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Regulation of the rhaEWRBMA Operon Involved in L-Rhamnose Catabolism through Two Transcriptional Factors, RhaR and CcpA, in Bacillus subtilis Kazutake Hirooka, Yusuke Kodoi, Takenori Satomura,* Yasutaro Fujita

ABSTRACT

The Bacillus subtilis rhaEWRBMA (formerly yuxG-yulBCDE) operon consists of four genes encoding enzymes for L-rhamnose catabolism and the rhaR gene encoding a DeoR-type transcriptional regulator. DNase I footprinting analysis showed that the RhaR protein specifically binds to the regulatory region upstream of the rhaEW gene, in which two imperfect direct repeats are included. Gel retardation analysis revealed that the direct repeat farther upstream is essential for the high-affinity binding of RhaR and that the DNA binding of RhaR was effectively inhibited by L-rhamnulose-1-phosphate, an intermediate of L-rhamnose catabolism. Moreover, it was demonstrated that the CcpA/P-Ser-HPr complex, primarily governing the carbon catabolite control in B. subtilis, binds to the catabolite-responsive element, which overlaps the RhaR binding site. In vivo analysis of the rhaEW promoter-lacZ fusion in the background of ccpA deletion showed that the L-rhamnose-responsive induction of the rhaEW promoter was negated by the disruption of rhaA or rhaB but not rhaEW or rhaM, whereas rhaR disruption resulted in constitutive rhaEW promoter activity. These in vitro and in vivo results clearly indicate that RhaR represses the operon by binding to the operator site, which is detached by L-rhamnulose-1-phosphate formed from L-rhamnose through a sequence of isomerization by RhaA and phosphorylation by RhaB, leading to the derepression of the operon. In addition, the lacZ reporter analysis using the strains with or without the ccpA deletion under the background of rhaR disruption supported the involvement of CcpA in the carbon catabolite repression of the operon. IMPORTANCE

Since L-rhamnose is a component of various plant-derived compounds, it is a potential carbon source for plant-associating bacteria. Moreover, it is suggested that L-rhamnose catabolism plays a significant role in some bacteria-plant interactions, e.g., invasion of plant pathogens and nodulation of rhizobia. Despite the physiological importance of L-rhamnose catabolism for various bacterial species, the transcriptional regulation of the relevant genes has been poorly understood, except for the regulatory system of Escherichia coli. In this study, we show that, in Bacillus subtilis, one of the plant growth-promoting rhizobacteria, the rhaEWRBMA operon for L-rhamnose catabolism is controlled by RhaR and CcpA. This regulatory system can be another standard model for better understanding the regulatory mechanisms of L-rhamnose catabolism in other bacterial species.

B

acillus subtilis is a soil-dwelling bacterium that has been extensively and closely studied as a Gram-positive model bacterium. B. subtilis species have often been isolated from the rhizosphere of various terrestrial plants; many of them have been shown to be plant growth-promoting rhizobacteria whose association with the plant roots enhances the adaptive potential of plants and increases their growth (1). Interestingly, B. subtilis is also a saprophytic bacterium that secretes various degrading enzymes, including those able to hydrolyze the polysaccharides that constitute the plant cell wall, although it does not prey on the living plant cells (2). Pectin, a polysaccharide contained in the primary plant cell wall, consists of homogalacturonan as a linear chain part and two types (I and II) of rhamnogalacturonans as branched-chain parts (3). L-Rhamnose (6-deoxy-L-mannose) is one of the major constituents of both rhamnogalacturonans. Here, “L-rhamnose” is referred to as “rhamnose,” unless otherwise noted. Rhamnose is also found in the mucilage secreted from some plant genera (4), and it is a common glycone component of various plant-derived glycosides such as glycosylated flavonoids (5). Hence, rhamnose is a potential carbon source for microorganisms in the rhizosphere when it is released by the degradation of those compounds. Actu-

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ally, it was reported that B. subtilis is equipped with a set of enzymes for degrading the main chain of rhamnogalacturonan type I (2). Based on their amino acid sequences and the organization of the gene clusters that include the genes of these enzymes (yesOPQRSTUVWXYZ and yteQRST), it was assumed that unsaturated rhamnogalacturonide, which is derived from the cleavage of the main chain by two rhamnogalacturonan lyases (YesW and YesX), is incorporated into the cytosol through an ABC sugar transport system involving YesOPQ and YteQ and then hydro-

Received 19 October 2015 Accepted 15 December 2015 Accepted manuscript posted online 28 December 2015 Citation Hirooka K, Kodoi Y, Satomura T, Fujita Y. 2016. Regulation of the rhaEWRBMA operon involved in L-rhamnose catabolism through two transcriptional factors, RhaR and CcpA, in Bacillus subtilis. J Bacteriol 198:830 – 845. doi:10.1128/JB.00856-15. Editor: A. Becker Address correspondence to Kazutake Hirooka, [email protected]. * Present address: Takenori Satomura, Department of Applied Chemistry and Biotechnology, Graduate School of Engineering, University of Fukui, Fukui, Japan. Copyright © 2016, American Society for Microbiology. All Rights Reserved.

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Department of Biotechnology, Faculty of Life Science and Biotechnology, Fukuyama University, Fukuyama, Hiroshima, Japan

Regulation of Rhamnose Catabolism in B. subtilis

parentheses.

lyzed by two unsaturated galacturonyl hydrolases (YesR and YteR) to provide rhamnose and unsaturated galacturonic acid (2, 6). The family Enterobacteriaceae, such as Escherichia coli and Salmonella enterica serovar Typhimurium, catabolize rhamnose to yield dihydroxyacetone phosphate and L-lactaldehyde with an enzyme set composed of L-rhamnose isomerase (RhaA, EC 5.3.1.14), L-rhamnulose kinase (RhaB, EC 2.7.1.5), and L-rhamnulose-1phosphate aldolase (RhaD, EC 4.1.2.19) (7). In E. coli, the resultant L-lactaldehyde is further converted to L-lactic acid by L-lactaldehyde dehydrogenase (EC 1.2.1.22) or to L-1,2-propanediol by L-lactaldehyde reductase (EC 1.1.1.77) under aerobic and anaerobic conditions, respectively (8) (Fig. 1). In addition, E. coli possesses L-rhamnose mutarotase (RhaM, EC 5.1.3.32), which facilitates the interconversion of the ␣ and ␤ anomers of L-rhamnopyranose, enhancing the rate of rhamnose catabolism (9); the catalytic mechanism of mutarotation can be envisaged by examining the crystal structures of the enzymes from E. coli and Rhizobium leguminosarum (9, 10). By analogy of a proposed catalytic mechanism of Pseudomonas stutzeri L-rhamnose isomerase, E. coli RhaA might preferentially isomerize ␤-L-rhamnopyranose to ␤-Lrhamnulofuranose (11). This assumption is supported by the crystal structure of the complex of E. coli RhaB with ␤-L-rhamnulofuranose and ADP (12) (Fig. 1). As for B. subtilis, Rodionova et al. recently reported that the RhaEW (formerly YuxG) protein is a bifunctional fusion of L-rhamnulose-1-phosphate aldolase and L-lactaldehyde dehydrogenase catalyzing two consecutive steps in rhamnose catabolism (7) (Fig. 1). The rhaEW gene is a constituent of the rhaEWRBMA (formerly yuxG-yulBCDE) gene cluster, in which the rhaA, rhaB, and rhaM genes encode proteins exhibiting significant homologies with E. coli RhaA (58% identity), RhaB (43% identity), and RhaM (57% identity), respectively, whereas the rhaR gene was predicted to encode a transcriptional regulator belonging to the DeoR family (Fig. 2). A recent study revealed that these five genes constitute an operon (13) although it remained unclear how the transcription of this operon is controlled. Since most of the DeoR

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family members reported thus far act as repressors (14–18), we supposed that the transcription of this operon is primarily regulated by RhaR acting as a repressor. With respect to the transcriptional regulation of the genes involved in rhamnose catabolism, the regulatory system of E. coli is most finely elucidated (19–23). It was reported that, in the E. coli genome, genes encoding the rhamnose transporter (RhaT), the rhamnose catabolic enzymes (RhaA, RhaB, and RhaD), and the rhamnose-responsive transcriptional activators (RhaS and RhaR) are assembled to form the rhaSR and rhaBAD operons and the rhaT gene, which are located adjacent to one another. Because the rhaM gene is situated just downstream of the rhaD gene in the same direction as the rhaBAD operon, the rhaM gene might be integrated into the operon members, although it has not been experimentally confirmed (9, 19). Unlike B. subtilis RhaR, both RhaS and RhaR from E. coli belong to the AraC/XylS family and act as activators; however, they have different functions. In response to rhamnose, RhaR activates the rhaSR operon, and the resulting RhaS associated with rhamnose, in turn, activates the rhaBAD (possibly rhaBADM) operon and the rhaT gene (20–22). The cyclic AMP receptor protein (CRP) functions as a coactivator by binding simultaneously with RhaS or RhaR to each of the regulatory regions (20, 22, 23). This involvement of CRP means that the expression of the genes for rhamnose catabolism is under the carbon catabolite repression. As in E. coli, it was likely that the transcription of the genes for rhamnose catabolism is under the carbon catabolite repression in B. subtilis; however, the mechanisms underlying it completely differ between the two bacteria. In many low-GC Gram-positive bacteria, including B. subtilis, carbon catabolite control occurs on the binding of the complex of catabolite control protein A (CcpA) and the seryl-phosphorylated form of a histidine-containing protein or its homolog (P-Ser-HPr or -Crh) to catabolite-responsive elements (cre’s) of the target genes; P-Ser-HPr or P-Ser-Crh was formed upon activation of HPr kinase/phosphorylase, which is triggered by an increase in the fructose-1,6-bisphosphate concen-

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FIG 1 Bacterial rhamnose catabolic pathway via phosphorylation. The relevant enzymes encoded in the B. subtilis rhaEWRBMA operon are indicated in

Hirooka et al.

their neighboring genes are indicated by large open arrows, and the rhaEW promoter and two hairpin structures likely functioning as transcription terminators are indicated by a bent arrow and stem-loops, respectively. aa, amino acids. An enlarged view of the regulatory region of the operon is presented in the lower part. The “⫺35” and “⫺10” sequences are underlined, and the transcription start site (⫹1) and the Shine-Dalgarno (SD) sequence are enclosed in boxes. The partial coding regions of the tlpB and rhaEW genes are indicated by lines. On DNase I footprinting, the regions protected by the RhaR protein and the CcpA/P-Ser-HPr complex in the coding and noncoding strands are indicated by dark gray and light gray bars, respectively. The cre sequence is shown in boldface type. Two imperfect direct repeats situated upstream (UDR) and downstream (DDR) are indicated by two sets of tandem arrows.

tration in the presence of certain carbohydrates, such as glucose, transported through the phosphoenolpyruvate-dependent phosphotransferase system (24). We found a candidate cre sequence in the putative regulatory region upstream of rhaEW, suggesting the carbon catabolite repression of this operon mediated by CcpA. In this study, we confirmed that the transcription of the B. subtilis rhaEWRBMA operon is induced in the presence of rhamnose. The recombinant RhaR protein specifically bound to an operator site of the regulatory region upstream of rhaEW, and the DNA binding of RhaR was effectively inhibited by L-rhamnulose1-phosphate, which had been prepared from commercially available L-rhamnulose and ATP using the recombinant E. coli RhaB (RhaBEc) protein. Here, “L-rhamnulose” and “L-rhamnulose-1phosphate” are referred to as “rhamnulose” and “rhamnulose-1phosphate,” respectively, unless otherwise noted. Moreover, the binding of the CcpA/P-Ser-HPr complex to the cre sequence in the regulatory region of the operon was confirmed by the DNA binding experiment. In addition to these in vitro data, the data of in vivo analysis using the rhaEW promoter-lacZ fusion under the background of ccpA deletion and/or the disruption of any of the operon members clearly indicate that RhaR acts as a repressor for the rhaEWRBMA operon and that rhamnulose-1-phosphate, produced from rhamnose through sequential reactions by RhaA and RhaB, is an effector molecule that antagonizes the DNA binding of RhaR, as well as indicating that CcpA plays an essential role in the carbon catabolite repression of the operon. MATERIALS AND METHODS Construction and cultivation of B. subtilis strains. The B. subtilis strains used in this study are listed in Table 1. B. subtilis strain 168 was used as the

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standard strain. Strains BFS1435, YULBd, YULCd, BFS1436, and YULEd, which had been constructed by the integration of plasmid pMUTIN2 (25) into the rhaEW, rhaR, rhaB, rhaM, and rhaA genes, respectively, of strain 168, were obtained from a collection of B. subtilis strains made as part of the National BioResource Project (NIG, Japan) (26). These mutant strains were transformed with genomic DNA of strain FU652 (⌬ccpA::cat) (27) to obtain resistance to chloramphenicol, which resulted in strains FU1198, FU1199, FU1200, FU1201, and FU1202. Strain FU1203 was constructed by ectopically introducing a region around the rhaEW promoter (bases ⫺144 to 147; base 1 is the transcription start base identified in this study) fused to the lacZ reporter gene [PrhaEW(⫺144 to 147)-lacZ] into the amyE locus of strain 168 as follows.

TABLE 1 B. subtilis strains used in this study Source or reference

Strain

Genotype

168 BFS1435 YULBd YULCd BFS1436 YULEd FU652 FU1198 FU1199 FU1200 FU1201 FU1202 FU1203

trpC2 rhaEW::pMUTIN2 trpC2 rhaR::pMUTIN2 trpC2 rhaB::pMUTIN2 trpC2 rhaM::pMUTIN2 trpC2 rhaA::pMUTIN2 trpC2 ⌬ccpA::cat trpC2 ⌬ccpA::cat rhaEW::pMUTIN2 trpC2 ⌬ccpA::cat rhaR::pMUTIN2 trpC2 ⌬ccpA::cat rhaB::pMUTIN2 trpC2 ⌬ccpA::cat rhaM::pMUTIN2 trpC2 ⌬ccpA::cat rhaA::pMUTIN2 trpC2 amyE::[cat PrhaEW(⫺144 to 147)-lacZ] trpC2

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Standard strain NIG, Japan (26) NIG, Japan (26) NIG, Japan (26) NIG, Japan (26) NIG, Japan (26) 27 This study This study This study This study This study This study

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FIG 2 Organization of the rhaEWRBMA operon and its regulatory region. Organization of the operon is depicted in the upper part. The operon members and

Regulation of Rhamnose Catabolism in B. subtilis

TABLE 2 Oligonucleotide primers used in this study Sequence (5=–3=)a

PrhaEW_XF PrhaEW_BR rhaRORF_NF rhaRORF_BR rhaAORF_XF rhaAORF_BR rhaBORF_XF rhaBORF_BR EcrhaB_XF EcrhaB_BR rhaEW_NF rhaEW_NR rhaR_NF rhaR_NR rhaB_NF rhaB_NR rhaM_NF rhaM_NR rhaA_NF rhaA_NR rhaEW_PE PrhaEW_delUF PrhaEW_delUR PrhaEW_delDF PrhaEW_delDR

ACATGGCTCTAGAACTTCAGGACAT CCCGGATCCAGTTGCAGACAGTGCG GAAGGGCGTTACATATGCTAGTAGC GGCAGTATAAAGGATCCTTTCACC GAGAGCTCGAGATGACCATAAAAGC TGCGGATCCGGATTTTTTATTAGAC GGGTCTCGAGATGATTTATACTGCC TTTGGATCCTTTTCACCTCTTCATC CAGCAGGATCTCGAGATGACCTTTC TCCCAGGCCGGATCCAGTTGAGTG GACCGCACTGTCTGCAACTGG GTCAACATGTTTGTATGGCAGAA GTCACAGAGGAAACCATTCGGC GCTGAGTTCAATAGCTGCTTTC CACAGGTTCGCTAACGGTTTCAG GTTGAAAGGCTGAAATTGTATTC TTGAAAAGAAAAGCCAGTATCATG AACTTCCTTTAGATCTATCGCAAC GTGCCCATTTCGATCCACTGC AAAATCTAAACCGAGACCAAGGTT GATCAGATTGGACCTGTACACC CTGCTTTCTTTTTTTGTTAAAAAGATATAATGAAAACGGATACAAA TTTGTATCCGTTTTCATTATATCTTTTTAACAAAAAAAGAAAGCAG GATTCAAAAACAAACAGAAATCATTATGGAGGAATATCAAATATGG CCATATTTGATATTCCTCCATAATGATTTCTGTTTGTTTTTGAATC

a

Restriction sites are underlined.

The corresponding DNA fragment was amplified by PCR with genomic DNA of strain 168 and primer pair PrhaEW_XF and PrhaEW_BR (Table 2), followed by trimming with XbaI and BamHI digestion. It was then cloned into the pCRE-test2 vector (28) that had been treated with the same restriction enzymes. Correct construction was confirmed by DNA sequencing. The resulting plasmid was linearized by PstI digestion and then integrated into the amyE locus of strain 168 through double-crossover transformation to obtain chloramphenicol resistance, which resulted in strain FU1203. B. subtilis cells were pregrown at 30°C overnight on tryptose blood agar base (BD Diagnostic Systems) plates supplemented with 0.18% glucose, which contained chloramphenicol (5 ␮g/ml) and/or erythromycin (0.3 ␮g/ml), according to the drug resistance of the cells. The cells were inoculated into a minimal medium (29) supplemented with a mixture of 16 amino acids (glutamine, histidine, tyrosine, and asparagine were omitted) (30) (MM⫹16aa) to give an optical density at 600 nm (OD600) of 0.05 and then incubated at 37°C with shaking. This medium contained glucose at 25 mM as a carbon source. The cells were also cultivated by using similar media, in which the 25 mM glucose was replaced with 25 mM rhamnose, xylose, or malic acid. When the strains carrying the pMUTIN2 integration were cultivated, expression of the genes located downstream of the pMUTIN2-inserted position was induced by adding isopropyl-␤D-thiogalactopyranoside (IPTG) to the medium to a final concentration of 1 mM before inoculation (25). Preparation of the recombinant proteins. The B. subtilis rhaR, rhaA, and rhaB coding regions were amplified by PCR with genomic DNA of strain 168 and the primer pairs rhaRORF_NF/rhaRORF_BR, rhaAORF _XF/rhaAORF_BR, and rhaBORF_XF/rhaBORF_BR, respectively, and the E. coli rhaB coding region was amplified by PCR with genomic DNA of strain DH5␣ and the primer pair EcrhaB_XF/EcrhaB_BR (Table 2). The rhaR fragment was digested with NdeI/BamHI and then cloned into the pColdIV vector (TaKaRa Bio, Japan), which had been treated with the same restriction enzymes, to yield expression plasmid pCold-rhaR. The other three fragments were digested with XhoI/BamHI and then cloned into the pColdI vector (TaKaRa Bio), which had been treated with the

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same restriction enzymes, to yield expression plasmids pCold-rhaA, pCold-rhaB, and pCold-EcrhaB. These pColdI-based plasmids were designed to produce each protein as an N-terminal in-frame fusion with the His6 tag (His-RhaA, His-RhaB, and His-RhaBEc). The correct cloning of each gene was confirmed by DNA sequencing. E. coli strain BL21 (Merck Millipore), transformed with pCold-rhaR, was grown with shaking in a Luria-Bertani medium (31) supplemented with ampicillin (50 ␮g/ml) at 37°C to an OD600 of 0.4. The culture was then refrigerated at 15°C and left to stand for 30 min. After IPTG had been added to a final concentration of 1 mM, shaking of the culture was resumed at 15°C and continued for another 24 h. The cells harvested from 6 liters of the culture were disrupted by sonication in buffer 1 (20 mM Tris-Cl buffer [pH 8.0] plus 10% [vol/vol] glycerol, 0.1 mM phenylmethylsulfonyl fluoride, and 1 mM dithiothreitol). After centrifugation (17,000 ⫻ g, 4°C, 20 min) and filtration (0.45-␮m pore size), the supernatant was subjected to ammonium sulfate precipitation. The fraction precipitated at 30 to 50% saturation was recovered by centrifugation (17,000 ⫻ g, 4°C, 20 min) and dialyzed against buffer 1 and then applied to a Toyopearl DEAE-650M column (Tosoh, Japan) equilibrated with buffer 1. The column was washed with buffer 1 and eluted with a gradient of 0 to 1 M NaCl. The RhaR fraction was collected and concentrated by ultrafiltration (Amicon Ultra-4 10K; Merck Millipore) and then subjected to gel filtration (Sephacryl S-200 HR; GE Healthcare) with buffer 1 containing 0.1 M NaCl at a flow rate of 0.8 ml/min. The RhaR fraction was collected and concentrated by ultrafiltration. To determine the molecular mass of the native form of RhaR, low-molecular-weight and high-molecular-weight gel filtration calibration kits (GE Healthcare) were used for the column calibration. E. coli strain BL21, bearing pCold-rhaA, pCold-rhaB, and pColdEcrhaB, respectively, was cultivated as mentioned above, from which crude lysates were similarly prepared. Each of the crude lysates, obtained from 500 ml of the culture, was subjected to a HisTrap FF column (1 ml, precharged with Ni2⫹; GE Healthcare) equilibrated with buffer 2 (20 mM Tris-Cl buffer [pH 8.0] plus 0.5 M NaCl) containing 10 mM imidazole. The column was washed with the same buffer and eluted with a stepwise

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Primer

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with genomic DNA of strain 168 and the primer pair PrhaEW_XF/ PrhaEW_BR. DNase I footprinting analysis. DNase I footprinting analysis was performed as described previously (35). The DNA probe for the footprinting was prepared by PCR with genomic DNA of strain 168 and the primer pair PrhaEW_XF/rhaEW_PE. Prior to PCR amplification, the 5= terminus of only one of the primers was labeled with [␥-32P]ATP, using a Megalabel kit. The DNA probe (0.04 pmol), labeled at the 5= end, was mixed with the RhaR protein and/or the CcpA/P-Ser-HPr proteins to obtain a DNAprotein complex, which was then partially digested with DNase I (TaKaRa Bio) in 50 ␮l of a reaction mixture and subjected to urea-PAGE with sequencing ladders. These ladders were prepared using the same template PCR product that was used for ladders of the primer extension analysis and each of the 5=-end-labeled primers (PrhaEW_XF and rhaEW_PE). Gel retardation analysis. Gel retardation analysis was performed essentially as described previously (36). The radiolabeled DNA probe (PrhaEW probe) that corresponds to the region around the rhaEW promoter (bases ⫺144 to 147) was prepared by PCR with genomic DNA of strain 168 and the primer pair PrhaEW_XF and PrhaEW_BR in the presence of [␣-32P]dCTP. As the XbaI and BamHI sites were adducted at the 5= and 3= ends, respectively, the length of the PrhaEW probe was 308 bp. To create two derivatives of the PrhaEW probe (PrhaEW_⌬UDR and PrhaEW_⌬DDR probes) that lack either of two imperfect direct repeats (the upstream and the downstream repeats were designated UDR and DDR, respectively), recombinant PCR (37) was performed with the internal overlapping primer pairs PrhaEW_delUF/PrhaEW_delUR and PrhaEW_delDF/PrhaEW_delDR, respectively (Table 2), together with the outer primer pair PrhaEW_XF/PrhaEW_BR and the genomic DNA of strain 168. The resultant two fragments were cloned into the pCR2.1TOPO vector (Life Technologies), which were confirmed by DNA sequencing to lack the UDR and DDR sequences correctly, and then used as the template for PCR with [␣-32P]dCTP and the primers PrhaEW_XF and PrhaEW_BR to produce the radiolabeled PrhaEW_⌬UDR and PrhaEW_⌬DDR probes, respectively. Each of the DNA probes (0.02 pmol) was mixed and incubated at 30°C for 10 min with various amounts of the RhaR protein in a 25-␮l binding mixture, and then the mixture was subjected to native PAGE. To evaluate the effect on the DNA binding of RhaR, rhamnulose-1phosphate and a phosphorylated rhamnulose were prepared using HisRhaBEc and His-RhaB, respectively, as described below. A total of 5 ␮l of each of the solutions, including these compounds, was added to the binding mixture (the total volume was adjusted to 25 ␮l), followed by native PAGE. Measurement of the enzymatic activities of His-RhaA, His-RhaB, and His-RhaBEc and preparation of rhamnulose derivatives. The rhamnose isomerization activity of His-RhaA was measured by a modification of a method described previously (38). The reaction mixture contained, in a total volume of 500 ␮l, 50 mM Tris-Cl (pH 8.0), 1 mM MnCl2, 10 mM L-rhamnose (Sigma-Aldrich), and 0.1 to 1 ␮g of the purified His-RhaA. A 450-␮l portion of the mixture without rhamnose was preincubated at 30°C for 3 min, and the reaction was initiated by adding 50 ␮l of 100 mM rhamnose solution. After incubation at 30°C for 20 min, the reaction was terminated by adding 50 ␮l of 10% trichloroacetic acid solution. A part of the mixture (110 ␮l; 1/5 volume) was taken and mixed with 20 ␮l of 1.5% cysteine-HCl solution, 600 ␮l of 70% (vol/vol) H2SO4 solution, and 20 ␮l of 0.12% carbazole solution dissolved with ethanol, followed by incubation at 35°C for 20 min for color development. By measuring the absorbance of the colored mixture at 545 nm, the amount of ketoses converted from rhamnose was quantified, based on the molar extinction coefficient at 545 nm (1,530 M⫺1 cm⫺1) determined by using L-rhamnulose (SigmaAldrich), which was dissolved at various concentrations with 50 mM Tris-Cl buffer (pH 8.0) plus 1 mM MnCl2, and colored by the same treatment for spectrophotometric measurement. The rhamnulose phosphorylation activity of His-RhaB and HisRhaBEc was measured as follows. The reaction mixture contained, in a

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gradient of 20 to 500 mM imidazole in buffer 2. The fractions of each of the recombinant proteins were collected and, as for His-RhaB and HisRhaBEc, the protein solutions were concentrated by ultrafiltration. B. subtilis CcpA and P-Ser-HPr proteins were prepared as reported previously (32). The CcpA protein, produced by E. coli strain JM109 bearing plasmid pCCPA19, was purified by two column chromatography steps (Toyopearl DEAE-650M and heparin-Sepharose 6FF [GE Healthcare]). The HPr protein, produced by E. coli strain BL21(DE3) (Merck Millipore) bearing plasmid pET-ptsH, was purified by one column chromatography step (Toyopearl DEAE-650M). An N-terminal in-frame fusion of B. subtilis HPr kinase/phosphorylase with the His6 tag (His-HPrK), produced by E. coli strain BL21(DE3) bearing plasmid pET-hprK, was purified by affinity column chromatography (Ni-nitrilotriacetic acid [NTA]-agarose; Qiagen). Ser-46 of the HPr protein was phosphorylated in vitro to form P-Ser-HPr by using the His-HPrK, and then the kinase was removed from the reaction mixture by its passage through an Ni-NTAagarose column. The purity of each of the recombinant proteins was evaluated by SDSPAGE with Coomassie brilliant blue (CBB) staining. His-RhaA, His-RhaBEc, HPr, and His-HPrK proteins were confirmed to have been purified to near homogeneity, while the purities of RhaR, His-RhaB, and CcpA proteins were estimated to be ca. 90%, as judged by measuring the band intensity on a gel using ImageJ (http://rsbweb.nih.gov/ij/). The percentage of phosphorylated HPr was estimated to be ca. 80%, which was calculated by measuring the band intensities of the phosphorylated and unphosphorylated HPrs on a gel of native PAGE with CBB staining; the mobility of the protein increased with increase in negative charge by the phosphorylation (data not shown). Protein concentrations were determined by the Bradford method with bovine serum albumin as the standard using a protein assay kit (Bio-Rad). RhaR, His-RhaA, CcpA, and P-Ser-HPr proteins were stored at ⫺80°C in buffers containing 5 to 20% (vol/vol) glycerol, while His-RhaB, His-RhaBEc, and His-HPrK solutions were stored at 4°C. It should be noted that, for dilution of the RhaR and His-RhaA protein solutions, buffers (50 mM Tris-Cl buffer [pH 7.5] plus 10% [vol/vol] glycerol and 0.1 mM dithiothreitol for RhaR; 50 mM Tris-Cl buffer [pH 8.0] for His-RhaA) containing 0.1% (vol/vol) Tween 20 were used; otherwise, the functional activities of these proteins were severely impaired. Northern blot analysis. For the analysis of transcripts from the rhaEW promoter, total RNAs were extracted and purified as previously reported (33) from the cells of B. subtilis strain 168 grown in MM⫹16aa and in a similar medium in which glucose was replaced with rhamnose. The cells were harvested when the OD600 of the culture reached 1.0. The RNA samples (10 ␮g) were electrophoresed in a glyoxal gel and transferred to a Hybond-N membrane (GE Healthcare). To prepare probes to detect transcripts carrying rhaEW, rhaR, rhaB, rhaM, and rhaA, DNA fragments (each 300 bp) were obtained by PCR with genomic DNA of strain 168 and primer pairs (rhaEW_NF/rhaEW_NR, rhaR_NF/rhaR_NR, rhaB_NF/ rhaB_NR, rhaM_NF/rhaM_NR, and rhaA_NF/rhaA_NR) (Table 2) and labeled using a BcaBEST labeling kit (TaKaRa Bio) and [␣-32P]dCTP (MP Biomedicals). The radiolabeled DNA probes were hybridized to the RNAs blotted onto the membrane, which was then washed as described previously (31). The autoradiograms, as well as those described below, were obtained and quantified using a Typhoon 9400 variable mode imager (GE Healthcare). Primer extension analysis. To determine the transcription start site from the rhaEW promoter, primer extension analysis was performed using the same RNA samples that were prepared for the Northern blot analysis. The RNA samples (45 ␮g) were annealed to 1 pmol of primer rhaEW_PE (Table 2), which had been 5= end labeled with a Megalabel kit (TaKaRa Bio) and [␥-32P]ATP (MP Biomedicals), and the primer extension reaction was then conducted with ThermoScript reverse transcriptase (Life Technologies), as described previously (34). For the dideoxy sequencing reaction for ladder preparation starting from the 5=-endlabeled rhaEW_PE primer, a template DNA was generated by PCR

Regulation of Rhamnose Catabolism in B. subtilis

MM⫹16aa containing glucose at 25 mM (lane 1) and from those grown in a similar medium, in which 25 mM glucose was replaced by 25 mM rhamnose (lane 2), and Northern blotting was performed. The blotted membranes were stained once with methylene blue to ensure equal amounts of the RNA samples; the bands corresponding to 16S rRNA are shown underneath. 32P-labeled DNA probes specific to each coding region (rhaEW, rhaR, rhaB, rhaM, and rhaA probes) were used. The positions of the RNA size markers (RNA Millennium markers; Life Technologies) are indicated by the arrows on the left.

total volume of 500 ␮l, 50 mM Tris-Cl (pH 8.0), 1 mM MnCl2, 1 mM MgCl2, 10 mM rhamnulose, 1 mM ATP, and 0.2 to 1 ␮g of the purified His-RhaB or 10 to 20 ng of the purified His-RhaBEc. A 450-␮l portion of the mixture without ATP was preincubated at 30°C for 3 min, and the reaction was initiated by adding 50 ␮l of 10 mM ATP solution. After incubation at 30°C for 30 min, the enzyme was inactivated by heat treatment at 95°C for 10 min, and then the mixture was quickly chilled on ice. Then, 40 ␮l of the 40-fold diluted mixture was mixed with 80 ␮l of a working reagent of the EnzyChrom kinase assay kit (BioAssay Systems) on a microtiter plate. After incubation at room temperature for 20 min, the fluorescence intensity was measured with the Typhoon 9400 variable mode imager (␭ex ⫽ 532 nm and ␭em ⫽ 580 nm) to detect ADP generated as a by-product. Because of the range of detection, the ATP concentration in the reaction mixture had been adjusted to be lower than that of rhamnulose. The amount of ADP production was quantified based on a standard curve given by a series of ADP standard solutions, by which the phosphorylation activity was calculated. To examine the substrate and product specificities of the enzymes, reaction mixtures with various combinations of the enzyme and substrate components were prepared which contained, in a total volume of 500 ␮l, 50 mM Tris-Cl (pH 8.0), 1 mM MnCl2, 1 mM MgCl2, 1 mM ATP, 10 mM concentrations of either sugar substrate (rhamnulose or rhamnose), and 10 ␮g of either kinase (His-RhaBEc or His-RhaB) with or without 1 ␮g of His-RhaA. The 450 ␮l of the mixture without ATP was preincubated at 30°C for 3 min, and the reaction was initiated by adding 50 ␮l of 10 mM ATP solution. Incubation of the reaction mixtures, heat inactivation of the enzyme(s), and detection of ADP production were performed as described above. For the gel retardation analysis, the preparation of rhamnulose-1phosphate and a phosphorylated rhamnulose was performed by using His-RhaBEc and His-RhaB, respectively, as follows. The reaction mixture contained, in a total volume of 250 ␮l, 50 mM Tris-Cl (pH 8.0), 1 mM MnCl2, 1 mM MgCl2, 50 mM ATP, 25 mM rhamnulose, and 20 ␮g of His-RhaBEc or His-RhaB. After incubation at 30°C for 18 h, followed by heat treatment (95°C, 15 min) and removal of the denatured proteins by centrifugation (17,000 ⫻ g, 4°C, 20 min), 200 ␮l of each supernatant was lyophilized, and the pellet was dissolved with 20 ␮l of ultrapure water to yield a 10-fold-condensed solution, 5 ␮l of which was added to the binding mixture of the gel retardation analysis to test the effect of these rhamnulose derivatives on the DNA binding of RhaR. To prepare the solution used as a control, 250 ␮l of the mixture containing 50 mM Tris-Cl (pH 8.0), 1 mM MnCl2, 1 mM MgCl2, 25 mM ATP, 25 mM rhamnulose, 25 mM ADP, 25 mM sodium phosphate, and 20 ␮g of either kinase, which

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had been heat treated at 95°C for 1 h for complete inactivation, was subjected to the same incubation and condensation processes. lacZ reporter analysis. B. subtilis strains carrying the promoter-lacZ fusion were grown in 50 ml of the MM⫹16aa and similar media containing either rhamnose, xylose, or malic acid instead of glucose at 37°C with shaking. After the OD600 reached 0.2, 1-ml aliquots of the culture were withdrawn at intervals of 1 h, and the ␤-galactosidase (␤-Gal) activity in crude cell extracts was spectrophotometrically measured using o-nitrophenyl-␤-D-galactopyranoside (Wako Pure Chemical Industries, Japan) as a substrate, according to the procedure described previously (30).

RESULTS

Northern blot analysis of transcripts carrying rhaEW, rhaR, rhaB, rhaM, and rhaA. Although it was reported that B. subtilis rhaEW, rhaR, rhaB, rhaM, and rhaA genes constitute an operon (13), it remained to be clarified how the transcription of this operon is controlled. Among the encoded proteins, only the function of RhaEW was experimentally confirmed to be a bifunctional enzyme catalyzing two sequential reactions of aldol cleavage and dehydrogenation, a downstream part of the rhamnose catabolic pathway after phosphorylation (7) (Fig. 1). A homology search predicted that the rhaB, rhaM, and rhaA genes encode enzymes involved in the steps upstream of the aldol cleavage, whereas the rhaR gene was predicted to encode a DeoR family transcriptional regulator. Hence, we supposed that the transcription of this operon is primarily regulated by RhaR and is induced when the cells are grown on rhamnose. We performed Northern blot analysis using the specific DNA probes and total RNAs that were prepared from the cells of strain 168 grown in MM⫹16aa, which contained glucose as a carbon source, and from cells grown in a similar medium, to which rhamnose was added instead of glucose. Bands corresponding to a 6-kb transcript were distinctly detected when each DNA probe was used with the RNA of cells grown in the rhamnose-containing medium but not with the RNA of those grown in the glucose-containing medium (Fig. 3). This result clearly indicates that the transcription is inducible by the presence of rhamnose, as well as confirming the organization of the operon. The extra bands with weak intensities could be derived from partial degradation of the 6-kb transcript. Determination of the transcription start site of the rhaEW promoter. To locate the transcription start site of the rhaEW pro-

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FIG 3 Northern blot analysis of transcripts carrying rhaEW, rhaR, rhaB, rhaM, and rhaA. RNA samples were prepared from the cells of strain 168 grown in the

Hirooka et al.

the rhaEW promoter. RNA samples from the cells of strain 168 grown in the MM⫹16aa containing glucose at 25 mM (lane 1) and from those grown in a similar medium containing rhamnose instead of glucose at 25 mM (lane 2) were used for the reverse transcription reaction to generate the runoff cDNA. Lanes G, A, T, and C contain products of the dideoxy sequencing reactions with the same primer that was used for reverse transcription. The runoff cDNA band is indicated by an arrow. The partial nucleotide sequence of the coding strand complementary to the ladders is shown, where the “⫺10” sequence is underlined, the transcription start site (⫹1) and the SD sequence are enclosed in boxes, and the cre sequence is shown in boldface type.

moter, we performed primer extension analysis using the same RNA samples that were used for the Northern blot analysis. As shown in Fig. 4, the specific band corresponding to runoff cDNA of the rhaEWRBMA transcript was detected with the RNA sample of the cells grown in the rhamnose-containing medium but not with the RNA of those grown in the glucose-containing medium; this supports the idea that the transcription of the operon is inducible by the presence of rhamnose. Based on the location of the transcription start site, we predicted the “⫺35” and “⫺10” sequences of the promoter, with a 17-bp spacer, to be TTCAAA and TATAAT (Fig. 2), which is likely recognized by the ␴A-RNA polymerase (39). Identification of the binding sites of RhaR and CcpA/P-SerHPr in the regulatory region of the rhaEWRBMA operon. To identify the RhaR binding site around the rhaEW promoter, DNase I footprinting analysis was performed by using the recombinant RhaR protein and the DNA probe corresponding to the region, including the rhaEW promoter (bases ⫺144 to 120; base 1 is the transcription start base). The RhaR protein was overexpressed in the E. coli cells and purified to be approximately homogeneous by ammonium sulfate precipitation, followed by two column chromatography steps. Upon gel filtration with protein standards to obtain the molecular mass, the RhaR protein exhibited a molecular mass of 68.9 kDa (data not shown). Since the molecular mass of the RhaR monomer is calculated to be 28.9 kDa, it was estimated that RhaR forms a dimer. When the RhaR protein was mixed with the DNA probe, followed by DNase I digestion and electrophoresis, a somewhat broad region protected against DNase I was detected, which covers the ⫺35 and ⫺10 sequences and the transcription start base (bases ⫺58 to 21 of the coding strand and bases ⫺57 to ⫺15/⫺5 to 20 of the noncoding strand) (Fig. 5, lanes 2 and 3). We found two imperfect direct repeats in the protected region, either or both of which were expected to be important for RhaR recognition. The

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FIG 4 Primer extension analysis to determine the transcription start site from

upstream and downstream direct repeats (UDR and DDR) are located successively and are composed of the 20-1-20 and 15-15 arrangements, respectively (UDR, CAAAAAtAAACAaAAAcgATT-CAAAAAcAAACAgAAAtcAT; DDR, ATAtAAtgAAAAcGG-A TAcAAatAAAAgGG; mismatched bases in the respective direct repeats are indicated by lowercase letters). A candidate cre sequence (ATGAAAACGGATACA), matched well with the consensus sequence of one subgroup of cre’s (WTG NAANCGNWWNCA; W and N stand for A or T and any base) (24), was found in the rhaEW promoter region, which includes the partial ⫺10 sequence and the transcription start base (Fig. 2). We examined whether the CcpA/P-Ser-HPr complex actually binds to this sequence by means of DNase I footprinting using the same DNA probe with recombinant CcpA and P-Ser-HPr proteins, prepared as reported previously (32). The CcpA/P-Ser-HPr complex, composed of two copies of each protein, was formed upon their addition at an equimolar ratio to the reaction mixture; the proportion of phosphorylated HPr was 80% of the total HPr protein. As a result, a region covering the candidate cre sequence was protected against DNase I upon the CcpA/P-Ser-HPr addition (bases ⫺7 to 16 of the coding strand and bases ⫺12 to 12 of the noncoding strand), indicating the specific interaction of the complex with this sequence as a cre (Fig. 5, lanes 4 and 5). As the newly identified cre sequence is included in the RhaR binding region, we examined whether RhaR and the CcpA/P-SerHPr complex are capable of binding simultaneously to the regulatory region by means of DNase I footprinting performed with the same DNA probe under the coexisting conditions of both regulatory factors. The patterns of the protected regions under these conditions were found to be identical to those obtained by RhaR alone, with respect to both of the coding and noncoding strands (Fig. 5, lanes 2, 3, 6, and 7). As for the noncoding strand, a part of the protected region observed in the presence of CcpA/ P-Ser-HPr alone was dissipated under the coexisting conditions, suggesting that, when both regulatory factors are active, RhaR predominantly binds to the regulatory region of the rhaEW promoter with the interruption of CcpA/P-Ser-HPr binding to the cre. Determination of the DNA element essential for the specific binding of RhaR. The RhaR binding site in the regulatory region of the rhaEW promoter, assigned by DNase I footprinting analysis, contains two imperfect direct repeats, i.e., UDR and DDR. The importance of UDR and DDR on the specific DNA binding of RhaR was evaluated by gel retardation analysis using various concentrations of the RhaR protein combined with the DNA probe corresponding to the region around the rhaEW promoter (PrhaEW probe; 308 bp, bases ⫺144 to 147 plus restriction sites at both ends) and two PrhaEW-probe derivatives carrying either the UDR deletion (PrhaEW_⌬UDR probe; 267 bp) or the DDR deletion (PrhaEW_⌬DDR probe; 278 bp). When the RhaR protein was mixed with the PrhaEW probe, band retardation was observed on a PAGE gel dependent on the RhaR concentration, which indicates that the RhaR protein specifically bound to this DNA probe. The apparent dissociation constant (Kd) value for the PrhaEW probe was estimated to be 31 ⫾ 3 nM (as a dimer; mean value ⫾ the standard deviation in two independent experiments) (Fig. 6, left), which was obtained by quantifying the band intensities corresponding to the RhaR-DNA complex and the free DNA probe in each lane. In contrast, when the PrhaEW_⌬UDR probe was used, no band retardation was detected, clearly demonstrating

Regulation of Rhamnose Catabolism in B. subtilis

rhaEWRBMA operon. A 5=-end-32P-labeled DNA probe (0.04 pmol), corresponding to the region around the rhaEW promoter, was incubated with each or both of the transcriptional regulators in the reaction mixture (50 ␮l), followed by partial digestion with DNase I and urea-PAGE. The products of the mixtures containing RhaR at 0.12 and 0.059 ␮M as a dimer were applied to lanes 2 and 3, respectively. The products of the mixture containing CcpA at 6.0 ␮M and HPr (phosphorylated and nonphosphorylated forms) at 7.5 ␮M and of the mixture containing CcpA at 3.0 ␮M and HPr at 3.8 ␮M, each as a monomer, were applied to lanes 4 and 5; the mixtures were assembled to contain the CcpA/P-Ser-HPr heterotetramer at 3.0 and 1.5 ␮M, respectively. The products of the mixtures containing both the RhaR protein and the CcpA/P-Ser-HPr complex were applied to lanes 6 and 7. The concentrations of the RhaR dimer and the CcpA/P-SerHPr complex were 0.12 and 3.0 ␮M in the mixture of lane 6 and 0.059 and 1.5 ␮M in the mixture of lane 7, respectively. The product of the mixture containing no recombinant protein was applied to lanes 1 and 8. To lanes G, A, T, and C, the products of the dideoxy sequencing reactions with the same 5=-labeled primers were applied. The regions protected by RhaR and CcpA/P-Ser-HPr are indicated on the right of each panel by dark gray and light gray bars, respectively. The “⫺35” and “⫺10” sequences are underlined, the transcription start site (⫹1) and the SD sequence are enclosed in boxes, and the cre sequence is shown in boldface type. The UDR and the DDR are indicated by two sets of tandem arrows.

that the UDR sequence is essential for the specific RhaR binding (Fig. 6, right). On the other hand, the combination of RhaR and the PrhaEW_⌬DDR probe resulted in band retardation due to the specific DNA binding, by which the Kd value for this probe was estimated to be 34 ⫾ 3 nM (Fig. 6, center). Since this Kd value was comparable to that obtained for the PrhaEW probe, the DDR deletion does not appear to affect significantly the DNA binding affinity of RhaR. However, the difference in mobility between the retarded bands corresponding to the RhaR-PrhaEW complex and the RhaR-PrhaEW_⌬DDR complex was distinctly larger than that between the bands of the two DNA probes alone. We speculate that the DDR deletion altered the DNA binding mode of RhaR, involving conformation of the RhaR-DNA complex or the number of the RhaR dimers binding to the DNA; therefore, the RhaRPrhaEW_⌬DDR complex migrated faster than did the RhaRPrhaEW complex. Determination of an effector molecule that antagonizes the DNA binding of RhaR. Most of the DeoR-type transcriptional regulators reported thus far act as repressors in sugar and nucleoside metabolism, and the effector molecules that antagonize their DNA binding are phosphorylated intermediates in the metabolic pathways that they control (14–18). By analogy with these common features, together with the location of the RhaR binding site (Fig. 2), we assumed that the binding of RhaR to the regulatory region of the rhaEWRBMA operon, acting as a repressor, is specifically inhibited by rhamnulose-1-phosphate, a phosphorylated

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intermediate in the B. subtilis rhamnose catabolic pathway (Fig. 1). To test this, gel retardation analysis was performed using binding mixtures, to which rhamnulose-1-phosphate was or was not added. Rhamnulose was phosphorylated at the C-1 position with ATP by E. coli L-rhamnulose kinase fused to the His6 tag at the N terminus (His-RhaBEc). The catalytic mechanism of this enzyme has been established in previous studies (12, 40). The specific activity of His-RhaBEc was determined to be 224 ⫾ 10 ␮mol min⫺1 mg⫺1 (mean value ⫾ the standard deviation in three independent experiments), which was somewhat higher than that of the tagfree RhaBEc obtained in the previous study (90 ␮mol min⫺1 mg⫺1) (40), likely due to differences in the reaction mixtures and the methods of measuring enzymatic activity. After formation of rhamnulose-1-phosphate, the reaction mixture was concentrated, and then an aliquot was added to a binding mixture containing a 0.12 ␮M concentration of the RhaR dimer and a 0.8 nM concentration of the PrhaEW probe, which were sufficient to cause complete band retardation under normal conditions. By measuring the amount of ADP given as a by-product using a kinase assay kit, it was estimated that phosphorylation of rhamnulose by His-RhaBEc was achieved with almost 100% efficiency and that rhamnulose-1-phosphate was present at 50 mM in the binding mixture for gel retardation. In addition to rhamnulose-1-phosphate, several substances such as ADP, ATP, and the residual heat-treated HisRhaBEc were present in the binding mixture. Thus, we prepared a mixture containing rhamnulose, ATP, ADP, sodium phosphate,

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FIG 5 DNase I footprinting analysis to identify the binding sites of the RhaR protein and the CcpA/P-Ser-HPr complex in the regulatory region of the

Hirooka et al.

rhaEW promoter (PrhaEW probe) and two PrhaEW derivatives lacking either the upstream direct repeat (PrhaEW_⌬UDR probe) or the downstream direct repeat (PrhaEW_⌬DDR probe) were 32P labeled, and 0.02 pmol of each probe was incubated in the binding mixture (25 ␮l) with various concentrations of the RhaR protein and then subjected to PAGE. The RhaR protein solution was diluted stepwise 2-fold, and an aliquot of each dilution was added to the binding mixture to obtain the concentrations used (as a dimer). The “no protein” lanes indicate that the applied mixture contained no RhaR protein. The bars indicate the bands obtained by the RhaR protein at the concentration around the Kd value. The experiments were repeated at least two times, and representative results are shown. The binding mixtures containing the PrhaEW and PrhaEW_⌬DDR probes were applied onto the same gel to compare the mobilities of the complex of RhaR with the PrhaEW probe and that with the PrhaEW_⌬DDR probe.

and the residual heat-inactivated His-RhaBEc, which simulated that the ␥-phosphate group of ATP was merely released by hydrolysis at an equimolar amount to rhamnulose without phosphoryl transfer to it. An aliquot of this control mixture was added to

FIG 7 Gel retardation analysis to evaluate the inhibitory effects of rhamnulose-1-phophate (A) and the phosphorylated rhamnulose produced by the His-RhaB-catalyzed reaction (B) on the DNA binding of RhaR. The 32P-labeled PrhaEW probe (0.02 pmol) and the RhaR protein (3 pmol as a dimer) were incubated in the binding mixture (25 ␮l) and then subjected to PAGE, which is sufficient to cause complete band retardation under normal conditions. (A) In addition to the PrhaEW probe and the RhaR protein, the binding mixture applied to lane 2 contained 50 mM rhamnulose-1-phophate, 50 mM ATP, 50 mM ADP, and the residual heat-treated His-RhaBEc, whereas the mixture applied to lane 3 contained 50 mM rhamnulose, 50 mM ATP, 50 mM ADP, 50 mM sodium phosphate, and the residual heat-treated His-RhaBEc. The binding mixtures applied to lanes 1 and 4 contained neither RhaR protein nor any substance added to the mixtures of lanes 2 and 3. (B) In addition to the PrhaEW probe and the RhaR protein, the binding mixture applied to lane 2 contained 23 mM phosphorylated rhamnulose given by His-RhaB, 27 mM rhamnulose, 77 mM ATP, 23 mM ADP, and the residual heat-treated HisRhaB, whereas the mixture applied to lane 3 contained 50 mM rhamnulose, 50 mM ATP, 50 mM ADP, 50 mM sodium phosphate, and the residual heattreated His-RhaB. The binding mixtures applied to lanes 1 and 4 contained neither RhaR protein nor any substance added to the mixtures of lanes 2 and 3.

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another binding mixture, which was subjected to PAGE, along with a binding mixture containing rhamnulose-1-phosphate formed by His-RhaBEc. As shown in Fig. 7A, the addition of the reaction mixture containing rhamnulose-1-phosphate significantly inhibited the DNA binding of RhaR (lane 2), whereas it was not inhibited by the addition of the control mixture (lane 3). This result clearly demonstrates that the DNA binding of RhaR was antagonized by rhamnulose-1-phoshate but not by rhamnulose, phosphate, ATP, or ADP. We also confirmed that rhamnose had no effect on the DNA binding affinity of RhaR by means of similar gel retardation analysis (data not shown). Functional analyses of B. subtilis RhaA and RhaB. It was assumed that, in the B. subtilis cells, rhamnulose-1-phosphate is provided by the isomerization of rhamnose to rhamnulose catalyzed by RhaA and the subsequent phosphorylation of rhamnulose with ATP catalyzed by RhaB. To verify the functional roles of RhaA and RhaB, their N-terminal fusions with the His6 tag (His-RhaA and His-RhaB) were prepared, and their enzymatic activities were measured. After the incubation of rhamnose with His-RhaA, the resultant product was clearly colored by the cysteine-carbazole-sulfuric acid method, indicating that rhamnose was converted to a ketose. Based on the molar extinction coefficient obtained by rhamnulose, the specific activity of HisRhaA for rhamnose was determined to be 58.5 ⫾ 6.2 ␮mol min⫺1 mg⫺1 (mean value ⫾ the standard deviation in three independent experiments), which was comparable to the value for the rhamnose of Mesorhizobium loti L-rhamnose isomerase (64.5 ␮mol min⫺1 mg⫺1) (41) and somewhat lower than the value for the rhamnose of P. stutzeri L-rhamnose isomerase (280 ␮mol min⫺1 mg⫺1) (38). As shown in Table 3, His-RhaBEc specifically phosphorylated rhamnulose, and it hardly accepted rhamnose as the substrate (I and II). We examined whether the ketose derived from rhamnose by His-RhaA is acceptable to His-RhaBEc as the substrate. When the reaction was carried out with a combination

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FIG 6 Gel retardation analysis to identify a sequence element essential for the specific binding of RhaR. A DNA probe corresponding to the region around the

Regulation of Rhamnose Catabolism in B. subtilis

TABLE 3 Relative phosphorylating activities of His-RhaBEc in the presence of various factorsa Reaction no.

His-RhaBEc

His-RhaA

Rhamnose

Rhamnulose

ATP

Mean relative activity (%) ⫾ SDb

I II III IV

⫹ ⫹ ⫹ –

– – ⫹ ⫹

– ⫹ ⫹ ⫹

⫹ – – –

⫹ ⫹ ⫹ ⫹

100.0 ⫾ 29.9 1.18 ⫾ 1.01 30.0 ⫾ 3.7 ND

a Each reaction was carried out at 30°C for 30 min in the presence (⫹) or absence (–) of 10 ␮g of His-RhaBEc, 1 ␮g of His-RhaA, 10 mM rhamnose, 10 mM rhamnulose, and 1 mM ATP in a total volume of 500 ␮l, and then an aliquot of the reaction mixture was taken for quantification of ADP generated as a by-product of phosphorylation as described in Materials and Methods. b The relative activities were based on the mean value of reaction mixture I (mean value ⫾ the standard deviation in two independent experiments). ND, not detected.

that B. subtilis RhaB is an L-rhamnulose kinase that phosphorylates rhamnulose at the C-1 position at the expense of ATP to yield rhamnulose-1-phosphate. Response of the rhaEW promoter activity to various carbon sources. The effects of various carbon sources on the in vivo rhaEW promoter activity in the presence of both RhaR and CcpA were examined by lacZ reporter analysis using B. subtilis strain FU1203 carrying the rhaEW promoter (bases ⫺144 to 147)-lacZ fusion introduced into the amyE locus (Table 1). Among the four carbon sources tested (glucose, rhamnose, xylose, and malic acid), only rhamnose significantly enhanced the ␤-Gal activity under the control of the rhaEW promoter (Fig. 8). It was assumed that, only in the presence of rhamnose as a carbon source, RhaR is relieved from binding to the regulatory region of the rhaEW promoter by interacting with rhamnulose-1-phosphate derived from rhamnose and that CcpA stays dissociated from the cre sequence in the rhaEW promoter, resulting in holo RNA polymerase access to the promoter. When glucose and malic acid were, respectively, used as a carbon source, CcpA formed a complex with P-Ser-HPr (24, 42), which was capable of binding to the cre sequence in the rhaEW promoter region. On the other hand, the inhibitory effect of rhamnulose-1-phosphate on the RhaR binding to the operator site that overlaps the cre sequence was not exerted under these culture conditions. Based on the DNase I footprinting result, we speculated that, in cases where both regulatory factors are active in binding to their target sequences, RhaR largely contributes to the repression of the rhaEW promoter by binding to the regulatory region preferentially (Fig. 5). When xylose was used as a carbon source, the rhaEW promoter was likely to be repressed only by the binding of RhaR to the operator site, without formation of the CcpA/P-Ser-HPr complex. We also examined the rhaEW promoter activity of the same reporter strain grown in a medium containing both glucose and rhamnose. In this case, the ␤-Gal activity remained low level during the logarithmic growth phase, gradually increasing from the stationary phase on. This suggests that, although the repression by RhaR was released from the beginning of cultivation, repression

TABLE 4 Relative phosphorylating activities of His-RhaB from B. subtilis in the presence of various factorsa Reaction no.

His-RhaB

His-RhaA

Rhamnose

Rhamnulose

ATP

Mean relative activity (%) ⫾ SDb

I II III IV

⫹ ⫹ ⫹ –

– – ⫹ ⫹

– ⫹ ⫹ ⫹

⫹ – – –

⫹ ⫹ ⫹ ⫹

100.0 ⫾ 10.2 0.275 ⫾ 0.389 29.2 ⫾ 5.7 ND

a Each reaction was carried out at 30°C for 30 min in the presence (⫹) or absence (–) of 10 ␮g of His-RhaB, 1 ␮g of His-RhaA, 10 mM rhamnose, 10 mM rhamnulose, and 1 mM ATP in a total volume of 500 ␮l, and then an aliquot of the reaction mixture was taken for quantification of ADP generated as a by-product of phosphorylation as described in Materials and Methods. b The relative activities were based on the mean value of reaction mixture I (mean value ⫾ the standard deviation in two independent experiments). ND, not detected.

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of rhamnose, ATP, His-RhaA, and His-RhaBEc, the phosphorylation proceeded at a rate of 30% of that obtained by the reaction with a combination of rhamnulose, ATP, and His-RhaBEc (III). No phosphorylating activity was detected in a reaction mixture containing rhamnose, ATP, and His-RhaA (IV). These results provide strong evidence that the ketose derived from rhamnose by His-RhaA is rhamnulose, i.e., B. subtilis RhaA catalyzes the isomerization of rhamnose to rhamnulose. We next examined the phosphorylating activity of His-RhaB. The specific activity of His-RhaB for rhamnulose was determined to be 10.7 ⫾ 0.3 ␮mol min⫺1 mg⫺1 (mean value ⫾ the standard deviation in three independent experiments), which was 21-fold lower than that of His-RhaBEc. However, like His-RhaBEc, HisRhaB specifically phosphorylated rhamnulose, and rhamnose was not accepted as the substrate. Moreover, 29% of the phosphorylating activity was detected by the reaction with a combination of rhamnose, ATP, His-RhaA, and His-RhaB compared to that obtained by the reaction with a combination of rhamnulose, ATP, and His-RhaB (Table 4). We also tested whether a phosphorylated rhamnulose produced in the reaction mixture containing rhamnulose, ATP, and His-RhaB is able to release the DNA binding of RhaR through gel retardation analysis. After the phosphorylation reaction, the reaction mixture was concentrated, and then an aliquot was added to the binding mixture, as in the above-mentioned case in which the inhibitory effect of rhamnulose-1-phosphate produced by HisRhaBEc on the DNA binding of RhaR was tested (Fig. 7A). It was estimated that 46% of the rhamnulose in the reaction mixture was phosphorylated by His-RhaB and that 23 mM the phosphorylated rhamnulose was present in the binding mixture. We used another control mixture, which was prepared by substituting His-RhaB for His-RhaBEc in the control mixture for testing the effect of rhamnulose-1-phosphate. The result of the gel retardation analysis shown in Fig. 7B clearly indicates that the phosphorylated rhamnulose produced in the His-RhaB-catalyzed reaction specifically inhibited the DNA binding of RhaR, which is sensitive to rhamnulose-1-phosphate. Thus, this result is consistent with the idea

Hirooka et al.

(amyE::PrhaEW-lacZ) was used to monitor the ectopically introduced rhaEW promoter in the presence of both RhaR and CcpA. The B. subtilis cells were cultivated in media containing glucose (circles), rhamnose (squares), both of these sugars (diamonds), xylose (triangles), or malic acid (crosses) at 25 mM. The open and filled symbols indicate the OD600 values and ␤-Gal activities, respectively. The ␤-Gal activities of the cells grown in the presence of malic acid are indicated by asterisks. The lacZ reporter experiments were repeated at least two times, and representative results are shown.

mediated by the CcpA/P-Ser-HPr complex continued until the stationary phase, at which time glucose became depleted in the medium, leading to derepression of the promoter activity. Regulation of the rhaEW promoter by RhaR and CcpA through rhamnose and glucose catabolisms, respectively. To verify that RhaR is primarily involved in the inducing effect of rhamnose on the rhaEW promoter and that CcpA is essential for the carbon catabolite repression of this promoter, we conducted lacZ reporter analysis using B. subtilis strains carrying the lacZ reporter fused to the rhaEW promoter with and without rhaR disruption and ccpA deletion (Table 1) that had been grown on each of three sugars (glucose, rhamnose, and xylose). The integration of plasmid pMUTIN2 causes disruption of the target gene, lacZ fusion to the promoter upstream of the target gene, and placement of the genes downstream of the target gene under the control of the IPTG-inducible Pspac promoter (25). It is noteworthy that the media contained IPTG for the forced expression of downstream genes and that the strains carrying a disruption in the genes encoding the rhamnose catabolic enzymes were able to grow normally even in the medium containing rhamnose as a carbon source, because this medium contained an abundance of amino acids, which were available as an alternative energy source. Strain BFS1435 (rhaEW::pMUTIN2) was used to monitor the rhaEW promoter in the presence of both RhaR and CcpA (Fig. 9, upper left). The ␤-Gal activity of this strain significantly increased only in the presence of rhamnose as a carbon source, showing a response property similar to that of strain FU1203, except that the induced activity of strain BFS1435 was much higher than that obtained by strain FU1203 (592 ⫾ 7 and 29.7 ⫾ 1.7 nmol min⫺1 OD600⫺1 at the maxima for BFS1435 and FU1023, respectively; mean value ⫾ the standard deviation in two independent experiments). Strain YULBd (rhaR::pMUTIN2) was used to monitor the rhaEW promoter in the absence of RhaR (Fig. 9, upper right). When this strain was grown in media containing either rhamnose or xylose, comparable ␤-Gal activities were obtained (136 ⫾ 7 and 120 ⫾ 15 nmol min⫺1 OD600⫺1 at the maxima in the presence of rhamnose and xylose, respectively), whereas the ␤-Gal activity in the presence of glucose remained lower than those obtained by the

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other two sugars. A comparison of the results of strains BFS1435 and YULBd indicates that, in strain YULBd, repression of the rhaEW promoter by RhaR in the absence of rhamnose was abolished by the rhaR disruption, although the CcpA-mediated carbon catabolite repression was still effective, resulting in partial repression of the promoter in the presence of glucose (Fig. 9, upper left and upper right). Furthermore, the ␤-Gal activity of strain YULBd in the glucose-containing medium gradually increased as the cell growth progressed, probably due to relief from the carbon catabolite repression associated with glucose consumption. In the rhamnose-containing medium, the ␤-Gal activity of strain YULBd was somewhat lower than that of strain BFS1435, which might be attributed to the difference in positions of the plasmid insertion. To monitor the rhaEW promoter in the absence of CcpA, strain FU1198 (⌬ccpA::cat rhaEW::pMUTIN2) was constructed, and its ␤-Gal activity in the presence of each sugar was measured (Fig. 9, lower left). The response pattern of the rhaEW promoter of this strain was almost the same as that observed in strain BFS1435, except that, in the presence of glucose, the ␤-Gal activity of strain FU1198 was slightly higher than that of strain BFS1435 (5.16 ⫾ 0.42 and 1.67 ⫾ 1.39 nmol min⫺1 OD600⫺1 at the maxima for FU1198 and BFS1435, respectively) (Fig. 9, upper left and lower left). This result indicates that, even under the ccpA deletion, RhaR alone sufficiently repressed the rhaEW promoter in the absence of rhamnose. Strain FU1199 (⌬ccpA::cat rhaR::pMUTIN2) was constructed to monitor the rhaEW promoter in the absence of both RhaR and CcpA (Fig. 9, lower right). When this strain was cultivated in the media containing any of the three sugars, comparable ␤-Gal activities were observed (119 ⫾ 24, 139 ⫾ 15, and 118 ⫾ 22 nmol min⫺1 OD600⫺1 at the maxima in the presence of glucose, rhamnose, and xylose, respectively), showing that the rhaEW promoter of this strain was completely released from the regulation of RhaR and CcpA. Effects of disruptions of the genes encoding the rhamnose catabolic enzymes on the rhamnose-dependent induction of the rhaEW promoter. To examine whether the rhamnose-dependent induction of the rhaEW promoter mediated by RhaR is affected by disrupting any of the rhaEWRBMA operon members encoding

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FIG 8 lacZ reporter analysis to examine the effects of various carbon sources on rhaEW promoter activity in the presence of both RhaR and CcpA. Strain FU1203

Regulation of Rhamnose Catabolism in B. subtilis

the enzymes for rhamnose catabolism, lacZ reporter analysis was further performed by using three B. subtilis strains carrying the pMUTIN2 insertion in any of the members (rhaB, rhaM, and rhaA), along with the ccpA deletion. No increase of ␤-Gal activity was observed in strains FU1200 (⌬ccpA::cat rhaB::pMUTIN2) and FU1202 (⌬ccpA::cat rhaA::pMUTIN2) grown in the presence of rhamnose (Fig. 10, upper and lower). On the other hand, the ␤-Gal activity of strain FU1201 (⌬ccpA::cat rhaM::pMUTIN2) was induced by rhamnose in the medium, although the induced activity was lower than that obtained by strain FU1198 (⌬ccpA::cat rhaEW::pMUTIN2) (53.9 ⫾ 16.2 nmol min⫺1 OD600⫺1 at the maxima for FU1201 grown in the presence of rhamnose), likely due to the positional effects of the plasmid insertion (Fig. 10, middle). These results, together with the result of strain FU1198 (Fig. 9, lower left), demonstrate that RhaA and RhaB are indispensable for the rhamnose-dependent induction of the rhaEW promoter; this is consistent with the functional roles of RhaA and RhaB presumed from the in vitro analyses, i.e., they supply rhamnulose-1phosphate as the effector molecule for RhaR through a sequence of isomerization and phosphorylation of rhamnose. DISCUSSION

In this study, we revealed that the B. subtilis rhaEWRBMA operon, composed of the genes encoding the rhamnose catabolic enzymes (RhaA, RhaB, RhaEW, and RhaM) and the DeoR-type transcriptional repressor (RhaR), is under the rhamnose-responsive control by RhaR and the carbon catabolite repression by CcpA. The RhaR binding region, which was determined by the DNase I footprinting analysis, contains two imperfect direct repeats

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(UDR and DDR) (Fig. 5). The gel retardation analysis using the deletion probes revealed that the UDR was essential for the highaffinity binding of RhaR, whereas the deletion of the DDR did not affect the DNA binding affinity of RhaR but altered the mobility of the RhaR-DNA complex (Fig. 6). Our ortholog clustering search (43; http://mbgd.genome.ad.jp/), as well as those conducted by Rodionov’s group (7, 44), found a gene cluster with the organization similar to the B. subtilis rhaEWRBMA operon in a subgroup of the genus Bacillus that includes B. atrophaeus, B. licheniformis, and B. halodurans, in addition to Terribacillus aidingensis, Thermobacillus composti, and some species of the genus Paenibacillus, including P. mucilaginosus, P. polymyxa, and P. terrae, although there are some differences in the order of the member genes and some clusters lack the rhaM ortholog. A comparison of the regulatory region of the B. subtilis operon with the regions upstream of the rhaEW orthologs of three Bacillus species (B. atrophaeus, B. licheniformis, and B. halodurans) revealed that a 9-bp conserved motif (CAAAAAWAA) is redundantly arranged in these regions and that the UDR of B. subtilis are composed of a quadruplet of the motifs (Fig. 11). Thus, we speculate that the RhaR homolog binds to the region, including this motif, to repress the putative operon for rhamnose catabolism in these Bacillus species. Although the DDR found in B. subtilis is not apparently conserved in the other species, we found the single 9-bp conserved motif in the DDR. According to the results of the DNase I footprinting and gel retardation analyses (Fig. 5 and 6), it is likely that the DDR interacts with the RhaR protein and contributes to the conformational arrangement of the RhaR-DNA complex, which possibly prevents the CcpA/P-Ser-HPr complex

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FIG 9 lacZ reporter analysis to monitor the rhaEW promoter in the presence and absence of RhaR and CcpA in B. subtilis cells grown in media containing glucose, rhamnose, or xylose. Strains BFS1435 (rhaEW::pMUTIN2), YULBd (rhaR::pMUTIN2), FU1198 (⌬ccpA::cat rhaEW::pMUTIN2), and FU1199 (⌬ccpA:: cat rhaR::pMUTIN2) were used to monitor the rhaEW promoter activity in the presence of both RhaR and CcpA (upper left), in the absence of RhaR (upper right), in the absence of CcpA (lower left), and in the absence of both RhaR and CcpA (lower right), respectively. Cells of each strain were cultivated in media containing any of glucose (circles), rhamnose (squares), and xylose (triangles) at 25 mM. The open and filled symbols indicate the OD600 values and ␤-Gal activities, respectively. Each medium contained 1 mM IPTG for the forced expression of genes located downstream of the pMUTIN2-inserted position. The lacZ reporter experiments were repeated at least two times, and representative results are shown.

Hirooka et al.

encoding rhamnose catabolic enzymes on the rhamnose-dependent induction of the rhaEW promoter. Strains FU1200 (⌬ccpA::cat rhaB::pMUTIN2), FU1201 (⌬ccpA::cat rhaM::pMUTIN2), and FU1202 (⌬ccpA::cat rhaA::pMUTIN2) were used to monitor the rhaEW promoter activity in the absence of any of RhaB (top), RhaM (center), and RhaA (bottom) under the ⌬ccpA background. Cells of each strain were cultivated in media containing either glucose (circles) or rhamnose (squares) at 25 mM. The open and filled symbols indicate the OD600 values and ␤-Gal activities, respectively. Each medium contained 1 mM IPTG for the forced expression of genes located downstream of the pMUTIN2-inserted position. The lacZ reporter experiments were repeated at least two times, and representative results are shown.

from binding to the cre included in the DDR. The 9-bp conserved motif in the DDR might be important for the interaction with RhaR. Based on the results of the reporter analysis and the DNase I footprinting (Fig. 5 and 9), we consider that the expression of the rhaEWRBMA operon is primarily and strictly regulated by RhaR in a rhamnose-responsive manner. In addition to this, if energetically favorable carbon sources such as glucose are available even in the presence of rhamnose, the CcpA/P-Ser-HPr complex presumably functions to repress the operon for prioritizing the use of more favorable carbon sources (Fig. 8). Rodionov’s group proposed a 19-bp direct repeat with the 9-1-9 arrangement (CAAAA AWAA-A-CAAANATRA; R stands for A or G) as a putative binding site of the RhaR homolog (7, 44). We also regard this sequence as a core region for binding of B. subtilis RhaR and its homolog.

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FIG 10 lacZ reporter analysis to examine the effects of disrupting the genes

Nevertheless, we deduce that the 9-bp motifs found in the positions other than this 19-bp direct repeat also interact with the RhaR homolog in the Bacillus species, as well as that in the DDR in B. subtilis. Many members of the DeoR family from various bacterial species have been characterized and were found to share several common features (45). They contain a highly conserved region near the N terminus that includes a helix-turn-helix DNA-binding motif, whereas their C-terminal region is assumed to be responsible for oligomerization and effector binding. Most of the members reported thus far act as repressors involved in sugar and nucleoside metabolism, and their effector molecules are usually phosphorylated intermediates in the metabolic pathways that they control. For instance, E. coli DeoR, E. coli GlpR, E. coli UlaR, B. subtilis IolR, and Lactococcus lactis FruR control the metabolic systems for deoxyribonucleoside, glycerol-3-phosphate, L-ascorbate, myo-inositol, and fructose, and their effector molecules are deoxyribose-5-phosphate, glycerol-3-phosphate, L-ascorbate-6phospate, 2-deoxy-5-keto-D-gluconate-6-phosphate, and fructose-1-phosphate, respectively (14–18). These DeoR members possess a conserved region near their C terminus, which is structurally related to E. coli D-ribose-5-phosphate isomerase, implying that this C-terminal region functions as the effector sensor (46). B. subtilis RhaR also possesses these conserved N-terminal and Cterminal regions, probably functioning as the DNA-binding domain and the domain for dimerization and effector binding, respectively. Our in vitro and in vivo experiments clearly demonstrated that RhaR serves as a repressor for the rhaERBMA operon and that its effector molecule is rhamnulose-1-phosphate, a phosphorylated intermediate in the B. subtilis rhamnose catabolic pathway (Fig. 7A, 9, and 10). These features are typical of DeoR family members but completely different from those of E. coli RhaS and RhaR. In E. coli, both RhaS and RhaR belong to a major subset of the AraC/XylS family, which share the C-terminal region for DNA binding and transcriptional activation and the N-terminal region for dimerization and/or effector binding. Rhamnose serves as an effector molecule for both RhaS and RhaR to cause their allosteric conformational changes suitable for DNA binding and transcriptional activation of the respective target genes (47). In B. subtilis and E. coli, rhamnose catabolic genes are also under carbon catabolite repression, which is exerted by modes of action different from each other. In B. subtilis, the CcpA/P-SerHPr complex acts as a repressor to bind to the cre sequence in the regulatory region of the rhaEWRBMA operon in the presence of preferred carbon sources. In contrast, in the case of E. coli, the CRP protein, complexed with cAMP, binds to each of the upstream regions of the rhaS and rhaB promoters, acting as a coactivator in the absence of the preferred carbon source. Therefore, the rhamnose catabolic gene clusters of B. subtilis and E. coli are totally different in the transcriptional regulatory mechanism involving the regulator types, the signaling molecules, and the action modes of the regulators, as well as in their gene organization, although the catabolic pathway and the catalyzing enzymes encoded in the targeted genes are quite similar, except for B. subtilis RhaEW and E. coli RhaD. Previous studies suggested that, in some Gram-negative bacteria, genes involved in rhamnose catabolism are regulated by a DeoR family member. In R. leguminosarum, rhamnose catabolism has an effect on its nodulation competitiveness, and the genes for rhamnose catabolism constitute two operons, i.e., the

Regulation of Rhamnose Catabolism in B. subtilis

rhaDI and rhaRSTPQUK operons, which are oriented divergently so that their promoters are proximally situated (48). The member genes are predicted to encode a set of rhamnose catabolic enzymes (RhaU, RhaK, RhaD, and RhaI), an ABC-type rhamnose transport system (RhaS, RhaT, RhaP, and RhaQ), and a DeoR-type transcriptional regulator (RhaR), among which the functions of RhaK and RhaSTPQ have been characterized (48–50). The participation of a DeoR family member in the gene regulation for rhamnose catabolism was also suggested in Chloroflexus aurantiacus (7) and Thermotoga maritima (51). However, the molecular mechanism underlying each regulatory system remains unclear. The information of the B. subtilis rhaEWRBMA operon should provide a useful clue for understanding these regulatory mechanisms, as well as those mediated by a DeoR-type regulator and/or the CcpA homolog distributed among various bacterial species. In E. coli, the genes for rhamnose catabolism constitute the rhaT gene, the rhaSR operon, and the rhaBAD (possibly rhaBADM) operon, which are contiguously arranged in this order; however, the direction of the rhaSR operon is opposite to those of the others so that the rhaS and rhaB promoters are close to each other (9, 19). This gene organization is widely conserved in the family Enterobacteriaceae, including the genera Enterobacter, Klebsiella, Pantoea, Salmonella, Shigella, Yersinia, and Escherichia, albeit to slight differences in the constituent members in some species (7, 43). Moreover, in the genomes of the genera Dickeya and Pectobacterium, rhaSR and rhaBADM clusters are present with the same arrangement as that of E. coli; however, the rhaT ortholog is replaced by a putative operon encoding an ABC transport system. Some Gram-negative bacterial species, such as Dickeya dadantii (alias Erwinia chrysanthemi), Pectobacterium carotovorum subsp. carotovorum, and Pantoea ananatis, are known as plant pathogens that possess the rhamnose catabolic genes likely regulated by the E. coli RhaS, RhaR, and CRP homologs (52), whereas B. subtilis and related species, such as B. licheniformis and P. polymyxa, are Gram-positive plant growth-

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promoting rhizobacteria, and their rhamnose catabolic genes are likely to be under the control of the B. subtilis-type regulatory system involving RhaR and CcpA. Since rhamnose catabolic enzymes are assumed to play a significant role in the invasion of such pathogens into plant tissues (52), an antagonist specific to E. coli-type RhaS and/or RhaR is expected to be an effective pesticide without preventing growth of the Gram-positive plant growth-promoting rhizobacteria. Moreover, in E. coli, the regulatory system of the rhaB promoter comprising RhaS, RhaR, and CRP is applied to the heterologous expression system, in which the expression of the target gene situated downstream of the rhaB promoter is capable of being strictly controlled by the addition of rhamnose or glucose (53). Similarly to this, it is possible that, in the cells of B. subtilis and related species, the expression of the introduced gene under the control of the rhaEW promoter can be finely regulated by RhaR and CcpA in the presence of rhamnose or glucose. We have confirmed that B. subtilis strain 168 can assimilate rhamnose by the observation that it grows in an S6 medium (54) plus 25 mM rhamnose and 0.02% yeast extract, while no growth was observed in the S6 medium supplemented only with 0.02% yeast extract (data not shown). This also supports that rhamnose is incorporated into the B. subtilis cells. A homology search showed that B. subtilis GlcU protein, which is predicted to be a glucose transporter, is modestly homologous to E. coli RhaT (18% identity and 62% similarity). GlcU might function to take up rhamnose in B. subtilis. Alternatively, it is possible that rhamnose is incorporated through the ABC transporter encoded in the gene clusters involved in the degradation of rhamnogalacturonan type I (2, 6). The functional characterization of these transporters would be required for the elucidation of rhamnose uptake in B. subtilis. ACKNOWLEDGMENTS We are grateful to Takayoshi Edahiro, Kosuke Kimura, and Masahiro Toratani for their help with the experiments.

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FIG 11 Comparison of the regulatory region of the B. subtilis rhaEWRBMA operon with the corresponding regions in the other Bacillus species. Each 9-bp conserved motif is indicated by a blue arrow. The result of a comparison of the 20 conserved motifs is shown as motif logos created with the B. subtilis Motif Location Search software (http://dbtbs.hgc.jp/motiflocationsearch.html).

Hirooka et al.

K.H. and Y.F. designed the study and wrote the manuscript. K.H. and T.S. constructed the plasmids and the B. subtilis strains. K.H. and Y.K. prepared the RhaR, His-RhaA, His-RhaB, and His-RhaBEc proteins. T.S. prepared the CcpA, HPr, and His-HPrK proteins and performed the phosphorylation of HPr and the purification of P-Ser-HPr. K.H. and Y.K. conducted the analyses of transcripts, the DNA binding experiments, the reporter assay, and the measurement of enzyme activities. All authors reviewed the results and approved the final version of the manuscript.

FUNDING INFORMATION

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This research received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.

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Regulation of the rhaEWRBMA Operon Involved in l-Rhamnose Catabolism through Two Transcriptional Factors, RhaR and CcpA, in Bacillus subtilis.

The Bacillus subtilis rhaEWRBMA (formerly yuxG-yulBCDE) operon consists of four genes encoding enzymes for l-rhamnose catabolism and the rhaR gene enc...
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