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(Received for publication, August 12, 1996, and in revised form, October 2, 1996)
From the Department of Molecular Biology and Biochemistry,
University of California, Irvine, California 92697
Binding of cAMP receptor protein (CRP) and CytR
mediates both positive and negative control of transcription from
Escherichia coli deoP2. Transcription is activated by CRP
and repressed by a multi-protein CRP·CytR·CRP complex. The latter
is stabilized by cooperative interactions between CRP and CytR. Similar
interactions at the other transcriptional units of the CytR regulon
coordinate expression of the transport proteins and enzymes required
for nucleoside catabolism. A fundamental question in both prokaryotic and eukaryotic gene regulation is how combinatorial mechanisms of this
sort regulate differential expression. To understand the combinatorial
control mechanism at deoP2, we have used quantitative footprint and gel shift analysis of CRP and CytR binding to evaluate the distribution of ligation states. By comparison to distributions for
other CytR-regulated promoters, we hope to understand the roles of
individual states in differential gene expression. The results indicate
that CytR binds specifically to multiple sites at deoP2,
including both the well recognized CytR site flanked by CRP1 and CRP2
and also sites coincident with CRP1 and CRP2. Binding to these multiple
sites yields both cooperative and competitive interactions between CytR
and CRP. Based on these findings we propose that CytR functions as a
differential modulator of CRP1 versus CRP2-mediated
activation. Additional high affinity specific sites are located at
deoP1 and near the middle of the 600-base pair sequence
separating P1 and P2. Evaluation of the DNA sequence requirement for
specific CytR binding suggests that a limited array of contiguous and
overlapping CytR sites exists at deoP2. Similar extended
arrays, but with different arrangements of overlapping CytR and CRP
sites, are found at the other CytR-regulated promoters. We propose that
competition and cooperativity in CytR and CRP binding are important to
differential regulation of these promoters.
In Escherichia coli, the enzymes and transport proteins
required for nucleoside catabolism and recycling are encoded by genes belonging to the CytR regulon. This gene family consists of at least
nine unlinked transcriptional units (for review, see Ref. 1).
Expression of these transcriptional units is coordinately regulated by
the interplay of two transcriptional regulatory proteins, CRP1 (also referred to as CAP) and the CytR
repressor. Transcription is activated in response to intracellular cAMP
levels by CRP, repressed by CytR, and induced by cytidine. A few of the
transcriptional units are also separately regulated by a second
repressor, DeoR (2, 3, 4), via an independent mechanism.
A key feature of the CytR regulon is that the individual cistrons are
differentially expressed. Extents of activation, repression, and
induction all vary among the different transcription units (cf. Ref. 5). This is achieved by nesting levels of local
repression, mediated by DeoR and CytR, on a more global regulation
mediated by CRP. This illustrates a process, common to both E. coli and higher order eukaryotes, in which complex patterns of
expression are controlled using a small number of regulatory proteins.
For example regulation of cell growth and differentiation often
combines tissue-specific or developmental stage-specific factors with
more global control elements. The mechanism of such broad regulatory programs is a fundamental issue in gene regulation. Presumably combinatorial mechanisms that rely on different local features of
different genes are involved.
Several of the CytR-regulated operons have been investigated at the
molecular level. The promoter deoP2 of the tetracistronic operon that directs the synthesis of purine and pyrimidine
phosphorylases and enzymes required for sugar utilization has generated
the greatest interest (6). Others that have been investigated include
tsx (7), encoding an outer membrane protein, cdd
(8), encoding cytidine deaminase, udp (9) encoding uridine
phosphorylase, cytr (10) encoding CytR, and most recently
nupG (11) encoding a membrane nucleoside transport protein.
In all cases, these studies have implicated interactions of CRP and
CytR with sequences located in the 80-100 bp immediately upstream of
the various transcription start sites and interactions between the
proteins as the basis for positive and negative gene regulation. Thus,
differential and coordinate gene regulation must depend on different
dispositions of CRP and CytR binding sites, different protein binding
affinities, and/or different levels of site-site interaction or
cooperativity.
Most CytR-regulated promoters contain tandem CRP sites. CRP1 (at about
The tandem CRP sites in different promoters are separated by DNA of
variable length and sequence. Based on footprinting analysis of
purified CytR binding to deoP2 and other promoters, this
intervening sequence is now thought to contain the CytR binding site
(6, 8, 16). A putative recognition motif has been identified (6, 17),
and pairs of such motifs, arranged as either direct or inverted
repeats, have been implicated as the CytR operator (9, 17). It is
ironic, given the results we report here, that until relatively
recently CRP and CytR were believed to compete to bind to the same
sites (cf. Ref. 4).
The mechanism of CytR-mediated repression is indirect. CytR has no
effect on basal level transcription but instead requires CRP binding to
mediate repression (16, 18). Of course, this necessarily means that
CytR functions only under conditions of CRP-dependent
activation. Under these circumstances CytR typically does not
completely reduce the activated level of expression to the basal level
(5). Despite this role of CytR as a functional antagonist of CRP, the
two proteins interact cooperatively, resulting in substantially
increased CytR binding affinity when CRP is present (6). The role of
cooperativity is widely thought to be to recruit CytR at otherwise
sub-saturating concentrations. The crucial role that cooperativity
plays in repression is highlighted by the fact that when CytR binds
cytidine, induction occurs as a result of the loss of cooperativity and
despite no effect of cytidine binding to CytR on intrinsic binding of
CytR to DNA (6).2
The complexity of these regulatory properties has generated confusion
about the nature of key molecular interactions. The most perplexing
questions involve the CRP-CytR cooperativity. For example, heterologous
cooperativity has been reported to require both CRP sites, from which a
lack of pairwise interactions is inferred (20). Yet, regulation of
cytRP, in which there is only one CRP site, also depends on
cooperative CytR and CRP binding (10). In addition, it was recently
observed (11, 21) that the apparent cooperativity, when assessed by the
effect of saturating CRP concentration on CytR affinity, is
substantially greater than when cooperativity is assessed by the effect
of saturating CytR concentration on CRP binding. This "one-way
stimulation" (11) represents an apparent conflict with the laws of
thermodynamics that still awaits a molecular explanation.
Several lines of evidence implicate protein-protein interactions as
providing the driving force for cooperativity. First, mutations have
been located on the surface of CRP in a putative protein interacting
domain that interfere with cooperativity with CytR and with
CytR-mediated repression (22). Second, CytR and CRP are reported to
mutually antagonize each other's protein binding induced bends in the
cytRP sequence (10). Since such coupled DNA structural
transitions necessarily contribute unfavorably to cooperativity, the
driving force for cooperativity in cytRP must be derived
from favorable protein-protein contacts. Third, a truncated CytR which
is lacking the DNA-binding helix-turn-helix motif was reported to bind
to deoP2 in the presence of CRP and further to do so with
only moderate reduction in overall affinity (23) compared with the
full-length protein. This would suggest that CytR is primarily a
protein-protein bridge, dependent on its interactions with CRP bound to
the flanking CRP sites for association with its operator. However, the
question remains open how even these favorable protein-protein contacts
could possibly compensate for the loss of a direct DNA binding
interaction with a Kd in the 10 nM
range.
Previous approaches used to investigate CytR-regulated promoters have
not been fully successful in developing an understanding of the
macromolecular interactions that regulate transcription. The confusion
stems in large part from the fact that only qualitative reasoning has
been used to address quantitative questions. For example, most of the
in vitro studies have been conducted as combinations of
plus/minus the various components (CRP, CytR, cAMP, cytidine, and
promoter elements) with insufficient understanding of the mutual
effects of interactions between these molecules to know what
concentrations are necessary to achieve particular effects. Our
immediate goal was to understand how the protein-DNA and
protein-protein interactions control the distribution of operator
configurations at deoP2. Our approach has been to use DNase
footprinting to obtain complete individual site isotherms for binding
of CytR and CRP to deoP2, at different configurations of
empty and filled sites for the other regulatory protein. From such
data, the complete population distribution of promoter configurations
can be determined as a function of both CRP and CytR concentration. We
anticipated being able to deduce connections between the individual
promoter configurations in the distribution and biologically functional states.
The results indicate a much more complex promoter structure than was
previously supposed. Multiple, specific CytR sites are arranged over an
extended region of DNA that includes both CRP1 and CRP2 as well as the
sequence they flank. CytR binding to these sites mediates both
cooperative and competitive interactions with CRP. Together these
explain quantitatively the apparent one-way stimulation. Similar
specific interactions of CytR with extended DNA sequences have been
identified at deoP1 and also near the middle of the
approximately 600-bp sequence separating P1 and P2. Comparison of the
DNA sequences of these sites further clarifies the CytR binding motif.
The distribution of such motifs, both at deoP2 and at other
CytR-regulated promoters, define arrays of contiguous and overlapping
CytR binding sites. This entirely new phenomenon is consistent with the
interpretation that CytR functions as a modulator of CRP-mediated
activation. We envision such a function operating at two levels: first
as a differential modulator of class I versus class II CRP
activation at individual promoters, and second as a differential
modulator of activation of the various CytR cistrons.
Crystalline adenosine 3 CRP was expressed from E. coli strain K12 CytR was expressed and purified as described.2 On
SDS-polyacrylamide gels, the purified material used in these studies
was at least 90% full-length CytR (Mr = 37,800). The remaining material was contained in two bands, with
apparent molecular weights of 31,000 and 27,000. This has comprised
from 5 to 20% of the total material in different CytR preparations.
Under native conditions, purified CytR elutes from a Pharmacia Superose
6 column in a single, sharp peak with apparent molecular weight
72,500 ± 2,500. This is consistent with sedimentation equilibrium
analysis that indicates CytR to be homogeneous dimer in
solution.3 This peak accounts for all
observable UV absorbing material. Thus, the lower molecular weight
bands appear to be products of endogenous proteases, as has been
observed with other members of the LacR repressor family (29, 30). We
conclude on this basis that the CytR is at least 95% pure.
Concentration was estimated using an extinction coefficient, Fig. 1 shows the deo
DNA fragments used. Plasmid pSS13322 contains the deo P1/P2
sequence from
Mutant promoters were generated in which site-specific CRP binding to
CRP1 (CRP1 Quantitative DNase I
footprint titrations were conducted as described (32, 34) in 10 mM bis-tris (pH 7.00 ± 0.01), 100 mM
NaCl, 0.5 mM MgCl2, 0.5 mM
CaCl2, 50 µg/ml bovine serum albumin, and 1 µg/ml calf
thymus-DNA. Binding reaction mixtures (200 µl) were equilibrated in a
water bath at 20 °C (±0.2 °C) for between 40 min and 2 h
prior to DNase I exposure. Measurements are independent of the
incubation time over this range. These were exposed to 2-6 ng of DNase
I, added in a 5.0-µl volume, for 12.0 min, and quenched by addition
of (null)/1;5 volume of 50 mM Na4EDTA before addition of stop solution (34). Two-dimensional optical scanning of
footprint titration autoradiograms and analysis of the digitized images
was as described (34).
Mobility-shift titrations were
conducted as described (35, 36) using 5% acrylamide gels (29:1,
acrylamide:bis) and 0.5 × TBE electrophoresis buffer (31, 33).
CytR and deoP2 DNA (10 pM; 285-bp fragment) were
incubated (40-60 min) at 20 °C (±0.1 °C) in the DNase I
footprint binding buffer but with 2 µg/ml CT-DNA and with 1.5%
Ficoll added to facilitate gel loading. Aliquots (20 µl) of
equilibrated binding reaction mixtures containing 1400 dpm of
32P were loaded onto 1.5-mm minigels in a Bio-Rad Mini
Protean II device that had been pre-electrophoresed for 5 min. Gels
were loaded with current on and electrophoresed at a constant 200 V for
35 min.
Dried gels were imaged using a Molecular Dynamics PhosphorImager 435SI.
Phosphor plates were exposed for 8-10 h and scanned at 176-µm
spatial resolution. Analysis of the digital images was conducted using
the program IPLabGel (Signal Analytics Corp.) or ImageQuant (Molecular
Dynamics, Inc.) essentially as described (35, 36). The combination of
long exposure and high specific radioactivity yielded ratios of average
pixel intensity to background of about 100, minimizing concerns about
local background variation (24, 32).
Binding data were analyzed by using the
nonlinear least squares parameter estimation program, NONLN (37). NONLN
estimates parameter values corresponding to a minimum in the variance,
and worst case joint confidence limits for each parameter corresponding to approximately one standard deviation. Simple, noncooperative binding
of a single protein to an individual DNA site is described by
Volume 271, Number 52,
Issue of December 27, 1996
pp. 33242-33255
©1996 by The American Society for Biochemistry and Molecular Biology, Inc.

41.5 bp from transcription start sites) and CRP2 (at about
92.5 bp)
appear to be class II and class I sites, respectively (12, 13). The
significance of these classifications is the suggestion that they
direct different kinetic mechanisms of activation. At class I sites,
CRP is proposed to increase the apparent affinity of RNAP for the
promoter, whereas at class II sites, CRP is proposed to increase the
rate of formation of the open transcription complex. At
deoP2, CRP1 alone, but not CRP2 alone, substantially
activates transcription (5). CRP1 has been proposed to substitute for the lack of a
35 consensus promoter sequence (14), whereas CRP2 is
proposed to be necessary only for CytR to bind. However, in the absence
of kinetic studies on any of these promoters there is little direct
evidence to suggest which kinetic mechanism of activation is involved
under any condition. The highest levels of expression are achieved with
both sites functional, suggesting some synergy in activation (15).
Reagents and Enzymes
-5
cyclic
monophosphate (cAMP) and crystalline cytidine (both >99%) were
purchased as free base from Sigma and as free acid
from ICN, respectively. Stock concentrations (in 50 mM
bis-tris base, pH 7.0, 1 mM EDTA) were determined, and purity was assessed spectrophotometrically by comparing observed spectra to published molar extinction coefficients and absorbance ratios (61). Bovine pancreas deoxyribonuclease I (DNase I, code D) from
Worthington was treated as described (24). [
-32P]dNTPs
(3000 Ci/mmol) were purchased from Amersham Corp. or ICN; unlabeled
dNTPs were from Life Technologies, Inc. Buffer components and reagents
were electrophoresis grade if available and reagent grade
otherwise.
H
trp transformed with the expression plasmid
pPLcCRP1 (25) and purified as described (26). No contamination is
detectable by Coomassie staining of overloaded SDS-polyacrylamide gels
from which we estimate at least 98% purity. CRP concentration was
estimated based on
(1%) = 9.2 at
max = 278 nm
(27).
= 0.30 ± 0.02 mg
1ml
1 at 280 nm.2
801 to +151 relative to the P2 start site for
transcription, cloned into the BamHI site of pUC13.
Insertion of an 8-bp NotI linker into the SmaI
site of the vector generated an 879-bp
NotI/HincII fragment in which the 32P-labeled NotI end is 192 bp downstream from
CRP1. A 285-bp NotI/SmaI fragment containing only
the P2 regulatory region was generated by inserting a NotI
linker into a BsmI site at
117 in the deo sequence. All DNA fragments were agarose gel purified after banding the
plasmid preparations twice in CsCl gradients. DNA was protein free, as
determined from A260/A280
(31). Fragments were labeled at their NotI sites using the
Klenow fill-in reaction as described (32).
Fig. 1.
Schematic of deo showing the
regulatory region used in these studies. A, complete operon
with promoters, P1 and P2. Enlarged view shows the deo
regulatory sequence cloned to make pSS1332. deo sequences
indicated by heavy lines; vector sequences and added linkers
by light lines. Filled circles, shaded
boxes, and the open hexagon indicate the known binding
sites for the transcriptional regulatory proteins, DeoR, CRP, and CytR,
respectively. P1 and P2 transcription start sites are indicated by
arrows. Coordinates are in bp numbered from +1 at the P2
start site. B, linear fragments used in binding studies
isolated from pSS1332 derivatives pLP03 and pSS1332-NotI,
respectively, as described in the text. C, enlargement of
CRP-1 and CRP-2 showing G/C to A/T transitions used to create reduced
CRP valency mutants CRP1
(pLP01) and CRP-2
(pLP02).
[View Larger Version of this Image (27K GIF file)]
; pLP01) or to CRP2 (CRP2
; pLP02)
was eliminated. The BamHI fragment from pSS1332 was
subcloned into pM13mp8, and single-stranded DNA was isolated as
described (33). Site-directed mutagenesis was conducted using the kit from Amersham Corp. Mutagenic oligonucleotides, 30 nucleotides in
length, were designed to produce symmetric G to A transitions in both
TGTGA, CRP recognition motifs of the mutated site. Sequences of the
mutants (Fig. 1) were confirmed by dideoxy DNA
sequencing. CRP1
and CRP2
operator
fragments (879 bp) were isolated as described.
where L is the concentration of free protein ligand, and
ki is the intrinsic association constant for binding
of the protein to the individual site. Binding of either CRP or CytR alone was analyzed to obtain the Gibbs free energy change corresponding to ki in Equation 1 (
(Eq. 1)
Gi =
RT lnki). In the analysis of footprint
titration data, where fractional protection rather than fractional
saturation is the quantity experimentally determined, it was also
necessary to fit the fractional protection end points as adjustable
parameters for each separate titration (24).
Fitting to Equation 1 was also used to estimate individual site loading
free energy changes
Gl,i (38) for binding experiments in which both CytR and CRP were present.
Gl,i is related to the integral of an individual
site binding curve and reflects the sum of free energy changes for both
intrinsic binding and the effect of all cooperative interactions. For
noncooperative binding,
Gl,i equals
Gi (Equation 1). Equation 1 can also provide an
accurate estimate of
Gl,i and of its confidence
limits when two different proteins interact cooperatively in binding to
different DNA sites, and when binding of one is titrated while the
concentration of the second is held constant (39). It is necessary that
the concentration held constant be saturating. In this limit, the
probability that the protein being titrated will bind to DNA that is
already liganded by the first protein is made arbitrarily close to
unity, and the shape of the binding curve is described by Equation 1.
Equations that describe the cooperative binding of CRP and CytR were derived by considering the relative probability of each of the deoP2 configurations (Table III) as specified by the Specific, and Nonspecific Additional Sites models that are introduced in the text. The probabilities are given by
|
(Eq. 2) |
Gs is the sum of free energy
contributions for configuration s (Table III), and where
i and j are the stoichiometries of bound
CRP(cAMP)1 complexes and CytR dimers in configuration s. The binding equation for any individual site is derived
by summing the relative probabilities for all configurations with protein bound to the site. For reduced valency, CRP1
and
CRP2
operators, configurations in which
CRP(cAMP)1 is bound to the mutated site were excluded from
the summation. Intrinsic binding of CytR was assumed to be unaffected
by base pair substitutions in CRP1 and CRP2.
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Global analysis of individual site CRP(cAMP)1 and CytR binding was conducted essentially as described (40). Each binding curve was first analyzed separately using Equation 1 to calculate normalized weighting factors for use in global analysis. Goodness of fit and internal consistency were evaluated based on two criteria: (i) comparison of the experimentally determined loading free energy changes for each of the individual sites in wild type and mutant operators, to values calculated from the model-dependent fitting parameters; and (ii) comparison of the ratio of variances for each individual site from the separate and global analyses, to the F statistic.
Footprint titrations of CRP
binding show the expected protected regions corresponding to CRP1 and
CRP2 (Fig. 2). Affinity for CRP2 is higher than for CRP1
as evidenced by the CRP concentration dependence of the protection. The
usual CRP protection pattern in which two bands become hypersensitive
to DNase I (cf. Refs. 9, 11) is observed in both sites but
is much more prominent in the higher affinity site, CRP2. The
fractional protections of CRP1 and CRP2 were separately determined and
analyzed using Equation 1 to obtain apparent free energy changes for
CRP binding to each of these sites. Separate analysis of the
hypersensitive bands and protected regions of CRP2 yielded
indistinguishable estimates of
G2, app,
indicating that the hypersensitivity and protection result from the
same molecular event: binding to CRP2.
G1,app =
11.9 ± 0.1 kcal/mol and
G2,app =
13.3 ± 0.1 kcal/mol.
Fraction of CRP as Functionally Active CRP(cAMP)1
The role of cAMP as an allosteric effector
of CRP functional states is well known (26, 41). Free CRP dimers have
three quaternary conformational states corresponding to the three cAMP ligation states. cAMP binding is negatively cooperative, thus favoring
the singly liganded species, CRP(cAMP)1, which is the functionally active, site-specific DNA binding form (42). However, as a
consequence of overlapping transitions for binding of the first and
second cAMPs, only a fraction of CRP is in this functionally active
form even at optimal cAMP concentration. Therefore it was necessary to
analyze the cAMP concentration dependence of CRP binding to
deoP2 to determine concentrations of active
CRP(cAMP)1 dimer in our experiments. Results are shown in
Fig. 3.
Gapp for binding to CRP1
(squares) and CRP2 (diamonds) relative to a 1 M total CRP dimer standard state. Individual
site binding curves from separate DNase I footprint titration
experiment at each [cAMP] were analyzed using Equation 1. Error
bars are confidence limits to fitted
Gapp. The solid curves drawn
through the points represent analysis of these data as described in the
text.
Consistent with previous reports, two overlapping transitions yield a
maximum in apparent affinity between 100 and 200 µM cAMP.
Affinities for CRP1 and CRP2 parallel each other over the entire range
of cAMP concentrations, consistent with the conclusion that cAMP is an
effector of pre-existing conformational states of free CRP dimers.
These data were analyzed quantitatively using Equation 4 of Heyduk and
Lee (41) to estimate the free energy changes corresponding to
macroscopic, step-wise association constants for binding of one
(K1) and two (K2) cAMPs
to free CRP dimers and for binding of CRP(cAMP)1 and
CRP(cAMP)2 to CRP1 and CRP2. The operator DNA binding
affinity of unliganded CRP is assumed equal to 0 in this model. This
analysis yielded K1 equal to (2.6 ×/
2.4) × 104 M
1 and
K2 equal to (1.8 ×/
1.8) × 103
M
1. These estimates at pH 7.0 are 6 times
less and 2 times greater than estimates at pH 7.8 (41) indicating both
lower intrinsic cAMP binding affinity and lower (negative)
cooperativity, i.e. weaker coupling between cAMP binding and
equilibria between quaternary conformational states.
Based on these values of K1 and K2, the fraction of CRP(cAMP)1 is calculated to reach a maximum of 0.635 (±0.021 by propagation of errors) of total CRP at 150 µM cAMP, the concentration used in all subsequent experiments. Although no unique estimate for the affinity of CRP(cAMP)2 for binding to either CRP1 or CRP2 was obtainable, the analysis did yield as an upper limit to the operator binding affinity of CRP(cAMP)2 a value 100-fold lower than that of CRP(cAMP)1. On this basis, the simplifying assumption was made that only CRP(cAMP)1 is functionally active in all subsequent analyses. Free energy changes reported for CRP binding use a standard state of 1 M dimeric CRP(cAMP)1. Estimates of the free energy changes for binding of CRP(cAMP)1 to CRP1 and to CRP2 are in Table I. Affinity for CRP2 is 10-fold higher than for CRP1, consistent with results reported at other experimental conditions (6).
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CytR binding to
deoP2 was also investigated by footprint titration. In
contrast to the CRP footprints, which are localized to the 22-bp CRP
binding sites, CytR binding protects an extended region from about bp
25 to
110 bp relative to the start site for transcription
(Fig. 4). This extended footprint includes not only the
previously identified CytR binding site (6) but also both flanking CRP
binding sites. Based on literature reports that CytR binds to a single
specific site flanked by CRP1 and CRP2, routine analysis of fractional
protection included only this sequence, i.e. approximately
bp
55 to
80 (see Fig. 1). Analysis of these data using the simple
binding model (Equation 1) yielded an apparent free energy change for
CytR binding to this site, equal to
10.4 ± 0.4 kcal/mol (Table
I).
235 and
590 bp (versus P2 start site) are labeled
as described in text. B, individual site curve for CytR
binding to deoP2 plotted as a function of CytR dimer
concentration. Solid curve fits Equation 1; this yields a
free energy change for intrinsic binding equal to
10.4 ± 0.4 kcal/mol.
Two additional CytR footprints were observed, one centered at about
235 bp and another at about
592 bp from the P2 transcription start
site. The latter is in the P1 regulatory region and overlaps DeoR
operator site, O1 (43). Hence, we denote this as the
"P1" CytR site. We denote the former as the "upstream" CytR
site. The CytR concentration dependences for protection at these sites
differ from that for the P2 site. Therefore, these represent distinct, specific CytR binding sites that have not been previously described. Quantitative analysis of the upstream site yielded the binding free
energy change of
11.6 ± 0.2 kcal/mol (mean of four
experiments). Thus, CytR's affinity for this site is nearly 10-fold
greater than its affinity for the P2 site. At 750 bp from the
32P-labeled end of the DNA fragment, the electrophoretic
resolution of bands in the P1 site is inadequate for quantitative
analysis.
To evaluate the possibility of cooperative interactions between CytR
bound to deoP2 and CytR bound to the upstream and/or P1
site(s), the P2 regulatory region and the upstream and P1 regions were
isolated on separate DNA fragments as described above. Footprint titration experiments conducted using these DNA fragments yielded apparent binding free energies for the deoP2 (
10.5 ± 0.1 kcal/mol; mean of five experiments) and upstream (
11.2 ± 0.3 kcal/mol; mean of nine experiments) CytR binding sites that are
indistinguishable from those obtained using the larger P1/P2 containing
DNA fragment. This result indicates no cooperative interaction between
CytR bound to P2 and CytR bound to the upstream site. We infer also no
interaction between P1 and P2 sites. Therefore, the apparent free
energy changes obtained in these analyses are intrinsic free energy
changes for binding of CytR to the local sites. Quantitative protection
data for the P1 site were obtained by labeling the P1-containing
fragment at the end near P1. Analysis of these data yielded an
intrinsic free energy change for CytR binding equal to
10.5 ± 0.6 kcal/mol (mean of eight experiments). Thus, affinities of CytR for
the P1 and P2 sites are approximately equal.
To evaluate cooperative interactions
between different proteins binding to DNA, we considered the
thermodynamic cycle for their simultaneous binding (cf. Ref.
44). Fig. 5 illustrates the approach based on the
deoP2 structure diagrammed in Fig. 1. The total free energy
change to fill an individual site with ligand, including the effects of
interactions with other ligands, is the individual site loading free
energy change,
Gl, (38) a model independent
quantity.
GCytR, the loading free energy change for CytR binding alone (no CRP) is equal to the intrinsic free
energy change,
G3 (Fig. 5). The loading free
energy change for CytR binding to deoP2 that is saturated by
CRP(cAMP)1 (
GCytRCRP)
includes contributions from both intrinsic binding and cooperativity, i.e.
G3 +
G123. Thus
G123 = (
GCytRCRP)
GCytR).
G123 can
also be evaluated by comparing CRP(cAMP)1 binding in the
presence versus the absence of saturating CytR. Cooperativity contributes unequally to CRP1 and CRP2, dependent on
their relative intrinsic affinities for CRP(cAMP)1 binding (
G1 and
G2; Fig.
5). Therefore, the CytR-mediated differences in loading free energy
changes for CRP(cAMP)1 binding to CRP1 and CRP2 are summed
to yield
G123 = (
GCRP1CytR
GCRP1) + (
GCRP2CytR
GCRP2). These two independent methods for
evaluation of
G123 provide a critical
control: if the molecular model properly accounts for all molecular
configurations and free energy states, the same value for
G123 must be obtained either way.
Gi,
i = 1, 2 or 3) and the Gibbs free energy changes for
pairwise or three-way cooperative interaction between liganded sites
(
Gij(k)). Cooperativity is defined thermodynamically as the difference between the total free energy change to saturate two or more sites simultaneously and the sum of
intrinsic free energy changes to fill them separately.
To evaluate the cooperative free energy change in this manner, loading free energy changes were determined for binding of CytR and CRP(cAMP)1 to deoP2, each in the presence and absence of a fixed concentration of the other (Table I). As a practical approximation to the limit of saturating CRP(cAMP)1, 0.1 µM CRP (total dimer) and 150 µM cAMP were used. Saturation of CRP1 and CRP2 are 0.97 and >0.99 at the resulting CRP(cAMP)1 concentration (64 nM). The fixed CytR concentration used was 0.5 µM, which yields 0.97 saturation. Table I reflects many repetitions of titration experiments on wild type deoP2. This is because titration experiments were conducted using both the longer, P1/P2 containing DNA fragment and the shorter P2 containing fragment (Fig. 1). Identical results were obtained for the two operator fragments, as described earlier. As an additional control, CytR titrations were conducted in the presence and absence of cAMP. At 150 µM, cAMP had no effect on intrinsic CytR binding.
G13 and
G23, which
pertain to pairwise cooperative interactions between CytR binding and
CRP bound to either CRP1 or CRP2, were similarly evaluated using
reduced valency mutants in which specific binding to either CRP1
(CRP1
) or CRP2 (CRP2
) was eliminated. The
mutants were produced by introducing a G to A transition into each of
the TGTGA motifs for either site (45, 46, 47). We mutated both TGTGA motifs
for each site because this is reportedly necessary to completely
abolish CRP activation of deoP2 (20).
No specific binding of CRP to either mutated site was observed.
Intrinsic binding to the remaining site (CRP1 or CRP2) was identical to
binding to CRP1 and CRP2 in the wild type operator (Table I). This is
consistent with the conclusion that CRP binding is noncooperative. CytR
titrations of CRP1
and CRP2
were conducted
to evaluate whether there was any effect of mutating either CRP1 or
CRP2. CytR binding to CRP1
, CRP2
, and wild
type deoP2 were identical (Table I) indicating no such
effect. We infer that there is also no effect of mutation of either CRP
site on the remaining CRP site.
Table II lists
Gij(k) values
obtained by taking differences between values in Table I, as described
above. Whereas the apparent cooperativity as evaluated from CytR
titrations is substantial, consistent with previous reports, the
apparent cooperativity as evaluated from CRP titrations is much weaker.
In particular, pairwise binding of CytR and CRP to either site appears
to be essentially noncooperative when evaluated from CRP titrations. This phenomenology is consistent with recent reports of
"unidirectional stimulation" in CRP and CytR interaction with
nupG (11). However, when couched in quantitative terms as in
Table II, it is evident that this phenomenology reflects the failure of
Fig. 5 to account for all configurations of bound CytR and/or cAMP-CRP.
Additional sites of interaction for one or both proteins are necessary
to explain these results. Control experiments cited above demonstrate that these effects are not the result of interactions with the upstream
or P1 CytR sites. Thus, we conclude that as yet uncharacterized sites
near deoP2 must be responsible.
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Considering it unlikely that high affinity CRP sites
would be overlooked since the sequence specificity for CRP binding is well known (45, 46, 47, 48), we decided to analyze the extended DNase I
protection pattern conferred by CytR binding (Fig. 4). The CytR
concentration dependence of separate regions of the extended CytR
footprint was evaluated systematically to determine whether the entire
footprint represents a single binding event or multiple binding of CytR
to separate sites. To obtain adequate electrophoretic resolution of the
CytR footprint for this purpose, CytR titrations were conducted using
the shorter, P2 DNA fragment (Fig. 1). Titration curves were
constructed from the fractional protection in a series of blocks of DNA
bands covering the sequence from bp
31 to
100 relative to the P2
start site. In defining blocks of bands for analysis, it is necessary
to choose well resolved DNA bands (24). As a consequence the blocks
analyzed represent a mixture of contiguous and overlapping blocks.
Results of this analysis are shown in Fig. 6.
10.8 ± 0.2 (included in average shown in Table I) and
9.7 ± 0.3 kcal/mol.
The apparent free energy change for the usual CytR block extending from
approximately bp
55 to
80 is consistent with the value listed in
Table I. Sub-partitions of this block yield values indistinguishable
from this, albeit with lower precision in some cases. This indicates
that the CytR concentration dependence of the protection over this
region is the same, consistent with a single molecular event, binding
of CytR to a single site. By contrast, the fractional protection that
extends over CRP1 and CRP2 shows a different CytR concentration
dependence as reflected in smaller apparent free energy changes for
CytR binding. Therefore, this protection reflects different molecular
events; it cannot be due to CytR binding to the
55 to
80 region. It
suggests additional CytR binding sites with somewhat lower affinity
than the previously identified CytR site (site 3; Fig. 5) and which
partially or wholly overlap CRP1 and CRP2. This extended protection
reflects specific binding of CytR to DNA. The more or less uniform
protection of the entire DNA fragment that results from nonspecific
binding is observed, but only at higher CytR concentrations than were used in these experiments.
To determine whether such interactions of CytR with DNA sequences overlapping and thereby occluding or competing with CRP binding to CRP1 and CRP2 can account quantitatively for both the loading free energy changes in Table I and apparent free energy changes in Fig. 6, two models that incorporate two additional CytR binding sites and which constitute opposing possibilities were evaluated. The possibilities are 1) that the additional CytR sites that overlap CRP1 and CRP2 are specific binding sites to which CytR binds noncooperatively and with defined affinity; and 2) that the additional binding is nonspecific but cooperative. Competitive binding was formulated as rules (constraints) that (i) CRP(cAMP)1 binding to CRP1 and CytR binding to the site overlapping CRP1 (denoted site 4) are mutually exclusive; and (ii) similarly, that CRP(cAMP)1 binding to CRP1 and CytR binding to the site overlapping CRP2 (denoted site 5) are mutually exclusive. In the second model, CytR bound to the high affinity site 3 nucleates nonspecific binding on either side. Since our data do not precisely define which base pairs constitute the additional CytR sites, the models define these sites only in terms of thermodynamic properties. Table III lists the operator configurations and free energy states that result.
The titration data represented by the
Gl values
in Table I were analyzed according to these models, which we denote Specific, and Nonspecific, Additional Sites. The concentrations of both
CytR and CRP(cAMP)1, whether used as titrant or as held constant, were the independent variables. The fractional protection of
individual sites was the dependent variable. No data for putative CytR
sites 4 and 5 were included in the analyses. Instead we used the data
for the known sites, 1-3, to determine (i) whether such additional
CytR binding sites could account quantitatively for the cooperative
free energy changes in Table II, (ii) whether in so doing, well bounded
parameter values,
G4 and
G5 (or
Gn.s. and
Gc) would be obtained, and (iii) whether the
parameters so obtained are consistent with the analysis of extended
protection in Fig. 6.
Including both the eight free energy contributions listed in Table III
and the individual site, fractional protection end points (24), 176 adjustable parameters are required for a global analysis of all 84 binding curves represented in Table I. Since this exceeds limitations
of our software and hardware, we instead analyzed two representative
titrations from each line in the table, 24 in all. In three cases these
were the only data. In all other cases, the criteria for selection were
(i) that the
Gl values for the pair of
experiments chosen reflect the mean and standard deviation of the
entire set so far as possible and (ii) that subject to criterion i, the
data be of the highest precision and best distribution of independent
variable available. Criterion ii ensures the greatest possible
sensitivity to systematic differences between experiments with
different titrants and different operators, thus imposing the most
critical possible standard on evaluation of goodness of fit.
To minimize effects of subjective bias in the selection of representative curves, we repeated the analysis using a second data subset in which different titrations were selected whenever possible. A third analysis was conducted in which the concentratio