INTRODUCTION

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The expression of heat shock genes in Escherichia coli is positively controlled at the transcriptional level by the product of the rpoH gene, the ³² subunit of RNA polymerase (RNAP) (5, 8, 30). Stress treatment of the cells, such as a sudden temperature upshift, induces transient heat shock gene expression until the cells have adapted to the applied stress. This heat shock response is mediated by increases in the translation of rpoH, stabilization of ³², and activation of ³² by sequestration of the DnaK chaperone and its DnaJ cochaperone from a complex with ³².

Under steady-state growth conditions, ³² is an extremely unstable protein with half-lives at 30 and 42°C of less than 1 and 4 min, respectively (17, 23, 26). Heat shock treatment of the cells by transfer from 30 to 42°C increases the half-life of ³² transiently to approximately 10 min (23). Degradation is mediated mainly by FtsH (HflB), an ATP-dependent metalloprotease associated with the inner membrane (10, 25, 27, 28). The interaction of ³² with the RNAP core enzyme prevents degradation, indicating that FtsH and RNAP compete for binding to ³² (28). The DnaK chaperone system plays an active but mechanistically unclear role in ³² degradation, since ³² is stabilized in dnaK and dnaJ mutant backgrounds (22, 24, 25, 28).

A conceptually interesting question with respect to ³² degradation is how this biologically active, and hence seemingly folded, protein can be subject to such efficient proteolysis. Either ³² carries specific degradation signals within its polypeptide, or it is thermodynamically unstable despite its biological activity. Earlier work showed that the in vivo half-life of fusions between N-terminal fragments of ³² and -galactosidase increased when a stretch of 23 residues (R122 to Q144), located between conserved regions 2 and 3 of ³² and termed region C, is deleted or replaced by another reading frame (17). However, subsequent analysis of ³² mutants altered in region C showed that this region is not essential for degradation but instead plays a role in ³² binding to RNAP (1). Another study identified a role for the C terminus of ³² in degradation by FtsH (3). In this previous study, rpoH genes with truncated 3' ends were cloned into an expression vector such that fusion proteins between C-terminally truncated ³² proteins (by 20 residues) and six amino acids from the vector sequence were generated. These fusion proteins were more stable than wild-type ³² in E. coli cells and in vitro. However, this approach did not exclude the possibility that the vector-encoded extra residues at the C terminus artificially stabilize the fusion proteins.

In this study we investigated further the role of the C terminus of ³² in the degradation process. Our analysis of C-terminally truncated ³² proteins shows that the authentic C-terminal residues are not essential for degradation.

	MATERIALS AND METHODS

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Strains, plasmids, and media. Cells of strains C600 (thr-1 leuB6 thi-1 lacY supE44 rfbD1 fhuA21) and BB2019 [GW1000 recA441 sulA11 (argF-lac)U169 supC(Ts) rpoH165(Am) pDMI,1] (6) were grown aerobically at 30 or 42°C in Luria broth (LB) or in M9 minimal medium supplemented with glucose (0.2%; M9-Glu), thiamine (20 µg/ml), and appropriate amino acids (50 µg/ml). Growth media were further supplemented with isopropyl--D-thiogalactopyranoside (IPTG; 1 mM), kanamycin (20 µg/ml), and ampicillin (50 µg/ml) when required. The wild-type rpoH gene cloned into plasmid pUHE21-2fd12 (7) was used as template for construction of 3'-truncated rpoH alleles. These alleles (rpoH-5aa, rpoH-11aa, rpoH-15aa, and rpoH-21aa) were generated by PCR using appropriate oligonucleotides with HindIII and PstI restriction sites for cloning of the coding sequences into pUHE21-2fd12. For production of hexahistidine-tagged ³² variants, wild-type and mutant alleles of rpoH were subcloned by inserting the HindIII-PstI fragments into pUHE212-1 (to encode N-His₆-³²). For immunoblotting and pulse-chase experiments, the rpoH alleles encoding wild-type ³², N-His₆-³², C-His₆-³² (7), and C-terminally truncated ³² proteins were subcloned by inserting the respective EcoRI-HindIII fragments into pFN476 (24). Plasmid pDMI,1 carrying lacI^q and encoding kanamycin resistance (14) was used to provide cells with a Lac repressor.

Overproduction of ³² proteins and ex vivo degradation assay. C600 cells carrying rpoH expression plasmids and pDMI,1 were grown at 30°C in 20 ml of LB containing ampicillin and kanamycin. IPTG (1 mM final concentration) was added to exponential-phase cultures (optical density at 600 nm [OD₆₀₀], 0.5 to 0.7) for 1 h, followed by harvesting and washing of the cells in buffer A (50 mM Tris-acetate [pH 8.0], 5 mM magnesium acetate, 2 mM -mercaptoethanol, 50 mM KCl, 5% glycerol). The cell pellet was dissolved (OD₆₀₀, 50) in buffer A containing 1 mM phenylmethylsulfonyl fluoride and sonicated. The lysate was transferred to Eppendorf tubes and centrifuged at 2,500 × g for 2 min, and the supernatant was subjected to ultracentrifugation at 100,000 × g for 2 h. The protein content of the soluble cytoplasmic fraction was quantified by Bradford assay using bovine serum albumin as a standard. A 5-µg portion of the cytoplasmic fraction was mixed with FtsH reaction buffer (50 mM Tris-acetate [pH 8.0], 5 mM magnesium acetate, 2 mM -mercaptoethanol, 50 mM KCl) to reach a final volume of 15 µl. Degradation assays were started by adding 1 µl of 100 mM ATP and 4 µl of 10 µM FtsH activated by the addition of Zn²⁺ (27) or buffer without FtsH as a control and incubated at 42°C for 1 h.

In vitro degradation of ³². Wild-type and C-terminally truncated N-His₆-³² proteins were purified and radiolabeled with N-succinimidyl[2,3-³H]propionate (Amersham) as previously described (7, 28). These proteins (1 µM each) were incubated with purified FtsH (2 µM) and tested for degradation as described previously (1, 28). After precipitation with trichloroacetic acid (TCA; 10%) followed by centrifugation (15,000 rpm, 3 min), the radioactive peptides generated by proteolysis were quantified in the supernatant by liquid scintillation counting. For assaying degradation of peptides, the final volume of the reaction was 60 µl. The ³²-derived peptides Q132-Q151-C (QRKLFFNLRKTKQRLGWFNQC) and A264-A284-C (AERVRQLEKNAMKKLRAAIEAC) (50 µM each) were mixed and incubated with FtsH (1). At various time points, aliquots of 18 µl were mixed with 92 µl of 0.5% trifluoroacetic acid to stop the reaction. Products were analyzed by reverse-phase chromatography using a 5-to-80% acetonitrile gradient in 0.1% trifluoroacetic acid.

Analysis of protein interactions. DnaK and RNAP core enzyme were purified as described previously (4, 15). Association of N-His₆-³² or C-terminally truncated N-His₆-³² with DnaK, DnaJ, and RNAP core was determined by gel filtration using a Superdex 200 column essentially as described previously (1, 7). To determine association of ³² with RNAP core, N-His₆-³² (1 µM) was incubated with RNAP core (1.5 µM) for 10 min at 30°C in transcription buffer (20 µl, final volume). To determine association of ³² with DnaK, DnaK (5 µM) was incubated for 2 h at 30°C in transcription buffer to disfavor oligomerization, mixed with N-His₆-³² (1 µM) in a final volume of 20 µl, and further incubated for 30 min at 30°C. These mixtures were placed on ice, adjusted to 100 µl by addition of transcription buffer, and loaded on a Superdex 200 column at 4°C. Labeled N-His₆-³² was detected in the elution fractions by liquid scintillation counting.

In vivo stability of ³². For determination of ³² stability in vivo, C600 cells carrying rpoH expression plasmids were grown at 30°C in M9-Glu until mid-log phase. One milliliter of culture was labeled for 1 min with 70 µCi of [³⁵S]methionine followed by chase with unlabeled methionine (200 µg/ml, final concentration). After 30 s (time for synthesis of ³²), aliquots of 200 µl were collected at various times, mixed with TCA (10%, vol/vol), and incubated for 15 min on ice. After centrifugation for 15 min at 14,000 rpm, the pellets were washed with acetone and resuspended in 50 mM Tris-HCl (pH 8.0), 1% sodium dodecyl sulfate (SDS), and 1 mM EDTA. Samples were subjected to immunoprecipitation using ³²-specific rabbit antiserum as described previously (25, 29).

SDS-PAGE, immunoblotting, and quantifications. Polyacrylamide gel electrophoresis (PAGE) was carried out as described by Laemmli (13) using SDS-12% polyacrylamide gels and staining with Coomassie brilliant blue. Immunoblotting was carried out according to standard procedures, using rabbit antisera specific for the relevant proteins as primary antibodies, and blots were developed with a Vistra ECF fluorescence Western blotting kit (Amersham) or 5-bromo-4-chloro-3-indolylphosphate-nitroblue tetrazolium color detection using alkaline phosphatase-conjugated anti-rabbit immunoglobulin G as the secondary antibody (Vector Laboratories, Inc.). Stained gels and developed immunoblots were scanned using a fluoroimager (FLA-2000) and quantified using MacBAS software (Fuji Film Co.).

	RESULTS

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Construction of truncated ³² mutant proteins. To investigate whether C-terminal residues of ³² constitute a destabilizing element, we generated ³² mutant proteins with truncations of their C termini by 5, 11, 15, and 21 residues (³²-5aa, ³²-11aa, ³²-15aa, and ³²-21aa) (Fig. 1). The truncations were chosen such that the new termini are at two positions within (³²-11aa and ³²-15aa), N-terminal to (³²-21aa), or C-terminal to (³²-5aa) an -helix predicted by a secondary-structure-predicting algorithm (Fig. 1). PCR fragments of the appropriately truncated rpoH genes were cloned into the expression vector pUHE21-2fd12 (7) such that a stop codon immediately follows the last codon, thus ensuring that no vector-encoded amino acids are fused to the ³² mutant proteins. The C termini of the truncated ³² mutant proteins do not resemble the consensus sequence of destabilizing C termini, as described by Sauer and coworkers (21), are not particularly hydrophobic, and do not constitute predicted DnaK binding sites (19).

View larger version (28K): [in this window] [in a new window]   FIG. 1.   Mutational alteration of 32. The locations of conserved regions 1 to 4 and details of the C-terminal region and its predicted secondary structure (using the PhD program) are shown. -helices are shown as cylinders. The end points of the C-terminal truncations of the 32 mutant proteins are indicated.

In vivo activity of truncated ³² mutant proteins. To determine the in vivo activity of the truncated ³² mutant proteins, we first tested the ability of pUHE21-2fd12-borne rpoH mutant alleles to complement the temperature-sensitive growth of rpoH165(Am) mutant cells on LB agar plates. The rpoH165(Am) mutant cells carry a temperature-sensitive amber suppressor mutation which is active and supports growth at 30°C. Only the rpoH-5aa mutant allele allowed complementation of growth at 42°C to an extent similar to that allowed by wild-type rpoH (data not shown). Even in the absence of IPTG, a condition in which read-through expression of the rpoH mutant alleles produced ³² levels only approximately fivefold above wild-type levels, ³²-5aa complemented the growth defects of rpoH(Am) mutants at 42°C (data not shown).

View larger version (99K): [in this window] [in a new window]   FIG. 2.   In vivo activity of C-terminally truncated 32 proteins. Cells of strain BB2019 [rpoH165(Am)] were transformed either with pUHE212-1 expressing wild-type or 3'-truncated rpoH alleles (producing N-His6-32 proteins) or with pUHE212-1 control vector. Cultures of these cells were induced with IPTG, followed by SDS-PAGE of equal amounts of total protein and staining of the gels with Coomassie brilliant blue (top) or immunoblotting using DnaK- and DnaJ-specific sera (bottom). Lanes: 1, vector control; 2, N-His6-32; 3, N-His6-32-5aa; 4, N-His6-32-11aa; 5, N-His6-32-15aa; 6, N-His6-32-21aa.

In vivo stability of truncated ³² mutant proteins. We determined the in vivo half-lives of two of the C-terminally truncated ³² mutant proteins (³²-11aa and ³²-15aa) by pulse-chase experiments followed by immunoprecipitation of ³². Such experiments were complicated by our finding that most of the C-terminally truncated ³² mutant proteins lack activity in vivo. Consequently, the synthesis of heat shock proteins is lower in cells producing the inactive mutant proteins from plasmids than in cells producing wild-type ³² from plasmids (Fig. 2). These heat shock proteins include DnaK and DnaJ, which are limiting for degradation of ³² (24, 25, 28). In the cells expressing the inactive ³² mutant proteins, ³² should be less subject to chaperone-mediated degradation. When the ³² proteins are overexpressed, the efficiency by which ³² mutant proteins are degraded in vivo will therefore be influenced by their biological activity. This complication led us to perform the ³² half-life determinations in cells of strain C600 expressing low levels of plasmid-encoded ³² proteins. Cells were transformed with plasmids expressing wild-type or mutant rpoH alleles under transcriptional control of a promoter specific for T7 polymerase. Since C600 cells lack T7 polymerase, only very little read-through from the T7 promoter occurs, just enough to produce approximately wild-type levels of ³² (Fig. 3). This fact allowed us to perform the half-life determinations of the ³² proteins under almost physiological conditions.

View larger version (24K): [in this window] [in a new window]   FIG. 3.   Cellular levels of plasmid-encoded 32 mutant proteins. Cells of strain C600 transformed with plasmid pFN476 expressing the appropriate rpoH allele were grown at 30°C in LB medium. Aliquots of exponential-phase cultures (OD600, 0.5 to 0.7) were collected, and equal amounts of total protein were analyzed by SDS-PAGE. Levels of 32 were determined by immunoblotting.

View larger version (16K): [in this window] [in a new window]   FIG. 4.   In vivo stability of 32 mutant proteins at 30°C. Cells of strain C600 which harbor pFN476 expressing rpoH-11aa, rpoH-15aa, or wild-type rpoH were grown at 30°C, pulse-labeled with [35S]methionine, and chased with unlabeled methionine. Aliquots were taken at the indicated times, followed by immunoprecipitation of 32 and quantification of the precipitated proteins relative to the largest value. Closed circles, wild-type 32; closed triangles, 32-11aa; open triangles, 32-15aa.

In vitro degradation of truncated ³² mutant proteins by FtsH. We investigated the susceptibility of N-His₆-tagged wild-type and C-terminally truncated ³² mutant proteins to degradation by FtsH in vitro. These degradation assays were performed only at 42°C, since FtsH is known to be poorly active at 30°C when tested in vitro in the presence of detergent (12). We first determined whether exogenously added FtsH is capable of degrading the ³² proteins present in extracts of C600 cells after IPTG-induced overproduction. The rationale for this experiment was that for these ex vivo degradation assays we could use authentic ³² mutant proteins without histidine tags, ruling out any contribution by the tag. All truncated and wild-type ³² proteins were degraded by FtsH in presence of ATP to similar extents (Fig. 5A).

View larger version (48K): [in this window] [in a new window]   FIG. 5.   (A) FtsH-dependent degradation of overproduced 32 in cytoplasmic fractions. Cells of strain C600 which harbor plasmid pUHE21-2fd12 expressing wild-type or truncated rpoH alleles mutant 32 or the control plasmid were grown at 30°C with or without (lanes 1 and 7) IPTG. The extracts of cells grown without IPTG served as a control to ensure that the endogenous chaperones, whose cellular concentrations increase upon IPTG-induced production of plasmid-encoded wild-type 32, do not interfere with the degradation of 32 in the extracts. Cytosolic extracts were prepared and assayed for degradation by exogenously added FtsH. Lane M, molecular weight marker. All other lanes show extracts of cells producing plasmid-encoded wild-type 32 supplemented with 2 µM purified wild-type 32 (since wild-type 32 could not be overproduced to large amounts) (lanes 1, 2, 7, and 8) or plasmid-encoded mutants 32-5aa (lanes 3 and 9), 32-11aa (lanes 4 and 10), 32-15aa (lane 5 and 11), and 32-21aa (lanes 6 and 12). The extracts were incubated with (+FtsH) or without (FtsH) added protease. (B) In vitro degradation of 32 mutant proteins by FtsH. 3H-labeled N-His6-32 proteins were incubated with FtsH in the presence of 5 mM ATP, followed by TCA precipitation at the indicated times. The curves represent the percentage of radioactivity in the supernatants which contain the proteolytic fragments. Closed circles, N-His-32; open circles, N-His6-32-5aa; closed triangles, N-His6-32-11aa; open squares, N-His6-32-21aa.

C-terminal residues of ³² do not constitute a substrate for FtsH. As an independent experimental approach to investigate the possibility that the C-terminal residues of ³² constitute a destabilizing element for FtsH-dependent degradation, we tested whether the C-terminal sequences themselves provide a substrate motif for FtsH. We designed a peptide comprising the C-terminal 21 residues of ³² and an additional cysteine at the C-terminal end (AERVRQLEKNAMKKLRAAIEAC) and tested whether it is a substrate for FtsH. The length of this 22-mer peptide was well above the minimal length of approximately 15 residues required for FtsH to degrade peptides (T. Tomoyasu and B. Bukau, unpublished results). As a positive control, we included a peptide of the same length and carrying a cysteine at the C terminus, derived from region C of ³² (QRKLFFNLRKTKQRLGWFNQC). This peptide has been shown previously to be a good substrate peptide for FtsH (1).

View larger version (21K): [in this window] [in a new window]   FIG. 6.   FtsH-mediated degradation of peptides derived from 32. Peptides from region C (32-Q132-Q151-C) and the C terminus (32-A264-A284-C) of 32 were mixed and incubated with FtsH. Degradation of the peptides at various times after mixing is shown as high-pressure liquid chromatography-reverse-phase chromatography data.

DnaK and DnaJ bind to truncated ³² mutant proteins in vitro. We tested whether DnaK binding to ³² is impaired by the C-terminal truncations. ³H-labeled wild-type ³² and mutant N-His₆-³² were incubated with DnaK followed by gel filtration to separate DnaK-³² complexes from free ³² (Fig. 7). Under the conditions used, approximately 75% of wild-type [³H]³² was recovered in complex with DnaK (eluting in fractions 12 to 17). The ³H-labeled C-terminally truncated N-His₆-³² mutant proteins (³²-5aa, ³²-11aa, and ³²-21aa) showed similar efficiencies of complex formation. Earlier work established that the interaction of DnaK with ³² does not occur through the histidine tag (6). Furthermore, no defects in chaperone binding were observed when the N-His₆-³² mutant proteins were incubated with DnaK together with DnaJ in the presence of ATP (data not shown). These data indicate that the authentic C terminus of ³² is not essential for interaction with DnaK and DnaJ.

View larger version (23K): [in this window] [in a new window]   FIG. 7.   Binding of 32 mutant proteins to DnaK. 3H-labeled N-His6-32, N-His6-32-5aa, N-His6-32-11aa, and N-His6-32-21aa were incubated with DnaK, followed by gel filtration of the reaction product. Labeled protein was quantified in the elution fractions. Open circles, wild-type 32; closed circles, 32-5aa; open squares, 32-11aa; closed squares, 32-21aa.

RNAP binding to truncated ³² mutant proteins in vitro. We considered the possibility that the truncated C-terminal segments of ³² are involved in the interaction with the RNAP core enzyme. We determined the efficiency of association of ³H-labeled N-His₆-³² with RNAP by gel filtration. The relative amounts of all truncated ³² proteins (³²-5aa, ³²-11aa, ³²-15aa, and ³²-21aa) recovered in association with RNAP (eluting in fractions 8 to 15) were lower than that of wild-type ³², with ³²-5aa showing the highest affinity for RNAP among all truncated proteins (Fig. 8). These results indicate a role for the C terminus of ³² in the binding to RNAP.

View larger version (22K): [in this window] [in a new window]   FIG. 8.   Binding of 32 mutant proteins to RNAP. 3H-labeled N-His6-32 proteins were incubated with RNAP, followed by gel filtration. The amount of labeled protein was quantified. Open circles, wild-type 32; closed circles, 32-5aa; open squares, 32-11aa; closed squares, 32-21aa.

Involvement of N-terminal segments of ³² in degradation by FtsH in vitro. Given our finding that the C terminus of ³² is not an essential degradation signal, we considered that perhaps N-terminal sequences play such a role. However, N-terminal sequences of ³² are not amenable to mutational analysis, since the efficiency of translation of rpoH mRNA drops dramatically upon mutational alteration of the downstream box located at the 5' end of the coding sequence (18).

View larger version (41K): [in this window] [in a new window]   FIG. 9.   Determination of in vivo degradation intermediates of 32. Cells of strain C600 which harbor plasmids expressing IPTG-regulated rpoH alleles encoding N-His-32, C-His-32, or authentic 32 were grown at 30°C in LB medium. Aliquots of exponential-phase cultures (OD600, 0.5 to 0.7) were collected, and equal amounts of total protein were analyzed by SDS-PAGE. The degradation intermediates were determined by immunoblotting using 32-specific antisera.

	DISCUSSION

Top Abstract Introduction Materials and Methods Results Discussion References

The aim of this study was to investigate the role of the C terminus of ³² in the degradation by FtsH. C termini frequently constitute the degradation determinants of naturally unstable proteins, which target them to proteolysis by AAA proteases (9, 21). Such a role has been also proposed for the C terminus of ³² on the basis of an analysis of C-terminal variants of ³² (3).

We show here that a series of C-terminally truncated ³² proteins are as unstable as wild-type ³² in vivo, in cell extracts supplemented with FtsH and ATP, and in a purified ATP-dependent degradation system with FtsH and ³² as the sole protein components. Furthermore, we show that the C-terminal truncations do not compromise the ability of ³² to associate with DnaK in vitro. The C-terminal truncations were chosen such that the local predicted secondary structures are considered and that the novel C termini do not constitute predicted DnaK binding sites (16, 19) and do not resemble the consensus sequence of protease targeting sites (9). Taken together, our results do not provide any evidence for an essential role of C-terminal sequences of ³² in the FtsH-dependent degradation process or in chaperone binding. Our findings agree well with the results of a recent study of truncated ³² proteins and hybrids of stable Bradyrhizobium japonicum and unstable E. coli proteins (2). That study mapped a region of 85 residues, located between residues 36 to 122 of ³², as being responsible for degradation of ³².

We do not know the reason for the discrepancy with a previous study claiming an essential role for the C terminus of ³² in degradation (3). It is possible that the additional six residues encoded by the vector which were fused to the C-terminally truncated ³² proteins in the former study generated fusion proteins which became stabilized for unknown reasons. Our results are, however, in agreement with the findings that (i) class I fusions between N-terminal segments of ³² and -galactosidase exhibit normal shutoff of the heat shock response and ³² instability (17) and (ii) blocking the N terminus, but not the C terminus, of ³² by fusion with green fluorescent protein resulted in stabilization of the fusion protein, indicating the importance of sequences at or near the N terminus of ³² in its degradation by FtsH (T. Tatsuta and T. Ogura, unpublished results). At this point we cannot exclude an involvement of internal sequences of ³² in degradation. Furthermore, our conclusion is supported by our finding that fragments of ³² proteins which were generated by FtsH-dependent degradation in vitro lack N-terminal segments but not C-terminal segments of ³². Although this finding should still be interpreted with caution, it suggests important roles for N-terminal segments of ³² in the degradation process. Further genetic and biochemical dissection is required to identify such a role.

Our analysis provided evidence for a role of the C-terminal sequences of ³² in association with RNAP. Only the mutant protein lacking five C-terminal residues remained partially proficient in binding to RNAP and in in vivo activity, whereas all mutant proteins with longer truncations showed no significant RNAP binding or in vivo activity. The molecular basis for this role of the C terminus of ³² in the association with RNAP is unclear. However, on the basis of several criteria (solubility and wild type-like partial proteolysis pattern), the C-terminally truncated ³² proteins are not perturbed in their overall structure. Thus, either local conformational changes induced by the truncations or the lack of C-terminal residues directly involved in the association process may account for the observed loss in affinity. In this respect it is interesting that many regions within the polypeptide chain of ³², including the regulatory region C and the conserved regions 2.1, 2.2, 3, and 4, have been implicated in the binding of RNAP (1, 3, 11, 20, 31). Structural analysis seems essential to solve this obvious complexity of the interaction between ³² and RNAP.