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J. Biol. Chem., Vol. 281, Issue 37, 27205-27215, September 15, 2006

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Transcriptional Specificity of RpoN1 and RpoN2 Involves Differential Recognition of the Promoter Sequences and Specific Interaction with the Cognate Activator Proteins^*

Sebastian Poggio, Aurora Osorio, Georges Dreyfus, and Laura Camarena¹

From the Departamento de Biología Molecular y Biotecnología, Instituto de Investigaciones Biomédicas, and Departamento de Genética Molecular, Instituto de Fisiología Celular, Universidad Nacional Autónoma de México, 04510 México D. F., México

Received for publication, February 23, 2006 , and in revised form, July 5, 2006.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The four RpoN factors of Rhodobacter sphaeroides are functionally specialized. In this bacterium, RpoN1 and RpoN2 are specifically required for the transcription of the nitrogen fixation and flagellar genes, respectively. Analysis of the promoter sequences recognized by each of these RpoN proteins revealed some significant differences. To investigate the functional relevance of these differences, the flagellar promoter fliOp was sequentially mutagenized to resemble the nitrogen fixation promoter nifUp. Our results indicate that the promoter sequences recognized by these sigma factors have diverged enough so that particular positions of the promoter sequence are differentially recognized. In this regard, we demonstrate that the identity of the -11-position is critical for promoter discrimination by RpoN1 and RpoN2. Accordingly, purified RpoN proteins with a deletion of Region I, which has been involved in the recognition of the -11-position, did not show differential binding of fliOp and nifUp promoters. Substitution of the flagellar enhancer region located upstream fliOp by the enhancer region of nifUp allowed us to demonstrate that RpoN1 and RpoN2 interact specifically with their respective activator protein. In conclusion, two different molecular mechanisms underlie the transcriptional specialization of these sigma factors.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
In eubacteria, the sigma subunit of the RNA polymerase is responsible for recognizing the promoter sequence to initiate transcription (1, 2). A large number of sigma factors have been described in eubacteria (1, 3, 4). In general, all of them can be grouped in two different families: the family of sigma-70 that in Escherichia coli includes ⁷⁰, ³², ²⁴, ^S, and ²⁸ and the family of sigma-54, of which ⁵⁴ is the only member (5-8). The RNA polymerase core (E) associated with a sigma factor of the ⁷⁰ family is capable of binding to a promoter and initiating transcription without any ancillary factor. In contrast, E⁵⁴ holoenzyme is unable to form open complex in the absence of an activator protein (9-11). E⁵⁴ binds to highly conserved promoters with the consensus sequence TGGCACN₅TTGCW, of which the most conserved positions are the GG and GC located at -24 and -12 nucleotides upstream from the transcription initiation site (12, 13). Mutation of the GG or GC dinucleotides in any ⁵⁴ promoter strongly reduces transcription, confirming its relevance in the transcriptional initiation process (14). The high degree of conservation of the ⁵⁴-dependent promoters in many bacteria allows the ⁵⁴ factor from Aquifex aeolicus to successfully recognize the ⁵⁴-dependent promoter glnHp2 from E. coli (15).

As mentioned above, E⁵⁴ strictly requires the presence of an activator protein to carry out the transition from closed to open complex. Experimental evidence obtained from in vitro studies using heteroduplex DNA fragments containing a ⁵⁴ promoter sequence mimicking a fork junction suggests that the inability of E⁵⁴ holoenzyme to spontaneously melt DNA is based on the interaction between the N-terminal region of ⁵⁴ and a repressive DNA fork formed transitorily by melted DNA at the -11- and -10-positions,² which are adjacent to the GC dinucleotide. This interaction inhibits the ability of the holoenzyme to spread melting and keeps it inactive to initiate transcription. Therefore, it has been proposed that the bases in the -12 box have a regulatory role in the isomerization process (16-20).

Transcriptional activators of E⁵⁴ usually bind to sequence motifs located 100 bp upstream of the promoter sequence and for this reason are commonly known as enhancer-binding proteins (EBPs).³ These proteins belong to the family of AAA+ ATPases and couple the energy derived from ATP hydrolysis to remodel the nucleoprotein complex formed by the N terminus of ⁵⁴ bound to the DNA fork structure, relieving the inhibitory interactions and allowing open complex formation (21-23).

By sequence alignment, ⁵⁴ has been divided in three regions. Region I is located at the N terminus of the protein and contains determinants for nucleating DNA melting, for inhibiting open complex formation, and to mediate the response to the EBPs (24, 25). Recently, it has been established that Region I makes a major contribution to bind the fork junction structure located at the -11-position in heteroduplex probes (16-20). Region II is variable, and although its role in transcription is still unclear, it has been implicated in assisting ⁵⁴ binding to DNA and melting (26, 27). At the C terminus, Region III shows a helixturn-helix motif and a very conserved amino acid sequence named the RpoN box; both have been proposed to interact with DNA (21). Recently, it was suggested that some residues in the RpoN box interact with the -24 promoter sequence (28, 29).

Results obtained using (p-bromoacetamidobenzyl)-EDTA Fe-⁵⁴ conjugates indicate that in addition to ⁵⁴ Region I, the residue Arg³³⁶, which lies in Region III, is also close to the -12 promoter box (20). This result agrees with previous observations, where mutants in Arg³³⁶, as well as mutants in Region I, were defective in controlling open complex formation. Therefore, two different structural elements of ⁵⁴ have been proposed to be part of the regulatory center that controls melting (30, 31).

The complete genome sequence of several microorganisms has shown that the rpoN gene (encoding the ⁵⁴ factor) is widely distributed among eubacteria. A few organisms have two copies of rpoN, such as Bradyrhizobium japonicum, Rhizobium etli (32, 33), Ralstonia solanacearum, and Burkholderia fungorum. In B. japonicum, these copies are highly similar, and they are functionally interchangeable (32). Rhodobacter sphaeroides is the only known bacterium with four copies of rpoN in its genome. This unusual high number of rpoN gene copies does not seem to be the result of recent events of duplication, since the products of these genes show a degree of similarity ranging from 48 to 51%. We have previously shown that these copies are not functionally interchangeable (34, 35). RpoN1 is involved in the expression of the genes required for nitrogen fixation, whereas RpoN2 is required for the transcription of the class II and class III flagellar genes. The molecular determinants that confer this specificity have not yet been investigated. In this work, we show evidence that the promoter sequences recognized by RpoN1 and RpoN2 have subtle differences that are used by these proteins to bind differentially; furthermore, our results indicate that each sigma factor is activated only by its cognate activator protein (EBP).

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Bacterial Strains, Plasmids, and Growth Conditions—Plasmids and strains of R. sphaeroides and E. coli used in this work are listed in Table 1. R. sphaeroides was grown in Sistrom's minimal medium (36) or in minimal medium with malate (0.4%) and glutamate as sole carbon and nitrogen sources, respectively (MG medium) (34). For the N⁺-aerobic growth conditions, a saturated culture was diluted 1:10 with fresh Sistrom's medium in a 250-ml Erlenmayer flask. The cultures were incubated in the dark with strong shaking (200 rpm). To achieve nitrogen fixing conditions, R. sphaeroides was grown as described before (34). Briefly, a culture grown under heterotrophic conditions was collected when it reached an A₆₀₀ of 0.4. After two washes with minimal medium, the cells were suspended in the original volume of MG medium. This culture was used to completely fill screw cap tubes that were incubated under constant illumination for a period of 12 h. Antibiotics were used at the following concentrations: for R. sphaeroides, spectinomycin (50 µg/ml), kanamycin (25 µg/ml), tetracycline (1 µg/ml), and gentamycin (5 µg/ml); for E. coli, spectinomycin (100 µg/ml), kanamycin (50 µg/ml), tetracycline (15 µg/ml), gentamycin (30 µg/ml), and ampicillin (100 µg/ml). E. coli strains were grown in LB medium at 37 °C with shaking at 200 rpm.

Site-directed Mutagenesis—Site-directed mutagenesis was performed according to the method of Kunkel (37) with a uracil-containing single-stranded DNA as template. The plasmid carrying the fliOp promoter consists of a 391-bp DNA fragment, cloned in pTZ19R. This DNA fragment carries 254 bp upstream of the transcriptional start site (38). The oligonucleotides used for mutagenesis were as follows: 5'-GTCCCCCTCCGCNGCAACATCCGTGCCG-3', 5'-CAACATCCGTGCCAAGCGCCCGCGTGAGGATC-3', and 5'-GTCCCCCTCCGCAGCACGATCCGTGCCAAGCGCC-3'. The presence of the desired mutation in the resultant plasmid was confirmed by sequencing. Finally, the fragments carrying the wild-type promoter or its derivatives were subcloned into pBBRMCS53 in the appropriate orientation.

Conjugation—Plasmid DNA was mobilized into R. sphaeroides by conjugation according to previously reported procedures (39).

-Glucuronidase Assay—-Glucuronidase was determined from sonicated cell-free extracts using 4-methyl-umbelliferyl--D-glucuronide as substrate and following a previously reported protocol (40). Briefly, cell-free extracts were incubated at 37 °C in reaction buffer. Samples of 100 µl were taken at three different time points and mixed with 0.9 ml of stop buffer (0.2 M Na₂CO₃). Fluorometric determinations were made in a PerkinElmer Life Sciences LA-5 apparatus (excitation wavelength, 360 nm; emission wavelength, 446 nm). The fluorometer was calibrated using 4-methylumbelliferone standards. Specific activities are expressed as nmol of 4-methylum-belliferone formed/min/mg of protein.

DNA and Proteins—Synthetic PAGE-purified oligonucleotides, corresponding to the -59 to +22 sequence of the fliOp promoter (38) or the -58 to +23 sequence of the predicted nifUp promoter were used to construct the heteroduplex molecules used in this work (see Fig. 4). DNA probes were prepared as follows. The top strand was labeled with [-³²P]ATP and mixed with the complementary strand. The mixture containing 4 pmol of ³²P-end-labeled DNA and 6 pmol of complementary strand in 20 mM Tris-HCl, pH 7.5, and 80 mM NaCl was boiled briefly and cooled slowly to room temperature. The resulting annealed probes were then diluted in TE (10 mM Tris-HCl, pH 8.0, 1 mM EDTA) containing 80 mM NaCl.

To obtain RpoN1 and RpoN2 proteins, the coding regions of rpoN1 and rpoN2 were amplified by PCR and cloned into pBADHis-C vector. The resultant plasmids were introduced to the LMG174-1 strain, and different growth temperatures and arabinose concentrations were tested in order to determine the optimal condition to overproduce these polypeptides. Both proteins remained in the insoluble fraction in all tested conditions, and for this reason, the proteins were purified from the inclusion bodies. The procedure for protein purification is based on previously reported methods (41) with minor modifications. Briefly, after 3 h of induction at 30 °C with 0.02% arabinose, the cells were lysed by sonication at 4 °C. The insoluble fraction was extracted with TGED buffer containing 1 M NaCl and 1 mM phenylmethylsulfonyl fluoride or a mixture of protease inhibitors manufactured by Roche Applied Science, and a second extraction with 1% Triton X-100 was carried out. Detergent was removed by washing three times with TGED. Finally, the pellet was carefully resuspended in 8 M urea buffered with TGED. The protein was dialyzed overnight at 4 °C against TGED buffer containing 250 mM NaCl, followed by a further 6-h dialysis with fresh TGED buffer containing 100 mM NaCl. Insoluble material was removed by centrifugation. The oligonucleotides 5'-AGATCTGGTGCGCGTGCCCGAGGGGGC-3' and 5'-AGATCTGCTGCGCCTGCCGCCGGCGCC-3' were used as forward primers to obtain the PCR product coding for RpoN1I and RpoN2I. These products were cloned into pBAD/His-C and introduced into LMG174-1 strain. To obtain the polypeptides corresponding to Region I of RpoN1 and RpoN2, the DNA region encoding the first 57 residues of these proteins was amplified by PCR using as reverse primers the 57Nif or 57Fli oligonucleotide (5'-GGAATTCTCACTCGAGGCAGGGATTTTC-3' or 5'-GAATTCTCATTCGATGAAAGGGTTTTC-3', respectively), which include TCA bases as the stop codon. These proteins were purified following the procedure described above.

Circular Dichroism—Previous to CD analysis, the proteins were dialyzed against 10 mM sodium phosphate buffer, pH 8.0, 10 mM NaCl, and 20% glycerol, with three changes of the dialysis buffer, each of 100 times the sample volume. Dialysis was carried out over a period of 24 h at 4 °C. Molar ellipticity values were obtained using an Aviv Biomedical CD spectrophotometer model 202-01. The resulting spectra were analyzed using CDPro (available on the World Wide Web at lamar.colostate.edu/~sreeram/CDPro/main.html).

Gel Mobility Shift Assay—Binding reactions were carried out at 30 °C in STA buffer (24) in a total volume of 20 µl. The reactions included 1 µg of poly(dI-dC) as nonspecific competitor, DNA probe labeled with ³²P, and a 50, 100, or 200 µM concentration of the indicated protein. After 20 min, the samples were loaded on a 4.5% native polyacrylamide gel and run at 12 mA in Tris-glycine buffer. After electrophoresis, radioactivity was visualized and quantified using a PhosphorImager apparatus (Bio-Rad) or by autoradiography using ImageJ software.

View larger version (37K): [in this window] [in a new window]   FIGURE 1. Logo sequences of the 54-dependent promoters from R. sphaeroides. The promoter sequences were obtained from the analysis of the complete genome of R. sphaeroides (59) (available on the World Wide Web at mmg.uth.tmc.edu/sphaeroides/). The nif consensus includes six sequences located upstream of genes involved in nitrogen fixation (nifB, nifH, nifE, orf-nifU, rnfA, and fprA). The fli consensus includes eight promoters located upstream of genes involved in flagellar biogenesis (fleT, fliK, fliO, flhA, orf-flgA, flgB, flgG, and motA). The initiation site for four of the fli promoters has been identified previously (35, 38). The logo was created using WebLogo (available on the World Wide Web at weblogo.berkeley.edu/) (60). The promoter numbering given in this work is taking the dinucleotide GC as the -13- and -12-positions, respectively.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

Evaluation of the Contribution of the Promoter Sequence Bias to Sigma Factor Specificity—To test if the differences in the promoter sequences mentioned above might be involved in the differential recognition and expression mediated by RpoN1 and RpoN2, we mutagenized the nucleotides of fliOp for those present in the sequence of nifUp. Transcriptional fusions of these promoter versions with the reporter gene uidA (encoding -glucuronidase) were constructed using pBBMCS53. The amount of -glucuronidase expressed from these plasmids was determined for WS8 cells grown under N⁺-aerobic conditions, which would maintain a low level of RpoN1 (43). Compared with the level of -glucuronidase produced by the wild-type fliOp promoter, a 5-fold reduction was observed when changes were introduced at positions -28, -27, and -26, and a 9-fold reduction was observed when changes were introduced at positions -15 and -16. When these changes where combined in a single construction, the promoter activity was reduced by only 5-fold (Fig. 2). It is noteworthy that when the wild-type A at the -11-position was substituted by any other nucleotide, a reduction of 50-100-fold in the promoter activity was observed, suggesting a major contribution of this particular nucleotide in the activity of the fliOp promoter. For simplicity, the changes at -28, -27, and -26 will be referred to hereafter as up-24, and the changes at -16 and -15 will be referred to as up-12.

All previous constructions were introduced into the SP7/pRS210 strain, which carries a mutation in rpoN2 but properly expresses the fleT and fleQ genes encoding the flagellar EBPs (35). As shown in Fig. 2, in this strain, the activities of all of these promoters were strongly reduced, indicating that RpoN2 was responsible for the activity detected in WS8 cells. As mentioned above, the expression of RpoN1 would be at a low level under the growth conditions used in the previous experiments (43). Therefore, to correctly evaluate if RpoN1 could promote transcription from these promoters, we cultured these strains under nitrogen-fixing conditions, as described under "Experimental Procedures." As an induction control, the plasmid carrying the fusion nifUp-uidA was included in the assay. All of the fliOp promoter versions tested in the WS8 strain showed a similar amount of -glucuronidase activity to that observed under N⁺-aerobic conditions (Fig. 3), indicating that these promoters are not significantly expressed by RpoN1. A slight increase in the activity level could be detected in SP7/pRS210 cells grown under nitrogen-fixing conditions, when the fliOp promoter more closely resembled nifUp (Fig. 3D), although the -glucuronidase activity is only 5% of that observed for nifUp under the same condition. From these results, we hypothesized that the RpoN1 holoenzyme was either not able to efficiently recognize any of these promoters or that the flagellar EBP bound to the enhancer region (UAS) was not able to remodel the regulatory center created by the RpoN1 holoenzyme and the promoter. To investigate these possibilities, we replaced the enhancer region of the fliOp wild-type and mutant promoter versions with the nifUp enhancer region, which is recognized by NifA; we will refer to the resultant promoters as fliOp-Nif. These chimeric promoters were introduced into WS8 and SP7/pRS210 strains, and the amount of -glucuronidase was determined from cells grown under nitrogen-fixing conditions or in the presence of ammonium in the culture medium. Remarkably, when the SP7/pRS210 strain was grown under nitrogenfixing conditions (Fig. 4D), the level of -glucuronidase promoted by fliOp-Nif (up-24, up-12, and -11T) was similar to that promoted by nifU. In contrast, fliOp-Nif (up-24 and up-12) and fliOp-Nif (-11T) showed a reduction of 90% as compared with the activity detected for nifUp. The wild-type fliOp-Nif showed a very low level of activity (only 0.28% of the activity detected for nifUp) (Fig. 4D). These results suggest that RpoN1 recognizes a different promoter sequence from that recognized by RpoN2 and that the flagellar and nitrogen fixation EBPs preferentially interact with or activate a particular RpoN protein. In contrast to what occurs with RpoN2, for RpoN1 the -11-position is as important as the nucleotides at -28, -27, and -26 and at -16 and -15, since mutagenesis of the -11-position has the same effect as mutagenesis of the other positions (compare the amount of -glucuronidase promoted by fliOp-Nif (-11T) with that promoted by fliOp-Nif (up-12 and up-24) in SP7/pRS210) (Fig. 4D).

View larger version (25K): [in this window] [in a new window]   FIGURE 2. Activities promoted by the wild-type and mutant versions of fliOp. Specific -glucuronidase activities were determined from different strains carrying the mutant versions or the wild-type fliOp promoter, fused to uidA. The conserved dinucleotides GG and GC located in the -24 and -12 promoter boxes are shaded. The asterisk indicates the -11-position. The sequence of the nifUp promoter is shown for comparison. The nucleotides mutagenized to resemble nifUp promoter are underlined in each case. Values are the mean of at least three independent culture determinations that showed <20% variation. The activities are expressed as nmol of 4-methylumbelliferone/min/mg of protein. The changes made at positions -28, -27, and -26 or at positions -16 and -15 are referred as up-24 and up-12, respectively.

View larger version (21K): [in this window] [in a new window]   FIGURE 3. Activity of the wild-type and mutant versions of fliOp under nitrogen-fixing conditions. Relative -glucuronidase activities determined from cells grown in nitrogen-fixing or N+-aerobic conditions. Values are given as a percentage of the activity promoted by fliOp in WS8 cells grown in N+-aerobic conditions (466 nmol of 4-methylumbelliferone/min/mg of protein). Values are the mean of at least three independent culture determinations that showed <20% variation.

View larger version (25K): [in this window] [in a new window]   FIGURE 4. Activity of the wild-type and mutant versions of fliOp-Nif. Relative -glucuronidase activities determined from cells grown in nitrogen-fixing or N+-aerobic conditions. Values are given as a percentage of the activity promoted by nifUp in WS8 cells grown in nitrogen-fixing conditions (511.8 nmol of 4-methylumbelliferone/min/mg of protein). Values are the mean of at least three independent culture determinations that showed <20% variation.

All of the promoters were practically inactive in SP13 strain (fleQ mutant) grown under N⁺-aerobic conditions (Fig. 4E), confirming that the activity observed in the WS8 strain grown under the same conditions was dependent on RpoN2 and FleQ/FleT. In nitrogen-fixing conditions, only the promoters recognized by RpoN1 showed a high level of reporter gene activity (Fig. 4F), although RpoN2 is present and must be bound to the fliOp-Nif (WT) and fliOp-Nif (up-24 and up-12) promoters. This result implies that NifA is not able to activate RpoN2 efficiently.

View larger version (17K): [in this window] [in a new window]   FIGURE 5. Activity of the wild-type and mutant versions of fliOp in the rpoN1, nifA double mutant. Relative -glucuronidase activities determined from SP17 cells grown in nitrogen-fixing or N+-aerobic conditions. Values are given as percentage of the activity promoted by fliOp in SP17 cells grown in N+-aerobic conditions (569.8 nmol of 4-methylumbelliferone/min/mg of protein). Values are the mean of at least three independent culture determinations that showed <20% variation.

The idea that fliOp-Nif (WT) and fliOp-Nif (up-24 and up-12) are activated from solution by FleQ/FleT is confirmed by the lack of activity of these promoters in the SP13 strain, whereas in SP16 these promoters show a level of activity similar to that in the WS8 strain.

To clearly determine the preference of RpoN2 by the wild-type and mutant versions of fliOp, we introduced these constructions into SP17 (carrying a mutation in nifA and rpoN1); in this strain, RpoN2 should be activated only by the flagellar EBP and would not compete with RpoN1 for promoter binding. As shown in Fig. 5, fliOp (-11T) and fliOp (up-24, up-12, and -11T), promoted a low -glucuronidase activity (1.6 and 3% of the activity detected for fliOp, respectively), whereas fliOp (up-24 and up-12) showed a 5-fold reduction regarding the activity promoted by fliOp. These results are in agreement with those observed in WS8 cells under N⁺-aerobic conditions (Fig. 3A).

Although the GG and GC nucleotides are central for promoter recognition by the ⁵⁴ factors (21, 38), our results allow us to conclude that additional information in the promoter sequence contributes to the observed specificity of the RpoN proteins of R. sphaeroides. In particular, RpoN2 shows a strong dependence on the identity of the -11-position and a minor dependence on the identity of the nucleotides located upstream of the -24- and -12-positions. In contrast, RpoN1 depends equally on the bases adjacent to the -24- and -12-positions and on the identity of the -11-position, although the -11- position seems to be by itself more relevant than any of the -24- and -12-positions independently. Transcription by the proper RpoN protein is further assured by the strict interaction of the activator proteins with their cognate sigma factor.

Purification of the RpoN1 and RpoN2 Proteins—To better understand the molecular mechanism of promoter discrimination by RpoN1 and RpoN2, we decided to study the in vitro properties of these proteins. For this purpose, we purified RpoN1 and RpoN2 fused to a His₆ tag. The coding region of each gene without the initiator methionine was cloned into pBADHisC plasmid producing a fusion of six residues of histidine at the N terminus of each protein. Both constructions were introduced in an E. coli strain carrying a lesion in rpoN to avoid any contamination with endogenous RpoN protein. RpoN1 and RpoN2 were induced, adding arabinose to exponentially growing cultures; both proteins remained in the insoluble fraction of the cell extract, even when the amount of arabinose was reduced (data not shown). The proteins were obtained from inclusion bodies and refolded as described previously for some RpoN mutant versions from Klebsiella pneumoniae (41). The same procedure was followed to obtain RpoN proteins lacking Region I. The purity of all of the proteins was verified by SDS-PAGE (supplemental Fig. 1).

Binding of RpoN1, RpoN2, RpoN1I, and RpoN2I to Early Melted Probes—We were not able to observe binding of RpoN1 and RpoN2 to fliOp as a homoduplex in electrophoretic mobility shift assays (data not shown). A similar situation has been reported for RpoN from enteric bacteria, where, depending on the promoter, RpoN binds with a low affinity or does not bind at all (48-50). From this, we conclude that RpoN1 and RpoN2 are not particularly deficient in the binding of homoduplex DNA promoters.

To analyze binding to early melted promoters, we initially used two different probes corresponding to the sequence of fliOp and nifUp. In both cases, the -11- and -10-positions of the nontemplate strand were changed to simulate the early melted state of these promoters (Fig. 6). It should be noticed that this would mean changing the bases that until now have been implicated in the specificity of the RpoN proteins. However, as previously mentioned, it has been shown that in the closed complex, RpoN binds tightly to the -11- and -10-positions of the template strand, exposed transitorily before activation (16, 17), which in these probes remain unchanged. However, according to the nomenclature for the identification of sequences in promoters, we refer to these bases by their identity at the nontemplate strand.

View larger version (37K): [in this window] [in a new window]   FIGURE 6. Relevant features of the probes used in the gel mobility shift assays. R. sphaeroides fliOp and nifUp promoter sequences from which synthetic oligonucleotides were designed to obtain heteroduplex probes. The sequences depicted at the top and bottom show a section of the wild-type promoter sequences of fliOp and nifUp promoters, respectively. The final structure of the heteroduplex probes near the -12 consensus element is shown. The consensus GG and GC dinucleotides are in boldface type and shaded, respectively. The unpaired nucleotides correspond to the -11- and -10-positions.

View larger version (43K): [in this window] [in a new window]   FIGURE 7. Gel mobility shift assays with RpoN1 or RpoN2 proteins. The heteroduplex probes used in these assays are indicated above each panel. RpoN1 and RpoN2 were included in the reaction at the following concentrations: 50 µM (+), 100 µM (++), and 200 µM (+++). The numbers above each lane represent the percentage of bound probe, calculated as the relation between total radioactivity in each lane and the radioactivity of the bound complexes. nd, not detected.

Since Region I of RpoN has been involved in the recognition of the -11-position, we evaluated if this region was involved in the differential binding. For this, we tested our probes with both RpoN proteins lacking Region I. As shown in Fig. 8, A and B, RpoN1I was able to bind to both probes to a comparable extent, suggesting that Region I of RpoN1 inhibits binding to fliOp. The retarded complex formed by this protein with both probes showed a faster migration than that formed with RpoN1 wild type or between RpoN2I and the fliOp probe (compare Fig. 8B, lanes 3 and 7), suggesting that the fast migrating complex formed by RpoN1I can be ascribed to a different conformation of the protein. Supporting this idea, we observed that RpoN1 forms a fast migration complex when it interacts with the Nif-Fli probe (see below; Fig. 10A). In contrast, RpoN2I was barely able to bind to fliOp, and no binding was detected with nifUp (Fig. 8, A and B). These results suggest that binding of RpoN2 is strongly dependent on the interactions made by Region I.

View larger version (38K): [in this window] [in a new window]   FIGURE 8. Binding of heteroduplex probes by RpoN1I or RpoN2I. The probes used in the gel mobility shift assay are indicated above each panel. RpoN1 and RpoN2 were included as controls. The amount of protein used in each reaction is as indicated in the legend to Fig. 7. The numbers above each lane represent the percentage of bound probe, calculated as the relation between total radioactivity in each lane and the radioactivity of the bound complexes. nd, not detected.

It has been reported that ⁵⁴I from K. pneumoniae does not bind efficiently to early melted DNA; however, binding is restored when Region I is added in the assay, indicating that Region I is able to complement binding of RpoNIin trans (19). The addition of the N-terminal region of RpoN1 or RpoN2 did not produce any change in the migration of the complexes previously detected, and no interaction could be detected between the N-terminal peptides and the fliOp or nifUp probes. Since the N-terminal peptides were purified from inclusion bodies, we measured their far-UV CD spectra to verify that they had folded appropriately. Both polypeptides had considerable amounts of secondary structure (data not shown). Nevertheless, in the absence of the solved structure of the RpoN protein, the possibility remains that the structure adopted by these proteins does not correspond to that found in the native protein.

View larger version (49K): [in this window] [in a new window]   FIGURE 9. Binding competition between RpoN1 and RpoN1I. Gel mobility shift assay showing the complexes formed by RpoN1 and RpoN1I with the nifUp heteroduplex probe. The amount of protein used in the reaction is as indicated in the legend to Fig. 7. The numbers above each lane represent the percentage of bound probe, calculated as the relation between total radioactivity in each lane and the radioactivity of the bound complexes.

Our first observation was that all of the proteins tested were less able to interact with these probes (Fig. 10), suggesting that the proper combination of promoter sequence and the identity of the bases at the fork structure is important for binding of RpoN1 and RpoN2. Nevertheless, we could still detect the interaction of RpoN1 or RpoN2 with these probes. Interestingly, binding of RpoN1 to the Nif-Fli probe yielded two complexes (Fig. 10A); one of them showed a similar migration to the complex formed with RpoN1I in the nifUp template (fast migration complex), whereas the other showed the usual migration of the complex formed by RpoN1 and the nifUp probe. The presence of the fast migration complex could indicate a defective interaction of Region I of RpoN1 with the fli bases at the fork. RpoN1I also binds to the Nif-Fli probe, but it only forms the fast migration complex (Fig. 10A). Remarkably, RpoN2 was able to form a complex with the Nif-Fli probe (Fig. 10A); since RpoN2 did not bind nifUp probe (see Fig. 7), this result supports the notion that recognition of the -11 and -10 nucleotides in the unpaired region makes an important contribution to the binding of RpoN2. As observed previously with the fliOp and nifUp probes, RpoN2I also binds poorly to the Nif-Fli and Fli-Nif probes, regardless of using a higher protein concentration (Fig. 10, A and B). As shown in Fig. 10B, neither RpoN1 nor RpoN2 bound efficiently to the Fli-Nif probe.

View larger version (48K): [in this window] [in a new window]   FIGURE 10. Binding of the RpoN and RpoNI proteins to the Nif-Fli and Fli-Nif probes. Gel mobility shift assay showing the complexes formed by RpoN1, RpoN2, RpoN1I, and RpoN2I with heteroduplex probes with interchanged sequences in the mismatch region. The probes used are indicated above each panel. The nucleotide sequence of these probes is shown in Fig. 4. The amount of protein used in the reaction is as indicated in the legend to Fig. 7. The numbers above each lane represent the percentage of bound probe, calculated as the relation between total radioactivity in each lane and the radioactivity of the bound complexes. nd, not detected.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

A closer inspection of the ⁵⁴ promoters revealed differences between the fli and nif promoters, which consisted in an extended consensus nif promoter and a marked differential preference at the -11-position (T for the nif and A for the fli promoters). The relevance of these variations for RpoN2-dependent transcription was tested in vivo by site-directed mutagenesis of the fliOp promoter. From these experiments, it was observed that substitution of the -11-position exerted the strongest negative effect, whereas substitution of the -28-, -27-, and -26-positions or -16- and -15-positions reduced RpoN2-dependent transcription less severely. The contribution of the bases upstream of the -24 and -12 boxes to the binding of RpoN2 was expected, since variations around the conserved dinucleotides GG and GC have been reported to affect RpoN-dependent transcription in a moderate degree (13, 51). In contrast, the remarkable dependence on the identity of the -11-position was surprising, since changes at this position were previously reported to affect RpoN-dependent transcription between 10 and 50%, depending on the identity of the substitution (51). In our case, the identity of the -11-position seems to be critical to allow RpoN2-dependent transcription, since any base different from A reduced the fliOp activity in more than 90% (Fig. 2).

To detect RpoN1-dependent transcription from the different fliOp promoter versions, it was necessary to substitute the upstream region with the one that is recognized by NifA. From these chimeric promoters, transcription dependent on RpoN1 and NifA was observed under nitrogen fixation conditions. The strongest activity (90% of that promoted by nifUp) was detected for the promoter carrying all of the substitutions (i.e. fliOp-Nif (up-24, up-12, and -11T), whereas only 10% of activity was detected for fliOp-Nif (up-24 and up-12) and fliOp-Nif (-11T), indicating that the identity of the -11-position has a strong influence on the activity promoted by RpoN1, but, in contrast to RpoN2, the nucleotides upstream of the -12- and -24-positions are together equally relevant. Interestingly, all nif promoters carry a T at the -11-position, whereas none of the flagellar promoters has this nucleotide at this position (Fig. 1). Therefore, the inference made from the logo sequences suggesting that RpoN1 recognizes an extended promoter sequence in comparison with that recognized by RpoN2 seems to be confirmed.

The second conclusion obtained from these results is that the flagellar EBP (FleQ/FleT) interacts specifically with RpoN2 to achieve transcription and that the same occurs between NifA and RpoN1. Consequently, the specific interaction between these sigma factors and its cognate activator protein is a second and equally effective mechanism that favors transcriptional specificity. It is interesting that two mechanisms that by themselves would be enough to achieve transcriptional specificity have evolved simultaneously. It is tempting to propose that, given the close functional relationship between the region I of RpoN with both the -11 promoter position and the activator protein, changes in either one of these interactions would affect the other, forcing them to evolve in conjunction.

It was recently reported that the identity of the -11-position in the fork structure is relevant, since, depending on it, either negative or positive interactions will be established with Region I of RpoN, which are then modulated by the EBP to achieve transcriptional control (52). In this regard, it could be hypothesized that the specificity of RpoN1 and RpoN2 is in part accomplished by using these interactions differentially.

Consistent with this idea, the gel mobility shift assays showed a strong preference of RpoN2 to bind the early melted fliOp probe, whereas RpoN1 was able to bind only the nifUp probe. In the absence of Region I, both sigma factors interact less efficiently with all of the probes, but whereas RpoN1I gained the ability to bind the fliOp probe, indicating an inhibitory effect of Region I on the binding of this probe, RpoN2I did not bind to nifUp and bound poorly to fliOp, suggesting that the strongest binding contacts of this sigma factor are made by Region I. These results are in agreement with the hypothesis of a differential interaction of RpoN1 and RpoN2 with the sequence in the fork, which is mediated by Region I.

In a competition experiment using the nifUp probe, RpoN1I was completely displaced by RpoN1, indicating that Region I of this protein, besides inhibiting binding to the early melted fliOp probe, contributes with positive interactions with its specific probe. This property would allow the binding to be favored or destabilized, depending on the identity of the -11-position.

In the case of RpoN2, Region I appears to be only a positive binding element, since RpoN2I did not bind efficiently even to its specific probe. This situation is similar to that reported for ⁵⁴I from K. pneumoniae, which does not bind efficiently to some sequence variants of early melted probes (19, 52).

Although we tested if Region I from any of the two RpoN proteins was capable of binding and discriminating between the fli and nif promoters, none of the polypeptides could bind to these probes by itself nor affect binding of the complete or truncated proteins. However, it remains to be analyzed if the purified Regions I of these two proteins are folded in the proper conformation, since, in contrast to the situation found in K. pneumoniae, these polypeptides were obtained from inclusion bodies.

We further analyzed the importance of the bases at the -11- and -10-positions and of the rest of the promoter in the binding of RpoN1 and RpoN2 by changing the nucleotides of the fork in the fliOp probe by those in the nifUp probe and vice versa. The fact that RpoN2 was able to bind to the nifUp probe with the fli fork (Nif-Fli probe), whereas RpoN2 could not bind nifUp, brings additional support to the idea that binding of RpoN2 strongly depends on interactions with specific nucleotides at the -11-position. In contrast, RpoN1 could bind to the Nif-Fli probe regardless of the presence of the fli -11, -10 fork. This result suggests that RpoN1 is able to bind this probe through its interaction with the promoter bases upstream of the -11-position (in agreement with the results obtained with RpoN1I); however, the absence of the positive interaction with the nif fork and the negative effect exerted by the fli fork results in weak binding as compared with the binding of RpoN1 to the nifUp probe and in the generation of a migration complex similar to the one produced by RpoN1I (compare Figs. 7A and 10A). In contrast, both proteins bind very poorly to the Fli-Nif probe, producing almost no retarded complex (Fig. 10B). This result can be explained by the requirement of RpoN2 for the particular sequence at the fli fork to bind stably, whereas RpoN1 requires making contact with the promoter sequence besides the sequence in the fork to form a stable complex.

We have also shown that each EBP is specific for the activation of a particular DNA-RpoN regulatory center; this differentiation could occur at any of the steps of the activation process. In agreement with this idea, it has been observed that the EBP, PspFHTH, could not activate transcription from the glnHp2 promoter using the ⁵⁴ factor from A. aeolicus, but an EBP from this bacterium was able to do it, suggesting that this ⁵⁴ and its activator have coevolved (15).

The molecular interactions between the EBP and RpoN proteins have been difficult to characterize, since the EBPs do not bind tightly to RpoN; however, in the simplest model, a first interaction between RpoN and the EBP takes place transitorily and independent of NTP hydrolysis. Subsequently, during hydrolysis, the central region of the EBP adopts a different conformation exposing the GAFTGA motif that interacts with Region I of RpoN. This step is central to bringing about an open complex formation. In this context, it will be interesting to determine if Region I is responsible for restricting the interaction to its cognate EBP in one of the steps of the activation process (53-57).

Promoter discrimination mediated by RpoN1 and RpoN2 seems to parallel the situation observed for the ⁷⁰ and ^S factors, since ^S recognizes a particular subset of genes whose promoter sequence is practically identical to that recognized by ⁷⁰. The resemblance of these promoters is such that in vitro a ^S-dependent promoter can be transcribed by ⁷⁰ and vice versa. Recently, it was reported that a C at the -13-position of the promoter sequence contributes importantly to ^S-dependent transcription. However, in the absence of -13 C, selectivity can be generated by other sequence elements such as an A/T-rich region downstream of the -10 box, a distal UP-element combined with a well conserved -35 box, etc. (for a recent review, see Ref. 58). In the case of RpoN1 and RpoN2, promoter discrimination must also have other sequence elements besides the -11-position, since at least two flagellar promoters (motA and fleT) have a C at this position. In these cases, other nucleotides could compensate for the absence of an A at the -11-position.

In conclusion, we propose that the specificity of the RpoN sigma factors of R. sphaeroides is accomplished by two mechanisms: (i) differential recognition of the promoter sequence involving relevant interactions between Region I of RpoN and the fork structure and (ii) the specific interaction of NifA with RpoN1 and of FleT/FleQ with RpoN2.

	FOOTNOTES

 
^* This study was supported by Consejo Nacional de Ciencia y Tecnología Grant 47172/A-1 and Dirección General de Asuntos del Personal Académico, Universidad Nacional Autónoma de México Grant IN222103-3. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. 1.

¹ To whom correspondence should be addressed: Dept. de Biología Molecular y Biotecnología, Instituto de Investigaciones Biomédicas, U.N.A.M. México, D. F., 04510 México. Tel.: 5255-562-23824; Fax: 5255-555-00048; E-mail: rosal{at}servidor.unam.mx.

² The promoter numbering given in this work considers the dinucleotide GC as the -13- and -12-positions, respectively.

³ The abbreviations used are: EBP, enhancer-binding protein; WT, wild type; UAS, upstream activation site.

⁴ S. Poggio, A. Osorio, G. Dreyfus, and L. Camarena, unpublished results.

	ACKNOWLEDGMENTS

 
We thank Teresa Ballado and Francisco Javier de la Mora for technical support.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 

1. Burgess, R. R., and Travers, A. A. (1970) FASEB J. 29, 1164-1169
2. 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]
3. Mittenhuber, G. (2002) J. Mol. Microbiol. Biotechnol. 4, 77-91[Medline] [Order article via Infotrieve]
4. Wosten, M. M. S. M. (1998) FEMS Microbiol. Rev. 22, 127-150[CrossRef][Medline] [Order article via Infotrieve]
5. Helmann, J. D., and Chamberlin, M. J. (1988) Annu. Rev. Biochem. 57, 839-872[CrossRef][Medline] [Order article via Infotrieve]
6. Ishihama, A. (2000) Annu. Rev. Microbiol. 54, 499-518[CrossRef][Medline] [Order article via Infotrieve]
7. Gross, C. A., Lonetto, M., and Losick, R. (1992) in Transcription Regulation (McKnight, S. L., and Yamamoto, K. R., eds) pp. 129-176, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
8. Merrick, M. J. (1993) Mol. Microbiol. 10, 903-909[Medline] [Order article via Infotrieve]
9. Popham, D. L., Szeto, D., Keener, J., and Kustu, S. (1989) Science 243, 629-635[Abstract/Free Full Text]
10. Sasse-Dwight, S., and Gralla, J. D. (1988) Proc. Natl. Acad. Sci. U. S. A. 85, 8934-8938