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J Bacteriol. Dec 2005; 187(24): 8411–8426.
PMCID: PMC1317013

The ClgR Protein Regulates Transcription of the clpP Operon in Bifidobacterium breve UCC 2003


Five clp genes (clpC, clpB, clpP1, clpP2, and clpX), representing chaperone- and protease-encoding genes, were previously identified in Bifidobacterium breve UCC 2003. In the present study, we characterize the B. breve UCC 2003 clpP locus, which consists of two paralogous genes, designated clpP1 and clpP2, whose deduced protein products display significant similarity to characterized ClpP peptidases. Transcriptional analyses showed that the clpP1 and clpP2 genes are transcribed in response to moderate heat shock as a bicistronic unit with a single promoter. The role of a clgR homologue, known to control the regulation of clpP gene expression in Streptomyces lividans and Corynebacterium glutamicum, was investigated by gel mobility shift assays and DNase I footprint experiments. We show that ClgR, which in its purified form appears to exist as a dimer, requires a proteinaceous cofactor to assist in specific binding to a 30-bp region of the clpP promoter region. In pull-down experiments, a 56-kDa protein copurified with ClgR, providing evidence that the two proteins also interact in vivo and that the copurified protein represents the cofactor required for ClgR activity. The prediction of the ClgR three-dimensional structure provides further insights into the binding mode of this protein to the clpP1 promoter region and highlights the key amino acid residues believed to be involved in the protein-DNA interaction.

Bifidobacteria aresome of the most common inhabitants of mammalian and animal gastrointestinal tracts (37). In the human gastrointestinal tract, their presence has been associated with beneficial health effects, such as the prevention of diarrhea, amelioration of lactose intolerance, and immunomodulation (21, 28, 37, 45). The preparation of Bifidobacterium containing products may require that the microbes survive industrial food manufacturing processes, such as freeze-drying, freezing, and spray drying, while remaining viable during storage. Such probiotic products reinforce the need for robust bifidobacteria to survive passage through the upper parts of the digestive tract, compete with the resident intestinal flora, preferably colonize the digestive tract, and express specific functions, probably under suboptimal growth conditions. Despite the commercial, and consequently, scientific interest, the genetics of bifidobacteria has not developed to a degree that can be compared to that of other high-G+C gram-positive bacteria (45), and therefore molecular tools that, for example, allow gene inactivation or in vivo transcription analysis are not yet feasible for bifidobacteria.

In order to resist stressful environmental challenges, cells synthesize protective proteins, including both chaperones and substrate-specific proteases, that primarily act to prevent the accumulation of misfolded proteins by performing various roles, such as protein folding, stabilization, renaturation, and resolubilization (22, 35, 39). Some of the genetic elements encoding these chaperones have been identified in bifidobacteria, including the groEL-groES (43) and dnaK (47) genes.

Recent studies of bacteria have focused on the expression of the Clp protein family, members of which are well conserved in both eukaryotic and prokaryotic organisms (30, 52). Many Clp proteins contain ATPase activity, and the number of ATP nucleotide-binding domains in such proteins has been used for classification purposes. It is widely accepted that Clp ATPases can function both as molecular chaperones and as regulator components of the proteolytic complex (52). In Escherichia coli, the Clp complex consists of two functionally distinct subunits: the larger ClpA protein functions as the ATP-binding regulatory subunit (25), conferring substrate specificity, whereas the smaller ClpP protein provides the proteolytic activity (25). On its own, ClpP possesses only peptidase activity, and it requires an association with ClpA to degrade polypeptides of longer than six amino acids. The Clp protease complex consists of two central heptameric rings of ClpP flanked by two hexameric rings of ClpA. In Firmicutes, with the exception of Bacillus thuringiensis (8), only a single chromosomal copy of the clpP gene has been found, whereas in Actinobacteridae up to five clpP paralogous genes have been identified (49, 51). So far, it is not known if the presence of multiple copies of the clpP gene is correlated with enhanced protection against certain stressful conditions. In Streptomyces lividans, the five identified clpP-like genes are organized into two operons, one that includes the clpP1 and clpP2 genes and one that encompasses the clpP3 and clpP4 genes, and a third monocistronic transcription unit harbors clpP5 (51).

In eubacteria, expression of the genes belonging to the clp family is subject to multiple modes of regulation. In Escherichia coli, the clpP, clpB, and clpX genes are controlled by the general heat shock sigma factor σ32 (11, 18). In contrast, in several gram-positive bacteria, including Bacillus subtilis, Listeria monocytogenes, Streptococcus salivarius, Enterococcus faecalis, Lactobacillus sakei, Lactococcus lactis, Oenococcus oeni, and Clostridium acetobutylicum, transcription of the single clpP gene is directed by the vegetative sigma factor σA and controlled by the CtsR repressor, which binds a heptanucleotide repeat that overlaps the −10 and −35 hexamers (6).

In high-G+C gram-positive bacteria like S. lividans, expression of the clpP1 clpP2 operon, as well as that of the clpC gene, is regulated by a transcriptional activator, ClgR (for clp gene regulator), which binds an imperfect consensus motif (CGCT-4N-GCGNAC) (2, 7). The expression of the second clpP operon (clpP3 clpP4) in Streptomyces lividans is regulated by a second transcriptional activator, designated PopR (49).

In this report, the clpP operon of Bifidobacterium breve UCC 2003 is described. The transcriptional induction of this operon upon exposure to stressful conditions was investigated, while the role of a ClgR homologue and an as-yet-unidentified cofactor protein in the regulation of the clpP operon was explored, revealing evidence for a novel heat shock-controlled regulatory mechanism in the genus Bifidobacterium.


Bacterial strains and culture conditions.

All Bifidobacterium strains used for this study are described in Table Table1.1. They were grown anaerobically in MRS (Difco, Detroit, MI) supplemented with 0.05% l-cysteine-HCl and were incubated at 37°C for 16 h. Escherichia coli was grown aerobically on a rotary shaker (150 rpm) at 37°C in LB medium or plated onto LB agar (Difco, Detroit, MI) plates when appropriate. Antibiotics were used at the following concentrations: ampicillin, 100 μg/ml; kanamycin, 25 μg/ml; and chloramphenicol, 2 μg/ml.

Strains used for this study, with clpP1 and clpP2 sequence accession numbers

DNA amplification.

Genomic DNAs used as templates for PCRs were extracted following a protocol described in a previous study (46).

PCR was used to amplify a DNA fragment corresponding to a 1,000-bp internal fragment of the clpP operon from all investigated Bifidobacterium strains, using the oligonucleotides clpP1-UNIV and clpP1-REV (Table (Table2).2). PCRs were carried out according to the standard procedure described by Sambrook et al. (32). The resulting amplicons were separated in a 1.5% agarose gel, followed by ethidium bromide staining. PCR fragments were purified using a PCR purification spin kit (QIAGEN, West Sussex, United Kingdom) and were subsequently sequenced.

Oligonucleotides used for this study

Phylogenetic analysis.

Phylogeny calculations, including distance calculations and the generation of phylogenetic trees, were performed using the PHYLIP package (9) and the ClustalX program. The numbers of synonymous substitutions between all possible pairs of the clpP1 and clpP2 genes were determined by applying the method of Nei and Gojobori (27), using the MEGA computer program (19). Correction for multiple substitutions was done according to the Jukes-Cantor formula (17).

Plasmids and plasmid constructions.

The E. coli pQE-30 vector (QIAGEN) was used for overproduction and purification of an N-terminally six-histidine-tagged bifidobacterial ClgR protein (h-ClgR). The clgR gene from B. breve UCC 2003 was amplified using the primers 903-uni and 903-rev, which contain a BamHI and a HindIII restriction site, respectively. The resultant 566-bp PCR fragment was digested with BamHI and HindIII and ligated into similarly restricted pQE30, using the T4 DNA ligase enzyme (Roche, Sussex, United Kingdom), to generate plasmid pQE-ClgR, which was introduced into E. coli M15 (QIAGEN, United Kingdom) as described by Sambrook et al. (32).

Plasmid pNZ272 (29), which contains a promoterless gusA gene system, was used as a reporter system. Various portions of the clpP promoter region were generated by PCR, using one fixed primer for the 3′ end (clp-Rev [complementary to sequences 9 bp upstream of the start codon of the B. breve UCC 2003 clpP1 gene]) and various primers for the 5′ end of this promoter region (Table (Table2).2). The resultant PCR amplicons were digested with BglII and PstI and ligated into similarly restricted pNZ272, which was used to transform E. coli M15 (QIAGEN, United Kingdom). From these transformant plasmids, pclp1, pclp6, pclp7, pclp8, and pclp3 were isolated, which carry the whole clpP promoter region (pclp1) or decreasing portions of it (pclp6, pclp7, pclp8, and pclp3). All of the above-mentioned plasmids were then introduced into B. breve UCC 2003 by electrotransformation. DNA sequences of all genetic constructs were confirmed by DNA sequencing (MWG Biotech, Ebersberg, Germany) and restriction analysis.

GUS assay.

B. breve UCC 2003 cultures with an inoculum level of 2% containing pNZ272 or a derivative thereof were grown exponentially at 37°C until an optical density at 600 nm (OD600) of 0.3 was reached, after which the temperature was shifted to 43°C or 50°C or NaCl was added to a final concentration of 0.7 M, and the cultures were incubated for 180 min. One milliliter of cells was then centrifuged at 7,000 rpm for 5 min, and β-glucuronidase (GUS) activity was determined as described by Platteeuw et al. (29).

Overproduction of h-ClgR in E. coli and protein purification.

A 300-ml culture of an E. coli M15 strain containing the pQE-ClgR plasmid was grown to an OD600 of 0.6 prior to induction by the addition of 1 mM IPTG (isopropyl-β-d-thiogalactopyranoside; Fluka, Germany). Three hours following induction, cells were harvested by centrifugation at 10,000 rpm for 10 min. Cell pellets were resuspended in lysis buffer (50 mM NaH2PO4, 10 mM Tris-HCl, 30 mM imidazole, pH 8.0) as recommended by the supplier (QIAGEN) and allowed to lyse by being shaken gently at 27°C for 2 h. Cell debris was eliminated from the lysate by centrifugation at 13,000 rpm for 10 min. The resulting supernatant was passed through a column containing 4 ml of Ni-nitrilotriacetic acid (Ni-NTA) agarose (QIAGEN), which had been preequilibrated with 10 ml of lysis buffer. The column was washed two times with 10 ml of wash buffer (50 mM NaH2PO4, 1 M NaCl, 30 mM imidazole, 0.5% [vol/vol] Triton X-100, 5 mM β-mercaptoethanol, pH 8.0) and then eluted using 10 ml of elution buffer (50 mM NaH2PO4, 1 M NaCl, 250 mM imidazole, pH 8.0). Protein concentrations were determined using a Bio-Rad protein assay in conjunction with a bovine serum albumin standard curve. The expected size and purity of the eluted h-ClgR protein were verified by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE).

h-ClgR cross-linking.

Cross-linking experiments were performed according to a previously published procedure (1). All samples were fractionated in loading buffer by SDS-PAGE.

RNA isolation and Northern blot analysis.

B. breve UCC 2003 cells were grown to an OD600 of 0.6. Temperature stress was applied by transferring the culture to either 20°C, 37°C, 43°C, 47°C, or 50°C, while osmotic stress was applied by the addition of 5 M NaCl-containing prewarmed medium to give a final concentration of either 0.5 M or 0.7 M. At various time points, 30-ml aliquots of culture were collected and briefly centrifuged to harvest cells. Total RNA was isolated using the macaloid acid method and then treated with DNase (Roche, United Kingdom). Briefly, cell pellets were resuspended in 0.5 ml of phenol, pH 7.5, and placed in a tube containing 0.18 g of macaloid acid (Sigma) and 0.8 g of glass beads (diameter, 106 μm; Sigma). The cells were lysed by shaking the mix at the maximum setting on a BioSpec homogenizer at 4°C for 2 min. The mixture was then centrifuged at 12,000 rpm for 15 min, and the upper phase containing the RNA sample was recovered. The RNA sample was further purified by phenol and ethanol precipitated according to the method described by Sambrook et al. (32). Slot blot hybridizations were carried out following a previously described protocol (48). RNA electrophoresis and Northern blot hybridization were carried out as described previously (42, 44). All slot blot and Northern hybridization experiments were performed at least twice.

Primer extension analysis.

The 5′ end of the clpP1 RNA transcript was determined using a protocol described in a previous study (44). The synthetic oligonucleotide used was named clpP1-prom (Table (Table22).

Gel mobility shift DNA binding assays.

A 377-bp DNA fragment corresponding to the clpP1 promoter region (from position −249 to position +128 with respect to the putative transcription start site) was amplified by PCR with primers P1-uni and P1-rev. The resultant amplicon was purified using a G50-Spin column (Amersham, Little Chalfont, United Kingdom) and then labeled using [γ-32P]dATP and T4 polynucleotide kinase (New England Biolabs, MA). The level of radioactive labeling was measured using a Beckman LS multipurpose scintillation counter (Fullerton, CA).

Binding reactions were performed according to a previously described protocol (48). Bands were visualized by autoradiography at −70°C, using Kodak Biomax MR film (Eastman-Kodak).

All gel retardation assays were performed at least twice.

Protease treatment of crude cell extract from B. breve UCC 2003.

Ten micrograms of crude cell extract from B. breve UCC 2003 was incubated with 20 U of pronase enzyme (Roche, United Kingdom) for 4 h at 37°C, as recommended by the supplier. The pronase was subsequently heat inactivated and/or chemically inactivated by incubating the mixture with 3 μl of 1× protease inhibitor cocktail (Roche, United Kingdom), as recommended by the supplier. The control sample was treated identically, except that no pronase was added.

Protein pull-down procedure.

The h-ClgR or h-HspR (48) protein was attached to a column containing 4 ml of Ni-NTA agarose (QIAGEN) which had been preequilibrated with 10 ml of lysis buffer. A crude extract from a UCC 2003 culture grown at 43°C or 50°C was then passed through the h-ClgR- or h-HspR-containing column. The column was treated as described above, except that the wash buffer did not contain Triton X-100 or β-mercaptoethanol and the elution buffer contained 500 mM imidazole. The expected size and purity of the coeluted protein were verified by SDS-PAGE.

DNase I footprint assays.

A 377-bp DNA fragment covering the clpP1 promoter region (from position −249 to position +128 with respect to the putative transcription start site) was obtained by PCRs using two primer combinations, in which one primer was an IRD800-labeled oligonucleotide (MWG Biotech, Germany) and the second primer was unlabeled in order to effect strand-specific labeling. The two primer combinations used for this purpose were P1-TOP IRD-800/P1-REV and P1-BOT/P1-uni (IRD800) (Table (Table22).

Binding reactions were performed as described above in the section outlining gel mobility shift DNA binding assays, using 0.15 pmol of labeled fragments, 25 to 100 pmol of h-ClgR, and 1 μg of crude lysate from B. breve UCC 2003 cultures grown at 43°C. The resulting reaction mixtures were then incubated for 30 min at 37°C, followed by the addition of 5 μl of 0.25-μg/ml DNase I in DNase I buffer (10 mM Tris-HCl, pH 8.0, 5 mM MgCl2, 5 mM CaCl2, 50 mM KCl, and 1 mM dithiothreitol), after which digestion was allowed to proceed for exactly 5 min at 37°C and stopped by the addition of 0.5 μl of 0.5 M EDTA, pH 8.0. After the addition of 500 μl of 95% ethanol, the DNA was precipitated overnight at −20°C and recovered by centrifugation (10,000 rpm for 15 min). The resulting DNA pellet was washed with 500 μl of 70% ethanol, air dried, and dissolved in 2.5 μl of water and 2.5 μl of loading solution (LicoR, Cambridge, United Kingdom). The reactions were separated as described for primer extension analysis (7).

ClgR 3D prediction.

To derive structure templates for the B. breve UCC 2003 ClgR protein, fold recognition techniques were carried out via the protein structure prediction Meta server (http://bioinfo.pl/Meta/). Several structure templates were used, including the tertiary structure of the DNA binding domain of SinR from Bacillus subtilis (20) (PDB entry 1b0n; chain A) and that of the CI repressor from bacteriophage λ (PDB entry 1rio; chain A). An alignment between the C-terminal portion (residues 94 to 170) of ClgR, 1b0n, and 1rio was generated by using the Fold and Function Assignment System (FFAS) (31), a profile-profile alignment algorithm. Finally, a three-dimensional (3D) model of this C-terminal domain of ClgR was obtained via the Swiss Model server (34).

Nucleotide sequence accession numbers.

The GenBank accession numbers for the partial clpP1 and clpP2 gene sequences generated in this study are reported in Table Table1.1. Nucleotide sequence data regarding the clpP operon of B. breve UCC 2003 have been deposited in GenBank under accession number AY955251. The nucleotide sequence of the B. breve UCC 2003 clgR gene has been deposited in GenBank under accession number AY837843.


Identification of the clpP operon in B. breve UCC 2003.

An analysis of the B. breve UCC 2003 genome sequence (S. Leahy, M. O'Connell-Motherway, J. A. Moreno-Muńoz, G. F. Fitzgerald, D. G. Higgins, and D. van Sinderen, unpublished data) revealed the presence of adjacent genes, designated clpP1 and clpP2, whose protein products displayed 53% and 48% identity to ClpP1 and ClpP2 of Streptomyces coelicolor A3, respectively. The residues Ser, His, and Asp, which constitute the catalytic triad of the serine protease ClpP in E. coli (25), are conserved in the ClpP1 sequences of B. breve UCC 2003 (positions 102, 121, and 172, respectively).

The structural organization and locations of the clpP1 and clpP2 genes in the chromosomes of B. breve UCC 2003 and other bacteria are schematically displayed in Fig. Fig.1,1, where the deduced amino acid sequences of the B. breve UCC 2003 clpP operon are aligned with those of high-G+C gram-positive bacteria and low-G+C gram-positive bacteria. This comparative analysis showed that the most similar proteins to the predicted B. breve ClpP1/ClpP2 proteases were the assumed ClpP1/ClpP2 proteins from Bifidobacterium longum (33). However, identity levels of ≥46% were still observed between the B. breve ClpP1 protein and the ClpP proteins of less-related bacterial taxa, such as Bacillus clausii. In contrast, the flanking DNA regions of the clpP1 and clpP2 genes were shown to be highly variable with respect to gene synteny, except between the two bifidobacterial species.

FIG. 1.
Comparison of the clpP operon in B. breve UCC 2003 and corresponding loci in various other bacteria. Each arrow indicates an open reading frame. The lengths of the arrows are proportional to the lengths of the predicted open reading frames. Orthologs ...

The clpP1 and clpP2 genes in bifidobacteria are preceded by the eriC gene, which putatively encodes a chloride channel protein, and are followed by clpX, a gene predicted to encode a ClpX protease (Fig. (Fig.11).

Phylogenetic analysis of the clpP operon in bifidobacteria.

The clpP1 and clpP2 DNA sequences from B. breve and B. longum were aligned and compared. Two identical regions corresponding to the 5′ and 3′ ends of the clpP1 and clpP2 genes were identified, and a pair of primers (clpP1-UNIV and clpP1-REV) was designed. These primers allowed the amplification of a 1,000-bp region encompassing part of the clpP1 and clpP2 sequences of nine Bifidobacterium species. Alignments of clpP1 and clpP2 DNA sequences were used to generate a phylogenetic tree by the neighbor-joining method (Fig. (Fig.2).2). These data were supported by the indicated bootstrap values (9). For completeness, we included in the analysis homologous DNA sequences from other strains belonging to different genera representing gram-positive bacteria with high and low G+C contents. This tree showed a clear separation into two major clusters representing the Firmicutes and the Actinobacteridae taxa. Furthermore, the two Actinobacteridae clpP paralogs separated into two distinct phylogenetic groups (Fig. (Fig.22).

FIG. 2.
Phylogenetic tree obtained using the clpP1 and clpP2 gene sequences. Bar, phylogenetic distances. Bootstrap values are reported for a total of 1,000 replicates. The clpP1 and clpP2 gene sequences are indicated. Bacteria belonging to the Firmicutes and ...

A comparison of the phylogenetic trees based on the clpP1 and clpP2 sequences shows very similar phylogenetic topologies in the short term of evolution, whereas some discrepancies in the branching order were depicted in bifidobacterial genus evolution (e.g., for the B. breve-Bifidobacterium suis-Bifidobacterium infantis subcluster and B. longum).

A phylogenetic tree which was constructed on the basis of the 16S rRNA gene sequences available in public databases was mostly similar to the clpP1- and clpP2-based phylogenies (data not shown). Moreover, the correlations (r2) between the pairwise distances for the 16S rRNAs and the synonymous distances for the clpP1 and clpP2 sequences were 0.795 and 0.823, respectively. Therefore, it can be concluded that the base substitutions occurring in the clpP1 and clpP2 sequences during the evolutionary process render these genes reliable molecular evolutionary clocks. Interestingly, closely related strains exhibit nearly identical 16S rRNA sequences, e.g., Bifidobacterium animalis subsp. animalis and B. animalis subsp. lactis occupy separate branches in the clpP1 and clpP2 sequence-based tree (Fig. (Fig.22).

Heat induction of the clpP operon in B. breve UCC 2003.

Expression of the clpP locus in high-G+C gram-positive bacteria such as Corynebacterium glutamicum (7) and Mycobacterium tuberculosis (36) is induced by a number of protein-denaturing stress treatments, such as heat and osmotic stress. To determine if the induction of the clpP operon occurs upon exposure to stressful conditions in B. breve UCC 2003, slot blot hybridization was used to analyze total RNAs isolated from B. breve cultures following exposure for up to 150 min to temperatures ranging from 20°C to 50°C and to NaCl concentrations of 0.5 M and 0.7 M (Fig. (Fig.3a3a).

FIG. 3.
Transcriptional analysis of the B. breve UCC 2003 clpP operon. (a) Slot blot hybridization using RNAs extracted from cells incubated for up to 150 min at various temperatures or with various NaCl concentrations (indicated in the left-hand margin). (b) ...

Based on the strength of the hybridization signal, the highest expression levels of the clpP1 gene occurred at 43°C following a 150-min exposure, whereas exposure to higher temperatures, high NaCl concentrations, or a low temperature (20°C) did not appear to significantly increase the level of clpP1 transcription (Fig. (Fig.3a).3a). Densitometric analysis of Northern slot blots revealed that the levels of clpP1 mRNA were increased 15-fold in cells that were subjected to heat shock at 43°C for 150 min compared to those in unstressed cells (Fig. (Fig.3b3b).

Characterization of clpP1 and clpP2 gene transcription activity by Northern blotting.

Northern hybridization experiments were performed in order to determine whether the clpP1 and clpP2 genes, and perhaps the downstream gene, are cotranscribed. Total mRNA was isolated from B. breve UCC 2003 grown at 37°C, following heat shock at 43°C or 50°C, or upon osmotic shock with 0.7 M NaCl. Transcription of the clpP1 gene was investigated by Northern blotting using an internal clpP1 probe. A 1.4-kb transcript was detected in RNAs extracted from 37°C and 43°C samples (Fig. 3c and d). The shift to heat shock conditions (43°C) strongly increased the strength of expression of the 1.4-kb transcript (Fig. (Fig.3d).3d). Also, when a probe spanning the flanking clpP2 gene was used, a signal of 1.4 kb was detected (data not shown). The transcriptional kinetics of the clpP2 gene were found to be identical to those of the clpP1 gene. Both genes increased their transcription level upon temperature shift and reached their maximum transcriptional level at 150 min. This result clearly demonstrated that the clpP1 and clpP2 genes form a bicistronic transcriptional unit. When Northern hybridization was performed with a probe corresponding to the clpX gene, no transcripts were detected (data not shown), showing that this gene is not part of the clpP operon and that its transcription is not induced by heat or osmotic shock. The latter finding is in contrast to the situation in E. coli, where clpP and clpX are part of a single transcriptional unit (11). In B. breve, two stem-loop structures (ΔG = −20.3 kcal and −12.9 kcal), which represent possible rho-independent transcription terminators, are present downstream of clpP2 and upstream of clpP1, respectively (Fig. (Fig.3c3c).

Identification of the clpP1 transcriptional start site.

To determine the transcriptional start point of the clpP1 gene, primer extension analyses were performed using RNAs extracted from B. breve cells which had been subjected to heat shock (see Fig. S1 in the supplemental material). An extension product was identified 30 nucleotides 5′ of the predicted translational start site of the clpP1 gene (see Fig. S1a and b in the supplemental material). The transcription start site was in the same position at 43°C and at 47°C (see Fig. S1a in the supplemental material). The analysis of the putative promoter region of clpP1 revealed a potential promoter-like sequence weakly resembling the previously found −10 and −35 bifidobacterial hexamers (42, 43, 47, 48). The predicted translational start site is preceded by a typical ribosome-binding-site sequence (AAGGAG) located eight nucleotides upstream of the putative translational start site.

The sequences of the region upstream of the clpP1 genes of both B. breve UCC 2003 and B. longum NCC 2705 were aligned in an attempt to identify putative regulatory elements. For completeness, we also identified by PCR analysis the putative promoter regions of the clpP1 genes from the closely related B. suis and B. infantis taxa and from two more distantly related Bifidobacterium species (B. dentium and B. globosum). As shown in Fig. S1c in the supplemental material, a large consensus promoter sequence can be deduced from the six sequences, which includes the putative −10 hexamers, −35 hexamers, the ribosome-binding-site region, and the transcriptional start site, which were conserved in all bifidobacterial sequences examined. Moreover, a number of other DNA motifs were shown to be conserved in all of these strains, including a 4-bp partially inverted repeat (IR; CGCT-4N-GCCNA) which is almost identical to a clpP operator site for the ClgR protein (CGCT-4N-GCGNAC) found in other members of the Actinobacteridae group (2, 7).

Regulation of clpP operon: h-ClgR binds to the clpP1 promoter region.

In several other members of the Actinobacteridae group, the ClgR protein was shown to bind the clpP1 promoter region, indicating that it acts as a transcriptional regulator of the clpP operon (2, 7).

An analysis of the B. breve UCC 2003 genome sequence showed a gene homologous to clgR from C. glutamicum and S. lividans (7), which in a similar manner, could be responsible for the regulation of some clp genes. We identified that in B. breve UCC 2003 and B. longum NCC 2705, the clgR gene and clpP operon are located in different chromosomal regions (data not shown). The clgR gene is located downstream of a predicted diacylglycerol-glycerol-3-phosphate-3-phosphatidyltransferase-encoding gene (pgsA3) and upstream of thepresumed recA gene (Fig. (Fig.44).

FIG. 4.
Organization of the clgR locus in different members of the Actinobacteridae. Each arrow indicates an open reading frame. The lengths of the arrows are proportional to the lengths of the predicted open reading frames. Orthologs are marked with the same ...

In order to determine if ClgR of B. breve UCC 2003 binds to the promoter region of the clpP1 gene, the UCC 2003 ClgR protein was overproduced in E. coli as a His-tagged version, purified, and designated h-ClgR (Fig. (Fig.5).5). The h-ClgR protein was then used in gel mobility shift DNA binding assays with 377-bp radiolabeled DNA fragments corresponding to the promoter region of the clpP1 gene (clpP1p). These experiments showed that 100 pmol of purified h-ClgR protein did not affect the mobility of the clpP1p fragment (Fig. (Fig.5b).5b). In contrast, when the binding assay was performed using 50 or 100 pmol of purified h-ClgR protein combined with 1 μg of crude cell extract from B. breve UCC 2003 cells, designated CX, that had been subjected to heat stress (43°C for 150 min), the mobility of the clpP1p fragment was clearly reduced (Fig. (Fig.5b).5b). Furthermore, no difference in migration was observed when the clpP1p fragment was incubated with 1 μg of CX without the purified h-ClgR protein (Fig. (Fig.5b5b).

FIG. 5.
Detection of ClgR binding to the clpP1 promoter of B. breve UCC 2003. (a) Overproduction and purification of ClgR. SDS-PAGE analysis was performed with the purified ClgR protein (lanes 1 to 3, containing 15, 20, and 8 μg of protein, respectively) ...

No binding activity was observed when 50 pmol or 100 pmol of h-ClgR protein was used in the presence of 1 μg of a crude extract of B. breve UCC 2003 cultures exposed to 50°C (Fig. (Fig.5b)5b) or grown in the presence of 0.7 M NaCl (Fig. (Fig.5b).5b). This suggests that the CX extract obtained from cells exposed to 43°C contains one or more cofactors required for ClgR binding activity.

In order to characterize the nature of this cofactor(s), protease treatment (with pronase) of the CX followed by pronase inactivation by thermal and/or chemical means was performed, and the reactions were subsequently analyzed in gel mobility shift assays (Fig. (Fig.5b).5b). Interestingly, when the equivalent amount of 1 μg of pronase-treated CX was incubated with 100 pmol of h-ClgR, no displacement of clpP1p was observed (Fig. (Fig.5b).5b). However, a clear retardation of the clpP1p fragment was observed in a control experiment employing the same amount of protein which had been treated in an identical manner but without the addition of pronase (Fig. (Fig.5b5b).

ClgR is a dimer.

To determine whether the purified ClgR protein exists in a multimeric form, we carried out in vitro cross-linking assays in the absence and presence of glutaraldehyde. Using SDS-PAGE, it was shown that in the absence of glutaraldehyde, the h-ClgR protein migrated as a single band at a position which corresponds to the molecular mass of the monomeric form. However, in the presence of glutaraldehyde, a fraction of the h-ClgR protein was shifted, with a molecular weight corresponding to a dimer (Fig. (Fig.6).6). The appearance of a cross-linked species with the apparent mobility of a dimer occurred as soon as 10 min after the addition of glutaraldehyde. After 80 min of glutaraldehyde treatment, the cross-linking process appeared to be completed.

FIG. 6.
Subunit composition of ClgR. An SDS-PAGE gel of glutaraldehyde-cross-linked and non-cross-linked h-ClgR is shown. The absence (−) or presence (+) of the glutaraldehyde cross-linking reagent and the time of the cross-linking reaction are ...

Purification of putative cofactor protein.

In order to determine whether the cofactor protein(s) required for ClgR binding to the clpP1 promoter region could form stable complexes with ClgR, a Ni-NTA column was first loaded with h-ClgR, after which the column was flooded with a crude cell extract from B. breve UCC 2003 cells that had been subjected to heat stress (43°C for 150 min) or with a crude cell extract from B. breve UCC 2003 cultures exposed to 50°C. The column was then washed, h-ClgR was finally eluted from the column, and the eluted fractions containing h-ClgR were analyzed by SDS-PAGE for coeluted proteins (Fig. (Fig.7a).7a). In the fraction where h-ClgR had been immersed with the UCC 2003 extract obtained from cells exposed to 43°C, h-ClgR was shown to coelute with a protein of ≈56 kDa (Fig. (Fig.7a).7a). This protein was absent in equivalent fractions from control experiments where ClgR was coeluted with UCC 2003 extract obtained from cells exposed to 50°C and/or in equivalent fractions where ClgR was coeluted with CX extract obtained from cells grown in the presence of 0.7 M NaCl (Fig. (Fig.7a).7a). Moreover, in a control experiment using another bifidobacterial transcription regulator, h-HspR (48) was subjected to the same experimental procedure and subsequently analyzed by SDS-PAGE. Only one band, of 21 kDa, which corresponds to the h-HspR protein, was detected (Fig. (Fig.7a7a).

FIG. 7.
Pull-down assay to identify the cofactor of ClgR (a) and DNA binding assay using activated and nonactivated h-ClgR molecules (b). (a) Lanes 1 and 2, SDS-PAGE of h-ClgR coeluted with CX obtained from a culture grown at 43°C; lane 3, SDS-PAGE of ...

Since we had shown that a proteinaceous cofactor contained in the UCC 2003 cell extract obtained from cultures grown at 43°C promotes the binding of h-ClgR to the clpP promoter region, we wanted to determine whether the coeluted 56-kDa protein and ClgR were able to interact with the clpP1 promoter region (clpP1p) in gel mobility shift assays. These experiments showed that h-ClgR plus the 56-kDa coeluate was capable of a complete mobility shift of the clpP1p fragment (in either the presence or absence of ATP), while no displacement of clpP1p was observed when a control h-ClgR coeluate, obtained with a UCC 2003 crude extract of cells exposed to 50°C or grown in the presence of 0.7 M NaCl, was used (Fig. (Fig.7b).7b). This result strongly suggests that the 56-kDa coeluted protein, which is uniquely present in UCC 2003 cell extracts exposed to 43°C, is in fact the cofactor required for ClgR binding activity.

Operator site of ClgR.

To precisely delineate the sequences that constitute the ClgR binding site, DNase I footprint assays were performed on the 377-bp DNA fragment that encompasses the clpP1 promoter region. As shown in Fig. 7a and b, using end-labeled template or nontemplate strands, the purified h-ClgR protein, in the presence of 1 μg of crude extract from B. breve UCC 2003 cultures exposed to 43°C, protects a region extending from positions −79 and −99 on the template (Fig. (Fig.8a)8a) and nontemplate (Fig. (Fig.8b)8b) strands, respectively. The protected region contains the partially inverted repeat (CGCT-4N-GCCNA) (Fig. (Fig.7c),7c), which resembles the ClgR operator site (CGCT-4N-GCGNAC) reported for the clpP1 promoter region of other members of the Actinobacteridae group (2, 7). The palindromic structure of the putative operator site of clpP1 in B. breve UCC 2003 is consistent with ClgR being present as a dimer in solution. Moreover, DNase I hypersensitivity sites, which are sites that become more susceptible to DNase I cleavage upon protein binding, were detected in the protected DNA region, which suggests that a distortion of the normal DNA structure had occurred as a result of ClgR binding.

FIG. 8.
DNase I footprints of the 347-bp fragment from the B. breve UCC 2003 clpP1 promoter region for the template strand (a) and the nontemplate strand (b). (c and d) β-Glucuronidase activities of various clpP1-gusA fusions in B. breve UCC 2003 grown ...

The involvement of the ClgR protected region, including the IR element, in gene expression of the B. breve UCC 2003 clpP operon was assessed by means of a number of transcriptional fusions between various clpP promoter fragments and a promoterless gusA reporter gene in the promoter probe vector pNZ272 (Fig. (Fig.8d).8d). The GUS activity from cells harboring the more complete clpP1 promoter region (pclp1) and grown at 43°C represented the same expression pattern as that determined by slot blot hybridization experiments. This heat shock regulation of β-glucuronidase is partially or completely lost in clones containing plasmid pclp6 or pclp7 or plasmid pclp8 or pclp3, respectively, where the DNA encompassing the binding site of ClgR, as determined by footprinting experiments, is partially (pclp6, pclp7, and pclp8) or completely (pclp3) removed (Fig. 8c and d). These data show that ClgR, possibly in association with a cofactor, can bind to the IR in a sequence-specific manner, thereby positively regulating the expression of the clpP operon at 43°C.

Structural investigation of ClgR of B. breve UCC 2003.

Using fold recognition prediction, it was possible to identify structural homologues of the ClgR protein that were limited to the C-terminal region only (from residues 94 to 170 of B. breve UCC 2003 ClgR). In contrast, the N-terminal part of ClgR did not display any significant matches. The most significant structural homologue for the C-terminal part of UCC 2003 ClgR is the SinR protein of B. subtilis (PDB entry 1b0nA) (20). This protein shares the highest sequence identity (31% identical residues for 65 matched positions) as well as the most significant sequence fold compatible score (FFAS score, −36.1). Therefore, the 1b0nA protein was employed as a structural template for 3D modeling. Moreover, it should be emphasized that the sequence alignment between ClgR and SinR generated from FFAS was highly consistent with those based on other methods, which implies that the alignment was highly reliable (data not shown).

The predicted C-terminal region of B. breve UCC 2003 ClgR contains five helices. Interestingly, helix 2 and helix 3 (residues 113 to 134) form an archetypal helix-turn-helix motif (Fig. (Fig.9a),9a), which is a recurrent substructure in different DNA binding proteins (16).

FIG. 9.
Predicted 3D model for the C-terminal end of ClgR. (a) In order to predict the mode of binding between ClgR and the DNA molecule, the DNA molecule (the ligand) from the crystal structure of 1rio was superimposed onto the predicted ClgR model and is depicted ...

The amphipathic region (residues 140 to 164), which includes helixes 4 and 5, may be involved in dimer formation of ClgR molecules as a consequence of its coiled-coil structure, as demonstrated for the homologous region in the SinR protein (10). By superimposing the ClgR C-terminal model with another structural homologue (i.e., the CI repressor protein of bacteriophage λ; PDB entry 1rio, chain A) (16), for which the structure of the bound DNA molecule is available, we identified eight amino acid residues (Arg-107, Leu-114, Arg-115, Ser-124, Leu-125, Arg-133, Lys-136, and Ser-139) which may be involved in ClgR-DNA binding (Fig. (Fig.9a).9a). Notably, of these eight amino acid residues, Ser-124, Leu-125, and Arg-133 are located in the recognition helix, implying a higher probability of their making base-specific interactions in the major groove. Furthermore, similar to what has been described for the 3D template molecule used (16), other amino acid residues (Arg-107,Leu-114, Arg-115, Lys-136, and Ser-139) could also be involved in DNA interactions by nonspecific contacts (e.g., contacts with the DNA backbone). The high conservation of this set of amino acid residues between various ClgR homologues (Fig. (Fig.9b)9b) may support this hypothesis.


This report describes the characterization of a bifidobacterial clpP operon which is activated upon heat stress treatments in bifidobacteria and gives evidence for the first time of the role of the transcriptional activator ClgR and a proteinaceous cofactor molecule in the regulation of such an operon in this group of bacteria. The clpP operon of B. breve UCC 2003 was identified on the basis of high levels of homology to previously described clpP operons (25, 49-51). Our results clearly indicate that the B. breve clpP operon consists of two genes, clpP1 and clpP2, in an organization which is identical to that of presumed clpP regions of other sequenced Bifidobacterium species and to other available sequences of members of the Actinobacteridae group, such as S. coelicolor A3 (4).

Phylogenetic analysis of the clpP1 and clpP2 paralogs revealed that the observed genetic constellation may have been derived from a duplication of the clpP gene during the evolutionary process from Firmicutes, representing a “true” paralogy; alternatively, they may have arisen as a consequence of a hidden paralogy, implying that the ancestor of the Firmicutes and Actinobacteria taxa once had multiple copies of the genes and that a differential loss of gene copies occurred, resulting in the current clpP gene distribution.

The clpP1 and clpP2 genes constitute ideal target candidates for diagnostic purposes because they are highly conserved and ubiquitous in bacteria (12-14, 23, 39). Thus, these genes may be used as alternative molecular markers to the rRNA sequences, which could be added to the current databases of alternative molecular markers such as the tuf, recA, groEL, atpD, grpE, and dnaK genes (24, 40-44, 47) and might corroborate and help to complete the evolutionary history of various bifidobacterial species (38, 41, 42).

Although molecular chaperone-encoding genes such as clpB, clpC, clpP, dnaK, groEL, and groES have been demonstrated to be induced by heat stress in bifidobacteria (39, 43, 47, 48; this study), nothing is known about a global regulation of this stress response in these bacteria.

The expression of the B. breve UCC 2003 clpP operon is induced at its highest level upon moderate heat shock regimens (ΔT of 6 K, where ΔT means the temperature difference between 37°C and the applied stress temperature) but not upon severe heat stress (ΔT of 10 to 13 K) or osmotic stress.

Notably, similar results have been found for another member of the B. breve UCC 2003 clp gene family, i.e., the clpC gene. The similarities between the transcription patterns of the clpP1/clpP2 and clpC genes are striking, not only with respect to stimulus induction but also with respect to the tuning systems which govern their expression, thus suggesting that in bifidobacteria the clpP operon and the clpC gene belong to the same regulon.

The clpP1 promoter region was only bound by the purified ClgR protein in the presence of a crude lysate of heat-stressed B. breve UCC 2003 cells, and this binding activity did not occur upon protease treatment of the crude lysate of heat-stressed B. breve UCC 2003 cells. In pull-down assays using whole-cell extracts from heat-stressed B. breve UCC 2003 cultures, a protein of 56 kDa was shown to copurify with ClgR. Moreover, we showed that the h-ClgR-56-kDa protein coeluate mixture was able to bind to the clpP1 promoter region without the assistance of any other cofactor, which would generate an h-ClgR-activated molecule. Taken together, these results represent a novel type of positive cofactor-mediated regulation of gene expression in high-G+C gram-positive bacteria. Preliminary results from mass spectrometric analysis identified the 56-kDa protein to be the B. breve UCC 2003 GroEL chaperone, and future experiments involving the overproduction and purification of B. breve UCC 2003 GroEL will be carried out in order to better elucidate the ClgR-GroEL interactions and to investigate if other molecules may be involved in this process.

We speculate that this cofactor is involved in assisting the proper folding of ClgR. To our knowledge, two other heat-induced transcriptional regulators which require the presence of a molecular chaperone as a cofactor to function properly have been previously described for B. subtilis. (26) and S. coelicolor (3). In B. subtilis, the GroEL chaperone machine modulates the activity of the transcriptional repressor HrcA (26), whereas in S. coelicolor the DnaK protein coregulates the activity of the heat-induced transcriptional repressor HspR (3). Thus, the ClgR-mediated regulation in bifidobacteria represents a third and novel cofactor-mediated activation of transcriptional acts involved in the heat response.

In order to provide evidence that, similar to other members of the Actinobacteridae group (2, 7), the bifidobacterial ClgR homologue is a transcriptional activator of the clpP operon, a reporter system which includes different portions of the clpP1 promoter region was used. Only clones harboring plasmids encompassing the putative ClgR operator site were shown to induce β-glucuronidase activity when subjected to moderate heat shock regimens. On the other hand, the levels of β-glucuronidase activity were drastically reduced in B. breve strains harboring a promoter probe vector in which the ClgR binding site was removed, thus suggesting that ClgR acts as a transcriptional activator of the clpP operon in B. breve UCC 2003 and other bifidobacteria.

The clpP1 promoter regions protected in DNase I footprint experiments contain a palindromic motif (CGCT-4N-GCCNA) which may represent the operator site for the ClgR protein in B. breve UCC 2003. This motif is highly similar to the consensus operator site (CGCT-4N-GCGNAC) of the ClgR regulons in Streptomyces, Mycobacterium, and Corynebacterium (2, 7). For all of these organisms, it has been shown that ClgR regulates many stress-induced genes, such as clpP1, clpP2, clpC, and lon (2). However, we have shown that in contrast to those of other high-G+C gram-positive bacteria, the bifidobacterial ClgR homologue requires a proteinaceous cofactor in order to acquire binding activity.

According to DNase I footprint analysis, the ClgR binding site in the clpP1 promoter region of B. breve UCC 2003 is located upstream of the transcription start site. Thus, we can hypothesize that ClgR activates expression by altering the promoter conformation. However, we cannot exclude that ClgR may alter the interaction with the RNA polymerase. Moreover, binding of ClgR may induce DNA bending, since several enhanced DNase I cleavage sites appear within the protected region of the clpP1 gene.

By analogy to other DNA binding proteins (10, 16, 53), the structure of B. breve UCC 2003 ClgR suggests that it may be active as a dimer or even as a multimer. Chemical cross-linking with glutaraldehyde (5) indicated that B. breve UCC 2003 ClgR is present as a dimer in solution and may thus bind to the clpP1 operator as a dimer. Although in vitro evidence of higher-order protein complexes was not obtained, it remains a possibility that the protein may bind DNA to form a higher-order multimeric complex.

There are very strong similarities between ClgR of B. breve UCC 2003 and its homologues in Streptomyces (2) and Corynebacterium (7), with 46% identity in the central region containing the predicted helix-turn-helix motif. Moreover, the prediction of the ClgR fold structure highlights the importance of eight amino acid residues, five of which are located within the helix-turn-helix motif, which supports its functional relationship with other transcriptional regulators (15, 53), although this requires experimental verification by nuclear magnetic resonance and/or crystallography structural studies. Prediction of a 3D model for B. breve UCC 2003 ClgR was possible only for the C-terminal part of the protein; the N-terminal domain did not reveal any homologous structures. Interestingly, the N-terminal domain of the bifidobacterial ClgR protein appears to be 72 to 92 amino acid residues longer than those of homologous proteins in other members of the Actinobacteridae group. Taken together, this finding along with the fact that the ClgR homologues in other members of the Actinobacteridae group do not require any cofactor molecules to bind to the clpP1 promoter region (2, 7) might suggest that this N-terminal extension found only in bifidobacterial ClgR interacts with the cofactor molecule. Therefore, this would represent a novel mechanism which is apparently unique to bifidobacteria of ClgR-mediated heat shock transcriptional regulation. One of the main tasks to be addressed in future studies will be gaining an understanding of ClgR regulation and activation in bifidobacteria. In fact, in other high-G+C gram-positive bacteria, ClgR was found to be regulated by its degradation by as yet unknown stimuli using the ClpP proteolytic complex (7). In S. lividans, the degradation of ClgR as well as another transcriptional regulator, PopR, by the ClpP proteolytic complex requires the C-terminal Ala-Ala residues (49). Since the B. breve UCC 2003 ClgR protein possesses C-terminal Ala-Ala residues, we speculate that bifidobacterial ClgR might also be degraded by ClpP. Thus, we postulate that UCC 2003 ClgR activity itself is controlled via regulated proteolysis by Clp protease.

The topic of stress responses in bifidobacteria is highly relevant to the food industry. Crucial aspects related to industrial applications, such as the preparation of cells using freeze-drying technologies and cell survival in products which present a hostile environment for bifidobacteria, make it essential to increase our knowledge of the molecular mechanisms and molecular actors (e.g., ClpP) involved in the heat stress response. More investigations on such molecular mechanisms are required in order to better understand the molecular basis of heat protection for bifidobacterial cells during food manufacturing and to select new heat stress-tolerant bifidobacterial strains.

Supplementary Material

[Supplemental material]


This work was financially supported by the Higher Education Authority Programme for Research in Third-Level Institutions, by the Science Foundation Ireland Alimentary Pharmabiotic Centre located at University College Cork, and by a Marie Curie Development host fellowship (HPMD-2000-00027) to M.V.

We thank the members of the B. longum DJO10A genome sequencing project funded by the U.S. Department of Energy Joint Genome Institute for making available the sequence of the clpP locus. Finally, we thank Valentina Bernini for helpful and constructive discussions.


Supplemental material for this article may be found at http://jb.asm.org/.


1. Ashok, K. D., C. S. Baker, K. Suzuki, A. D. Jones, P. Pandit, T. Romero, and P. Babitzke. 2003. CsrA regulates translation of the Escherichia coli carbon starvation gene cstA by blocking ribosome access to the cstA transcript. J. Bacteriol. 15:4450-4460. [PMC free article] [PubMed]
2. Bellier, A., and P. Mazodier. 2004. ClgR, a novel regulator of clp and lon expression in Streptomyces. J. Bacteriol. 186:3238-3248. [PMC free article] [PubMed]
3. Bucca, G., A. M. E. Brassington, H. J. Schonfeld, and C. P. Smith. 2000. The HspR regulon of Streptomyces coelicolor: a role for the DnaK chaperone as a transcriptional co-repressor. Mol. Microbiol. 38:1093-1103. [PubMed]
4. Crecy-Lagard, V., P. Servant-Moisson, J. Viala, C. Grandvalet, and P. Mazodier. 1999. Alteration of the synthesis of the Clp ATP-dependent protease affects morphological and physiological differentiation in Streptomyces. Mol. Microbiol. 32:505-517. [PubMed]
5. Derre, I., G. Rapoport, and T. Msadek. 1999. CtsR, a novel regulator of stress and heat shock response, controls clp and molecular chaperone gene expression in gram-positive bacteria. Mol. Microbiol. 31:117-131. [PubMed]
6. Derre, I., G. Rapoport, and T. Msadek. 2000. The CtsR regulator of stress response is active as a dimer and specifically degraded in vivo at 37°C. Mol. Microbiol. 38:335-347. [PubMed]
7. Engels, S., J. E. Schweitzer, C. Ludwig, M. Bott, and S. Schaffer. 2004. clpC and clpP1P2 gene expression in Corynebacterium glutamicum is controlled by regulatory network involving the transcriptional regulators ClgR and HspR as well as the ECF sigma factor σH. Mol. Microbiol. 52:285-302. [PubMed]
8. Fedhila, S., T. Msadek, P. Nel, and D. Lereclus. 2002. Distinct clpP genes control specific adaptive responses in Bacillus thuringiensis. J. Bacteriol. 184:5554-5562. [PMC free article] [PubMed]
9. Felsenstein, J. 1993. PHYLIP (Phylogeny Inference Package), version 3.5c. Distributed by the author. University of Washington, Seattle, Wash.
10. Gaur, N. K., J. Oppenheim, and I. Smith. 1991. The Bacillus subtilis sin gene, a regulator of alternative developmental processes, codes for a DNA-binding protein. J. Bacteriol. 173:678-686. [PMC free article] [PubMed]
11. Gottesman, S. 1996. Proteases and their targets in Escherichia coli. Annu. Rev. Genet. 30:465-506. [PubMed]
12. Gupta, R. S. 1995. Evolution of the chaperonin families (Hsp60, Hsp10 and Tcp-1) of proteins and origin of eukaryotic cells. Mol. Microbiol. 15:1-11. [PubMed]
13. Gupta, R. S. 2001. The branching order and phylogenetic placement of species from completed bacterial genomes, based on conserved indels found in various proteins. Int. Microbiol. 4:187-202. [PubMed]
14. Gupta, R. S. 2002. Phylogeny of bacteria: are we now close to understanding it? ASM News 68:284-291.
15. Harrison, S. C., and A. K. Aggarwal. 1990. DNA recognition by proteins with helix-turn-helix motif. Annu. Rev. Biochem. 59:933-969. [PubMed]
16. Jain, D., B. E. Nickels, L. Sun, A. Hochschild, and S. A. Darst. 2004. Structures of a ternary transcription activation complex. Mol. Cell 13:45-53. [PubMed]
17. Jukes, T. H., and C. R. Cantor. 1969. Evolution of protein molecules, p. 21-132. In H. N. Munro (ed.), Mammalian protein metabolism. Academic Press, New York, N.Y.
18. Kroh, H. E., and L. D. Simon. 1990. The ClpP component of the Clp protease is the sigma 32-dependent heat shock protein. J. Bacteriol. 172:6026-6034. [PMC free article] [PubMed]
19. Kumar, S., K. Tamura, and M. Nei. 1993. MEGA: molecular evolutionary genetics analysis, version 1.01. Pennsylvania State University, College Park, Pa.
20. Lewis, R. J., J. A. Brannigan, W. A. Offen, I. Smith, and A. J. Wilkinson. 1998. An evolutionary link between sporulation and prophage induction in the structure of a repressor:anti-repressor complex. J. Mol. Biol. 283:907-912. [PubMed]
21. Lievin, V., I. Pfeiffer, S. Hudault, F. Rochat, D. Brassart, J. R. Neeser, and A. L. Servin. 2000. Bifidobacterium strains from resident infant human gastrointestinal microflora exert antimicrobial activity. Gut 47:646-652. [PMC free article] [PubMed]
22. Lindquist, S., and E. A. Craig. 1988. The heat-shock proteins. Annu. Rev. Genet. 22:631-677. [PubMed]
23. Ludwig, W., and K. H. Schleifer. 1999. Phylogeny of bacteria beyond the 16S rRNA standard. ASM News 65:752-757.
24. Masco, L., M. Ventura, R. Zink, G. Huys, and J. Swings. 2004. Polyphasic taxonomic analysis of Bifidobacterium animalis and Bifidobacterium lactis reveals relatedness at subspecies level: reclassification of Bifidobacterium animalis as Bifidobacterium animalis subsp. animalis comb. nov. and Bifidobacterium lactis as Bifidobacterium animalis subsp. lactis comb. nov. Int. J. Syst. Evol. Microbiol. 54:1137-1143. [PubMed]
25. Maurizi, M. R., W. P. Clark, S. H. Kim, and S. Gottesman. 1990. ClpP represents a unique family of serine proteases. J. Biol. Chem. 265:12546-12552. [PubMed]
26. Mogk, A., G. Homuth, C. Scholz, L. Kim, F. X. Schmid, and W. Schumann. 1997. The GroE chaperonin machine is a major modulator of the CIRCE heat shock regulon of Bacillus subtilis. EMBO J. 16:4579-4590. [PMC free article] [PubMed]
27. Nei, M., and T. Gojobori. 1986. Simple methods for estimating the numbers of synonymous and nonsynonymous nucleotide substitutions. Mol. Biol. Evol. 3:418-426. [PubMed]
28. Ouwehand, A. C., S. Salminen, and E. Isolauri. 2002. Probiotics: an overview of beneficial effects. Antonie Leeuwenhoek 82:279-289. [PubMed]
29. Platteeuw, C., G. Simons, and W. de Vos. 1994. Use of the Escherichia coli β-glucuronidase (gusA) gene as a reporter gene for analyzing promoters in lactic acid bacteria. Appl. Environ. Microbiol. 60:587-593. [PMC free article] [PubMed]
30. Porankiewicz, J., J. Wang, and A. K. Clarke. 1999. New insights into the ATP-dependent Clp protease: Escherichia coli and beyond. Mol. Microbiol. 21:449-458. [PubMed]
31. Rychlewski, L., L. Jaroszewski, W. Li, and A. Godzik. 2000. Comparison of sequence profiles. Strategies for structural predictions using sequence information. Protein Sci. 9:232-241. [PMC free article] [PubMed]
32. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
33. Schell, M. A., M. Karmirantzou, B. Snel, D. Vilanova, B. Berger, G. Pessi, M. C. Zwahlen, F. Desiere, P. Bork, M. Delley, R. D. Pridmore, and F. Arigoni. 2002. The genome sequence of Bifidobacterium longum reflects its adaptation to the human gastrointestinal tract. Proc. Natl. Acad. Sci. USA 99:14422-14427. [PMC free article] [PubMed]
34. Schwede, T., J. Kopp, N. Guex, and M. C. Peitsch. 2003. SWISS-MODEL: an automated protein homology-modeling server. Nucleic Acids Res. 31:3381-3385. [PMC free article] [PubMed]
35. Squires, C., and C. L. Squires. 1992. The Clp proteins: proteolysis regulators or molecular chaperones? J. Bacteriol. 174:1081-1085. [PMC free article] [PubMed]
36. Stewart, G. R., L. Wernisch, R. Stabler, J. A. Mangan, J. Hinds, and K. G. Laing. 2002. Dissection of the heat-shock response in Mycobacterium tuberculosis using mutants and microarrays. Microbiology 148:3129-3138. [PubMed]
37. Tannock, G. W. 1994. The acquisition of the normal microflora of the gastrointestinal tract, p. 1-16. In S. A. Gibson (ed.), Human health: the contribution of microorganisms. Springer, London, United Kingdom.
38. Vandamme, P., B. Pot, M. Gillis, P. de Vos, K. Kersters, and J. Swings. 1996. Polyphasic taxonomy, a consensus approach to bacterial systematics. Microbiol. Rev. 60:407-438. [PMC free article] [PubMed]
39. Van de Guchte, M., P. Serror, C. Chervaux, T. Smokvina, S. D. Ehrlich, and E. Manguin. 2002. Stress responses in lactic acid bacteria. Antonie Leeuwenhoek 82:187-216. [PubMed]
40. Ventura, M., and R. Zink. 2002. Rapid identification, differentiation, and proposed new taxonomic classification of Bifidobacterium lactis. Appl. Environ. Microbiol. 68:6429-6434. [PMC free article] [PubMed]
41. Ventura, M., and R. Zink. 2003. Comparative sequence analysis of the tuf and recA genes and restriction fragment length polymorphism of the internal transcribed spacer region sequences supply additional tools for discriminating Bifidobacterium lactis from Bifidobacterium animalis. Appl. Environ. Microbiol. 69:7517-7522. [PMC free article] [PubMed]
42. Ventura, M., C. Canchaya, D. van Sinderen, G. F. Fitzgerald, and R. Zink. 2004. Bifidobacterium lactis DSM 10140: identification of the atp (atpBEFHAGDC) operon and analysis of its genetic structure, characteristics, and phylogeny. Appl. Environ. Microbiol. 70:3110-3121. [PMC free article] [PubMed]
43. Ventura, M., C. Canchaya, R. Zink, G. F. Fitzgerald, and D. van Sinderen. 2004. Characterization of the groEL and groES loci in Bifidobacterium breve UCC 2003: genetic, transcriptional and phylogenetic analysis. Appl. Environ. Microbiol. 70:6197-6209. [PMC free article] [PubMed]
44. Ventura, M., C. Canchaya, V. Meylan, T. R. Klaenhammer, and R. Zink. 2003. Analysis, characterization, and loci of the tuf genes in Lactobacillus and Bifidobacterium and their direct application for species identification. Appl. Environ. Microbiol. 69:6908-6922. [PMC free article] [PubMed]
45. Ventura, M., D. van Sinderen, G. F. Fitzgerald, and R. Zink. 2004. Insights into the taxonomy, genetics and physiology of bifidobacteria. Antonie Leeuwenhoek 86:205-223. [PubMed]
46. Ventura, M., R. Reniero, and R. Zink. 2001. Specific identification and targeted characterization of Bifidobacterium lactis from different environmental isolates by a combined multiplex-PCR approach. Appl. Environ. Microbiol. 67:2760-2765. [PMC free article] [PubMed]
47. Ventura, M., R. Zink, G. F. Fitzgerald, and D. van Sinderen. 2005. Gene structure and transcriptional organization of the dnaK operon of Bifidobacterium breve UCC 2003 and its application in bifidobacterial tracing. Appl. Environ. Microbiol. 71:487-500. [PMC free article] [PubMed]
48. Ventura, M., J. G. Kenny, Z. Zhang, G. F. Fitzgerald, and D. van Sinderen. 2005. The clpB gene of Bifidobacterium breve UCC 2003: transcriptional analysis and first insights into stress induction. Microbiology 151:2861-2872. [PubMed]
49. Viala, J., and P. Mazodier. 2002. ClpP-dependent degradation of PopR allows tightly regulated expression of the clpP3clpP4 operon in Streptomyces lividans. Mol. Microbiol. 44:633-643. [PubMed]
50. Viala, J., and P. Mazodier. 2003. The ATPase ClpX is conditionally involved in the morphological differentiation of Streptomyces lividans. Mol. Genet. Genomics 268:563-569. [PubMed]
51. Viala, J., G. Rapoport, and P. Mazodier. 2000. The ClpP multigenic family in Streptomyces lividans: conditional expression of the clpP3 clpP4 operon is controlled by PopR, a novel transcriptional activator. Mol. Microbiol. 38:602-612. [PubMed]
52. Wawrzynow, A., B. Banecki, and M. Zylicz. 1996. The Clp ATPases define a novel class of molecular chaperones. Mol. Microbiol. 21:895-899. [PubMed]
53. Wintjens, R., and M. Rooman. 1996. Structural classification of HTX DNA binding domains and protein-DNA interaction modes. J. Mol. Biol. 262:294-313. [PubMed]

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