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Journal of Bacteriology, May 1999, p. 3039-3050, Vol. 181, No. 10
0021-9193/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Identification and Transcriptional Control of the
Genes Encoding the Caulobacter crescentus ClpXP
Protease
Magne
Østerås,
Agathe
Stotz,
Stefanie Schmid
Nuoffer, and
Urs
Jenal*
Division of Molecular Microbiology,
Biozentrum, University of Basel, CH-4056 Basel, Switzerland
Received 19 January 1999/Accepted 2 March 1999
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ABSTRACT |
The region of the Caulobacter crescentus chromosome
harboring the genes for the ClpXP protease was isolated and
characterized. Comparison of the deduced amino acid sequences of the
C. crescentus ClpP and ClpX proteins with those of their
homologues from several gram-positive and gram-negative bacteria
revealed stronger conservation for the ATPase regulatory subunit (ClpX)
than for the peptidase subunit (ClpP). The C. crescentus
clpX gene was shown by complementation analysis to be functional
in Escherichia coli. However, clpX from E. coli was not able to substitute for the essential nature
of the clpX gene in C. crescentus. The
clpP and clpX genes are separated on the
C. crescentus chromosome by an open reading frame pointing in the opposite direction from the clp genes, and
transcription of clpP and clpX was found to be
uncoupled. clpP is transcribed as a monocistronic unit with
a promoter (PP1) located immediately upstream of the 5' end
of the gene and a terminator structure following its 3' end.
PP1 is under heat shock control and is induced upon entry
of the cells into the stationary phase. At least three promoters for
clpX (PX1, PX2, and
PX3) were mapped in the clpP-clpX intergenic
region. In contrast to PP1, the clpX promoters
were found to be downregulated after heat shock but were also subject to growth phase control. In addition, the clpP and
clpX promoters showed different activity patterns during
the cell cycle. Together, these results demonstrate that the genes
coding for the peptidase and the regulatory subunits of the ClpXP
protease are under independent transcriptional control in C. crescentus. Determination of the numbers of ClpP and ClpX
molecules per cell suggested that ClpX is the limiting component
compared with ClpP.
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INTRODUCTION |
In Caulobacter
crescentus, cell differentiation is an integral part of each cell
cycle (18). Asymmetric cell division gives rise to two
distinct progeny, only one of which, the stalked cell, is competent for
a new round of DNA replication and cell division. The other daughter
cell, the swarmer cell, has to go through an obligatory differentiation
step into a stalked cell to regain its replicative ability. Recent
studies have revealed that in this organism, specific proteolytic
events are key control mechanisms for cell differentiation as well as
for proper cell cycle progression (1, 6, 17, 20, 51). For
example, CcrM, an essential DNA methyltransferase, is synthesized only
in C. crescentus late predivisional cells, where it is
required to fully methylate the newly replicated chromosomes (43,
44, 55). To avoid early methylation of the new chromosomes at the
beginning of the S phase, DNA methylation activity is restricted to
late predivisional cells. This restriction is accomplished by rapid
degradation of CcrM, resulting in its presence in cells only when it is
actively synthesized. The instability of CcrM and, thus, part of its
temporal control are dependent on the C. crescentus Lon
protease (51). Several other C. crescentus
proteins have been shown to be degraded in a cell cycle-dependent way;
these include the flagellar anchor protein FliF (17); the
McpA chemoreceptor (1); CtrA, a cell cycle transcriptional
regulator (6); and FtsZ, a tubulin-like GTPase required for
cytokinesis (20). To understand how the proteolytic turnover
of these key regulatory, metabolic, and structural proteins is
controlled by the cell cycle, it is necessary to identify the
corresponding protease(s) and to understand the regulation of its
activity. However, none of the proteases responsible for the turnover
of these proteins has been isolated so far.
Short stretches of amino acids at the carboxyl termini of the CtrA,
FliF, and McpA proteins have been identified as turnover signals and
shown to be strictly required for the cell cycle-dependent degradation
of these proteins (1, 6, 17). Similarly, in Escherichia coli, several proteins are tagged for
degradation by the ATP-dependent ClpXP proteases by a short C-terminal
domain (12, 24, 28). This possible parallel in the mechanism
of substrate recognition led us to test the hypothesis that in C. crescentus, ClpXP might be responsible for the degradation of some
of the proteins mentioned above. Here we report the isolation and
characterization of the C. crescentus chromosomal locus that contains the genes for the peptidase and the ATPase regulatory subunits
of the ClpXP protease. In a parallel study, we showed that both the
clpP and the clpX genes are essential for growth, viability, and cell cycle progression (16). In addition,
ClpXP was shown to be required for cell cycle-dependent degradation of
the CtrA response regulator protein. To better understand the physiological role of the ClpXP protease in Caulobacter, the
promoters and transcripts of the clpP and clpX
genes were mapped and their transcriptional control was investigated.
The analysis of heat shock induction as well as the cell cycle- and
growth phase-dependent activities of the mapped promoters demonstrated
that clpP and clpX are controlled differently at
the transcriptional level. The significance of this finding is
discussed with respect to the stoichiometry of ClpP and ClpX in cells
and the role of the ClpXP protease in Caulobacter cell cycle progression.
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MATERIALS AND METHODS |
Bacterial strains and growth conditions.
The bacterial
strains and plasmids used in this study are listed in Table
1. E. coli DH10B and S17-1
were used as the host strain for molecular cloning experiments and as
the donor strain for conjugation experiments, respectively. E. coli strains were cultured at 37°C in Luria-Bertani (LB) broth
(39) supplemented with ampicillin (100 µg/ml), kanamycin
(30 µg/ml), or tetracycline (10 µg/ml) as necessary. C. crescentus strains were grown on either PYE complex medium
(35) or M2 minimal glucose medium (7) supplemented with nalidixic acid (20 µg/ml), kanamycin (20 µg/ml), or tetracycline (2 µg/ml) as necessary. Synchronizable C. crescentus NA1000 was used as the wild-type strain in all
experiments. Cell cycle synchronization was carried out with this
strain by Ludox density gradient centrifugation as described previously
(8).
DNA manipulations and sequence analysis.
DNA preparation and
manipulation techniques used in this study were as previously described
(2, 39). Transformations of E. coli were done by
electroporation, and plasmids were transferred from E. coli
S17-1 to C. crescentus by conjugation as described previously (7). E. coli clones for sequencing
were obtained by subcloning specific DNA fragments into vector
pBluescript SK(+) (Stratagene, La Jolla, Calif.). The DNA sequence of
the C. crescentus chromosomal clp region was
determined for both strands by the dideoxy chain termination method
(40).
ClpX activity assay with E. coli.
The clpX
genes from E. coli and C. crescentus were
amplified by PCR with specific primers (5'-CGA CTC TAG AGC ATA TGA CAG ATA AAC GCA AAG ATG GCT C-3', 5'-CGC GGA TCC CCT TTT TGG TTA ACT TAT
TGT ATG GG-3', 5'-AAA CAT ATG ACG AAA GCC GCG AGC-3', and 5'-AAA GGA
TCC GCT TCG AAA GCA CGC GCT-3') and cloned as
NdeI-BamHI fragments into pET21b to introduce a
ribosome-binding site upstream of clpX. The resulting
XbaI-BamHI fragments were cloned into the low-copy-number vector pMR20, resulting in plasmids pSSN3 (E. coli) and pSSN6 (C. crescentus) with the
clpX gene under the control of the lacZ promoter.
ClpX activity was determined by the cell killing assay based on the
bacteriophage P1 plasmid addiction module Phd-Doc (
25).
The
stable cellular toxin Doc is inhibited by the antidote protein
Phd,
which is unstable in
E. coli due to its rapid degradation
by
the ClpXP protease (
26). ClpX activity was determined by
monitoring the growth of
E. coli cells containing the
phd and
doc genes on a plasmid with a
temperature-sensitive replicon (pGB2ts::
phd-doc)
after a shift to the nonpermissive temperature. While the progressive
loss of the plasmid stops the new synthesis of Phd and Doc, functional
ClpX will degrade the antidote protein Phd, allowing the toxic
protein
Doc to stop cell growth. In the absence of ClpX, both
Phd and Doc are
stable and growth is not
affected.
For the plasmid addiction assay, combinations of plasmid pGB2ts or
pGB2ts::
phd-doc with plasmid pSSN3 or pSSN6 were
introduced
into strains W3110 and SSN1. Cultures in LB broth
supplemented
with the appropriate antibiotics were grown at 30°C and
successively
diluted so as to maintain logarithmic growth conditions.
The assay
was started by diluting the cultures into antibiotic-free LB
broth
at 40°C and monitoring cell growth by measuring the absorbance
at 600
nm.
Promoter mapping and transcriptional activity assays.
DNA
fragments containing promoters for clpP, clpX,
and cicA were identified by cloning restriction fragments
spanning the entire chromosomal clp region (see Fig. 4) into
vector pRKlac290, generating transcriptional fusions to the
lacZ reporter gene. The resulting plasmids were transferred
into C. crescentus NA1000 wild-type cells, and overnight
cultures of the resulting strains were diluted in fresh PYE medium and
grown to an optical density at 660 nm (OD660) of 0.5 to
0.6. 
-Galactosidase activity was then determined as described
previously (33).
For primer extension assays,
C. crescentus NA1000 was grown
in PYE medium to an OD
660 of 0.5, and total RNA was
isolated as
described previously (
38). Residual DNA was
removed by precipitating
the RNA twice with 3 volumes of sodium acetate
(pH 7, 3 M) at
20°C for 6 h before centrifugation for 15 min.
Specific primers
complementary to the 5' end of the
clpP
gene (clpPPE1, 5'-TCG
ACC ACC ATC GGC ACC AGG TTC AT-3'; clpPPE2,
5'-ATG ATC CGT TCC
TTC AAC AGG CGC GA-3') and the
clpX gene
(clpXPE1, 5'-GGC TTT
CGT CAT GAT CGC TTC TCA CA-3'; clpXPE2, 5'-GCT TGC
GCA CCT CAT
GTT GGC TCT TT-3') were end labeled with
[
-
33P]ATP (370 MBq/ml; Amersham) and T4 polynucleotide
kinase (NEN
BioLabs). Radiolabeled primer (5 × 10
5
dpm) was annealed to 40 µg of
C. crescentus total RNA, and
the
extension reaction was carried out with a SuperScript II kit
(Bethesda
Research Laboratories). A
35S-labeled DNA
sequencing reaction with the desired primer and
pUJ138 DNA as a
template was carried out with a ThermoSequenase
cycle sequencing kit
(Amersham) and served as a standard to identify
the transcriptional
start
site.
The promoter activities of
clpP and
clpX
promoters throughout the cell cycle and following heat shock were
determined by pulse-labeling
of cultures harboring plasmid-encoded
lacZ reporter gene fusions.
Pulse labeling and
immunoprecipitation of
-galactosidase were
carried out as described
previously (
32).
Immunoblotting.
Immunoblotting was performed as described
previously (17). Polyclonal sera against ClpP or ClpX were
diluted 1:10,000 and 1:5,000, respectively, before use. Secondary
antibody (goat anti-rabbit immunoglobulin G [IgG] coupled to
horseradish peroxidase [HRP]) was used at a 1:10,000 dilution.
Immunoblots were developed with a Renaissance kit from DuPont NEN by
following the manufacturer's instructions. The cellular levels of ClpP
and ClpX were estimated by comparison to a dilution range of purified
His-tagged Clp proteins (16). Protein concentrations were
determined by the method of Lowry with a Bio-Rad DC protein assay kit
and bovine serum albumin as the standard. Signals from the immunoblot
analysis were quantitated by use of scanned images and ImageQuant software.
Nucleotide sequence accession number.
The DNA sequence for
clp has been deposited in the EMBL nucleotide sequence
database under accession no. AJ010321.
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RESULTS |
Cloning of the genes for the subunits of the
Caulobacter ClpXP protease.
We isolated the C. crescentus clpP and clpX genes on the basis of their
proximity to the lon gene, coding for the Lon protease (51). In E. coli, the clpP and
clpX genes are located immediately upstream of the
lon gene (Fig. 1)
(11). Assuming a conserved gene order, we isolated and
analyzed the chromosomal region upstream of the C. crescentus
lon gene. The genes coding for the ClpP and ClpX homologues were
identified together with the tig gene, coding for the
homologue of the E. coli trigger factor (Fig. 1). The gene
order tig-clpP-clpX is conserved in 7 of 10 bacteria
analyzed so far, with the exception of Rhodobacter
capsulatus, Helicobacter pylori, and Bacillus
subtilis, in which one of the three genes is located elsewhere on
the chromosome (Fig. 1). The C. crescentus clp locus
contains an additional piece of DNA of about 1 kb between clpP and clpX; this piece is not present in any
of the organisms listed in Fig. 1. This region contains an open reading
frame, cicA (clp intergenic region in
Caulobacter), transcribed in the direction opposite that of
the clp genes. Database searches revealed a weak similarity
between the deduced amino acid sequence of cicA and that of
a family of bacterial proteins with unknown function (data not shown).
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FIG. 1.
Schematic diagram of the clpP-clpX regions of
gram-positive and gram-negative bacteria. The genes coding for the
trigger factor (tig), the ClpP peptidase subunit
(clpP), the ClpX ATPase subunit (clpX), and the
Lon protease (lon) are indicated. In most organisms analyzed
so far, the gene order tig-clpP-clpX is conserved, with the
lon gene located immediately downstream. In C. crescentus, the clp genes are separated by
cicA, a gene of unknown function; in Haemophilus
influenzae, the lon gene is not linked to the
clp genes, which are followed by the secE
(preprotein translocase subunit) and nusG (transcription
antiterminator protein) genes (9); in R. capsulatus, neither tig nor lon is linked to
the clp genes, which are flanked by genes of unknown
function (?) (35a); in H. pylori, the
clpX and lon genes, although linked, are not
found at the same location as the tig and clpP
genes, which are followed by the def (polypeptide
deformylase) gene (45); and in B. subtilis,
clpP is not linked to the tig, clpX,
and lon genes. The lon gene has undergone
duplication in B. subtilis (23); in
Synechocystis and Mycobacterium tuberculosis, no
gene coding for a homolog of the Lon protease is found. However, two
clpP genes are present in M. tuberculosis, and
four clpP genes are found in Synechocystis, one
of them linked to the tig and clpX genes
(19). Two genes are present between the clpP and
clpX genes of M. tuberculosis, and the deduced
sequences of their products are similar to those of a methyltransferase
and a drug efflux protein (4). In Aquifex
aeolicus, the lon gene is not linked to the
tig, clpP, and clpX genes and is found
elsewhere on the chromosome (5).
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It is interesting to note that the ClpX ATPase subunit seems
significantly more conserved in bacteria than the ClpP peptidase
subunit. The
C. crescentus ClpP amino acid sequence showed
between
31.6 and 65.6% similarity and the ClpX amino acid sequence
showed
49.9 to 78.3% similarity with homologues from other bacteria.
For both proteins, the weakest similarity was observed with ClpX
and
ClpP of the spirochete
Borrelia burgdorferi, while the
strongest
match was with ClpX and ClpP of
R. capsulatus,
like
C. crescentus a member of the
-purple group of
gram-negative bacteria. The
similarity of the
Caulobacter
ClpP protein with its counterparts
extends over the entire protein
sequence, and the Ser, His, and
Asp residues of the catalytic triad
(
31,
49) are conserved
(Fig.
2A). Similarly, the ClpX sequences are
conserved throughout
the entire protein length, with the highest
homology around the
ATP-binding boxes (
48) and the
C-terminal signature sequences
(
41) (Fig.
2B). A Zn
finger-like motif, CXXC(X
18)CXXC, is also
conserved in all
ClpX sequences shown in Fig.
2B (
11). The weakest
similarity
among all ClpX amino acid sequences is found in the
tandem C-terminal
PDZ-like (PSD-95, Dlg, and ZO-1 proteins, where
the domain was first
identified) domains that were proposed to
specifically bind target
proteins (Fig.
2B) (
27).
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FIG. 2.
Sequence alignment of ClpP (A) and ClpX (B) from
C. crescentus (Ccr) with homologous sequences
from R. capsulatus (Rca) (35a),
E. coli (Eco) (11, 30), B. subtilis (Bsu) (23), and A. aeolicus (Aae) (5). The amino acid residues
identical to those of ClpP or ClpX from C. crescentus are
indicated by white letters on a black background. A putative Zn
finger-binding site, the ATP-binding (Walker box and ATP-bind.) motifs,
and the proposed PDZ-like domains of ClpX (27) are marked.
The serine, histidine, and aspartate residues involved in ClpP activity
(31, 49) are boxed. Gaps are represented by dashes and were
introduced to maximize the alignment. The alignment was generated with
the Megalignment program of the DNAstar program package (DNAstar,
Madison, Wis.).
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ClpXCc is functional in E. coli.
C.
crescentus ClpX (ClpXCc) showed 66.8% similarity to
E. coli ClpX (ClpXEc). To test if
ClpXCc is a functional homologue of the ClpXEc
chaperone (50), we tried to complement the E. coli
clpX mutant SSN1 with the gene for ClpXCc
(clpXCc). For this purpose, the
clpXCc coding region was fused to the E. coli Plac promoter with efficient translational start
signals (see Materials and Methods), and the fusion was cloned into the
low-copy-number vector pMR20. As a test system for functionality, we
used the cell killing assay based on the bacteriophage P1 plasmid
addiction module Phd-Doc (25) (see Materials and Methods).
The growth of E. coli with the temperature-sensitive
replicon that does not contain the doc and phd
genes (pGB2ts) was unaffected. While wild-type strain W3110 with
pGB2ts::phd-doc stopped growing at a nonpermissive temperature (Fig. 3A), the growth of
clpX mutant strain SSN1 was not affected by the loss of
plasmid pGB2ts::phd-doc (data not shown),
demonstrating that the basis of plasmid stabilization by the Phd-Doc
module is the degradation of Phd by the ClpXP protease (26).
Growth was affected, however, when the clpX mutant strain was complemented with a plasmid-encoded copy of the ClpXEc
gene (clpXEc) (pSSN3). Similarly, expressing
clpXCc from plasmid pSSN6 in a clpX
mutant background resulted in growth inhibition after a temperature
shift (Fig. 3A). Growth inhibition in cells expressing ClpXCc was slightly less severe than that in cells
expressing the homologous ClpXEc protein (Fig. 3A). This
result could be due to reduced activity of the ClpXCc
protein in the heterologous host, as immunoblot analysis with an
anti-ClpX antibody (16) confirmed that ClpXEc
and ClpXCc were present in comparable amounts in the
clpX mutant test strain (Fig. 3B). These results suggest that ClpXCc not only is able to recognize the Phd protein
but also can interact with the proteolytic ClpP component of E. coli. Interestingly, while a plasmid-encoded copy of
clpXCc (pSSN6) was able to support the growth of
a C. crescentus mutant with a disrupted chromosomal
clpX copy, clpXEc (pSSN3) could not
rescue the mutant phenotype (data not shown). This result suggested
that ClpXEc is not able to interact with the C. crescentus ClpP subunits, potential substrates, or accessory
proteins.
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FIG. 3.
clpXCc is able to complement an
E. coli clpX mutant. (A) The activity of ClpX was monitored
by its ability to degrade, together with ClpP, the antidote protein Phd
of the P1 plasmid addiction module Phd-Doc (26). Loss of the
phd gene results in Doc-dependent cell killing if the Phd
protein is degraded by the ClpXP protease. Growth of cultures
containing the phd and doc genes on a plasmid
with a temperature-sensitive replicon
(pGB2ts::phd-doc) was monitored after a shift to
the nonpermissive temperature. Growth is shown as the log
OD600. The time after the temperature increase is indicated
in hours. Cessation of growth 3 to 5 h after the temperature shift
was an indicator of the rapid disappearance of the antidote protein Phd
and thus of ClpX activity. The following plasmids were used: plasmid
pGB2ts is temperature sensitive for replication;
pGB2ts::phd-doc is identical to pGB2ts except that
it contains the plasmid addiction genes phd and
doc; pSSN6 contains clpXCc; and pSSN3
carries clpXEc. Growth of the following E. coli strains was monitored: W3110/pGB2ts/pSSN6 plus
isopropyl--D-thiogalactopyranoside (IPTG) ( ; negative
control); W3110/pGB2ts::phd-doc/pSSN6 plus
IPTG ( ; positive control); SSN1/pGB2ts/pSSN6 plus IPTG ( );
SSN1/pGB2ts::phd-doc/pSSN3 plus IPTG ( );
SSN1/pGB2ts::phd-doc/pSSN6 plus IPTG ( ); and
SSN1/pGB2ts::phd-doc/pSSN6 ( ). (B) Immunoblot
analysis with an anti-ClpX serum and extracts of E. coli
strains expressing ClpXCc and ClpXEc. Equal
amounts of total protein from the following strains were analyzed:
W3110/pMR20 (lane 1); W3110/pSSN3 (lane 2); W3110/pSSN6 (lane 3);
SSN1/pMR20 (lane 4); SSN1/pSSN3 (lane 5); and SSN1/pSSN6 (lane 6). The
band corresponding to ClpX is marked by an arrow.
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The clpP and clpX genes are transcribed
from independent promoters.
In E. coli, the
clpP and clpX genes are transcribed as an operon
from the same promoter, indicating that the syntheses of the regulatory
and proteolytic subunits of the ClpXP protease are coupled
(11). Three GC-rich inverted repeats followed by several
thymidine residues, located downstream of the tig (T1, 5'-GGCGCGGCTCGCGAGGGCCGCGCCTTTTT-3'), clpP (T2,
5'-ACAAAGCCGCCGGCCAGGAGGTCGGCGGCTTTTTT-3'), and
clpX (T3, 5'-CGCCATCATCGGATGGCGCGCTTTT-3') genes,
could act as rho-independent transcriptional terminators, implying that these genes are expressed independently from each other (Fig. 4). Also, the large intergenic region
between clpP and clpX (Fig. 4) suggested that
clpP transcription and clpX transcription are not
coupled in C. crescentus.
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FIG. 4.
Identification of promoter regions in the C. crescentus clp locus. A schematic of the chromosomal
clp region is shown at the top. The tig (trigger
factor), clpP (ClpP peptidase), cicA (unknown
function), clpX (ClpX ATPase), and lon (Lon
protease) genes are indicated by black arrows. The approximate
locations and orientations of the promoters identified
(PP1, PX1, PX2, and
PX3) are marked by short open arrows, and putative
transcriptional terminator structures (T1, T2,
and T3) are indicated as stem-loop outlines. Fragments that
were cloned into the lacZ reporter plasmid pRKlac290 (see
Materials and Methods) are indicated as open bars below the schematic,
with the filled triangles marking the location and orientation of the
lacZ reporter gene. The names of the corresponding
constructs are on the left, and the number on the right indicates the
-galactosidase activity (Miller units) generated by each fusion
construct. All measurements were determined in triplicate, and average
numbers are presented. The following abbreviations are used for
recognition sites of restriction enzymes: RI, EcoRI; P,
PstI; S, SalI; X, XhoI; RV,
EcoRV; A, ApaI; M, MluI; B,
BamHI; and F, FspI.
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To analyze the transcription of the
clp genes and to
localize their promoter regions, we cloned several fragments of the
clp locus into vector pRKlac290, generating transcriptional
fusions
between potential promoter fragments and the
lacZ
reporter gene.
The transcriptional activities of the corresponding
fragments
were determined by introducing the resulting constructs into
C. crescentus NA1000 wild-type cells and measuring
-galactosidase
activities. A
SalI-
XhoI
fragment (pAS2; Fig.
4) containing the
5' end of
clpP and
its upstream region generated 1,809 Miller
units. This fragment most
likely contains the promoter for
clpP (P
P1; see
below). The 3' part of the
tig gene generated only
background
levels of
-galactosidase activity and thus does not carry
a promoter
(pAS1; Fig.
4). However, we cannot exclude the possibility
that
some transcriptional activity originating upstream of
tig reads
through the putative terminator structure T1
located immediately
downstream of the
tig gene (Fig.
4).
Comparison of the Miller
units generated by constructs pAS3 and pAS29
(Fig.
4) suggested
that the majority of the transcripts that originate
at P
P1 are
terminated by the stem-loop structure T2
downstream of the
clpP gene. Thus, P
P1 does not
contribute significantly to the expression
of downstream
genes.
The region between the
clpP and
clpX genes
generated a total transcriptional activity corresponding to 3,443 Miller units
and reading into the
clpX gene (pAS24; Fig.
4).
This total activity
could be assigned to at least three promoter
regions represented
by the constructs pAS64 (P
X3, 690 Miller units), pAS26 (P
X2, 1,422
Miller units), and pAS6
(P
X1, 1,399 Miller units). Since the activities
of these
constructs can be added up to the activity found for
the entire
clp intergenic region (pAS24), we assume that the
transcripts
originating in all three promoter regions extend into the
clpX gene. In addition, construct pAS23 (3,005 Miller units;
Fig.
4)
combined the transcriptional activities of P
X1 and
P
X2, and the
activity generated by construct pAS27 was
equal to the sum of
the transcriptional activities of P
X2
and P
X3 (2,246 Miller units;
Fig.
4). The expression of
clpX in
C. crescentus is therefore
controlled by
at least three different promoter regions located
in the
clpPX intergenic region. It is not clear if and to what
extent these promoters contribute to the expression of the adjacent
lon gene.
clpX is followed by a putative
rho-independent terminator
(T3) that could uncouple the expression of
the
clpX and
lon genes
(Fig.
4).
The
clpPX intergenic region contains several potential open
reading frames, the longest,
cicA, pointing in the direction
opposite
that of
clpP and
clpX (Fig.
4). To
identify a potential promoter
region for
cicA, we created a
lacZ fusion to the
ApaI-
FspI fragment
in the direction opposite that in construct pAS23. While this
piece of
DNA contains P
X1 and P
X2, generating about
3,000 Miller
units reading into
clpX, it also gives rise to
about 2,100 Miller
units reading in the opposite direction (pAS63; Fig.
4). Speculating
that this activity drives the expression of
cicA, we call this
promoter region P
cicA (Fig.
4). In contrast to that generated
by construct pAS63, the activity
generated by construct pAS25
was only marginally above the background
of about 200 Miller units
found for vector pRKlac290 alone. This result
suggested that the
transcripts originating from P
cicA do
not read into the
clpP gene
but are terminated at the
stem-loop structure T2 (Fig.
4). T2
is localized about 20 bp downstream
of the
clpP stop codon and
30 bp downstream of the stop
codon for the potential
cicA open
reading frame. This
observation and the observation that the T2
inverted repeat is flanked
at both ends by several thymidine residues
on the transcribed strain
indicate that this structure could act
as a bidirectional
transcriptional
terminator.
On the basis of the genetic mapping of the promoter regions, we
performed primer extension experiments (see Materials and
Methods) to
precisely localize the transcriptional initiation
sites for
clpP and
clpX. With primers clpPPE1 and clpPPE2
(see
Materials and Methods), two close start sites were found to be
positioned 59 and 62 bp upstream of the ATG start codon of
clpP (Fig.
5). A
10 box and
a
35 box designating P
P1 were identified
upstream of the
transcriptional initiation sites for
clpP (Fig.
5B). Two
separate transcriptional start sites were found directly
upstream of
clpX by use of two different primers, clpXPE1 and
clpXPE2
(see Materials and Methods). One was positioned 103 bp
upstream of the
proposed translational start codon of
clpX and
immediately
downstream of the
BamHI site (Fig.
4 and
5). Two weaker
start signals seen in Fig.
5A (P
X1) were observed only with
the
clpXPE1 primer, suggesting that they are artifacts. The second
transcriptional start site was located 196 bp upstream of the
clpX ATG start codon and immediately upstream of the
MluI site
(Fig.
4 and
5). Consensus sequences for
10 and
35 promoter boxes
were identified upstream of both transcriptional
start sites (Fig.
5B). The locations of these two transcriptional start
sites downstream
of the
BamHI site and upstream of the
MluI site suggested that
the former corresponds to
P
X1 (pAS6 and pAS7; Fig.
4) and that
the latter corresponds
to P
X2 (pAS26; Fig.
4). It is difficult
to unambiguously
assign the identified promoter regions to
C. crescentus
promoter consensus sequences. As depicted in Fig.
5B,
the
10 and
35
boxes of P
P1, P
X1, and P
X2 show
some similarity
to the consensus sequences of both
32-dependent heat shock promoters (
37,
53)
and
73-dependent housekeeping promoters (
29).
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|
FIG. 5.
Promoter analysis of the C. crescentus clpP
and clpX genes. (A) Primer extension products obtained with
the clpX- and clpX-specific primers are shown in
the rightmost lanes. Sequencing reactions generated with the same
primers are shown in lanes T, C, G, and A. The relevant sequence of the
coding strand is shown to the right of each gel, and the positions of
the major extension products are indicated by arrows. (B) Sequence
alignment of PP1, PX1, and PX2. The
35 and 10 (boxed) and +1 (circled) regions are indicated, and the
distance between the transcriptional start site and the presumed
translational start codon is shown (Nx). The C. crescentus consensus sequences for 73-dependent
(29) and 32-dependent (37, 53)
promoters are shown in boldface below the three clp
promoters.
|
|
Heat shock control of clpP and clpX
promoters.
The components of the ClpXP protease in E. coli (22) and the ClpP peptidase in B. subtilis (15, 47) are induced by heat shock, implying a
role for the corresponding proteases in the degradation of unfolded and
misfolded proteins. To examine if the C. crescentus clpP and
clpX genes are also under heat shock control, we assayed the
activities of the clpP and clpX promoters as well
as the relative concentrations of the ClpP and ClpX proteins after
shifting the temperature from 30 to 42°C. Cells of wild-type strain
NA1000 carrying plasmid pAS2 (PP1-lacZ) or
plasmid pAS24 (PX1-PX2-PX3-lacZ) were
grown to the mid-log phase at 30°C, and aliquots of the cultures were
shifted to the higher temperature. Promoter activity was assayed by
pulse labeling of cells with [35S] Met at different times
after the temperature shift and subsequent immunoprecipitation of the
-galactosidase synthesized. The activity of PP1
increased about 2.5 times during the initial 5 to 10 min after heat
shock, with transcriptional activity remaining high even after
prolonged exposure of the cells to heat (Fig.
6A). In contrast, clpX
promoter activity very rapidly decreased by about 50% after heat shock
(Fig. 6A). The levels of the ClpP and ClpX proteins followed a similar
trend, as the ClpP protein level increased about two times upon heat
shock induction, while the ClpX protein level decreased to below 50%
its initial value (Fig. 6B). These results suggested that
PP1 is under heat shock control and possibly dependent on
the C. crescentus sigma factor RpoH (36, 53).
Since the clpP gene has recently been shown to be essential
for the growth and viability of C. crescentus
(16), a requirement for RpoH to activate PP1
could explain the finding that even at low temperatures, the C. crescentus rpoH gene could not be inactivated (36).
View larger version (16K):
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|
FIG. 6.
Heat shock control of ClpP and ClpX expression. (A) The
transcription of clpP ( ) and clpX () was
determined with cultures of NA1000/pAS2
(clpP::lacZ) and NA1000/pAS24
(clpX::lacZ) by
[35S]methionine labeling, -galactosidase
immunoprecipitation, and quantitation at different times after a shift
in the temperature from 30 to 42°C. (B) Cellular concentrations of
ClpP () and ClpX () after the temperature shift from 30 to
42°C, as determined by immunoblot analysis for the same samples as
those analyzed in panel A. The values are relative to the level of
transcription or protein at 30°C.
|
|
Cell cycle- and growth phase-dependent transcription of
clpP and clpX.
Both clpP and
clpX are essential genes in C. crescentus, and
their product, the ClpXP protease, is involved in cell cycle control in
this organism (16). To examine a possible cell
cycle-dependent activity pattern of the clpP and
clpX promoters, all four clpP and clpX
promoter fragments fused to the lacZ reporter gene (pAS2, pAS6, pAS26, and pAS64; Fig. 4) were assayed by pulse labeling at
different short intervals during the cell cycle. Subsequent immunoprecipitation with an anti--galactosidase antibody allowed us
to quantitate promoter activity at any given time of the cell cycle.
Cell cycle fluctuations of promoter activity were observed only for
PX1 and PX3 (Fig.
7A). The activity of PP1 and
PX2 did not change significantly during the cell cycle. The
activity of PX1 and PX3 peaked in predivisional
cells (Fig. 7A). However, while PX1 activity was low in
stalked cells and high in swarmer cells, the pattern for
PX3 activity was the opposite (Fig. 7A). Even though these
activity patterns are reproducible, their significance is not clear.
Immunoblot analysis with anti-ClpP and anti-ClpX antibodies had
revealed that the concentrations of both proteins did not fluctuate
significantly during the cell cycle (16).
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FIG. 7.
Cell cycle- and growth phase-dependent expression of
clpP and clpX. (A) The relative activities of
PX1 () and PX3 () were determined during
the cell cycle by pulse labeling of synchronized cultures of strain
NA1000 containing plasmids pAS6 (PX1) and pAS64
(PX3), respectively, with [35S]methionine for
5 min at different intervals and determining -galactosidase
synthesis during the pulse time by immunoprecipitation (see Materials
and Methods). The promoter activity is shown relative to maximal
activity. Progression of the cell cycle is indicated schematically at
the bottom. (B and C) Growth phase-dependent expression (B) and
cellular levels (C) of ClpP and ClpX. (B) Stationary-phase overnight
cultures were diluted in fresh PYE complex medium (NA1000, NA1000/pAS2,
and NA1000/pAS24) or PYE complex medium plus 0.2% xylose
(NA1000/pCS225), and growth was monitored by determining the
OD660 (). At different intervals, samples were removed
from the cultures, and -galactosidase activity (Miller units) was
determined as described in Materials and Methods for the following
strains: NA1000/pAS2 (PP1) (), NA1000/pAS24
(PX1, PX2, and PX3) (), and
NA1000/pCS225 (xylX promoter) (). (C) Relative cellular
levels of ClpP () and ClpX () in wild-type strain NA1000 were
determined as a function of the growth phase by immunoblot analysis.
, OD660.
|
|
During the analysis of the
clp promoter fragments, we found
that the strength of the
clp promoters was dependent on the
growth
phase of the cells. To determine the influence of the growth
phase
on the activity of the
clpP and
clpX
promoters,
-galactosidase
production from constructs pAS2 and pAS24
(Fig.
4) was monitored
through the logarithmic and stationary phases.
When cells from
an overnight culture were diluted into fresh complex
medium, the
activity of P
P1 was initially high but
decreased to about 50%
during exponential growth (Fig.
7B). When the
cells reached the
late logarithmic phase, the activity of
P
P1 increased coincidently
with a decrease in growth rate.
Eventually, when the cells entered
the stationary phase, the activity
of P
P1 reached its original
value (Fig.
7B). A similar
behavior was also observed for the
clpX promoters, although
the initial decrease occurred more rapidly
(Fig.
7B). As a control, the
activity of the
xylX promoter (
32)
was assayed as
a function of growth phase in the presence of the
inducer xylose. In
contrast to that of the
clp promoters, the
activity of the
xylX promoter did not change over the course of
exponential
growth and the stationary phase, demonstrating that
promoter induction
in the stationary phase is not a general phenomenon
in
C. crescentus. The levels of the ClpP and ClpX proteins showed
a
similar, but more pronounced, fluctuation (Fig.
7C). These results
suggested that in
C. crescentus, the
clpP and
clpX promoters are
induced in the late exponential and
stationary phases, resulting
in increased ClpP and ClpX levels in
cells.
Determination of the number of ClpP and ClpX molecules per
cell.
Specific antibodies against ClpP and ClpX were used for an
approximate determination of the number of molecules per cell for both
components of the protease (see Materials and Methods). Different concentrations of purified His-tagged ClpP and ClpX proteins
(16) were analyzed in immunoblot experiments, and the
signals obtained were compared quantitatively with the signals from
total cellular protein (Fig. 8). On the
basis of the assumption that both antibodies react similarly with the
full-length His-tagged and native proteins, the numbers of molecules
per cell could be calculated from the data. Averages of 23,000 and
41,000 molecules per cell were determined for ClpP in logarithmically
growing cells and in the stationary phase, respectively. The
corresponding numbers for ClpX were 5,000 molecules per cell in the
logarithmic growth phase and 9,000 molecules per cell in the stationary
phase. Under the condition that all of the protein pools in the cell
are assembled into stable tetradecamers (ClpP) (21, 49) and
hexamers (ClpX) (14), the numbers of complexes were about
1,650 ClpP structures and 830 ClpX rings in the logarithmic growth
phase and about 3,000 ClpP structures and 1,500 ClpX rings in the
stationary phase.
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|
FIG. 8.
Estimation of the number of ClpP and ClpX molecules per
cell. Immunoblot analysis was done with anti-ClpP and anti-ClpX sera
and with specific amounts (picomoles) of purified His-tagged ClpP and
ClpX proteins, respectively, and total proteins from NA1000. The band
intensities of total proteins correspond to the amounts of ClpP and
ClpX present in 2 × 107 cells and 1 × 108 cells, respectively, during exponential growth (Log)
and in 1 × 107 cells and 5 × 107
cells, respectively, during the stationary phase (Stat).
|
|
| |
DISCUSSION |
We have isolated the genes coding for the ClpXP protease in
C. crescentus. ClpXP is a two-component protease with a
peptidase subunit (ClpP) and an ATP-dependent regulatory chaperone
subunit (ClpX). The peptidase by itself does not have any substrate
specificity and requires ClpX for substrate recognition and
interaction. Thus, the presence of the regulatory subunit is crucial
for the specific activity of the protease. In E. coli, an
alternative ATPase, ClpA, is able to interact with and compete for ClpP
subunits (14), with ClpXP and ClpAP having distinct
substrate spectra (13). However, the molecular basis for
substrate selection by the regulatory proteins is not known. In some
cases, substrate-specific mediator proteins are required for the
interaction between the Clp ATPase and the substrate protein (46,
54). In other cases, a direct substrate interaction is mediated
by PDZ-like domains in the C-terminal part of the chaperone
(27). In support of this notion, the ClpX C termini are
strongly conserved in all bacterial ClpX sequences, indicating a
conserved substrate recognition mechanism for ClpX homologues in
different organisms. The results of our complementation experiments are
in agreement with such a prediction. A copy of the C. crescentus
clpX gene can functionally substitute for clpX in
E. coli. This fact strongly suggests that the
ClpXCc protein is able to interact with at least some of
the ClpXEc substrates and with the ClpP peptidase in
E. coli. On the other hand, clpXEc was not able to complement a clpXCc mutation.
Unlike clpXEc, clpXCc has
been shown to be essential for cell cycle progression and thus for
growth (16). Thus, one or several substrate proteins for
which ClpXP-dependent degradation is indispensable must exist in
Caulobacter (16). The failure of
ClpXEc to support growth in Caulobacter could be
due to its inability to interact with ClpPCc or to
inadequate recognition of some or all of the ClpXP substrates.
Alternatively, lack of proper control of the
ClpXEc-substrate interaction might be the reason for the
deficient function of ClpXEc in C. crescentus.
Since ClpXP plays a role in controlling the C. crescentus
cell cycle, tight temporal control of its activity and thus ClpX
substrate accessibility is postulated.
In E. coli, the clpP and clpX genes
are coregulated at the transcriptional level (11). In
particular, heat shock results in a twofold induction of transcription
of the clpPX operon (11), suggesting a role for
the ClpXP protease in the stress response in E. coli. In
contrast, we found that in C. crescentus, transcription of
the clpP and clpX genes is not linked and only
the clpP gene is subject to heat shock induction.
PP1 was mapped by primer extension analysis and shown to
resemble the previously identified 32-dependent heat
shock promoters (3, 36, 37, 52, 53). However, based on the
proposed consensus sequences (29, 37, 53), it is difficult
to unambiguously assign PP1 to either the 32-dependent or the 73-dependent family
of promoters. We have observed a twofold increase of PP1
activity in the presence of an additional, plasmid-encoded copy of the
rpoH gene (data not shown), indicating that
32 at least contributes to PP1 activity.
Most transcripts that originated at PP1 terminated
immediately downstream of the clpP gene, probably at a
transcription terminator-like structure. Thus, heat-inducible
PP1 is not responsible for the transcription of the
clpX gene. The transcription of clpX is directed by at least three independent promoters in the clpP-clpX
intergenic region. This region of about 1 kb was shown to code for an
open reading frame (cicA) pointing in the direction opposite
that of clpP and clpX. Data obtained recently
have confirmed that cicA codes for a protein that, like ClpP
and ClpX, is essential for the growth of C. crescentus
(50a). Two of the three clpX promoters were
localized to the region between the 5' ends of cicA and
clpX. Both show some similarity with the postulated
consensus sequence for 73-dependent promoters
(29). The third clpX promoter was located further
upstream, in the coding region of cicA. In contrast to the
data for clpP, heat shock did not result in an increase but rather resulted in a decrease in total clpX transcription.
This finding clearly emphasizes the independent control of the
clpP and clpX promoters. Since the change in the
concentrations of ClpP and ClpX after heat shock was very similar to
the change in the transcription of both genes, we concluded that heat
shock control of these genes is exerted primarily at the
transcriptional level. It is not clear which mechanism is responsible
for the reduction of clpX transcription following heat
shock. Since downregulation did not occur for the xylX
promoter (15a), the observed reduction of clpX
transcription is likely to be the result of a specific regulatory
mechanism rather than a general phenomenon. Thus, unlike ClpXEc, ClpXCc does not appear to be involved
in the heat shock response. A reduction of ClpX synthesis after heat
shock would allow the use of the available ClpP pool more specifically
for stress-associated functions. One way to accomplish this would be
via an exchange of ClpP-associated ATPases, in which ClpX is replaced
by another regulatory ATPase subunit, i.e., ClpA. In vitro, the
affinities of the E. coli ClpA and ClpX proteins for ClpP
have been shown to be comparable, suggesting that ClpA and ClpX compete
for ClpP in vivo (14). The clpACc
gene has recently been cloned and shown to be dispensable for growth at
a normal temperature (34a). We are currently testing the
hypothesis that ClpA is involved in the heat shock response in C. crescentus.
The fact that clpX has three promoters with different cell
cycle controls suggested that Caulobacter very carefully
regulates the expression of this gene and thereby the cellular
concentration of the ClpX protein. While the physiological significance
and the individual control of each promoter remain to be elucidated, tight regulation of the clpX gene is consistent with our
finding that both the depletion and the overexpression of the ClpX
protein are highly toxic for the cell and result in cessation of growth and loss of viability (16). In contrast, the overexpression of the ClpP protein has no obvious physiological consequences for the
cell. Assessment of the molar ratio of ClpP to ClpX has revealed that
ClpP is present in a molar excess in Caulobacter. Under the
assumption that both ClpP and ClpX exist predominantly in their
complexed form, about 1,650 assembled ClpP double rings and 830 ClpX
rings are present in a cell. Since each ClpX hexamer can complex with
two opposite sides of a ClpP tetradecamer in E. coli
(14) and assuming that ClpX and ClpP assemble with the same
stoichiometry in C. crescentus, the available ClpX pool can saturate at most one-fourth of the theoretically accessible peptidase sites. In comparison, E. coli ClpP has been shown to be
limiting compared with ClpX and ClpA combined (14), a
finding that is reasonable in view of clpP and
clpX being coexpressed and thus most likely present in
similar amounts. In Caulobacter, tight control of ClpX
expression therefore might be necessary because ClpX is the limiting
factor for the degradation of one or several key proteins by the ClpXP
protease (16). Alternatively, it is conceivable that ClpX,
in addition to its essential role in protein degradation, fulfills
critical cellular ClpP-independent functions as a chaperone
(50). One of the main future challenges will thus be to
characterize the different activities of ClpX in Caulobacter and to identify the proteins with which it is able to interact.
| |
ACKNOWLEDGMENTS |
We thank the members of the laboratory for helpful comments and
critical reading of the manuscript.
This work was supported by Swiss National Science Foundation fellowship
31-46764.96 to U.J.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Division of
Molecular Microbiology, Biozentrum, University of Basel, CH-4056 Basel, Switzerland. Phone: 41-61-267-21-35. Fax: 41-61-267-21-18. E-mail: jenal{at}ubaclu.unibas.ch.
| |
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Journal of Bacteriology, May 1999, p. 3039-3050, Vol. 181, No. 10
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Copyright © 1999, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
