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Journal of Bacteriology logoLink to Journal of Bacteriology
. 2008 May 23;190(15):5153–5161. doi: 10.1128/JB.00437-08

Complex Regulation of the DnaJ Homolog CbpA by the Global Regulators σS and Lrp, by the Specific Inhibitor CbpM, and by the Proteolytic Degradation of CbpM

Matthew R Chenoweth 1, Sue Wickner 1,*
PMCID: PMC2493262  PMID: 18502857

Abstract

CbpA is a DnaJ homolog that functions as a DnaK cochaperone. Several cellular processes, including growth at low and high temperatures and septum formation during cell division, require either CbpA or DnaJ. CbpA is encoded in an operon with the gene for CbpM, which is a specific in vivo and in vitro inhibitor of CbpA. Here, we have cooverexpressed CbpA with CbpM in a ΔcbpAM ΔdnaJ strain and examined the resulting phenotypes. Under these conditions, sufficient free CbpA activity was present to support growth at low temperatures, but not at high temperatures. Defects in cell division and in λ replication were also partially complemented by CbpA when cooverexpressed with CbpM. Utilizing reporter fusions, we demonstrated that the cbpAM operon was maximally transcribed at the transition from exponential growth to stationary phase. Transcription was controlled by the σS and Lrp global regulators, and both leucine availability and growth temperature influenced transcription. CbpA and CbpM accumulated to similar levels in stationary phase, ∼2,300 monomers per cell. When not bound to CbpA, CbpM was unstable and was degraded by the Lon and ClpAP proteases. These data demonstrate that CbpA activity is controlled at multiple levels.


Chaperones assist in numerous cellular functions during both normal growth and times of stress. The DnaK/Hsp70 system is a universally conserved chaperone machine and is composed of DnaK and two cochaperones, DnaJ and GrpE, in Escherichia coli (8). DnaK is an energy-dependent chaperone that promotes remodeling of substrate polypeptides. DnaJ assists DnaK by facilitating substrate delivery and by stimulating the intrinsically weak ATPase activity of DnaK. GrpE stimulates nucleotide exchange, thereby assisting substrate release.

In addition to dnaJ, the E. coli chromosome contains genes for five other DnaJ homologs: cbpA, djlA, hscB, ybeS, and ybeV. CbpA (curved DNA binding protein A) functions not only as a multicopy suppressor for dnaJ mutations in vivo, but also as a cochaperone for the DnaK system in vitro (9, 27, 28). Unlike DnaJ, CbpA binds DNA efficiently and is associated with the nucleoid of stationary-phase cells (1, 27). CbpA is encoded in an operon that codes for a second protein, CbpM, which interacts specifically with CbpA to inhibit both the cochaperone and DNA binding activities of CbpA (9). CbpM homologs exist in a number of diverse bacteria, including the intracellular human pathogen Coxiella burnetii and the iron-reducing environmental organism Geobacter sulfurreducens. As in E. coli, CbpM homologs in other organisms are typically encoded in an operon that also codes for a CbpA-like protein (9). The common genetic organization suggests a conserved interaction between the CbpA and CbpM homologs. The fact that these homologs exist in diverse organisms with widely varying lifestyles implies that CbpA and CbpM fulfill basic physiological roles.

Previously, we demonstrated that CbpM binds to CbpA in vivo, resulting in an inactive complex (10). Overexpression of CbpM generates a ΔcbpA phenocopy for all known ΔcbpA phenotypes, which are evident only in ΔdnaJ strains due to functional overlap between CbpA and DnaJ (10). However, CbpM was present at much higher levels than CbpA in the experiments that detailed this inhibition. Since CbpA and CbpM are encoded by the same operon (9), we wanted to determine if CbpM inhibits CbpA in vivo when the two proteins are coexpressed. Also, we examined the regulation of the operon and the levels of CbpA and CbpM within the cell. CbpM partially inhibited CbpA when the two genes were coexpressed, and some phenotypes were more affected than others. The cbpAM operon was controlled at the level of transcription by σS and Lrp, and the two proteins accumulated to similar levels. Finally, we found that CbpM was degraded in a lon- and clpP-dependent manner in vivo and by Lon and ClpP in vitro when it was not bound to CbpA.

MATERIALS AND METHODS

Strains and culture conditions.

Strains, plasmids, and phages used in this study are listed in Table 1. All strains used were derivatives of BW27784, a derivative of MG1655 (15). Transductions with P1vir were performed as described previously (19). The lon::Tn10, clpP::kan, and rpoS::kan alleles were acquired from Brill et al. (6), Sledjeski et al. (24), and Bohannon et al. (4), respectively. Unless noted otherwise, all strains were grown in LB and incubated at 30°C.

TABLE 1.

Strains utilized in this study

Strain, plasmid, or phage (Parental strain) description Reference or source
Strains
    BW27784 (BW25113) DE(araFGH) φ(ΔaraEp PCP18-araE) 15
    DY330 (W3110) ΔlacU169 gal490 λcI857 Δ(cro-bioA) 31
    MC105 (BW27784) ΔcbpA3::cat 9
    MC108 (BW27784) ΔcbpM3::cat 9
    MC125 (MC105) lon::Tn10 This study
    MC127 (MC105) ΔclpP::kan This study
    MC143 (BW27784) ΔcbpAM3::cat 9
    MC144 (MC143) ΔdnaJ::kan 9
    MC150 (BW27784) ΔdnaJ::kan 9
    MC153 (BW27784) λMRC133 (cbpA1::lacZ) This study
    MC155 (BW27784) λMRC135 (cbpAM1::lacZ) This study
    MC158 (BW27784) λMRC138 (cbpAM2::lacZ) This study
    MC160 (BW27784) λMRC140 (cbpA2::lacZ) This study
    MC163 (MC153) ΔrpoS::tet This study
    MC247 (BW27784) λMRC141 (cbpM1::lacZ) This study
    MC342 (BW27784) Δlrp3::cat This study
    MC351 (MC153) Δlrp3::cat This study
    MC357 (MC163) Δlrp3::cat This study
    MC367 (BW27784) λDDS314 (dsrB1::lacZ) This study
    MC371 (MC367) Δlrp3::cat This study
Plasmids
    pBAD24 14
    pcbpA+ pBAD24 with cbpA insert 9
    pcbpM+ pBAD24 with cbpM insert 9
    pcbpAM+ pBAD24 with cbpAM insert This study
    pRS1551 Plasmid for construction of translational fusions 23
    LacZ
        pRS1553 Plasmid for construction of transcriptional fusions 23
    lacZ
        pMRC03 pRS1553 with cbpA1::lacZ This study
        pMRC05 pRS1553 with cbpAM1::lacZ This study
        pMRC08 pRS1551 with cbpAM2::lacZ This study
        pMRC10 pRS1551 with cbpA2::lacZ This study
        pMRC11 pRS1553 with cbpM1::lacZ This study
Phage
    λRS468 Phage containing ω fragment of lacZ 23
    λDDS314 imm21dsrB1::lacZ 24
    λMRC133 imm21cbpA1::lacZ λRS468 × pMRC03
    λMRC135 imm21cbpAM1::lacZ λRS468 × pMRC05
    λMRC138 imm21cbpAM2::lacZ λRS468 × pMRC08
    λMRC140 imm21cbpA2::lacZ λRS468 × pMRC10
    λMRC141 imm21cbpM1::lacZ λRS468 × pMRC11

Strain and plasmid construction.

To construct plasmid pcbpAM+, cbpAM was amplified by PCR using primers listed in Table 2 and digested with EcoRI and HindIII. The inserts were ligated into pBAD24 (14). The ligated constructs were electroporated into DH5α, purified, sequenced, and electroporated into the BW27784 derivatives.

TABLE 2.

Oligonucleotides used for plasmid and strain construction and for Northern blotting

Oligonucleotide Sequence (5′-3′)
Construction of plasmids
    cbpAMfor CGCGGAATTCACCATGGAATTAAAGGATTATTACGCCATCATG
    cbpAMrev CGCCTGCAGTCACGGATGAGCTACAAACCGGGAAAGCCG
Chromosomal mutations
    Δlrp3for CAGACAGGAGTAGGGAAGGAATACAGAGAGACAATAATATTACGCCCCGCCCTGCCACTC
    Δlrp3rev TTAGCGCGTCTTAATAACCAGACGATTACTCTGCTTGACTTACGCCCCGCCCTGCCACTC
Construction of lacZ fusions
    MC.026 CGCGAATTCTCAACTATCAAAAATCGCTCACCC
    MC.028 CGCGGATCCCGACGTGGATCAAAAGACGACTGGGCG
    MC.030 CGCGGATCCCGATGAGCTACAAACCGGGAAAGCCG
    MC.046 CGCGAATTCCATCGCAACGATCCGCAATTTAACCGT
Biotinylated probes for Northern blots
    cbpA probe CCACGGGCTAACCGGCACCACAATTTCCAGATCCTGGCCG
    cbpM probe CCAGTTCATGACGCAGGCGTACCGCGCGTTGCACCACAAT
    crl probe CGTTGACGCATACAGCCAGACAATCGAAAAAGAATCGATT
    dps probe TGTTCCAGTGCGCTTGTTTGGTAATCAAAGAAAGATCAAT

lacZ reporter constructs were made using plasmids pRS1551 and pRS1553, which encode the LacZ α-peptide and are derivatives of vectors pRS551 and pRS552 (23). Transcriptional (cbpA1::lacZ) and translational (cbpA2::lacZ) cbpA fusions were created with PCR products of cbpA that included 200 bp upstream of the ATG start codon and the first 900 bp of the 921-bp cbpA gene. The primers used for amplification were MC.026 and MC.028 (Table 2). Transcriptional (cbpAM1::lacZ) and translational (cbpAM2::lacZ) cbpM fusions were created with PCR products of cbpAM that included 200 bp upstream of the ATG start codon, the entire cbpA gene, and the first 300 bp of the 306-bp cbpM gene. The primers utilized were MC.026 and MC.030 (Table 2). Additionally, a cbpM transcriptional fusion lacking the cbpAM promoter (cbpM1::lacZ) was created with a PCR product that included bases 211 to 921 of cbpA and the first 300 bp of cbpM. The primers utilized were MC.046 and MC.030 (Table 2). The amplified products were cloned into the EcoRI and BamHI sites of pRS1551 and pRS1553 for translational and transcriptional fusions, respectively (Table 1). The resulting plasmids were crossed with λRS468, which contains the ω fragment of lacZ (23), to obtain the corresponding transducing phage. As such, the transducing phage carries either cbpA or cbpAM fragments fused to a complete lacZ gene. The recombinant phage were used to lysogenize BW27784 (Table 1). Monolysogens were identified as described previously (20).

The lrp deletion/insertion strain was created as previously described (31). Chloramphenicol cassettes were amplified using the oligonucleotide primer pair Δlrp3for and Δlrp3rev (Table 2). The resulting Δlrp3 allele removed all sequence between the lrp start and stop codons and was confirmed by sequencing.

Lambda replication.

Plaquing efficiency (2) and burst size (16) were determined as described previously. A multiplicity of infection of ∼3 was used to determine burst size in the single-step infections.

Microscopy.

Cell morphology was examined by phase-contrast as previously described (10).

Western blotting.

Cultures were adjusted to an optical density at 595 nm (OD595) of 0.4, precipitated with 10% trichloroacetic acid (vol/vol), separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), and transferred to nitrocellulose. The protein of interest was detected by probing with specific rabbit serum using a Western immunodetection kit (WesternBreeze; Invitrogen, Carlsbad, CA).

RNA isolation and Northern blotting.

RNA was isolated by hot-phenol extraction, and DNA was removed (17). Northern blotting was performed as described previously (17). The biotinylated probes used for detection of cbpA, cbpM, crl, and dps are listed in Table 2. Detection was performed with a BrightStar Biodetect kit (Ambion, Austin, TX) according to the manufacturer's instructions.

β-Galactosidase assays.

Samples from strains containing lacZ fusions were lysed in Fastbreak lysis buffer (Promega, Madison, WI), and the levels of β-galactosidase activity were determined by utilizing a β-galactosidase enzyme assay system (Promega). Activity was determined as Miller units (22).

Quantitative Western blots.

Stationary-phase culture samples were precipitated with trichloroacetic acid, and multiple dilutions of the experimental sample were loaded onto the gel with known concentrations of the purified protein (1 to 15 ng). After SDS-PAGE separation and Western immunodetection, the film was scanned and analyzed with ImageJ software (http://rsb.info.nih.gov/ij/). A standard curve was constructed using the purified proteins, and the amount of CbpA or CbpM per sample was calculated in the linear range of the standard curve. These data were used in conjunction with plate counts to determine the number of monomers of each protein per cell.

Protein purification.

CbpA (27), CbpM (9), ClpP (18), ClpA (18), and ClpX (18) were isolated as described previously. Lon was a gift from Michael Maurizi (National Institutes of Health, Bethesda, MD).

RESULTS

Cooverexpression of CbpM with CbpA in a ΔdnaJ ΔcbpAM strain inhibits growth at high but not low temperatures.

When overexpressed from a plasmid, CbpM inhibits CbpA's ability to support growth at high and low temperatures in a ΔdnaJ strain (10). Since cbpA and cbpM are located in the same operon, we wanted to know whether CbpM exerts its inhibitory effect when cooverexpressed with CbpA. To address this question, the plating efficiencies of ΔdnaJ ΔcbpAM triple-deletion strains constitutively expressing CbpA or both CbpA and CbpM from a plasmid were examined over a range of temperatures. The ΔdnaJ ΔcbpAM strain grew at 25 and 30°C, although growth was reduced by ∼100-fold compared to the wild type (10) (Fig. 1A, columns 1 and 2). Growth of the triple-deletion strain was reduced ∼10,000-fold at 16 and 37°C compared to the wild type, and there was no detectable growth at 42°C. When CbpA was reintroduced to the ΔdnaJ ΔcbpAM strain via a multicopy plasmid, growth at 16, 25, 30, and 37°C was restored to wild-type levels (Fig. 1A, column 3). At 42°C, CbpA overexpression was only sufficient to allow low-level growth, ∼10,000-fold lower than the wild type (Fig. 1A, column 3). Overexpression of CbpA and CbpM together from the pcbpAM+ multicopy plasmid in the ΔdnaJ ΔcbpAM strain yielded unexpected results. At 16, 25, and 30°C, this strain exhibited wild-type growth, indicating that there was sufficient CbpA activity to allow growth at low temperatures (Fig. 1A, column 4). However, at 37 and 42°C, cooverexpression of CbpA and CbpM in the ΔdnaJ ΔcbpAM strain resulted in severe growth defects, similar to those evident in the ΔdnaJ ΔcbpAM triple mutant (Fig. 1A, column 4). Taken together, these results indicate that when CbpM is present with CbpA, there is sufficient CbpA activity to support growth at low temperature, but not at high temperature.

FIG. 1.

FIG. 1.

In vivo effects of cooverexpression of CbpA and CbpM. (A) E. coli strains were grown in LB for 24 h at 25°C and serially diluted as indicated, and 10 μl of each dilution was plated on LB agar. The plates were incubated at the indicated temperatures. Ampicillin (50 μg/ml) and arabinose (0.02%) were added to the plates for strains containing plasmids. The relevant genotypes are shown at the top. Column 1, wild type; column 2, ΔdnaJ ΔcbpAM; column 3, ΔdnaJ ΔcbpAM pcbpA+; column 4, ΔdnaJ ΔcbpAM pcbpAM+. (B) E. coli strains were grown in LB overnight (16 h) at 25°C, and the cell morphologies were examined by phase-contrast microscopy. Ampicillin (50 μg/ml) and arabinose (0.02%) were added to cultures of cells containing plasmids.

Cooverexpression of CbpA and CbpM partially suppresses the cell division defect of ΔdnaJ ΔcbpAM cells.

Our previous work showed that (i) a ΔdnaJ ΔcbpAM strain exhibits filamentous cell morphology and overexpression of CbpA restores the wild-type rod-shaped morphology and (ii) overexpression of CbpM in a ΔdnaJ strain results in a filamentous morphology like that of a ΔdnaJ ΔcbpA or a ΔdnaJ ΔcbpAM strain (10). We wanted to determine if CbpA could repair the cell division defect in ΔdnaJ ΔcbpAM cells when it was cooverexpressed with CbpM at 25°C. Interestingly, this resulted in slightly elongated cell morphology compared to the wild type (Fig. 1B, images 1 and 2). ΔdnaJ ΔcbpAM cells bearing the pcbpAM+ plasmid did not form the filaments typical of ΔdnaJ ΔcbpAM cells (Fig. 1B, image 3) or short rods typical of a ΔdnaJ ΔcbpAM strain overexpressing CbpA alone (Fig. 1B, image 4). Thus, cooverexpression of CbpM with CbpA in a ΔdnaJ ΔcbpAM strain at 25°C results in sufficient CbpA activity to correct the defect in cell division to nearly the wild-type state.

Overexpression of CbpA with CbpM only partially suppresses the growth defect of bacteriophage λ on ΔdnaJ strains.

Because cooverexpression of CbpM with CbpA prevented CbpA from participating in cell growth under some conditions but not others, we were interested to know how λ replication would be affected in a ΔdnaJ background when both cbpA and cbpM were coexpressed on a multicopy plasmid. Previous work showed that λ is unable to grow on dnaJ mutant strains (25) (Table 3) due to a defect in DNA replication (21, 30) and that multicopy cbpA complements the defect (27, 28) (Table 3). Interestingly, when we cooverexpressed CbpM with CbpA in a ΔdnaJ strain, the plaquing efficiency of λ was not significantly reduced (Table 3). However, the plaques formed on the ΔdnaJ pcbpAM+ strain were noticeably smaller than those produced on either the wild-type or ΔdnaJ pcbpA+ strain.

TABLE 3.

Lambda plaquing efficiencies and burst sizes

Parameter Value
WT ΔdnaJ ΔdnaJ pcbpA+ ΔdnaJ pcbpAM+
PFUa 1.2 ± 0.2 × 1010 <103 7.3 ± 2.8 × 109 6.0 ± 1.4 × 109
Efficiency (%)b 100 <10−5 62 51
Burst sizec 170 ± 70 1.4 ± 0.4 147 ± 64 26 ± 4.5
a

PFU is the average ± standard deviation of three independent experiments.

b

The efficiency of the wild type (WT) was set to 100%, and those of the other strains are relative to it.

c

Burst size is the average ± standard deviation of three independent experiments.

To investigate the difference in plaque size, the burst sizes were measured. The ΔdnaJ cells overexpressing CbpA produced nearly wild-type levels of phage, but ΔdnaJ cells overexpressing both CbpA and CbpM produced about sixfold-lower levels of phage (Table 3). Therefore, in the absence of DnaJ, CbpM limits λ replication when CbpA and CbpM are coexpressed from a plasmid, indicating partial inhibition of CbpA activity.

CbpA and CbpM are coexpressed during the growth cycle.

As it is perplexing that the gene for an inhibitor of CbpA appears in the same operon with cbpA, we wanted to know if there are some situations in vivo when CbpA is in excess of CbpM.

To determine if cbpA and cbpM may be differentially regulated, the expression of these two genes was examined by Northern blot analysis using oligonucleotides specific for cbpA and cbpM (see Fig. S1 in the supplemental material). Both genes were transcribed on a 1.4-kb mRNA, indicating that they are likely cotranscribed (Fig. 2A). The expression of cbpA and cbpM reached a maximum during the transition from exponential-phase growth to stationary phase, and then transcription of both genes rapidly decreased to almost undetectable levels (Fig. 2A). This pattern of transcription was also displayed by dps, a σS-dependent stationary-phase gene, and by crl, a σS-independent stationary-phase gene (see Fig. S2 in the supplemental material), suggesting that cbpA and cbpM transcription is typical of stationary-phase genes. Importantly, the pattern of expression of cbpA was indistinguishable from that of cbpM.

FIG. 2.

FIG. 2.

Expression of cbpA and cbpM during growth and dependence upon global regulators. (A) Northern analysis of cbpA and cbpM transcription. Wild-type E. coli was grown in LB at 30°C, and RNA samples were collected at the indicated cellular densities during the growth cycle. Transcripts were detected by Northern blotting utilizing biotinylated oligonucleotides specific for cbpA (top) or cbpM (bottom). (B) Transcriptional analysis of cbpA and cbpM utilizing lac fusions. E. coli strains bearing cbpA or cbpAM transcriptional fusions were grown in LB at 30°C, and β-galactosidase activity from cbpA1::lacZ (circles) or cbpAM1::lacZ (squares) was determined throughout the growth cycle at the indicated time points. Each time point represents the average ± standard deviation (SD) of three independent assays. Diamonds, OD595 measurements of the culture. (C) Roles of σS, Lrp, and leucine in the transcription of cbpA. E. coli strains containing cbpA1::lacZ fusions were grown at 30°C in MOPS (morpholinepropanesulfonic acid) minimal medium supplemented with 0.2% glucose, 0.4 mM valine, and 0.4 mM isoleucine (26) to late stationary phase (OD595, ∼2.5; 24 h of growth) and assayed for β-galactosidase activity. Combinations of rpoS and lrp deletions were introduced as indicated. The light-gray bars indicate that no leucine was added to the medium (− leu), and the dark-gray bars indicate 0.4 mM leucine was added to the medium (+ leu). The values represent the average ± SD of at least three independent assays. (D) Role of growth temperature in the transcription of cbpA in exponential phase. Cultures were grown to mid-exponential phase (OD595, ∼0.3; 7 h of growth), and the transcriptional activity of the cbpA promoter was monitored as for panel C. Combinations of lrp deletion and leucine addition were introduced as indicated. (E) Role of growth temperature in the transcription of cbpA in stationary phase. Transcription of cbpA in late-stationary-phase cultures (OD595, ∼2.5; 24 h of growth) grown at either 30 or 37°C was determined as in panel D.

To further monitor the expression of cbpA and cbpM, transcriptional lacZ gene fusions were made to cbpA and to cbpAM and inserted into the chromosome in single copies. Using these fusions, we found that the transcriptional patterns of cbpA and cbpAM were similar (Fig. 2B). With both reporter fusions, transcription was low during exponential growth, increased sharply as cells transitioned to stationary phase, and leveled off in stationary phase (Fig. 2B), similar to the previously reported cbpA transcriptional profile (29). Approximately 40% more β-galactosidase was produced by the cbpAM1::lacZ fusion than by the cbpA1::lacZ fusion, possibly due to differences in mRNA stability. However, the ratio of β-galactosidase produced by the cbpAM fusion compared to that produced by the cbpA fusion was nearly constant at 1.4 ± 0.1 during the growth period. Additionally, a cbpM transcriptional fusion (cbpM1::lacZ) lacking the promoter upstream of cbpA failed to produce β-galactosidase activity, indicating that cbpM probably does not have a separate promoter (see Fig. S3 in the supplemental material). These observations, together with the Northern blot analysis, show that cbpA and cbpM follow similar transcription profiles throughout growth.

The roles of σS and Lrp in cbpAM transcription.

Previously, transcription of cbpA was shown to be induced by σS during stationary phase (29) and repressed by Lrp during exponential growth (26). We wanted to further study the effects of these global regulators on the cbpAM operon to determine if there was interplay between the two regulatory systems. After 24 h of growth (OD595, ∼2.5), a ΔrpoS strain produced ∼80% less β-galactosidase activity from the cbpA1::lacZ transcriptional fusion than from the wild-type strain (Fig. 2C). Introduction of the Δlrp3 allele to the wild type resulted in a ∼40% decrease in cbpA promoter activity (Fig. 2C). Interestingly, introduction of the Δlrp3 allele into a strain already bearing the rpoS deletion had no further effect upon cbpA transcription (Fig. 2C). The effects of these deletions upon cbpM transcription followed the same patterns seen for cbpA (data not shown). In a control experiment, the lrp deletion had no effect on another σS-dependent gene, dsrB (data not shown). This demonstrates that the effect of Lrp upon cbpAM is specific for cbpAM and depends upon σS, which is the main regulator of cbpAM transcription.

Since transcriptional regulation by Lrp is complex and can be either leucine dependent or independent (7), the role of leucine in the Lrp-dependent regulation of cbpAM was probed. We repeated the above-mentioned experiments with the addition of 0.4 mM leucine to the growth medium. In wild-type cells, addition of leucine resulted in an ∼40% decrease in cbpA transcription compared to transcription in the absence of leucine (Fig. 2C). Addition of leucine had no effect on cbpA transcription in either ΔrpoS or Δlrp cells (Fig. 2C). Furthermore, the decrease in cbpA transcription caused by adding leucine to wild-type cultures was almost identical to the decrease in transcription seen in Δlrp cells grown in the absence of leucine (Fig. 2C). This demonstrates that Lrp activates the cbpAM promoter only in the absence of leucine. In a control experiment, addition of leucine had no effect on dsrB transcription (data not shown). Taken together, these data show that σS and Lrp are both necessary for the full activation of the cbpAM promoter during stationary phase and that Lrp activates cbpA transcription only in cells starved for leucine.

After determining that Lrp is a transcriptional activator of cbpA in stationary phase at 30°C, we attempted to replicate a previous study's finding that Lrp represses cbpA transcription in exponential phase (26). At 37°C, the temperature used in the prior study, neither leucine nor Lrp had a significant effect on cbpA transcription (Fig. 2D). At 30°C, addition of leucine or deletion of lrp each reduced cbpA transcription by ∼20%, and the effects were additive (Fig. 2D). However, the growth temperature did affect cbpA transcription; in the absence of leucine, the wild-type cells grown at 37°C transcribed ∼40% less cbpA than cells grown at 30°C (Fig. 2D).

We next examined the effects of Lrp and leucine on stationary-phase cells grown at 30 and 37°C. In contrast to our finding that at 30°C addition of leucine or deletion of lrp reduced cbpA transcription, at 37°C, either adding leucine or deleting lrp resulted in a small but reproducible increase in cbpA transcription (Fig. 2E). The transcriptional effects were additive when the conditions were combined (Fig. 2E). Interestingly, wild-type transcription of cbpA in the absence of leucine was ∼50% lower at 37°C than at 30°C (Fig. 2E). Together, these data demonstrate that Lrp can activate or repress cbpA depending upon the conditions. Also, increased temperatures result in decreased transcription of cbpA in both exponential and stationary phases.

Translation and accumulation of CbpA and CbpM.

Since there was no apparent difference between the transcriptional patterns of cbpA and cbpM, we wanted to determine if there were effects at the level of translation that might lead to higher levels of CbpA than CbpM, thereby allowing CbpA to function at its full capacity. Translational fusions of lacZ to cbpA and cbpM were constructed in a manner similar to that for the transcriptional constructs and used to determine that the translational patterns of cbpA and cbpM were similar to the transcriptional patterns (Fig. 3A). For both genes, translation was very low during exponential growth, increased sharply during the transition to stationary phase, and reached a plateau as the cells entered stationary phase. Under the conditions used, the ratio of β-galactosidase activity produced from the CbpM-LacZ relative to the CbpA-LacZ fusion remained stable at 1.5 ± 0.3, very similar to the ratio seen with the transcriptional fusions.

FIG. 3.

FIG. 3.

Translation and accumulation of CbpA and CbpM. (A) Translational analysis of CbpA and CbpM utilizing Lac fusions. E. coli strains bearing translational fusions were grown in LB at 30°C, and the β-galactosidase activities of strains producing CbpA2-LacZ (open circles) or CbpM2-LacZ (open squares) was determined in cultures throughout the growth cycle. Each time point represents the average ± standard deviation (SD) of three independent assays. Diamonds, OD595 measurements. (B) Quantification of CbpA monomers per cell. Wild-type E. coli was grown in LB at 30°C for 24 h and analyzed for CbpA content. Varying culture volumes and known amounts of purified protein were separated by SDS-PAGE, detected by Western blotting with specific antisera, and analyzed by densitometry. The resulting data were used to determine the amount of CbpA (ng) per μl of culture. CFU were determined by plating the cells on LB. A representative blot and analysis are shown. The numbers of CbpA monomers per cell were calculated, and the average ± SD of at least three assays is shown in the AVG column. (C) Experiments to quantify CbpM were performed and analyzed as for panel B.

After determining that cbpA and cbpM transcription and translation followed similar patterns, we wanted to know if the proteins accumulated in the cell to similar levels. Using quantitative Western blot analysis, the levels of CbpA and CbpM in stationary-phase cells were directly determined. We found that cells from 24-h cultures had 2,500 ± 300 CbpA monomers per cell and 2,100 ± 300 CbpM monomers per cell (Fig. 3B and C). This indicates that after being transcribed and translated, CbpA and CbpM remain in the cell at similar levels, suggesting that neither is preferentially removed from the cytoplasm under the conditions tested. Based on the quantitative Western data and the LacZ translational-fusion data, we calculate that approximately 200 monomers each of CbpA and CbpM were present in late-exponential-phase cells prior to the transition to stationary phase.

Taken together, these data demonstrate that regulation of cbpA and cbpM occurs at the level of transcription and that CbpA and CbpM accumulate rapidly within the cells as growth ceases and then remain at relatively constant levels. There is no evidence for asynchrony in either transcription or translation of cbpA and cbpM that would lead to different protein levels of CbpA and CbpM. Indeed, our data demonstrate that CbpA and CbpM exist in approximately equal amounts in stationary-phase cells.

CbpM is unstable in the absence of CbpA.

In wild-type cells, CbpA and CbpM levels remained constant after the termination of protein synthesis by chloramphenicol treatment (Fig. 4A), consistent with the observation that they remained constant throughout stationary phase despite low levels of transcription (Fig. 2A and data not shown). We wanted to determine if they were stable under other conditions, since differential degradation would provide a possible rationale for the coexpression of CbpA and CbpM. Single-deletion strains were used to examine the level of CbpA in the absence of CbpM and the level of CbpM in the absence of CbpA. Wild-type amounts of CbpA were present in ΔcbpM cells at both early (OD595, 2.5; 6 h of growth) and late (OD595, 5.0; 24 h of growth) stationary phase (Fig. 4B). Interestingly, CbpM was unstable in the ΔcbpA strain (Fig. 4C). When CbpA was supplied via a multicopy plasmid, CbpM was stabilized in late stationary phase (Fig. 4C). Induction with low levels of arabinose that produced CbpA in quantities similar to wild-type levels was sufficient to prevent almost all CbpM degradation (data not shown). Taken together, these results show that CbpA is stable in the absence of CbpM and is required to stabilize CbpM.

FIG. 4.

FIG. 4.

Stability of CbpA and CbpM. (A) In vivo stability determined by chloramphenicol chase. Cultures were grown in LB at 30°C for 24 h, and then chloramphenicol (100 μg/ml) was added to halt protein synthesis. Samples were precipitated with trichloroacetic acid (TCA) at the indicated time points, and the CbpA (circles) and CbpM (squares) levels were determined by Western blotting, followed by densitometry. The values are the average ± standard deviation (SD) of at least three independent experiments. (B) Stability of CbpA in the presence and absence of CbpM in vivo. Wild-type (WT) or ΔcbpM cells were grown to early (6 h) or late (24 h) stationary phase in LB at 30°C, TCA precipitated, separated by SDS-PAGE, and analyzed by Western blotting for CbpA levels. (C) Stability of CbpM in the presence and absence of CbpA in vivo. Wild-type, ΔcbpA, and ΔcbpA pcbpA+ cells were grown to early (6 h) or late (24 h) stationary phase, separated by SDS-PAGE, and analyzed by Western blotting for CbpM levels. The strain bearing the pcbpA+ plasmid was supplemented with 50 μg/ml ampicillin and induced with 0.02% arabinose. (D) In vivo roles of Lon and ClpP in the degradation of CbpM. E. coli strains were grown to late stationary phase (24 h) in LB at 30°C, and the CbpM content was determined by Western blotting with CbpM antiserum. The relevant genotypes are indicated below lanes 1 to 4. (E) In vitro degradation of CbpM by ClpAP. Reaction mixtures (50 μl) containing CbpM (352 pmol) were incubated with (open squares) or without (filled squares) 181 pmol CbpA dimers in the presence of 17 pmol ClpP tetradecamers, 10.3 pmol ClpA hexamers, 5 mM ATP, and 20 mM MgCl2 in buffer A (20 mM Tris-HCl, pH 7.5, 100 mM KCl, 0.1 mM EDTA, 10% glycerol, 5 mM dithiothreitol [DTT], 0.005% Triton X-100) at 25°C. Reaction mixtures containing 13 pmol ClpX hexamers instead of ClpA were carried out under identical conditions (filled triangles). Samples were precipitated with TCA at the indicated times, separated by SDS-PAGE, stained with Coomassie blue, and analyzed by densitometry using ImageJ software. The values are the averages ± SD of at least three independent experiments. (F) In vitro degradation of CbpM by Lon. Reaction mixtures (20 μl) containing CbpM (317 pmol) were incubated with (open squares) or without (closed squares) 181 pmol CbpA dimers in the presence of 3.8 pmol Lon hexamers, 4 mM ATP, 10 mM MgCl2, 1 mM DTT, and an ATP-regenerating system (10 mM phosphocreatine and 0.2 mU creatine phosphokinase) in 50 mM Tris-HCl, pH 8.0. Samples were precipitated with TCA at the indicated times, and CbpM was quantified as in panel E. The values are the average ± SD of three independent experiments.

After observing that CbpM was unstable in the absence of CbpA, we wanted to determine which proteases were responsible for the degradation of CbpM. Lon, one of the major proteases in E. coli, and ClpP, the peptidase component of the ClpAP and ClpXP proteases, were tested to determine if they played a role. Deletions of the lon and clpP genes were transduced into a ΔcbpA background, resulting in ΔcbpA Δlon and ΔcbpA ΔclpP double-deletion strains. Compared to cells lacking only CbpA, cells lacking both CbpA and Lon contained significant amounts of CbpM in late stationary phase, although ∼5-fold less than the wild type (Fig. 4D, lanes 1 to 3). A lesser, but significant, amount of CbpM remained stable in the ΔcbpA ΔclpP strain (Fig. 4D, lane 4). These results suggest that CbpM is likely recognized and acted upon by a number of proteases when it is not protected by CbpA.

To confirm the in vivo results, the degradation of CbpM was examined in vitro. We found that ClpAP degraded CbpM at a rate of ∼1.9 pmol/min (Fig. 4E). The addition of CbpA to the reaction mixtures reduced the degradation rate of CbpM ∼5-fold, to ∼0.4 pmol/min (Fig. 4E). In a control experiment, CbpA did not significantly inhibit ClpAP degradation of another ClpAP substrate, green fluorescent protein-SsrA (data not shown). CbpM was not detectably degraded by ClpXP (Fig. 4E). Next, we tested Lon and found that it degraded CbpM at a rate of 1.6 pmol/min (Fig. 4F). Addition of CbpA to the reaction mixtures prevented CbpM degradation by Lon. These results indicate that the protection of CbpM from proteolysis by CbpA was likely due to a specific interaction between CbpA and CbpM. Taken together with the in vivo data, this suggests that free CbpM is a substrate for proteolysis but that CbpM complexed to CbpA is not.

DISCUSSION

To study the regulation of CbpA activity by CbpM, we examined several phenotypes of a ΔdnaJ ΔcbpAM strain coexpressing CbpA and CbpM from a plasmid with the expectation of finding conditions under which CbpA might be active in the presence of CbpM. Cooverexpression of CbpA and CbpM restored viability at low temperatures but was not sufficient to support growth at high temperatures. Additionally, cooverexpression of CbpA and CbpM partially relieved the cell division defect in ΔdnaJ ΔcbpAM cells and supported limited λ replication in ΔdnaJ cells, suggesting that cooverexpression of cbpM with cbpA results in incomplete inhibition of the CbpA activity. Taken together, these data imply that CbpM functions more to modulate CbpA activity than to shut it down completely.

Three of the E. coli DnaJ homologs, DnaJ, CbpA, and DjlA, have significant functional overlap, although it is unclear why this redundancy exists (11-13, 27). Conceivably, each homolog may recognize its own specific set of substrates that are preferentially targeted for remodeling by DnaK. If this is the case, then CbpM inhibition of CbpA may prevent DnaK from interacting efficiently with CbpA's set of substrates under certain growth or stress conditions. In this way, CbpA may act as an adaptor for the DnaK system, facilitating delivery of particular high-affinity substrates at specific times. CbpM may act as an antiadaptor, protecting those same substrates from DnaK action at other times. In this scenario, CbpM holds CbpA in a state of limited activity until a cellular or environmental signal leads to dissociation. When the CbpA-CbpM complex separates, fully active CbpA is released and CbpM could be removed by selective degradation by cellular proteases. It is tempting to speculate that other DnaJ homologs have their own inhibitory partners to further adjust the DnaK/Hsp70 chaperone system, although none have been isolated yet.

Recently, another adaptor-antiadaptor pair was described in E. coli. This system is comprised of RssB and IraP, and it controls the turnover of σS (5). RssB acts as an adaptor that targets σS for degradation by ClpXP. In response to phosphate starvation, IraP binds to RssB and prevents it from delivering σS to the protease, thereby stabilizing σS. Such systems add yet another layer to the levels of regulation available to control cellular processes.

Our examination of expression of the cbpAM operon showed that CbpA and CbpM, like other stationary-phase proteins, were mainly expressed at the end of the exponential phase as cells ceased active growth. This is in agreement with a previous transcriptional study of cbpA (29) but contrasts with a different study, which found that CbpA accumulated only in very late stationary phase (3). The reason for the disparity is unclear, although strain and technique variations likely are involved.

We determined that CbpA and CbpM accumulation was controlled at the level of transcription and that the genes were transcribed from the same promoter. Previously, transcription of cbpA was shown to be driven by σS (29). Here, we demonstrated that the cbpAM promoter was dependent upon both the σS and Lrp global regulators for full activity. σS activated cbpAM transcription independently of Lrp, but Lrp required σS to activate cbpAM transcription, suggesting that the Lrp effect is indirect and occurs upstream of σS. Activation of the cbpAM promoter by Lrp was also dependent on leucine, a gauge of amino acid availability (7). Additionally, transcription of the cbpAM operon was dependent upon the growth temperature, with more transcription at 30°C than at 37°C. This was likely due to the increased stability of σS at lower temperatures (24).

Our data showed that CbpA accumulated ∼2,500 monomers per cell and CbpM accumulated ∼2,100 monomers per cell in late stationary phase. The number of molecules per cell differed significantly from a previously published estimate of 15,000 CbpA monomers per cell (1). The strains and techniques used to arrive at the different numbers were not identical, possibly explaining the discrepancy.

Although CbpA and CbpM were both stable in strains expressing both proteins under the conditions tested, CbpM was degraded in the absence of CbpA by Lon, either ClpAP or ClpXP or both, and possibly other proteases. This lends support to our model, which proposes that the limited posttranslational control CbpM exerts over CbpA is alleviated by selective degradation of the inhibitor. Studies are ongoing to determine what signal leads to the dissociation of the CbpA-CbpM complex and which substrates are specifically recognized by CbpA.

Supplementary Material

[Supplemental material]

Acknowledgments

We thank Joel Hoskins, Susan Gottesman, and Nadim Majdalani for helpful discussions and technical assistance. We thank Michael Maurizi for the gift of Lon. We thank Jodi Camberg, Shannon Doyle, Joel Hoskins, and Danielle Johnston for critical reading of the manuscript.

This research was supported by the Intramural Research Program of the NIH National Cancer Institute Center for Cancer Research.

Footnotes

Published ahead of print on 23 May 2008.

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

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