INTRODUCTION

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 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).

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 K_d 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.

MATERIALS AND METHODS

Crystalline adenosine 3-5 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.

CRP was expressed from E. coli strain K12 Htrp 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).

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 (M_r = 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,  = 0.30 ± 0.02 mg¹ml¹ at 280 nm.²

Fig. 1 shows the deo DNA fragments used. Plasmid pSS1332² contains the deo P1/P2 sequence from 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).

Mutant promoters were generated in which site-specific CRP binding to CRP1 (CRP1; 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.

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

(Eq. 1)

where L is the concentration of free protein ligand, and k_i 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 k_i in Equation 1 (G_i = RT lnk_i). 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 G_l,i (38) for binding experiments in which both CytR and CRP were present. G_l,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, G_l,i equals G_i (Equation 1). Equation 1 can also provide an accurate estimate of G_l,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)

where G_s 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.

Table III. Operator configurations and free energy states for CRP and CytR binding to deoP2: specific and nonspecific models for higher order CytR binding deoP2 configurations with sites denoted as filled (CRP or CytR) or empty (). The total Gibbs free energy of each configuration relative to the unliganded reference state is given as a sum of contributions from eight free energy changes for intrinsic binding of CRP and CytR and for cooperative interaction between liganded sites. Intrinsic binding and cooperative interactions among CRP1, CRP2, and the intervening CytR site are defined in Fig. 5. Two phenomenological models for competition between CytR and CRP define additional CytR binding sites that occlude CRP1 and CRP2 in terms of thermodynamic properties. These are 1) Specific Binding, CytR binds specifically, but non-cooperatively to two additional operators that occlude CRP1 and CRP2, with intrinsic free energy changes G4 and G5, respectively; and 2) Non-specific (n.s.), Cooperative (C), Binding, CytR binds non-cooperatively with intrinsic free energy change, Gn.s., and binding to directly adjacent sequences is weakly cooperative, with free energy change, Gc. Operator configurations Free energy contributions if inhibition is due to CRP2   (CytR) CytR   (CytR) CRP1 Additional specific CytR binding Non-specific, cooperative CytR binding 1       Reference state Reference state  Gs,1 2     CRP  G1  G1  Gs,2 3 CRP      G2  G2  Gs,3 4   CytR    G3  G3  Gs,4 5     CytR  G4  Gn.s.  Gs,5 6 CytR      G5  Gn.s.  Gs,6 7 CRP   CRP  G1 + G2  G1 + G2  Gs,7 8   CytR CRP  G1 + G3 + G13  G1 + G3 + G13  Gs,8 9 CytR   CRP  G1 + G5  G1 + Gn.s.  Gs,9 10 CRP CytR    G2 + G3 + G23  G2 + G3 + G23  Gs,10 11 CRP   CytR  G2 + G4  G2 + Gn.s.  Gs,11 12   CytR CytR  G3 + G4  G3 + Gn.s. + Gc  Gs,12 13 CytR CytR    G3 + G5  G3 + Gn.s. + Gc  Gs,13 14 CytR   CytR  G4 + G5 2Gn.s.  Gs,14 15 CRP CytR CRP  G1 + G2 + G3 + G123  G1 + G2 + G3 + G123  Gs,15 16 CRP CytR CytR  G2 + G3 + G4 + G23  G2 + G3 + Gn.s. + G23 + Gc  Gs,16 17 CytR CytR CRP  G1 + G3 + G5 + G13  G2 + G1 + Gn.s. + G13 + Gc  Gs,17 18 CytR CytR CytR  G3 + G4 + G5  G3 + 2(Gn.s. + Gc)  Gs,18

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.

RESULTS

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 G_{2, app}, indicating that the hypersensitivity and protection result from the same molecular event: binding to CRP2.

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.

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 M¹ and K₂ equal to (1.8 ×/ 1.8) × 10³ M¹. 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).

Table I. Loading free energy changes for binding of CRP and CytR to deoP2 regulatory region Free energy changes for saturation of deoP2 operator sites CRP1 and CRP2 with CRP(cAMP)1 or the intervening CytR site with CytR, in the presence or absence of effector ligands as indicated. Values of Gl (kcal/mol) determined by analysis of individual site binding curves as described in the text. deoP2 valency Titrant Effectorsa No. expts.b Operator site CRP2 CytR CRP1 Wild-type CRP cAMP 11  13.0  ± 0.2  11.7  ± 0.2 CRP1 CRP cAMP 5  13.1  ± 0.3 CRP2 CRP cAMP 5  11.6  ± 0.1 Wild-type CRP CytR, cAMP 7  13.5  ± 0.3  13.0  ± 0.4 CRP1 CRP CytR, cAMP 2  13.6  ± 0.3 CRP2 CRP CytR, cAMP 4  12.2  ± 0.5 Wild-type CytR None or cAMP 14  10.4  ± 0.4 CRP1 CytR None or cAMP 3  10.4  ± 0.2 CRP2 CytR None or cAMP 2  10.5  ± 0.5 Wild-type CytR CRP, cAMP 9  13.1  ± 0.2 CRP1 CytR CRP, cAMP 3  12.4  ± 0.3 CRP2 CytR CRP, cAMP 2  12.4  ± 0.5 a  Effector concentrations: CRP, 0.1 µM (total dimer); cAMP, 150 µM; CytR, 0.4 µM (dimer). b  Gl values shown are means of multiple determinations (±S.D. of mean). Number of experiments represented in the means are indicated. Mean values for binding to wild type deoP2 reflect approximately equal numbers of titration experiments conducted using 285 bp of P2 containing and 879 bp of P1/P2 containing DNA fragments. There was no difference in results obtained using these two fragments.

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).

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, G_l, (38) a model independent quantity. G_CytR, 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 (G_CytR^CRP) includes contributions from both intrinsic binding and cooperativity, i.e. G₃ + G₁₂₃. Thus G₁₂₃ = (G_CytR^CRP)  G_CytR). 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 = (G_CRP1^CytR  G_CRP1) + (G_CRP2^CytR  G_CRP2). 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.

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 G_ij(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.

Table II. Apparent cooperativity as assessed by differences in individual site loading free energy changes Gij(k)app was calculated as model-independent (Gl) from values in Table I (see text). Confidence limits were estimated by error propagation. Association of these energetics with particular pair wise or three-way cooperativity is as described by the model in Fig. 5. Titrant  G123app  G13app  G23app CytR  2.7  ± 0.4  1.9  ± 0.7  2.0  ± 0.4 CRP  1.8  ± 0.6  0.6  ± 0.4  0.5  ± 0.4

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.

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 G_l 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 G_n.s. and G_c) 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 G_l 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