• DNA binding;
  • phosphorylation;
  • Streptomyces;
  • two-component system SenS–SenR


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

The two-component system SenS–SenR from Streptomyces reticuli has been shown to influence the production of the redox regulator FurS, the mycelium-associated enzyme CpeB, which displays heme-dependent catalase and peroxidase activity as well as heme-independent manganese peroxidase activity, and the extracellular heme-binding protein HbpS. In addition, it was suggested to participate in the sensing of redox changes. In this work, the tagged cytoplasmic domain of SenS (SenSc), as well as the full-length differently tagged SenR, and corresponding mutant proteins carrying specific amino acid exchanges were purified after heterologous expression in Escherichia coli. In vitro, SenSc is autophosphorylated to SenSc∼P at the histidine residue at position 199, transfers the phosphate group to the aspartic acid residue at position 65 in SenR, and acts as a phosphatase for SenR∼P. Bandshift and footprinting assays in combination with competition and mutational analyses revealed that only unphosphorylated SenR binds to specific sites upstream of the furS–cpeB operon. Further specific sites within the regulatory region, common to the oppositely orientated senS and hbpS genes, were recognized by SenR. Upon its phosphorylation, the DNA-binding affinity of this area was enhanced. These data, together with previous in vivo studies using mutants lacking functional senS and senR, indicate that the two-component SenS–SenR system governs the transcription of the furS–cpeB operon, senSsenR and the hbpS gene. Comparative analyses reveal that only the genomes of a few actinobacteria encode two-component systems that are closely related to SenS–SenR.


electrophoretic mobility shift assay


liquid chromatography


response regulator


phosphorylated SenR


cytoplasmic domain of SenS


phosphorylated SenSc


sensor histidine kinase


two-component system

One of the major signal transduction systems governing bacterial responses and adaptation to environmental changes is the two-component system (TCS). A typical TCS consists of an autophosphorylating sensor histidine kinase (SK) and a cognate response regulator (RR) [1]. SKs detect stimuli via an extracellular input domain or intracellular signals via cytoplasmic regions, or use transmembrane regions and sometimes additional short extracellular loops for sensing [2]. In addition to the N-terminal input domain, SKs contain a C-terminal portion representing the transmitter module, with several blocks of amino acid residues being conserved among these kinase types. Phosphorylation within a typical SK usually takes place at a conserved histidine residue; the phosphoryl group of the SK is subsequently transferred to a conserved aspartic acid residue within the receiver domain of the RR. As a result, its C-terminally located output domain has an altered DNA-binding capacity for the regulatory region of target gene(s) or operons [3,4]. The well-studied receiver domain within the nitrogen regulatory protein C − controlling the transcription of genes involved in nitrogen metabolism − has been shown to change its topology upon activation by phosphorylation [5]. Generally, the signaling pathway includes a phosphatase that returns the RR to the nonphosphorylated state. The phosphatase can exist as an individual protein, or reside on a module, which is linked either to the RR or to the kinase. A combination of kinase and phosphatase activity ensures rapid coordination of the cell response [6].

Streptomycetes are Gram-positive and G + C-rich bacteria with a complex developmental life cycle. Germination of spores and subsequent elongation of germ tubes lead to a network of vegetative hyphae. In response to nutritional stress and extracellular signaling, aerial hyphae develop, in which spores mature [7]. As soil-dwelling organisms, streptomycetes need to respond to highly variable conditions. The range of environmental stimuli to which a bacterium can respond is expected to correlate with the number of functional SKs and RRs. These are assumed to have evolved by selection pressure for different ecophysiologic properties of the different strains [8]. The complete genome sequence of Streptomyces coelicolor A3(2) comprises 84 SK genes and 80 RR genes [9]. The physiologic roles of only a few of them have been investigated experimentally. For instance, the AbsA1–AbsA2 system negatively regulates the production of several antibiotics [10,11], and the VanR–VanS system activates the expression of vancomycin resistance [12,13]. Phosphate control of the production of actinorhodin and undecylprodigiosin in S. lividans and S. coelicolor A3(2) is mediated by the two-component PhoR–PhoP system, which also controls the alkaline phosphatase gene (phoA) and other phoA-related genes [14,15]. To date, however, the phosphorylation cascade between a Streptomyces SK and its cognate RR leading to altered DNA-binding affinity of the RR has not been analyzed in detail.

The cellulose degrader S. reticuli has been reported to contain the neighboring genes senS and senR, which encode an SK and an RR, respectively. SenS (42.2 kDa) comprises five predicted membrane-spanning portions. SenR (23.2 kDa) has a C-terminal region with a predicted helix–turn–helix motif, which is characteristic for different DNA-binding proteins [16]. It was concluded that SenR is the cognate RR for the SK SenS. Comparative transcriptional and biochemical studies with a designed S. reticuli senSsenR chromosomal disruption mutant showed that the presence of SenS–SenR influences the transcription of the furS–cpeB operon encoding the redox regulator FurS and the catalase-peroxidase CpeB, and the hbpS gene for the secreted HbpS, representing a novel type of heme-binding protein [16]. Physiologic studies showed that the production of HbpS is positively influenced by hemin in S. reticuli; this correlated with increased hemin resistance. Interestingly, the presence of HbpS leads to enhanced synthesis of the heme-containing CpeB [17].

In this study, we describe the in vitro phosphorylation cascade between the purified cytoplasmic domain of SenS (SenSc) and SenR. Using designed mutant proteins, the phosphorylation sites within SenSc and SenR have been investigated. Bandshift and footprinting analyses have allowed the characterization of the DNA-binding properties in response to phosphorylation by the sensorkinase SenS.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

Cloning of wild-type and mutant senSc and senR genes and purification of fusion proteins

As shown previously, overexpression of the full-length senS gene resulted in the synthesis of an insoluble protein in Escherichia coli[16]. To obtain a truncated SenS (comprising its predicted cytoplasmic portion; see Experimental procedures) with an N-terminal Strep-tag (SenSc), the corresponding portion of senS was cloned into the plasmid pASK-IBA7. Furthermore, using site-directed mutagenesis, a mutant gene was designed and cloned into plasmid pASK-IBA7 (see Experimental procedures), leading to the mutant SenScH199A, which carried an alanine residue in place of the histidine residue in position 199. After induction with anhydrotetracycline, each of the corresponding E. coli XL1-Blue transformants produced a SenSc fusion type in a soluble form within the cytoplasm. Using streptactin affinity chromatography, the SenSc and the SenScH199A fusion protein, both with a predicted molecular mass of 27.1 kDa, were obtained (96 nmol per 1 L of culture) in high purity (Fig. 1). After proteolytic treatment with trypsin, each protein was analyzed by liquid chromatography/mass spectrometry (LC-MS), and was found to comprise the correct N-terminal and internal peptides (data not shown).


Figure 1.  Expression and purification of SenSc and SenR proteins. Soluble protein extracts containing SenSc obtained from E. coli XL1-Blue pASK2 (lane 1) after induction with anhydrotetracycline (lane 2) were loaded onto a streptactin column. After washing (see Experimental procedures), SenSc was eluted with buffer W containing 2.5 mm desthiobiotin (lane 3). SenSCH199A was purified in the same manner (lane 4). To obtain SenR, a cytoplasmic protein extract (lane 5) containing SenR obtained from E. coli BL21(DE3)pLys pETR1 after induction (lane 6) was loaded onto an Ni2+–nitrilotriacetic acid-containing agarose column. Bound SenR was eluted with solution A containing 250 mm imidazole (lane 7) as described under Experimental procedures. SenRD60A (lane 8) and SenRD65A (lane 9) were purified in the same manner. The molecular masses of the protein markers (S) are indicated.

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The full-length senR gene and mutant senR genes (carrying designed codon exchanges) were cloned into the plasmid pET21a. The resulting wild-type protein carrying a C-terminal His-tag (SenR) with a predicted molecular mass of 24.3 kDa was purified to homogeneity from an E. coli BL21(DE3)pLys transformant after induction with isopropyl thio-β-d-galactoside by Ni2+–nitrilotriacetic acid affinity chromatography (Fig. 1). Correspondingly, the mutant SenRD60A and SenRD65A fusion proteins (24.3 kDa), which carried an alanine instead of the original aspartic acid residue at position 60 or 65, were purified to homogeneity from the corresponding E. coli BL21(DE3)pLys transformants by Ni2+–nitrilotriacetic acid affinity chromatography (Fig. 1). Surprisingly, SenRD60A seemed to be partially degraded and aggregated. From 1 L of E. coli culture, about 144 nmol of each SenR type was purified.

SenSc acts as a histidine autokinase in vitro

SenSc exhibited time-dependent autophosphorylation during incubation with [32P]ATP[γP]. The highest signal intensity was already achieved after 5 min of incubation (Fig. 2A). The subsequent addition of an excess of unlabeled ATP resulted in a constant level of phosphorylated SenSc (SenSc∼P) over a relatively long period (at least 20 min; Fig. 2B). Sequence alignments showed that the histidine residue at position 199 within SenS is predicted to be the phosphorylation site [16]. To corroborate this assumption, the corresponding H199 codon was replaced by one for alanine using site-directed mutagenesis (see Experimental procedures). The purified SenScH199A (Fig. 2C, left) failed to undergo autophosphorylation after incubation with [32P]ATP[γP] (Fig. 2C, right). Chemical stability tests were applied to characterize the nature of the phospholigand. Thus, after treatment of SenSc∼P with 1 m HCl, the labeled phosphate group was lost from the protein, but it was retained in the presence of 1 m NaOH (Fig. 2D). This is the characteristic feature of a phosphoamidate, which is stable under alkaline conditions but is sensitive to acidic conditions, under which rapid aminolysis at pH < 5.5 is induced [18]. Taken together, the presented data show clearly that SenS is a histidine autokinase.


Figure 2.  Phosphorylation analysis of SenSc. (A) To test its autokinase activity, the purified SenSc protein (74 pmol) was incubated in kinase buffer containing 0.05 µCi of [32P]ATP[γP] at 30 °C for the indicated period. Each sample was then separated by SDS/PAGE; subsequently, the gel was dried and exposed on an X-ray-sensitive film. (B) After 4 min of self-phosphorylation of SenSc, an excess of unlabeled ATP was added to the samples. Each reaction was terminated by adding an equal amount of 4 × sample buffer. After electrophoresis, the gel was dried and exposed on an X-ray-sensitive film. (C) SenSc (148 pmol) or SenScH199A (148 pmol) was incubated in the kinase buffer with 0.05 µCi of [32P]ATP[γP] for 5 min at 30 °C. After the addition of 4 × sample buffer, the reaction was stopped, and the mixture was subsequently subjected to SDS/PAGE. The gel was stained with Coomassie Brilliant Blue (left), or alternatively dried and exposed on an X-ray-sensitive film (right). (D) After autophosphorylation of 74 pmol of SenSc with 0.05 µCi of [32P]ATP[γP] in kinase buffer for 5 min at 30 °C, the reaction was terminated by adding 4 × sample buffer and subjected to SDS/PAGE. Each gel was treated with the indicated solutions, dried, and exposed on an X-ray-sensitive film.

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SenSc phosphorylates and dephosphorylates SenR

As SenR was predicted to be the cognate RR of the SK SenS, the transfer of radiolabeled phosphate from SenSc to SenR was investigated. For this purpose, the purified SenR was added to the 32P-autophosphorylated SenSc (see previous section). Very rapid (within 5–10 s) labeling of SenR was observed, together with a concomitant reduction of the phospholabel within SenSc(Fig. 3A,B). Autophosphorylation activity of SenR using [32P]ATP[γP] or the phosphodonor acetylphosphate could not be detected (data not shown). The deduced SenR comprises aspartic acid residues at position 60 (D60) and position 65 (D65), each of which is a candidate to participate in the phosphorylation process [16]. Site-directed mutagenesis showed that each of the two residues was replaced by an alanine. SenRD60A and SenRD65A were subsequently purified from corresponding E. coli transformants (see above). Further transphosphorylation analysis revealed that the presence of SenSc∼P provoked phospholabeling of wild-type SenR and SenRD60A. In contrast, the mutant protein SenRD65A was not found to be phosphorylated by SenSc∼P (Fig. 3C). D65 is therefore the phosphorylation site within SenR.


Figure 3.  Phosphotransfer from SenSc to SenR, SenRD60A or SenRD65A. (A, B) Purified SenSc (184 pmol) was incubated with 0.05 µCi of [32P]ATP[γP] for self-phosphorylation. After 4 min, equal amounts of purified SenR were added and incubated for the indicated period at 30 °C. The reactions were terminated by adding 4 × sample buffer. After SDS/PAGE, the gel was dried and exposed on an X-ray-sensitive film (A) or quantified by detection of the radioactivity emitted by SenR∼P (▪) or SenSc∼P (◆) using a PhosphorImager (B).  (C) The wild-type SenR or SenR mutant proteins (SenRD60A or SenRD65A), in each case 330 pmol of protein, were mixed with 260 pmol of SenSc∼P in transphosphorylation buffer for 1 min at 30 °C. Reactions were terminated with 4 × sample buffer, subjected to SDS/PAGE, and stained with Coomassie Brilliant Blue (left), or alternatively the gel was dried and exposed on an X-ray-sensitive film (right).

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As demonstrated by quantitative analysis (using a PhosphorImager system), during the transphosphorylation reaction dephosphorylation of phosphorylated SenR (SenR∼P) occurred after aproximately 3 min of incubation (Fig. 3B); during this period, no rephosphorylation of SenSc was recorded. To investigate this process in more detail, phospholabeled SenR (carrying a His-tag) was separated immediately after phosphorylation from SenSc (carrying a Strep-tag) by Ni2+–nitrilotriacetic acid affinity chromatography. The addition of dephosphorylated SenSc to a reaction mixture containing phospholabeled SenR provoked a rapid (within 60 s) loss of the phosphoryl group from SenR (Fig. 4A,B). In the absence of SenSc, autodephosphorylation of SenR∼P occurred only after a longer (> 120 s) period of incubation (data not shown). These data show that SenSc also acts as a phosphatase for SenR∼P.


Figure 4.  Dephosphorylation rate of SenR∼P. (A) SenR was first phosphorylated by SenSc∼P in a transphosphorylation reaction, and subsequently separated from it by Ni2+–nitrilotriacetic acid affinity chromatography. Purified SenR∼P (∼ 82 pmol) was incubated at 30 °C alone (top) or with (bottom) 148 pmol of dephosphorylated SenS for the indicated times. Each reaction was stopped by adding an equal amount of 4 × sample buffer, and the products were analyzed by SDS/PAGE. Gels were dried and exposed on an X-ray-sensitive film. (B) Dried gels were further analyzed using a PhosphorImager. The diagram shows the quantified results representing the measured radioactivity at the indicated times (▪) with SenR∼P alone or for the mixture (◆) of SenR∼P and SenSc.

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DNA-binding properties of SenR depend on its phosphorylated state

Comparative analysis of wild-type S. reticuli and the senSsenR disruption mutant showed that the presence of SenS–SenR correlates with a significant reduction of transcripts (furS–cpeB and hbpS) and the corresponding proteins [16]. For further analyses, different DNA fragments (Fig. 5A) corresponding to the upstream region (310 bp, named up-furS1) of the furS–cpeB operon or the upstream region (548 bp, named up-hbpS1) located between hbpS and senS were amplified by PCR. Electrophoretic mobility shift assays (EMSAs) were performed with labeled DNA (5200 pmol of up-furS1 or 2900 pmol of up-hbpS1) and increasing quantities (0–16 pmol) of the purified SenR or SenR∼P. Interestingly, in contrast to SenR∼P, SenR interacted with up-furS1 (Fig. 5B). The addition of 12 pmol of SenR to the reaction mixture led to an ∼ 84% decrease of free up-furS1, whereas the same amount of SenR∼P provoked only a ∼ 10% reduction (Fig. 5D). The presence of small quantities (4 and 8 pmol) of SenR led to one type of retarded DNA species (Fig. 5B, arrow b); an additional one was formed if the protein concentration (12 and 16 pmol) was increased (Fig. 5B, arrow a). These data suggested the presence of multiple SenR-binding sites. The specificity of this interaction was verified by competition using constant amounts of SenR and additional increasing amounts of unlabeled up-furS1 (Fig. 5B, third box from left). Furthermore, SenR was not found to interact with the upstream region (up-cpeB) of the cpeB gene (Fig. 5B, fourth box from left).


Figure 5.  Gene organization and EMSAs with isolated SenR proteins. (A) The gene organization of furS–cpeB, hbpS, senS and senR is indicated. The labeled DNA regions are marked in gray. (B, C) The upstream region of the furS–cpeB operon (5200 pmol of up-furS1) (B) or the intergenic region between hbpS and senS (2900 pmol of up-hbpS1) (C) was incubated without or with increasing amounts (0, 4, 8, 12 or 16 pmol; black triangle) of SenR or SenR∼P in incubation buffer (see Experimental procedures). For competition experiments, labeled up-furS1 (5200 pmol) was incubated with constant amounts (16 pmol) of SenR and increasing amounts of unlabeled up-furS1 (0, 5200, 7800, 10 400 or 13 000 pmol; open triangle) (B, third box from left). In the same manner, unlabeled up-hbpS1 (0, 2900, 4350, 5800 or 7250 pmol; open triangle) was added to the mixture comprising labeled up-hbpS1 (2900 pmol) and constant amounts (16 pmol) of SenR (C, third box from left). For further corroboration of the binding specificity, SenR (0–16 pmol; black triangle) was incubated with the upstream region of cpeB (up-cpeB, 5500 pmol) (B, fourth box from left). After incubation at 30 °C for 15 min, the mixtures were separated on 5% polyacrylamide gels, and then subjected to autoradiography. The retarded DNA fragments are indicated (a, b, c and d). The control DNA in mixtures without SenR is everywhere marked as lane 0. (D) In addition, gels were dried and analyzed by a PhosphorImager System. The radioactivity level of the DNA probe alone was set at 100%. The reaction products up-furS1 + SenR (▪), up-furS1 + SenR∼P (□), up-hbpS1 + SenR (◆), and up-hbpS1 + SenR∼P (◊) as well as the quantities of SenR used are indicated.

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EMSAs (also known as bandshift assays) with up-hbpS1 and varying amounts (0–16 pmol) of SenR or SenR∼P showed that the DNA-binding affinity was enhanced after phosphorylation (Fig. 5C). This was indicated by the observation that SenR (4 pmol) led to a ∼ 63% decrease in free up-hbpS1, whereas the same amount of SenR∼P (4 pmol) enhanced it to ∼ 94% (Fig. 5D). Interestingly, SenR induced the formation of two shifted species (Fig. 5C, arrows c and d), suggesting the existence of at least two binding sites within up-hbpS1. One of them (marked as d) was only observed in the presence of small quantities (4 pmol) of SenR but not with SenR∼P. Further EMSAs using different quantities of proteins showed that, to obtain a 50% decrease in free up-hbpS1, at least three times as much SenR as SenR∼P was required (Table 1). Competition studies using constant amounts of SenR and increasing amounts of unlabeled up-hbpS verified the specificity of the SenR–up-hbpS1 interaction (Fig. 5C, third box from left). Taken together, these data revealed that SenR binds specifically to up-furS1 and up-hbpS1, and the phosphorylation of SenR by SenS∼P substantially alters its DNA-binding characteristics.

Table 1.   Relative binding affinity of wild-type and mutated SenR proteins for 32P-labeled DNA-fragments. EMSAs were done (as described in Experimental procedures) using increasing (0–100 pmol) amounts of the mentioned proteins and analysis was done with a PhosphorImager. The indicated amount (in pmol) of each protein is required to obtain a 50% decrease of the intensity of free DNA (up-furS1 or up-hbpS1). For this purpose, the radioactivity level of the sample without protein was set at 100%. The experiments were repeated four times; the obtained data were reproducible.
 Dephosphorylated proteinPhosphorylated protein
up-furS16.2> 4111.134.6> 41

Further bandshift assays using different amounts of purified SenR proteins demonstrated that each of the SenR mutant proteins (SenRD60A and SenRD65A) has reduced binding affinity for up-furS1 and up-hbpS1 (Table 1).

Identification of the SenR-binding sites

To identify the exact DNA-binding site(s) within up-furS1 and up-hbpS1, DNaseI footprinting experiments with the purified RRs SenR and SenR∼P, after their phosphorylation in the presence of ATP by SenSc, were performed. Footprinting experiments with radioactively labeled up-furS1 showed that SenR protected a region spanning 9 bp (I, AACTTGGGG) against DNaseI cleavage (Fig. 6A, left). In addition, a short region (marked by a white block) upstream of region I was protected, implicating probable binding sites (as observed by bandshift experiments), or a change in DNA topology being induced after interaction with SenR. Increasing amounts of SenR neither extended nor altered the extent of the protection. SenR∼P had no affinity for this DNA region, even at high concentrations (up to 60 pmol) (Fig. 6A, right). A truncated up-furS1 fragment (∼ 100 bp, named up-furS2) comprising site I (I, Fig. 7A) still interacted with SenR, as shown by bandshift assays (Fig. 7B). Studies with this fragment having a deleted site I (ΔI) (Fig. 7A) showed that it was targeted neither by SenR nor by SenR∼P (Fig. 7B). The specificity of the SenR–up-furS2 interaction was further corroborated by competition using constant amounts of SenR and increasing amounts of unlabeled up-furS2ΔI (Fig. 7D).


Figure 6.  Footprinting studies. (A) up-furS1 (6900 pmol) and (B) up-hbpS1 (5800 pmol) were incubated without SenR or SenR∼P, or with increasing amounts (20.5, 41 and 61.5 pmol) of SenR or SenR∼P in 10 mm Tris/HCl (pH 7.9), 5 µg·mL−1 sonicated salmon sperm DNA, 5% glycerol, 40 mm KCl, 2 mm MgCl2 and 2 mm dithiothreitol. After treatment with DNaseI, analyses were performed with 6% polyacrylamide-urea gels, and autoradiography. The protected DNA regions (I, II and III) are indicated by black blocks. The additional protected region within up-furS1 is indicated by a small, open rectangle.

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Figure 7.  EMSAs with mutated DNA regions up-furS and up-hbpS. (A) Portions of the DNA fragments (up-furS2 or up-hbpS2, see below) containing the complete (I, II, III and II + III; underlined) or deleted (ΔI, ΔII, ΔIII and ΔII + ΔIII; dotted lines) binding motifs, or the complete (PIII, marked by >>> <<<) or deleted (ΔPIII, dotted lines) perfect inverted repeat overlapping the binding site III are shown. (B) The ∼ 100 bp upstream region of the furS–cpeB operon (up-furS2) (14 200 pmol) and the corresponding mutated region (up-furS2ΔI) (14 200 pmol) were incubated without or with increasing amounts (8, 16 and 24 pmol; black triangle) of SenR (left) or SenR∼P (right) in incubation buffer at 30 °C for 15 min. (C) The intergenic region (∼ 100 bp) between hbpS and senS (up-hbpS2) (9400 pmol) and the mutated counterparts (up-hbpS2ΔII; ΔIII; ΔPIII; ΔII + ΔIII) (9400 pmol) were incubated without or with increasing amounts (8, 16 and 24 pmol; black triangle) of SenR (top) or SenR∼P (bottom), in incubation buffer. (D) For competition experiments, labeled up-furS2 (14 200 pmol) was incubated with constant amounts (24 pmol) of SenR and increasing amounts of unlabeled up-furS2ΔI (0, 14200, 21300 or 28400 pmol; open triangle) (left). In the same manner, unlabeled up-hbpS2ΔII + ΔIII (0, 9400, 14 100 or 18 800 pmol; open triangle) was added to the mixture comprising labeled up-hbpS2 (9400 pmol) and constant amounts (24 pmol) of SenR (right). The control DNA in the mixture without SenR is (B, C, D) marked as lane 0. The analyses were performed with 5% polyacrylamide gels, and autoradiography.

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DNaseI protection assays using the amplified intergenic region between hbpS and senS (up-hbpS1) revealed two SenR-binding sites (II and III) spanning 21 bp (II: ACCTCCAGTAGAGCCTGGGCT) and 19 bp (III: GGACCGGGCCGCGTCCCGT) (Fig. 6B). Site II is located near to the start codon of hbpS, and site III is relatively distant from it. After incubation with SenR∼P, the ends of both sites became hypersensitive to DNaseI treatment. This was accompanied by an apparent extension of region II as well as of region III (Fig. 6B). Further bandshift assays showed that the shortened up-hbpS1 fragment (∼ 100 bp, named up-hbpS2) containing both motifs (II + III, Fig. 7A) was still targeted by SenR and SenR∼P (Fig. 7C). The up-hbpS2 fragment lacking the perfect inverted repeat (ΔPIII; Fig. 7A) interacts only slightly with SenR∼P (Fig. 7C). Analyses with fragments with either site II (ΔIII or site III (ΔIII) or both (ΔII + ΔIII) deleted revealed that both binding sites are required for interaction with SenR, independent of its phosphorylation status (Fig. 7C). The specificity of the SenR–up-hbpS2 interaction was further corroborated by competition using constant amounts of SenR and increasing amounts of unlabeled up-hbpS2ΔII + ΔIII (Fig. 7D).

Taken together, these data confirm the specificity of the SenR-binding sites and corroborate the assumption that phosphorylation of SenR by SenS∼P alters its DNA-binding characteristics.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

The designed cloning procedures allowed us to obtain the cytoplasmic domain of SenS carrying a Strep-tag (SenSc) and the full-length SenR protein with a His-tag (SenR) at high purity as a basis for in vitro studies. SenSc was found to function as an efficient autokinase. Chemical stability assays revealed that the ligand within the phosphorylated SenSc (SenSc∼P) must be a phosphoamidate, which is extremely acid-labile but relatively base-stable. This feature discriminates all phosphorylated amino acid residues (except arginine) from phosphoamidates [18]. Additional mutational investigations demonstrated that SenSc requires the histidine residue at position 199 for autokinase activity. The in vitro transfer of the phosphate group from SenSc to SenR (dephosphorylated SenR) occurred very rapidly, but did not occur in a designed mutant SenR protein carrying a substitution of the aspartic acid residue at position 65 (D65). The kinase SenSc was found to act additionally as a phosphatase for SenR∼P (phosphorylated SenR).

Bandshifts revealed that SenR, but not SenR∼P, binds specifically to a region (site I) upstream of the furS–cpeB operon encoding the redox regulator FurS and the catalase-peroxidase CpeB. The deletion of site I abolishes the interaction with SenR. Interestingly, this site is located within the previously determined FurS operator [19] and overlaps with its central region. The data imply that, in addition to FurS, SenR participates in regulating the transcription of the furS–cpeB operon. This conclusion is supported by the earlier finding that the absence of a functional furS or senR gene provokes enhanced transcription of the furS–cpeB operon [16,20]. Overlapping DNA-binding sites have also been described for other known regulators. Depending on the physiologic condition, either the activator NhaR or the RR RcsB from E. coli interacts with overlapping motifs within the upstream region of osmC. This gene encodes a predicted envelope protein that is required for resistance to organic peroxides and also for long-term survival in the stationary phase [21,22]. The regulator PutR and the activator CRP from Vibrio vulnificus bind simultaneously to overlapping sites but probably to opposite faces. This process leads to activation of the transcription of the operon encoding a proline dehydrogenase and a proline permease [23]. PutR has been suggested to facilitate the DNA binding of CRP by direct protein–protein interaction or to induce a change in DNA topology that allows more efficient recruitment of CRP.

Comparative transcriptional and biochemical studies have revealed that SenS–SenR modulates the transcription of the furS–cpeB operon as well as the hbpS gene encoding a novel heme-binding protein. Interestingly, SenR has a high affinity for the intergenic region between hbpS and senS, spanning 21 bp (site II) and 19 bp (site III). Both became hypersensitive to DNaseI treatment at their ends after incubation with SenR∼P, indicating altered DNA topology. The phosphorylated form of RRs has been shown to provoke oligomerization and to bind cooperatively to target DNA sequences [24,25]. Altered DNA binding upon phosphorylation was observed, for example, for the RR RegR of the RegS–RegR system from Bradyrhizobium japonicum, controlling the expression of numerous genes, the products of which are either directly involved in nitrogen fixation or in functions associated with the microaerobic lifestyle of this symbiont [26]. A corresponding observation was also made for MisR of the TCS MisR–MisS from Neisseria meningitides, which is required for its pathogenicity [24], and for NtrC of the NtrB–NtrC system, which controls the expression of genes involved in nitrogen metabolism in Rhodobacter capsulatus[27].

The two SenR-binding sites II and III share a common motif CNTCCNGT in the same orientation. Additionally, binding site III is localized within a region (CGGCCCGGACCGGGCCG) representing a perfect inverted repeat (Fig. 7). The use of DNA fragments lacking either binding site II, binding site III or both showed that each of them is necessary for specific targeting by SenR. Further single replacements within each binding site (I, II or III) may reveal the essential role of single nucleotides in the specific interaction with SenR. The position of the SenR operator (sites II and III) indicates that the transcription of senSsenR is autoregulated. As reported previously [16], SenS–SenR shows similarity to the ChrS–ChrA system from Corynebacterium diphtheriae. ChrA has so far not been purified, but it has been predicted to modulate the transcription of the heme oxygenase gene (hmuO) [28,29]. On the basis of our data, we identified a DNA region upstream of hmuO with high similarity to SenR-binding site II and an additional shared sequence (GGGCGTCGG) near to its 3′-end (data not shown). This is in accordance with the fact that the helix–turn–helix DNA-binding domains of SenR and ChrA share 61% amino acid identity (data not shown).

The designed SenRD60 and SenRD65A proteins showed reduced DNA-binding affinity for up-furS as well as for up-hbpS. D60 and D65, together with other aspartic acid residues (in positions 19 and 20), in SenR correspond to those that have been predicted to form an acidic pocket within RRs containing a CheY-like receiver domain [30]. Mutations at any of the acidic pocket aspartates result in loss of functionality [31]. SDS/PAGE analysis of purified SenR proteins revealed that SenRD60A appeared to be partially degraded and aggregated (Fig. 1). Both SenRD60A and SenRD65A seem to be perturbed in their conformation and hence show altered DNA-binding abilities.

Our previous data revealed that the presence of SenS–SenR considerably enhances the resistance of S. reticuli to hemin or the redox cycling compound plumbagin, suggesting its relevance in sensing of redox changes [16]. Further preliminary comparative analysis (data not shown) revealed that under different redox-stress conditions, the presence of SenS–SenR is required for the production of additional extracellular proteins, whose characteristics remain to be clarified. One key part of the sensing processes is expected to be orchestrated by the heme-binding protein HbpS [17]. How it participates in delivering signals to SenS will be explored.

Sequence comparisons showed that the relative organization of senS and senR is identical to those of homologous genes within the Scoelicolor A3(2) genome [32]; these genes are also preceded by an uncharacterized gene that is closely related to hbpS[16]. Further sequence alignments revealed the presence of other predicted TCSs showing high amino acid identity with SenS–SenR within Rhodococcus sp. RHA1 [33] and Arthrobacter aurescens TC1 [34]. Interestingly, each of the corresponding homologous genes is also preceded by a close homolog of hbpS, the organization of which is identical to that within the S. reticuli genome. The fact that each corresponding intergenic region comprises motifs that are related to the SenR-binding sites (II and III) (data not shown) indicates that these homologous systems are also autoregulated. According to these findings, it could be assumed that HbpS and SenS–SenR, and probably also the corresponding homologs from the other mentioned actinobacteria, interact together to mediate an appropriate response to environmental changes. Such a mode of interaction has been recently postulated for the lipoprotein LpqB and the TCS MtrA–MtrB, which together might form an actinobacterial three-component system [35]. The elucidation of the exact role of accessory proteins for the modulation of bacterial TCSs might give new insights into the complex network of signaling processes.

Taking the presented and previous data into account, the TCS SenS–SenR from S. reticuli is a model that is well suited to elucidate the role of other related TCSs from different actinobacteria.

Experimental procedures

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

Bacterial strains, plasmids, media and culture conditions

The plasmid pUC18 [36] was a gift from J Messing (State University of New Jersey, Piscataway, NJ, USA). The E. coli strains DH5α[37], BL21(DE3)pLys (Novagen, Darmstadt, Germany) and XL1-Blue [38], and the plasmids pET21a (Novagen) and pASK-IBA7 (IBA, Göttingen, Germany), were used for routine cloning purposes. The constructs pUKS10 (pUC18 derivative) and pWKB1 (pWHM3 derivative), which contain the furS–cpeB operon, have been described previously [16,20]. E. coli strains were grown in LB medium at 37 °C [36].

Chemicals and enzymes

Chemicals for SDS/PAGE were obtained from Serva (Heidelberg, Germany). Molecular weight markers were supplied by Sigma (Steinheim, Germany). Restriction enzymes, T4 ligase, T4 polynucleotide kinase, DNaseI and Pfu DNA polymerase for PCR were obtained from New England Biolabs (Frankfurt am Main, Germany), Roche (Mannheim, Germany), or Promega (Mannheim, Germany).

Isolation of DNA and transformations

Plasmids were isolated from E. coli with the aid of a mini plasmid kit (Qiagen, Hilden, Germany). E. coli DH5α and XL1-Blue were transformed with plasmid DNA by electroporation [39], whereas BL21(DE3)pLys was transformed with the CaCl2 method as previously described [36].

PCR, DNA sequencing and computer analysis

PCR was performed with Pfu DNA polymerase. To test the correctness of cloned genes, sequencing was done using the Ready Reaction mix and ABI PRISM equipment (PE Biosystems, Foster City, CA, USA) by the departmental sequence service (U Coja, FB Biologie, University of Osnabrueck). For DNaseI footprinting studies, sequencing was done using the AutoRead Sequencing Kit (Amersham Biosciences, Freiburg, Germany); however, radioactively labeled primers corresponded to those utilized for the EMSAs (see below). Sequence entry, primary analysis and ORF searches were performed using clone manager 5.0. Database searches using the PAM120 scoring matrix were carried out with blast algorithms (blastx, blastp and tblastn) on the NCBI file server [40]. Multiple sequence alignments were generated by means of the clustalw (1.74) program [41].

Site-directed mutagenesis within senS and senR

A point mutation in the senS gene on plasmid pQS1 [16] was introduced using the QuikChange Site-Directed Mutagenesis kit (Stratagene, Amsterdam, The Netherlands) with the following specific primers: H199A1, 5′-CCCGGGAGATCGCCGACACCCTCGC-3′; and H199A2, 5′-GCGAGGGTGTCGGCGATCTCCCGGG-3′. These oligonucleotides were designed to replace the selected histidine codon at position 199 (H199) with an alanine codon (underlined). The constructs were transformed into Ecoli XL1-Blue. Subsequently, each of the cloned inserts in the plasmid DNA, isolated from several individual transformants, was analyzed with restriction enzymes and by sequencing. The resulting correct plasmid was named pQS1H199A.

For introducing specific mutations in the senR gene, a different strategy was used. First, the subconstruct pUR1 was created by ligation of the longer SphI–BamHI fragment of pUC18 with the 1.8 kb SphI–BamHI fragment (containing senR) of pWKB1. The plasmid pUR1 was then used for PCRs and further ligation. The desired PCR products were obtained using the forward primer R1NSph (5′-CAGCGCATGCTGCTCCAGGCAGCCGAC-3′), and one of the reverse primers R1D65Pst (5′-CGAGCTGCAGGGCCATCAGGACGACGTC-3′) or R1D60Pst (5′-CGAGCTGCAGGTCCATCAGGACGACGGCCGGGGCGGTCTTGC-3′). The PstI restriction site is in bold type within R1D60Pst and R1D65Pst, and the codon for alanine replacing the original codon (GTC) for aspartic acid is underlined; the SphI restriction site within R1NSph is in bold type. The cleaved PCR product resulted in a 0.76 kb DNA fragment containing a part of the mutagenized senR gene (D60A or D65A). Each of these was then ligated with the large SphI–PstI fragment (3.75 kb) of plasmid pUR1. The resulting constructs (isolated from E. coli DH5α transformants) were named pUR1D60A or pUR1D65A, respectively. The correctness of the in-frame replacement was controlled by restriction and sequencing.

Cloning of genes in E. coli

The DNA region (from the nucleotide at position 505 to that at position 1197 of senS) encoding the cytoplasmic part (comprising the segment from the aspartic acid at position 169 to the arginine at position 398) of SenS was amplified by PCR with the following primers: SenS3, 5′-CTAGAATTCGACGACCTGGTC-3′, harboring an EcoRI restriction site (in bold type), and SenS4, 5′-GATCTGCAGTCATCTCGGCTC-3′, containing the PstI restriction site (in bold type). The plasmids pWKB1 [16] and pQS1H199A were used as DNA templates. Each PCR product was digested with EcoRI and PstI and ligated with EcoRI–PstI-digested pASK-IBA7. The resulting construct pASKS2 or pASKS2H199A was transformed into E. coli XL1-Blue, recovered, and sequenced. Transformants containing the correct constructs and producing the designed fusion protein with Strep-tag codons (SenSc or SenScH199A) were selected.

The senR-coding region of plasmid pWKB1, pUR1D60A or pUR1D65A was amplified by PCR with following primers: SenR1, 5′-CCCATATGACCCCCACCCCGCAGCCGCC-3′, consisting of an NdeI restriction site (in bold type), followed by the sequence encoding the N-terminal amino acids of SenR, and SenR2, 5′-CGCGCTCGAGAGACAGGAGGCGTTGTTC-3′, determining the C-terminal amino acids of SenR, followed by the XhoI restriction site (in bold type). The PCR products were digested with NdeI and XhoI, ligated with the NdeI–XhoI-cleaved pET21a, and subsequently transformed into E. coli XL1-Blue. Having sequenced several of the resulting plasmids pETR1, pETR1D60A or pETR1D65A, we verified the correctness of the senR gene and the in-frame fusion with the His-tag codons.

Purification of the fusion proteins

An E. coli XL1-Blue transformant containing the pASKS2 or pASKS2H199A plasmid was inoculated in LB medium with ampicillin (100 µg·mL−1) and cultivated at 37 °C. The synthesis of the SenSc or the SenScH199A fusion protein was induced (at a D600 of 0.6) by adding anhydrotetracycline (200 ng·mL−1). The cells were grown for 4 h, harvested, washed with buffer W (100 mm Tris/HCl, pH 8.0, 150 mm NaCl, 1 mm EDTA), and disrupted in the same buffer W by ultrasonication (10 × 10 s, with 10 s intervals) using a Branson sonifier (Danbury, CT, USA). The fusion protein was purified directly from a cleared cell lysate using streptactin affinity chromatography according to the instructions of the manufacturer (IBA).

Plasmid pETR1, pETR1D60A or pETR1D65A was transformed into E. coli BL21(DE3)pLys. A selected transformant containing the desired plasmid was inoculated in LB medium containing ampicillin (100 µg·mL−1) and chloramphenicol (34 µg·mL−1) at 37 °C. The synthesis of the protein was induced (at a D600 of 0.6) by adding 1 mm isopropyl thio-β-d-galactoside for 4 h. The collected cells were washed and resuspended in chilled buffer A (10 mm Hepes, 60 mm KCl, pH 8.0). The extracts were subsequently sonicated and clarified by centrifugation at 16 000 g for 30 min (Avanti J-25 centrifuge; Beckman Coulter, Palo Alto, CA, USA). The supernatant containing soluble proteins and additional 25 mm imidazole was mixed with Ni2+–nitrilotriacetic acid agarose (Qiagen) on a wheel at 4 °C for up to 3 h. The agarose column containing bound proteins was washed with solution A supplemented with 25 mm imidazole. His-tagged proteins were released with the same solution in the presence of 250 mm imidazole. The protein concentration of the samples was determined by a previously established method [42]. The quality of the proteins was analyzed by electrophoresis in 12.5% SDS/PAGE gels according to the method of Laemmli [43].


The N-terminal amino acids and internal peptides within SenSc were determined by LC-MS (Bruker Daltonics, Karlsruhe, Germany) analysis. For this purpose, the purified protein was first separated by SDS/PAGE. After staining (with Coomassie Brilliant Blue), the SenSc-containing band was excised. The gel piece was subsequently cleaned, dehydrated, dried, and digested with trypsin (modified, sequencing grade from Roche) following standard procedures [44,45].

In vitro phosphorylation assays

For the autokinase reaction, the SenSc protein was incubated in phosphorylation buffer (P-buffer), which contained 50 mm Tris/HCl (pH 7.5), 10% glycerol, 20 mm MgCl2, 1 m NaCl, 2 mm dithiothreitol, 1 mm ATP, and 0.05 µCi of [32P]ATP[γP] (Amersham Biosciences). The reaction mixture was incubated at 30 °C for different time intervals, and the reaction was terminated by the addition of 4 × SDS/PAGE loading buffer. To test the transphosphorylation reaction, autophosphorylated SenSc (SenSc∼P) was incubated with equimolar amounts of SenR-His-tag (SenR) without labeled ATP in P-buffer. The reaction mixture was incubated at 30 °C for different time intervals. Samples were resolved by 12.5% SDS/PAGE. After drying of the gels, products were visualized by autoradiography and quantified by a PhosphorImager with imagequant 5.0 software. Alternatively, the phosphorylation of SenR by acetylphosphate was tested. Therefore, 32P-labeled acetylphosphate was generated by incubation of acetate kinase with acetate (60 mm KOAc) and [32P]ATP[γP] in 25 mm Tris/HCl (pH 7.5) and 10 mm MgCl2 for 30 min at 25 °C or as a control without acetate kinase. SenR and P-buffer were added, and the mixture was incubated at 30 °C for different time intervals. Samples were resolved by SDS/PAGE, and after drying, the gels were analyzed by autoradiography.

In vitro dephosphorylation assays

Purified SenR was first transphosphorylated by SenSc to SenR∼P as described above. Then, the complete kinase reaction mixture was loaded onto an Ni2+–nitrilotriacetic acid agarose column and subjected to affinity chromatography. Thus, His-tagged SenR∼P was separated from SenSc with a Strep-tag. The freshly purified SenR∼P was incubated alone or with equal amounts of SenSc in the dephosphorylation buffer (50 mm Tris/HCl, pH 7.5, 10% glycerol, 100 mm MgCl2, 2 mm dithiothreitol and 50 mm KCl) at 30 °C for different time intervals. Proteins within each sample were separated by 12.5% SDS/PAGE. After drying of the gels, labeled proteins were visualized by autoradiography, and quantified by a PhosphorImager with imagequant 5.0 software.

Chemical stability assays

To detect a phosphohistidine [46], 32P-labeled SenSc (three individual samples) was fractionated by 12.5% SDS/PAGE. One sample was then treated for 30 min at room temperature with 1 m NaOH, the second with 1 m Tris/HCl (pH 7.5), and the third with 1 m HCl. Subsequently, the gels were dried and exposed to an X-ray film.

DNA and oligonucleotide labeling

PCR fragments and oligonucleotides were labeled at the 5′-end, using T4 polynucleotide kinase and [32P]ATP[γP].


The upstream region of the furS–cpeB operon was amplified from plasmid pWKB1 using primers F1 (5′-GTCCGGGGCCACATGATGCG-3′) and F2c (5′-GCGACGCGAGCTGCCGTCACG-3′). The intergenic region between hbpS and senS (cloned within the plasmid pWKB1) was obtained by PCR with primers S1 (5′-CGACGACACCGGCACCGA-3′) and S2c (5′-CGGGGCCAGGACGACGAGCA-3′). Each of the fragments was end-labeled with [32P]ATP[γP] using T4 polynucleotide kinase. An aliquot of the labeled fragment was incubated in DNA-binding buffer (10 mm Tris/HCl, pH 7.9, 5 µg·mL−1 sonicated salmon sperm DNA, 5% glycerol, 40 mm KCl, 1 mm MgCl2 and 2 mm dithiothreitol) with varying quantities of SenR or SenR∼P at 30 °C for 15 min. Bandshifts were analyzed by subsequent electrophoresis on a 5% polyacrylamide gel. Gels were run at 100 V for 3 h, and dried, and products were visualized by autoradiography and quantified by a PhosphorImager with imagequant 5.0 software.

To determine whether SenR had already been in a phosphorylated form within E. coli, the obtained protein was incubated with SenSc in dephosphorylation buffer and separated by Ni2+–nitrilotriacetic acid affinity chromatography. Subsequently, EMSAs were performed; these revealed that both (treated and untreated with SenSc) SenR types possess identical binding activities (data not shown).

DNaseI footprinting studies

Using the labeled primers F1 and F2c, or S1 and S2c, and the plasmid pUKS10, labeled fragments corresponding to the upstream region of furS–cpeB or the intergenic region between hbpS and senS were generated by PCR. Each DNA sample was incubated for 15 min at 30 °C in buffer (as used for bandshift assays, but containing 2 mm MgCl2) with varying quantities of SenR or SenR∼P. Each sample was then treated with 20 mU of DNaseI at room temperature for 40–120 s, and the reaction was terminated by adding EDTA (final concentration 5 mm). The product was precipitated with 2.5 volumes of ethanol in the presence of sodium acetate (final concentration 300 mm), and subsequently washed with 75% ethanol. The pellet was suspended in 1 µL of H2O and 4 µL of formamide-containing sequencing buffer, and applied to a 6% polyacrylamide/urea gel.

Mutagenesis of SenR-binding sites

The identified (by footprinting) SenR-binding sites (I, II and III) or the region representing a perfect inverted repeat overlapping binding site III (PIII) (Fig. 7A) were deleted by insertion of a restriction site (EcoRI) using overlapping primers (Table 2) and the pUKS10 plasmid [19] as template for PCR. For this purpose, the different generated PCR fragments (A–I) were first amplified and then cleaved with EcoRI. Restricted fragments A and B (to delete site I), C and D (to delete site II), E and F (to delete site III), G and H (to delete the perfect inverted repeat) and I and F (to delete simultaneously sites II and III) were ligated to each other. The resulting DNA fragments (∼ 450 bp) were cleaved with HindIII and PstI, and ligated with the longer HindIII–PstI fragment of the plasmid pUC18. Each of the resulting constructs was transformed in E. coli DH5α by electroporation. The correctness of the expected mutations (Fig. 7A) was confirmed by restriction analysis and sequencing.

Table 2.   List of primers used to obtain mutated SenR-binding sites.
FragmentPrimer name Primer sequence (5′- to 3′; restriction sites in bold type)
AIHinforCATGAAGCTTGCATGGCCGGGGCC (located upstream of furS)
IEcorevCGAGAATTCGAAAACGAACGGTGC (located upstream of furS and ending at the 5′-end of binding site I)
BIEcoforGTTGAATTCTCGTGTTTATGAGGG (located upstream of furS and beginning at the 3′-end of binding site I)
IPstrevGGAGCTGCAGCCCACGCGATCGCG (located downstream of the start codon of furS)
CIIHinforGGTAAGCTTCTCCAGGGTCAGATG (located upstream of hbpS)
IIEcorevGCCGAATTCTCCTCAGCATGTCCAG (located upstream of hbpS and ending at the 5′-end of binding site II)
DIIEcoforCGAGAATTCGGGGGCGTCGGTCGC (located upstream of hbpS and beginning at the 3′-end of binding site II)
IIPstrevCACCTGCAGGGCGCGCGGGTGTCGTC (located downstream of the start codon of hbpS)
EIIHinfoFor sequence and characteristics, see above
IIIEcorevGCGGAATTCGGGCCGAGGATCGG (located upstream of hbpS and ending at the 5′-end of binding site III)
FIIIEcoforGCCGAATTCCGCCGGACCGGATG (located upstream of hbpS and beginning at the 3′-end of binding site III)
IIPstrevFor sequence and characteristics, see above
GIIHinforFor sequence and characteristics, see above
REcorevGACGGAATTCAGGATCGGTTCCGG (located upstream of hbpS and ending at the 5′-end of the inverted repeat)
HREcoforGCTCGAATTCCGTCCCGTCGCCGG (located upstream of hbpS and beginning at the 3′-end of the inverted repeat)
IIPstrevFor sequence and characteristics, see above
IIIHinforFor sequence and characteristics, see above
II/IIIEcorevGGGAATTCGGGCCGAGGATCGGTTCCGGGAGCGACCGACGCCCCCTCCTCAGCATGTCC  (located upstream of hbpS and ending at the 5′-end of binding site III, and containing a deleted binding site II)

To test the consequences of the deleted sites for SenR binding, EMSAs were done. For this purpose, a ∼ 100 bp fragment (containing or lacking binding site I) upstream of furS (up-furS2) was amplified from each of the corresponding plasmids, using primers P1 (5′-GAGGTGTACGGCGGGTGACGACAG-3′) and P1rev (5′-GTGGTCGGTGTCGGGGAGGCGG-3′). A second ∼ 100 bp fragment (containing or lacking binding sites II or III or the inverted repeat PIII) upstream of hbpS (up-hbpS2) was amplified from the corresponding plasmids with primers P2 (5′-GAGGACGCGGGTACGGCGGGACGG-3′) and P2rev (5′-CGGGGTCATCCGATCGGATGACCC-3′). Fragments were end-labeled with [32P]ATP[γP] using T4 polynucleotide kinase as a prerequisite for binding studies with SenR or SenR∼P.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References

We are very grateful to Dr Stefan Walter (in our group) for analyzing the correctness of SenSc peptides by LC-MS. G. Bogel and the running costs were financed in part by funds to H. Schrempf and in part from a grant from the Deutsche Forschungsgemeinschaft to D. Ortiz de Orué Lucana.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Experimental procedures
  6. Acknowledgements
  7. References
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