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The heat shock response in bacterial cells is characterized by rapid induction of heat shock protein expression, followed by an adaptation period during which heat shock protein synthesis decreases to a new steady-state level. In this study we found that after a shift to a high temperature the Clp ATPase (ClpE) in Lactococcus lactis is required for such a decrease in expression of a gene negatively regulated by the heat shock regulator (CtsR). Northern blot analysis showed that while a shift to a high temperature in wild-type cells resulted in a temporal increase followed by a decrease in expression of clpP encoding the proteolytic component of the Clp protease complex, this decrease was delayed in the absence of ClpE. Site-directed mutagenesis of the zinc-binding motif conserved in ClpE ATPases interfered with the ability to repress CtsR-dependent expression. Quantification of ClpE by Western blot analysis revealed that at a high temperature ClpE is subjected to ClpP-dependent processing and that disruption of the zinc finger domain renders ClpE more susceptible. Interestingly, this domain resembles the N-terminal region of McsA, which was recently reported to interact with the CtsR homologue in Bacillus subtilis. Thus, our data point to a regulatory role of ClpE in turning off clpP gene expression following temporal heat shock induction, and we propose that this effect is mediated through CtsR.

 
ClpC and ClpE belong to the highly conserved HSP100/Clp family of ATPases that are widely distributed in prokaryotic and eukaryotic cells. These proteins have been implicated in a variety of biological processes either as parts of proteolytic complexes that also include the ClpP protease or as molecular chaperones (reviewed in reference 41). Members of the ClpC subfamily are found in gram-positive bacteria and plants, and they have been shown to be important for controlling growth at high temperatures, sporulation, competence, and virulence (31, 34, 35, 38, 39, 47). The ClpE subfamily is characterized by the presence of an amino-terminal zinc-binding motif, and so far alleles have been identified only in gram-positive bacteria (11, 19, 32). The typical feature of the ClpE (11, 19, 32) and ClpX (50) subfamily proteins is an N-terminal zinc-binding domain, a so-called zinc finger, whose presence in certain proteins was first noted by Miller and coworkers (28). While the function of this domain in ClpE is unknown, such motifs are often involved in DNA binding and protein-protein interactions (5, 24, 25, 43). Inactivation of clpE alleles has generally had minor phenotypic effects (11, 19), although a Listeria monocytogenes clpE mutant had a higher growth rate at elevated temperatures and showed attenuated virulence (32).

Expression of the clp genes is regulated by the negative regulator CtsR. Homologues of CtsR have been identified in a number of gram-positive bacteria, and CtsR has been shown to bind to well-conserved DNA-binding sites present in the promoter regions of target genes (12, 22, 33). In the absence of stress expression of the CtsR regulon is repressed by CtsR binding; however, when cells are stressed, CtsR binding is released and expression is temporally induced. In the continued presence of stress the activity of CtsR is restored, and genes belonging to the CtsR regulon are re-repressed. This pattern of temporal derepression followed by repression has been observed in other stress regulatory systems. In Bacillus subtilis the heat shock regulator, HrcA, requires the GroE chaperonin for DNA binding (29). When stress is encountered, GroE is titrated by the accumulation of misfolded proteins, and HrcA is unable to bind DNA. As the concentration of chaperones is increased as part of the heat shock response, free GroE becomes available to promote binding of HrcA to DNA (30). In B. subtilis and L. monocytogenes it has been observed that expression of the CtsR regulon is derepressed in the absence of clpC (11, 12, 32), suggesting that ClpC could be a modulator of CtsR activity. In this study, we investigated the role of the ClpC and ClpE ATPases in controlling expression of the CtsR-regulated clpP gene in the gram-positive bacterium Lactococcus lactis, which is widely used in production of a variety of dairy products. Our results show that in L. lactis ClpE is involved in restoring the repressed state of clpP expression following a heat shock, and we propose that this effect is mediated through an interaction between CtsR and the zinc-binding motif in the N-terminal region of ClpE. To our knowledge, this is the first report of a role for this motif that is conserved in the ClpE subfamily of Clp ATPases.

 
Bacterial strains, plasmids, and growth conditions. The strains and plasmids used in this study are listed in Table 1. L. lactis strains were grown in M17 (44) supplemented with 0.5% glucose (GM17 medium). Escherichia coli XL1-Blue (Stratagene) was grown in Luria-Bertani broth. When needed, tetracycline (8 µg/ml for E. coli and 2 µg/ml for L. lactis), erythromycin (200 µg/ml for E. coli and 2 µg/ml for L. lactis), or chloramphenicol (25 µg/ml for E. coli and 6 µg/ml for L. lactis) was added. For clpP'-gusA expression studies saturated overnight cultures of L. lactis strains grown at 30°C were diluted 1:1,000 in GM17 medium preheated at 30°C. Strains were grown in water baths at different temperatures, and growth was monitored by determining the optical density at 600 nm (OD₆₀₀). Cell samples (1.5 ml) were collected at appropriate intervals, and cell pellets were stored at -80°C until they were used for ß-glucuronidase (GusA) activity determinations.

 
General DNA techniques, transformation, and DNA amplification. Molecular cloning techniques were performed essentially as described by Sambrook et al. (40). Restriction enzymes, T4 DNA ligase, and deoxyribonucleotides were obtained from New England Biolabs and were used according to the instructions of the supplier. Chromosomal DNA isolation from and transformation of L. lactis were performed essentially as described previously (2, 18). For PCR amplification, the reaction conditions recommended by the manufacturer of DynaZyme DNA polymerase (Finnzymes, Espoo, Finland) were used. PCR products were purified with a Qiagen PCR purification kit.

Construction of chromosomal clpC and clpE deletion mutants. A replacement recombination technique was used to construct two recombinant strains of L. lactis MG1363 with deletions in the clpC or clpE gene. Gene replacement vectors were constructed by using plasmid pGh8 (26) with the thermosensitive pG⁺host origin of replication (6). For the clpC replacement vector, a 2.5-kb region from MG1363 chromosomal DNA was amplified by using primers p1 (5'-ATAAGATATCACTGACAGAACGTGAAG-3') and p2 (5'-ATTTGTCGACTTAATCTCACCCGAGAG-3'). The resulting fragment was digested with EcoRV and SalI, and this was followed by ligation with SmaI/SalI-digested pBluescriptII SK+ (Stratagene). A deletion was made in the 2.5-kb insert by removing a 1.2-kb internal EcoRI fragment. The resulting 1.3-kb insert was cloned as a BamHI-SalI fragment into the corresponding sites of pGh8 to obtain pHI1910, the final clpC replacement vector. The clpE deletion vector was made by amplifying two regions with primers p3 (5'-CATCTCTAGAGGCAGCAGTTGACCAACTC-3') and p4 (5'-GACCTGCAAAGCACTGAAGATATCG-3') and with primers p5 (5'-ATCCTGCAATGGGTGCAAG-3') and p6 (5'-TGATGTCGACTTATCATCTGGTTGGGAAC-3'). The resulting products were cut with XbaI/SspI and SspI/SalI, respectively, and ligated with XbaI/SalI-digested pG⁺host8. The resulting clpE deletion vector (pHI1909) carried 700-bp fragments in both sides of a 2-kb deletion. For integration of replacement vectors, the transformed, tetracycline-resistant L. lactis colonies were grown overnight at 37°C in GM17 broth containing 2 µg of tetracycline per ml and plated on GM17 agar containing 2 µg of tetracycline per ml, and this was followed by incubation overnight at 37°C. To allow excision of the integrated vectors from the chromosome, the integrants were grown overnight at 28°C and plated on GM17 agar plates containing tetracycline (2 µg/ml); this was followed by incubation at 28°C. The excised plasmid was cured by incubating the strains at 37°C in GM17 medium with no antibiotic. Tetracycline-sensitive colonies were tested by PCR for the presence of the wild-type gene or a gene carrying an internal deletion. The clone carrying a 1.2-kb deletion in clpC and the clone carrying a 2-kb deletion in clpE were designated L. lactis HI1924 and HI1931, respectively.

Construction of L. lactis strains with a chromosomal clpP'-gusA transcriptional fusion. To monitor expression of the clpP gene, a 330-bp DNA fragment that included the region from nucleotide -160 to nucleotide 170 with respect to the transcriptional start site of clpP transcription (14) was amplified by PCR by using primers p7 (5'-AACAGATCTAGAGGCCAAAAATCATCG C-3') and p8 (5'-ATATCTGCAGCACGTTCACCACGG-3'). The amplified product was cloned as an XbaI/PstI fragment into the corresponding site in promoter-reporter integration vector pLB85 (9) to obtain pPV33. The pPV33 plasmid was analyzed by sequencing prior to integration into the phage attachment site in L. lactis MG1363 (wild type) and its mutant derivatives PV1 (ctsR) (48), HI1931 (clpE) (this study), and HI1924 (clpC) (this study) to obtain L. lactis PV28, PV29, PV30, and PV31, respectively.

ß-Glucuronidase assays. GusA activity in L. lactis strains was qualitatively assayed on GM17 agar plates containing 0.5 mM 5-bromo-4-chloro-3-indolyl-ß-D-glucuronic acid (X-Gluc) (Biosynth AG, Staad, Switzerland). Fresh colonies were streaked on X-Gluc plates, and the GusA activity was determined visually by accumulation of a blue color after 24 h of incubation at 30 or 37°C.

For quantitative GusA assays cells were grown exponentially to an OD₆₀₀ of 0.3 to 0.4, and samples were harvested and frozen. For quantification cells were thawed on ice and disrupted with glass beads (diameter, 0.1 mm; Sigma) in a homogenizer (Fastprep FB 120; Savant) for 45 s. Disrupted cells were placed on ice and resuspended in 300 µl of ice-cold GusA buffer (34). Cell debris and glass beads were removed by centrifugation at 12,000 x g for 5 min at 4°C. Determination of GusA specific activity in the cell extracts was performed essentially as described previously (37) by using 4-nitrophenyl-ß-D-glucuronic acid (Biosynth AG) at a concentration of 1.5 mM in reaction buffer. The protein concentrations in cell extracts were determined as described by Bradford (8) by using the Bio-Rad protein assay with bovine serum albumin as the standard. Statistical comparisons were made by using Student's t test.

RNA methods. For total RNA isolation from L. lactis strains, the cells were grown exponentially at 25°C in GM17 medium to an OD₆₀₀ of 0.3 to 0.4, after which heat stress was applied by transferring the tubes to 38.5°C. Total RNA was isolated from cell samples incubated for 0, 5, 20, or 45 min at 38.5°C by using an RNeasy Mini kit (Qiagen) as described previously (48). RNA gel electrophoresis, blotting, and hybridization were performed as described previously (36, 48). The clpP- and dnaK-specific probes were obtained by PCR by using primers p9 (5'-CAAATTCTATCATTGCC-3') and p10 (5'-GAGCGATTAGAATTATCAGCAAGG-3') and primers p11 (5'-CTGCTGAAAGCTACCTTGGCG-3') and p12 (5'-CAGCTGGTTGATTATCAGCGG-3'), respectively. Probes were labeled with [-³²P]dATP (>3,000 Ci/mmol; Amersham Pharmacia Biotech). Following hybridization and washes the membranes were scanned and quantified by using a PhosphorImager (Storm system; Molecular Dynamics) and ImageQuaNT (version 4.2; Molecular Dynamics). The amounts of RNA on the membranes were corrected by probing the membranes with a probe specific for L. lactis 16S rRNA obtained by PCR performed with primers p13 (5'-TACGGYTACCTTTGTTACGACT-3') and p14 (5'-AGAGTTTGATCMTGGCTCAG-3').

Site-directed mutagenesis and complementation studies. To complement the chromosomal clpE deletion and to study the role of the putative zinc finger in ClpE, two pCI372 (17) vector-based recombinant plasmids, pPV50 and pPV52, were constructed. The 3.1-kb insert in pPV50 and pPV52 covers a chromosomal region from 700 nucleotides upstream of the ClpE translational start codon, including the putative promoter (19, 48), to 170 nucleotides downstream of the ClpE translation stop codon (19). The inserts of pPV50 and pPV52 were obtained by PCR by using primer p15 (5'-TCTCTAGAGCAGGCAGCAGTTG-3') binding upstream of the putative clpE promoter, as well as primers pzinc1 (5'-ACAGATCTATTTGTTTTTTCTGACCATTTAC-3'), pzinc2 (5'-AAATAGATCTGTGCCAAAACTGTTATCAAA-3'), pzinc3 (5'-AAATAGATCTGAGCCAAAACTGTTATCAAA-3'), and p16 (5'-TCTCGGTCGACTTGATGAGTGGATTGACGA-3'). Primers pzinc1 to pzinc3 bind to the putative zinc finger coding region of ClpE, pzinc1 is a minus-strand primer, and pzinc2 and pzinc3 are plus-strand primers. Primers pzinc1, pzinc2, and pzinc3 contain a new BglII restriction site (underlined) as a silent mutation. In addition, primer pzinc3 contains one nucleotide change (boldface type) compared to pzinc2, which results in to a change from a Cys codon (TGC) to a Ser codon (AGC). The 760- and 2,340-bp PCR products obtained with primers p15 and pzinc1 and with primers pzinc2 and p16, respectively, were digested with XbaI/BglII and BglII/SalI, respectively. The fragments were ligated to pCI372 digested with XbaI/SalI to obtain pPV50 encoding wild-type ClpE. Plasmid pPV52 that encodes ClpE with Ser at position 29 instead of Cys was obtained like pPV50, except that primer pzinc3 was used instead of pzinc2 in the PCR. The 3.1-kb inserts of pPV50 and pPV52 were sequenced, and no additional mutations were observed. The pPV50 and pPV52 plasmids were transformed into clpE strain L. lactis PV30, and expression of clpP'-gusA was measured with cells growing exponentially at 30 or 38.5°C. The effect of pPV50 on heat shock induction of clpP and dnaK was studied by performing a Northern blot analysis after transformation in L. lactis HI1931 (clpE).

ClpE purification, antibody production, and Western blot analysis. For purification of His₆-ClpE the clpE gene was amplified with primers p17 (5'-AAAGGATCCCTTTGTCAAAATTGTAATATTAATG-3') and p16 and cloned as a BamHI/SalI fragment into the pQE30 QIAexpress vector (Qiagen) to obtain pPV53. Recombinant N-terminal His-tagged ClpE was expressed and purified by standard procedures described by the manufacturer (Qiagen) and was used for antibody production in rabbits. For Western blot analysis cells were grown in GM17 medium at an elevated temperature (usually 38.5°C; the exception was the clpP mutant strain, which was grown at 37°C) until the OD₆₀₀ reached 0.4 to 0.5, and cell samples were harvested. Protein samples were separated by using the NuPAGE bis-Tris electrophoresis system (Invitrogen) and blotted onto nitrocellulose filters (pore size, 0.45 µm; Bio-Rad). Colorimetric detection of ClpE was carried out by using a rabbit polyclonal antibody (1:3,000) raised against His₆-ClpE as the primary antibody and an anti-rabbit immunoglobulin G-avidin-horseradish peroxidase conjugate (1:3,000) (Bio-Rad) as the secondary antibody. The colorimetric reaction for the filter was carried out according to the instructions provided by the manufacturer (Bio-Rad). Alternatively, a 1:40,000 dilution of the anti-ClpE antibody and a 1:50,000 dilution of the horseradish peroxidase-conjugated goat anti-rabbit immunoglobulin G (Bio-Rad) was used when the detection system based on chemiluminescence was used. Visualization of ClpE was performed with a SuperSignal West Dura extended duration kit used according to the instructions provided by the manufacturer (Pierce). Membranes were scanned with a GS-525 molecular imager system (Bio-Rad) and were analyzed with MultiAnalyst and QuantityOne software (Bio-Rad).

 
L. lactis ClpE affects expression of clpP at a high temperature. In L. lactis expression of clpP is negatively regulated by CtsR (48). To investigate if the Clp ATPases, ClpC and ClpE, play a role in controlling L. lactis clpP gene expression, we constructed a transcriptional fusion of the clpP promoter region to a reporter gene, gusA encoding ß-glucoronidase (20), and inserted it into the chromosomal attachment site (attB) of phage TP901-1 (10). When we introduced this fusion into wild-type cells (MG1363) and ctsR mutant cells (PV29), we found that at 30°C clpP gene expression was increased sixfold in the absence of CtsR, which confirmed the CtsR-dependent regulation of clpP expression (data not shown). When we subsequently introduced the fusion into PV31, from which the clpC gene had been deleted, measurements of GusA activity revealed that clpP expression was unaffected by the clpC mutation at both 30 and 38.5°C (data not shown). However, in cells carrying the clpP'-gusA fusion from which clpE was deleted (PV30), the calculated mean value for the GusA specific activity, 0.124 U/mg (standard deviation, 0.035 U/mg), was significantly (P < 0.0001, as determined by Student's t test) higher than the value measured for wild-type cells growing at 38.5°C, 0.050 U/mg (standard deviation, 0.020 U/mg) (Fig. 1). At 30°C clpP expression was unaffected by the clpE deletion (data not shown). These data suggest that ClpE represses clpP gene expression at an elevated temperature.

 
ClpE is required for efficient re-repression of clpP expression following heat shock. To further examine the role of ClpE in clpP gene expression at a high temperature, we determined the amount of clpP mRNA by Northern blot analysis using RNA isolated from wild-type, PV1 (ctsR), and HI1931 (clpE) mutant cells. Figure 2 shows that before heat induction equal amounts of clpP mRNA were present in wild-type and HI1931 cells, while the amount was greater in ctsR mutant cells. After cells were shifted to 38.5°C, clpP expression in the wild-type and HI1931 strains rapidly increased to about the same level found in PV1 before and after the shift. When incubation was continued at a high temperature, the induction of clpP expression in wild-type cells was followed by repression, which was not observed in ctsR mutant cells. Interestingly, this repression was greatly delayed in clpE mutant cells, which resulted in sevenfold more clpP transcript in HI1931 cells than in wild-type cells 45 min after the shift to 38.5°C. To confirm that the prolonged induction of clpP expression in the clpE mutant was caused by the absence of the clpE gene product, the clpE deletion mutant (HI1931) was transformed with a plasmid encoding ClpE (pPV50) and was analyzed by Northern blotting. Figure 3A shows that the patterns of heat shock induction of clpP were similar in wild-type cells carrying the pCI372 vector and in HI1931 complemented with pPV50, while HI1931 carrying the pCI372 vector showed prolonged induction of clpP expression of heat shock.

 
Absence of ClpE does not affect expression of an HrcA-controlled gene. One way that ClpE might influence ClpP gene expression is if it acts as a general chaperone. The absence of ClpE could thus lead to accumulation of misfolded proteins to which CtsR responds and subsequent induction of the CtsR regulon (15). To examine this possibility, we investigated whether the clpE deletion affected the expression of dnaK, which belongs to the HrcA regulon, which is also known to respond to misfolded proteins (30). Northern blot analysis revealed that the clpE deletion did not affect dnaK expression following a heat shock, suggesting that the effect is confined to the CtsR regulon (Fig. 3B).

Stability of clpP mRNA is not elevated in the clpE mutant cells at 38.5°C. To examine the possibility that the clpE deletion affects the stability of clpP mRNA at a high temperature, the rate of clpP mRNA decay was investigated in both wild-type and clpE mutant cells. Both strains were grown at 38.5°C to an OD₆₀₀ of 0.3, and following inhibition of transcription by rifampin samples were withdrawn and the time-dependent decay of the clpP mRNA was determined by Northern blot analysis (Fig. 4A). A regression analysis of the data (SigmaPlot program; SPSS Inc., Chicago, Ill.) is shown in Fig. 5B. The results of two independent experiments indicated that the half-life of clpP mRNA in both wild-type cells (MG1363) and cells from which clpE was deleted was approximately 2 min. Thus, deletion of clpE does not affect the stability of the clpP transcript.

 
ClpE zinc finger motif is required for re-repression of clpP expression. Members of the ClpE family of proteins have a highly conserved zinc finger motif in the N-terminal domain (11, 19). In order to determine the role of this motif in ClpE-mediated regulation of clpP gene expression, we constructed an additional plasmid carrying a ClpE derivative in which cysteine residue 29 was replaced by a serine (pPV52), which disrupted the zinc finger motif. At 30°C both pPV50 (ClpE) and pPV52 (ClpE-C29S) had only minor effects on the steady-state level of clpP expression monitored by using the chromosomally located clpP-gusA fusion (Fig. 5A). At 38.5°C introduction of pPV50 into the clpE deletion strain, PV30, resulted in repression of the clpP'-gusA fusion to a level comparable to that in wild-type cells carrying the vector alone (Fig. 5B). However, when the vector was introduced into PV30 and when pPV52 expressing the mutated ClpE was introduced into PV30, the GusA activities were 2.5- and 2-fold greater than the activity in wild-type cells, respectively (Fig. 5B), suggesting that the ClpE zinc finger motif is required for restoring the ClpE-mediated repression of clpP gene expression.

ClpP-dependent processing of ClpE. In order to determine the amount of ClpE expressed at an elevated temperature (38.5°C), we expressed and purified a His-tagged ClpE and used this tagged ClpE to raise ClpE-specific antibodies. When we analyzed cell lysates from wild-type and ClpE mutant cells by Western blotting, we observed two ClpE-specific bands (Fig. 6A, lanes 1 and 2) in which the amount of the higher-molecular-weight ClpE (ClpE1) greatly exceeded the amount of the lower-molecular-weight ClpE (ClpE2). In cells lacking ClpE two cross-reacting bands still remained (Fig. 6A, lane 2). The sizes of these bands correspond to the sizes of the two ClpB proteins observed in a previous study (18). This finding was confirmed as the bands were not produced by a clpB deletion strain (Fig. 6A, lane 4).

 
When we determined the amount of ClpE in clpE mutant cells carrying either pPV50 expressing ClpE or pPV52 expressing ClpE-C29S (Fig. 6B, lanes 3 and 4), we found that both plasmids restored the total amount of ClpE (ClpE1 plus ClpE2) to a level greater than that in the wild type (Fig. 6B, lane 1), showing that the inability of ClpE-C29S to repress clpP expression (Fig. 4) is not due to expression of less protein. Interestingly, we observed that in cells producing ClpE-C29S the amount of ClpE2 was greatly increased compared to the amount ClpE1 (Fig. 6B, lane 4) and that in cells lacking the Clp protease gene, clpP (14), the amount of ClpE2 was less than the detection limit (Fig. 6A, lane 5). These results indicate that a small amount of ClpE is processed in a ClpP-dependent manner and that the zinc finger protects the protein against such processing. To confirm this, we introduced pPV52 into the wild-type strain and the clpP deletion strain and verified that processing of ClpE-C29S was eliminated (Fig. 6C, lanes 2 and 4). The processing of ClpE appears to be independent of ClpC and ClpB since mutations in either of the corresponding genes affected the ClpE2/ClpE1 ratio (Fig. 6A, lanes 3 and 4).

In addition to the specific processing of ClpE, the clpP mutation also increased the total amount of ClpE present (Fig. 6A, lane 5, and Fig. 6C, lanes 3 and 4), and the increase was accompanied by increased transcription of the gene (data not shown). Since clpE expression is negatively regulated by CtsR, our data suggest that L. lactis CtsR is a target for the ClpP protease, as has been observed in B. subtilis (12, 21).

 
Transcriptional regulators and alternative factors play crucial roles in the survival of bacteria in stress situations. In several gram-positive bacteria two major negative regulators have been identified as part of the heat shock regulatory network. HrcA controls expression of the chaperone homologues DnaK, DnaJ, GrpE, and GroEL, while CtsR primarily regulates expression of the genes encoding the Clp ATPases and the ClpP protease (12, 42). Recently, several studies have reported that the Clp ATPases influence CtsR activity. In L. monocytogenes deletion of clpC greatly increased expression of the CtsR-regulated clpE gene in the absence of heat shock, suggesting that there is regulatory cross talk between the Clp ATPases (32). Additionally, ClpC appears to be involved in transcriptional regulation of a number of other genes in this organism (34). In B. subtilis the ClpCP complex degrades CtsR at elevated temperatures following modification by the McsB arginine kinase, while CtsR is stabilized in the absence of heat shock by interaction with the zinc-binding protein McsA (23). At low temperatures CtsR is a target of the ClpXP complex, possibly to ensure low-level expression of the CtsR regulon (13).

In the present study we investigated the role of the ClpC and ClpE ATPase homologues in modulating the expression of clpP, which belongs to the CtsR regulon in L. lactis. We found that while ClpC does not affect clpP expression, ClpE is required for the normal repression of clpP expression that occurs following a heat shock. The N-terminal region of ClpE, including a putative zinc finger (19), is important for this activity as replacement of the cysteine at residue 29 with a serine resulted in prolonged heat shock induction. The effect of ClpE on clpP expression is likely to be mediated through CtsR, as heat shock induction in the absence of CtsR and heat shock induction in the absence of ClpE induced clpP equally (Fig. 2). Thus, ClpE could be a chaperone that modulates the activity of CtsR. However, if this is true, ClpE appears to have a narrow substrate specificity as the absence of ClpE did not lead to induction of the HrcA regulon known to respond to misfolded protein (30). Alternatively, ClpE could interact with CtsR in a reaction resembling the reaction of McsA and CtsR in B. subtilis. In fact, searches of the genome sequences of L. lactis (7), Streptococcus pyogenes (4), Streptococcus pneumoniae (45), Streptococcus mutans (1), Streptococcus agalactiae (46), and Lactobacillus plantarum (21) showed that while these organisms encode CtsR homologues, they appear to lack McsA counterparts (data not shown). Intriguingly, the N-terminal zinc finger domains of ClpE of these bacteria exhibit 31 to 41% identity to the first 32 amino acids of B. subtilis McsA and its homolog in Staphylococcus aureus (Fig. 7). As both ClpE and MscA have positive effects on the activity of CtsR, it is tempting to speculate that ClpE is a functional homologue of McsA.

 
While measuring the amount of ClpE by Western blotting, we observed two forms of ClpE, one whose size corresponded to the size of full-length ClpE (ClpE1) and the other approximately 6-kDa smaller (ClpE2). In cells lacking ClpP the smaller form of the protein was absent, whereas in cells expressing the mutated ClpE-C29S the ClpE2 form was predominant. These results suggest that the zinc-binding site and possibly zinc binding in the N-terminal region of ClpE protect the protein against ClpP-dependent processing. While ClpP target proteins are normally degraded to small peptides, a few examples of ClpP-dependent processing exist. In E. coli autoprocessing of ClpP removes the N-terminal 14 amino acids (27), and in Streptomyces coelicolor that contains four ClpP homologues, ClpP1 and ClpP2 appear to cross-process each other from the N terminus (49). At this time, the physiological role of the ClpP-dependent processing of ClpE remains obscure; however, the small amount of ClpE2 observed in cells expressing wild-type ClpE shows that the processing takes place under normal growth conditions. Also, it remains to be determined if it is only the processed form of ClpE-C29S that is defective in repression of clpP expression. However, our data show that ClpE is important in reestablishing repression of a CtsR-regulated gene following heat shock. Furthermore, as shown previously for ClpX (3), the zinc-binding site is required for full ClpE activity.

 
We thank D. Frees, K. Savijoki, M. Kilstrup, and A. K. Nielsen for helpful discussions throughout this work. We are grateful to C. Rasmussen for excellent technical assistance.

The Danish Dairy Research Foundation, The Danish Food Research Programme (FØTEK-2) through The Centre for Advanced Food Studies (LMC), and the Academy of Finland financed this work.

 
* Corresponding author. Mailing address: University of Helsinki Faculty of Veterinary Medicine, Department of Basic Veterinary Sciences, Section of Microbiology, P.O Box 57, 00014 Helsinki University, Finland. Phone: 358 9 19149787. Fax: 358 9 19149799. E-mail: pekka.varmanen@helsinki.fi.

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