell poles of C. glutamicum (61). Overexpression of the translational fusion divIVA-egfp2 (from plasmid pEAG2) in C. glutamicum led to an altered morphology: the cells were rounded, larger, and swollen and tended to grow in a polar fashion, with the DivIVA-EGFP2 product located mainly at the cell poles. It has been suggested that DivIVA participates in the maintenance of cell morphology in C. glutamicum (61).
We constructed plasmids pEPDG and pEKDG (Table 1), in which PgntP and PgntK controlled the expression of the divIVA-egfp2 translational fusion. The plasmids were transferred by conjugation to C. glutamicum ATCC 13032 for comparison with our previous results (61). Transconjugants containing either plasmid were grown in TSB or MM containing different sugars, and the expression levels relative tothe morphological changes were compared to that of divIVA-egfp2 under its own promoter (plasmid pEAG2; Table 1). C. glutamicum(pEAG2) displayed the previously described altered morphology (Fig. 4) independently of the medium used for growth (TSB or MM). However, when C. glutamicum(pEPDG) and C. glutamicum(pEKDG) were grown in TSB, their morphologies were similar to that of the wild type, suggesting low levels of expression of DivIVA-EGFP2 under the control of PgntP or PgntK (Fig. 4). This is supported by the low level of apical fluorescence detected from C. glutamicum(pEPDG) or C. glutamicum(pEKDG).
C. glutamicum(pEPDG) and C. glutamicum(pEKDG) were also grown in MM in the presence of gluconate (1%) or fructose (4%) to control the expression of divIVA-egfp2 under the control of PgntP or PgntK. As can be observed in Fig. 4, larger and swollen cells were observed in the presence of gluconate, but the cell size and the fluorescence level were not affected as much by the presence of fructose. It can also be observed that the fluorescence of the DivIVA-EGFP2 fusion under PgntK was stronger than that under PgntP, confirming our results regarding promoter strength (see above).
Cloning and expression of the glxR gene. As indicated above, no typical binding domains for GntR interaction were found in the promoter region of gntP or gntK; however, consensus sequences corresponding to the CBS described for catabolite repression in gram-negative bacteria were found upstream from gntP and gntK. We then performed a search for a hypothetical crp gene on the genome sequences of C. glutamicum, C. efficiens, and C. diphtheriae (GenBank accession numbers BX927147, BA4035, and BX248353, respectively). We were able to identify a sequence encoding a common 227-amino-acid CRP protein, previously described as the glyoxylate regulator GlxR (37). This protein showed significant homology with putative transcriptional regulators from Mycobacterium tuberculosis and Streptomyces coelicolor and with CRP regulators from E. coli and Vibrio cholerae. To test the possible participation of the GlxR protein on the regulation of the gnt genes, the C. glutamicum crp1 gene (also called the glxR gene herein) was PCR amplified and subcloned into pNPro (pNProCRP), and the BglII cassette was cloned into the bifunctional E. coli-corynebacteria expression vector pEM2 (Fig. 1C), yielding the plasmid pEMCRP, which contains the glxR gene under the kan promoter (Pkan-glxR). With the glxR gene present on a multicopy plasmid, C. glutamicum(pEMCRP) showed delayed growth, in agreement with the results previously published by Kim et al. (37). This strain was grown in MMG (1%) or MMGn (1%), and the expression of gntP and gntK was evaluated by qRT-PCR; the transcription levels of the gnt genes in C. glutamicum(pEMCRP) were slightly lower than those of C. glutamicum(pEM2) and C. glutamicum without plasmid, which were used as CT control values (Table 3).
The suicide plasmid pKCRP, which contains a 300-bp internal fragment from glxR (Table 1), was repeatedly transferred from E. coli to C. glutamicum by conjugation, but kanamycin-resistant transconjugants were never obtained; this result might indicate that glxR is essential for the viability of C. glutamicum, as was described previously (37).
Purification of the GlxR protein and binding analysis to the gnt promoter regions. A His-tagged GlxR protein was purified from E. coli(pETCRP) and analyzed by SDS-PAGE (Fig. 5A); its molecular mass (27 kDa) coincides with the expected size deduced from the gene sequence.
To determine if purified protein (His-tagged GlxR) binds to the gnt promoter regions, PgntP and PgntK were used as probes in gel shift mobility assays. We used the promoter region of the arsenic detoxification operon ParsB1 (53) as the negative control. Starting from the protocol described by Kim et al. (37), we optimized the reaction conditions for each probe and obtained gel-retarding bands only for probes PgntP and PgntK. The presence of retarding bands was directly dependent on the presence of cAMP in the reaction mixture (Fig. 5B); bands were detectable when the cAMP values were between 0.05 and 0.6 mM (and above). No retarding bands were observed in the absence of cAMP or His-tagged GlxR (Fig. 5B). Treating His-tagged GlxR with thrombin to remove the His tag had no effect on binding (data not shown). When the ParsB1 probe was used, no retardation was observed (data not shown). In addition, the binding specificity of GlxR to PgntP or PgntK was confirmed by adding increasing amounts of unlabeled probe (Fig. 5B). No differences were found on the levels of retarding bands from the gel shift assay when cAMP was present or absent in the SDS-PAGE and/or in the electrophoresis buffer.
DISCUSSION
C. glutamicum can metabolize numerous carbon sources, including glucose, fructose, sucrose, ribose, mannose, lactate, acetate, and gluconate. In general, gluconate is incorporated into bacterial cells by a gluconate permease (GntP) and is then converted to 6-phosphogluconate by a gluconate kinase (GntK); 6-phosphogluconate is channeled through the pentose phosphate pathway and through the Entner-Doudoroff pathway, if present (59). Owing to the presence of edd and the absence of eda, which encode the Entner-Doudoroff enzymes 6-phospho-gluconate dehydratase and 2-keto-3-deoxy-6-phosphogluconate-aldolase, respectively, in the genome of C. glutamicum (as it was deduced from the database), it may be assumed that gluconate/6-phosphogluconate is metabolized through the pentose phosphate pathway. To our knowledge, no studies specifically addressing gluconate metabolism in corynebacteria have been conducted, except for those regarding the use of gluconate as a secondary carbon source in nongrowing cells of C. glutamicum (7, 43).
Microorganisms able to metabolize gluconate can contain a typical gnt operon which includes genes for the repressor protein (gntR), for gluconate permease (gntP or the equivalent), and for gluconate kinase (gntK). This organization is present in the E. coli GntI system (72) and in B. subtilis (20), Pseudomonas aeruginosa (accession number, NC002516), Staphylococcus aureus (NC002952), Streptomyces coelicolor A3 (2) (NC003888), and Propionibacterium acnes (NC006085), as shown in Fig. 6. A gnt operon structure lacking gntR is present on the genomes of Neisseria meningitidis (NC003116) and Corynebacterium diphtheriae (NC002935). The most frequent situation is the presence of one or two genes of the gnt system scattered on the chromosome, as occurs in Clostridium perfringens (NC003366), Enterococcus faecalis (NC004668), Haemophilus influenzae (NC007146), Lactobacillus acidophilus (NC006814), Corynebacterium efficiens, and C. glutamicum (Fig. 6).
A second hypothetical gluconate kinase gene (gntK2) was located in a recent sequence analysis of the C. glutamicum and C. efficiens genomes (Fig. 6); comparative analysis of the ORFs showed that the gntK2 gene is larger than gntK (494 and 156 amino acids, respectively) and also showed certain similarities to the gntK gene from Bacillus (513 amino acids); moreover, phylogenetic analysis of bacterial GntK performed with the MEGA3 program displays different associations for gntK and gntK2 (see Fig. S1 in the supplemental material); the former is well positioned close to the actinobacteria, whereas gntK2 (absent in C. diphtheriae) is related to the low-G+C-content gram-positive bacteria. In addition, the evidence obtained from this work suggests that gntK2, if functional, is not involved in gluconate catabolism in C. glutamicum, since C. glutamicum GNTK (a mutant with gntK disrupted) was unable to grow in MM containing gluconate. A similar analysis was performed with gntP (see Fig. S2 in the supplemental material) and gntR (see Fig. S3 in the supplemental material) and showed a high level of conservation (likeness) in the sequences of both genes among the analyzed bacteria (even though in this analysis no specific gntR was found in C. glutamicum or within the Corynebacterineae group).
Northern blotting analysis showed that gntP and gntK are transcribed in C. glutamicum as single monocistronic units of 1.5 and 0.6 kb, respectively. The expression of the gntK and gntP genes is not induced by gluconate, since equivalent levels of expression were obtained when acetate was the only carbon source present in the media (Fig. 2; Table 3). An important repressor effect was observed in the presence of several assimilable carbon sources, such as glucose, fructose, and sucrose (Table 3). The repression caused by sucrose and fructose is the strongest and highest when the sugar concentration is increased (Table 3). In all cases, the basal level of expression of the gntK gene was higher than that in gntP (Fig. 2; Table 3). The repressor effect of glucose or fructose did not revert when gluconate was added to the culture; on the contrary, a slight increase of gnt gene repression was continuously observed by qRT-PCR (Table 3) and by mel (Fig. 3A) and gfp (Fig. 3B) expression analysis. A slight repressor effect was also observed when cells were grown in MMGn containing 4% gluconate rather than 1% (Table 3 and Fig. 3B).
Parallel gene expression results were obtained when C. glutamicum strains containing bifunctional promoter probe vectors with mel and egfp2 under the control of gnt promoters (PgntP and PgntK) were assayed; in addition to the results described above, we observed repression of the reporter geneswhen ribose or mannose was present (Fig. 3A and B). Strains containing plasmids with the fully expressed mel operon are easily visualized by its typical dark color. The assay was performed in complex medium (TSA) (Fig. 3A), since phenotypical differences when cells were grown in MM were not appreciated due to the high level of expression of the mel operon in these culture conditions. In addition, to evaluate the fluorescence levels from cultures containing pEGFP derivatives grown at different conditions, we applied a fast and confident method in which the promoter fusion results were correlated to those obtained by qRT-PCR without the tedious process of mRNA isolation. The data differences seen in these two procedures could be due to the presence of several copies of the promoter probe vector in the cells, which contrasts with the quantification by qRT-PCR of single-copy gene expression.
Several putative regulators with partial similarity to CcpA were found in Streptomyces (73) and Corynebacterium (54). However, C. glutamicum does not have genetic information for HPr kinase/phosphatase, the central enzyme of CCR in low-GC-content gram-positive bacteria (54); therefore, due to the presence of typical CBS upstream from gntP and gntK, we decided to look for CRP-type regulators. We searched the C. glutamicum genome for a putative crp gene using the crp homologues of Mycobacterium and Streptomyces and phylogenetic analysis (see Fig. S4 in the supplementary material). We found that the crp1 gene (CG0350; Fig. 6) was exactly the same as glxR as described by Kim et al. (37) for the strain C. glutamicum ATCC 13059. These authors mentioned the heterologous complementation of E. coli CRP mutants by GlxR protein and suggested a possible interaction of GlxR with other promoters (37). glxR seems to be essential for cell viability, in agreement with the results previously mentioned (37) and in contrast to all transcriptional regulators described so far for C. glutamicum, where the corresponding encoded genes were not essential (24, 41, 64). We indicate the presence (based on database gene alignment) of the crp and ccpA genes located on the chromosome of the above-mentioned bacteria (Fig. 6).
Due to the presence of typical CBS upstream from gntP and gntK, we looked for such sites around the promoter region of C. glutamicum genes shown to be regulated by carbon sources, such as aceA and aceB (37), acn (41), gluA (40), and cspB (70). The aceA and aceB genes coding for isocitrate lyase and malate synthase, respectively, are transcriptionally regulated by the RamB repressor; the effect of RamB was evaluated in media containing glucose (24). In addition, GlxR has been described as a carbon catabolite transcriptional regulator involved in aceB gene expression (37), although an equivalent mechanism is mentioned for aceA. The expression of acn, which codes for aconitase, is modulated by the transcriptional repressor AcnR (41). However, in acnR mutants of C. glutamicum, an increase in aconitase activity (1.5- to 2.3-fold) in cultures grown on acetate, citrate, or propionate compared to the activity of cultures grown on glucose was observed. Therefore, an additional carbon-dependent regulatory mechanism has been suggested (41). gluA is the first gene from the gluABCD operon involved in glutamate uptake in C. glutamicum (40); a repressor effect on glutamate uptake was established when cells were grown in the presence of different carbon sources (such as glucose or fructose) and compared to cells grown on glutamate as the sole carbon source (54). In the case of cspB (coding for the S-layer protein PS2), the catabolite repressor effect was measured by analyzing the amount of PS2 present in cultures grown in the presence of lactose or glucose or by measuring the galactosidase activity from C. glutamicum strains containing the cspB promoter fused to lacZ (PcspB-lacZ) grown in media containing lactate or glucose. Both methods showed a repressive effect of glucose (70). The CBS consensus sequence, TGTGA-N6-ACACT, matched with the analyzed carbon catabolite-regulated promoters from C. glutamicum (Fig. 7). The physical position of CBS on PgntP and PgntK could be a determining factor for the modulation of gene expression, as with CRE sequences (48). In the case of PgntP, the hypothetical CBS is partially included in the –35 box, and therefore its inherent promoter activity could be lower and could strengthen the repressor effect of the sugars (Fig. 1A; Table 3). The contrary may be true for PgntK, where the CBS is located upstream of the –35 box (Fig. 1B; Table 3). Therefore, based on our results for the expression of the gnt genes (Fig. 2 and Table 3) and the amount of EGFP2 produced from gnt promoters (Fig. 3B), we can conclude that gntP is expressed in C. glutamicum at lower levels and is more sensitive to catabolite repression than gntK.
EMSA using purified GlxR from C. glutamicum and PgntP and PgntK showed a specific DNA-GlxR interaction; therefore, the gnt genes would be subject to carbon catabolite regulation, with the binding of GlxR to the promoters mediated by the intermediate regulator, cAMP. In C. glutamicum, high levels of cAMP are present when cells are grown on glucose medium, while low levels are present when cells are grown on acetate (37, 45). The repressor effect observed in C. glutamicum grown in media containing sucrose or fructose was high compared with that seen with other sugars (ribose, glucose, or mannose) or organic acids (acetate, lactate, or gluconate); this could be due to the specific or unspecific mechanism of sugar uptake (25, 49) and/or to the generation of certain glycolytic intermediates. These intermediates could be involved directly or indirectly with cAMP production in catabolite repression. We also observed a higher repressor effect when C. glutamicum was grown in complex medium (TSB) than when it was grown in defined medium (MM) (Fig. 2; Fig. 3A and B). In B. subtilis, the inositol dehydrogenase gene (idn) is repressed by the presence of carbohydrates (51); analogous results, described by Deutscher et al. (13) for a B. subtilis mutant, showed that the inositol dehydrogenase activity was almost completely repressed when the strain was grown in rich medium but only partially repressed when the grown was in MM (23-fold and 4-fold, respectively). We do not know the possible mechanism involved in the increased repressor effect of TSA/TSB medium, but equivalent results were obtained when Luria-Bertani complex medium (lacking glucose) was used. Perhaps the presence of very little sugar in the complex medium would be responsible for a certain repressor effect, but it is more probable that additional unknown factors are involved.
In this work, we describe the fact that gntP and gntK, which are involved in gluconate metabolism in C. glutamicum, are subject to carbon catabolite regulation mediated by cAMP andGlxR; expression of the glxR gene does not seem to be thelimiting factor, as its overexpression [in C. glutamicum(pEMCRP)] decreased the expression of the gnt genes slightly (Table 3). However, the presence of additional regulatory mechanisms involved in gnt gene expression is possible. The promoters of the gluconate system in corynebacteria display a weak to strong repression as a function of the media (complex or minimal) and of the presence of certain sugars (mannose, glucose, ribose, fructose, and sucrose). The effect of catabolic repression is higher when the repressor agents, sugars, are present in large quantities.
The availability of regulated promoters in corynebacteria will allow us to control the expression of essential genes, particularly those studied in basic and medical research. An example of the regulation of an essential gene from C. glutamicum which is involved in cell division is reported in this work (Fig. 4); the overexpression of divIVA from C. glutamicum leads to an aberrant cell morphology. We were able to modify the cell morphology from large and irregular to normal by use of vectors containing the divIVA-egfp2 gene fusion product under the control of the gnt promoters and specific nutritional conditions.
ACKNOWLEDGMENTS
We thank Brett Geissler for the critical revision of the manuscript.
M. Letek, N. Valbuena, E. Ordoez, and A. Ramos were recipients of fellowships from the Junta de Castilla y Leon (E.O. and A.R.) and Ministerio de Educacion y Ciencia (M.L. and N.V.). This work was supported by grants from the Junta de Castilla y Leon (LE 24/01), Diputacion de Leon (2003FMC), Universidad de Leon (ULE-2001-03), and Ministerio de Ciencia y Tecnologia (BIO 2002-03223).
Supplemental material for this article may be found at http://jb.asm.org/.
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