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INTRODUCTION |
DNA replication is a tightly regulated process to guarantee that
genetic information is precisely copied once, and only once, each cell
cycle (1). This is achieved by means of complex circuits that are
distinct in prokaryotes and eukaryotes, although some similarities have
become evident recently (2-4). Common essential elements are a
cis-acting DNA sequence (the origin of replication) and a
trans-acting factor (the initiator). DNA replication
initiators are single proteins, or multisubunit complexes, that bind
specifically to the origin of replication, where they usually assemble
into oligomers. Initiators play two roles: (i) to melt the strands of
DNA and (ii) bring to the so-created replication bubble a number of
other protein factors. These are required for further extension of the
replication fork (helicases), synthesis of an RNA primer (primases),
and copying the template (DNA polymerases) (reviewed in Refs. 1 and 2).
Any initiator, to be enabled to bind and melt origin DNA, requires an
activation step whose molecular basis is a central subject in research
on both DNA replication and the cell cycle.
A trait common to initiators is that their function is controlled by
ATP binding and hydrolysis (3, 5, 6) defining active or silent
conformational states. In the bacterial chromosomal initiator, DnaA
(reviewed in Ref. 7), ATP is not strictly required for specific binding
to double-stranded origin DNA (oriC). However, it is
essential for melting oriC AT-rich repeats, binding to the resulting single-stranded sequences (6) and then for loading DnaB
helicase. Subsequent ATP hydrolysis, stimulated by the processivity factor of DNApol III (
-clamp) and other cellular factors, renders DnaA unable to initiate further replication rounds. DnaA can then be
reactivated by either acidic phospholipids or DnaK chaperone, which
exchange ADP by ATP (7). On their side, Orc1 and Orc5 subunits of the
eukaryotic initiator ORC1
also bind ATP, which is required for specific recognition of origin
sequences (8). Single-stranded DNA, generated after origin melting,
stimulates ATP hydrolysis and exerts a change in the overall shape of
ORC (5).
Plasmids are extrachromosomal DNA molecules that borrow from their
hosts most of the factors required for replication. However, they often
encode their own initiator, termed Rep (reviewed in Refs. 9 and 10). In
Gram-negative bacteria, Rep proteins usually bind to directly repeated
sequences (iterons) to establish the initiation complex. In addition,
some Rep proteins also bind to an inversely repeated sequence
(operator) that overlaps with the promoter of the rep genes,
thus acting as self-repressors. Dimers of Rep bind to the operator,
whereas monomers bind to the iterons (11-17).
RepA is the initiator protein of pPS10, a plasmid isolated from
Pseudomonas (18). Mutations in an LZ-like sequence motif, found at the N terminus of RepA (19), enhance dimer dissociation (14).
However, a proof for a direct role of LZ in RepA dimerization is still
lacking. An helix-turn-helix motif at the protein C terminus is the
main determinant of RepA binding to both operator and iteron DNA
sequences (20). We have proposed recently (21) that RepA consists of
two WH domains (reviewed in Ref. 22). Furthermore, a similar WH fold is
found at the C terminus of the eukaryotic/archaeal initiators Orc4/Cdc6
(4, 23), underlining the relevance of studying the molecular mechanism
of RepA activation. We had proposed that dissociation of RepA dimers
into monomers would result in a structural change altering the relative
arrangement and compaction of its two WH domains (21). In the dimers,
the C-terminal domain (WH2) binds to the operator through the major
groove, whereas the N-terminal domain (WH1) acts as dimerization
interface. In the monomers, WH2 binds to iteron DNA as it does to the
operator, whereas the remodeled WH1 binds to the opposite end of iteron through both the phosphodiester backbone and the minor groove (21).
This model has been then confirmed by the crystal structure of the
monomer of a related initiator, RepE54, bound to iteron DNA (24). It
shows that the
-helix that might resemble an LZ in the dimers is
found in the monomers split into two helical portions, packed together,
and buried in the hydrophobic core of WH1 (24).
A feature of Rep-type initiators is that, unlike DnaA and ORC, they do
not bind ATP. Thus, although Rep proteins can promote some structural
transitions in DNA, most plasmids require DnaA to aid in origin melting
(17, 25-30) and helicase loading (28, 29). However, they still
experience conformational activation (reviewed in Ref. 30). Molecular
chaperones, either the triad DnaK-DnaJ-GrpE or ClpA, have been
implicated in the activation of the Rep-type initiator proteins of
plasmids P1 (11, 31-34) and F (35). ClpX chaperone has a role in the
activation of the initiator of RK2 plasmid (36). In addition, ClpA
unfolds P1 Rep to be proteolyzed by ClpP (37, 38). We have proposed
recently (4) that Hsp70 chaperones, the eukaryotic homologues of DnaK, are implicated in the assembly of ORC. Two alternative functions have
been proposed for chaperones in P1 replication; either Rep dimers would
be so stable that chaperones are required to dissociate them (31, 32),
or they would modify Rep conformation (39). Thus, some monomeric
mutants still require the action of chaperones in vivo (33,
34), although they have higher association rates to iterons in
vitro and increased initiation frequency (40). In addition, dimers
of a few Rep-type proteins dissociate spontaneously by dilution (34,
41, 42). The current view is that chaperones would induce in Rep both
monomerization and a conformational change, whose exact nature remains
to be determined (30, 39, 43).
We have searched for determinants of the activation of pPS10 RepA. We
show in this paper that micromolar amounts of a single iteron DNA
sequence actively induce in vitro both the dissociation of
RepA dimers into monomers and a conformational change. On the contrary,
binding of RepA dimers to the operator sequence neither dissociates
them nor changes their conformation. The ligand-induced monomerization
of RepA dimers with a coupled conformational change, reported in this
paper, would thus be a case for the allosteric effect of DNA substrates
in the structure of their binding domains (44, 45) and a novel player
in the activation of plasmid DNA replication initiators. Furthermore,
we also present a detailed biophysical characterization of RepA-2L2A, a
mostly monomeric species of RepA obtained by site-directed mutagenesis
of two Leu residues in the LZ-like motif. Our data suggest that it
resembles a transient folding intermediate in the way from dimers to
active monomers. In RepA-2L2A, the mutations disable the first of the two
-helical portions of the putative LZ to fold back into the second one, thus disrupting hydrophobic interactions at the core of
monomeric WH1.
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EXPERIMENTAL PROCEDURES |
Cloning, Expression, and Purification of
RepA-2L2A--
His6-RepA-WT was expressed and purified,
and when required, its tag was removed as described previously (21).
Mutant repA-2L2A gene was obtained by PCR, using as template
the expression vector pRG-recA-NHis-repA-WT (23)
and primers including the following: (i) a SacII site plus
sequences coding for the 5' end of repA but with mutated
codons (GCC for Ala-12 and GCG for Ala-19) replacing those for leucines
in the putative LZ and (ii) a HindIII site plus a TGA stop
codon and sequences complementary to the 3' end of repA.
Mutations were verified by automated DNA sequencing. Expression of
His6-RepA-2L2A was carried out at 30 °C in
Escherichia coli SG22097 (clpXP), as
performed for RepA-WT (21). His6-RepA-2L2A was purified to
homogeneity from inclusion bodies using the same procedure developed
for deletion mutants in RepA (21). Protein stocks were kept in 0.5 M (NH4)2SO4, 50 mM NH4-acetate, pH 6.0, 10 mM
-MeEtOH, 0.1 mM EDTA, 10% glycerol.
RepAs concentrations were calculated based on their absorption at 280 nm in 5.6 M GuHCl, considering a molar extinction
coefficient of 17210 M
1·cm
1
(www.expasy.ch/tools/protparam.html).
Testing in Pseudomonas aeruginosa the Effect of the
His6 Tag and the 2L2A Mutation in RepA--
pPSEC, a new
series of shuttle vectors including the pPS10 and ColE1 replicons, was
constructed as follows. pRG9B (46) was digested with BamHI
to give a 4-kbp fragment including the pPS10 replicon (18) and the Kn
resistance determinant. This fragment was then filled in with Klenow
and ligated to a 759-bp DraI-AflIII (ori) fragment from pUC18, in which the latter site had also
been made blunt-ended. The resulting plasmid (pPSEC1) was then modified to substitute the 5' half of the repA-WT gene (a 317-bp
EcoRI-SphI fragment) by versions encoding a
His6 tag (pPSEC2) and the 2L2A mutations (pPSEC3). This was
carried out by PCR on the corresponding pRG-recA-NHis-repA-WT/2L2A templates (see above),
using as primers the following: (i) a 41-bp tail with the sequence
between the EcoRI site and the ATG initiation codon in
repA (18), plus the encoded His6 tag and (ii)
sequences from the complementary non-coding strand comprising the
SphI site in repA. The three pPSEC constructs were rescued in E. coli JM109, checked by DNA
sequencing, and then transformed into P. aeruginosa PAO1024
(18). Relative plasmid copy numbers were determined from cultures of
PAO1024/pPSECs in LB medium, supplemented with Kn to 50 µg/ml, at
30 °C. 1.5-ml aliquots were harvested at A600 = 0.5, and total lysates were obtained as described (18). 60 µl
(1/10) of the lysates were then loaded into 0.8% agarose-Tris acetate
EDTA gels, run at 30 V for 12.5 h, and stained with EtBr.
Southern blotting was carried out by transferring the gels to nylon
membranes (18) and then hybridized with a 954-bp EcoRI
repA fragment from pRG9B, radiolabeled by random priming
with Klenow and 40 µCi of [
32P]dCTP (Amersham
Biosciences). X-ray films (AGFA) were exposed and then both the
hybridized plasmid bands and the EtBr-stained chromosome were
quantified using GelDoc (Bio-Rad). The amount of intracellular RepA in
P. aeruginosa cells carrying pPSEC plasmids was estimated by
Western blotting, as described (47). 200-µl culture aliquots were
taken at different A600 values. Total lysates were obtained, resolved by SDS-PAGE, and transferred to nitrocellulose membranes by electroblotting. Membranes were incubated with an anti-RepA rabbit polyclonal antiserum (1:1000) and then revealed with
horseradish peroxidase-conjugated donkey anti-rabbit IgG (1:10000) by
the ECL procedure (Amersham Biosciences). RepA amounts were estimated
by comparing the intensities of the specific luminescent bands in
pPSEC-containing cells with those in lanes including extracts from
plasmid-free cells, the latter supplemented with known amounts of pure
RepA. The number of cells contributing to the loaded extracts was
estimated after serial dilutions of the cultures by counting at a
microscope. Viables were then determined by plating on LB-agar
supplemented with Kn. Molar concentration of RepA was calculated
considering for a P. aeruginosa cell the same volume
determined previously for E. coli (4.6 × 10
10 µl) (48).
EMSA--
DNA oligonucleotides containing either the operator or
the four iteron sequences of pPS10 (18) were synthesized and then phosphorylated with T4 polynucleotide kinase. After annealing with
their complementary strands, they were cloned into the SmaI site of pUC18 and sequenced. Plasmids were cut at the XbaI
site and then labeled with 2 units of Klenow and 30 µCi of
[
-32P]dCTP for 30 min at 25 °C. Fragments were
excised with EcoRI and purified by electrophoresis in
polyacrylamide gels (13). Increasing amounts of RepA proteins (WT or
2L2A) were incubated in ice with radiolabeled DNAs (4000 cpm) in a
20-µl volume of 20 mM Hepes·NH4, pH 7.8, 5 mM
-MeEtOH, 0.1 mM EDTA, 6% glycerol, 50 µg/ml bovine serum albumin, and
(NH4)2SO4, provided by the supplied
protein, to 0.2 M. Samples, assembled in ice, were then transferred to room temperature for 30 min before loading into 6%
polyacrylamide (29:1)-0.5× TBE gels. Electrophoresis was run at 150 V
and 4 °C. Gels were then dried out and exposed to x-ray films.
Analytical Ultracentrifugation--
Sedimentation equilibrium
experiments were performed in a Beckman XL-A analytical
ultracentrifuge. RepA stocks were dialyzed against 0.25 M
(NH4)2SO4, 50 mM
NH4-acetate, pH 6.0, 0.1 mM EDTA. 60-µl
samples, with different protein concentrations, were displayed into
six-channel centrifuge cells, with a 1.2-cm optical path and
centerpieces of epon charcoal. Sedimentation equilibrium gradients were
formed at 5 °C by spinning at either 13000 or 15000 rpm. Radial
scans were taken at different wavelengths. Baseline offsets were
measured at 50000 rpm. Sedimentation velocity of RepA, either alone or
complexed with DNA fragments, was performed at 50000 rpm and 5 °C
with 350-µl samples displayed into double sector cells. Data from
both types of experiments were processed as described (49). The
sedimentation coefficient distributions for the RepA-DNA complexes
(Fig. 6C, inset) were calculated by direct linear
least squares boundary modeling of the sedimentation velocity data
(ls-g*(s)) using SEDFIT (50).
Spectroscopic Assays--
Steady state fluorescence spectroscopy
was performed in a Fluorolog Jobin Yvon-Spex spectrofluorimeter.
350-µl samples of His6-RepA-WT or
His6-RepA-2L2A (5 µM) in 0.25 M
(NH4)2SO4, 50 mM
NH4-acetate, pH 6.0, 0.1 mM EDTA buffer were
displayed into 0.2 × 1.0-cm path length quartz cuvettes and left
to equilibrate at 5 °C. Trp-94 in RepA was then selectively excited
(2-nm slit) at 295 nm (21), and emission spectra were acquired between
300 and 450 nm (3-nm slit, 
= 1 nm). For extrinsic
fluorescence measurements, bis-ANS (Sigma) was supplied, from a stock
in methanol, to a final concentration of 10 µM. Samples
were left to equilibrate for 30 min at 5 °C and then were excited at
395 nm. Emission spectra were acquired in the interval of 400 to 600 nm. The contribution of the buffer was subtracted to all spectra.
Circular dichroism analysis of His6-RepA-WT,
His6-RepA-2L2A, and their complexes with DNAs was performed
in a Jasco-720 spectropolarimeter, using 0.1-cm path length quartz
cuvettes. The DNA oligonucleotides tested in the binding assays (RepA
sites underlined) were as follows: 1IR (operator),
5'-GAACAAGGACAGGGCATTGACTTGTCCCTGTCCCTTAAT-3' (39-mer); 1DR (iteron),
5'-ATACCCGGGTTTAAAGGGGACAGATTCAGGCTGTTATCCACACCC-3' (45-mer); TEL (unrelated),
5'-GATCCCACACCCACACACCCACACACCCACACACCCAG-3' (38-mer); plus their
complementary strands. Oligonucleotides were purified, and their
concentrations were determined as described (21). Annealing was carried
out by slow cooling in Tris-EDTA buffer, and double-stranded DNA
stocks (50 µM) were stored at
20 °C. Binding
reactions (200 µl) were assembled in ice, in a solution containing
0.145 M (NH4)2SO4, 50 mM NH4-acetate, pH 6.0, 0.1 mM
EDTA, 1 mM
-MeEtOH, 5% glycerol, plus 5 µM protein and/or double-stranded DNA. Spectra were
acquired at 5 °C from 320 to 200 nm, using a bandwidth of 1 nm, 4-s
response, 50 nm/min scan speed (0.2-nm steps), and 10 millidegrees of
sensitivity. Five to ten spectra were accumulated for averaging. The
spectrum of the buffer was subtracted, and raw ellipticity data (in
millidegrees) were transformed to molar ellipticity ([
])
(in degrees·cm2·dmol
1). No differences in
the spectra were appreciated when reactions were incubated for variable
times, ranging from 15 min to 3 h, indicating that binding
equilibrium is reached rather quickly (42). Secondary structure
estimates were performed with CDNN (51) at its Web server
(bioinformatik.biochemtech.uni-halle.de/cdnn/java/Started.html). Thermal denaturation analyses were performed on the same samples, overlaid with mineral oil. Ellipticity at 228 nm was measured while
temperature of the cell holder was increased, using a
computer-interfaced water bath, from 5 to 90 °C (0.2 °C steps,
20 °C/h; scans performed at 50 °C/h gave similar curves).
Size Exclusion Chromatography--
Gel filtration assays were
performed in a Superdex-200 HR10/30 column, assembled in an ÅKTA
basic-10 HPLC equipment (Amersham Biosciences). The column was
equilibrated at room temperature in 0.145 M
(NH4)2SO4, 50 mM
NH4-acetate, pH 6.0, 0.1 mM EDTA, 1 mM
-MeEtOH, 5% glycerol. Duplicates (200 µl) of the
samples assayed previously by CD spectroscopy (see above) were then
injected. Chromatography was carried out at 0.5 ml/min in the same
buffer, measuring the absorption of the eluate at 254 nm to optimize
DNA detection.
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RESULTS |
Rational Design and Functional Testing of a Monomeric RepA Mutant
(RepA-2L2A)--
We had described previously that point mutations,
changing to Val the Leu residues of the N-terminal LZ-like sequence
motif found in RepA, resulted in the displacement of dissociation
equilibrium from dimers toward monomers when RepA was fused to MBP
(14). This observation pointed to some (direct or indirect) role for the putative LZ in RepA dimerization. Such a hypothesis received additional support by the finding that an N-terminal deletion of the
LZ-like motif (
N37) results in an increase in the monomeric fraction
of that RepA fragment (21). However, we have since noticed the
following. (i) MBP-RepA fusions, though capable of binding the four
iterons found in pPS10 origin of replication (14), have reduced binding
cooperativity and fail to be wrapped by iteron DNA (not shown). (ii)
When expressed in the absence of the fusion with MBP, those RepA Leu
Val mutants yielded insoluble proteins, which we have been unable
to refold in vitro. To clarify both the role of the LZ-like
motif in RepA dimerization and the conformational change intrinsic to
RepA monomerization (21) we have used the crystal structure of a
monomeric homologous protein, RepE54 (24), to design RepA-2L2A. This is
a new RepA mutant in which the first and second Leu residues of the
hydrophobic LZ heptad (Leu-12 and Leu-19) have been substituted by Ala
(Fig. 1). In RepE54 monomers, the LZ-like
motif is found dislocated into two
-helices, resembling a folded
jack-knife. The shortest one (
1) includes the first Leu residue
(Leu-12 in RepA) and the largest (
2) the third and fourth (Leu-26
and Leu-33), whereas the second (Leu-19) is found in the intervening
turn (Fig. 1B). The rationale behind the new Leu
Ala
mutations is as follows. (i) They would impair a possible dimerization
through the putative LZ, because the side chain of Ala is smaller and
less hydrophobic than that of Leu. (ii) They would necessarily disrupt
the hydrophobic network linking
1, through Leu-12 and Leu-19, to the
rest of the WH1 domain, mainly Trp-94. (iii) Mutations would thus allow
1 to move freely, resulting in an extended conformation of the proposed jack-knife.

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Fig. 1.
A structural framework for the design of RepA
mutants affected in dimerization and conformational activation.
A, overview of the WH1 domain of the monomeric RepE54
initiator from mini-F plasmid (Protein Data Bank entry 1REP) (24), with
its -helical elements highlighted in red; 3 and 4
constitute the helix-turn-helix motif, in which the latter is the DNA
recognition helix. B, slab view removing the
helix-turn-helix to uncover 1 and 2, which include Leu residues
conforming to an LZ-like sequence motif (19). The first two of such
leucines (Leu-12 and Leu-19), substituted by Ala to generate the
RepA-2L2A mutant, are depicted in green. In cyan
is the single Trp in RepA (Trp-94) that is a crucial part of the
network of hydrophobic interactions (distances displayed in
white) to which Leu-12 and Leu-19 also contribute. These
three residues, absolutely conserved in RepA, RepE54, and other Rep
proteins (9, 10, 24), are numbered according to RepA sequence, but the
others (in orange) keep the original labeling from RepE54
(24). A dashed purple arrow indicates the expected
jack-knife movement that would undergo the N-terminal 1-helix around
the hinge including Leu-19, once the hydrophobic network is disrupted
by the mutations in RepA-2L2A that, in addition, would hamper any
possible dimerization through the leucines.
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RepA-2L2A mutant was constructed by site-directed mutagenesis on an
expression vector including repA-WT gene fused to
His6 (21) (see "Experimental Procedures"). With the aim
of testing the effect of the mutations on pPS10 replication in
vivo, repA-2L2A was then transferred to a new
pPS10-ColE1 shuttle vector (Fig. 2A, pPSEC1), replacing the
parental repA-WT gene, to get pPSEC3. To check the effect of
the His6 tag present in RepA-2L2A, a plasmid in which the
repA-WT gene includes the sequence coding for the tag was
also constructed (pPSEC2). After being rescued in E. coli, plasmids were transformed into P. aeruginosa. The
transformation frequency of the repA-2L2A carrying plasmid
decreased by three orders of magnitude when compared with those
encoding for the WT proteins, whereas no differences were apparent
between the tagged and untagged versions of RepA. After four rounds
(~80 generations) of replica plating with no selective pressure, the
Kn resistance marker had been stably inherited in all clones. However,
Southern blot analysis of total lysates from several Kn-resistant
pPSEC3 (repA-2L2A) colonies showed that the plasmid was
integrated in the chromosome, instead of being replicated autonomously
(not shown). This observation suggests that the mutant RepA-2L2A
protein is inactive as DNA replication initiator. Growth curves for
cells carrying plasmids encoding repA-WT show no significant
differences between constructs with or without the His6 tag
(Fig. 2B), confirming that the fused protein is fully
functional as initiator. This was further proved by Southern blotting
total lysates from mid-log phase cells, showing that there are no
significant differences in plasmid copy number either (Fig.
2C). Western blot was performed, with anti-RepA polyclonal
antiserum, on cells harvested at different stages of growth (Fig.
2B), showing similar protein levels for both versions of
RepA-WT. Thus, besides being active as initiator, His6-RepA
is also able to regulate its own synthesis (Fig. 2D). Approximated quantification of the RepA amounts yields about 860 protein molecules (or 430 dimers) per cell. This value is around the
estimations for the Rep proteins of pSC101 (500 molecules) (41) and RK2
(300 molecules) (52) plasmids but far from the those reported for R6K
(about 20-fold higher) (53) and P1 (160 molecules) (47) initiators. It
is relevant for the results presented later in this paper to note that
pPS10 RepA is found at concentrations over 5 µM across
all the growth curves (Fig. 2D).

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Fig. 2.
Testing the functionality of RepA-2L2A and
the effect of the N-terminal His6 tag in
vivo. A, scheme of the shuttle plasmid
series pPSEC, including a ColE1-type replicon, for maintenance in
E. coli, and the pPS10 replicon, for propagation in
Pseudomonas. The pPS10 origin of replication
(oriV) and sequences 5' to the repA gene are
enlarged to highlight the following differences between plasmids:
having His6 tags encoded in repA (pPSEC2/3) or
not (pPSEC1) and including the repA-2L2A mutation (pPSEC3)
or not (pPSEC1/2). All the clones transformed with pPSEC3 were found to
have this plasmid integrated in the chromosome (not shown) and thus are
not included in subsequent panels. B, growth curves for
P. aeruginosa cells carrying the plasmids pPSEC1 or pPSEC2.
C, Southern blotting of total lysates from mid-log P. aeruginosa cells, carrying the plasmids pPSEC1 or pPSEC2.
Upper panel shows the chromosome bands stained with EtBr,
whereas the lower part corresponds to the signal of
32P-labeled repA gene hybridized with the
plasmids. The relative plasmid copy number is indicated
below each track, corrected for loading bias by chromosome
density and normalized to the value of pPSEC1. D, Western
blotting, with polyclonal anti-RepA serum, of total cell extracts
obtained at different points of the growth curve in B. First two tracks, purified RepA standards loaded together
with a total lysate from plasmid-free P. aeruginosa cells.
The estimated amount of RepA, the number of cells contributing to the
load, and the concentration of RepA in the cells are indicated
below each track.
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EMSAs were performed to test whether the failure of RepA-2L2A to act as
initiator is due to have its DNA binding properties altered.
Radiolabeled DNA fragments, including either repA operator sequence (Fig. 3A) or the four
iterons found at pPS10 origin of replication (Fig. 3B), were
incubated with increasing amounts of pure RepA protein, either WT
(His6-tagged or untagged) or mutant 2L2A. The latter was
used with the His6 tag attached, because this increases its
solubility and the tag, by itself, does not substantially affect DNA
replication in vivo (Fig. 2, panels B and
C). RepA-WT binding to operator DNA results in a typical
pattern of two bands. These correspond to complexes including one or
two protein dimers (Fig. 3A, D1 and D2, respectively) (13,
21), to finally yield large protein-DNA aggregates that remain in the well of the gel (W). However, only the first of such complexes, that with a single RepA dimer (D1), is observed with the 2L2A mutant.
Its apparent dissociation constant (Kd(app), the
protein concentration at which 50% of the DNA probe is bound) is at
least 40-fold higher (
0.7 µM) than that for the
His6-tagged RepA-WT (18 nM). Thus, although
with a substantially decreased affinity, the RepA protein in which two
Leu at its LZ-like motif (Leu-12 and Leu-19) were substituted by Ala
seems able to dimerize upon binding to operator DNA. This observation
points to the existence of an additional dimerization interface in
RepA. It would most probably be the
-sheet in WH1, as proposed for
RepE, based in the crystal structure of its monomer (24). Moreover,
binding contacts of RepA-2L2A with the operator DNA must be unaffected, because they are made through WH2 (21) that it is intact in the mutant.
The affinity of the tagged protein for the operator sequence appears to
decrease; about 6-fold more His6-RepA-WT than RepA-WT is
required to get a similar amount of bound DNA (Fig. 3A,
lanes 1 and 3). In addition, the complex
corresponding to the binding of a second RepA dimer (D2) does not
appear. Thus, because the in vivo data show that both
versions (tagged and untagged) of RepA-WT are equally self-regulated
(Fig. 2D), the formation of complex D2 seems not essential
for repA promoter repression. D1 complexes, established by
His6-RepA dimers, show a bit lower electrophoretic mobility
than those for the untagged protein, probably because of the extra
4-kDa mass coming from the His6 tag. Concerning the binding
of RepA monomers to the iterons at the origin (Fig. 3B)
(14), a sharp transition to large protein-DNA complexes that remain in
the well of the gel (W), is observed. This occurs even with the minimal
amount of untagged RepA-WT tested (7.5 nM), whereas only a
tiny fraction of DNA is found in discrete complexes (M1-M4). Although
untagged RepA appears to bind cooperatively to iterons, its
His6 version does not so much; DNA fragments with one to
four iterons bound co-exist (M1-M4), and the complexes stacked in the
well appear at a higher protein concentration (~50 nM).
However, the observed differences in binding cooperativity between
tagged and untagged RepA-WT have no significant effect on their
function as DNA replication initiators in vivo, because the
copy numbers of plasmids coding for them are very similar (Fig.
2C). On the contrary, RepA-2L2A fails to bind stably to iterons (M1 and M2 complexes appear at protein concentrations about
200-fold those operational with RepA-WT; not shown). This is not
surprising, because the domain altered in the mutant (WH1) was found in
RepA-WT to bind an end of iteron sequence stably (21, 24). The failure
of RepA-2L2A to bind iterons in vitro is in accordance with
the fact that the pPS10 derivative coding for this mutant (pPSEC3)
cannot be established autonomously in vivo (see above).
Because the precise nature of the structural alteration induced by the
mutations in RepA remained to be determined, we have performed a
combined biophysical approach to further characterize the structure of
RepA-2L2A.

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Fig. 3.
EMSA on the complexes established by RepA,
and its His6 and 2L2A variants, at the operator and origin
sequences. A, titration of a 32P-labeled
DNA fragment (0.17 nM) including the operator inverted
repeat. F, unbound probe; D1, complex between a
RepA dimer and the operator sequence; D2, complex including
a second RepA dimer-bound (13); W, RepA-DNA aggregates that
stay in the well. B, titration of a radiolabeled fragment
(0.10 nM) comprising the four origin iterons. F,
unbound DNA. M1-M4, complexes including 1-4 RepA
monomers bound to 1-4 iteron sequences (14); W, larger
RepA-DNA complexes in the well. Protein concentrations (nM)
in both experiments were as follows: lane 1, 7.5; lane
2, 18.8; lane 3, 47.2; lane 4, 117.3;
lane 5, 293.1; lane 6, 732.9.
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Physicochemical Characterization of RepA-2L2A as a Metastable
Folding Intermediate--
One of the expected effects of the mutations
designed in RepA-2L2A is to interfere with protein dimerization by
altering the hydrophobic spine in the putative LZ
-helix (see above)
(Fig. 1B). To test the association state of RepA, we have
performed sedimentation equilibrium experiments in an analytical
ultracentrifuge. We have also addressed whether the N-terminal
His6 tag affects RepA-WT dimerization (Fig.
4, panels A and B),
before determining the association state of RepA-2L2A (Fig.
4C). In this protein the His6 fusion was kept
attached for improving solubility (see above). Ultracentrifugation runs
were performed with a RepA range between the minimal concentration
giving a reasonable signal to noise ratio with the absorption optics of
the ultracentrifuge (1-2 µM) and a maximum close to the
limits of RepA solubility (25 µM). RepA-WT remains
essentially dimeric through all the concentration range tested (not
shown), with a net tendency of the protein to assemble further, but no
sign of dissociation. We had reported previously (14) for a fusion with
MBP that RepA-WT, in the lowest concentration range tested here, was
close to the dissociation equilibrium between dimers and monomers. Now,
in light of our new data, that observed dissociation is revealed as a
possible steric hindrance effect of the fused MBP moiety, probably
caused by its large size (47 kDa). His6-tagged RepA-WT
shows a similar behavior to its untagged counterpart, with a slightly
more marked tendency to associate beyond dimers (not shown). This is in
accordance with data shown above pointing to similar in vivo
and in vitro activities for both proteins (see Figs. 2 and
3). Therefore, the rest of experiments described in this paper (see
Figs. 5-7) were carried out with the
His6-tagged version of RepA-WT, to be compared straightforward with those performed with His6-RepA-2L2A.
Sedimentation velocity analysis of His6-tagged and untagged
RepA-WT, at either 5 or 15 µM, allowed to determine a
sedimentation coefficient (s20,w) for
the RepA particle of 4.2 S and a frictional ratio
(f/f0) of 1.2 (not shown). These
values are again compatible with being the most abundant RepA species
dimers, with nearly spherical shape. Thus it is relevant to underline
that, based on the present analytical centrifugation analyses and on
the conditions used in the spectroscopic studies described below,
RepA-WT protein is largely found as stable dimers. Sedimentation
equilibrium analysis on RepA-2L2A shows that this mutant protein is
polydisperse; at the lower concentrations tested (2-7
µM), a significant monomeric fraction is found (Fig. 4C), together with large aggregates. These appear to become
the major species at higher protein concentrations (not shown).
Aggregation, described previously (21) for deletion mutants affecting
the putative LZ in RepA, can be attributed to the exposure of
hydrophobic surfaces (probably an additional dimerization interface;
see above) to the solvent. Both the disruption of RepA dimers and the
exposure of hydrophobic residues, otherwise buried in the core of the
folded N-terminal WH, were expected to arise from the double mutation designed in RepA-2L2A (Fig. 1B).

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Fig. 4.
The molecular masses of the distinct RepA
species determined by sedimentation equilibrium.
Symbols represent the experimental gradients (UV absorption
of the sample versus radial position in the centrifuge cell)
at sedimentation equilibrium for 5 µM of RepA-WT
(A), His6-RepA-WT (B), and
His6-RepA-2L2A (C). Solid lines are
the best fit gradients for single sedimenting species with molecular
masses of 53, 58, and 29 kDa, respectively. Dashed lines
show the theoretical gradients of the monomer and dimer species of the
three RepA proteins.
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Fig. 5.
Steady state fluorescence spectroscopy
highlights differences in the folds of His6-tagged RepA-WT
and 2L2A mutant. A, fluorescence emission spectra
( excit = 295 nm) of Trp-94 in RepA-WT and RepA-2L2A (both at 5 µM), whose association states and domain compactness are
represented schematically. Dotted vertical lines indicate
the emission maxima for the WT (327 nm) and mutant (348 nm) proteins.
B, fluorescence emission spectra ( excit = 395 nm)
for the extrinsic fluorophore bis-ANS (10 µM) incubated
with either RepA-WT or RepA-2L2A proteins at the same experimental
conditions assayed in A. Dashed vertical line
marks the emission maximum at 472 nm.
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The possible structural changes induced by the 2L2A mutations were then
explored by means of steady state fluorescence spectroscopy. Trp-94,
the single tryptophan residue in RepA (18, 21), is a key node in the
network attaching Leu-12 and Leu-19 to the hydrophobic core of WH1
(Fig. 1B). Thus, it is expected to be a suitable
spectroscopic sensor for any structural modification in its
environment. RepA-WT has nine tyrosines (18). Comparing the excitation
and emission spectra of RepA with those for free tyrosine and
N-acetyl-tryptophanamide solutions we have verified that, at
295 nm, no Tyr residue is excited in RepA. On the contrary, that
wavelength falls in the tail of the excitation spectrum of Trp-94 (not
shown). Thus, Trp-94, either in RepA-WT or in RepA-2L2A (5 µM), was selectively excited at 295 nm and then
fluorescence emission spectra were acquired (Fig. 5A). The
maximum emission was achieved at 327 nm (RepA-WT) or 348 nm
(RepA-2L2A). The extra band at 335 nm observed in RepA-2L2A emission
suggests the presence of two major populations of different rotamers
for Trp-94 side chain. The red-shifted emission found for RepA-2L2A is
characteristic of exposed Trp residues, and it is compatible with the
increased solvent accessibility expected for Trp-94 after the release
of its hydrophobic linkage to the bipartite
-helix (Fig.
1B). On the contrary, the emission of Trp-94 in RepA-WT,
close to 320 nm, is typical of Trp residues buried in a protein core or
in a contacting interface (21). The wavelength for the emission maximum
in RepA-WT did not change after serial dilutions of the protein sample
(tested up to 78 nM; not shown), providing further evidence
for its dimeric association state in a broad concentration range.
Another clue for a structural change affecting Trp-94 in RepA-2L2A is
its enhanced fluorescence emission intensity compared with that for the
WT protein (Fig. 5A). To get additional proof for the
presence of exposed hydrophobic residues in RepA-2L2A, an extrinsic
fluorescence probe was used: bis-ANS, a naphthalene derivative that
binds to solvent-accessible hydrophobic patches in proteins, enhancing
its fluorescence emission over 450 nm (21). Incubation of bis-ANS (10 µM) with RepA-WT or 2L2A (5 µM) results in
spectra with emission maxima around 472 nm (Fig. 5B). The
fluorescence intensity at this wavelength is 61% higher for Rep-2L2A
than for the WT protein, confirming that the former has a larger
hydrophobic surface exposed to the solvent.
As a summary of the results shown so far, RepA-WT is a stable dimer
in vitro in a broad (µM) concentration range
(Fig. 4A), similar to that found in P. aeruginosa
cells (Fig. 2D). In terms of RepA association state (Fig.
4B), DNA binding properties (Fig. 3), or even its activity
as a DNA replication initiator in vivo (Fig. 2), there is no
significant difference in having a His6 peptide fused to
the RepA N terminus. RepA-2L2A, in which two Leu residues in the
putative LZ (Leu-12 and Leu-19) were substituted by Ala (Fig. 1),
exposes hydrophobic residues to the solvent, as revealed by
fluorescence studies (Fig. 5) and by its enhanced tendency to
aggregation. Thus, it exhibits the biophysical properties expected for
a metastable monomeric (Fig. 4C) protein folding intermediate (54).
Binding to Iteron DNA Dissociates RepA Dimers into Monomers and
Acts as Allosteric Effector on Protein Conformation--
With the aim
of deepening our understanding of the structural basis of the
recognition of operator and iteron DNA sequences by RepA dimers and
monomers, respectively, we have performed a CD spectroscopy
study on RepA-DNA complexes (see Figs. 6
and 7). 5 µM
His6-tagged RepA-WT protein were incubated with equimolar amounts of double-stranded oligonucleotides, including either the
repA operator sequence (1IR), a single iteron (1DR), or an unrelated sequence of similar size (yeast telomeric TEL). Fig. 6A shows overlaid CD spectra of the oligonucleotides alone
and in complex with RepA-WT. The bands with positive ellipticity
([
]) in the near UV range of the spectra (320-250 nm) arise from
the stacking of DNA bases, with nearly null contribution from protein aromatic residues. In the far UV (250-200 nm) the predominant contribution comes from the protein moiety. Having no substantial differences in the spectra, between 320 and 250 nm, for the free and
protein-bound states of a given DNA can be interpreted as a sign of no
significant structural alterations in the DNA (55). Because that is the
case for the distinct RepA-DNA complexes in Fig. 6A, the
spectra of free DNAs can be subtracted from those of their complexes
with RepA, thus focusing on possible structural changes on the protein
side. The result of this algebraic operation is depicted in Fig.
6B, in which two types of protein spectra become evident:
(i) Curves with double minima at about 208 and 222 nm, a signature for
a significant proportion of
-helical secondary structure (33%), are
one type. These correspond to free RepA-WT and in complex with the
operator oligonucleotide 1IR. (ii) Curves with a broader minimum around
215 nm, attributable to an increase in the
-sheet component (by
about 4%) at the expense of former
-helices, as observed for
RepA-WT in complex with the iteron oligonucleotide 1DR, are another
type. It is relevant that RepA-WT does not change its spectrum and thus
its structure to a detectable extent, when it is free in solution or
bound to the operator. This is coincident with the fact that RepA is
essentially an homogeneous dimer in solution at this concentration
(Fig. 4B) and that it binds to that DNA sequence as a dimer
(Fig. 3A) (14). However, when it binds to the iteron DNA as
a monomer (Fig. 3B) (14) its spectrum resembles that of the
mostly monomeric RepA-2L2A mutant (Fig. 6B). We have failed
to acquire clean spectra for complexes between this mutant protein and
any DNA, because a cloudy precipitate appears. Although less severe,
protein aggregation is also observed, in the form of noisy
spectra, for the nonspecific complexes between RepA-WT and DNAs of
unrelated sequence, such as TEL (Fig. 6), other mixed sequence
oligonucleotides, and poly(dI-dC) (data not shown). As a summary, our
spectroscopic studies show that RepA has a distinct secondary structure
composition when bound to the inversely repeated operator sequence or
to the directly repeated iteron DNA.

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Fig. 6.
Iteron DNA as an allosteric effector on both
RepA structure and association state. A, near and far
UV CD spectra of dsDNA oligonucleotides (5 µM;
filled symbols) and their complexes with equimolar amounts
of His6-tagged RepA-WT (empty symbols):
inversely repeated operator (1IR, in red),
directly repeated iteron (1DR, in blue), and
unrelated yeast telomeric (TEL, in green)
sequences. B, far UV CD spectra of His6-RepA-WT
and 2L2A proteins at 5 µM (circles), plotted
together with the result (diamonds) of subtracting the
spectra of naked DNAs to the protein-DNA complexes. C, HPLC
gel filtration elution profiles of duplicates of the samples in
A, maintaining the same color and symbol codes. Drawings
close to the peaks interpret the different DNA fragments and their
complexes with His6-RepA-WT. Inset shows the
sedimentation coefficient distributions calculated for both protein-DNA
complexes, which were spun down in an analytical ultracentrifuge just
after peak elution.
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Fig. 7.
Binding to operator or iteron DNAs increases
the thermal stability of His6-RepA-WT. Thermal
denaturation profiles were acquired for duplicates of the samples in
Fig. 6 (the same colors and symbols are adopted), measuring the
evolution of molar ellipticity ([ ]) at 228 nm (protein secondary
structure) with increasing temperature. Inset contains
thermodynamic parameters calculated from the curves.
Tm, temperature for half transition between
folded (low values) and unfolded ( getting close to 0) states.
Tm, variation of the melting temperature of
His6-RepA-WT in the complexes with DNA, compared with that
for the protein alone.
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To correlate the observed conformational change with the association
state of RepA, we carried out EMSAs (not shown), size exclusion
chromatography (Fig. 6C), and sedimentation velocity (Fig.
6C, inset) on the same samples tested by CD.
EMSAs consistently result in that the RepA-operator complex slightly
has lower electrophoretic mobility than the RepA-iteron band (not
shown). Taking into account that the free iteron oligonucleotide (1DR)
is 6 bp larger than the operator one (1IR), this result is compatible
with having a larger protein mass in the latter complex (a dimer) than
in the former (a monomer). However, EMSAs are not conclusive, because the minute difference in the electrophoretic migration between both
types of complexes could be also because of disparate bending behavior
of the bound DNA fragments. For the sample including RepA and the
unrelated oligonucleotide TEL, no discrete retarded band is visible but
some smearing, indicative of low affinity, nonspecific binding (not
shown). When the samples were injected into a size exclusion column the
specific RepA-oligonucleotide complexes remain stable, and their
hydrodynamic behavior (Fig. 6C) was unambiguous. The
relative elution position of the peaks corresponding to bound DNAs is
the reverse of that for the unbound species (the assay is sensitive
enough to detect the aforementioned slight difference in the sizes of
free oligonucleotides). This fact can be explained if the protein mass
associated with the smaller operator DNA piece (1IR) is substantially
larger (a dimer) than that associated with the longer iteron (1DR)
fragment (a monomer). The possibility of having just one subunit in a
RepA dimer bound to the iteron, as proposed for the Rep protein of R6K
plasmid (16), is thus very unlikely. Such a complex would be expected
to elute just before the RepA-operator one. To characterize further the
nature of the complexes eluted from the gel filtration column,
analytical ultracentrifugation was performed. Unfortunately, RepA-DNA
complexes dissociate during the time course required to achieve
sedimentation equilibrium (not shown). However, the interpretation
proposed above is in agreement with the results of sedimentation
velocity experiments performed, immediately after the gel filtration
runs, with the peak fractions. There is a substantial difference in the
sedimentation coefficients for the RepA-1IR (6.5 ± 0.2 s) and the RepA-1DR (3.8 ± 0.2 s)
complexes. These sedimentation coefficients are only compatible with
being the RepA-1IR complex significantly more compact
(f/f0 = 1.2) than RepA-1DR
(f/f0 = 2.0). The latter value
fits with the elongated shape of a complex between a RepA monomer and
an oligonucleotide with identical length to 1DR, modeled on the crystal
structure of RepE54 (24) (not shown). Therefore, our results clearly
show that the structural changes in RepA, coupled with binding to
iteron DNA (Fig. 6B), are linked to dissociation of the
otherwise stable protein dimers (Fig. 4B) into monomers
(Fig. 6C).
It is noteworthy that, for unbound RepA, we have not found any sign of
dissociation of dimers after incubation times ranging from 30 min
(spectroscopy; see Figs. 5 and 6) to 16 h (sedimentation equilibrium; see Fig. 4). On the contrary, upon incubation with origin
DNA RepA dissociation (Fig. 6C) and the coupled structural transformation (Fig. 6B) occur within a few minutes. Because
serial dilutions of unbound RepA exhibit the fluorescence emission
spectrum characteristic of dimers (Fig. 5A) at least up to
low nanomolar concentrations (not shown), the possibility that at 5 µM a small fraction of monomeric RepA, in equilibrium
with dimers, would bind to the iteron sequence and that this event
would drive monomer formation seems unlikely. In summary, there are
solid basis to affirm the following. (i) Because the
Kd for RepA dimers appears to be in the low
nanomolar-subnanomolar range, in our experimental (micromolar)
conditions the equilibrium is well displaced toward the dimeric
species. (ii) Along broad time courses no monomeric unbound fraction
has been detected for RepA. Thus the reported induction of RepA
dissociation by iteron DNA seems to be genuine.
Thermal denaturation analyses performed by optical techniques, such as
CD spectroscopy, provide valuable information on the thermodynamic
stability of proteins (4, 21) and their complexes with DNA (55). We
have carried out this kind of approach to study the effect on RepA
stability of binding to the DNA sequences described above (Fig. 6).
Inspection of the CD spectra in Fig. 6A reveals that, at 228 nm, the contribution of the oligonucleotides to the ellipticity
([
]) of their complexes with RepA is essentially null, whereas
that from the
-helical and
-sheet structural elements in RepA
(Fig. 6B) is significant. Thus, we plotted the variation of
228 with temperature for each kind of RepA-DNA complex (Fig. 7) and
then compared the resulting curves. Because thermal denaturation of
RepA and its complexes is irreversible, a detailed thermodynamic analysis of the CD profiles was precluded. The temperature at which
50% of protein molecules unfold (Tm) was
determined, because it is essentially unaffected by irreversibility and
gives an idea on protein stability. Binding to operator DNA (1IR)
stabilizes RepA, relative to its unbound state, by 12.4 °C, whereas
binding to iteron DNA has a smaller effect (4.8 °C). The more
pronounced slope of the curve for 1IR complexes indicates that RepA
denaturation becomes more cooperative upon operator binding by the
dimers. This effect is also observed for the complexes of RepA monomers with iteron DNA (1DR), although to a lesser extent. However, binding to
nonspecific sequences (TEL) severely decreases the stability of RepA
(by 11.5 °C), approaching the Tm values
measured for the mostly monomeric folding intermediate RepA-2L2A (Fig.
7) and for an N-terminal partial deletion derivative (
N37) reported
previously (21). This observation suggests that, as described for some restriction endonucleases (56), nonspecific DNA could, to some extent,
exert conformational changes in RepA (Fig. 6B) enabling it
to scan for specific iteron sequences. However, the resulting RepA
molecules seem to be unstable folding intermediates that, because of
the exposure of hydrophobic residues prone to aggregation (see above),
would recruit molecular chaperones. Therefore, iteron DNA not
only acts as allosteric conformational effector on RepA dimers (Fig.
6B) but also has a role in stabilizing the structure of the
resulting monomers.
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DISCUSSION |
RepA-2L2A: a Folding Intermediate in the Pathway from Repressor
Dimers to Initiator Monomers--
In this paper we have described the
design of RepA-2L2A, a mutant in the protein that initiates DNA
replication of the Pseudomonas plasmid pPS10, based on the
available three-dimensional structure of the monomer of a homologous
initiator (24). RepA-2L2A carries a double Leu
Ala substitution
(Leu-12 and Leu-19) in the LZ-like motif found in the protein (14, 19,
21). These mutations were expected to enhance dissociation of an
otherwise dimeric protein into monomers and to destabilize the
hydrophobic core of the latter (Fig. 1). Functional characterization of
RepA-2L2A indicates that it is inactive as initiator of replication
(Fig. 2). RepA-2L2A has affected its binding to iteron DNA sequences (Fig. 3B) and reduced its affinity for operator DNA (Fig.
3A), where RepA binds as a dimer (13, 21). Hydrodynamic
studies (Fig. 4) show that RepA-2L2A can be largely found as a monomer in a narrow micromolar range, forming nonspecific aggregates at higher
concentrations. Dimerization of WH domains (23) through hydrophobic
residues in the
-helices equivalent to
2 and
4 in Rep proteins
(Fig. 1) has been described for the complex between the eukaryotic
transcription factors E2F4 and DP2 (57). However, a role in
dimerization for the antiparallel
-sheet found at the N-terminal WH
domain of RepE54 is also possible (24). To determine whether the
LZ-like motif has a direct contribution to the interprotomeric contacts
in RepA dimers, or whether it rather favors RepA association indirectly
(e.g. stabilizing the dimeric conformation), will require solving the three-dimensional structure of a Rep protein dimer. Spectroscopic evidence (Fig. 5) supports that RepA-2L2A resembles a
transient folding intermediate, with hydrophobic residues partially exposed to the solvent, expected to occur in the pathway from repressor
dimers to initiator monomers. This is the first report on the isolation
and characterization of a conformational intermediate in a Rep-type
initiator protein. In addition, we have shown that short
His6 fusions to RepA have a minor effect on the association state of the protein in vitro (Fig. 4B), which
remains as a dimer in a broad micromolar range. The His6
tag does not alter RepA initiator activity in vivo (Fig. 2),
despite showing some reduction in its binding cooperativity to iteron
sequences in vitro (Fig. 3B).
The Allosteric Effect of Iteron Binding on RepA Conformation and
Association State--
At the same concentration in which RepA is a
dimer (Fig. 4) with no sign of dissociation into monomers, binding to
an oligonucleotide encoding for a single iteron origin sequence results
in dissociation of RepA into monomers (Fig. 6C) and a change
in protein secondary structure, which becomes similar to that for the
monomeric intermediate RepA-2L2A (Fig. 6B). On the contrary,
the structure of RepA dimers appears to be unaltered by binding to the
inversely repeated operator sequence (Fig. 6B).
Interestingly, binding to both types of DNA stabilizes RepA against
thermal denaturation, although to a different extent (Fig. 7). This
fact also points to differences in the structures of RepA dimers and
monomers. In the same experimental conditions, nonspecific DNA
oligonucleotides result in precipitation of RepA and in destabilization
of the structure of the protein (Fig. 7). Structural changes in a
number of DNA-binding proteins upon specific ligand recognition have
been previously reported (44, 45, 56) as cases of allosterism. However,
apart from being found in some viral initiators (58, 59) and in ORC
with single-stranded DNA (5), they are new events in the initiation of
DNA replication in bacterial plasmids.
The precise mechanism through which iteron DNA exerts the allosteric
effect on RepA structure described in this paper remains to be
determined in its molecular details. It is worth noting that in the
eukaryotic transcription factor Ets1 an
-helix (
I-2) is found at
the N terminus of the three
-helices of its WH domain (60), packing
against the first helix very much as
1 and
2 do in Rep proteins
(Fig. 1B). A phosphate in DNA backbone triggers an
allosteric transition in Ets1
I-2 by establishing a hydrogen bond
with the amide NH of a Leu residue at
1 N terminus. This corresponds in RepE54 monomers with Arg-33 (Fig. 1B), which
also binds to iteron phosphate backbone in the crystal structure (24), whereas modeling of Rep dimers suggests that it must be displaced apart
from the operator phosphate backbone (24). Arg-33 in RepE54 consistently aligns with hydrophobic residues (mainly Leu, as in Ets1)
in RepA, other members of its family (9, 10, 24), and the eukaryotic
and archaeal initiators Orc4/Cdc6 (4). Thus a way is opened to a
possible general role of this residue in triggering allosteric
transitions in WH initiator proteins.
The Role of Chaperones and Specific Origin Sequences in the
Structural Activation of Rep Proteins--
At the concentrations it is
found in Pseudomonas cells in vivo (Fig.
2D), RepA is dimeric (Fig. 4), acting as transcriptional repressor (13, 20). It has been found for P1 (34, 42) and pSC101 (41)
plasmids that their Rep proteins can spontaneously dissociate into
monomers just by dilution to low/sub-micromolar concentration. However,
besides pPS10 RepA, the initiators of F (12), R6K (16, 61), and
RK2 (15, 36) plasmids have their Kd values in
the low nanomolar range. For the latter the requirement for an active
mechanism in Rep dissociation, and in the coupled conformational
change, thus remains as a bottleneck. Our findings on the ability of an
iteron sequence to induce structural transitions in RepA confer a
property to origin DNA that was attributed previously to chaperone
action alone (reviewed in Ref. 30).
Based on data presented in this paper, we propose that the structural
changes in Rep that are coupled to dimer dissociation would imply the
following: (i) releasing protein-protein interactions between
protomers, involving the LZ-like motif and/or the
-sheet in the
first domain (21, 24); (ii) remodeling the hydrophobic core of WH1,
where the relevant Leu residues (Leu-12 and Leu-19) are tightly packed
with Trp-94 (Fig. 1B); (iii) an increase in
-strand
structure at the expense of the
-helical components (Fig.
6B). The rearrangement experienced by Rep would necessarily generate transient intermediates with properties similar to those of
the RepA-2L2A mutant, namely having hydrophobic sequence patches exposed to the solvent and a loosely folded core (Fig. 5), both features of molten globules (54). This explains why monomeric RepA, when free in solution and in the absence of iteron DNA, is prone
to aggregation through WH1 (see above) (21). To cope with these
challenges to Rep activation, different, albeit complementary, molecular mechanisms can be envisaged.
Chaperones can directly dissociate Rep dimers and simultaneously change
the conformation of the monomers, to make them competent for iteron
binding. This has been reported for the DnaK-DnaJ-GrpE triad (11, 31,
33-35), ClpA (32, 43), and ClpX (36). These could target WH1 in Rep;
ClpA recognizes a region at the N terminus of P1 Rep (residues 10-70)
(62) as the first step toward unfolding, either for its activation as
initiator or for degradation by the associated ClpP protease (37, 38).
DnaK, on its side, recognizes a hydrophobic patch (residues 36-49) in the same Rep protein (63), which in pPS10 RepA corresponds to a
sequence (residues 91-105) comprising the Trp-94 residue discussed above. We had shown previously (64) that pPS10 replication in vitro is sensible to DnaK levels. Although they physically
interact (4), if DnaK is acting on pPS10 RepA dissociation remains to be determined. For those Rep dimers that dissociate spontaneously, chaperones could bind to exposed hydrophobic regions (65) in the
monomeric intermediates, thus protecting them from falling into local
energy minima that would act as kinetic traps in the pathway leading to
Rep folding and iteron binding. Otherwise, large Rep aggregates could
arise from the accumulation of monomeric intermediates, but it seems
feasible that the combined action of several chaperones, such as ClpB
plus DnaK-DnaJ-GrpE (66), could rejuvenate them into active initiators.
The allosteric conformational changes induced in RepA by iteron DNA
(Fig. 6) would constitute another means to get the monomeric species,
specially for those Rep proteins that are stable dimers at the
concentrations they are found in vivo (Fig. 2D)
(16, 52, 53). This way could be favored by the cooperativity of Rep
binding to DNA (Fig. 3B). Alternatively, iteron DNA could also capture and stabilize the monomeric folding intermediates produced
after spontaneous dissociation of Rep dimers, as proposed above for
chaperones. An implication of this new activation mechanism is that
only a fraction of the total amount of Rep molecules, coincident with
the number of iteron sequences, would be eventually activated whereas
the others would remain dimeric. On the contrary, there is no obvious
means to limit the number of active Rep monomers generated by the
chaperone-mediated route that could be accumulated in excess over the
levels required for regulated initiation. In addition, the allosteric
binding of iteron DNA to RepA implies that the structural changes
associated to monomerization are intrinsically accessible to Rep
proteins and not necessarily the exclusive product of chaperone action.
In plasmids containing iteron sequences at their origins of
replication, the most favored model for negative control of initiation implies pairing distant Rep-bound iteron repeats ("handcuffing") (30, 67). Except in R6K plasmid (16) handcuffing appears to be mediated
by Rep monomers. However, in some Rep proteins, a number of mutants
have been isolated that, being monomeric hyperactive initiators, fail
to pair iterons (40, 68). Most of such mutations fall in a putative
additional dimerization interface in Rep that, besides the LZ-like
motif, we have discussed for RepA-2L2A. Thus, Rep proteins activated by
the allosteric route resemble the behavior of those hyperactive
mutants. The minimal requirements for pairing are two DNA fragments
containing each a single iteron repeat (69), but we have failed to
detect handcuffing even at 5 µM RepA-iteron complex
concentration (Fig. 6C). Our model leaves open the
possibility for the existence of two kinds of Rep monomers, correlated
with their opposite functions as initiators or negative regulators of
plasmid DNA replication. Thus, unwinding of origin DNA after the
formation of the Rep-iteron nucleoprotein complex might induce a
further conformational change in the monomers, as described for the
chromosomal initiators ORC in eukaryotes (5) and DnaA in bacteria (6).
RepA would then become competent for origin pairing but disabled for a
new round of initiation on recently replicated DNA.