Regulated communication between the upstream face of RNA polymerase and the β′ subunit jaw domain

Abstract

We used bacteriophage T7-encoded transcription inhibitor gene protein 2 (gp2) as a probe to study the contribution of the Escherichia coli RNA polymerase (RNAP) β′ subunit jaw domain—the site of gp2 binding—to activator and ATP hydrolysis-dependent open complex formation by the σ⁵⁴-RNAP. We show that, unlike σ⁷⁰-dependent transcription, activated transcription by σ⁵⁴-RNAP is resistant to gp2. In contrast, activator and ATP hydrolysis-independent transcription by σ⁵⁴-RNAP is highly sensitive to gp2. We provide evidence that an activator- and ATP hydrolysis-dependent conformational change involving the β′ jaw domain and promoter DNA is the basis for gp2-resistant transcription by σ⁵⁴-RNAP. Our results establish that accessory factors bound to the upstream face of the RNAP, communicate with the β′ jaw domain, and that such communication is subjected to regulation.

Introduction

The central enzyme of gene expression, the DNA-dependent RNA polymerase (RNAP), is a complex and highly regulated molecular machine that is conserved between eukaryotes, archaea and bacteria (Ebright, 2000). In Escherichia coli, the catalytically competent core RNAP (E, subunit composition α₂ββ′ω) binds one of the seven sigma (σ) subunits, at the so-called ‘upstream face' of the core RNAP, to form an RNAP holoenzyme (Eσ) that is capable of promoter-specific transcription (Murakami et al, 2002; Chung et al, 2003; Bushnell et al, 2004).

There are two distinct types of σ factors. RNAP holoenzymes containing the major σ⁷⁰-type factors (named after the prototypical housekeeping σ of E. coli, σ⁷⁰) recognize promoters characterized by conserved sequence elements centered around 35 and 10 base pairs upstream of the transcription start site at +1. Closed promoter complexes formed by RNAP holoenzymes containing σ⁷⁰-type factors readily isomerize into transcription-competent open promoter complexes. RNAP holoenzymes containing evolutionarily unrelated σ⁵⁴-type factors recognize promoters characterized by conserved sequence elements located 24 and 12 base pairs upstream of the transcription start site and form closed complexes that cannot spontaneously isomerize into open complexes (Buck et al, 2000). Conversion of Eσ⁵⁴ closed complexes to open complexes requires ATP hydrolysis by a specialized activator protein belonging to the AAA (ATPases Associated with various cellular Activities) family (Zhang et al, 2002). The activator binds DNA upstream of the σ⁵⁴ promoter, interacts with the Eσ⁵⁴ closed complex and induces conformational changes that lead to the formation of a transcription-competent open complex (Buck et al, 2000; Zhang et al, 2002).

Core RNAP surfaces that are responsible for formation and maintenance of open promoter complexes are not yet fully characterized. For σ⁷⁰-dependent transcription, the interaction between the β′ jaw domain which is located at the so-called ‘downstream face' of the RNAP (residues 1149–1190 of E. coli β′ subunit; Figure 1A) and the double-stranded DNA downstream of the transcription initiation start point has been shown to be important for stable open complex formation (Ederth et al, 2002). Binding of the bacteriophage T7-encoded gene 2 protein (gp2) to the β′ jaw domain strongly inhibits promoter complex formation by Eσ⁷⁰, further underscoring the importance of the β′ jaw domain in transcription (Nechaev and Severinov, 1999). In eukaryotic RNAPII, the jaw lobe of RPB1, a homologue of β′, forms part of the DNA-binding cleft and contacts bases in the downstream DNA (Bushnell et al, 2004). The role of the β′ jaw during σ⁵⁴-dependent transcription is not known. In this work, we have used gp2 as a probe to study the contribution of the E. coli RNAP β′ jaw domain to transcription by Eσ⁵⁴. We show that, unlike Eσ⁷⁰ transcription, activator-dependent Eσ⁵⁴ transcription is not inhibited by gp2. We provide evidence that activation induces a promoter DNA-dependent conformational change involving the β′ jaw domain, which allows Eσ⁵⁴ to escape inhibition by gp2 and to engage promoter DNA.

Figure 1.

Proteins used in this study. (A) The crystal structure of the T. aquaticus core RNAP with the β′ jaw domain highlighted in green space fill and the E1188K mutation indicated in yellow. The β, β′, ω and αI and αII subunits are color coded as shown. The red sphere indicates the catalytic center. The downstream and upstream faces of the RNAP are labeled. (B) Domain organization of K. pneumoniae σ⁵⁴. The major activator, core RNAP and promoter DNA-binding determinants are shown. In Region III, residues 329–346 have been shown to UV-crosslink to DNA (X-link) (Cannon et al, 1994), residues 366–386 constitute the putative helix–turn–helix DNA-binding motif (HTH) (Merrick and Chambers, 1992) and residues 454–463 constitute a highly conserved patch (RpoN box) that interacts with the −24 consensus promoter element of σ⁵⁴-dependent promoters (Burrows et al, 2003).

Mutations in σ⁵⁴ regulatory regions I and III (Figure 1B) result in Eσ⁵⁴ that is capable of transcription in the absence of the activator (Casaz et al, 1999; Chaney and Buck, 1999; Wang and Gralla, 2001; Wigneshweraraj et al, 2002). We show that the integrity of Region I is important for the binding of gp2 to the β′ jaw domain, and thereby present evidence for a functional and/or structural link between σ⁵⁴ Region I and the β′ jaw domain. Like Eσ⁷⁰ transcription, activator-independent transcription by mutant Eσ⁵⁴ is inhibited by gp2. We demonstrate that gp2 inhibits activator-independent transcription by preventing the step(s) leading to DNA melting. Overall, we present evidence for an internal conformational signaling pathway in Eσ⁵⁴ that involves σ⁵⁴ Region I and the β′ jaw domain, and provide clues as to how σ⁵⁴ bound at the upstream face of the RNAP could regulate the activity of RNAP by controlling the β′ jaw function.

Results

Activator-dependent Eσ⁵⁴ transcription is not inhibited by T7 gp2

We compared the effect of gp2 on Eσ⁷⁰ and Eσ⁵⁴ transcription in a single-round transcription assay using the lacUV5 (for Eσ⁷⁰ transcription) and the Sinorhizobium meliloti nifH (for Eσ⁵⁴ transcription) promoters. To activate Eσ⁵⁴ transcription, we used a fragment of the E. coli σ⁵⁴ transcription activator phage shock protein F (PspF_1–275), which lacks the DNA-binding domain and efficiently activates transcription from solution (Cannon et al, 2003). RNAP holoenzymes were reconstituted with purified σ proteins and combined with gp2 prior to the addition of promoter DNA. As a control, we used holoenzymes reconstituted with core RNAP harboring the E1188K substitution in the β′ jaw domain. This mutation prevents gp2 binding and makes Eσ⁷⁰ transcription resistant to gp2 (Nechaev and Severinov, 1999; Figure 1A). As expected, transcription from lacUV5 was strongly inhibited by gp2 in reactions containing wild-type Eσ⁷⁰, but not in reactions containing ^E1188KEσ⁷⁰ (Figure 2A). In contrast, gp2 had no obvious effect on activated transcription from the S. meliloti nifH promoter by Eσ⁵⁴ (Figure 2B). Eσ⁵⁴ transcription from two other σ⁵⁴-dependent promoters (E. coli glnHp2 and pspA) was also resistant to gp2 (data not shown). The order of addition experiments did not reveal any detectable inhibition of Eσ⁵⁴-dependent transcription regarding whether gp2 was added before RNAP holoenzyme formation, after closed complex formation, or after open complex formation (data not shown). It therefore appears that activated Eσ⁵⁴ transcription is fully resistant to gp2.

Figure 2.

Eσ⁵⁴ is insensitive to inactivation by gp2. (A) Single-round transcription by Eσ⁷⁰ from the lacUV5 promoter in the presence of increasing amounts of gp2. (B) Single-round transcription by Eσ⁵⁴ from the S. meliloti nifH promoter in the presence of PspF_1–275, ATP and increasing amounts of gp2. In (A) and (B), empty boxes indicate the use of Eσ reconstituted with the gp2-resistant E1188K mutant core RNAP and filled boxes Eσ reconstituted with the gp2-sensitive wild-type core RNAP. The error range for the values graphed in (A) and (B) is within ±7%. (C) An autoradiograph of a 4.5% (w/v) native gel showing binding of ³²P-labeled gp2 to E and Eσ⁵⁴. The migration positions of gp2 (lane 1), E:gp2 (lane 2) and Eσ⁵⁴:gp2 (lane 4) are indicated. Protein components in each lane are indicated on top. (D) T7 gp2 binds to Eσ⁵⁴-closed promoter complexes. An autoradiograph of a 4.5% (w/v) native gel showing the binding of ³²P-labeled gp2 to E and Eσ⁵⁴ in the presence of streptavidin-labeled S. meliloti nifH early-melted promoter probe (lane 4). The positions of gp2 (lane 1), E:gp2 (lane 2), Eσ⁵⁴:gp2 (lane 3) and Eσ⁵⁴:DNA:gp2 (lane 4) complexes are shown and the protein and DNA components of each lane indicated (top of the figure). A cartoon of the biotin (B)- and streptavidin (S)-tagged early-melted promoter probe is shown. The positions of the transcription start site (+1) and the start-site distal (−24) and proximal (−12) consensus promoter elements of σ⁵⁴-dependent promoters are shown.

We considered the possibility that gp2 might not be able to interact with the β′ jaw domain in the presence of σ⁵⁴. However, a native gel assay using ³²P-labeled gp2 confirmed that gp2 interacted stably and specifically with Eσ⁵⁴ (Figure 2C). A radioactive band observed in reactions containing the wild-type core RNAP and gp2 corresponded to the RNAP core–gp2 complex (Figure 2C, lane 2) that was converted into a slightly slower migrating complex in reactions containing core RNAP, gp2 and σ⁵⁴ (Figure 2C, lane 4). As expected, no such complexes were observed in reactions that contained core RNAP or Eσ⁵⁴ harboring the E1188K substitution in β′ (Figure 2C, lanes 3 and 5). SDS–PAGE analysis of complexes shown in Figure 2C (lanes 2 and 4) confirmed the presence of σ⁵⁴ in the complex seen in lane 4, but not lane 2 (data not shown).

Additional experiments using gp2 modified with lysine-specific protein-cleaving reagent 2IT-FeBABE (Datwyler and Meares, 2000) confirmed that gp2 was proximal to the β′ jaw domain in the context of RNAP core, Eσ⁷⁰, and Eσ⁵⁴ (data not shown). We therefore conclude that, even though gp2 specifically interacts with the β′ jaw domain in the presence of either σ⁷⁰ or σ⁵⁴ (Nechaev and Severinov, 1999; Figure 2C), it only inhibits Eσ⁷⁰ transcription. The result thus suggests that the β′ jaw domain may make different interactions in transcription initiation by Eσ⁵⁴ compared to Eσ⁷⁰. Possibly, within Eσ⁵⁴ promoter complexes, the interaction between the downstream DNA and the β′ jaw causes gp2 to dissociate from RNAP.

T7 gp2 binds to Eσ⁵⁴ closed promoter complexes

To determine whether gp2 is able to bind to closed Eσ⁵⁴ promoter complexes, we combined ³²P-labeled gp2 with Eσ⁵⁴ complexes formed on an early-melted promoter probe that contains a heteroduplex segment near the −12 consensus promoter element and thus mimics the conformation of DNA in the closed complex (Cannon et al, 2003) (Figure 2D). An excess of early-melted promoter probe over Eσ⁵⁴ was used to ensure that most Eσ⁵⁴ was in the DNA-bound form. As can be seen, the mobility of Eσ⁵⁴ bound to ³²P-labeled gp2 changed in the presence of DNA (Figure 2D, compare lanes 3 and 4), indicating that ³²P-labeled gp2 binds to the preformed Eσ⁵⁴ closed promoter complex. Similar results were obtained when ³²P-labeled gp2 was incubated with Eσ⁵⁴ prior to the addition of the early-melted probe (data not shown). The binding results thus indicate that, unlike Eσ⁷⁰ closed promoter complexes (Nechaev and Severinov, 1999), Eσ⁵⁴ closed promoter complexes are not disrupted by gp2.

T7 gp2 binds Eσ⁵⁴ activator complex

We investigated whether gp2 remains bound to Eσ⁵⁴ upon interaction with the activator. Stable, nucleotide-dependent binding of activator to Eσ⁵⁴ can be measured by native PAGE in the presence of ADP-aluminum fluoride (ADP-AlF_x, where x is either 3 or 4), an analog of ATP in the transition state of hydrolysis (Chaney et al, 2001). We incubated the activator with ADP-AlF_x and Eσ⁵⁴ in the presence of ³²P-labeled gp2, and resolved the reaction products by native PAGE. As shown in Figure 3A, ³²P-labeled gp2 interacted with Eσ⁵⁴ when added either before or after Eσ⁵⁴-PspF_1–275:ADP-AlF_x complex formation (compare lanes 5 and 7). Similarly, ³²P-labeled gp2 interacted with Eσ⁵⁴ when ATP was substituted for ADP-AlF_x (data not shown). Therefore, gp2 has no effect on Eσ⁵⁴ activator interaction. Further, this result also excludes the site of gp2 binding on the β′ jaw domain as a major site or determinant of activator interaction.

Figure 3.

T7 gp2 does not bind an intermediate Eσ⁵⁴ promoter complex. (A) Activator–Eσ⁵⁴ interaction is not disrupted by T7 gp2. Autoradiograph of a 4.5% (w/v) native gel showing the ability of ³²P-labeled gp2 to bind a ternary Eσ⁵⁴:PspF_1–275:ADP-AlF_x complex (lanes 5 and 7). In lane 5, gp2 was added prior to Eσ⁵⁴:PspF_1–275:ADP-AlF_x complex formation and, in lane 7, gp2 was added after Eσ⁵⁴:PspF_1–275:ADP-AlF_x complex formation. The protein components in each lane and the migration positions of gp2, E:gp2, Eσ⁵⁴:gp2 and Eσ⁵⁴:PspF_1–275:ADP-AlF_x:gp2 are indicated. (B) T7 gp2 does not bind an early intermediate promoter complex. Silver-stained 4.5% (w/v) native gel (top panel) and the autoradiograph of the same gel (bottom panel) showing that gp2 does not interact with an early intermediate Eσ⁵⁴ promoter complex on streptavidin-tagged early-melted promoter probe in the presence of ³²P-labeled gp2 (see Materials and methods). The arrow in lanes 5 and 11 points to the migration position of Eσ⁵⁴:DNA:PspF_1–275:ADP-AlF_x complex. The migration positions of other protein–protein and protein–DNA complexes are as indicated. The protein and DNA components in each lane are shown on the top of the figure. (C) Single-round transcription assays from the supercoiled S. meliloti nifH promoter using ATP and either wild-type PspF_1–275 (lanes 1 and 2) or ^T86SPspF_1–275 (lanes 3 and 4) for activation. The reactions in lanes 1–4 were conducted using the gp2-sensitive wild-type core RNAP, whereas the reactions in lanes 4–8 were conducted using the gp2-resistant E1188K mutant core RNAP. The protein components in each lane are indicated on the top of the figure.

T7 gp2 does not bind intermediate Eσ⁵⁴ promoter complex

To determine whether activator-induced conformational changes in the Eσ⁵⁴ closed complex allow Eσ⁵⁴ transcription to escape inhibition by gp2, we tested whether gp2 is able to interact with intermediate Eσ⁵⁴ promoter complexes. The Eσ⁵⁴:DNA:PspF_1–275:ADP-AlF_x complex represents an early intermediate en route to open complex formation by Eσ⁵⁴ (Chaney et al, 2001; Cannon et al, 2003). We formed the Eσ⁵⁴:PspF_1–275:ADP-AlF_x complex in the presence of the early-melted promoter probe (present at a four-fold molar excess over Eσ⁵⁴ to ensure that most Eσ⁵⁴ was in the DNA-bound form), added ³²P-labeled gp2 and analyzed the reaction by native PAGE. Silver staining of the native gel revealed a band (Figure 3B, lane 5, marked with the arrow) whose formation was dependent on the presence of DNA (Figure 3B, lane 6) and PspF_1–275:ADP-AlF_x (Figure 3B, lane 4). Additional control reactions in the absence of core RNAP and σ⁵⁴ did not result in the formation of the band marked with the arrow in Figure 3B, lane 5 (data not shown). Therefore, we conclude that this band (marked with the arrow in Figure 3B, lane 5) must correspond to Eσ⁵⁴:DNA:PspF_1–275:ADP-AlF_x complex. However, the autoradiograph of the native gel did not reveal any radioactivity in this band (Figure 3B, compare lanes 5 and 11, marked with the arrow). We therefore conclude that ³²P-labeled gp2 is not able to stably interact with the Eσ⁵⁴:DNA:PspF_1–275:ADP-AlF_x complex. Identical results were obtained when Eσ⁵⁴ was incubated with ³²P-labeled gp2 prior to the addition of DNA and PspF_1–275:ADP-AlF_x (data not shown). Overall, the result indicates that activator-induced and promoter-dependent conformational changes in Eσ⁵⁴ prevent the interaction of gp2 with the β′ jaw domain and may thereby allow activated transcription by Eσ⁵⁴ to escape inhibition by gp2. Clearly, promoter DNA plays a critical role in inhibiting binding of gp2, since Eσ⁵⁴:PspF_1–275:ADP-AlF_x complex binds gp2 well (Figure 3A, lanes 5 and 7).

Activator triggers conformational change(s) involving the β′ jaw domain

Evidence that the interactions between the Eσ⁵⁴ closed complex and the activator somehow change the interaction between gp2 and the β′ jaw domain and allows Eσ⁵⁴ to escape inhibition by gp2 prompted us to investigate whether ATP hydrolysis-dependent ‘remodeling' of the Eσ⁵⁴ closed complex is the reason for the gp2 resistance of Eσ⁵⁴ transcription. We conducted single-round transcription assays from the S. meliloti nifH promoter using PspF_1–275 carrying a point substitution at the conserved position Thr⁸⁶ (T86S; ^T86SPspF_1–275). The mutant activator is defective for interaction with the Eσ⁵⁴ closed complex and is less efficient in ATP hydrolysis-dependent remodeling of Eσ⁵⁴ closed complexes (Chaney et al, 2001). As shown in Figure 3C, we found that gp2 inhibited Eσ⁵⁴ transcription activated by ^T86SPspF_1–275 by 6–7-fold (compare lanes 3 and 4). Similar levels of inhibition were observed in reactions using different activator mutants impaired for nucleotide binding and/or hydrolysis or Eσ⁵⁴ interaction (data not shown). The inhibition was not detected in control reactions containing gp2-resistant ^E1188KEσ⁵⁴ (Figure 3C, compare lanes 7 and 8). As expected, in reactions containing wild-type PspF_1–275, no inhibition of Eσ⁵⁴ transcription was observed (Figure 3C, compare lanes 1 and 2). Since the β′ jaw domain is not a direct target for the activator (Figure 3A and Wigneshweraraj et al, in preparation), we infer that, during open complex formation, the activator drives an ATP hydrolysis-dependent conformational change in the Eσ⁵⁴ closed complex that causes, directly or indirectly, dissociation of gp2. The mutant activator (^T86SPspF_1–275) is less efficient in coupling ATP hydrolysis to conformational changes in Eσ⁵⁴ closed complexes, which leads to gp2 sensitivity. Overall, the results point to activator- and ATP hydrolysis-dependent use of the β′ jaw domain during open complex formation by Eσ⁵⁴, which likely involves a β′ jaw domain-DNA interaction.

Activator-bypass transcription by Eσ⁵⁴ is inhibited by T7 gp2

The observation that activator and ATP hydrolysis-driven conformational changes involving the β′ jaw domain enable Eσ⁵⁴ to escape inhibition by gp2 prompted us to investigate whether activator-independent (termed activator-bypass) transcription by Eσ⁵⁴ is inhibited by gp2. We expected that, since no activator and ATP hydrolysis-driven conformational changes would occur during activator-bypass transcription, gp2 will be able to inhibit activator-bypass transcription by Eσ⁵⁴.

Certain substitutions in σ⁵⁴ regulatory regions I and III allow Eσ⁵⁴ in vitro transcription to occur without activation. In contrast to open promoter complexes formed by wild-type Eσ⁵⁴, which become heparin-resistant prior to initiation of RNA synthesis, activator-bypass mutant Eσ⁵⁴ promoter complexes become heparin-resistant only after the initiation of RNA synthesis (Wang and Gralla, 1996). To check the effect of gp2 on activator-bypass transcription, we conducted single-round transcription assays using S. meliloti nifH promoter and Eσ⁵⁴ reconstituted using an activator-bypass σ⁵⁴ carrying the R336A substitution in Region III (Figure 1A, Chaney and Buck, 1999; Wang and Gralla, 2001; Wigneshweraraj et al, 2001). The first step of activator-bypass transcription assay from the S. meliloti nifH promoter involves the incubation of Eσ⁵⁴ with the DNA template and GTP. Since the first three bases of the nifH RNA are G (Sundaresan et al, 1983), activator-bypass transcription initiation can occur under these conditions. In the second step of the assay, heparin and the remaining nucleotides are added to allow elongation of any initiated transcripts.

Activator-bypass transcription by Eσ⁵⁴_R336A was strongly inhibited by gp2 (Figure 4A) and the inhibition was as effective as that with Eσ⁷⁰ (compare Figures 2A and 4A). The inhibition of Eσ⁵⁴_R336A transcription is specific, since a double mutant Eσ⁵⁴ reconstituted from ^E1188KE and σ⁵⁴_R336A was resistant to gp2 (Figure 4A). The order of addition experiments revealed that gp2 inhibited Eσ⁵⁴_R336A transcription when added either before or after closed complex formation (Supplementary data Figure 1). Inhibition of transcription was not due to direct inhibition of closed complex formation by gp2, since ³²P-labeled gp2 was able to stably bind Eσ⁵⁴_R336A closed promoter complexes formed on the early-melted promoter probe (Figure 4B, lane 1). Once activator-bypass transcript initiation had occurred, gp2 had no effect on elongation by Eσ⁵⁴_R336A (Figure 4C). gp2 had a similar effect on activator-bypass transcription by RNAPs reconstituted with two additional Region III mutants, σ⁵⁴_F318A (Wigneshweraraj et al, 2002) and σ⁵⁴_K388A (Wang and Gralla, 2001) (data not shown). We therefore conclude that, during activator-bypass transcription by Eσ⁵⁴ with lesions in σ⁵⁴ Region III, gp2 binding interferes with the β′ jaw domain function during one or more steps that lead to activator-bypass open complex formation.

Figure 4.

Activator-bypass transcription from supercoiled S. meliloti nifH promoter by Eσ⁵⁴_R336A is inhibited by T7 gp2. (A) Titration of gp2 against Eσ⁵⁴_R336A reconstituted with gp2-resistant E1188K mutant core RNAP (empty boxes) and gp2-sensitive wild-type core RNAP (filled boxes). The error range for the values shown in graph is ±5%. An autoradiograph of a 4% (w/v) denaturing gel showing the inhibition of activator-bypass transcription by Eσ⁵⁴_R336A by gp2 is shown. (B) An autoradiograph of a 4.5% (w/v) native gel showing the binding of ³²P-labeled gp2 to Eσ⁵⁴ (lane 3) and Eσ⁵⁴_R336A (lane 1) promoter complexes formed on the S. meliloti nifH early-melted DNA probe. The migration positions of E:gp2, Eσ⁵⁴:gp2 and Eσ⁵⁴:DNA:gp2 are indicated. The protein and DNA components in each lane are shown on the top. (C) An autoradiograph of a 4% (w/v) denaturing gel showing that activator-bypass transcription by Eσ⁵⁴_R336A is not inhibited when gp2 is added after transcription initiation. Reactions in lanes 1 and 2 were conducted using the gp2-sensitive wild-type core RNAP, whereas the gp2-resistant E1188K mutant RNAP was used in reactions in lanes 3 and 4. The arrow indicates the 470 nucleotide transcript from the S. meliloti nifH promoter on pMKC28. The reaction schematic is shown on the top of each figure (see Materials and methods). (D) An autoradiograph of a 4% (w/v) denaturing gel showing that activated transcription by Eσ⁵⁴_R336A is not inhibited by gp2. The gel is labeled as in (C).

Activation allows Eσ⁵⁴_R336A to escape inhibition by gp2

We next conducted the reciprocal experiment and investigated whether activation allows Eσ⁵⁴_R336A to overcome inhibition by gp2. In this experiment, escape from inhibition was interpreted to mean that activator and ATP hydrolysis drive conformational changes within the promoter complex that prevent gp2 action, most likely by changing gp2 binding to the β′ jaw domain (see above). As can be seen in Figure 4D, activator-dependent transcription by Eσ⁵⁴_R336A was slightly inhibited by gp2 when gp2 was added either before (compare lanes 1 and 2) or after closed complex formation (Supplementary data Figure 2). However, in titration assays using increasing amounts of gp2 in the presence of Eσ⁵⁴_R336A, ATP and activator, the extent of inhibition of activator-dependent transcription by Eσ⁵⁴_R336A was significantly reduced (compare Figure 4A, lane 6 with Figure 4D, lane 2 and data not shown). Similarly, the addition of gp2 after activator-dependent open complex formation had no effect on Eσ⁵⁴_R336A transcription (Supplementary data Figure 2). Identical results were obtained with Eσ⁵⁴ reconstituted with F318A and K388A activator-bypass σ⁵⁴ proteins (data not shown). The results thus further confirm that, during open complex formation, activator-induced conformational changes lead to the loss of gp2 binding to the β′ jaw domain.

T7 gp2 prevents activator-bypass DNA melting by Eσ⁵⁴_R336A

The results presented above point to an activator- and ATP hydrolysis-dependent use of the β′ jaw domain during DNA melting and open complex formation, and suggest that gp2 could inhibit activator-bypass transcription by preventing RNAP conformational changes and DNA interactions required for efficient DNA melting. To test this idea, we formed Eσ⁵⁴_R336A promoter complexes on supercoiled S. meliloti nifH promoter in the absence or presence of gp2 under conditions where activator-bypass DNA melting should occur, and detected DNA melting with potassium permanganate (KMnO₄) (see Materials and methods). In all experiments described below, gp2 was added prior to promoter complex formation. As shown in Figure 5A, no promoter DNA melting was detected in reactions that contained wild-type Eσ⁵⁴, but lacked PspF_1–275 (compare lanes 4 and 5). In contrast, activator-bypass DNA melting (at position −8 on the nontemplate strand and position −12 on the template strand) was detected within promoter complexes formed with Eσ⁵⁴_R336A in the presence of GTP (recall that the first three bases transcribed from the S. meliloti nifH promoter are G) (Figure 5A, lane 7). PspF_1–275 moderately increased the efficiency of DNA melting by Eσ⁵⁴_R336A (Figure 5A, lane 8). In the presence of gp2, no DNA melting was detected in reactions containing Eσ⁵⁴_R336A and GTP (Figure 5A, lane 14). The extent of activator-dependent promoter melting by the wild-type Eσ⁵⁴ was unaffected by gp2 (Figure 5A, lane 12). In agreement with the observation that activation allows Eσ⁵⁴_R336A to escape inhibition by gp2, we are able to detect DNA melting by Eσ⁵⁴_R336A in the presence of gp2 and PspF_1–275 (Figure 5A, compare lanes 8 and 15). However, we note that the extent of DNA melting has been reduced in reactions containing gp2 and PspF_1–275 (lane 15) when compared to reactions containing no gp2 (lane 8).

Figure 5.

T7 gp2 prevents activator-bypass DNA melting within promoter complexes formed by Eσ⁵⁴_R336A. (A) An autoradiograph of a 10% (w/v) denaturing gel showing KMnO₄ probing of S. meliloti nifH promoter complexes (on pMKC28) (reaction conditions as shown on the top of the figure) formed by Eσ⁵⁴ and Eσ⁵⁴_R336A in the absence (lanes 1–8) and presence (lanes 9–16) of gp2. In (A), the DNA sequence of the S. meliloti nifH promoter is shown, with the asterisks indicating the thymine residues at position −8 on the nontemplate strand and −12, −9 and −6 on the template strand. (B) An autoradiograph of a 10% (w/v) denaturing gel showing DNase I probing of S. meliloti nifH promoter complexes (on pMKC28) (reaction conditions as shown on the top of the figure) formed by Eσ⁵⁴ and Eσ⁵⁴_R336A in the absence (lanes 1–6) and presence (lanes 7–11) of gp2. In (A) and (B), lanes 16 and 12, respectively, contain chain termination DNA-sequencing reactions conducted with pMKC28 and ddTTP. In (B), lane 6, the dotted line indicates the extended protection within Eσ⁵⁴_R336A promoter complexes in the presence of GTP. (C) An autoradiograph of a 10% (w/v) denaturing gel showing run-off transcription from a linear S. meliloti nifH promoter probe containing a heteroduplex segment between positions −12 and −1. The arrow indicates the 28 nucleotide run-off product. The marker lane contains a mixture of end-labeled S. meliloti nifH promoter DNA fragments.

DNase I footprinting of complexes formed by Eσ⁵⁴_R336A on supercoiled S. meliloti nifH showed clear protection of promoter DNA, both in the absence and presence of gp2 (Figure 5B, compare lanes 6 and 11). Thus, gp2 did not disrupt promoter complexes formed by Eσ⁵⁴_R336A under the assay conditions. In the footprint of the Eσ⁵⁴_R336A promoter complex, in the absence of gp2, a moderate extension of the footprint downstream of position −8 was observed (Figure 5B, lane 6; marked by the dotted line). This extended footprint is indicative of the activator-bypass isomerization of Eσ⁵⁴_R336A promoter complex in the presence of GTP (Figure 5B; compare lanes 5 and 6), and is also seen within wild-type Eσ⁵⁴ promoter complexes in response to activation (data not shown; Wang and Gralla, 1996; Wigneshweraraj et al, 2002). No extended footprint was observed in the presence of gp2 in reactions containing Eσ⁵⁴_R336A and GTP (Figure 5B; compare lanes 6 and 11). Thus, gp2 binding to the β′ jaw domain interferes with activator-bypass isomerization and stable promoter DNA melting by Eσ⁵⁴_R336A. In agreement with this, run-off activator-bypass transcription by Eσ⁵⁴_R336A from a heteroduplex S. meliloti nifH promoter (Cannon et al, 2000), where the DNA was stably pre-melted between positions −12 and −1, was unaffected by gp2 (Figure 5C, compare lanes 4 and 8).

Overall, the results strongly suggest that gp2 inhibits activator-bypass transcription by preventing step(s) leading to stable DNA melting and, by extension, suggest that the β′ jaw domain is required for DNA melting (possibly stabilizing the melted DNA) during Eσ⁵⁴ transcription.

σ⁵⁴ Region I determines β′ jaw conformation

The observation that the β′ jaw domain is involved in activator and ATP hydrolysis-dependent conformational changes during Eσ⁵⁴ transcription, but is not a direct binding target for the activator, prompted us to investigate the role of regulatory regions of σ⁵⁴ in determining the β′ jaw domain function during activator-dependent transcription. The amino-terminal regulatory Region I of σ⁵⁴ constitutes a binding site for the activator (Chaney et al, 2001) and undergoes major activator-dependent conformational changes during open complex formation (Casaz and Buck, 1999). Removal of, or mutations within, Region I confers an activator-bypass phenotype upon Eσ⁵⁴ (Wang and Gralla, 1996; Casaz et al, 1999). We used σ⁵⁴ lacking Region I (σ⁵⁴_ΔRI) or containing a triple alanine substitution at Region I positions R24, L26 and L26 (σ⁵⁴_ala24–26) to reconstitute Eσ⁵⁴ and investigate whether the removal of or mutations within Region I affected the β′ jaw domain–gp2 interaction. Native-PAGE analysis revealed that little or no stable complex between ³²P-labeled gp2 and Eσ⁵⁴_ΔRI or Eσ⁵⁴_ala24–26 was formed (Figure 6A). Since the free core RNAP, E:gp2, Eσ⁵⁴ and Eσ⁵⁴:gp2 complexes can be distinguished on native-PAGE gels, we silver-stained the native gel shown in Figure 6A to confirm that gp2 did not affect binding of σ⁵⁴_ΔRI or σ⁵⁴_ala24–26 to the RNAP (data not shown).

Figure 6.

T7 gp2 does not bind Eσ⁵⁴ reconstituted with σ⁵⁴-containing lesions in Region I. (A) An autoradiograph of a 4.5% (w/v) native gel showing the binding of ³²P-labeled gp2 to E (lane 2), Eσ⁵⁴ (lane 3), Eσ⁵⁴_ΔRI (lane 5) and Eσ⁵⁴_ala24–26 (lane 7). The migration positions of gp2 (lane 1), E:gp2 (lane 2) and Eσ⁵⁴:gp2 (lane 3) are indicated. Protein components in each lane are indicated on the top of the figure. (B) An autoradiograph of a 4.5% (w/v) native gel showing that ³²P-labeled gp2 can only detectably interact with Eσ⁵⁴_ΔRI in the presence of σ⁵⁴ Region I. Lanes 5 and 6 contain 2 and 4 μM σ⁵⁴ Region I, respectively, that was added in trans to the reaction prior to the addition of ³²P-labeled gp2. The migration positions of gp2, E:gp2, Eσ⁵⁴:gp2 and Eσ⁵⁴_ΔRI:RI:gp2 are as indicated. The protein components in each lane are indicated on top of the gel.

We considered the possibility that the addition of a peptide corresponding to σ⁵⁴ Region I residues 1–56 in trans may restore the binding of gp2 to Eσ⁵⁴_ΔRI. This expectation was fulfilled as ³²P-labeled gp2 formed a stable complex with Eσ⁵⁴_ΔRI when σ⁵⁴ Region I peptide was combined with RNAP holoenzyme prior to the addition of ³²P-labeled gp2 (Figure 6B, lanes 5 and 6). By isolating the Eσ⁵⁴_ΔRI:gp2 complex (from Figure 6B, lane 6) and analyzing the protein components by SDS–PAGE, we confirmed that this complex contained core RNAP, σ⁵⁴ Region I, σ⁵⁴_ΔRI and ³²P-labeled gp2 (data not shown). In trans addition of σ⁵⁴ Region I could somehow change the conformation of σ⁵⁴_ΔRI in such a manner that the gp2-binding site on the β′ jaw domain is revealed, thus allowing gp2 to stably bind. Control reactions demonstrated that gp2 did not bind Region I peptide or that the presence of σ⁵⁴ Region I in trans did not alter the native gel migration properties of the E:gp2 complex (data not shown). Clearly, the results demonstrate that σ⁵⁴ Region I, which is located at the ‘upstream face' of the RNAP (Wigneshweraraj et al, 2000), affects the conformation of the gp2-binding site on the β′ subunit (located at the ‘downstream face' of the RNAP; Figure 1a) and, by extension, determines the function of the β′ jaw domain during open complex formation.

σ⁵⁴ Region I residues 24–26 and 42–47 have a role in determining β′ jaw domain conformation

To identify Region I determinants important for gp2 binding to Eσ⁵⁴, we analyzed, by native PAGE, the binding of ³²P-labeled gp2 to Eσ⁵⁴ reconstituted from σ⁵⁴ Region I mutants in which residues 6–50 have been systematically changed to alanines, three consecutive residues at a time (Casaz et al, 1999). Densitometry analysis of the gel presented in Figure 7A revealed that Eσ⁵⁴ reconstituted with σ⁵⁴_ala24–26, σ⁵⁴_ala27–29, σ⁵⁴_ala33–35, σ⁵⁴_ala36–38, σ⁵⁴_ala42–44 and σ⁵⁴_ala45–47 bound gp2 60–90%less efficiently than wild-type Eσ⁵⁴ (Figure 7B), suggesting that σ⁵⁴ residues 24–29, 33–38 and 42–47 are important for gp2 binding to Eσ⁵⁴.

Figure 7.

σ⁵⁴ Region I residues 24–26 and 42–47 are important for T7 gp2 binding to Eσ⁵⁴. (A) An autoradiograph of a 4.5% (w/v) native gel showing the ability of ³²P-labeled gp2 to bind Eσ⁵⁴ reconstituted with an array of σ⁵⁴ Region I mutants. The migration positions of gp2, E:gp2 and Eσ⁵⁴:gp2 are indicated. At the bottom of the gel, ‘+' indicates that the corresponding mutant Eσ⁵⁴ is active and ‘−' indicates that it is not active for activator-bypass transcription. The error range for the graphed values is within ±9%. (B) Graph showing the efficiency of ³²P-labeled gp2 binding to the Eσ⁵⁴ mutants relative to its binding to the wild-type Eσ⁵⁴ (see text). (C) An autoradiograph of a 4% (w/v) denaturing gel showing the effects of gp2 on activator-bypass transcription by Eσ⁵⁴ reconstituted with selected σ⁵⁴ Region I mutants (see text). The arrow indicates the 470 nucleotide transcript from the S. meliloti nifH promoter on pMKC28. The reaction schematic is shown on the top of the gel.

Since σ⁵⁴ Region I residues important for gp2 binding to Eσ⁵⁴ result in strong deregulation of Eσ⁵⁴ when mutated (Casaz et al, 1999), we conducted activator-bypass transcription assays to check whether activator-bypass transcription by Eσ⁵⁴'s reconstituted with σ⁵⁴_ala24–26, σ⁵⁴_ala27–29, σ⁵⁴_ala33–35, σ⁵⁴_ala36–38, σ⁵⁴_ala42–44 and σ⁵⁴_ala45–47 was resistant to inhibition by gp2. As shown in Figure 7C and, as expected, activator-bypass transcription by Eσ⁵⁴_ΔRI (lanes 1 and 2), Eσ⁵⁴_ala24–26 (lanes 5 and 6), Eσ⁵⁴_ala42–44 (lanes 13 and 14) and Eσ⁵⁴_ala45–47 (lanes 15 and 16) was not inhibited by gp2, consistent with the fact that these mutant Eσ⁵⁴'s bound gp2 with 10–30% efficiency compared to the wild-type Eσ⁵⁴ (Figure 7A and B). Surprisingly, activator-bypass transcription by Eσ⁵⁴_ala27–29, Eσ⁵⁴_ala33–35 and Eσ⁵⁴_ala36–38, which was deficient in gp2 binding in a native-PAGE assay (Figure 7A and B), was inhibited by gp2 (Figure 7C, lanes 8, 10 and 12). We explain this by suggesting that binding of gp2 to Eσ⁵⁴_ala27–29, Eσ⁵⁴_ala33–35 and Eσ⁵⁴_ala36–38 may not be sufficiently stable to survive native PAGE, but is stable enough to prevent activator-bypass transcription in solution. Indeed, reducing the incubation time with gp2 (from 5 min to 30 s) before the addition of promoter DNA to the transcription reaction resulted in a detectable recovery of activator-bypass transcription by Eσ⁵⁴_ala27–29, Eσ⁵⁴_ala33–35 and Eσ⁵⁴_ala36–38, while transcription in the control reaction containing Eσ⁵⁴_R336A was efficiently inhibited by gp2 under these conditions (data not shown).

Overall, the results support the view that σ⁵⁴ Region I, particularly Region I residues 24–26 and 42–47, are important for gp2 binding to Eσ⁵⁴ and may contribute to the β′ jaw domain function in the context of wild-type Eσ⁵⁴.

Discussion

Multisubunit RNAPs must exist in several different functional states for the process of transcription initiation to progress. Bacterial RNAP containing σ⁵⁴ relies on an activator- and ATP-hydrolysis-driven remodeling step for transcription initiation. We have used bacteriophage T7-encoded transcription inhibitor of σ⁷⁰-dependent transcription, the gp2 protein, as a tool to study the contribution of the β′ jaw domain—the gp2-binding site—to transcription initiation by Eσ⁵⁴. Our analysis has revealed that even though gp2 binds to the same β′ site in σ⁵⁴- and σ⁷⁰-containing holoenzymes, activator- and ATP hydrolysis-dependent Eσ⁵⁴ transcription is resistant to inhibition by gp2. We have shown that unlike Eσ⁷⁰ closed complexes, Eσ⁵⁴ closed complexes are not disrupted by gp2. Using an analogue of ATP in the transition state of hydrolysis, we demonstrated that gp2 is not able to stably interact with β′ jaw domain within an early intermediate promoter complex. Thus, we suggest that the basis for the resistance of activated Eσ⁵⁴-dependent transcription to gp2 is due to activator- and ATP hydrolysis-driven conformational changes that disrupt inhibitory interaction(s) between gp2 and the β′ jaw domain during normal open complex formation. Supporting this view, the use of activator mutants defective in efficient coupling of energy derived from ATP hydrolysis to conformational changes in Eσ⁵⁴ resulted in some inhibition of Eσ⁵⁴ transcription by gp2. Further, in assays using the wild-type activator, reducing the incubation time with the activator (from 5 min to 30 s) resulted in significant inhibition of Eσ⁵⁴ transcription in reactions containing the wild-type core RNAP, but not the gp2-resistant E1188K mutant core RNAP (our unpublished observations).

The requirement for σ⁵⁴ Region I for the binding of gp2 to Eσ⁵⁴ suggests a functional and/or structural link(s) between σ⁵⁴ Region I and the β′ jaw domain. The precise nature of this link is currently unclear, but must have a role in regulating the functionality of the RNAP. Region I constitutes the major activator-binding site (Chaney et al, 2001) and it undergoes activator- and ATP hydrolysis-dependent conformational changes during open complex formation (Casaz and Buck, 1999; Wigneshweraraj et al, 2001; Burrows et al, 2003; 2004). Thus, an attractive possibility is that ‘activating signals' from the ‘upstream face' of the RNAP (where Region I is located; Wigneshweraraj et al, 2000) are somehow relayed to the β′ jaw domain during open complex formation. Since the β′ jaw domain is modeled to contact the downstream DNA within the crystal structure of the Thermus aquaticus Eσ^A promoter complex (Murakami et al, 2002) and is suggested to make stabilizing interactions with downstream DNA during open promoter complex formation by Eσ⁷⁰ (Ederth et al, 2002), we suggest that the β′ jaw domain, upon activation, becomes engaged with downstream DNA during open complex formation by Eσ⁵⁴. Similarly, the homologue of the β′ jaw domain in eukaryotic RNAP II, the RPB1 jaw lobe, undergoes a conformational change and moves towards DNA upon binding of transcription factor TFIIB (Bushnell et al, 2004).

In agreement with the activator- and ATP hydrolysis-dependent use of the β′ jaw domain during Eσ⁵⁴ open complex formation, gp2 is able to inhibit activator-bypass transcription by some activator-bypass forms of Eσ⁵⁴ as efficiently as Eσ⁷⁰ transcription. Notably, activator-bypass mutant Eσ⁵⁴_R336A is gp2-resistant in activator-dependent assays, but not in activator-bypass assays. The striking difference in gp2 sensitivity between activator-dependent and activator-bypass transcription further underscores the activator-dependent engagement of the β′ jaw with DNA in a σ⁵⁴ Region I-dependent manner during Eσ⁵⁴ transcription. Since gp2 inhibited activator-bypass Eσ⁵⁴ transcription by preventing stable DNA melting, we suggest that the β′ jaw domain has a role in stable DNA melting during open complex formation by Eσ⁵⁴. Supporting this idea, Eσ⁵⁴ reconstituted using a mutant RNAP core form lacking the β′ jaw domain (β′Δ1149–1190) is defective for DNA melting and transcription (Wigneshweraraj et al, in preparation).

In bacterial RNAP, access to the cleft containing the catalytic center is restricted by the β′ clamp domain (Murakami et al, 2002). This conformation is referred to as the ‘closed clamp state'. Promoter DNA must enter the cleft, either as melted DNA or as DNA ‘to be melted' during or after the ‘clamp-opening' event. Several groups (Murakami et al, 2002; Armache et al, 2003; Bushnell and Kornberg, 2003) proposed that initiation of DNA melting ‘outside' the cleft may allosterically induce conformational changes that result in ‘clamp opening' and propagation of melting. The β′ clamp and jaw domains are integral parts of the β′ subunit and thus are likely to undergo interdependent conformational changes during ‘clamp opening'. Supporting this view, recent structural information on yeast RNAPII revealed that the RPB1 clamp and jaw-lobe domains undergo inter-dependent conformational changes upon interaction with transcription factor TFIIB (Bushnell et al, 2004). Based on different gp2 sensitivities of Eσ⁷⁰ and Eσ⁵⁴ transcription, below, we discuss the role of the β′ jaw domain in ‘clamp opening' and transcription by bacterial RNAP below.

σ⁵⁴ can initiate DNA melting in response to activation in the absence of RNAP core (Cannon et al, 2001). Our results suggest that activation somehow results in conformational changes in the β′ jaw domain and allows the β′ jaw domain to engage downstream DNA, which leads to stable melting. Based on the two-step model proposed by Wang and Gralla (1996) for open complex formation by Eσ⁵⁴, we suggest that the first activation step results in a core RNAP-independent initiation of DNA melting, which occurs in the ‘closed clamp' conformation outside the catalytic cleft (Cannon et al, 2001). During the second activation step, conformational changes in the β′ jaw domain trigger ‘clamp opening' and so facilitate the entry of the partly melted DNA into the RNAP catalytic cleft to allow propagation of melting in an RNAP core-dependent manner. Within the RNAP catalytic cleft, the RNAP catalytic site proximal downstream DNA contacts are made by the β′ clamp domain, whereas the RNAP catalytic site distal downstream DNA contacts are made by the β′ jaw domain (Murakami et al, 2002). We envisage that the gp2 dissociates from the β′ jaw domain upon activator- and ATP hydrolysis-dependent loading of DNA onto the β′ jaw domain. Consistent with the view that activator ‘drives' the DNA onto the β′ jaw domain, the interaction between core RNAP and gp2 is disrupted by heparin or nonspecific DNA (our unpublished observations). Similarly, the interaction between gp2 and Eσ⁵⁴ is highly heparin-sensitive (see Supplementary data Figure 3). In the case of activator-bypass transcription, the DNA enters the RNAP catalytic cleft and contacts the β′ jaw domain independently of the activation event. Since activator-bypass open complexes are significantly more heparin sensitive than activator-dependent open complexes, we argue that during activator-bypass transcription the DNA-β′ jaw domain contacts are weak and cannot result in dissociation of gp2.

In contrast, σ⁷⁰ strictly relies on the use of core RNAP subunits for DNA melting (Young et al, 2004). Hence, core RNAP-dependent DNA melting may allosterically induce ‘clamp opening' and subsequently facilitate further DNA melting within the catalytic cleft. Therefore, as proposed by Ederth et al (2002), the presence of gp2 could prevent β′ jaw–DNA contacts during promoter DNA melting and thereby destabilize intermediate open promoter complexes. However, once complete promoter DNA melting and ‘clamp opening' has occurred, gp2 is not able to bind open promoter complexes and therefore has no effect on Eσ⁷⁰ open promoter complexes (Nechaev and Severinov, 1999)—presumably because the gp2-binding site on the β′ jaw domain is bound by DNA. Clearly, accessory factors such as σ factors which bind the upstream face of the RNAP are able to control the activity of other parts of the RNAP (Figure 1A), allowing control of RNAP functionality (Chung et al, 2003; Bushnell et al, 2004 and this work).

Materials and methods

Protein purification

All wild-type and mutant Klebsiella pneumoniae σ⁵⁴ and E. coli PspF_1–275 were purified as described in Wigneshweraraj et al (2003). T7 gp2 was purified from E. coli 7009(DE3) cells essentially as detailed in Nechaev and Severinov (1999). The protocol for the overexpression and purification of wild-type E. coli σ⁷⁰ is described in Nechaev and Severinov (1999). Wild-type E. coli core RNAP was purchased from Epicentre Technology (Madison, WI). T7 gp2-resistant E. coli core RNAP containing the E1188K mutation was purified from E. coli 7009 cells as detailed in Nechaev and Severinov (1999).

In vitro transcription assays

Single-round activator-dependent and activator-bypass transcription assays from supercoiled (pMKC28) and linear S. meliloti nifH templates were conducted as detailed in Wigneshweraraj et al (2003). The protocol for Eσ⁷⁰-dependent transcription from pOMlacUV5 is described in Colland et al (1999). Where indicated, gp2 was present in four-fold molar excess over Eσ (unless otherwise stated). The final concentrations of template DNA, Eσ (reconstituted using 1:4 molar ratio of core RNAP over σ), PspF_1–275 (where indicated) and ATP (where indicated) were at 20, 100 nM, 4 μM and 4 mM, respectively. The elongation mix (abbreviated as ‘El.mix' in the figures) contained 1 mM ATP, CTP, 0.05 mM UTP, 3 μCi of [α-³²P]UTP and 100 μg/ml heparin. The point at which gp2 was added to the reactions is specified in the figure and legends thereof.

T7 gp2-Eσ⁵⁴-binding assays

All binding reactions (10 μl) were conducted in STA (25 mM Tris-acetate, pH 8.0, 8 mM Mg-acetate, 10 mM KCl, 3.5% w/v PEG 6000) buffer at 37°C. Eσ⁵⁴ was present at 100 nM (reconstituted as above) and ³²P-gp2 at 800 nM. Where indicated, the streptavidin–biotin-labeled S. meliloti nifH promoter probe (positions −35 to +1) was present at 400 nM. Biotin-tagged (at the 5′ end of the nontemplate strand) S. meliloti nifH promoter probe was labeled with streptavidin (Sigma) by incubating with four-fold streptavidin for 5 min at 37°C in STA buffer. Where indicated, σ⁵⁴ Region I residues 1–56 were present at the indicated concentrations (see figure legends). Assays were started by reconstituting the Eσ⁵⁴ for 5 min, followed by adding of gp2 and incubating for a further 5 min. Where indicated, a streptavidin-labeled promoter probe was added for 5 min. The reactions were stopped and analyzed for native PAGE as in Wigneshweraraj et al (2003).

Activator interaction assays

The Eσ⁵⁴:gp2 or Eσ⁵⁴:DNA:gp2 complexes were formed as described above. The ADP-AlF_x-dependent activator-binding assays were conducted as described in Chaney et al (2001) using final concentrations of 5 μM PspF_1–275, 0.2 mM ADP, 0.2 mM AlCl₂ and 5 mM NaF to form the PspF_1–275:ADP-AlF_x complex.

DNA melting and footprinting assays

Promoter complexes were formed on pMKC28 in the absence and presence of gp2 at concentrations as outlined above for the transcription assays. KMnO₄ and/or DNase I footprinting reactions were conducted as detailed in Cannon et al (2001) and the reactions were processed as detailed in Burrows et al (2003).

Supplementary Material

Supplementary data Figure 1

Supplementary data Figure 2

Supplementary data Figure 3