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J Bacteriol. 2009 Jul; 191(13): 4251–4258.
Published online 2009 May 1. doi:  10.1128/JB.00303-09
PMCID: PMC2698506

The ldhA Gene, Encoding Fermentative l-Lactate Dehydrogenase of Corynebacterium glutamicum, Is under the Control of Positive Feedback Regulation Mediated by LldR


Corynebacterium glutamicum ldhA encodes l-lactate dehydrogenase, a key enzyme that couples l-lactate production to reoxidation of NADH formed during glycolysis. We previously showed that in the absence of sugar, SugR binds to the ldhA promoter region, thereby repressing ldhA expression. In this study we show that LldR is another protein that binds to the ldhA promoter region, thus regulating ldhA expression. LldR has hitherto been characterized as an l-lactate-responsive transcriptional repressor of l-lactate utilization genes. Transposon mutagenesis of a reporter strain carrying a chromosomal ldhA promoter-lacZ fusion (PldhA-lacZ) revealed that ldhA disruption drastically decreased expression of PldhA-lacZ. PldhA-lacZ expression in the ldhA mutant was restored by deletion of lldR, suggesting that LldR acts as a repressor of ldhA in the absence of l-lactate and the LldR-mediated repression is not relieved in the ldhA mutant due to its inability to produce l-lactate. lldR deletion did not affect PldhA-lacZ expression in the wild-type background during growth on either glucose, acetate, or l-lactate. However, it upregulated PldhA-lacZ expression in the sugR mutant background during growth on acetate. The binding sites of LldR and SugR are located around the −35 and −10 regions of the ldhA promoter, respectively. C. glutamicum ldhA expression is therefore primarily repressed by SugR in the absence of sugar. In the presence of sugar, SugR-mediated repression of ldhA is alleviated, and ldhA expression is additionally enhanced by LldR inactivation in response to l-lactate produced by LdhA.

Lactate is a fermentation end product and a carbon source for growth of bacteria. Lactate dehydrogenases (LDHs) are enzymes that enable both production and utilization of lactate (7, 22, 41). Soluble, NAD-linked LDHs are responsible for the formation of lactate from pyruvate under fermentative conditions (41, 42). This reaction consumes one NADH molecule per three carbon atoms, thus balancing out the NADH produced in the glyceraldehyde-3-phosphate dehydrogenase step of glycolysis. In contrast, respiratory LDHs involved in utilization of lactate are membrane-bound proteins which use quinone as an electron acceptor and are coupled to the respiratory chain (7, 22).

Regulation of expression of LDHs is most studied for the gram-negative bacterium Escherichia coli, which expresses three LDHs: one fermentative LDH encoded by the ldhA gene and two respiratory LDHs encoded by the lldD and dld genes (2, 4, 12, 41, 42, 49). The ldhA gene is induced in anaerobically grown cultures (3, 28). Expression of the ldhA gene is affected both by the redox sensor ArcAB and by carbohydrate metabolism regulators, CsrAB and Mlc (17), although direct binding of the regulators and their regulatory roles involved have not been elucidated. The lldD gene is included in the lldPRD operon (4). lldP encodes an l-lactate-specific permease, and lldR encodes a transcriptional regulator. Expression of the lldPRD operon is regulated by LldR in response to the presence of l-lactate (1) and repressed under anaerobic conditions by the ArcAB system (27).

Corynebacterium glutamicum is a nonpathogenic high-GC gram-positive bacterium which is widely used in the industrial production of amino acids (13, 20). We have also demonstrated use of C. glutamicum for production of lactate (15, 33, 34). While LdhA, encoded by the ldhA gene, is the NAD-dependent LDH responsible for production of l-lactate in C. glutamicum (15), LldD, encoded by the lldD gene, is the membrane-bound, quinone-dependent LDH essential for growth on l-lactate (36, 37). The lldD gene forms an operon with cgR_2818, which encodes a permease (37). We recently demonstrated that the DeoR-type transcriptional regulator, SugR, which is a transcriptional repressor of genes involved in the phosphoenolpyruvate-dependent phosphotransferase system (PTS) and the glycolytic genes (5, 6, 8, 40, 43), represses ldhA expression in the absence of sugar and that sugar phosphates have a negative effect on its repressor activity (45). On the other hand, expression of the cgR_2818-lldD operon is repressed by the GntR-type transcriptional regulator, LldR (CgR_2816), and the repression by LldR is relieved in the presence of l-lactate (10).

LldR binds to the ldhA promoter region (45), but its role in regulation of ldhA expression remains unknown. Transcriptome analyses (9, 10) indicate that LldR represses not only the cgR_2818-lldD operon but also the fruR-fruK-ptsF operon, which is indispensable for fructose utilization, whereas overexpression of LldR reduces ldhA expression (10).

In this study, we report the role of LldR in ldhA expression in C. glutamicum. We use transposon mutagenesis to identify genes affecting ldhA expression and show that disruption of ldhA decreases expression of a chromosomal ldhA promoter-lacZ fusion. The ldhA promoter activity reduced in the ldhA mutant is restored by deletion of the lldR gene, indicating that LldR acts as a transcriptional repressor of ldhA and that the repression is alleviated in response to l-lactate, a product of the LdhA reaction. The positive feedback regulation of lactate-producing LDH gene has not been hitherto reported for bacteria. The coordinated roles of two transcriptional repressors SugR and LldR in ldhA expression are discussed.


Bacterial strains and plasmids.

The strains and plasmids used in this study are listed in Table Table11.

Bacterial strains and plasmids used in this study

Culture conditions.

For genetic manipulation, E. coli strains were grown at 37°C in Luria-Bertani (LB) medium. C. glutamicum strains were grown at 33°C in nutrient-rich medium (A medium) (16) with 4% glucose. When appropriate, the media were supplemented with antibiotics. The final antibiotic concentrations for E. coli were 50 μg of ampicillin ml−1, 50 μg of kanamycin ml−1, and 50 μg of chloramphenicol ml−1; for C. glutamicum, kanamycin (50 μg ml−1) and chloramphenicol (5 μg ml−1) were used. Growth experiments with C. glutamicum were performed using A medium containing 1% glucose, acetate, or lactate as described previously (43).

DNA manipulations.

The oligonucleotides used in this study (Table (Table2)2) were obtained from Gene Design (Osaka, Japan). Plasmid DNA was isolated with the QIAprep spin miniprep kit (Qiagen, Hilden, Germany). Chromosomal DNA was isolated from C. glutamicum using Genomic Prep (GE Healthcare Bioscience, Piscataway, NJ), modified by using 4 mg ml−1 lysozyme at 37°C for 2 h. Restriction endonucleases were purchased from Takara (Osaka, Japan) and used according to the manufacturer's instructions. The PCR was carried out using a GeneAmp PCR system (Applied Biosystems, Foster City, CA) using LA Taq polymerase (Takara) or Phusion polymerase (New England Biolabs). The resulting PCR fragments were purified with the QIAquick PCR purification kit (Qiagen). E. coli was transformed by the CaCl2 procedure, as described by Sambrook et al. (35). C. glutamicum was transformed by electroporation as described previously (47). The nucleotide sequence of cloned DNA fragments was determined by using a 3130xl genetic analyzer (Applied Biosystems).

Oligonucleotide primers used in this study

Transposon mutagenesis.

Transposon mutagenesis using Tn5-based minitransposon (EZ::Tn<Kan2> transposome system; Epicenter, Wisconsin) was performed as described previously (39). The EZ::Tn<Kan2> transposome was inserted into the genome of reporter strains KT14 and KT15, carrying the ldhA promoter-lacZ fusion on the genome (45), by electroporation (46). Cells were incubated in A medium containing 4% glucose for 2 h at 33°C and subsequently spread plated on A medium containing 50 μg ml−1 kanamycin and 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside (X-gal) and incubated for 1 to 2 days at 33°C.

Determination of transposon insertion sites.

To determined transposon insertion sites, thermal asymmetric interlaced PCR using the primers AP1, GSP1, and GSP2 and the genomic DNA as a template was performed as described previously (26, 39). The resulting amplicon was used for direct sequencing using the primer GSP3. The sequence obtained was searched by BLAST against the C. glutamicum R genomic DNA sequence (50).

Construction of mutants.

A DNA fragment containing the lldR gene was amplified using the primer lldR FWSph and the primer lldR RVSal. The PCR product was cloned into pCRA725 (14), a suicide vector for markerless gene disruption, yielding pCRC617. An internal 621 bp of the lldR gene was removed by inverse PCR using the primers lldR invFW and lldR invRV and pCRC617 as a template. The PCR product was digested with BglII and self-ligated, yielding pCRC618. C. glutamicum R was transformed by electroporation with pCRC618, and screening for the deletion mutants was performed as described previously (14). Deletion of the lldR gene was checked by PCR. The obtained mutant strain was designated KT19. Similarly, the lldR gene in the ldhA and sugR mutants was deleted, resulting in strains KT17 and KT21, respectively.

β-Galactosidase assay.

β-Galactosidase activity in C. glutamicum was determined as described previously (16). Cells permeabilized with toluene were incubated with o-nitrophenyl-β-galactosidase, and activity was measured in Miller units as previously described (29). Assays were carried out in triplicate for each sample, and results are presented as means ± standard deviations.

Overexpression and purification of His-tagged LldR protein.

The lldR gene was amplified from chromosomal DNA of C. glutamicum R by PCR with the primers lldR His FWNde and lldR His RVXba. The PCR product was cloned into the expression vector pColdI (Takara), yielding pCRC616. E. coli BL21(DE3) transformed with pCRC616 was grown at 37°C in LB medium to an optical density at 600 nm of 0.5. The culture was incubated for 30 min at 15°C, and then expression of the His-tagged LldR was induced by addition of 0.5 mM isopropyl-β-d-thiogalactopyranoside. The culture was shaken overnight at 15°C. The cells were harvested by centrifugation and frozen at −80°C until ready for use. The His-tagged protein was purified by affinity chromatography on nickel-nitrilotriacetic acid agarose (Qiagen) according to the instruction manual. For desalting, the eluted protein was loaded into the PD-10 column (GE Healthcare Bioscience) and eluted with buffer A (50 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 1 mM EDTA, and 1 mM dichlorodiphenyltrichloroethane). The concentration of the purified protein was determined with a Bio-Rad protein assay (Bio-Rad Laboratories) using bovine serum albumin as a standard.


Cy3-labeled DNA fragments used in electrophoretic mobility shift assays (EMSAs) were generated by PCR and purified using a PCR purification kit (Qiagen). The ldhA promoter fragments P1, P2, and P3 were generated using the primers pldhA FW1, pldhA FW2, and pldhA FW5, together with the primer pldhA RV Cy3. EMSA with His-tagged LldR was performed as described previously (44). The resulting DNA-protein complexes were loaded onto a 5% polyacrylamide gel. Electrophoresis was performed at room temperature and at 150 V using 0.5× TBE (44.5 mM Tris base, 44.5 mM boric acid, 1 mM EDTA) as the electrophoresis buffer. DNA and DNA-protein complexes were visualized by using a Typhoon TRIO variable-mode imager (GE Healthcare Bioscience).

DNase I footprinting.

DNase I footprinting was performed as described previously (44). Labeled DNA probes were obtained by amplification with 5′-IRD-700-labeled oligonucleotides. The primer pair pldhA IRD FW/pldhA RV1 or pldhA IRD RV/pldhA FW8 was used to generate the coding- or noncoding-strand-labeled DNA fragment. The DNA sequencing reactions were set up using the same IRD-700-labeled primers and pCRC615 (45) as a template and a DYEnamic direct cycle sequencing kit with 7-deaza-dGTP (GE Healthcare).


Expression of ldhA requires functional ldhA gene.

We used a transposon mutagenesis system, which we previously developed for C. glutamicum R (39), to identify genes involved in the regulation of ldhA expression. We applied the system to strain KT14, which carries the ldhA promoter-lacZ translational fusion (PldhA-lacZ) in the wild-type genome (45) and forms blue colonies on a plate containing glucose and X-Gal. We screened for mutants forming white or light blue colonies, in which ldhA expression was expected to be decreased. The transposon was inserted in the lacZ gene in strains forming white colonies on the plate. Mutants forming light-blue colonies contained transposon insertions in pgi (cgR_0966), pfk (cgR_1327), zwf (cgR_1626), ptsH (cgR_1797), ldhA (cgR_2812), a putative aldehyde dehydrogenase gene (cgR_2330), or dnaK (cgR_2690). The genes ptsH, pgi, pfk, and zwf, encoding the general component of PTS, phosphoglucoisomerase, phosphofructokinase, and glucose-6-phosphate dehydrogenase, respectively, are involved in sugar uptake and the upper glycolytic pathway. It is possible that disruption of any of these genes decreases sugar phosphates and results in downregulation of PldhA-lacZ expression, since sugar phosphates act as inducers of ldhA expression by inhibiting activity of the transcriptional repressor SugR (45).

To elucidate the SugR-independent regulation of ldhA expression, we applied the transposon mutagenesis system to the sugR mutant strain KT15, which also carries PldhA-lacZ on the genome. Unlike the wild-type strain, KT15 expressed ldhA even in the absence of sugar due to the absence of the SugR repressor, hence forming blue colonies on a plate containing acetate and X-Gal. We screened for mutants which form light-blue colonies on the plate. The transposon disrupted the ptsI (cgR_1763) or ldhA (cgR_2812) gene in the mutants forming light-blue colonies. The ptsI gene encodes a general component of PTS. It is not clear what, if any, role PtsI plays in ldhA expression in the absence of sugar. The fact that the disruption of the ldhA gene reduced ldhA promoter activity in both the wild-type and sugR mutant backgrounds suggested that a lack of l-lactate production has an inhibitory effect on ldhA expression.

ldhA promoter activity is reduced in the ldhA mutant.

To confirm the results of the transposon mutagenesis studies, we integrated PldhA-lacZ into the genome of the ldhA mutant, resulting in strain KT16. We compared its β-galactosidase activity to that in the wild-type background. Expression of PldhA-lacZ in strain KT14 (wild type) was increased at the onset of the stationary phase of growth on glucose (Fig. (Fig.1).1). The PldhA-lacZ expression level in strain KT16 (ldhA mutant) was 30% of that in strain KT14 at the onset of the stationary phase, confirming requirement of the ldhA gene for proper ldhA expression. This conclusion was also supported by the fact that strain KT16 transformed with pCRA715 carrying the C. glutamicum R ldhA gene under the control of the constitutive promoter showed higher PldhA-lacZ expression than a control strain transformed with the vector pCRB1 only (Fig. (Fig.22).

FIG. 1.
ldhA promoter activity in strain KT14 (wild type; white circles), strain KT16 (ldhA mutant; black circles), or strain KT18 (ldhA-lldR mutant; triangles) during growth on A medium with 1% of glucose. Activity is expressed as means from at least ...
FIG. 2.
ldhA promoter activity in strain KT16L (transformed with pCRA715, which carries the l-lactate-producing LDH gene; triangles), strain KT16D (transformed with pCRB203, which carries the d-lactate-producing LDH gene; circles), or strain KT16C (transformed ...

We have previously reported production of d-lactate by an ldhA mutant of C. glutamicum transformed with pCRB203 carrying the Lactobacillus delbrueckii ldhA gene encoding d-lactate-producing LDH (34). Unlike the effect of introduction of pCRA715 carrying an l-lactate-producing LDH described above, pCRB203 introduction did not increase, but rather decreased, the PldhA-lacZ expression of strain KT16 (Fig. (Fig.2,2, results for KT16D). The low PldhA-lacZ expression of strain KT16 can thus be attributed to its incapacity for l-lactate production but not to the absence of NADH-dependent pyruvate reduction.

Deletion of the lldR gene restores ldhA promoter activity in the ldhA mutant.

Previously, we identified LldR as a protein binding to the ldhA promoter region (45). It has been reported elsewhere that LldR acts as a transcriptional repressor of the cgR_2818-lldD operon, involved in l-lactate utilization, and that LldR-mediated repression is relieved in the presence of l-lactate but not in the presence of d-lactate (10). We posited that LldR also represses ldhA expression and that the repression by LldR is not relieved in the ldhA mutant because of its inability to produce l-lactate. To test this assumption, we deleted the lldR gene in the ldhA mutant and compared the ldhA promoter activity in the ldhA-lldR mutant to that in the ldhA mutant. Expression of PldhA-lacZ in strain KT18 (ldhA-lldR mutant) was comparable to that in strain KT14 (wild type) during growth on glucose (Fig. (Fig.1),1), demonstrating that the decrease of expression of PldhA-lacZ by disruption of ldhA in strain KT16 occurs because repression of the ldhA gene by LldR is not relieved.

ldhA expression is primarily repressed by SugR.

Expression of ldhA is also repressed by SugR, a global repressor of sugar utilization genes (5, 45). To elucidate how these two transcriptional regulators, LldR and SugR, control ldhA expression, we compared PldhA-lacZ expression in the wild-type strain against that in lldR, sugR, and lldR-sugR mutants during growth on each of glucose, acetate and l-lactate. During growth on glucose, PldhA-lacZ expression in the lldR mutant was comparable to that in the wild-type strain (Fig. (Fig.3A).3A). PldhA-lacZ expression in the sugR mutant was higher than that in the wild-type strain, as described earlier (Fig. (Fig.3A)3A) (45). During growth on either acetate or l-lactate, we detected no PldhA-lacZ expression in either the wild-type strain or the lldR mutant, whereas PldhA-lacZ expression was markedly upregulated in the sugR mutant (Fig. 3B and C). Thus, ldhA derepression by LldR inactivation is probably suppressed by SugR. In fact, expression of PldhA-lacZ in the sugR-lldR mutant was significantly higher than that in the sugR mutant during growth on acetate (Fig. (Fig.3B).3B). However, during growth on glucose or l-lactate, no difference in PldhA-lacZ expression was observed between the strains (Fig. 3A and C). These results suggest that l-lactate generated in the glucose-grown cells or provided in the medium inactivates LldR in the sugR mutant, while in the acetate-grown cells, ldhA derepression in the sugR mutant is not enough to relieve LldR-mediated repression.

FIG. 3.
ldhA promoter activity in strain KT14 (wild type; circles), strain KT15 (sugR mutant; triangles), strain KT20 (lldR mutant; white squares) or strain KT22 (lldR-sugR mutant; black squares) during growth on A medium with 1% of glucose (A), acetate ...

Binding of LldR to ldhA promoter region.

To examine binding of LldR to the ldhA promoter region, we performed an EMSA using a DNA fragment containing the ldhA promoter region encompassing the region from position −104 to +92 with respect to the transcriptional start point (TSP) of the ldhA gene and recombinant His-tagged LldR. The probe DNA was shifted by LldR in a protein concentration-dependent manner (Fig. (Fig.4A,4A, P1), demonstrating that LldR specifically binds to the ldhA promoter region.

FIG. 4.
Binding of LldR to the ldhA promoter region. (A) EMSA using ldhA promoter fragments and the LldR protein. The regions contained in the DNA fragments (P1 to P3) are indicated by position with respect to the TSP of ldhA at the top of the gels. Each well ...

To locate the LldR binding site more precisely, we performed DNase I footprinting assays. We labeled each of the strands of a DNA fragment encompassing the region from position −203 to +146, incubated them with increasing quantities of LldR, and then hydrolyzed them using DNase I. Figure Figure4B4B shows that LldR protected a 17-bp sequence on both strands. The region protected on the coding strand extends from position −28 to −44, whereas the protected region on the noncoding strand extends from position −31 to −47 with respect to the TSP. This region includes the putative binding site for LldR, TNGTNNNACNA (10) (Fig. (Fig.4C).4C). We observed no band shift upon deleting the putative LldR binding site (between positions −43 and −33 with respect with the TSP) (Fig. (Fig.4A,4A, compare P2 with P3), confirming that LldR recognizes the site for binding.

SugR and LldR independently bind to the ldhA promoter.

The region protected by SugR from DNase I attack (between positions −38 and +20) (45) overlaps the region protected by LldR (Fig. (Fig.4B).4B). To examine the effect of binding of one regulator on that of another, we performed DNase I footprinting assays in the presence of both transcriptional regulators. The same region was protected from DNase I attack regardless of whether SugR was added before LldR, after LldR, or simultaneously with LldR, indicating that the two regulators are able to bind to the ldhA promoter region independently (Fig. (Fig.5).5). However, an additional region between positions −52 and −48 was protected only in the presence of both SugR and LldR, suggesting cooperative binding of the two regulators to the ldhA promoter.

FIG. 5.
DNase I footprinting analysis of the binding of LldR and SugR to the ldhA promoter regions examined on the coding and the noncoding strands. A DNA fragment (10 nM) was incubated with increasing amounts of SugR and/or LldR: lanes 1 and 11, no protein; ...


Previously, we demonstrated that ldhA expression is repressed by SugR in the absence of sugar (45). The results obtained in the current study indicated that LldR acts as a transcriptional repressor of ldhA in the absence of l-lactate. No effect of deletion of the lldR gene on ldhA expression was observed in the wild-type background, but a positive effect on ldhA expression was observed in the sugR mutant background (Fig. (Fig.3),3), indicating that ldhA expression is primarily repressed by SugR. This may be ascribed to the fact that the binding site of SugR is located around the −10 region of the ldhA promoter and that of LldR is located from position −43 to −33 with respect to the TSP (Fig. (Fig.4C).4C). In contrast, the cgR_2818-lldD operon, involved in l-lactate utilization, is derepressed by deleting the lldR gene alone (10). During growth on l-lactate as the sole carbon and energy source, ldhA expression is repressed by SugR, even though repression of the cgR_2818-lldD operon by LldR can be relieved. This enables the cell to efficiently utilize l-lactate for growth. During growth in the presence of sugar, repression of ldhA by SugR is partially relieved at the onset of the stationary phase and l-lactate produced by LdhA alleviates the repression of ldhA by LldR. To the best of our knowledge, this is the first example of fermentative LDH requiring lactate for its proper expression in bacteria. This indicates that ldhA expression in C. glutamicum is subjected to the positive feedback regulation mediated by LldR.

Repression of genes involved in l-lactate production (ldhA) and utilization (lldD) by a common repressor, LldR, seems to create a futile cycle between pyruvate and l-lactate. The futile cycle, consisting of cytoplasmic (LdhA) and membrane-bound (LldD) LDHs, has been suggested to function in reoxidization of NADH in a strain carrying a mutation in the ndh gene, which encodes NADH dehydrogenase (31); LdhA reduces pyruvate to l-lactate with oxidation of NADH, whereas LldD oxidizes l-lactate to pyruvate with reduction of menaquinone. Since oxidation of NADH by respiration would be limited at the onset of the stationary phase, where oxygen tension is decreased by high cell density, the LdhA-LldD system, instead of respiration, may function to reoxidize NADH under these conditions. It has also been reported that cytoplasmic malate dehydrogenase and membrane-bound malate dehydrogenase function as a NADH reoxidation system in the ndh mutant of C. glutamicum (30). In addition, in E. coli, overexpression of membrane-bound LDH (Dld) was able to complement the ndh mutant, which excretes large quantities of d-lactate, unlike the wild-type strain, due to an inability to reoxidize NADH by respiration (49).

The ldhA mutant strain complemented with the L. delbrueckii ldhA gene, whose product converts pyruvate to d-lactate, not l-lactate, exhibited lower ldhA promoter activity than the control strain transformed with vector only (Fig. (Fig.2).2). Therefore, accumulation of substrates (NADH and/or pyruvate) may be required for upregulation of ldhA expression at the onset of the stationary phase. In Bacillus subtilis, the fermentative LDH-encoding gene, ldh, is repressed by Rex, a NADH/NAD+ sensor (24). The DNA binding ability of Rex is inhibited at a high NADH/NAD+ ratio and increased at a low NADH/NAD+ ratio (11, 48). Although no Rex homolog is encoded in the genome of C. glutamicum, there might be a factor monitoring the cytosol redox balance, as Rex does in B. subtilis. A novel transcriptional regulator, ArnR, which regulates the nitrate reductase operon, narKGHJI, responsible for anaerobic respiration, is one of the candidates because it possesses an oxygen-sensing cofactor, such as an Fe-S cluster (32).

Finally, SugR and LldR can simultaneously bind to the ldhA promoter region, and binding of both the regulators extended the region protected from DNase I attack (Fig. (Fig.5).5). This indicates that the simultaneous binding of both SugR and LldR has an additional regulatory role for ldhA expression. A putative binding site of GlxR, a cAMP receptor protein-type global transcriptional regulator controlling various physiological reactions (18, 19, 25), has been found in the ldhA promoter between positions −70 and −55, and GlxR is able to interact with the ldhA promoter in vitro (21). The GlxR binding site flanks the region protected only by simultaneous binding of both SugR and LldR, suggesting that the two repressors affect binding of GlxR to the ldhA promoter. However, the physiological function of GlxR in ldhA expression and the interaction of the three transcriptional regulators on the ldhA promoter remain to be investigated.


We thank Crispinus A. Omumasaba (RITE) for critical reading of the manuscript.

This work was partially supported by a grant from the New Energy and Industrial Technology Development Organization (NEDO).


Published ahead of print on 1 May 2009.


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