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INTRODUCTION |
The plasmid addiction system of bacteriophage P1 consists of two
proteins: a toxin known as Doc (death on
cure) and an antidote named Phd (prevent
host death) (1). Bacteriophage P1 lysogenizes Escherichia coli as a low copy plasmid that is inherited in
a remarkably stable fashion, in part, because the plasmid addiction system kills cells that have lost the P1 genome (1-5). The mechanism of Doc toxicity is unknown. Phd is, however, known to be degraded in a
fashion dependent on the host ClpXP protease system, and continual
synthesis of Phd is therefore required to counteract Doc toxicity (3).
In daughter cells that have lost the P1 genome, degradation of Phd
eventually results in reduction of antidote activity to a level where
the host is killed. Phd may exert its antitoxin activity by binding Doc
and physically blocking its interaction with cellular target molecules
(1) or by acting in an allosteric fashion to alter Doc structure and
toxin activity. Alternatively, Phd might exert its antidote activity
indirectly by activating another protein that neutralizes Doc.
Some of the regulatory and biochemical properties of Phd have
been established. For example, Phd is a DNA-binding protein that
represses transcription of its own gene as well as that of Doc by
binding to an operator DNA site that overlaps the addiction promoter
(4). Four molecules of Phd bind to the intact operator with dimers
binding to adjacent 10-base pair subsites (6). Phd has a predominantly
-helical structure when bound to DNA or at low temperatures but the
free monomeric protein has a Tm of 25 °C and is
largely denatured at 37 °C (6).
Addiction systems involving a stable toxin and proteolytically unstable
antidote are also used by other low copy plasmids and may be involved
in the regulation of programmed cell death in E. coli
(7-13). Although direct interaction between the toxin and its antidote
is assumed to be the mechanism of inhibition for each of these systems,
experimental evidence for complex formation has been presented only for
the CcdA antidote and CcdB toxin of the F plasmid of E. coli
(14). In the study of the phage P1 addiction proteins presented here,
we demonstrate that Phd binds directly to Doc, forming a trimeric
complex (P2D) with one molecule of Doc and two molecules of
Phd. Complex formation appears to be accompanied by changes in
secondary structure. P2D complexes form at low micromolar
concentrations and are very short lived, ensuring a dynamic equilibrium
between bound and free forms within the cell.
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MATERIALS AND METHODS |
Protein Expression and Purification--
E. coli
strain X90/pRDM032, which expresses Phd and Doc under
Ptac control (4), was grown at 37 °C to an
A600 of approximately 0.7 in Luria-Bertani broth
containing 100 µg/ml ampicillin, and protein expression was induced
by the addition of 100 µg/ml
isopropyl-1-thio-
-D-galactopyranoside for 30 min. Cells
were harvested by centrifugation (3,500 × g for 30 min
at 4 °C), and the pellets were frozen at
80 °C before
purification. Cell pellets were thawed at 4 °C, resuspended in
buffer A (50 mM NaCl, 50 mM Tris-HCl (pH 7.4),
1 mM EDTA), and phenylmethylsulfonyl fluoride was added to
a final concentration of 50 µM. Cells were lysed by
sonication, and the resulting solution was centrifuged at 4 °C for
10 min at 13,000 × g. The supernatant was applied to a
MonoS HR 5/5 FPLC column (Amersham Pharmacia Biotech) equilibrated in
buffer A, which was eluted with gradient from 50 to 1,000 mM NaCl in 50 mM Tris-HCl, pH 7.4. Elution was
monitored by A280. A peak containing Phd and Doc
eluted at approximately 600 mM NaCl. This peak was applied
to a semi-preparative reverse-phase C18 column, and Phd and
Doc were separated using a gradient from 0 to 80% acetonitrile in
0.1% trifluoracetic acid. Following this step, the proteins were
greater than 95% pure as assessed by Coomassie staining of
SDS-polyacrylamide gels.
Characterization of Purified Protein--
The masses of the
purified proteins were determined by
MALDI-TOF1 mass spectrometry
using a Voyager-DE STR Biospectrometry Workstation (PerSeptive
BioSystems) with sinapinic acid (3,5-dimethoxy-4-hydroxycinnamic acid)
as the matrix, linear mode operation, a grid voltage at 91.0%, and a
pulse delay time of 150 ns. Protein concentrations were determined by
measuring tyrosine absorbance at both pH 7 and in 0.1 M KOH
using a Hewlett Packard 8452A diode array spectrophotometer with
extinction coefficients per tyrosine of 1,394 M
1 cm
1 (274 nm at pH 7) and
2377 M
1 cm
1 (294 nm in 0.1 M KOH) for Phd (1 Tyr) and Doc (4 Tyr). Circular dichroism
spectra were obtained using an AVIV 60DS spectrapolarimeter equipped
with a temperature-controlled sample holder and a 10-mm pathlength
cuvette. Thermal denaturation experiments were performed in a buffer
containing 50 mM Tris-HCl (pH 7.4), 100 mM
NaCl, and 0.1 mM EDTA; samples were equilibrated at 1 °C
temperature intervals for 1 min, and the ellipticity at 222 nm was
averaged for 2 min.
Fluorescence Labeling and Spectroscopy--
Purified Phd protein
was reacted with 2 equivalents of carboxytetramethylrhodamine
succinimidyl ester (Molecular Probes, Eugene OR) in 0.5 M
HEPES (pH 7), 100 mM NaCl, for 24 h at room
temperature in the dark (15). Doc was labeled in the same way except
using carboxyfluorescein succinimidyl ester. Unreacted dyes were
separated from labeled protein by gel filtration chromatography on a
QuickSpin G-25 Sephadex column (Roche Molecular Biochemicals).
Equilibrium binding, assayed by fluorescence resonance energy transfer,
was performed in 50 mM Tris-HCl (pH 7.4), 100 mM NaCl, and 1 mM EDTA using a FluoroMax2
fluorescence spectrophotometer (Jobin Yvon
Spex, Edison, NJ).
Kinetic experiments were performed using an Applied Photophysics
DX17.MV stopped-flow instrument and were monitored by changes in
fluorescence using a bandpass filter (520 ± 10 nm).
Crystallization and Analysis of the Phd·Doc
Complex--
Crystals of a Phd·Doc complex grew after several days
at 4 °C from solutions containing purified protein at concentrations of 5 mg/ml or higher in 50 mM Tris·Cl (pH 8.0), 500 mM NaCl, and 1 mM EDTA. Individual crystals
were thoroughly washed, boiled in SDS sample buffer, and analyzed by
SDS-polyacrylamide gel electrophoresis, confirming the presence of both
Phd and Doc in the crystals at 99% or higher purity. Amino acid
analysis of washed and redisolved crystals was performed by the MIT
Biopolymers laboratory.
Ultracentrifugation and Gel Filtration--
Analytical
ultracentrifugation experiments were performed using a Beckman Optima
XL-A centrifuge. Solutions containing Phd and Doc from dissolved
crystals were centrifuged overnight at 15,000 rpm. After reaching
equilibrium (established by unchanged A274
readings at 1 h intervals), absorbance profiles were determined at
274 nm, and five scans were averaged for analysis. Data from experiments in which ln(A) versus r2
plots showed curvature were fitted by nonlinear least-squares methods
to a 2P + D
P2D equilbrium model using Eq. 1.
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(Eq. 1)
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where Cr is the total concentration and the
terms Cb,x represent the concentrations of each
species at a reference radial position. Ax = (1
X
)
2/2RT, where
X
is the partial specific volume of each species (calculated from its
sequence),
is the solution density,
is the rotor angular
velocity in radians per second, R is the gas constant, and
T is the absolute temperature. Kd is the
equilibrium dissociation constant, Mx is the
molecular mass of each species, r is the radial position,
rb is a reference radial position, and
is a
small baseline correction term. This equation takes the relative
extinction coefficients of Phd (one), Doc (four), and the
P2D complex (six) into account.
Gel filtration experiments were performed by chromatographing 200-µl
samples on a 10 × 300 mm Superdex 75 HR 10/30 column (separation
range 3-70 kDa) using a flow rate of 0.5 ml/min. Absorbance was
monitored at 280 nm, and the column was calibrated using gel filtration
molecular weight standards (Bio-Rad).
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RESULTS |
Copurification of Phd and Doc--
As a first step in studying
potential interactions between Phd and Doc, both proteins were purified
from E. coli strain X90/pRDM032, a cell expressing the phage
P1 addiction proteins under Ptac promoter control
(4). As shown in Fig. 1A, Doc
protein and the major portion of the Phd protein coeluted during
ion-exchange chromatography on a MonoS column; a smaller quantity of
Phd alone also eluted at a lower salt concentration than the Phd·Doc
complex. During purification from cells without Doc, Phd eluted only in the low salt position. Further purification by reverse-phase HPLC chromatography under denaturing conditions separated Doc from Phd and
resulted in purification of each protein to greater than 95%
homogeneity (Fig. 1B). The identity of each protein was
confirmed by MALDI-TOF mass spectrometry (Fig.
2). The observed masses (8,130 Da for
Phd; 13,578 Da for Doc) are within 1 or 2 Da of those expected from the
protein sequences, indicating that the formyl group of the N-terminal
fMet residue is removed post-translationally for each protein.

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Fig. 1.
Purification. A, Phd and Doc
coelute during chromatography on a MonoS ion-exchange column developed
with a gradient of NaCl; B, separation of Phd and Doc on a
C18 HPLC reverse-phase column developed with a gradient of
acetonitrile.
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Fig. 2.
MALDI-TOF mass spectrometry.
HPLC-purified proteins have masses within 2 Da of those expected from
the unmodified Phd (8,128 Da) and Doc (13,579 Da) protein
sequences.
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Phd and Doc Form a 2:1 Complex in Crystals--
Solutions
containing high concentrations of purified Phd and Doc formed
needle-shaped crystals that grew to a length of several millimeters
after 24-48 h at 4 °C (Fig.
3A). Following washing, SDS-polyacrylamide gel electrophoresis showed that individual crystals
contained both Phd and Doc (Fig. 3B). Densitometry of the
Coomassie Blue-stained gel indicated that the Phd and Doc proteins were
present in the crystal in a 2:1 molar ratio (Fig. 3C). Amino
acid analysis of protein from washed crystals also supported a 2:1
molar ratio of Phd:Doc in the crystals (Table I). Hence, in the crystalline state, Phd
and Doc appear to form a macromolecular complex containing twice as
many Phd molecules as Doc molecules. The smallest oligomer consistent
with this stoichiometry is a P2D trimer, but
P4D2 hexamers, P6D3
nonamers, et cetera are also consistent with these data.

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Fig. 3.
Crystals of Phd·Doc complex.
A, photograph of crystals; B, SDS-polyacrylamide
gel electrophoresis of washed and dissolved crystals shows presence of
both Phd and Doc; C, densitometry of the Coomassie
Blue-stained polyacrylamide gel is most consistent with a 2:1 ratio of
Phd:Doc.
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Oligomeric State in Solution--
In gel filtration chromatography
on Superdex 75, purified Doc eluted at an approximate volume expected
for a monomer (data not shown); Phd had been previously shown to be
monomeric in solution (6). To determine the solution form of Phd·Doc
complexes, protein from dissolved cocrystals was also chromatographed
on Superdex 75. At high loading concentrations (80 µM in
Doc monomer equivalents), a major peak of approximately 30 kDa was
observed (Fig. 4A). This value
is consistent with formation of a P2D solution trimer
(expected molecular mass, 29.8 kDa). Some trailing asymmetry was
evident in the Fig. 4A column profile, suggesting that a
small fraction of P2D complexes dissociated during
chromatography. Indeed, when the sample was diluted 10-fold immediately
before chromatography (loading concentration 8 µM in Doc
monomer equivalents), less of the P2D peak was present and
more prominent, lower molecular weight peaks corresponding to Doc and
Phd alone were observed (data not shown). These experiments suggest
that Phd·Doc complexes dissociate in the micromolar concentration
range with kinetics on the time scale of minutes or faster. Gel
filtration of a mixture of Phd:Doc in a 4:1 ratio revealed no species
larger than 30 kDa but showed significant free Phd (data not shown).
These results show that the P2D complex is the major
solution form, even in the presence of excess Phd.

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Fig. 4.
Oligomeric state. A, the
Phd·Doc complex chromatographs at an apparent molecular mass of 30 kDa in gel filtration of dissolved cocrystals (80 µM in
Doc equivalents) on a Superdex 75 column. The elution positions of
ovalbumin (44 kDa) and myoglobin (17 kDa) are indicated. B,
equilibrium distribution of Phd·Doc complex absorbance (loading
concentration 80 µM in Doc equivalents) during analytical
centrifugation at 15,000 rpm, 20 °C. Calculated distributions for
P2D complexes (solid line),
P4D2 complexes (dashed line), and PD
complexes (dashed-dotted line) are shown. C,
equilibrium distribution of Phd·Doc complex absorbance (loading
concentration 20 µM in Doc equivalents) during analytical
centrifugation at 15,000 rpm, 20 °C. The solid line is calculated
for the reaction 2P + D P2D with a
Kd of 0.96 µM2.
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As a second assay of solution oligomeric form, protein from dissolved
Phd·Doc cocrystals (80 µM in Doc equivalents) was
analyzed by analytical ultracentrifugation. These data were fit best by a 2:1 ratio of Phd:Doc and showed significant deviations from the
behavior expected for 1:1 or 2:4 complexes of Phd:Doc (Fig. 4B). Equilibrium centrifugation experiments performed using
lower protein concentrations (loading concentration of 20 µM in Doc equivalents) showed curvature in the
ln(A274) versus
r2 plot, indicative of some complex
dissociation. Fitting of this data to a 2P + D
P2D
model gave an equilibrium dissociation constant of 1.0 ± 0.3 µM2 at 20 °C (Fig. 4C).
Changes in Secondary Structure Induced by Complex
Formation--
The circular dichroism (CD) spectrum of purified Doc
had features expected for a protein containing approximately 50%
-helix (Fig. 5A). Melting
experiments in a neutral buffer at protein concentrations of either 1 or 15 µM had similar shapes and midpoints (Fig.
5B), as expected if Doc is monomeric. The native structure of Doc (Tm 60 °C) is quite stable in comparison
to Phd (Tm 25 °C) (6). In fact, Phd is almost
completely denatured at 37 °C, whereas Doc is native at this
temperature.

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Fig. 5.
Doc secondary structure and stability.
A, far ultraviolet CD spectrum of purified Doc (5 µM) at 4 °C suggests an -helical content of
approximately 50%. B, Doc at concentrations of 1 and 15 µM undergoes cooperative thermal denaturation with a
Tm of 60 °C as monitored by changes in CD
ellipticity at 222 nm. The CD signals for the two protein
concentrations were normalized to allow comparison. The buffer for the
experiments shown in panels A and B was 50 mM Tris-HCl (pH 7.4), 100 mM NaCl, and 0.1 mM EDTA.
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By comparing the CD spectra of a mixture of 2 µM Phd and
1 µM Doc with the summed spectra of the uncomplexed
proteins, secondary structural changes induced by complex formation
were assessed. At 4 °C, where Phd and Doc by themselves are both
native, the mixture had slightly less ellipticity than the isolated
proteins, suggesting that complex formation leads to some change in
structure (Fig. 6A). At
37 °C, where Phd by itself is primarily unfolded, the
P2D complex had greater ellipticity than the individual
proteins, suggesting an increase in secondary structure upon binding
(Fig. 6B). In the latter instance, it seems probable that at
least a portion of Phd folds as it binds to Doc. Fitting of titration experiments, monitored by changes in CD ellipticity at 37 °C, gave
an equilibrium dissociation constant of 0.7 ± 0.1 µM2 (Fig. 6C).

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Fig. 6.
Changes in CD ellipticity accompany complex
formation. A, CD spectra, at 4 °C, of 1 µM Doc and 2 µM Phd before and after mixing
in a tandem cuvette; B, mixing experiment at 37 °C. Other
conditions identical to panel A. C, increasing
quantities of Doc were added to 2 µM Phd protein and the
change in CD ellipticity at 222 nm was monitored. The solid
line is calculated for the reaction 2P + D P2D
with a Kd of 0.72 µM2. The
buffer for all experiments was 50 mM Tris-HCl (pH 7.4), 100 mM NaCl, and 0.1 mM EDTA.
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Affinity and Kinetics of Complex Formation--
To study the
affinity of complex formation by another method, we performed
fluorescence resonance energy transfer experiments by titrating
increasing amounts of rhodamine-labeled Doc (the acceptor molecule)
against a constant amount of fluorescein-labeled Phd (the donor
molecule). Because energy transfer depends on the inverse sixth power
of the distance between the fluorescent dyes, any change in
fluorescence should arise from interactions between the proteins. Fig.
7A shows fluorescence spectra,
and Fig. 7B shows the change in fluorescence at 25 °C as
a function of Phd concentration. These experimental data were fit best
by a binding curve calculated for a model
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(Eq. 2)
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that is second-order in Phd and first-order in Doc, with an
equilibrium dissociation constant of 0.83 ± 0.02 µM2.

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Fig. 7.
Resonance energy transfer. A,
fluorescence spectra of rhodamine-labeled Phd (0.8 µM),
fluorescein-labeled Doc (0.02 µM), and a mixture of both
proteins; B, complex formation assayed by resonance energy
transfer. The solid line is calculated for the reaction 2P + D P2D with a Kd of 0.81 µM2. C, kinetics of dissociation
of complexes containing rhodamine-labeled Phd and fluorescein-labeled
Doc monitored by fluorescence after addition of a 250-fold excess of
unlabeled Phd. The solid line is a single exponential fit
with a rate constant of 42 s 1. Experiments in all panels
were performed at 25 °C in a buffer containing 50 mM
Tris-HCl (pH 7.4), 100 mM NaCl, and 1 mM
EDTA.
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To determine the half-life of complex dissociation, fluorescently
labeled P2D complexes were mixed with excess unlabeled Phd in a stopped-flow instrument, and the change in fluorescence was monitored (Fig. 7C). The rate constant of this exchange
process, which should be limited by dissociation was 42 (± 7)
s
1 indicating that complex dissociation must occur very
rapidly. Similar results were obtained when the dissociation reaction
was initiated by mixing fluorescently labeled P2D complexes
with unlabeled Doc. The rapid dissociation rate is consistent with the
tailing seen in gel filtration. From the equilibrium constant and a
dissociation rate constant, a third-order association rate constant of
approximately 6·1013 M
2
s
1 can be calculated.
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DISCUSSION |
Efficient inhibition of the toxic activity of Doc by the Phd
antidote is essential for maintaining the viability of bacteriophage P1
lysogens of E. coli (1) but little concrete information concerning the molecular mechanism of this inhibition has been available. The results presented here show that Phd and Doc interact directly to form a heterotrimeric complex in solution. In the absence
of more detailed information, it is not possible to know whether
formation of the Phd·Doc complex blocks the toxic activity of Doc by
sterically occluding interactions with its cellular target or if
formation of the complex alters the structure of Doc in a way that
prevents this interaction. It is intriguing, however, that formation of
the Phd·Doc complex appears to result in a small change in secondary
structure. Were this alteration of structure to involve a portion of
Doc, then this could serve as the mechanism of toxin neutralization.
The Phd·Doc complex is a P2D heterotrimer. Heterotrimers
are not common in biological systems and must be assembled
asymmetrically. A well known example with a 2:1 ratio of components is
the heterotrimer of growth hormone with two extracellular receptor
domains, in which each receptor binds to a distinct surface of growth
hormone (16). The oligomeric form of the antidote-toxin complex for the
ccd system is hexameric with two dimers of the toxin and one dimer of the antidote (14), but this architecture also requires asymmetric construction and, indeed, can be considered as a
heterotrimer of dimers. We find it intriguing that there are 2:1 or 1:2
ratios of antidote:toxin in both the P1 addiction system and F
ccd system. Whether these and other addiction systems share
common structural features is not known.
Because simultaneous three-body collisions between protein monomers of
Phd and Doc is not a statistically viable mechanism of assembly,
formation of the P2D complex must proceed either through a
P2 or a PD intermediate. Neither of these dimeric species has been detected in solution, implying that if either species can form
then it must be unstable and poorly populated at equilibrium. We note,
however, that two monomers of Phd bind cooperatively to operator
subsites, suggesting that Phd subunits interact with each other in the
DNA-bound state (6). It is also worth noting that the P2D
solution complex may also be an active operator binding species; using
a mutant form of Doc, Magnuson and Yarmolinsky (5) have suggested that
a 2:1 ratio of Phd:Doc night occur in operator complexes.
The P2D complex of Phd and Doc is unstable by comparison
with many macromolecular complexes. Kd values for
complex dissociation of 0.7-1.0 µM2 were
obtained from analytical ultracentrifugation at 20 °C, from fluorescence resonance energy transfer at 25 °C, and from changes in
CD ellipticity at 37 °C. Dissociation of unlabeled Phd·Doc complexes to free components was also observed in the micromolar concentration range by gel filtration. P2D complexes
equilibrate with free subunits on a subsecond time scale in
vitro, which should ensure rapid equilibration of the bound and
free intracellular pools of Phd and Doc.
There is an approximate 3-fold ratio of Phd to Doc in overproducing
cells. Because the phd and doc genes are part of
the same operon, a similar ratio would be expected in P1 lysogens.
Moreover, because Phd represses its own synthesis and that of Doc
(1-5), P1 lysogens are unlikely to contain very high concentrations of either protein. To ensure 90-99.9% binding of Doc with a
Kd of 0.7-1.0 µM2, we
calculate that the free Phd concentration would need to be from 2.2 to
36 µM (
2,200 to 36,000 molecules/cell). However, under
these conditions there would still be free Doc concentrations of
260-22 nM. Hence, it is important to view the Phd-Doc
interaction as buffering the free concentration of Doc rather than
eliminating free Doc from the cell. If the assumptions that underlie
these rough calculations are sound, then free Doc at submicromolar
levels must be relatively innocuous to cells. Otherwise, P1 lysogens would have a significant growth disadvantage relative to nonlysogens. However, higher levels of free Doc (presumably in the micromolar range)
must be lethal for the plasmid addiction mechanism to function. These
considerations make it extremely unlikely that the mechanism of Doc
toxicity involves single hit enzymatic lethality and suggest that Doc
may reversibly inhibit some required cellular activity in a cooperative
fashion that results in a steep concentration dependence of toxicity.
In P1 lysogens, Phd binding spares the cell from Doc-mediated killing
(3). However, Doc binding induces structure in denatured Phd at
physiological temperatures, and thus it seems likely that Doc binding
would also help protect Phd from degradation. Degradation of Phd is
mediated directly or indirectly by the ClpXP protease (3), which
consists of ClpP protease subunits and regulatory ClpX ATPase subunits
(17-19). ClpX is responsible for substrate recognition by ClpXP and
has been shown to recognize specific peptide sequences at the C termini
of several substrate proteins (20-22). If ClpXP directly degrades Phd,
then C-terminal substrate recognition is probably also involved. By
this model, formation of the P2D complex with Doc could
prevent degradation of Phd by sequestering C-terminal sequences
required for ClpX recognition and perhaps by stabilizing at least some
parts of Phd in a native structure. We note, however, that the rapid
dissociation rate and relatively high equilibrium dissociation constant
for P2D complexes makes it likely that a pool of free Phd,
in a largely denatured form, is present even when Doc is present in P1
lysogens. If Phd were too good a substrate for ClpXP, then degradation
of the free pool and mass action to produce more free Phd could lead to
Doc toxicity even in lysogens. Hence, it seems likely that Phd should
be a rather poor substrate for ClpXP. Indeed, in ClpXP+
strains, Phd is degraded relatively slowly over the course of several
cell generations (3)