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Journal of Bacteriology, October 2007, p. 7408-7416, Vol. 189, No. 20
0021-9193/07/$08.00+0     doi:10.1128/JB.00791-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

The Alcohol Dehydrogenase Gene adhA in Corynebacterium glutamicum Is Subject to Carbon Catabolite Repression

Annette Arndt and Bernhard J. Eikmanns^*

Institute of Microbiology and Biotechnology, University of Ulm, 89069 Ulm, Germany

Received 22 May 2007/ Accepted 4 August 2007

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Corynebacterium glutamicum has recently been shown to grow on ethanol as a carbon and energy source and to possess high alcohol dehydrogenase (ADH) activity when growing on this substrate and low ADH activity when growing on ethanol plus glucose or glucose alone. Here we identify the C. glutamicum ADH gene (adhA), analyze its transcriptional organization, and investigate the relevance of the transcriptional regulators of acetate metabolism RamA and RamB for adhA expression. Sequence analysis of adhA predicts a polypeptide of 345 amino acids showing up to 57% identity with zinc-dependent ADH enzymes of group I. Inactivation of the chromosomal adhA gene led to the inability to grow on ethanol and to the absence of ADH activity, indicating that only a single ethanol-oxidizing ADH enzyme is present in C. glutamicum. Transcriptional analysis revealed that the C. glutamicum adhA gene is monocistronic and that its expression is repressed in the presence of glucose and of acetate in the growth medium, i.e., that adhA expression is subject to catabolite repression. Further analyses revealed that RamA and RamB directly bind to the adhA promoter region, that RamA is essential for the expression of adhA, and that RamB exerts a negative control on adhA expression in the presence of glucose or acetate in the growth medium. However, since the glucose- and acetate-dependent down-regulation of adhA expression was only partially released in a RamB-deficient mutant, there might be an additional regulator involved in the catabolite repression of adhA.

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Corynebacterium glutamicum is a gram-positive, aerobic soil bacterium that is widely used in the biotechnological production of amino acids such as L-glutamate and L-lysine (30, 36). The organism grows on a variety of carbohydrates and organic acids as single or combined sources of carbon and energy, and we recently also showed the growth of C. glutamicum on ethanol (3). This substrate is oxidized via acetaldehyde to acetate, which enters the tricarboxylic acid cycle after activation to acetyl coenzyme A (acetyl-CoA). As in the case of the growth of C. glutamicum with acetate as the sole carbon source, the glyoxylate cycle plays an essential role in the anaplerotic function during growth on ethanol (3). Whereas the acetate-activating enzymes acetate kinase (AK) and phosphotransacetylase (PTA) and, also, the key enzymes of the glyoxylate cycle, isocitrate lyase (ICL) and malate synthase (MS) and their respective genes (ack, pta, aceA, and aceB) have been intensively investigated (reviewed in reference 20), much less is known about the alcohol and acetaldehyde dehydrogenases (ADH and ALDH, respectively), which are involved in ethanol's oxidation to acetate. And although the genome sequence of C. glutamicum has been determined and annotated (26, 29), the genes encoding these two enzymes have not been identified and a functional characterization has not yet been performed.

In contrast to the growth of C. glutamicum on many substrate mixtures, such as glucose plus acetate, lactate, pyruvate, or fructose (7, 12, 52), the growth of this organism on a mixture of glucose and ethanol is biphasic, with glucose consumption in the first and ethanol consumption in the second exponential growth phase (3). This biphasic growth behavior is probably due to relatively low ADH and ALDH activities in the first and much higher ADH and ALDH activities (and thus, high ethanol oxidation activity) in the second growth phase (3). Diauxic growth of C. glutamicum and the sequential utilization of carbon sources have been reported so far only for its growth in medium containing glucose and glutamate, and here the biphasic growth behavior is due to the repression of the glutamate uptake system in the presence of glucose (32, 33). Interestingly, so far there is no evidence for a global carbon catabolite repression system in C. glutamicum, such as the CcpA-dependent carbon control system in Bacillus subtilis and other low-GC, gram-positive bacteria (38, 51) or the cyclic AMP (cAMP)-dependent cAMP receptor protein system in Escherichia coli (6, 45). However, Letek et al. (35) recently reported on the sugar-mediated repression of genes (gntP and gntK) involved in gluconate metabolization in C. glutamicum and showed that GlxR, a cAMP-dependent transcriptional regulator, is involved in this regulation mechanism. However, biphasic growth or the sequential utilization of glucose and gluconate has not been shown for the growth of C. glutamicum in medium containing glucose plus gluconate, and thus, the significance of GlxR for a sequential utilization of two or more carbon sources remains to be investigated.

Recently, we identified two novel regulatory proteins, designated as regulators of acetate metabolism A and B, i.e., RamA and RamB (10, 19). Both proteins were shown to be transcriptional regulators of the AK, PTA, ICL, and MS genes, RamA being an activator in the presence of acetate and RamB being a repressor when C. glutamicum grows on glucose as the sole carbon and energy source. Both regulators are subject to negative autoregulation, and additionally, ramB expression is subject to carbon source-dependent positive control by RamA (8, 9). RamA binds to single or tandem stretches of A/C/TG_4-6T/C or AC_4-5A/G/T, whereas RamB specifically binds to a highly conserved 13-bp motif (AA/GAACTTTGCAAA). Due to their regulatory function as transcriptional regulators of the AK, PTA, ICL, and MS genes and since all these enzymes are also involved in ethanol metabolism, both RamA and RamB are likely candidates to also be involved in the regulatory control of the genes encoding the ethanol-oxidizing enzymes.

In the present study, we identified the ADH gene (adhA), which is involved in the growth of C. glutamicum on ethanol. Furthermore, we analyzed the transcription of adhA and provide evidence for carbon source-dependent transcriptional regulation of this gene. Finally, we investigated the regulatory functions of RamA and RamB in adhA expression control.

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Bacteria, plasmids, oligonucleotides, and culture conditions. Bacterial strains, plasmids and oligonucleotides, as well as their relevant characteristics and sources, are given in Table 1. TY medium (42) was used as complex medium for C. glutamicum and Escherichia coli. The minimal medium used for C. glutamicum has been described previously (14) and contained ethanol, glucose, or acetate or a mixture of those carbon sources in the concentrations indicated in Results. When appropriate, the medium contained kanamycin (50 µg ml^–1 for strains carrying plasmids and 15 µg ml^–1 for integration mutants and for transformant selection). C. glutamicum was grown aerobically at 30°C and E. coli at 37°C in 50-ml cultures in 500-ml baffled Erlenmeyer flasks on a rotary shaker at 120 rpm. The growth of the bacteria was followed by measuring the optical density at 600 nm.

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TABLE 1. Strains, plasmids, and oligonucleotides used in this study

DNA preparation and transformation. The isolation of chromosomal and plasmid DNA from C. glutamicum was performed as described previously (14), and plasmid isolation from E. coli was carried out according to the method of Birnboim (5). DNA transfer into C. glutamicum was performed by electroporation, and recombinant strains were selected on LBHIS agar plates (48). Transformation of E. coli was carried out according to the method of Inoue et al. (27).

PCR techniques. The PCR experiments were performed in a Biometra personal cycler (Biotron) with Taq polymerase (MBI-Fermentas). The oligonucleotides were obtained from MWG-Biotech or from biomers.net GmbH. The cycling times and temperatures were chosen according to fragment length and primer constitution. The PCR products were separated on agarose gels and purified using a Nucleospin extract kit (Macherey Nagel).

DNA manipulation and Southern hybridization. Restriction enzymes, T4 DNA ligase, calf intestinal phosphatase, RNase A, and proteinase K were obtained from MBI-Fermentas and used as instructed by the manufacturer. DNA hybridization experiments were performed as described previously (14). An adhA-specific 880-bp DNA fragment was amplified from the chromosomal DNA of wild-type (WT) C. glutamicum, labeled with digoxigenin-dUTP by PCR with primers adhA-SB-hin and adhA-rev, and used as a probe. The labeling, hybridization, washing, and detection were performed by using a nonradioactive DNA labeling and detection kit from Roche Diagnostics, following the manufacturer's instructions.

Construction of a C. glutamicum adhA integration mutant. The inactivation of the chromosomal adhA gene in C. glutamicum was performed by integration of the vector pDrive into the adhA gene. An internal fragment covering the region between base pair 321 of the 5' end and base pair 208 of the 3' end of the adhA gene was generated by using primers adhA-hin and adhA-rev and ligated via the A/T overhangs into pDrive, and the ligation mixture was transformed into E. coli. The identification of positive E. coli clones was performed by blue/white screening on TY agar plates containing kanamycin, isopropyl-ß-D-thiogalactopyranoside (IPTG; 50 µM), and 5-bromo-4-chloro-3-indolyl-ß-D-galactopyranoside (X-Gal; 80 µg/ml). Recombinant plasmids were isolated from E. coli and electroporated into WT C. glutamicum. The integration of pDrive into the chromosomal adhA locus of the resulting strain, C. glutamicum AA1, was verified by Southern blot analysis. For this purpose, a labeled adhA probe was hybridized to HindIII-restricted and size-fractioned chromosomal DNA from WT C. glutamicum and C. glutamicum AA1. The hybridization resulted in one signal of about 10 kb with DNA from the WT strain and in two signals of about 11 kb and 3.3 kb with DNA from the mutant. According to the restriction map of the adhA locus, these sizes were expected.

Cloning of adhA promoter fragments. The adhAp₁ (401 bp), adhAp₂ (236 bp), and adhAp₃ (213 bp) promoter fragments were amplified from the chromosomal DNA of WT C. glutamicum. The PCR products were digested with SalI and BamHI, ligated into the SalI/BamHI-restricted plasmid pET2, and transformed into E. coli. The recombinant plasmids pET2-adhAp₁, pET2-adhAp₂, and pET2-adhAp₃ were isolated from E. coli and introduced into C. glutamicum.

RNA techniques. For RNA isolation, C. glutamicum cells were grown in minimal medium containing glucose or ethanol, harvested in the exponential growth phase (optical density at 600 nm of about 4.5), and treated with 1 volume of ice-cold killing buffer (20 mM Tris-HCl, pH 8.0, 20 mM NaN₃, 5 mM MgCl₂). The isolation procedure was performed as described previously (43), and aliquots of the RNA were stored at –70°C until use.

For Northern (RNA) hybridization, an 1,149-bp DNA fragment was amplified from the chromosomal DNA of WT C. glutamicum by PCR with primers adhA-SB-hin and adhA-NB-rev, digested with SalI and XbaI, purified with a Nucleospin extract kit, and used as a probe. The [-³²P]dATP labeling of the probe was performed with Klenow fragment (MBI-Fermentas). The separation of unincorporated nucleotides was done by using MicroSpin G-25 columns (Amersham Biosciences). For the hybridization, about 10 µg of total RNA from WT C. glutamicum was separated on an agarose gel containing 17% (vol/vol) formaldehyde and transferred onto a nylon membrane (14). Filter hybridization and detection were accomplished with conventional protocols (42).

The primer extension reactions were carried out as described previously (43) with IRD800-labeled primers CM4 and CM5 and RNA from ethanol-grown C. glutamicum(pET2-adhAp₂) cells, and the primer extension products were analyzed with an automatic sequencer (LI-COR 4000L; Licor, Inc.) using a 6% (wt/vol) polyacrylamide gel at 1,500 V and 50°C. For the exact localization of the transcriptional start site, sequencing reactions using plasmid pET2-adhAp₂ and the same oligonucleotide used for the respective primer extension reaction were coelectrophoresed in the sequencing reaction mixture.

Promoter binding assays with His-tagged RamA and RamB fusion protein. The construction of plasmids for the preparation of His-tagged RamA and RamB and the synthesis and purification of these proteins have been described previously (10, 19). The binding of purified RamA or RamB to the adhA promoter region was tested by DNA electrophoretic mobility shift assays (EMSAs) using the fragments adhAp₁, adhAp₂, and adhAp₃. The EMSA experiments were carried out as described previously (10).

Enzyme assays. For the determination of enzyme activities in cell extracts, C. glutamicum cells were grown in minimal medium containing the respective carbon source(s) and harvested in the exponential growth phase. Cell extracts were prepared as described previously (3).

ADH activity was analyzed in the acetaldehyde-forming direction as described previously (3). Chloramphenicol acetyltransferase (CAT) activity was determined as described by Schreiner et al. (43).

Computational analysis. For the analysis of the G°' value (free energy under standard conditions) of the adhA terminator structures, the program Clone Manager 7 (Sci Ed Central) was used. Databank searches and alignments were carried out using BLAST, CLUSTAL W, and BioEdit software (1, 24, 47).

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Identification and characterization of the adhA gene. We previously found relatively high specific activities of ADH in ethanol-grown cells of C. glutamicum (3). Five genes (cg0273, cg0387, cg0400, cg2714, and cg3107) were annotated as potential ADH genes in the genome of C. glutamicum (GenBank accession numbers NC_003450 and NC_006958) (26, 29). Amino acid sequence alignments of the deduced polypeptides with ADH enzymes from organisms known to utilize ethanol indicated Cg3107 to be the most likely candidate as ADH involved in the ethanol oxidation of C. glutamicum. Cg3107 showed 47% identity to the functionally analyzed ADHs of Pichia stipidis and Saccharomyces cerevisiae, 49% identity to the thermostable ADH of the gram-positive Bacillus stearothermophilus, and 57% identity to the NAD⁺-dependent ADH of the gram-negative Pseudomonas aeruginosa. The other four proteins (Cg0273, Cg0387, Cg0400, and Cg2714) showed less similarity (<40%) to all these ADHs. Therefore, cg3107 was inactivated and the resulting mutant C. glutamicum AA1 was analyzed for growth on ethanol. In fact, this mutant was not able to grow in minimal medium containing ethanol as the single carbon source, whereas its growth on glucose or acetate was not affected (data not shown). For confirmation, the specific ADH activities in cell extracts of WT C. glutamicum and AA1 were determined. In contrast to the WT strain, the mutant did not show any ADH activity on any substrate tested (Table 2). This result and the inability of the mutant to grow on ethanol indicate that cg3107 in fact represents the ADH gene, and therefore, we designated it as adhA.

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TABLE 2. Specific ADH activities of different C. glutamicum strains

The adhA gene of C. glutamicum consists of 1,038 bp and is preceded by a typical ribosomal binding site (AGAAGG). A region of dyad symmetry is located 193 bp downstream of the adhA stop codon, and the mRNA hairpin loop predicted from this sequence has a G°' value of –21.5 kcal/mol at 25°C. This structure indicates transcriptional termination downstream of adhA. According to the nucleotide sequence, adhA encodes a polypeptide of 345 amino acid residues with a predicted molecular mass of 36.8 kDa. Upstream and downstream of adhA, there are genes encoding hypothetical or putative proteins of unknown functions.

Amino acid sequence alignments and BLAST analysis with the C. glutamicum ADH revealed homologous proteins in Corynebacterium efficiens YS-314 (GenBank accession no. CE0053) and Corynebacterium diphtheriae NCTC 13129 (GenBank accession no. DIP2114), showing identities of 72% and 74%, respectively, whereas no significant similarities to proteins of Corynebacterium jeikeium K411 were detected. Further comparisons of the C. glutamicum enzyme with E. coli proteins revealed no significant similarity to the bifunctional AdhE, which physiologically catalyzes ethanol formation from acetyl-CoA (22).

Transcriptional analysis of the adhA gene. To determine the size of the adhA transcript and to test whether the different specific ADH activities observed in C. glutamicum cells grown on either glucose or ethanol are due to transcriptional control, Northern blot hybridization experiments were performed. As shown in Fig. 1A, the hybridizations revealed their main signals at about 1.5 kb. With RNA from ethanol-grown cells, the signal was much more intense than that obtained with RNA from glucose-grown cells (Fig. 1A). These results indicate that the C. glutamicum adhA gene is monocistronic and that it is subject to carbon source-dependent transcriptional regulation.

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FIG. 1. Transcriptional characterization of the adhA gene. (A) Northern blot analysis after growth of C. glutamicum in minimal medium containing ethanol (lanes 1 to 3) or glucose (lanes 4 to 6) as carbon sources. Total RNA (5 µg in lanes 1 and 4; 10 µg in lanes 2 and 5; 15 µg in lanes 3 and 6) was electrophoresed and probed with a radioactively labeled adhA-specific DNA probe. Sizes are shown on the right. (B) Primer extension analysis of the transcriptional start site in front of the adhA gene. The primer extension product is shown in lane 5. Lanes A, C, G, and T represent the products of sequencing reactions with the same primer (CM4) used for the primer extension reaction. The relevant DNA sequence (coding strand) is shown on the right, and the transcriptional start site is indicated by an asterisk.

To further investigate the carbon source-dependent transcription of the adhA gene, the three promoter fragments adhAp₁, adhAp₂, and adhAp₃ (Fig. 2A) were tested for activity. For this purpose, the plasmids pET2-adhAp₁, pET2-adhAp₂, and pET2-adhAp₃ were transformed into WT C. glutamicum, and the specific CAT activities in the resulting strains were determined during growth in minimal medium containing ethanol, glucose, acetate, and the substrate mixtures ethanol plus glucose or ethanol plus acetate. As shown in Table 3, WT C. glutamicum(pET2-adhAp₁) showed specific CAT activities of 6.89 U/mg protein on ethanol and about fourfold lower activities on glucose or ethanol plus glucose. Interestingly, on acetate or acetate plus ethanol, the specific CAT activities were even about 14-fold lower than on ethanol. In WT C. glutamicum(pET2-adhAp₂) containing the shortened adhA promoter fragment, the specific CAT activities on all substrates were five- to ninefold lower than in WT C. glutamicum(pET2-adhAp₁). WT C. glutamicum(pET2-adhAp₃) showed no CAT activity at all. These results indicate (i) that the adhA promoter resides in the adhA upstream region covered by fragment adhAp₂ (position –218 to +1 with respect to the translational start codon), (ii) that the adhA promoter drives the carbon source-dependent expression of adhA, (iii) that fragment adhAp₃ does not contain a functional promoter, and (iv) that the region covered by fragment adhAp₃ contains an activating element leading in a carbon source-independent manner to an increased adhA promoter activity.

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FIG. 2. Genomic locus of the adhA promoter region and DNA fragments used for mapping the relevant RamA and RamB binding sites (A) and representative EMSAs using hexahistidyl-tagged RamA (B) and RamB (C) proteins. cg3108 codes for a putatively secreted, unknown protein. The transcriptional start site is denoted as TS. The black boxes represent the putative RamA binding sites and the grey boxes the RamB binding sites. Their respective localizations relative to the transcriptional start site are given. The fragments used for the binding assays are given as bars and designated as indicated to the left, whereas the respective sizes are indicated to the right. The fragments used in the EMSAs with hexahistidyl-tagged RamA (B) and RamB (C) are indicated below the different parts of the gels. Lanes 1 to 4 show EMSAs using 0, 0.25, 0.5, and 1 µg of RamA or RamB, respectively. Lane 5 shows a negative control using 1 µg of bovine serum albumin protein with the respective probe.

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TABLE 3. Specific CAT activities of C. glutamicum strains carrying different adhA promoter fragments in plasmid pET2

To investigate the carbon source-dependent expression of adhA in more detail, we performed successive growth experiments with C. glutamicum(pET2-adhAp₂) and C. glutamicum(pET2-adhAp₁) in minimal medium containing glucose (1%), a glucose-ethanol substrate mixture (0.125% and 1%, respectively), ethanol (1%), and glucose again (1%) and determined the CAT activities (i.e., adhA promoter activities) in the course of the cultivations. Representative results for C. glutamicum(pET-adhAp₂) are shown in Fig. 3. During growth on glucose, the specific CAT activity was relatively low (Fig. 3A). In the subsequent culture containing glucose plus ethanol, the growth was biphasic; CAT activity was low in the glucose-metabolizing growth phase and increased in the ethanol-metabolizing growth phase to a maximum of about 1 U/mg protein (Fig. 3B). When the strain was then grown on ethanol as the single carbon source, CAT activity was high in the exponential growth and slightly lower in the stationary phase (Fig. 3C). The following cultivation on glucose led to a decrease of the specific CAT activity to the level observed in the first cultivation with glucose as the sole substrate (Fig. 3D). The CAT activities determined during the cultivations with C. glutamicum(pET-adhAp₁) correlated well with those of C. glutamicum(pET-adhAp₂) except that they were five- to ninefold higher under all conditions. The CAT activities observed in all these cultivations also correlate well with the ADH activities of WT C. glutamicum previously observed in identical successive growth experiments (3). Taken together, the results corroborate that the carbon source-dependent regulation of ADH in C. glutamicum and, also, the growth phase-dependent regulation of ADH on ethanol as the sole substrate take place at the transcriptional level.

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FIG. 3. adhA promoter activity during growth of WT C. glutamicum(pET2-adhAP2) on 1% glucose (A), 0.125% glucose and 1% ethanol (B), 1% ethanol (C), and 1% glucose (D). Black circles, growth; black bars, specific CAT activity.

To precisely localize the transcriptional initiation site of the C. glutamicum adhA gene, primer extension experiments were performed. With the use of oligonucleotide CM4 and 50 µg total RNA from C. glutamicum(pET2-adhAp₂), a signal corresponding to a G residue 130 nucleotides upstream of the adhA translational start codon ATG was observed (Fig. 1B). This result was confirmed in an independent experiment with a different primer (CM5) (data not shown). Centered 10 bp upstream of the adhA transcriptional initiation site, the sequence TAGGGT was found and designated as the –10 region. However, there is only relatively weak similarity to the consensus –10 sequence (TAT/CAAT) of corynebacterial promoters (39, 40). No apparent –35 region could be recognized, which is a common feature of C. glutamicum promoters (39, 40).

RamA and RamB bind to the adhA promoter region. AK, PTA, ICL, and MS are essential enzymes for acetate metabolism in C. glutamicum and are also required for the growth of C. glutamicum on ethanol (3). The expression of the genes encoding these four enzymes has been shown to be transcriptionally controlled by RamA and RamB (see the introduction). Interestingly, we observed three potential RamA binding sites in the promoter region of adhA, i.e., AGGGGGT, located 164 bp upstream of the transcriptional start, and CGGGGGT, located 112 bp and 57 bp upstream of the transcriptional start (Fig. 2A). Furthermore, two potential RamB binding sites were detected, located 132 bp upstream (TATTTTTCGAAAT; completely conserved nucleotides underlined) and 96 bp downstream (AGCAATTTGCCAA) of the adhA transcriptional start site (Fig. 2A). These observations suggested a direct interaction of RamA and RamB with the adhA promoter region and prompted us to perform EMSAs with His-tagged RamA and RamB proteins and the adhAp₁, adhAp₂, and adhAp₃ fragments (Fig. 2B and C). The incubation of adhAp₁ and adhAp₃ with RamA resulted in a three-step retardation of the probe (Fig. 2B), probably due to the binding of RamA to the three postulated RamA binding sites. The incubation of RamA with adhAp₂ resulted in a one-step retardation of the probe, probably due to the binding of RamA to the single RamA binding site 57 bp upstream of the transcriptional start. The incubation of adhAp₁ with RamB led to a two-step retardation signal (Fig. 2C), whereas the incubation of RamB with adhAp₂ and adhAp₃ led to a one-step retardation of the respective probes. We also tested for an interaction of RamA and RamB during the binding to the adhAp₁, adhAp₂, and adhAp₃ fragments. However, neither combined incubation with RamA and RamB nor the prebinding of RamA or RamB blocked or enhanced the binding of the other regulator (data not shown). These results are in agreement with the postulated RamA and RamB binding sites and indicate a direct participation of RamA and of RamB in adhA gene expression control in C. glutamicum.

RamA and RamB control the adhA gene expression. To test for a regulatory function of RamA and RamB in adhA expression, the specific ADH activities were determined in the ramB-deficient mutant C. glutamicum RG1 and in the ramA-deficient mutant C. glutamicum RG2 (Table 2). When grown in minimal medium containing ethanol, C. glutamicum RG1 showed about the same specific ADH activity as WT C. glutamicum. When grown on glucose or ethanol plus glucose and, also, when grown on acetate or a mixture of ethanol and acetate, C. glutamicum RG1 showed significantly (two- to fivefold) higher specific ADH activities than WT C. glutamicum. In contrast to C. glutamicum RG1, the RamA-deficient strain RG2 was not able to grow on ethanol, on acetate, or on ethanol plus acetate and showed no ADH activity on the other substrates tested (Table 2).

To study the direct influence of RamA and RamB on adhA promoter activity, the specific CAT activities were determined in the respective mutant strains containing plasmids pET2-adhAp₁ and pET2-adhAp₂ (Table 3). The RamA-negative strains C. glutamicum RG2(pET2-adhAp₁) and C. glutamicum RG2(pET2-adhAp₂) showed no promoter activities at all. However, in C. glutamicum RG1(pET2-adhAp₁), we detected twofold higher specific CAT activities on glucose and on ethanol plus glucose and about threefold higher activities on acetate and ethanol plus acetate than in WT C. glutamicum(pET2-adhAp₁). The specific CAT activities on ethanol were similar in both WT C. glutamicum(pET2-adhAp₁) and C. glutamicum RG1(pET2-adhAp₁) (Table 3). Interestingly, the specific CAT activities in C. glutamicum RG1(pET2-adhAp₂) were similar to the activities determined in WT C. glutamicum(pET2-adhAp₂) on all substrates tested. Taken together, the results indicate an involvement of both RamA and RamB in the expression control of adhA. RamA is absolutely required for the expression of the C. glutamicum adhA gene and, thus, for ADH activity. RamB exerts a negative control when the cells grow in the presence of glucose or acetate as the sole or as an additional carbon source. However, the results also indicate that neither RamA nor RamB alone is responsible for the carbon source-dependent ADH regulation, i.e., for complete adhA repression in the presence of glucose or acetate in the growth medium. It is feasible that different RamA abundance within the cells (and thus, different adhA activation), together with RamB, controls adhA expression during growth on different substrates. Alternatively, C. glutamicum might possess an additional transcriptional regulator(s) controlling adhA expression and, together with RamB, being responsible for catabolite repression in medium containing ethanol and glucose or acetate.

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C. glutamicum is able to grow on ethanol as the single source of carbon and energy, and it has been shown that the enzymes AK, PTA, ICL, and MS are essential for growth on this substrate (3). Here we identified and characterized the C. glutamicum adhA gene encoding an ADH which is essential for ethanol oxidation. ADH proteins are ubiquitous enzymes and widespread in all life forms, fulfilling quite different physiological functions. Some are involved in the formation and others in the degradation of alcohols. The best-studied ADHs are the long-chain, zinc-dependent enzymes of group I, represented by the ADHI of S. cerevisiae (reviewed in reference 41). Group II ADHs consist of short-chain, zinc-independent enzymes, and group III contains iron-activated ADHs, represented by the CoA-dependent bifunctional enzymes from E. coli (AdhE) and Clostridium acetobutylicum (16, 22). BLAST analysis showed high similarities of the C. glutamicum adhA gene product with group I ADHs, and sequence alignments revealed highly conserved residues and sequence motifs in those regions suggested to be essential for cofactor binding and catalytic activity. Among these were the GXGXXG pattern (position 180 to 185) in the coenzyme-binding fold and a highly conserved aspartate (Asp-203) (28, 34, 41, 46). This aspartate residue corresponds well with the finding that the C. glutamicum ADH activity was specific for NAD⁺ (3). Furthermore, the GHEXXGXXXXXGXXV motif containing the second ligand of the catalytic zinc (i.e., His-69) (28, 41) is highly conserved (position 68 to 82). Taken together, all these data indicate that the ADH of C. glutamicum is a member of the group I ADHs.

The transcriptional characterization of the C. glutamicum adhA revealed a monocistronic organization and a carbon source-dependent expression. It was induced in the presence of ethanol and repressed by glucose. The repression of adhA was also observed when both carbon sources were present in the medium, in agreement with the low specific ADH activity detected on the substrate mixture. As shown previously, we also observed low ALDH, AK, PTA, ICL, and MS activities on ethanol-glucose (3), suggesting a repression of all the corresponding genes in the presence of both these carbon sources. The biphasic growth behavior observed on the ethanol-glucose mixture is thus obviously due to the repression of all the genes in the presence of glucose and derepression when glucose was exhausted.

Interestingly, adhA was also repressed in the presence of acetate. Acetate-mediated adhA repression is corroborated by DNA microarray experiments showing lower adhA-RNA levels in acetate-grown C. glutamicum cells than in glucose-grown cells (37). Since the ADH and the ALDH reactions in principle are reversible, it can be speculated that the acetate-mediated repression of adhA prevents ethanol formation from acetate. Above all, we detected a biphasic growth behavior of C. glutamicum during growth on the ethanol-acetate substrate mixture, with acetate utilization preferred (data not shown). The favored acetate metabolization in the presence of ethanol was unexpected, since the results of Wendisch et al. (53) indicated a parallel substrate metabolization of acetate and other carbon sources, such as lactate, succinate, or glutamate. Furthermore, in contrast to the adhA gene, the pta-ack operon, aceA, and aceB showed high levels of expression in the presence of acetate (reviewed in reference 20). These results led us to conclude that the observed biphasic growth behavior on ethanol-acetate is due to the repression of the adhA gene in the presence of acetate and derepression after acetate consumption. However, the preferred utilization of acetate or other organic acids is not uncommon in other soil bacteria, as shown for, e.g., pseudomonads, Azotobacter vinelandii, Acinetobacter baylyi ADP1, or Ralstonia eutropha (2, 11, 13, 18).

As shown here, ethanol utilization is tightly regulated in C. glutamicum, and this has also been shown for other organisms. The genes encoding the components of the ethanol oxidation system of P. aeruginosa are positively regulated by AgmR. Furthermore, the two-component system ExaDE controls the expression of exaA, the gene for the pyrroloquinoline quinone-dependent ADH (21, 23). ADH2, the ethanol-oxidizing ADHII gene of S. cerevisiae, was repressed in the presence of glucose or other fermentative carbon sources and activated by Adr1 when glucose was exhausted (17, 54). In Aspergillus nidulans, the alc genes of the ethanol utilization pathway are also subject to two regulatory mechanisms: the positive transcriptional regulation mediated by AlcR and the negative control of CreA in the presence of more favorable growth substrates, such as glucose or acetate (reviewed in reference 15).

In contrast to the well-characterized regulation of ethanol metabolism in some other organisms, so far nothing was known on the regulation of ethanol utilization in C. glutamicum. However, the results obtained in this study show that the two regulators of acetate metabolism, RamA and RamB, are involved in adhA regulation. Similar to the regulation of the pta-ack operon, aceA, and aceB (10), RamA activates the adhA gene and, in fact, is essential for adhA expression on any carbon source tested. As previously shown for the control of the expression of the RamA gene itself (9), the level of adhA expression is dependent on the number of RamA binding sites in the promoter region. A dependence of the binding-site number on the level of gene expression has been also assumed for genes controlled by AmtR, the repressor of nitrogen control in C. glutamicum, and for the CreA-mediated regulation of alcR in A. nidulans (4, 15). In contrast to RamA, RamB was found to exert a negative control on adhA expression. Similar to the control of the AK, PTA, ICL, and MS genes (19), RamB represses adhA expression in the presence of glucose. RamB also represses adhA in the presence of acetate, and this is in contrast to the control of the former genes, which are activated in the presence of acetate. However, the adhA gene in the RamB-deficient mutant C. glutamicum RG1 was not completely derepressed in the presence of glucose or acetate. This finding might be due to a lower intracellular abundance of RamA and thus, in fact, to an incomplete activation of adhA expression under these conditions. Alternatively, RamB might not be the only regulator involved in the glucose- and acetate-dependent repression of adhA. Interestingly, although in vitro binding of RamB to both potential RamB binding sites was shown, only the motif 132 bp upstream of the transcriptional start site exhibited a physiological function under the conditions tested. All these results led us to suppose that the RamB-mediated repression of adhA is different from that of the pta-ack operon, aceA, and aceB. We propose the requirement of other effector molecules and, possibly, the involvement of other proteins in adhA repression. A candidate for involvement might be GlxR, a cAMP-dependent regulator (31), since we observed a sequence motif (TGTTG-N₆-ACACA) with similarity to the proposed consensus sequence of GlxR (TGTGA-N₆-ACACT) (35) in the adhA promoter region, centered 21 bp downstream of the adhA transcriptional initiation site. GlxR has been shown to be involved in the glucose-induced repression of the genes of the gluconate metabolism (gntP and gntK) (35), and due to the observations of potential binding sites, it has been supposed to be involved in the carbon source-dependent regulation of other genes, such as acn (encoding aconitase) or gluA (encoding a component of the glutamate uptake system) (35). However, further studies should focus on the involvement of GlxR in the regulation of adhA.

As outlined in the introduction, carbon catabolite repression is an important global regulatory system that has been investigated in several organisms (6, 45). In contrast, very little is known about the carbon catabolite repression mechanism in C. glutamicum and closely related Corynebacterineae, such as other corynebacteria or mycobacteria. The investigation of the regulation of ethanol metabolism in C. glutamicum and the identification of the participating regulators should help to elucidate catabolite repression in these organisms.

 
The support of the BMBF (grants 0313105 "GenoMik" and 0313805G "GenoMik-Plus") is gratefully acknowledged.

 
* Corresponding author. Mailing address: Institute of Microbiology and Biotechnology, University of Ulm, 89069 Ulm, Germany. Phone: 49 731 502 2707. Fax: 49 731 502 2710. E-mail: bernhard.eikmanns{at}uni-ulm.de

Published ahead of print on 10 August 2007.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

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Journal of Bacteriology, October 2007, p. 7408-7416, Vol. 189, No. 20
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