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Proc Natl Acad Sci U S A. 2005 January 11; 102(2): 285–290. Published online 2004 December 30. doi: 10.1073/pnas.0405779102. | PMCID: PMC544287 |
Copyright © 2005, The National Academy of Sciences Biochemistry The effects of upstream DNA on open complex formation by Escherichia coli RNA polymerase Caroline A. Davis,* Michael W. Capp,† M. Thomas Record, Jr,*† and Ruth M. Saecker†‡ Departments of *Biochemistry and †Chemistry, University of Wisconsin, Madison, WI 53706 Received August 6, 2004; Accepted November 16, 2004. Binding of activators to upstream DNA sequences regulates transcription initiation by affecting the stability of the initial RNA polymerase (RNAP)–promoter complex and/or the rate of subsequent conformational changes required to form the open complex (RPO). Here we observe that the presence of nonspecific upstream DNA profoundly affects an early step in formation of the transcription bubble. Kinetic studies with the λPR promoter and Escherichia coli RNAP reveal that the presence of DNA upstream of base pair -47 greatly increases the rate of forming RPO, without significantly affecting its rate of dissociation. We find that this increase is largely due to an acceleration of the rate-limiting step (isomerization) in RPO formation, a step that occurs after polymerase binds. Footprinting experiments reveal striking structural differences downstream of the transcription start site (+1) in the first kinetically significant intermediate when upstream DNA is present. On the template strand, the DNase I downstream boundary of this early intermediate is +20 when upstream DNA is present but is shortened by approximately two helical turns when upstream DNA beyond -47 is removed. KMnO4 footprinting reveals an identical initiation bubble (-11 to +2), but unusual reactivity of template strand upstream cytosines (-12, -14, and -15) on the truncated promoter. Based on this work, we propose that early wrapping interactions between upstream DNA and the polymerase exterior strongly affect the events that control entry and subsequent unwinding of the DNA start site in the jaws of polymerase. Early steps in transcription initiation require interactions with DNA sequences located upstream from the start site (+1). In eukaryotes, assembly of the RNA polymerase (RNAP) II preinitiation complex hinges on the recruitment of the TATA-binding protein to sequences typically located ≈25 bp upstream from +1 (see, for example, ref. 1). In Escherichia coli, the specificity factor σ 70 brings the polymerase machinery to a promoter via interactions with the so-called -35/-10 elements ( 2, 3). The rates and equilibria of steps involved in binding and opening the DNA are further modulated by proteins (activators, enhancers, and repressors) that bind to sites located even further upstream ( 1, 4). Strikingly, high-resolution structures of bacterial RNAP and yeast RNAP II reveal a conserved architecture that dictates that upstream interactions cannot place downstream DNA directly in the cleft containing the active site ( 5– 7). Instead, significant DNA deformations must occur to access the cleft. Additionally, the width of the cleft fundamentally determines whether single- or double-stranded DNA can enter. How do interactions with upstream DNA sequences influence these critical steps or other conformational changes that occur in forming the open complex? A paradigmatic example of the role of upstream DNA sequences in regulating transcription is found in the lysis/lysogeny decision of λ phage after infection of E. coli ( 8). DNA sequences separating two divergent phage promoters (λP R and λP RM) contain three sites for the phage protein λcI. In a precise balance between λcI concentration, site affinity, and position, transcription is repressed from λP R and activated at λP RM ( 9– 11). λcI activates λP RM by facilitating the rate-limiting conformational change (isomerization) in open complex formation, a step that occurs after polymerase binds ( 12– 14). λcI activation of λP RM has been hypothesized to depend critically on the formation of a correctly aligned λcI-σ interface ( 15, 16), recently defined by x-ray crystallography ( 16). How does formation of such an interface increase the rate-limiting isomerization rate constant ( k2)by ≈10-fold at 37°C, with no net effect on the stability of the preceding intermediate ( 12, 14)? Additional puzzles are presented by activation of a polymerase containing a mutant σ 70 by WT λcI, which increases the stability of the first intermediate by ≈10-fold, with only a 2-fold increase in k2 ( 13). Intriguingly, modeling indicates that the transition from a nonproductive to a productive interface may alter the path of the DNA upstream of λcI (approximately -50) ( 16). These data, along with other studies, suggest to us that the trajectory of upstream DNA is defined in the earliest steps of open complex formation, and that this trajectory and the resulting interactions between upstream DNA and polymerase dictate subsequent downstream events. For E. coli RNAP, interactions between σ 70 and the -35 element likely set the initial trajectory; binding to the -35 consensus sequence by the homologous σ from Thermus aquaticus changes the DNA helical axis by 36 degrees ( 17). Changes to or reinforcements of this initial trajectory presumably occur when the C-terminal domains (CTD) of the α subunits ( 18) and/or proteins such as catabolite activator protein (CAP), λcI, IHF, or FIS bind upstream, bend their DNA sites ( 16, 19– 21), and as a consequence activate transcription initiation. Possibly more significantly, the phasing of these activator sites with respect to the promoter critically affects their function ( 22– 24). For example, repositioning of CAP at the gal P1 promoter by 1 bp, from a site centered at -41.5 to -40.5, converts CAP from an activator to a repressor ( 23). Do these experiments indicate that upstream DNA plays a direct role in activation, possibly by contacting core RNAP, or does it serve primarily as an assembly platform that mediates protein–protein interactions? Several lines of evidence suggest that upstream DNA wraps around E. coli RNAP during initiation, even in the absence of transcription factors. In particular, hydroxyl radical footprinting studies of the open complex (RP O) formed at several promoters (λP R, lacUV5, and T7 A1) reveal a pattern of periodic protection of upstream DNA extending to approximately -70 ( 25– 27). To reconcile the length and periodicity of the RP O footprint with the dimensions of E. coli RNAP determined by electron microscopy ( 28), upstream DNA was proposed to wrap in a surface groove of RNAP ( 25, 28). A similar model of RP O at λP R was deduced from atomic force microscopy data ( 29). However, for RP O formed at lacUV5, only the flexibly tethered αCTDs were found to crosslink to DNA positions -43 to -93 ( 30). If this crosslinking pattern is general, and αCTD binding explains upstream protection in RP O ( 18, 25– 27, 31– 34), then it is necessary to propose rapidly reequilibrating nonspecific binding of the αCTDs to the minor groove of DNA throughout this region ( 30). Consistent with, but by no means a proof of, this hypothesis, deletion or mutation of the αCTDs eliminates protection from -40 to -70 in RP O ( 18, 27, 35, 36). Although it is evident that interactions with upstream DNA influence formation of RPO, little quantitative data exist regarding upstream contacts in the intermediates that precede it. How do these interactions influence the individual steps that form and then convert the initial polymerase–DNA complexes into RPO? To elucidate the role of upstream DNA in open complex formation, we compared a λPR promoter truncated at position -47 [upstream truncated (UT)] with full-length (FL) λPR using kinetic and footprinting experiments. The kinetics of formation and dissociation of RPO complexes at the λPR promoter require a minimal three-step mechanism: where R is free polymerase; P is promoter DNA; I 1 and I 2 are the first and second kinetically significant intermediates, respectively; and RP O is the final open complex ( 37– 41). ( K1 and K3 are equilibrium constants for the first and third steps, and k2 and k-2 are the forward and reverse rate constants for the rate-determining interconversions of I 1 and I 2). Our studies demonstrate that DNA sequences upstream of -47 do not greatly affect the stability of the first intermediate or the kinetics of dissociation of RP O at λP R. Instead, we find that they greatly accelerate the conversion of the first to the second intermediate by a factor of 35- to 60-fold. DNase I footprints reveal that the presence of upstream DNA permits downstream DNA to fully enter the active-site cleft of polymerase early in the process. These results motivate our proposal that contacts to upstream DNA sequences directly influence movements in the transcriptional machinery that place the start site in the jaws of polymerase. Buffers. RNAP storage buffer ( 25), binding buffer ( Table 1) ( 41), wash buffer ( 41), 90 mM Tris/64.6 mM boric acid/2.5 mM EDTA, pH 8.3 (TBE buffer) ( 25), urea loading buffer ( 25), and KMnO 4 stop buffer ( 40) have been described. DNase I stop buffer is 200 mM EDTA and 2 M NaCl. | Table 1.Kinetic quantities for RNA polymerase interactions with FL and UT λPR |
RNAP. E. coli K12 RNAP holoenzyme (Eσ 70) was purified by modification of the Burgess and Jendrisak procedure ( 42, 43) and stored at -70°C in storage buffer; working stocks were stored at -20°C. Promoter-binding activities of the RNAP preparations were determined as described ( 41) and ranged from 36% to 77%. Binding activities were determined in parallel with the experiments described here and did not change significantly over the course of 1 month. All RNAP concentrations reported refer to the active RNAP. λPR Promoter DNA. The WT λP R promoter sequence from -60 to +20, centered in a FL 190-bp fragment (-110 to +80), was isolated from plasmid pBR81 and labeled on the nontemplate (nt) strand as described ( 25). A 191-bp FL λP R fragment (-102 to +89) was isolated from a pBR80 derivative by using XbaI and SmaI, and the upstream end of the template (t) strand was labeled with 32P-dCTP by using Sequenase DNA polymerase. The 5′ overhang was filled in with dNTPs. FL λP R used in association assays performed at 37°C was an 898-bp fragment, as described ( 41). We observed no difference in the kinetics of association with the 898- and 187-bp fragments (data not shown). A UT promoter, consisting of the WT λP R sequence from -42 to +20, centered in a 136-bp fragment (-47 to +89), was isolated from plasmid pRLG936 ( 31) by cleavage with EcoRI and SmaI. The upstream end of the t strand was labeled with 32P-dATP. To label the downstream end of the nt strand, an UT λP R promoter fragment (-47 to +64) was generated by cutting pRLG936 with EcoRI and filling in the 5′ overhang with dNTPs. The fragment was then cut with XhoI, and the 3′ end of the nt strand was labeled with 32P-dTTP. All λP R fragments were isolated and purified as described ( 25). To label the nt strand of UT λP R for KMnO 4 probing, pRLG936 was cut with EcoRI. The resulting 5′ γ phosphate was removed and replaced with γ 32 phosphate as described ( 25). Proteins were phenol-extracted, and pRLG936 was cut with SmaI. The λP R fragment was precipitated, and the upstream overhang was filled in with dNTPs. UT λP R was isolated and purified as described in ref. 25. Association Kinetics. Nitrocellulose filter-binding assays were performed in binding buffer by manual mixing ( 41). RNAP concentrations ranged from 2 to 112 nM; DNA concentrations ranged from 0.2 to 0.5 nM. To test that heparin (50 μg/ml final) was a nonperturbing competitor for free RNAP, representative experiments were repeated by using 100 and 250 μg/ml heparin. Equilibrium occupancy of competitor-resistant (CR) complexes and values of kobs exhibited no dependence on heparin concentration, indicating that it is an inert competitor (data not shown). Dissociation Kinetics. RNAP and DNA were mixed in binding buffer and incubated at 17 or 37°C until binding equilibrium was reached. At 37°C, 12 nM RNAP was used for FL and UT λP R; at 17°C, 70 nM RNAP was used with UT λP R and 12 nM RNAP with FL λP R. Irreversible dissociation was initiated by the addition of heparin (50 μg/ml final), and 100 μl of the reaction was removed and filtered at recorded time intervals ( 41). Heparin is not a perturbing competitor for the dissociation of CR complexes at FL ( 44) and UT λP R (data not shown). Data Analysis. Observed first-order rate constants (β CR) for reversible and irreversible association to form CR complexes (RP O, I 2) were determined as described ( 41). The irreversible association rate constant (α CR) was obtained from β CR and the equilibrium fraction of promoter DNA in the form of CR complexes (  ):  , where  was calculated as described ( 41). The dependence of α CR on RNAP concentration ([R] T) is hyperbolic and α CR = k2K1[R] T/(1+ K1[R] T) [( 41); see ]. The product k2K1 is the composite second-order association rate constant ( ka). All fits were weighted by 1/σ 2, where σ 2 is the variance of α CR. t1/2 for isomerization is given by t1/2 = 0.693/ k2. First-order dissociation rate constants ( kd), where kd = k-2/(1 + K3) ( 41), were determined by plotting cpm corr [counts per minute at a given time ( t) corrected for background retention] as a function of time. Dissociation reactions were fit to a first-order decay equation: cpm corr= (cpm 0) e-kdt, where cpm 0 is the initial ( t = 0) counts per minute. All kinetic data were analyzed by using sigmaplot 2000 (Systast, Point Richmond, CA). DNase I Footprinting. RNAP (70 nM) and promoter DNA (0.4–1 nM) were mixed and incubated at 17°C. To characterize I 1, samples were incubated for 15 (FL) or 350 s (UT); RP O was characterized at 2,500 (FL) or 10,000 s (UT). At the appropriate time, heparin (50 μg/ml final) or H 2O was added, and DNase I (0.3 μg/ml final) was added 10 s later; DNase I cleavage was stopped after 10 s. Samples were prepared and loaded on sequencing gels as described in ref. 25. All DNase I footprints were reproduced in three to seven independent experiments. Fractions of DNA in CR and I1 Complexes. The fraction of promoter DNA in CR complexes (  ) at any given time in an association experiment at 17°C and 70 nM RNAP was determined from the integrated relaxation rate equation  . At 350 s after mixing, for UT λP R,  ; at 15 s after mixing, for FL λP R,  . The fraction of promoter DNA present as I 1 complexes (θ I1) was calculated from θ I1 = (1 -  ) [K 1[R] T/(1 + K 1[R] T)]. At 17°C and 70 nM RNAP, θ I1 = 0.68 at 350 s for UT λP R and θ I1 = 0.55 at 15 s for FL λP R. Approximately 24% of total UT λP R was uncomplexed at 350 s, and 33% of total FL λP R was uncomplexed at 15 s. KMnO4 Footprinting. RNAP (20 nM) and promoter DNA (≈8 nM) were incubated at 37°C for 30 min and at 17°C for 2,500 s (FL) or 10,000 s (UT), then challenged for 10 s with heparin (50 μg/ml) or H 2O. KMnO 4 (0.5–3.5 mM) was added and stopped after 10 s. KMnO 4 modification reactions were followed by piperidine cleavage ( 40). Piperidine was removed from the samples as described ( 40); final samples were resuspended in 3 μl of 1 × 10 mM Tris/1 mM EDTA, pH 8.0 (TE buffer) and 4 μl of urea loading buffer and loaded onto an 8% acrylamide gel. KMnO 4 footprints were reproduced in 10 independent experiments run on seven gels. PhosphorImager Analysis. Gel images were obtained by using a Molecular Dynamics PhosphorImager and analyzed as described ( 25) by using imagequant Ver. 4.2 (Molecular Dynamics). Transcription Assays. Single round transcription assays were performed by using unlabeled FL and UT λPR. RNAP (20 nM) and promoter DNA (10 nM) were mixed and, after a 30-min incubation at 37°C, 32P-UTP (50 μM), NTPs (500 μM ATP, 10 μM UTP, 100 μM GTP, and CTP) and heparin (50 μg/ml) were added simultaneously to the reaction. Reactions were stopped after 30 s with urea loading buffer and run on a 15% polyacrylamide gel. Presence of Upstream DNA Results in Faster Kinetics of Association of RNAP at the λPR Promoter. To examine the role of upstream DNA in association at λP R, rates of forming CR complexes, assumed to be primarily RP O, at the UT and FL λP R promoters were measured at 17 and 37°C in excess polymerase ([RNAP] >> [promoter]). At both temperatures and at all RNAP concentrations, CR complexes form much more quickly at FL λP R than at UT λP R (; Fig. 4, which is published as supporting information on the PNAS web site). In contrast to FL λP R, not all UT promoter DNA is converted to CR complexes at 17°C, even though RNAP is in vast excess over promoter DNA. Instead, the equilibrium fraction of promoters in CR complexes (  ) increases with increasing RNAP concentration (reversible association), such that  at the highest RNAP concentration investigated (112 nM). However, at 37°C, formation of CR complexes at UT λP R goes to completion (  ; irreversible association) at all RNAP concentrations investigated. Observation of single exponential kinetics indicates that free RNAP and DNA rapidly equilibrate with a first kinetically significant intermediate (generically abbreviated I 1) on the time scale of its conversion to a CR complex ( 45). [Because formation of I 1 from reactants is a rapidly established equilibrium, the kinetics of forming I 1 are not relevant for the total observed kinetics (α CR). Hence, facilitated diffusion makes no contribution to α CR.] Analysis of the irreversible rate constant α CR as a function of RNAP concentration () allows one to determine the overall composite association rate constant ka and to dissect it into its contributions from the binding ( K1) and isomerization ( k2) steps. The presence of DNA upstream of -47 increases ka significantly (≈36-fold increase at 17°C and ≈7-fold increase at 37°C; Table 1). Unexpectedly, we find that this large effect on ka results primarily from the isomerization step ( Table 1). At 17°C, the presence of upstream DNA greatly reduces the time required for the isomerization of I 1, from a half time of 3,000 s (UT) to 50 s (FL)! At 37°C, the half time for isomerization is reduced from 35 s for UT λP R to 1 s on FL. Equally surprising is the fact that the equilibrium constant for forming I 1 ( K1) increases ≈2-fold upon deletion of upstream DNA ( Table 1). Similar results have been observed for a series of upstream deletions of the lacUV5 promoter ( 46). Similar Kinetics of Dissociation of RNAP from UT and FL λPR. Given the large effects on the kinetics of association, we next examined whether the dissociation of CR complexes was similarly affected. Irreversible dissociation of CR complexes at UT and FL λP R was induced by the addition of heparin (50 μg/ml final), which prevents free RNAP from rebinding to the promoter. The decay kinetics of both UT and FL λP R complexes are single exponential and are significantly slower at 37°C than at 17°C (Fig. 5, which is published as supporting information on the PNAS web site). This behavior demonstrates the existence of a second kinetically significant intermediate (I 2) in rapid equilibrium with RP O on the time scale of the slow step in dissociation ( 38, 47). Fits of these data reveal only minor differences in kd for FL and UT λP R. At 17°C, kd is slightly (1.2-fold) greater for FL than for UT λP R; at 37°C, kd is ≈2-fold greater for UT than for FL λP R ( Table 1). DNase I Footprinting of I1 at UT and FL λPR and Comparisons with RPO. Because the presence of upstream DNA dramatically affects the isomerization of I 1 to I 2, we asked whether the structure of I 1 at FL λP R (  ) differs from that at UT λP R (  ). To address this question, DNase I footprinting was performed at 17°C at times in the association process when the principal polymerase–promoter complex is predicted to be I 1. DNase I footprints of CR complexes (primarily RP O) formed at binding equilibrium were also obtained. Analyses of these footprints reveal that  is competitor-sensitive and has a continuous footprint with downstream boundaries of +20 on the t strand (; Fig. 6, which is published as supporting information on the PNAS web site) and ≈+25 on the nt strand (; Fig. 7, which is published as supporting information on the PNAS web site). Both the sensitivity to heparin and the downstream boundaries of the footprint agree with previous characterizations of a  at equilibrium at 0°C ( 40). In sharp contrast, the downstream DNase I boundary of  extends only to +2 (t) and +7 (nt), with an isolated site of protection at +27 (nt) (, respectively; Figs. 8 and 9, which are published as supporting information on the PNAS web site). In the footprints of  and  , any protection observed after the 10 s heparin challenge is attributed to the small fraction of CR complexes (see Experimental Procedures) present in these experiments. No end-bound complexes were detected in these footprints. Because the structures of the first kinetic intermediates differ, we continue to designate them  and  . On the t strand, protection of DNA in  is continuous to position -8; partial protection is observed from positions -1 to +2. Despite the lack of strong DNase I cleavage sites between positions -7 and -2 on the free DNA, comparison of weak signals with and without RNAP from at least three independent experiments suggests that this region is not protected by RNAP. Enhancements at positions -15 (t) and -16 (t) of the UT promoter are reproducibly observed; these positions are protected in the footprint of  . On the nt strand, protection of  is continuous to -4, with partial protection from +1 to +7, followed by cleavage of the backbone from +8 to +26. The DNA backbone at +27 is reproducibly protected in all footprints of  . Gels run for short times to resolve bands downstream from +27 demonstrate that this isolated site of protection is not the result of an end-bound RNAP (data not shown). In the upstream direction, DNase I protection of  extends at least to -45 (nt), essentially the end of the fragment. The protection of  extends to at least -52 (t) and -45 (nt). In  , DNase I enhancements are located at positions -23 (nt) and -34 (nt); in  , enhancements occur at -22 and -45 (nt). Fractional protection of the upstream (-35 to -3) and downstream (+2 to +27) regions of the  footprint is the same within error (see Figs. 6 and 7). Thus under these conditions, as well as at equilibrium at 0°C ( 40), DNase I footprinting detects a homogeneous population of competitor-sensitive complexes with extended downstream footprints on FL λP R. At binding equilibrium, RP O formed on FL and UT λP R (Figs. 6–9 and Fig. 10, which is published as supporting information on the PNAS web site) have the same downstream boundaries: +20 (t) and ~+25 (nt). As reported ( 25), DNase I enhancements occur at -38 (t), -48 (t), and -45 (nt) in RP O formed on the FL promoter, indicative of DNA deformations (e.g., bending); no enhancements are observed at any position in RP O formed on the UT promoter. No end-bound complexes were detected on UT or FL λP R. Differences in UT and FL RPO as Detected by KMnO4. Equilibrium complexes formed at 37°C and 17°C were next probed with KMnO 4 to estimate the span of the initiation bubble and the pyrimidines (T >> C) that are solvent-accessible in RP O. For both FL and UT promoters, thymines at +1, -8, -9, and -11 (t) and at -10, -4, -3 and +2 (nt) [; ( 40)] are oxidized by KMnO 4 and are therefore solvent-accessible. Reactivity of the thymine at -13 (nt) is not detected at either UT or FL λP R. Surprisingly, cytosines at -12, -14, and -15 (t) in RP O on the UT λP R promoter are KMnO 4 reactive at both 37°C () and 17°C (data not shown), whereas these positions are completely unreactive for FL λP R (). Does this result indicate additional opening of the transcription bubble at the UT promoter? To address this, we probed the t strand of the UT promoter with DMS, which methylates single-stranded cytosines ( 48). In both FL and UT promoters, only cytosines at -2, -5, and -6 are reactive; positions -12, -14, and -15 (t) are not reactive (Fig. 11 and Supporting Text, which are published as supporting information on the PNAS web site) ( 49). The lack of dimethyl sulfate reactivity suggests that these cytosines are distorted but remain base paired or are unpaired but interact with RNAP in such a way that the N3 is protected. Although the structural details of RP O formed at FL and UT λP R appear to differ, both are competent to transcribe DNA upon addition of nucleotides (data not shown). Creation of a transcriptionally competent open complex occurs through a series of steps, whose rates and equilibria can be tuned by regulatory factors. In both prokaryotes and eukaryotes, activators often stabilize an early intermediate (e.g., I 1 or its precursors) at least in part by favorable protein–protein interactions. For promoters where the rate-limiting step occurs after binding, activation by driving subsequent conformational changes provides the most direct method of accelerating open complex formation. Upstream binding of the bacterial protein CAP and the phage protein λcI alters the rate of the isomerization step in open complex formation; at 37°C, CAP binding at -41.5 increases k2 by ≈7-fold at gal P1 ( 23), whereas the activating effect of λcI on k2, when bound immediately upstream of the -35 hexamer, is ≈10-fold at λP RM ( 12, 14). Changing the αCTD-binding sites upstream of -40 from nonspecific DNA sequences to AT-rich “UP” sequences ( 50) also accelerates this “bottleneck” step, with effects on k2 ranging from 2- to 10-fold, depending on the temperature and promoter studied ( 51– 53). Although these effects have been quantified, the precise nature of the critical isomerization step remains unknown. Because DNase I protection of sequences upstream of -38 suggests that both αCTDs are nonspecifically bound to regions centered at -43 (t) and -54 (t) in  , we expected that deletion of DNA upstream of -47 would reduce the stability of I 1. Instead, we find that deletion of this upstream DNA from FL λP R has little effect on the stability of this first kinetically significant intermediate but instead profoundly reduces the rate constant k2 for the subsequent rate-determining conformational change in open complex formation. Under the conditions studied, these effects ( Table 1) are significantly larger than those reported previously for CAP or λcI ( 12, 14, 23) or alterations in the DNA sequence of the αCTD-binding site ( 51– 53). For the lengths studied here, the presence of upstream DNA increases the rate constant k2 for formation of CR complexes (I 2 and RP O) from I 1 by 60-fold at 17°C and 33-fold at 37°C. Analogous effects are reported for kinetics of association between WT RNAP and a series of upstream DNA deletions at the lacUV5 promoter in the accompanying paper ( 46): upstream truncation of DNA at -63 and -42 increases K1 by 20- and 10-fold, respectively, but decreases k2 by ≈30- and 50-fold, respectively, relative to lacUV5 promoters extending to -100 and -130. However, when the αCTDs are absent, the presence of upstream DNA increases k2 by only 2.5-fold ( 46). These results clearly indicate that the αCTDs are necessary to observe the effect of upstream DNA. Discerning whether upstream DNA merely provides additional nonspecific αCTD-binding sites or whether the αCTDs set a trajectory required for wrapping interactions between upstream DNA and other elements of RNAPinI 1 or other early complexes awaits further experiments. When upstream DNA is present, downstream DNA is continuously protected from DNase I to +20 (t) and ≈+25 (nt) in  , presumably due to a cascade of events accompanying upstream wrapping and/or αCTD–DNA interactions. When upstream DNA is removed, promoter DNA is protected only to +2 (t) and +7 (nt) in  with an isolated site of protection at +27 (nt). Therefore, downstream contacts that extend this initial protection to continuous protection to +20 (t) and +27 (nt) in RP O on UT λP R must form during or subsequent to the conversion of I 1 to the (I 1–I 2) ‡ transition state. If such contacts are established in forming (I 1–I 2) ‡, we speculate that these additional events are responsible for dramatically reducing k2 on UT λP R relative to FL λP R and for concomitantly increasing the activation enthalpy [-Rdlnk 2/d(1/T)] required to drive I 1 to (I 1–I 2) ‡ from ≈34 to ≈40 kcal. Surprisingly, even though a much smaller polymerase–promoter DNA interface is formed in  , its stability (-RTlnK 1) is comparable to that of the larger one formed at the FL promoter. We suggest that under these conditions, the loss of upstream and downstream contacts in  is nearly compensated for by a reduction in the energetic costs required to make these contacts (e.g., DNA distortions). One implication of this proposal is that the absence of large effects of activators on K1 should not be interpreted to mean that the activator has no effect on the structure of I 1. That is, activators observed to affect only k2 may do so by altering the interface in I 1, even if not detected in K1. Similarly, differences in the structure of RP O observed by KMnO 4 reactivity may offset differences in upstream interactions and cause kd to remain the same even when upstream DNA is removed. Comparison of KMnO 4 and dimethyl sulfate footprints indicates the presence of DNA distortions at the upstream end of the initiation bubble [-12 to -15 (t)] on UT but not FL λP R. The DNA backbone in this region is also distorted in  . Although the KMnO 4 reactivity detected at these positions could result from a small population of off-pathway complexes formed on UT λP R, the lack of a similar shift in KMnO 4 reactivity of the nt strand suggests that this is unlikely. A model of  proposes that formation of a sharp kink at -11/-12 directs downstream DNA into the jaws created by the β and β′ subunits of RNAP ( 41), thus protecting it from DNase I cleavage to ≈+25 ( 40). The observation that the DNase I footprint of I 1 on the UT promoter is shorter by two helical turns indicates that either the DNA is not properly placed in the jaws and/or the jaws are not in the appropriate conformation to make extensive downstream contacts. Complexes formed at low temperature at other promoters, thought to correspond to early competitor-sensitive intermediates (referred to as RP C), exhibit even shorter downstream DNase I footprints (to ≈-5) ( 7, 23, 26, 54). Darst and coworkers ( 7) inferred from these data and their structure of the bacterial RNAP bound to a 33-bp forked-junction promoter fragment (from -40 to -7) that DNA downstream of the -10 region in RP C is not yet bent into the jaws but instead passes above them. A superposition of the DNA backbone bound in the active site of DNase I [Protein Data Bank ID code file 1DNK ( 55)] onto the DNA bound in RP C ( 7) demonstrates that any position downstream of -5 can be cleaved. To explain protection extending to +2(t)/+7 (nt) in  , we hypothesize that downstream DNA must be bent into the jaws, but that its disposition fundamentally differs from that in  . The isolated site of protection at +27 (nt) in  may also be consistent with an altered downstream DNA trajectory or change in the position of the downstream jaw (see below). Interestingly, the downstream DNase I boundary of  is very similar to that of the unusual open complex formed at the T7 A2 promoter DNA by a variant E. coli RNAP [+1 (t) and +8 (nt); ( 56)]. This variant contains a large deletion in β (Δ186–433) that essentially removes the downstream lobe of the jaw. The downstream footprint of  may also reflect an altered conformation of the downstream jaw resulting from the loss of upstream contacts. In free holoenzyme, the negatively charged N-terminal domain of σ 70 (region 1.1) has been proposed to bind in this region of the jaws ( 57) and thereby prop them open ( 58, 59). Possibly the absence of upstream contacts somehow impedes the expulsion of region 1.1 from the active site cleft, a transition required to form RP O. In this scenario, which is currently being investigated, downstream DNA would be bent into the jaws in  , but prevented from descending into them by σ 70. We thank W. Ross and R. Gourse (University of Wisconsin, Madison) for providing pRLG936, for helpful discussions, and for sharing unpublished data. We are grateful to W. Kontur for labeling some λPR used in this study, S. Darst (The Rockefeller University, New York) for providing coordinates for the model of RPC, C. Bingman (University of Wisconsin, Madison) for valuable discussion, and the reviewers for their comments. This work was supported by National Institutes of Health Grant GM23467. 1. Burley, S. K. & Roeder, R. G. (1996. ) Annu. Rev. Biochem. 65, 769-799. [PubMed] 2. McClure, W. R. (1985. ) Annu. Rev. Biochem. 54, 171-204. [PubMed] 3. Record, M. T., Jr., Reznikoff, W. S., Craig, M. L., McQuade, K. L., Schlax, P. J. (1996. ) in Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology, ed. Neidhardt, F. C. (Am. Soc. Microbiol., Washington, DC), Vol. 1, pp. 792-821. 4. Ebright, R. H. (1993. ) Mol. Microbiol. 8, 797-802. [PubMed] 5. Bushnell, D. A., Westover, K. D., Davis, R. E. & Kornberg, R. D. (2004. ) Science 303, 983-988. [PubMed] 6. Vassylyev, D. G., Sekine, S., Laptenko, O., Lee, J., Vassylyeva, M. N., Borukhov, S. & Yokoyama, S. (2002. ) Nature 417, 712-719. [PubMed] 7. Murakami, K. S., Masuda, S., Campbell, E. A., Muzzin, O. & Darst, S. A. (2002. ) Science 296, 1285-1290. [PubMed] 8. Ptashne, M., Johnson, A. D. & Pabo, C. O. (1982. ) Sci. Am. 247, 128-130, 132, 134-140. [PubMed] 9. Hershberger, P. A. & deHaseth, P. L. (1991. ) J. Mol. Biol. 222, 479-494. [PubMed] 10. Hershberger, P. A., Mita, B. C., Tripatara, A. & deHaseth, P. L. (1993. ) J. Biol. Chem. 268, 8943-8948. [PubMed] 11. Ackers, G. K., Johnson, A. D. & Shea, M. A. (1982. ) Proc. Natl. Acad. Sci. USA 79, 1129-1133. [PubMed] 12. Hawley, D. K. & McClure, W. R. (1982. ) J. Mol. Biol. 157, 493-525. [PubMed] 13. Li, M., McClure, W. R. & Susskind, M. M. (1997. ) Proc. Natl. Acad. Sci. USA 94, 3691-3696. [PubMed] 14. Shih, M. C. & Gussin, G. N. (1983. ) Proc. Natl. Acad. Sci. USA 80, 496-500. [PubMed] 15. Dove, S. L., Huang, F. W. & Hochschild, A. (2000. ) Proc. Natl. Acad. Sci. USA 97, 13215-13220. [PubMed] 16. Jain, D., Nickels, B. E., Sun, L., Hochschild, A. & Darst, S. A. (2004. ) Mol. Cell 13, 45-53. [PubMed] 17. Campbell, E. A., Muzzin, O., Chlenov, M., Sun, J. L., Olson, C. A., Weinman, O., Trester-Zedlitz, M. L. & Darst, S. A. (2002. ) Mol. Cell 9, 527-539. [PubMed] 18. Ross, W., Gosink, K. K., Salomon, J., Igarashi, K., Zou, C., Ishihama, A., Severinov, K. & Gourse, R. L. (1993. ) Science 262, 1407-1413. [PubMed] 19. Schultz, S. C., Shields, G. C. & Steitz, T. A. (1991. ) Science 253, 1001-1007. [PubMed] 20. Rice, P. A., Yang, S., Mizuuchi, K. & Nash, H. A. (1996. ) Cell 87, 1295-1306. [PubMed] 21. Pan, C. Q., Finkel, S. E., Cramton, S. E., Feng, J. A., Sigman, D. S. & Johnson, R. C. (1996. ) J. Mol. Biol. 264, 675-695. [PubMed] 22. Newlands, J. T., Josaitis, C. A., Ross, W. & Gourse, R. L. (1992. ) Nucleic Acids Res. 20, 719-726. [PubMed] 23. Lavigne, M., Kolb, A. & Buc, H. (1992. ) Biochemistry 31, 9647-9656. [PubMed] 24. Adhya, S., Gottesman, M., Garges, S. & Oppenheim, A. (1993. ) Gene 132, 1-6. [PubMed] 25. Craig, M. L., Suh, W. C. & Record, M. T., Jr. (1995. ) Biochemistry 34, 15624-15632. [PubMed] 26. Schickor, P., Metzger, W., Werel, W., Lederer, H. & Heumann, H. (1990. ) EMBO J. 9, 2215-2220. [PubMed] 27. Kolb, A., Igarashi, K., Ishihama, A., Lavigne, M., Buckle, M. & Buc, H. (1993. ) Nucleic Acids Res. 21, 319-326. [PubMed] 28. Polyakov, A., Severinova, E. & Darst, S. A. (1995. ) Cell 83, 365-373. [PubMed] 29. Rivetti, C., Guthold, M. & Bustamante, C. (1999. ) Embo J. 18, 4464-4475. [PubMed] 30. Naryshkin, N., Revyakin, A., Kim, Y., Mekler, V. & Ebright, R. H. (2000. ) Cell 101, 601-611. [PubMed] 31. Ross, W., Aiyar, S. E., Salomon, J. & Gourse, R. L. (1998. ) J. Bacteriol. 180, 5375-5383. [PubMed] 32. Ross, W., Schneider, D. A., Paul, B. J., Mertens, A. & Gourse, R. L. (2003. ) Genes Dev. 17, 1293-1307. [PubMed] 33. Benoff, B., Yang, H., Lawson, C. L., Parkinson, G., Liu, J., Blatter, E., Ebright, Y. W., Berman, H. M. & Ebright, R. H. (2002. ) Science 297, 1562-1566. [PubMed] 34. Estrem, S. T., Ross, W., Gaal, T., Chen, Z. W., Niu, W., Ebright, R. H. & Gourse, R. L. (1999. ) Genes Dev. 13, 2134-2147. [PubMed] 35. Minakhin, L. & Severinov, K. (2003. ) J. Biol. Chem. 278, 29710-29718. [PubMed] 36. Aiyar, S. E., Gourse, R. L. & Ross, W. (1998. ) Proc. Natl. Acad. Sci. USA 95, 14652-14657. [PubMed] 37. Roe, J. H., Burgess, R. R. & Record, M. T., Jr. (1984. ) J. Mol. Biol. 176, 495-522. [PubMed] 38. Roe, J. H., Burgess, R. R. & Record, M. T., Jr. (1985. ) J. Mol. Biol. 184, 441-453. [PubMed] 39. Roe, J. H. & Record, M. T., Jr. (1985. ) Biochemistry 24, 4721-4726. [PubMed] 40. Craig, M. L., Tsodikov, O. V., McQuade, K. L., Schlax, P. E., Jr., Capp, M. W., Saecker, R. M. & Record, M. T., Jr. (1998. ) J. Mol. Biol. 283, 741-756. [PubMed] 41. Saecker, R. M., Tsodikov, O. V., McQuade, K. L., Schlax, P. E., Jr., Capp, M. W. & Record, M. T., Jr. (2002. ) J. Mol. Biol. 319, 649-671. [PubMed] 42. Burgess, R. R. & Jendrisak, J. J. (1975. ) Biochemistry 14, 4634-4638. [PubMed] 43. Gonzalez, N., Wiggs, J. & Chamberlin, M. J. (1977. ) Arch. Biochem. Biophys. 182, 404-408. [PubMed] 44. Suh, W. C., Leirmo, S. & Record, M. T., Jr. (1992. ) Biochemistry 31, 7815-7825. [PubMed] 45. Tsodikov, O. V. & Record, M. T., Jr. (1999. ) Biophys. J. 76, 1320-1329. [PubMed] 46. Ross, W. & Gourse, R. L. (2005. ) Proc. Natl. Acad. Sci. USA 102, 291-296. [PubMed] 47. Buc, H. & McClure, W. R. (1985. ) Biochemistry 24, 2712-2723. [PubMed] 48. Kirkegaard, K., Buc, H., Spassky, A. & Wang, J. C. (1983. ) Proc. Natl. Acad. Sci. USA 80, 2544-2548. [PubMed] 49. Suh, W. C., Ross, W. & Record, M. T., Jr. (1993. ) Science 259, 358-361. [PubMed] 50. Gourse, R. L., Ross, W. & Gaal, T. (2000. ) Mol. Microbiol. 37, 687-695. [PubMed] 51. Strainic, M. G., Jr., Sullivan, J. J., Velevis, A. & deHaseth, P. L. (1998. ) Biochemistry 37, 18074-18080. [PubMed] 52. Rao, L., Ross, W., Appleman, J. A., Gaal, T., Leirmo, S., Schlax, P. J., Record, M. T., Jr. & Gourse, R. L. (1994. ) J. Mol. Biol. 235, 1421-1435. [PubMed] 53. Tang, Y., Murakami, K., Ishihama, A. & deHaseth, P. L. (1996. ) J. Bacteriol. 178, 6945-6951. [PubMed] 54. Kovacic, R. T. (1987. ) J. Biol. Chem. 262, 13654-13661. [PubMed] 55. Weston, S. A., Lahm, A. & Suck, D. (1992. ) J. Mol. Biol. 226, 1237-1256. [PubMed] 56. Severinov, K. & Darst, S. A. (1997. ) Proc. Natl. Acad. Sci. USA 94, 13481-13486. [PubMed] 57. Mekler, V., Kortkhonjia, E., Mukhopadhyay, J., Knight, J., Revyakin, A., Kapanidis, A. N., Niu, W., Ebright, Y. W., Levy, R. & Ebright, R. H. (2002. ) Cell 108, 599-614. [PubMed] 58. Murakami, K. S., Masuda, S. & Darst, S. A. (2002. ) Science 296, 1280-1284. [PubMed] 59. Mooney, R. A. & Landick, R. (2003. ) Genes Dev. 17, 2839-2851. [PubMed]
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