FIS (factor for inversion stimulation) is a small dimeric DNA‐bending protein which both stimulates DNA inversion and activates transcription at stable RNA promoters in Escherichia coli. Both these processes involve the initial formation of a complex nucleoprotein assembly followed by local DNA untwisting at a specific site. We have demonstrated previously that at the tyrT promoter three FIS dimers are required to form a nucleoprotein complex with RNA polymerase. We now show that this complex is structurally dynamic and that FIS, uniquely for a prokaryotic transcriptional activator, facilitates sequential steps in the initiation process, enabling efficient polymerase recruitment, untwisting of DNA at the transcription startpoint and finally the escape of polymerase from the promoter. Activation of all these steps requires that the three FIS dimers bind in helical register. We suggest that FIS acts by stabilizing a DNA microloop whose topology is coupled to the local topological transitions generated during the initiation of transcription.
FIS (factor for inversion stimulation) is a small homodimeric DNA‐bending protein from Escherichia coli which both facilitates DNA inversion (Huber et al., 1985; Johnson and Simon, 1985; Kahmann et al., 1985) and activates transcription from stable RNA promoters (Nilsson et al., 1990; Ross et al., 1990). Both stable RNA transcription and DNA inversion are stimulated strongly by negative supercoiling of DNA (Mertens et al., 1984; Lamond, 1985; Bowater et al., 1994) and involve the initial formation of a complex nucleoprotein assembly followed by DNA untwisting at the transcription startpoint and crossover sites respectively (Ohlsen and Gralla, 1992a; Klippel et al., 1993).
The promoters of stable RNA (tRNA and rRNA) operons of E.coli can achieve the highest rates of initiation of all bacterial promoters. Under physiological conditions, these promoters are probably not saturated by RNA polymerase (Zhang and Bremer, 1995), and their regulation, which reflects the importance of their products for essential cellular functions, allows the rate of initiation to be varied over a wide range. The stable RNA promoters contain two important regulatory elements, a GC‐rich discriminator between the −10 region and the transcription startpoint (Travers, 1980a) and an upstream activating sequence (UAS) which extends to ∼120–150 bp upstream of the startpoint and is required for optimal expression (Lamond and Travers, 1983; Gourse et al., 1986; van Delft et al., 1987). The discriminator is a necessary response element for a stringent control system which abrogates stable RNA synthesis in response to amino acid starvation (Cashel et al., 1996) and is mediated by the nucleotide ppGpp in vivo and in vitro (Travers, 1980b; Lamond and Travers, 1985; Hernandez and Cashel, 1995; Josaitis et al., 1995; Zhang and Bremer, 1995).
The UAS DNA is anisotropically flexible (Drew and Travers, 1985; Gourse et al., 1986; Plaskon and Wartell, 1987) and contains, in addition, three binding sites for the FIS protein positioned in helical register (Nilsson et al., 1990; Ross et al., 1990; Condon et al., 1992; Lazarus and Travers, 1993), suggesting that bending of the UAS is necessary for transcriptional activation. Consistent with this notion, the UAS can function in vitro both with and without FIS (Newlands et al., 1991; Zacharias et al., 1992; Gaal et al., 1994). However, bending of the UAS DNA by FIS, although necessary, is not sufficient for activation in vivo since a class of FIS mutants has been isolated which bind and bend DNA but fail to activate transcription (Gosink et al., 1993). Some of these mutants are impaired in cooperative binding to DNA, indicating that transcriptional activation in vivo may require the participation of more than one FIS dimer (L.Lazarus, O.Ninnemann, R.Kahmann and A.A.Travers, unpublished results). In agreement with this observation, we have shown recently that, in vitro, FIS forms a specific nucleoprotein complex at the UAS which recruits polymerase to the tyrT promoter (Muskhelishvili et al., 1995), an effect which requires all three FIS‐binding sites positioned in helical register. On this basis, we proposed that the UAS forms a microloop which is stabilized by FIS.
The formation of the transcription initiation complex at bacterial promoters is a sequential process in which the initial formation of a closed polymerase–promoter complex is followed by structural transitions in both the enzyme and DNA, which eventually result in the untwisting of DNA at the transcription startpoint (Buc and McClure, 1985). It is this latter step which is antagonized by ppGpp (Ohlsen and Gralla, 1992a). The polymerase then initiates transcription and escapes from the promoter. Each of these steps is potentially rate‐limiting and subject to control by transcriptional activators. There is substantial evidence that at the rrnB P1 promoter FIS recruits RNA polymerase into a closed complex and thus increases the KB (Bokal et al., 1995). However, other experiments indicate that FIS may also activate subsequent steps in the initiation pathway. In particular, FIS overrides the inhibitory action of ppGpp on tyrT transcription (Lazarus and Travers, 1993) and at rrnD P1 FIS facilitates the transition to the elongating complex (Sander et al., 1993). In this study, we show directly that FIS affects sequential steps on the initiation pathway, thereby optimizing the interaction of polymerase with the promoter and facilitating high rates of initiation.
Kinetics of FIS–RNA polymerase complex formation at the tyrT promoter
Surface plasmon resonance (SPR) techniques measure small local changes in refractive index at a surface containing a fixed ligand, and can be used to monitor relative affinities of proteins binding to immobilized DNA fragments (Fisher et al., 1994; Buckle et al., 1996). A unique advantage of this technique is the ability to study the real‐time kinetics of very early steps in the initiation process. To examine the effects of FIS on ternary complex formation at the tyrT promoter, we immobilized biotin end‐labelled promoter fragments containing the FIS sites to streptavidin surfaces in a BIAcore SPR machine (BIAcore AB). Two fragments were used in this study: a 197 bp wild‐type sequence containing the three FIS sites in helical register upstream of the tyrT promoter and a 203 bp mutant fragment with a 5 bp insertion at position −98 immediately upstream of FIS site II (Figure 1A). This insertion weakens the central FIS‐binding site (site II) and disrupts the helical register of sites I and III. Consequently FIS should no longer induce a coherent bend in the UAS. Functionally the mutation prevents the formation of a FIS‐dependent polymerase–promoter complex, as observed by gel retardation (Lazarus and Travers, 1993; Muskhelishvili et al., 1995).
Binding of proteins was monitored after their injection into the flowcell containing the surface‐immobilized DNA fragments. SPR analysis of the binding of RNA polymerase alone to the wild‐type and mutant fragments revealed that the enzyme has a 10‐fold higher affinity for the wild‐type than for the mutant promoter. This is illustrated by an enhanced overall association rate, leading, however, to final complexes of comparable stability (Table I). We therefore conclude that the rate of polymerase–promoter complex formation at the wild‐type promoter is 10 times as rapid as at the mutant promoter and that this difference is due uniquely to the presence of a 5 bp insertion at position −98 in the UAS.
By analysing the kinetics of FIS binding (Figure 1B), we assumed that the three FIS sites in the UAS are characterized by two distinct affinities (sites I and III being of higher affinity than site II, Lazarus and Travers, 1993). The calculated binding constants are shown in Table II. The results are consistent with FIS saturating sites I and III on both fragments but only poorly binding site II on the mutant fragment.
The formation of a ternary complex between FIS, polymerase and the promoter DNA reached a steady‐state equilibrium at the mutant promoter (Figure 1C), but at the wild‐type promoter an anomalous profile was obtained in which after reaching a maximum value the signal then decreased during the injection of proteins. Such a profile may be indicative of an evolving interaction in which the rapidly attained steady‐state shifts to a final equilibrium state that is different from that originally established. In this particular case, the ternary complex between FIS, polymerase and the wild‐type promoter forms more rapidly than at the mutant promoter and then undergoes a transition to a more stable complex. This kinetic profile was observed only on the simultaneous addition of FIS and polymerase. With polymerase alone, a profile consistent with normal steady‐state binding was obtained (data not shown). We note that the observed reduction in signal measured by SPR takes place in the continued presence of free FIS and polymerase and is greater than that observed during the dissociation phase at the end of injection. This phenomenon could be due either to an alteration of the conformation of the complex or to an effectively irreversible dissociation of one or more of the components of the complex.
The data obtained by SPR are consistent with our previous findings (Muskhelishvili et al., 1995) that the wild‐type and +5 mutant tyrT promoters differ in their ability to support FIS‐dependent trapping of polymerase. In addition, these data imply sequential and unidirectional effects of FIS at the tyrT promoter: an initial facilitation of polymerase binding followed by a structural change in the complex.
Destabilization of polymerase–promoter complexes by FIS
To investigate further the nature of the transition observed after the initial formation of the polymerase–FIS–DNA ternary complex at the wild‐type tyrT promoter, we carried out DNase I footprinting of FIS–polymerase complexes under experimental conditions close to those used for the SPR measurements. In the time‐course experiment, we observed a substantial weakening of the protection by polymerase but not by FIS (Figure 2A). The lessening of protection by polymerase proceeded more rapidly at the wild‐type than at the mutant promoter (compare lanes at 30 s for the wild‐type with the same for the mutant). This effect was not due to the occlusion of the promoter by FIS because no FIS‐specific hypersensitive sites within the promoter region were observed (G.Muskhelishvili, unpublished observations). Displacement of stably bound polymerase molecules did not, however, preclude contacts made by polymerase in the vicinity of the −35 region, as indicated by the retention of the strong DNase I hypersensitivity at position −37 (Figure 2A).
If FIS destabilizes polymerase, this should be reflected in reduced amounts of transcript produced if the transcription were initiated with a delay after addition of FIS. We tested this possibility in a runoff assay by adding all four nucleoside triphosphates to incubation mixtures for a fixed time but at different intervals after mixing FIS and polymerase with the promoter DNA (Figure 2B). This experiment showed that within 20 s, FIS reduced transcription from the wild‐type promoter by nearly 60%, a value that was only attained at later times at the mutant promoter. Taken together, these results suggest that the initial recruitment of polymerase by FIS at the tyrT promoter is followed by a rapid weakening of polymerase–promoter contacts in a majority of the complexes formed.
Both the SPR measurements and the solution transcription and cleavage protection experiments indicate that complexes at the mutant promoter are more resistant to destabilization by FIS than those at the wild‐type promoter. However, the apparent extent of this difference appears to depend on the method used. We note that the local environment of the immobilized DNA in the SPR experiments is significantly different from that of DNA free in solution, and this difference could contribute to observed differences in apparent residence times. Thus, although both conditions show qualitatively that the ternary complex is destabilized, precise quantitative comparisons between SPR and other methods may not, in this case, be justified.
FIS activates sequential steps in the initiation process
The experiments described above indicate that the formation of a FIS–polymerase–DNA ternary complex at the wild‐type promoter is followed by changes in the structure of the complex and provide direct evidence that FIS can affect sequential steps in the dynamic transitions undergone by the complex. To assess the relevance of these changes to the initiation process, we chose different conditions that allowed us to distinguish the effects of FIS on the initial binding of polymerase, on promoter opening and finally on polymerase escape.
We first analysed polymerase–promoter complex formation at 30°C and elevated salt concentrations (140 mM), conditions known to impair the transition from the closed to open complex at the rrnB P1 promoter (Ohlsen and Gralla, 1992b). Using DNase I as a probe for complex formation, we observed that under these conditions the interaction of polymerase with both the wild‐type and mutant promoter fragments was characterized solely by an enhanced DNase I cleavage at position −37, with little or no protection apparent within the remainder of the polymerase‐binding site (Figure 3A and B, arrowheads). However, upon addition of FIS, protection was apparent at the wild‐type but not the mutant promoter (Figure 3B), although in the latter case the enhancement of cleavage at −37 was increased. The downstream limit of the observed protection varied in different experiments between positions +8 and +17 as mapped by using DNA fragments of different lengths. The former value is consistent with the limit of the initial or closed complex formed at the rrnB P1 promoter but the latter does not extend to the +25 limit of the open complex on the same promoter (Ohlsen and Gralla, 1992a). This result confirms our previous conclusion that under restrictive conditions FIS site I alone is insufficient to stabilize polymerase binding at the tyrT promoter (Muskhelishvili et al., 1995).
To test whether FIS affected subsequent steps in the initiation process, we then monitored the effect of FIS on promoter opening. On addition of the two nucleoside triphosphates necessary for the synthesis of the first dinucleotide bond, RNA polymerase forms comparatively stable complexes at both the rrnB P1 (Gourse, 1988; Ohlsen and Gralla, 1992a) and tyrT (Küpper et al., 1975; Debenham, 1979) promoters. These complexes, termed initiation complexes, are characterized by a high reactivity of the promoter DNA in the −10 hexamer region to permanganate, a reagent that is specific for untwisted DNA (Gralla et al., 1993), and by a DNase I footprint extending to near position +25 (Gourse, 1988; Ohlsen and Gralla, 1992a; G.Muskhelishvili, unpublished observations).
By using a high molar ratio of RNA polymerase to DNA in the presence of initiating nucleoside triphosphates so that initial complex formation at both promoters was independent of FIS, we asked whether FIS influenced the reactivity to potassium permanganate of thymine residues around the −10 region and transcription startpoint. At 140 mM salt concentration, addition of FIS substantially increased permanganate reactivity of thymines within the −10 hexamer region (Figure 4A, positions −12 and −9) at the wild‐type but only to a slight extent at the +5 mutant promoter (Figure 4B). The observation that FIS increases the accessibility of this region to permanganate suggests an increase in the extent of untwisting of DNA within the −10 region necessary for promoter opening. Again, this effect requires all three FIS sites to be positioned in helical register.
The regulatory nucleotide ppGpp inhibits promoter opening at the rrnB P1 promoter (Ohlsen and Gralla, 1992a) but FIS is known to override the negative effect of ppGpp on transcription initiation at the tyrT promoter (Lazarus and Travers, 1993). We therefore asked whether FIS could overcome the effect of ppGpp on promoter opening at the tyrT promoter. We observed that the addition of ppGpp prevented the enhancement of the permanganate reactivity in the −10 region by polymerase alone and that FIS partially overcame the negative effect of ppGpp (Figure 4C). This effect of FIS was apparent at both the wild‐type and +5 mutant promoters.
We next asked whether FIS could affect any reaction steps subsequent to the formation of an initiation complex. Since the addition of heparin destabilizes the binding of FIS to site II (G.Muskhelishvili, unpublished observations), we could not use this compound to remove unstable pre‐initiation complexes. We therefore pre‐formed initiation complexes by the addition of the two nucleoside triphosphates, GTP and CTP, necessary for the synthesis of the first dinucleotide bond. To the pre‐formed initiation complexes we added UTP to allow more extensive RNA synthesis, up to a nonanucleotide (Figure 5A). Addition of this nucleotide alone further increased the permanganate reactivity of the bases within the −10 hexamer region at the wild‐type promoter and increased the permanganate reactivity of the base at position +1 (Figure 5B and C), indicating a conformational alteration of the complex. Quantitation of the extent of permanganate reactivity within the −10 hexamer region (Figure 5C) showed that on addition of UTP the signal obtained after 10 s for the bases at −9 and −12 with polymerase alone (2.4 ± 0.6) significantly increased in the presence of FIS (3.7 ± 0.7) at the wild‐type but not at the mutant tyrT promoter. These results suggest that in the presence of UTP, binding of FIS to helically phased sites in the UAS facilitates a conformational transition of initiation complexes.
To confirm that this effect of FIS was related to the efficiency of transcription initiation, we carried out a runoff transcription assay under similar conditions. First, initiation complex formation was allowed in the presence of GTP and CTP and then [α‐32P]UTP and ATP were added. FIS markedly increased the amount of the synthesized product at the wild‐type, but not at the mutant promoter (Figure 5D). This result is consistent with FIS stimulating a rapid transition of the complexes to the elongation mode. Again, this effect requires the wild‐type configuration of three FIS‐binding sites in UAS.
We have demonstrated that the FIS–polymerase nucleoprotein complex formed at the tyrT promoter is a dynamic structure which undergoes conformational transitions driven by FIS dimers bound to the UAS. It thus appears that, in contrast to other prokaryotic transcriptional activators, FIS activates transcription initiation by enabling efficient polymerase recruitment and also by facilitating promoter opening and subsequent post‐initiation events.
Sequential effects of FIS on transcription initiation
We have shown previously that FIS forms a nucleoprotein complex with RNA polymerase at the tyrT promoter, a process which requires the participation of three FIS dimers (Muskhelishvili et al., 1995). We have now shown that under restrictive conditions (30°C, 140 mM KCl) FIS promotes the establishment of a polymerase–promoter complex at the wild‐type, but not the +5 mutant promoter (Figure 3). Similarly, SPR measurements show that the overall rate of formation of a FIS–polymerase complex is higher at the wild‐type than at the mutant promoter. These results confirm our previous findings and show that under these conditions FIS recruits RNA polymerase to the tyrT promoter. This observation is similar to that of Bokal et al. (1995) who showed that FIS facilitated the initial binding of polymerase to the rrnB P1 promoter. However, whereas recruitment at the rrnB P1 promoter required only the proximal FIS‐binding site, this site, especially under restrictive conditions, is not sufficient at the tyrT promoter. Although the properties of the two promoters clearly differ in this respect, it is unclear whether the observed difference is biologically relevant or is simply a consequence of differences in assay conditions.
FIS also facilitates a second step in the initiation process, the untwisting of DNA in the −10 region. Again this effect is strong at the wild‐type but barely apparent at the +5 mutant promoter. Since the extent of untwisting is similar to that observed in other polymerase initiation complexes, we infer that FIS is promoting initiation complex formation. This view is also consistent with the antagonistic effects of FIS and ppGpp, a nucleotide which is known to block the transition to the initiation complex at the rrnB P1 promoter (Ohlsen and Gralla, 1992a). FIS partially counteracts the negative effect of ppGpp on untwisting but, interestingly, this effect is observed with both wild‐type and mutant promoters, suggesting that the intact UAS may not be required in the presence of the inhibitory nucleotide. Further genetic studies are under way to clarify this point.
At a higher temperature (37°C) FIS weakens the interaction of polymerase with the promoter DNA, an effect again requiring the participation of all three FIS‐binding sites in the UAS. In the absence of nucleoside triphosphates, this results in the dissociation of bound polymerase. However, under conditions which allow RNA chain elongation, FIS facilitates both post‐initiation structural changes in the −10 region and also transcription itself. These effects are quantitatively similar to the FIS‐induced enhancement of transition of open to transcribing complexes observed at the rrnD P1 promoter (Sander et al., 1993).
The ability of FIS to stimulate sequential steps in the initiation process at the tyrT promoter in vitro is consistent with the otherwise disparate observations that it promotes initial complex formation at the rrnB P1 promoter (Bokal et al., 1995) but increases the rate of both promoter opening and polymerase escape at the rrnD P1 promoter (Sander et al., 1993). More compellingly, this property provides an explanation for the observation that in vivo FIS stimulates expression from both down and up polymerase‐binding site mutants but not from the wild‐type tyrT promoter (Lazarus, 1992; Lazarus and Travers, 1993; H.Auner and G.Muskhelishvili, unpublished observations). We surmise that in the absence of FIS, initiation at the wild‐type promoter is finely tuned so that under optimum conditions the different steps in the initiation process are kinetically coordinated, i.e. no one step is strongly rate‐limiting. The role of FIS in such a situation would be to act as a facultative activator overcoming any kinetic bottlenecks caused by substrate or polymerase limitation. Similarly both up and down promoter mutations could also create kinetic blocks (Ellinger et al., 1994a) which again could be relieved by FIS.
Certain prokaryotic activators have the potential to activate different steps dependent on their placement with respect to the polymerase‐binding sites. For example, the cAMP receptor protein (CRP) accelerates polymerase recruitment at the lac promoter (Malan et al., 1984), isomerization to the open complex at the gal promoter (Herbert et al., 1986) and polymerase escape at the malT promoter (Menendez et al., 1987). However, to our knowledge, FIS is the first example of a prokaryotic transcriptional activator that is involved throughout the initiation process.
Active role of DNA microloops
As measured by SPR in the absence of FIS, the wild‐type tyrT promoter has an ∼10‐fold higher affinity for RNA polymerase than the +5 mutant promoter. This result is comparable with the 14‐fold enhancement of association rate conferred by an intact UAS at the rrnB P1 promoter (Newlands et al., 1991) and implies that at the tyrT promoter, sequences upstream of position −98 are necessary for full factor‐independent UAS function in vitro. One interpretation of this extended sequence requirement is that the tyrT UAS forms a microloop making an additional contact with RNA polymerase upstream of the 5 bp insertion point (Muskhelishvili et al., 1995). The existence of such loops has been inferred from the enhancement of promoter activity by upstream curved DNA (Bracco et al., 1989; Gartenberg and Crothers, 1991; Ellinger et al., 1994b) and from the activation of the λ pL and malT promoters by the DNA‐bending protein IHF (Giladi et al., 1990; Déthiollaz et al., 1996). More direct evidence for an upstream polymerase contact at the lac UV5 promoter has also been presented (Buckle et al., 1992). We suggest that the 5 bp insertion mutation alters the phasing of the anisotropically flexible tyrT UAS region (Drew and Travers, 1985) and so reduces, but does not necessarily eliminate, the probability of loop formation.
How does FIS mediate its effects on the transcription initiation process? The coherent DNA bending induced by FIS in the UAS could increase both the probability of forming a microloop and its subsequent stability. Such an effect would be consistent with the inability of the +5 mutant to support the formation of a FIS–polymerase complex (Muskhelishvili et al., 1995) or to promote FIS‐dependent DNA untwisting in the −10 region. Similarly, the FIS dependence of post‐initiation events at the wild‐type promoter implies that the integrity of the loop is maintained during the initial stages of transcription elongation. Mechanistically, the role of FIS in facilitating the initiation process could be explained most easily by assuming that FIS stabilizes a left‐handed writhe. In this model, the writhed microloop captures the polymerase in the initial complex, and then a rotation of RNA polymerase writhes the loop in a right‐handed sense, thereby generating torsion in the microloop (Figure 6). FIS subsequently drives a reversion to left‐handed writhe. This motion both transmits untwisting to the separate topological domain formed by the initiation bubble and accommodates the negative superhelicity generated upstream by the movement of the elongation bubble. In this model, torsional transmission could be mediated by either direct FIS–polymerase contacts (Muskhelishvili et al., 1995) or polymerase contacts with UAS DNA or, alternatively, by both types of contacts.
We and others (Gosink et al., 1993; Muskhelishvili et al., 1995) have observed previously that high concentrations of FIS can compete with RNA polymerase for its binding site at the rrnB P1 and tyrT promoters. We have now shown here that FIS can destabilize pre‐formed complexes, as indicated by a reduction in the SPR signal (Figure 1), by the loss of an extensive polymerase footprint and by loss of transcriptionally productive complexes (Figure 2). However, under these conditions, the enhanced DNase I cleavage immediately upstream of the −35 region suggests that polymerase can still interact with and distort the DNA at this position. Unlike protection, a protein‐induced enhanced DNase I cleavage signal may only require a transient distortion to be detectable and is not necessarily indicative of high occupancy by the protein. It seems unlikely that the FIS‐induced destabilization of polymerase binding we have reported here is a consequence of competition between FIS and polymerase since we observe no FIS‐related footprint within the polymerase‐binding region under our assay conditions. At higher FIS concentrations, invasion of this region by FIS is readily apparent (G.Muskhelishvili, unpublished observations). An alternative possibility is that the structural transitions in the nucleoprotein complex that occur between the initial and initiation complexes may directly drive the observed FIS‐dependent destabilization of polymerase binding.
The rapid synthesis of stable RNA species is a prerequisite for the efficient growth of E.coli. Such optimized synthesis requires a concomitant optimization of the initiation process, from the initial capture of polymerase by the promoter to its subsequent escape as an actively transcribing enzyme. The ability of FIS to overcome the barriers to differing rate‐limiting steps in initiation is consistent with the notion that the primary biological role of FIS is to optimize the rate of transcription initiation at stable RNA promoters under otherwise non‐ideal conditions (Lazarus and Travers, 1993; Muskhelishvili et al., 1995). However, if in vivo conditions were sufficiently unfavourable, for example if concentrations of the initiating triphosphates were low, FIS potentially could abort initiation by forcing the dissociation of bound polymerase. Taken together, these results suggest that FIS functions as a molecular machine which optimizes the turnover of polymerase holoenzyme at the tyrT promoter. The ability of FIS to stimulate both the assembly of the transcription complex and the subsequent promoter opening parallels its function in promoting Gin‐mediated recombination. The binding of FIS to the recombinational enhancer is thought to facilitate both the assembly of the synaptic complex (Merker et al., 1993) and the subsequent DNA untwisting at the sites of strand exchange (Klippel et al., 1993). We note that the mechanism of torsional transmission inferred for promoting transcription initiation would provide a means for channelling the free energy of negative supercoiling, thereby localizing untwisting at biologically relevant sites.
Materials and methods
Biotinylated DNA substrates
The uniquely end‐biotinylated wild‐type and the +5 mutant tyrT extended promoter fragments (positions −150 to +47 and −155 to +47 respectively) were obtained by PCR (Saiki, 1989) using the 5′‐biotinylated primer R‐bio (5′‐CACCACGGGGTAATGCTTT‐3′), the primer UAS‐L (5′‐CTTTGTTTACGGTAATCGAACG‐3′) and the tyrT promoter constructs ptyrTΔ150 and ptyrTΔ150+5 (Lazarus, 1992; Lazarus and Travers, 1993) as templates for amplification respectively. In these fragments, the biotinylated terminus was downstream of the transcription startsite. The +5 mutant refers to the promoter construct bearing a 5 bp insertion at position −98 which impairs the FIS site II and changes the helical phasing between FIS sites I and III (Lazarus and Travers, 1993; Muskhelishvili et al., 1995).
Surface plasmon resonance (SPR)
SPR measurements were conducted using a BIAcore instrument from BIAcore AB. The units of measurement are expressed in resonance units (RUs) where a change of 10−4 degrees is equivalent to a change of 1 RU and the machine has an effective dynamic range from 3–4 RUs to 30 000 RUs. The actual response in RU as a function of the change in surface molecule depends to an extent upon the differential refractive index of the solute, but for many globular proteins 1 kRU is equivalent to a change in surface concentration of ∼1 ng/mm2.
Immobilization of DNA fragments. The uniquely end‐biotinylated 197 bp wild‐type and the 203 bp +5 mutant tyrT extended promoter fragments (0.125 μg/ml) in 75 μl were injected independently across streptavidin‐pre‐treated dextran sensor surfaces in situ in the BIAcore apparatus at 5 μl/min in 20 mM Tris–HCl, pH 7.9, 50 mM NaCl, 50 mM KCl, 0.1 mM dithiothreitol (DTT), 0.005% surfactant P20 (BIAcore AB), at 37°C. In the experiments shown, ∼7.39×10−16 mol of wild‐type promoter DNA was immobilized at the surface (equivalent to an effective concentration in the dextran of 7.4 μM) and 1.18×10−15 mol (11.8 μM) of the mutant DNA.
Protein binding. FIS or RNA polymerase singly or in combination were applied at various concentrations to the different immobilized surfaces in 20 mM Tris–HCl, pH 7.9, 50 mM NaCl, 50 mM KCl, 0.1 mM DTT, 0.005% surfactant P20 (Biosensor Pharmacia), at 37°C. The surface was regenerated by washing with a 10 ml pulse of 1 M NaCl for 2 min, which removed all bound protein.
Interpretation of sensorgrams. In order to obtain the rates associated with the formation (ka) and dissociation (kd) of a given complex, sensorgrams were fitted to the algorithms provided by the BIAcore instrumentation. For the dissociation process (kd), the rate of change of resonance units (R in RUs) as a function of time was fitted to a simple exponential (Rt= R0exp + Rdrift). The association phase (ka) was described by the equation:
The expected response Rt as a function of maximal analyte binding capacity (Rmax) is calculated as a function of the concentration (C) of added soluble protein. The bulk contribution is made by the sample refractive index (Rbulk). Careful temperature control minimized the baseline drift (Rdrift).
DNase I footprinting
DNase I footprinting was performed with tyrT promoter fragments uniquely radiolabelled at the bottom strand as previously described (Muskhelishvili et al., 1995). The 197 bp wild‐type and 203 bp mutant DNA fragments were uniquely end‐labelled by PCR amplification using radioactively 5′ end‐labelled primer R3 (5′‐CACCACGGGGTAATGC‐3′) and primer UAS‐L (see above). The primers R3 and S90 were radiolabelled using [γ‐32P]ATP (NEN; 3000 Ci/mmol) and T4 polynucleotide kinase. The ptyrΔ50 and ptyrΔ50+5 constructs (see above) were used as templates in these PCR reactions. The fragments obtained were purified by PAGE using a neutral 0.5× TBE gel. Unless otherwise indicated, the incubation mixtures contained 10 mM Tris–HCl, pH 7.9, 0.1 mM DTT, 0.005% Triton X‐100, NaCl (as indicated) and various concentrations of polymerase and FIS in a 20 μl total volume. The reaction was initiated by adding polymerase, or FIS and polymerase, to a mixture containing DNA and other ingredients. Before mixing, all the components were pre‐equilibrated for 5 min at the required temperature. After incubation for different time intervals, a freshly prepared mixture of DNase I and MgCl2 (adjusted to the required temperature) was added to 5 μg/ml and 10 mM final concentrations respectively. The reaction was stopped after 10 s by adding 80 μl of the solution containing 0.5% SDS and 50 mM EDTA. After digestion by proteinase K for 45 min at 42°C, the samples were deproteinized by phenol extraction and the aqueous phase precipitated with ethanol. The pellets were washed with 70% ethanol, dried, dissolved in the loading dye and analysed on 6% sequencing gels.
Potassium permanganate reactivity assay
The reactions for potassium permanganate reactivity assays were assembled and processed similarly to those used for DNase I footprinting unless otherwise indicated. GTP and CTP were added to 1 mM each and, where used, UTP to 50 μM and ppGpp to 100 μM. The reaction was initiated by adding only polymerase, or FIS and polymerase, to a mixture containing radiolabelled DNA. Before mixing, all the components were pre‐equilibrated at the required temperature. After the incubation, 2 μl of 100 mM permanganate solution was added to 20 μl reaction mixtures containing DNA and proteins for either 10 s or 1 min as indicated in the figure legends. The reactions were stopped by addition of 2 μl of 14 M β‐mercaptoethanol, 8 μg of sonicated salmon sperm DNA and sodium acetate to 0.3 M, precipitated with 3 volumes of ice‐cold ethanol and washed with 70% ethanol. The pellets were resuspended in 100 μl of 10% piperidine and incubated at 90°C for 20 min. Then LiCl was added to 0.5 M, the DNA precipitated with 3 volumes of ice‐cold ethanol and washed at least twice with 100% ethanol. The pellets were dried, dissolved in the loading dye and analysed on 6% sequencing gels. The signals due to permanganate reactivity of bases were quantified by using the PhosphorImager (Storm 840, Molecular Dynamics). The absolute values of the signals obtained by this procedure may vary and need to be normalized for comparative analysis. We normalized the reactivity of bases in different lanes by using the ratios of the sum of signals obtained for bases at −9 and −12 divided by the value obtained for the base at −14 (which is the first thymine outside of the −10 region) in each lane. The ratios obtained were averaged and subjected to statistical analysis. The ratio obtained for the naked DNA at both promoters was similar and varied within a narrow range (1.79 ± 0.22).
In vitro transcription assay
The 299 bp wild‐type and 304 bp mutant tyrT DNA templates used in the runoff assay were obtained by EcoRI–NsiI digestion of the ptyrTΔ150 DNA and ptyrTΔ150+5 DNA followed by agarose gel purification of the respective fragments. The runoff transcription assays were performed at 37°C in a buffer containing 10 mM Tris–HCl, pH 7.9, 0.1 mM DTT, various concentrations of NaCl, 10 mM MgCl2, 5 nM of the EcoRI–NsiI tyrT DNA fragment, various concentrations of polymerase and FIS, 1 mM each of GTP and CTP, 0.05 mM [α‐32P]UTP and 0.4 mM ATP. The reactions were stopped after different time intervals by directly adding equal amounts of the formamide loading dye to aliquots of incubation mixtures. The reaction products (145 bp) were analysed on 6% sequencing gels and quantified by using the PhosphorImager (Storm 840, Molecular Dynamics).
This work was supported by the Deutsche Forschungsgemeinshaft through SFB190.
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