Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Schaubach, O. L.
Right arrow Articles by Dombroski, A. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Schaubach, O. L.
Right arrow Articles by Dombroski, A. J.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

J Biol Chem, Vol. 274, Issue 13, 8757-8763, March 26, 1999


Transcription Initiation at the Flagellin Promoter by RNA Polymerase Carrying sigma 28 from Salmonella typhimurium*

Olivia Lee Schaubach and Alicia J. DombroskiDagger

From the Department of Microbiology and Molecular Genetics, University of Texas Health Science Center, Houston, Texas 77030

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The sigma  subunit of RNA polymerase is a critical factor in positive control of transcription initiation. Primary sigma  factors are essential proteins required for vegetative growth, whereas alternative sigma  factors mediate transcription in response to various stimuli. Late gene expression during flagellum biosynthesis in Salmonella typhimurium is dependent upon an alternative sigma  factor, sigma 28, the product of the fliA gene. We have characterized the intermediate complexes formed by sigma 28 holoenzyme on the pathway to open complex formation. Interactions with the promoter for the flagellin gene fliC were analyzed using DNase I and KMnO4 footprinting over a range of temperatures. We propose a model in which closed complexes are established in the upstream region of the promoter, including the -35 element, but with little significant contact in the -10 element or downstream regions of the promoter. An isomerization event extends the DNA contacts into the -10 element and the start site, with loss of the most distal upstream contacts accompanied by DNA melting to form open complexes. Melting occurs efficiently even at 16 °C. Once open complexes have formed, they are unstable to heparin challenge even in the presence of nucleoside triphosphates, which have been observed to stabilize open complexes at rRNA promoters.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Prokaryotic RNA polymerase (RNAP)1 is a multimeric enzyme consisting of the core subunits, alpha 2beta beta ' (E), and a sigma  factor (1). The sigma  factor is responsible for directing the recognition and binding of specific promoter DNA consensus sequences by holoenzyme (alpha 2beta beta 'sigma ) and for facilitating transcription initiation (2, 3). Many different sigma  factors have been identified, and they fall into two major categories, those similar to Escherichia coli sigma 70 and those similar to sigma 54 (4). The sigma 70 family can be further subdivided into primary and alternative sigma  factors (5, 6). Primary sigma  factors are essential and required for the expression of housekeeping genes in vegetative cells, whereas alternative sigma  factors are activated in response to environmental changes and in programming developmental pathways (4, 6, 7). Promoters recognized by the sigma 70 family consist of conserved hexamers at -35 and -10 with respect to the start site of transcription (8). Additional DNA contacts are made at some promoters by the C-terminal domain of the alpha  subunit of RNAP at an AT-rich region between -38 and -59 called the UP element (9).

Comparison of the amino acid sequences of members of the sigma 70 family reveals four highly conserved regions with subdomains that have been implicated in specific functions (Fig. 1) (4). Region 1.1 inhibits sigma 70 from binding to the DNA in the absence of the core subunits (10) and is also required for efficient progression from the earliest RNAP·DNA complex to a transcriptionally active complex during initiation (11). Deleting regions 1.1 and 1.2 of sigma 70 results in transcriptional arrest after initial binding of RNAP to the promoter (11). Regions 2.1, 2.2, and 3.2 are important for core binding (12-14), and region 2.3 has been implicated in promoter melting (15-18). Both holoenzyme and the sigma  factor interact with non-template bases in the -10 element to stabilize the open complex (19-21). Regions 2.4 and 4.2 are responsible for contacting the -10 and -35 promoter recognition elements, respectively, and for positioning holoenzyme for initiation (10, 22-26). Regions 1.2, 2, and 4 are found in almost all sigma  factors, but region 1.1 is found only in the primary sigma  factors (4). Interestingly, both regions 1.1 and 1.2 are absent in the Salmonella typhimurium alternative sigma  factor required for flagellum biosynthesis, sigma 28, hinting at potential variation in the structure of Esigma 28·DNA complexes or in the mechanism of transcription initiation.


View larger version (6K):
[in this window]
[in a new window]
  		Fig. 1.   Linear schematic diagram of sigma 70 and sigma 28. Primary sigma  factors, such as sigma 70, possess four highly conserved regions of amino acids. Each shaded section represents further delineation into subregions. Region 1.1 (black) is encountered specifically on primary sigma  factors. It controls DNA binding by free sigma  and is required for efficient isomerization to the open complex during initiation. Region 2.1 (gray) is involved in core binding. Region 2.4 (white) interacts with the -10 promoter consensus sequence, and region 4.2 (vertical stripes) interacts with the -35 promoter consensus sequence. Regions 1.2, 2, and 4 are found in almost all sigma  factors; however, for S. typhimurium sigma 28, region 1.2 is absent.

Initiation can be described as a series of sequential steps, which have been well characterized for both Esigma 70 and Esigma 32 from E. coli (27). By varying the conditions, several intermediates can be visualized using footprinting methods. RNAP (R) binds to the promoter (P) to form an initial closed complex, RPC1, which protects the DNA from approximately -60 to -5 relative to the start point of transcription. RPC1 isomerizes to a second closed complex (RPC2) that maintains the upstream contacts and extends further downstream of the transcription start site to +20. RPC2 then undergoes strand opening to form an open complex, RPO, whereas the length of the footprint is unchanged. There is evidence for more than one open complex, which is dependent on the presence or absence of Mg2+ (28, 29). RPO enters the initiation stage, or RPinit, in the presence of initiating nucleotides. Short transcripts of 2-12 nucleotides in length (abortive products) are synthesized while RNAP remains at the promoter. RNAP then enters into the elongation phase of transcription upon promoter clearance and sigma  factor release. Several of these intermediate complexes have been visualized by performing DNase I and KMnO4 footprinting experiments over a range of temperatures, with the rationale that temperature-dependent complexes represent time-dependent events (30-32).

The S. typhimurium flagellar operon can be divided into three classes of genes (I-III) based on their transcriptional hierarchy in flagellar assembly (33). sigma 28, encoded by the fliA gene, is expressed late in flagellum biosynthesis and is required for transcription of all class III genes including fliC, encoding flagellin, the primary component of the flagellar filament (34), and flgM, encoding the sigma 28 anti-sigma factor (35, 36). In this report, we have characterized transcription initiation by RNA polymerase carrying the S. typhimurium sigma  factor, sigma 28, on variants of the flagellin (fliC) promoter. We detected intermediate complexes formed by Esigma 28 during initiation that are distinct from those characterized for other holoenzymes. Initial binding to the promoter does not require the -10 element, but further extension of the RNA polymerase-DNA contacts and isomerization to the open complex demand the presence of the -10 sequence. We propose an alternative mode of promoter recognition and binding as compared with other well characterized holoenzymes.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Overproduction and Purification of sigma 28 and Reconstitution of Holoenzyme-- The fliA gene, encoding sigma 28, was inserted into plasmid pET15b (Novagen, Inc.) to generate pKH439 (a gift from K. Hughes), which resulted in the addition of six histidines at the amino terminus. Hexahistidine-tagged sigma 28 was overproduced and purified using the method described by Wilson and Dombroski (11). Holoenzyme was reconstituted by adding 1.0 pmol of E. coli core RNA polymerase (E) (Epicentre Technologies Corp.) to 5.0 pmol of sigma 28 in protein dilution buffer (10 mM Tris-HCl (pH 8.0), 10 mM KCl, 10 mM beta -mercaptoethanol, 1 mM EDTA, 0.4 mg/ml bovine serum albumin, and 0.1% Triton X-100) and incubating on ice for 15 min. Because of the amino acid sequence conservation among the core subunits, heterologous holoenzymes have been used to assess the behavior of alternative sigma  factors (37-40). Additionally, we compared the amino acid sequences of the alpha  and beta  subunits of E. coli and S. typhimurium RNA polymerases (11) (data not shown). There is 100% identity between the alpha  subunits and 98% identity between the beta  subunits.

Constructions and Generation of Promoter Fragments-- pfliCDelta 35 was constructed by annealing two oligonucleotides (Integrated DNA Technologies) of 122 bases in length (41). The DNA was designed to incorporate HindIII and BamHI restriction sites near the 5'- and 3'-ends, respectively. The double-stranded DNA was digested with HindIII and BamHI, ligated into the same sites in pBluescriptII KS+ (Stratagene), and transformed into E. coli strain DH5alpha (Life Technologies, Inc.). The clones were sequenced to confirm the deletion using the fmolTM DNA sequencing system (Promega).

pfliCDelta 10 was constructed using an oligonucleotide with a deletion of the -10 element as a polymerase chain reaction primer with plasmid pMC72, containing the wild-type fliC promoter, as the template. The resulting DNA fragment was ligated into pCR2.1 and transformed into Invalpha F' One Shot competent cells from the original TA cloning kit (Invitrogen). The clones were sequenced to confirm the deletion using the fmolTM DNA sequencing system.

32P-5'-End-labeled primers were generated for use in synthesizing labeled fliC promoter DNA (10). Oligonucleotide primers were from BioServe Biotechnologies, Genosys Biotechnologies, Inc., or Integrated DNA Technologies.2 Plasmid pMC72 (a gift from K. Hughes) and the two plasmids containing pfliCDelta 10 and pfliCDelta 35, as described above, were used as the template DNA to generate the fliC promoter DNA and derivatives. Both radiolabeled and unlabeled fliC promoters were synthesized using the polymerase chain reaction to generate a 230-base pair fragment. Each polymerase chain reaction contained 10× Taq Buffer A (Fisher), 50 pmol of each primer, 40 ng of template DNA, and 2.5 units of Taq polymerase (Fisher) in a final volume of 100 µl. A Perkin-Elmer Thermocycler was set for 35 cycles with 95 °C for denaturation, 50 °C for annealing, and 72 °C for extension. The products were purified using the Qiaquick polymerase chain reaction DNA purification kit (QIAGEN Inc.).

DNase I Footprinting-- 32P-End-labeled DNA promoter fragment and DNase I buffer (20 mM sodium Hepes (pH 7.5), 10 mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 1 mM dithiothreitol, and 200 µg/ml bovine serum albumin) were combined in a volume of 50 µl. Holoenzyme was added in 10-fold excess over the DNA unless noted otherwise. Additional processing of these samples has previously been described (40). Several variations of the DNase I footprinting are described below.

Heparin Competition-- The Esigma 28-promoter complexes were allowed to form at 37 °C for 15 min. In some cases, nucleoside triphosphates (NTPs) were added to a final concentration of 0.2 mM for 1 min, and then heparin was added (25 µg/ml final concentration), followed by incubation for another minute. The complexes were treated with 0.5 units of DNase I (Promega) for 30 s and processed as already described.

Temperature Variation-- The Esigma 28-promoter complexes were allowed to form at 0, 4, 16, 25, and 37 °C for 15 min. The complexes formed at 0 and 4 °C were subjected to DNase I digestion (2 units of DNase I) for 35 and 30 min, respectively. Complexes formed at 16 °C were digested for 4 min (1.5 units of DNase I), and the 25 and 37 °C samples were digested for 1 min (1.0 unit of DNase I).

KMnO4 Footprinting-- Esigma 28 and 0.1 pmol of end-labeled DNA promoter fragment were combined in KMnO4 buffer (20 mM sodium Hepes (pH 7.5), 10 mM MgCl2, 100 mM NaCl, 0.1 mM EDTA, 0.2 mM dithiothreitol, and 200 µg/ml bovine serum albumin) to a final volume of 50 µl and then treated with 2.5 µl of 50 mM KMnO4 (Sigma) for 2 min at the temperatures indicated. Additional processing has been described (42).

Nucleotide Stabilization Assay in Vitro-- The nucleotide stabilization assay was performed as outlined by Wilson and Dombroski (11) with the following modifications. The Esigma 28-promoter complexes were allowed to form for 15 min, and then 0.1 mM ATP, CTP, and GTP were added to the preformed complexes for 30 s, prior to filtration and washing with 0.8 M NaCl.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Promoter Binding and Transcription in Vitro-- RNAP was formed by mixing hexahistidine-tagged sigma 28 with purified E. coli core RNAP. Because of the high degree of similarity between the core subunits of E. coli and S. typhimurium RNAPs, a heterologous system was used as reported by others to assess the behavior of alternative sigma  factors (37-40). Runoff transcription assays were performed to analyze the overall efficacy of transcription initiation by Esigma 28. The promoter chosen for this analysis, pfliC, drives the expression of flagellin, one of the late flagellar gene products. A transcript of the expected size was observed. No difference in behavior was noted in the presence or absence of the hexahistidine tag (data not shown). Deletion of the AT-rich sequence just upstream of the -35 element had a slight effect on transcription (2-fold reduction), but was not attributable to the presence of an UP element since holoenzyme containing a truncation of the carboxyl-terminal domain of the alpha  subunit resulted in the same transcriptional behavior as the wild-type enzyme, independent of the upstream DNA sequence (data not shown).

DNase I footprinting was used to characterize the interaction of Esigma 28 with pfliC (Fig. 2). A DNA fragment of 230 base pairs in length containing the fliC promoter was 5'-end-labeled on the template strand with 32P. Increasing amounts of reconstituted Esigma 28 were added, followed by DNase I digestion (Fig. 3). Continuous protection was observed from -24 to +17, with partial protection from -46 to -24 relative to the transcription start site. Additional weak interactions between +17 and +20 could be discerned. Overall, this footprint is shorter in the upstream region than those typical for Esigma 70 or Esigma 32 under similar conditions.


View larger version (12K):
[in this window]
[in a new window]
  		Fig. 2.   Variants of the fliC promoter used in this study. The promoters used in this study, wild-type pfliC, pfliCDelta 35, and pfliCDelta 10, are shown. The -10 and -35 regions for sigma 28 are indicated in boldface type with underlining. The positions where the -10 and -35 elements were deleted are indicated by asterisks.


View larger version (29K):
[in this window]
[in a new window]
  		Fig. 3.   Binding of Esigma 28 to pfliC using DNase I footprinting. 5'-Radiolabeled pfliC (template strand) was incubated with increasing amounts of Esigma 28 for 15 min at 37 °C and subsequently treated with DNase I. All samples were analyzed by electrophoresis on a denaturing 8% polyacrylamide gel. The extent of each footprint is indicated on the side of each footprinting ladder. Lane 1, DNA alone; lane 2, 0.01 pmol of Esigma 28; lane 3, 0.02 pmol of Esigma 28; lane 4, 0.04 pmol of Esigma 28; lane 5, 0.08 pmol of Esigma 28; lane 6, 0.1 pmol of Esigma 28; lane 7, 0.2 pmol of Esigma 28.

Intermediate Complexes Formed during Transcription Initiation-- We characterized the intermediate Esigma 28·DNA complexes during the process of transcription initiation that can be visualized by manipulating the temperature of incubation. Esigma 28 was incubated with 5'-end-labeled pfliC (template strand) for 15 min, at 0, 4, 16, 25, and 37 °C. Protection was observed from -65 to -54 and from -48 to -19 at 0 and 4 °C (Fig. 4A). At 16 °C, a hypersensitive band appeared at -46, and partial protection began to extend downstream toward the -10 region and the start site (-46 to +17), with the reappearance of bands at -33 and -24. By 25 °C, Esigma 28 fully occupied the region from -46 to +17. At 37 °C, the upstream contacts from -65 to -54 disappeared, but strong protection from -46 to +17 remained. Thus, Esigma 28 appeared to initially bind primarily to the upstream region of the promoter since -10 protection was absent at lower temperatures and then shifted downstream to contact the -10 element up to the +17 region as the temperature was increased. We cannot rule out the possibility that some minor contacts in the upstream region are maintained. However, it is clear that a major rearrangement takes place as a function of temperature. The same pattern of contacts was observed with footprinting performed using DNA labeled on the non-template strand (data not shown).


View larger version (47K):
[in this window]
[in a new window]
  		Fig. 4.   DNase I and KMnO4 footprinting of Esigma 28-promoter complexes as a function of temperature. 5'-Radiolabeled pfliC (template strand) was incubated with Esigma 28 at the temperatures indicated below for 15 min, and the complexes were subsequently treated with DNase I or KMnO4 and piperidine. DNA Alone shown here was obtained at 37 °C; however, the same pattern of digestion was obtained at 0 °C (data not shown). All samples were analyzed by electrophoresis on a denaturing 8% polyacrylamide gel. A, DNase I protection. Arrows on the side of each footprint indicate the extent of protection. First lane, DNA alone; second lane, 0 °C; third lane, 4 °C; fourth lane, 16 °C; fifth lane, 25 °C; sixth lane, 37 °C. B, KMnO4 reactivity. Arrows indicate the reactive bases in the open complex. First lane, DNA alone; second lane, 0 °C; third lane, 4 °C; fourth lane, 16 °C; fifth lane, 25 °C; sixth lane, 37 °C.

Open Complex Formation-- Extended DNase I footprints at 37 °C are usually indicative of open complexes for Esigma 70 (27, 43-46). We used KMnO4 sensitivity to determine which Esigma 28-promoter complexes in the DNase I temperature series were open complexes. KMnO4 chemically modifies thymine residues in single-stranded DNA, which renders modified positions sensitive to cleavage upon piperidine treatment (47). Esigma 28-promoter complexes were formed for 15 min at 0, 4, 16, 25, or 37 °C and subjected to KMnO4 treatment and piperidine cleavage (Fig. 4B).

From 0 to 4 °C, the DNA remained base-paired as demonstrated by lack of reactivity to KMnO4. Open complexes were fully established at 16, 25, and 37 °C with reactivity to KMnO4 at +1, -6, -7, and -9. Thus, similar to RPO for Esigma 70 and Esigma 32, the Esigma 28 DNase I footprints that extend into the -10 region and start site represent open complexes. Surprisingly, however, Esigma 28 formed open complexes at temperatures as low as 16 °C, whereas Esigma 70 and Esigma 32 form primarily extended closed complexes (RPC2) under these conditions (27, 30-32, 46, 48). We did not observe an RPC2-type complex for Esigma 28 under any conditions.

Promoter Variants Lacking the -10 or -35 Element-- If Esigma 28 initially binds to the -35 region, releases upstream interactions, and shifts its contacts to include the -10 and downstream regions, then removal of the -35 element should preclude any binding by Esigma 28. Likewise, removal of the -10 consensus sequence should permit binding of only Esigma 28 in the -35 region. We constructed deletions of the TAAA sequence at -35 (pfliCDelta 35) and the GCCGATA sequence at -10 (pfliCDelta 10) (Fig. 2) and analyzed the DNase I footprints over a range of temperatures. Deletion of the -10 element still allowed normal binding of Esigma 28 at 0 and 4 °C, as expected based on the presence of the predicted initial binding site at -35 (Fig. 5A). However, upon raising the temperature, binding to pfliCDelta 10 was gradually abolished, consistent with the idea that polymerase shifts its contacts downstream during open complex formation to interact with the -10 element and downstream sequences. In the case of pfliCDelta 10, these sequences have been removed, and Esigma 28 is unable to remain stably bound to the upstream sequences and results in dissociation.


View larger version (78K):
[in this window]
[in a new window]
  		Fig. 5.   DNase I footprinting of Esigma 28 on pfliC variants. 5'-Radiolabeled pfliCDelta 10 (template strand) or pfliCDelta 35 (non-template strand) was incubated with Esigma 28 at the temperatures indicated below for 15 min and subjected to DNase I treatment. All samples were analyzed by electrophoresis on a denaturing 8% polyacrylamide gel. Arrows denote the locations of the protected regions. A, pfliCDelta 10. The position of the deletion of the -10 element is indicated by an asterisk. First lane, DNA alone; second lane, core RNAP; third lane, 0 °C; fourth lane, 4 °C; fifth lane, 16 °C; sixth lane, 25 °C; seventh lane, 37 °C. B, pfliCDelta 35. The position of the deletion of the -35 element is indicated by an asterisk. First lane, DNA alone; second lane, core RNAP; third lane, 0 °C; fourth lane, 4 °C; fifth lane, 16 °C; sixth lane, 25 °C; seventh lane, 37 °C.

Also as expected, Esigma 28 was unable to bind to pfliCDelta 35 even at 0 °C, presumably due to lack of the initial binding site in the -35 region and upstream portions of the promoter (Fig. 5B). KMnO4 footprinting showed no open complexes at either pfliCDelta 10 or pfliCDelta 35 at any temperature (data not shown). Taken together, these results support the idea that initial binding occurs from -65 to -19, followed by isomerization to relinquish major contacts between -65 and -54 and establishment of new contacts extending to +20 with concomitant DNA melting.

Stability of Esigma 28-Promoter Complexes-- Previous studies have shown that Esigma 70 typically forms a heparin-stable open complex (48-51). One exception is the ribosomal operon promoter, rrnBP1, which requires the addition of NTPs to confer heparin stability (52). We used heparin challenge as a tool to examine the stability of Esigma 28·DNA complexes. The binding of Esigma 28 to pfliC was sensitive to 25 µg/ml heparin, even in the presence of initiating NTPs (Fig. 6). Thus, Esigma 28 open complexes (RPO and RPinit) appear to be generally less stable than Esigma 70 open complexes. RNAP with the primary sigma  factor from Bacillus subtilis, Esigma A, also forms heparin-sensitive complexes in the presence or absence of NTPs (53).


View larger version (28K):
[in this window]
[in a new window]
  		Fig. 6.   Determination of Esigma 28-promoter complex stability. 5'-Radiolabeled pfliC (template strand) was incubated with Esigma 28 for 15 min at 37 °C, and the complexes were subsequently treated with DNase I. All samples were analyzed by electrophoresis on a denaturing 8% polyacrylamide gel. Arrows show the locations of the protected regions. First lane, DNA alone; second lane, Esigma 28; third lane, Esigma 28 + 25 µg/ml heparin (Hep); fourth lane, Esigma 28 + 2 mM ATP, CTP, and GTP (ACG); fifth lane, Esigma 28 + 2 mM ATP, CTP, and GTP + heparin.

RPinit complexes for Esigma 70 can be distinguished from RPO complexes by resistance to challenge with high salt in the presence of NTPs (11, 50, 54). We used an NTP stabilization assay to determine whether RPinit complexes formed by Esigma 28 on pfliC were stable to salt challenge. Esigma 28·fliC complexes were formed in the absence and presence of NTPs and were then filtered through nitrocellulose and subjected to a 0.8 M NaCl wash. In the presence of NTPs, 52% of the complexes were retained on the filter, as expected since RPinit is in equilibrium with RPO, whereas in the absence of NTPs, only 5% were retained. Therefore, Esigma 28 RPinit complexes on pfliC, like Esigma 70·DNA complexes, are stable to 0.8 M NaCl, indicating a equivalent conformational change and stabilization upon NTP binding.

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Transcription initiation has been characterized as a multistep process in studies in which intermediates on the pathway to formation of an initiated complex have been detected by manipulating the temperature of incubation and varying the level of Mg2+ for the RNAP-promoter interactions (30, 31, 43, 46, 50, 51). There is evidence for two different closed complexes (RPC1 and RPC2), which differ in the extent of their contacts with the promoter DNA. RPC1, which extends from approximately -60 to -5, is only observed at low temperatures (0 °C) for Esigma 70 (30) and Esigma 32 (31, 32). However, we have recently shown that derivatives of sigma 70 with region 1.1 or both regions 1.1 and 1.2 removed slow the initiation process such that RPC1 can be detected at 37 °C (11).

We have investigated the mechanism of transcription initiation by holoenzyme carrying sigma 28, the alternative sigma  factor required for flagellum biosynthesis in S. typhimurium. Our analysis was conducted using the promoter for fliC, encoding flagellin, a late gene in this pathway. The identity and stability of transcription complex intermediates that we identified during transcription initiation by Esigma 28 are distinctive from those previously identified for Esigma 70 and Esigma 32.

Based on our results, we suggest the following pathway for the mechanism of Esigma 28 transcription initiation (Fig. 7). Esigma 28 forms a short complex with pfliC at 0 °C that protects primarily the upstream region of the promoter (-65 to -19) from DNase I digestion. This suggests that the initial binding of Esigma 28 is mediated mainly through interactions in the vicinity of the -35 consensus sequence. Esigma 28 then isomerizes to make additional contacts that extend to +20. During this progression, some upstream contacts are lost as evidenced by the reappearance of bands from -65 to -54 on the DNase I footprints at 25 and 37 °C. Thus, Esigma 28 appears to initially bind in the -35 region and then shifts downstream to contact the -10 region, start site, and downstream sequences to +20 while relinquishing some of the original upstream contacts. This represents a notable difference in binding intermediates from those observed for Esigma 70 or Esigma 32.


View larger version (15K):
[in this window]
[in a new window]
  		Fig. 7.   Schematic of the mechanism for transcription initiation by Esigma 28. RNA polymerase (R) binds to the promoter DNA (P) to form a closed complex (RPC), which extends from -65 to -19. Extension of the downstream contacts to +17 and release of upstream contacts occur concomitantly with open complex formation (RPO). Regions of full protection are indicated by black bars. Partial protection is indicated by dashed lines. The thymidine residues sensitive to KMnO4 modification in the open complex are indicated (triangle ).

The identity of RPC versus RPO complexes was assessed by KMnO4 sensitivity, which probes for DNA strand separation. Typically, significant open complex formation for Esigma 70 or Esigma 32 requires temperatures above 16 °C (27, 30, 31). One distinctive characteristic of Esigma 28·fliC complexes is that they are optimally strand-separated between -9 and +1 even at 16 °C. Thus, the extended DNase I protection observed at 16 °C is representative of RPO, rather than an extended closed complex like RPC2. Thus, one major difference between Esigma 28 intermediates and those found for Esigma 70 or Esigma 32 from E. coli is the absence of a detectable RPC2-like complex under the conditions that we used. Esigma 28 appears to form a single closed complex characterized by a short DNase I footprint at 0-4 °C and then progresses to RPO. If an RPC2 complex forms, it may be too rapid or unstable to be detected with the methods employed in this study.

Because Esigma 28 is functionally similar to the B. subtilis holoenzyme that is required for flagellin biosynthesis, Esigma D, we compared the transcription initiation complexes formed by these two enzymes at their respective flagellin promoters. The flagellin gene promoter (phag) from B. subtilis is very similar to pfliC, except it contains an UP element between -60 and -40 that provides additional contact sites for RNAP through the alpha  subunit (57). Esigma D forms complexes that are more similar to Esigma 70 and Esigma 32 because, at 4 °C, the DNase I footprint occupies from -73 to +1 (55). This protection then extends to +9 and +21 at 23 and 37 °C, respectively. Disappearance of the upstream contacts, which is observed for Esigma 28, does not occur. Thus, DNA binding primarily in the -35 and upstream regions in the earliest detectable complex, with lack of contact in the -10 region, appears to be novel for Esigma 28.

With respect to open complex formation, however, Esigma 28 and Esigma D share the ability to generate strand-separated promoter regions at low temperatures. At 16 °C, Esigma 28 displays the same degree of open complex formation that is observed at 37 °C, but forms only closed complexes at 4 °C. Esigma D begins to show DNA distortions in the -10 region at temperatures as low as 4 °C; however, the region of strand melting propagates unidirectionally as a function of temperature, forming three distinct open complex intermediates (55), whereas the reactivity of bases in the Esigma 28 open complex remains the same from 16 to 37 °C, with no indication of directional movement.

We used challenge with the polyanionic competitor heparin as a tool to assess the relative stability of open (RPO) and initiated (RPinit) complexes. Esigma 28·fliC open complexes are unstable to even low levels of heparin in the presence or absence of initiating NTPs. In contrast, many Esigma 70 open complexes are stable to heparin even in the absence of NTPs (11, 48-51). Esigma 28 appears to form transcriptionally competent complexes that remain dissociable throughout the initiation process. We cannot rule out that Esigma 28 may form heparin-resistant complexes on other sigma 28-dependent promoters or that heparin is actively destabilizing these complexes. However, the information from these studies is useful in comparing the relative stability of holoenzymes carrying different sigma  factors.

In the case of sigma 70, it has been shown that amino-terminal region 1.1 is involved in inhibition of DNA binding by sigma  (10, 56) and is required for efficient open complex formation by holoenzyme. Region 1.2, which is typically present in all primary and alternative sigma  factors, may also be involved in open complex formation for Esigma 70 (11). sigma 28 is a very unusual member of the sigma 70 family of proteins since it lacks any homology to region 1.2, leading to some speculation regarding how this difference in structure may translate into a difference in function. The flagellar biosynthesis sigma  factor from B. subtilis, sigma D, retains homology to region 1.2. Esigma D forms two closed complexes and isomerizes through several open complex intermediates (55). sigma 28, on the other hand, lacks both regions 1.1 and 1.2. Esigma 28 forms an unusually short RPC1, and we did not observe an RPC2-like intermediate. Additionally, many Esigma 70 open complexes are stable to heparin either in the absence or presence of NTPs, and open complexes do not form efficiently at low temperatures, whereas the opposite is true in both cases for Esigma 28. The composition of the amino terminus of the sigma  factor may affect the nature of the RNAP·DNA complexes that can be discerned during initiation. sigma 28 appears to facilitate transcription initiation at pfliC very efficiently by utilizing a mechanism with few intermediate complexes, but with reduced stability of the open complex. Whether this mechanism is a general phenomenon for all sigma 28-dependent promoters or whether it is characteristic for the flagellin gene promoter remains to be determined.

    ACKNOWLEDGEMENTS

We thank Dr. K. Hughes for plasmids containing the fliA and fliC genes. We are grateful to Dr. W. Ross for sharing footprinting protocols. We thank Drs. R. Gourse and T. Gaal for core RNA polymerase with a C-terminal deletion of the alpha  subunit. We thank the other members of the laboratory, N. Baldwin, B. Johnson, and C. Wilson Bowers, for helpful discussions and critical reading of the manuscript. We are grateful to M. Schaubach for assistance with computer graphics.

    FOOTNOTES

* This work was supported by Research Grant NP-902 from the American Cancer Society and Research Grant GM56453-01 from the National Institutes of Health.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed: Dept. of Microbiology and Molecular Genetics, University of Texas Health Science Center, 6431 Fannin, Houston, TX 77030. Tel.: 713-500-5442; Fax: 713-500-5499; E-mail: dombros{at}utmmg.med.uth.tmc.edu.

2 The sequences are available upon request.

    ABBREVIATIONS

The abbreviations used are: RNAP, RNA polymerase; Esigma 28, sigma 28 holoenzyme.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
  1. Burgess, R. R., Travers, A. A., Dunn, J. J., and Bautz, E. K. F. (1969) Nature 221, 43-46[CrossRef][Medline] [Order article via Infotrieve]
  2. Travers, A. A., and Burgess, R. R. (1969) Nature 222, 537-540[CrossRef][Medline] [Order article via Infotrieve]
  3. Losick, R., and Pero, J. (1981) Cell 25, 582-584[CrossRef][Medline] [Order article via Infotrieve]
  4. Lonetto, M., Gribskov, M., and Gross, C. (1992) J. Bacteriol. 174, 3843-3849[Free Full Text]
  5. Stragier, P., Parsot, C., and Bouvier, J. (1985) FEBS Lett. 187, 11-15[CrossRef][Medline] [Order article via Infotrieve]
  6. Helmann, J. D., and Chamberlin, M. J. (1988) Annu. Rev. Biochem. 57, 839-872[CrossRef][Medline] [Order article via Infotrieve]
  7. Gross, C. A., Lonetto, M., and Losick, R. (1992) in Transcriptional Regulation (McKnight, S. L., and Yamamoto, K. R., eds), Vol. 1, pp. 129-176, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  8. Harley, C. B., and Reynolds, R. P. (1987) Nucleic Acids Res. 15, 2343-2361[Abstract/Free Full Text]
  9. Estrem, S. T., Gaal, T., Ross, W., and Gourse, R. L. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 9761-9766[Abstract/Free Full Text]
  10. Dombroski, A. J., Walter, W. A., Record, M. T., Jr., Siegele, D. A., and Gross, C. A. (1992) Cell 70, 501-512[CrossRef][Medline] [Order article via Infotrieve]
  11. Wilson, C., and Dombroski, A. J. (1997) J. Mol. Biol. 267, 60-74[CrossRef][Medline] [Order article via Infotrieve]
  12. Lesley, S. A., and Burgess, R. R. (1989) Biochemistry 28, 7728-7734[CrossRef][Medline] [Order article via Infotrieve]
  13. Zhou, Y. N., Walter, W. A., and Gross, C. A. (1992) J. Bacteriol. 174, 5005-5012[Abstract/Free Full Text]
  14. Joo, D. M., Ng, N., and Calendar, R. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 4907-4912[Abstract/Free Full Text]
  15. Jones, C. H., and Moran, C. P., Jr. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 1958-1962[Abstract/Free Full Text]
  16. Jones, C. H., Tatti, K. M., and Moran, C. P., Jr. (1992) J. Bacteriol. 174, 6815-6821[Abstract/Free Full Text]
  17. Juang, Y.-L., and Helmann, J. D. (1994) J. Mol. Biol. 235, 1470-1488[CrossRef][Medline] [Order article via Infotrieve]
  18. Rong, J. C., and Helmann, J. D. (1994) J. Bacteriol. 176, 5218-5224[Abstract/Free Full Text]
  19. Roberts, C. W., and Roberts, J. W. (1996) Cell 86, 495-501[CrossRef][Medline] [Order article via Infotrieve]
  20. Huang, X., Lopez de Saro, F. J., and Helmann, J. D. (1997) Nucleic Acids Res. 25, 2603-2609[Abstract/Free Full Text]
  21. Marr, M. T., and Roberts, J. W. (1997) Science 276, 1258-1260[Abstract/Free Full Text]
  22. Gardella, T. H., Moyle, H., and Susskind, M. M. (1989) J. Mol. Biol. 206, 579-590[CrossRef][Medline] [Order article via Infotrieve]
  23. Siegele, D. A., Hu, J. C., Walter, W. A., and Gross, C. A. (1989) J. Mol. Biol. 206, 591-603[CrossRef][Medline] [Order article via Infotrieve]
  24. Zuber, P., Healy, J., Carter, H. L., III, Cutting, S, Moran, C. P., Jr., and Losick, R. (1989) J. Mol. Biol. 206, 605-614[CrossRef][Medline] [Order article via Infotrieve]
  25. Daniels, D., Zuber, R., and Losick, R. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 8075-8079[Abstract/Free Full Text]
  26. Waldburger, C., Gardella, T., Wong, R., and Susskind, M. M. (1990) J. Mol. Biol. 215, 267-276[CrossRef][Medline] [Order article via Infotrieve]
  27. Record, M. T., Jr., Reznikoff, W. S., Craig, M. L., McQuade, K. L., and Schlax, P. J. (1996) in Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology (Neidhardt, F. C., ed), 2 Ed., Vol. 1, pp. 792-820, ASM Press, Washington, D. C.
  28. Suh, W.-C., Ross, W., and Record, M. T., Jr. (1993) Science 259, 358-361[Abstract/Free Full Text]
  29. Zaychikov, E., Denissova, L., Meier, T., Gotte, M., and Heumann, H. (1997) J. Biol. Chem. 272, 2259-2267[Abstract/Free Full Text]
  30. Kovacic, R. T. (1987) J. Biol. Chem. 262, 13654-13661[Abstract/Free Full Text]
  31. Cowing, D. W., Mecsas, J., Record, M. T., Jr., and Gross, C. A. (1989) J. Mol. Biol. 210, 521-530[CrossRef][Medline] [Order article via Infotrieve]
  32. Mecsas, J., Cowing, D. W., and Gross, C. A. (1991) J. Mol. Biol. 220, 585-597[CrossRef][Medline] [Order article via Infotrieve]
  33. Kutsukake, K., Ohya, Y., and Iino, T. (1990) J. Bacteriol. 172, 741-747[Abstract/Free Full Text]
  34. Ohnishi, K., Kutsukake, K., Suzuki, H., and Iino, T. (1990) Mol. Gen. Genet. 221, 139-147[Medline] [Order article via Infotrieve]
  35. Gillen, K. L., and Hughes, K. T. (1991) J. Bacteriol. 173, 6453-6459[Abstract/Free Full Text]
  36. Ohnishi, K., Kutsukake, K., Suzuki, H., and Iino, T. (1992) Mol. Microbiol. 6, 3149-3157[Medline] [Order article via Infotrieve]
  37. Shorenstein, R. G., and Losick, R. (1973) J. Biol. Chem. 248, 6170-6173[Abstract/Free Full Text]
  38. Chen, Y.-F., and Helmann, J. D. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 5123-5127[Abstract/Free Full Text]
  39. Aiyar, S. E., Juang, Y.-L., Helmann, J. D., and deHaseth, P. L. (1994) Biochemistry 33, 11501-11506[CrossRef][Medline] [Order article via Infotrieve]
  40. Johnson, B. D., and Dombroski, A. J. (1997) J. Biol. Chem. 272, 31029-31035[Abstract/Free Full Text]
  41. Dombroski, A. J. (1997) J. Biol. Chem. 272, 3487-3494[Abstract/Free Full Text]
  42. Newlands, J. T., Ross, W., Gosink, K. K., and Gourse, R. L. (1991) J. Mol. Biol. 220, 569-583[CrossRef][Medline] [Order article via Infotrieve]
  43. Craig, M. L., Suh, W.-C., and Record, M. T., Jr. (1995) Biochemistry 34, 15624-15632[CrossRef][Medline] [Order article via Infotrieve]
  44. Duval-Valentin, G., and Ehrlich, R. (1986) Nucleic Acids Res. 14, 1967-1983[Abstract/Free Full Text]
  45. Hofer, B., Muller, D., and Koster, H. (1985) Nucleic Acids Res. 16, 5995-6013
  46. Spassky, A., Kirkegaard, K., and Buc, H. (1985) Biochemistry 24, 2723-2731[CrossRef][Medline] [Order article via Infotrieve]
  47. Gralla, J. D., Hsieh, M., and Wong, C. (1993) in Footprinting of Nucleic Acid-Protein Complexes (Revzin, A., ed), pp. 107-128, Academic Press, Orlando, FL
  48. Duval-Valentin, G., and Ehrlich, R. (1987) Nucleic Acids Res. 15, 575-594[Abstract/Free Full Text]
  49. Melancon, P., Burgess, R. R., and Record, M. T., Jr. (1982) Biochemistry 21, 4318-4331[CrossRef][Medline] [Order article via Infotrieve]
  50. Roe, J., Burgess, R. R., and Record, M. T., Jr. (1984) J. Mol. Biol. 176, 495-521[CrossRef][Medline] [Order article via Infotrieve]
  51. Buc, H., and McClure, W. R. (1985) Biochemistry 24, 2712-2723[CrossRef][Medline] [Order article via Infotrieve]
  52. Gourse, R. L. (1988) Nucleic Acids Res. 16, 9789-9809[Abstract/Free Full Text]
  53. Whipple, F. W., and Sonenshein, A. L. (1992) J. Mol. Biol. 223, 399-414[CrossRef][Medline] [Order article via Infotrieve]
  54. Taylor, W. E., and Burgess, R. R. (1979) Gene (Amst.) 6, 331-365[CrossRef][Medline] [Order article via Infotrieve]
  55. Chen, Y.-F., and Helmann, J. D. (1997) J. Mol. Biol. 267, 47-59[CrossRef][Medline] [Order article via Infotrieve]
  56. Dombroski, A. J., Walter, W. A., and Gross, C. A. (1993) Genes Dev. 7, 2446-2455[Abstract/Free Full Text]
  57. Fredrick, K., and Helmann, J. D. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 4982-4987[Abstract/Free Full Text]

Copyright © 1999 by The American Society for Biochemistry and Molecular Biology, Inc.
Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J. Bacteriol.Home page
L. Shen, X. Feng, Y. Yuan, X. Luo, T. P. Hatch, K. T. Hughes, J. S. Liu, and Y.-x. Zhang
Selective Promoter Recognition by Chlamydial {sigma}28 Holoenzyme
J. Bacteriol., November 1, 2006; 188(21): 7364 - 7377.
[Abstract] [Full Text] [PDF]

Home page
 J. Bacteriol.Home page
J. R. Givens, C. L. McGovern, and A. J. Dombroski
Formation of Intermediate Transcription Initiation Complexes at pfliD and pflgM by {sigma}28 RNA Polymerase
J. Bacteriol., November 1, 2001; 183(21): 6244 - 6252.
[Abstract] [Full Text] [PDF]

Home page
 Microbiol. Mol. Biol. Rev.Home page
G. S. Chilcott and K. T. Hughes
Coupling of Flagellar Gene Expression to Flagellar Assembly in Salmonella enterica Serovar Typhimurium and Escherichia coli
Microbiol. Mol. Biol. Rev., December 1, 2000; 64(4): 694 - 708.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Schaubach, O. L.
Right arrow Articles by Dombroski, A. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Schaubach, O. L.
Right arrow Articles by Dombroski, A. J.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 1999 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement