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Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Unusually among bacteria, actinobacteria possess myo-inositol 1-phosphate synthase (mIPS). In the developmentally complex Streptomyces coelicolor, the mIPS-encoding gene (inoA) is cotranscribed with a putative regulatory gene (inoR). The inoRA transcript was more abundant in an inoR in-frame deletion mutant, and InoR formed different complexes in vitro with an extensive region around the inoRA promoter. Binding was relieved by adding glucose 6-phosphate. Thus, InoR is a metabolite-sensitive autorepressor that influences inoA expression, and hence the level of inositol, by controlling transcription from PinoRA. Disruption of inoA resulted in inositol-dependent growth and development, with full phenotypic correction at 0.1 mM inositol: at lower inositol concentrations differentiation was arrested at intermediate stages. This pattern may partly reflect increased demand for membrane phospholipids during sporulation septation. A corresponding sharp upregulation of inoRA transcription coincident with sporulation was dependent on a developmental regulator, WhiI. A truncated form of WhiI could bind two sites downstream of PinoRA, and one of the WhiI-binding sites overlapped the InoR-binding site. The combined action of a metabolic regulator and a developmental regulator at the simple PinoRA promoter is a previously undescribed strategy for the differential provision of developmentally appropriate levels of a substance required during the formation of spore chains.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Streptomyces spp. are filamentous, mainly soil-dwelling, members of the Actinobacteria (high-G+C Gram-positive bacteria). On solid medium, they undergo complex multicellular differentiation: germinating spores give rise to substrate mycelium that, probably partially in response to nutrient limitation, produces branches that grow away from the medium to form aerial mycelium. The aerial hyphae elongate for a while, then the long tip compartments undergo multiple sporulation septation, eventually forming chains of up to c. 100 spores (Flärdh and Buttner, 2009). The sporulation process has been analysed via developmental white (whi) mutants that produce aerial hyphae but cannot form grey-pigmented spores (Chater, 2001). Studies of one of the whi genes, whiI, led to the work described in this paper.

The whiI product is an atypical paralogue of the response regulators found in bacterial two-component systems, and is directly and completely dependent on the sporulation-specific sigma factor encoded by whiG (Ainsa et al., 1999). Based on the terminal phenotypes of site-directed mutations, WhiI appears to act at two stages during sporulation: one activity is needed for sporulation septation and the other, which depends on some undefined post-translational modification or ligand-binding, is required for spore maturation (Tian et al., 2007). Transcriptome analysis revealed 45 genes that were affected in a whiI null mutant, some of which were also affected in constructed whiI point mutants expected to specify WhiI locked in one form or the other (Tian et al., 2007). Thus, the adjacent genes SCO3899 and SCO3900, which are the subject of this paper, were poorly expressed compared with the wild-type during sporulation of a whiI null mutant and a mutant expected to express a WhiI form fixed in the initial state (whiI D27A), suggesting that SCO3899 and SCO3900 were upregulated in response to a sporulation-associated change in WhiI. A first follow-up mutational study of SCO3900, which encodes a PadR-like putative regulatory protein, indicated that it was cotranscribed with SCO3899 and two further downstream genes (SCO3898, encoding a possible membrane transport facilitator protein, and SCO3897, of unknown function), with the SCO3900 gene product playing an autorepressing role (Zhang et al., 2010). A mutation in SCO3900 caused a partial block in development.

SCO3899 appears to encode an orthologue of the Mycobacterium tuberculosis myo-inositol-1-phosphate synthase (mIPS), which uses glucose 6-phosphate as substrate (Movahedzadeh et al., 2004), suggesting that inositol might be required during development in S. coelicolor. Although inositol is important in eukaryotic biology, it is absent from most bacteria. However, it is found in many actinomycetes. Inositol is normally synthesized in two steps. First, glucose 6-phosphate is converted to inositol-1-phosphate by inositol-1-phosphate synthase, and then dephosphorylated by inositol monophosphate phosphatase to produce inositol. From information in the KEGG (http://www.genome.jp/kegg/) resource and published literature, inositol mainly serves as a precursor of three kinds of molecule in the small number of prokaryotes in which it occurs: mycothiol (MSH), which is the primary reducing thiol compound in actinobacteria to protect against oxygen toxicity, a role played by glutathione in many other microorganisms (Newton et al., 1996; Fahey, 2001); phosphatidylinositol (PI), an essential phospholipid of eukaryotic cells whose occurrence in prokaryotic cells is largely confined to some actinobacteria (Jackson et al., 2000; Michell, 2008); and di-myo-inositol-phosphate (DIP) (Rodionov et al., 2007), a major osmoprotecting metabolite in a number of hyperthermophilic species of archaea and bacteria (Santos and da Costa, 2002).

Here we show that a SCO3899 mutant is inositol-dependent, and that different inositol metabolite levels are needed for S. coelicolor growth and development. These levels are shown to depend on the combined regulatory effects of the SCO3900 and WhiI proteins on the promoter of the SCO3900-3897 (inoRAXY) operon.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

A SCO3899 mutant depends on inositol to different extents for growth and differentiation

SCO3899 encodes a 360-amino-acid protein with 78% identity with the well-studied ino1 product (mIPS) of Mycobacterium tuberculosis (Movahedzadeh et al., 2004). Similar genes are generally present in Streptomyces and many other actinomycetes, always next to a SCO3900-like gene (data not shown). In the light of this similarity, and of results in this paper, SCO3899 was named inoA. To investigate the function of inoA, we set out to generate an inoA disruption mutant. A disruption plasmid, pKC1132::ΔinoA::neo, was constructed in E. coli. In this construct, the inoA coding sequence from +22 to +1039 nt was replaced by the neo gene. The disruption plasmid was introduced into Scoelicolor M145 by conjugation via ET12567/pUZ8002. Kanamycin-resistant colonies were selected, and then replicated to MS medium containing both kanamycin and apramycin to identify kanR/aprS double cross-over disruptants, but none were obtained. However, when 10 mM inositol was added to the medium, kanR/aprS colonies could be found (eight independent isolates), and these were confirmed as inoA disruption mutants by Southern analysis (data not shown). Thus, a role for inoA in inositol biosynthesis was confirmed, and inositol appeared to be important for the growth of S. coelicolor. One of these mutants (inoADM, as inoAdisruption mutant) was randomly selected for further study.

When re-streaked on MM medium with mannitol or glucose as the carbon source, strain inoADM showed weak growth and failed to produce aerial hyphae after 5 days, in contrast to the robust growth and grey-pigmented spores of M145 (Fig. 1A). Similar results were obtained on R2YE medium containing glucose as the carbon source (Fig. 1B). However, on the soy flour-containing MS (which presumably contains some inositol), strain inoADM grew more strongly, and produced fuzzy white aerial mycelium after incubation for 3 days, but was arrested at this stage even after incubation for more than 7 days (Fig. 1C). These results indicated that inositol might be required in higher amounts for differentiation than for vegetative growth. In complementation experiments, segments containing inoA inserted into the integrating vector pSET152 were introduced into strain inoADM (Fig. 1D). pHK102, which carries inoA coupled with its adjacent upstream gene inoR and 308 bp of DNA upstream of inoR, gave full complementation on MM and MS without inositol supplement, but poor complementation was seen on R2YE medium; while pHK101, which contains an entire inoA and its 237 nt upstream sequence, gave very poor complementation on MM. This result suggested that the smaller DNA insert in pHK101 did not contain a significantly active promoter driving inoA expression, and was consistent with previous data (Zhang et al., 2010), indicating that inoA is part of an inoRAXY operon, extending from SCO3900 to SCO3897. Because we did not study inoX and inoY any further, the operon is referred to as the inoRA operon in the rest of this paper.

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Figure 1. Phenotype of strain inoADM and its complemented derivatives after 5 days on different agar media. A. MM with mannitol as carbon source. B. R2YE. C. MS. D. Schematic diagram of complementation strategy. Strains: a, wild-type M145; b, inoADM (inoA disruption mutant); c, inoADMC1 (inoADM/pHK101); d, inoADMC2 (inoADM/pHK102); e, inoADM with pSET152 as control.

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To confirm that there is a higher inositol requirement for sporulation than for earlier stages of growth and development, spores of strain inoADM were spread on solid MM medium containing mannitol, and a filter disc containing 0.4 µmol inositol was placed in the centre of the Petri dish. After 5 days, the disc was surrounded by three distinct zones of growth (Fig. 2A). Near the centre, the growth was vigorous and covered with abundantly sporulating grey aerial mycelium. Outside this zone there was a further zone in which the aerial growth was white, and still further out there was little aerial growth. SEM confirmed that spore chains were abundant in the grey zone (Fig. 2B), but that few, and only immature, spore chains were present in the white zone (Fig. 2C). Beyond the white zone, only vegetative mycelium was seen (Fig. 2D). When incubation was continued, the zone boundaries were maintained. Additional confirmatory results were obtained when strain inoADM was incubated on MM medium with mannitol supplemented with different concentrations of inositol (1, 10, 100 and 1000 µM). The addition of 1 µM inositol led to moderate growth and the formation of sparse aerial hyphae; while strong, but still white and non-sporulating, aerial growth was seen on 10 µM inositol. With the addition of 100 µM inositol, the aerial mycelium was grey. Complete restoration of development to wild-type levels required 1000 µM inositol (Fig. 3).

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Figure 2. Effect of inositol on growth and differentiation. A. Strain inoADM (inoA disruption mutant) was incubated for 5 days on MM containing mannitol with 0.4 µmol of inositol on the filter disc in the centre of the 8.5 cm diameter Petri dish. Three distinctive phenotypes corresponding to zones I, II and III were observed in response to the inositol gradient. B–D. Morphology of inoADM examined by SEM. Samples from zones I, II and III of (A) corresponded to (B), (C) and (D) respectively.

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Figure 3. Effect of different concentrations of added inositol on the inoADM phenotype. Strain inoADM was grown on MM mannitol medium supplemented with different concentrations of inositol and photographed after incubation for the indicated times. The inoculated strains are indicated on the top left plate: a, M145; b, inoADM (inoA disruption mutant); c, inoADMC2 (inoADM/pHK102); d, inoADM/pSET152. Strain inoADM gave poor growth and little aerial mycelium phenotype in the absence of inositol; stronger growth but weak white aerial mycelium on 1 µM inositol; strong vegetative growth but aerial mycelium remaining pale on 10 µM inositol; mutant nearly indistinguishable from wild-type on 100 µM inositol; full restoration to wild-type phenotype on 1000 µM inositol.

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The inoRA promoter is upregulated in a WhiI-dependent manner during sporulation

Microarray data had indicated a role for the whiI gene product in a sporulation-associated increase in inoR and inoA transcript levels (Tian et al., 2007). To reinforce these results, and to determine the location of the promoter(s) involved, we used S1 nuclease protection assays to evaluate inoA transcript levels in RNA isolated from M145 (wild-type) and two phenotypically different whiI mutants: a whiI null mutant in which development was arrested at the pre-septation stage of sporulation, and a whiI (D27A) point mutant, which formed pale grey spores but could not undergo spore maturation (Tian et al., 2007). For mRNA isolation, the strains were grown for various time periods on MM with mannitol. The probe was designed to detect both transcripts with 5′ ends close to inoA, and those with 5′ ends further upstream (including those emanating from the inoR promoter region), referred to as readthrough transcripts (Fig. 4A, Experimental procedures). Transcription of the constitutive hrdB gene was used as a control. In wild-type M145, the strongest transcriptional signal was from readthrough transcripts, and there were two weaker signals with 5′ ends in the inoR gene, close to the start of inoA. Although clear-cut evidence is missing, we suspect that the two mRNA 5′ ends immediately upstream of inoA (Fig. 4B) may result from cleavage of the readthrough transcript, either in vivo or in the course of the S1 nuclease treatment. The respective transcriptional signals were fairly constant at all time points, except for a strong transient increase of all three signals at 72 h, the time at which sporulation was occurring most abundantly, and markedly diminished levels in the final sample (84 h). In both whiI mutants, the developmental up-shift and the subsequent diminution were absent, but levels in the earlier samples were otherwise similar to those in M145 (Fig. 4B).

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Figure 4. Transcriptional assay of inoA in M145 (WT), J2676 (ΔwhiI) and J2671 (whiID27A). A. The sketch indicates the design of the hybrid probe carrying a non-homologous tail from pHK101. B. The hybrid probe was used to detect inoA transcripts. RNAs from M145, J2676 and J2671 grown on MM mannitol plates for 18, 36, 48, 72, 84 h were hybridized with inoA probe for S1 mapping. A probe for hrdB mRNA was used as an internal control for RNA quality and loading.

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To establish whether the readthrough inoA transcript included the whole of inoR and inoA, RT-PCR was performed with mRNA isolated from M145. As predicted, using the specific reverse-transcribed cDNA as the template, a DNA species of about 1.8 kb spanning from the start codon of inoR to the stop codon of inoA was detected by agarose gel electrophoresis. In PCR controls with the same primer pair, the same sized product was obtained using genomic DNA as the template, whereas no such product was detected with the DNA-free mRNA as the template (data not shown). Therefore, the developmental up-shift in inoA mRNA abundance was attributable to changes in promoter activity upstream of inoR. Taken together with the inositol dependence of development in the inoADM mutant, it seems likely that the increase in transcript abundance is implicated in the provision of extra inositol metabolites during sporulation. This up-shift depends on the developmental regulator WhiI.

InoR is auto-repressing and binds to its own promoter

Our previous results indicated that InoR played a negative regulatory role in the differentiation of S. coelicolor through its effects on the expression of the inoRA operon: overexpression of inoR in the wild-type M145 caused poor growth and development on MM similar to the phenotype of strain inoADM (Zhang et al., 2010). We found subsequently that the phenotype of the overexpression construct (M145/pIJ8600::inoR) was fully corrected by adding 0.1 mM inositol to the medium (data not shown). This indicated that the detrimental effect of inoR overexpression was due to inositol/inositol-1-P depletion, caused by super-repression of the chromosomal inoRA operon by the trans-acting plasmid-encoded repressor.

These results strongly suggested that InoR should bind to DNA in or adjacent to the operon. This was investigated by in vitro band-shift assays. InoR-His6 was overexpressed in E. coli, and the protein was purified by Ni-NTA agarose chromatography. The protein behaved as a dimer during gel filtration (data not shown). Band-shifts were done with two [γ-32P]-ATP-labelled probes: a fragment extending 235 bp upstream of the inoA start codon, and a 314 bp DNA fragment containing the inoR promoter region. Binding of InoR to the inoR promoter caused clear band-shifts at protein concentrations ranging from 12.5 nM to 3.75 µM, giving rise to at least five InoR-probe binding complexes as the protein concentration increased (Fig. 5). The labelled probe was totally shifted into protein–DNA complexes at 0.375 µM InoR. Specificity of binding was demonstrated by showing that 30- and 100-fold excess unlabelled probe released InoR (0.375 µM) from the labelled probe. On the other hand, InoR could not form specific complexes with the DNA immediately upstream of inoA, although a weak shift band appeared when the concentration of InoR reached 1.25 µM; a concentration at which InoR exhibited non-specific binding to a DNA fragment including the promoter of hrdB (Fig. S1). Based on these results, we conclude that InoR influences growth and development through its auto-regulatory effects on levels of the readthrough transcript from inoR to inoA, consequently affecting inositol/inositol-1-P biosynthesis.

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Figure 5. Electrophoretic mobility shift assays to determine the binding of InoR to the PinoR region. The 32P end-labelled PinoR probe was incubated with increasing concentrations of InoR. EMSAs with 375 nM InoR and 30-, 100-fold excess unlabelled cold probe were used to detect the specificity of the binding.

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InoR binding protects a long region of DNA around the inoRA transcription start point

The inoR transcription start point was determined as ‘G’ at 48 nt upstream of the translational start codon by high resolution S1 mapping (Fig. 6A). To investigate the InoR-binding site in the inoR promoter, a DNase I footprinting experiment was performed. The result revealed that InoR has an extensive binding region overlapping the inoR sense strand sequence from −28 nt to +30 nt with respect to the transcriptional start site of inoR, and the antisense strand sequence from −20 nt to +38 nt (Fig. 6B). In the protected region, a long perfect inverted repeat (5′-CGTGCTGATGTATCGACTCGATACATCAGCACG-3′) shares some similarity with the typical PadR binding site (ATGT-8N-ACAT), and includes the transcriptional start point (Fig. 6C). Thus, InoR seems likely to negatively regulate its own expression and that of inoA by inhibiting binding of RNA polymerase to the inoR promoter. An identical sequence was present in the corresponding position in the inoRA operons of the six other Streptomyces genomes in StrepDB (http://strepdb.streptomyces.org.uk), but no similar sequence was present anywhere else in any of the genomes, indicating that InoR is likely to regulate inoRA expression specifically.

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Figure 6. Determination of inoR transcription start point (TSP) and DNase I footprinting of InoR on PinoR. A. RNA from M145 grown on MM mannitol for 72 h was used to identify the TSP of inoR by high resolution S1 mapping. The bent arrow indicates the TSP and the transcriptional orientation. B. The PinoR fragment was labelled in the top and bottom strands respectively. Regions protected against DNase I digestion upon binding to increasing concentrations of InoR are indicated by positions relative to the TSP of inoR. G, A, T and C indicate the nucleotide sequence ladder of PinoR. C. Nucleotide sequence of the inoR promoter and InoR binding sites. The numbers indicate the distance in nt from the TSP of inoR. The TSP of inoR is indicated by a bent arrow. The InoR binding sites on the top and bottom strand are indicated by solid lines and dashed lines respectively, in which a 16 bp perfect inverted repeat sequence is marked by convergent arrows. The putative ribosome binding site (RBS) and −10 and −35 regions are marked by boxes. The coding sequence of inoR is in italics.

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WhiI-C binds to sequences in the inoRA promoter region that overlap with the InoR-binding site

To determine whether direct interaction of WhiI with the inoRA promoter region might account for the WhiI-dependence of the developmental up-shift in inoRA, we tried, but failed, to express soluble and active full-length WhiI protein of different types: N- or C-terminally His-tagged protein (protein was insoluble), and glutathione-S-transferase (GST) fusion protein (protein was soluble, but failed to complement the developmental deficiency of a whiI mutant of S. coelicolor, or to bind the inoRA promoter region). However, we were able to obtain the truncated WhiI C-terminal domain as a soluble N-terminally His-tagged protein (WhiI-C) that exhibited DNA-binding activity in vitro. This protein consists of 103 amino acids (excluding the His-tag). It has a theoretical molecular weight of 10.63 kD, and iso-electric point value of 12.05.

In band-shift assays, purified WhiI-C showed distinct banding patterns with PinoRA DNA (Fig. 7A), whereas no specific band-shift was seen with a non-cognate promoter (PhrdB, the promoter of hrdB) (Fig. S2). The band-shift pattern could be released by using excess unlabelled cold PinoRA probe, implying that binding was specific. The shifted band was super-shifted by the further addition of anti-WhiI-C polyclonal antibody (Fig. 7B). Therefore, we concluded that the inoR promoter is a direct target of WhiI. To our knowledge, no other direct target of WhiI has yet been identified in the literature.

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Figure 7. Electrophoretic mobility shift assays to determine the binding of WhiI-C to the PinoR region. A. EMSAs were performed with 0.5 ng 32P end-labelled PinoR probe, increasing concentrations of the C-terminal of truncated WhiI protein (WhiI-C) and 1 µg of poly-(dI-dC). EMSA with 30 nM of WhiI-C and 250-fold excess unlabelled cold probe was used to detect the specificity of the binding. B. EMSAs were performed as in (A), but 2000-fold diluted anti-WhiI-C antibodies in final concentration were added to give super-shift complexes.

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To compare the position(s) of WhiI-binding to that of InoR in the PinoRA region, DNase I footprinting was carried out on the WhiI-C/PinoRA complex. WhiI-C protected two regions of the probe DNA. One of these, site I, overlapped the inoR sense strand sequence from +19 nt to +42 nt (Fig. 8A) and the antisense strand sequence from +21 nt to +39 nt (Fig. 8B) with respect to the transcriptional start site. The other, site II, was within the coding region of inoR, and overlapped the inoR sense strand sequence from 77 nt to 96 nt (Fig. 8A) and the antisense strand sequence from 78 nt to 95 nt (Fig. 8B) with respect to the translational start codon. Site I partially overlapped the inverted repeat suggested as a potential InoR recognition sequence (Fig. 8C). It therefore appears that transcription of inoRA is regulated by two proteins, InoR and WhiI, both of which interact directly with DNA in or close to the inoRA promoter; and, since the binding sites overlap that the two proteins may compete with each other for binding.

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Figure 8. DNase I footprinting of WhiI-C on PinoR. A. About 30 ng PinoR fragment was labelled in the top strand. Two regions protected against DNase I digestion upon binding to increasing concentrations of WhiI-C (140, 210, 420, 840, 1400 nM respectively), are indicated by solid bent lines as site I and site II. G, A, T and C indicate the nucleotide sequence ladder of PinoR. B. Similar amounts of PinoR fragment labelled in the bottom strands were employed in footprinting experiments with the same amounts of WhiI-C as above. Two protected regions are indicated by dashed bent lines. C. Nucleotide sequence of the inoR promoter and WhiI-C binding sites, indicating the relationship with InoR binding sites. The numbers indicate the distance in nt from the TSP of inoR. The TSP of inoR is indicated by a bent arrow. The WhiI-C binding sites on the top and bottom strand are indicated by solid bent lines and dashed bent lines respectively. InoR binding sites on the top and bottom strands are indicated by straight and dashed lines. The 16 bp perfect inverted repeat sequence is marked by convergent arrows. The putative ribosome binding site (RBS) and −10 and −35 regions are marked by boxes. The coding sequence of inoR is in italic.

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Glucose 6-phosphate affects the binding of InoR to the inoRA promoter

Evidence from the phenotypes of site-directed whiI mutations previously indicated that WhiI might respond to some unknown developmental cue to change the spectrum of genes subject to its influence, including inoRA (Tian et al., 2007). We were interested to determine whether InoR might also respond to some signal, since it appears to form an auto-repression circuit on the inoRA operon. We therefore tested whether the substrate, product or any metabolically related molecules might affect this regulatory circuit. Thus, glucose, glucose 6-P, fructose 6-P, inositol-1-P or inositol was directly added to the binding system. In the band-shift assay, a moderate amount of InoR protein was used to give an intermediate band-shift, thereby maximizing sensitivity. The most striking effects were seen when glucose 6-P was added: at final concentrations of 17.5 to 20 mM glucose 6-P, InoR was dissociated from the inoRA promoter (Fig. 9A). Of glucose, fructose 6-P, inositol-1-P and inositol, only glucose showed any effect, 25 mM glucose causing partial dissociation of InoR-DNA complexes (Fig. 9B and C). The effect of glucose 6-P was clearly seen in DNase I footprinting experiments (Fig. 9D).

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Figure 9. Effect of ligands on the binding activity of InoR. EMSAs were performed with 0.5 ng 32P end-labelled PinoR probe, 15 nM InoR (final concentration), and different amounts of potential ligands. A. The effects of glucose 6-P at final concentrations of 1, 5, 10, 12.5, 15, 17.5 and 20 mM on the binding activity of InoR to PinoR. A binding assay without InoR but with 10 mM glucose 6-P was used as the control. B. Similar binding assay as in (A), but glucose replaced glucose 6-P as the ligand. C. Similar binding assay as in (A), but different concentrations of fructose 6-P, inositol and inositol-1-P were used as the ligands. The first lane contains the free DNA probe. D. Effects of increasing concentrations of glucose 6-P on binding of InoR to PinoR detected by DNase I footprinting. About 30 ng probe, with labelled top strand, was used. Different concentrations of InoR and glucose 6-P are indicated at the top of the gel. With the dissociation of InoR from PinoR at glucose 6-P concentrations from 17.5 to 25 mM, new cleavage fragment signals emerged that were similar to the control (non-protein bound DNA).

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These results indicated that feedback repression by inositol-1-P or inositol was not involved in inoRA regulation, in contrast to the situation in yeast (Hirsch and Henry, 1986). On the other hand, there was an intriguing suggestion of feed-forward regulation by high concentrations of the mIPS substrate glucose 6-P.

The phenotypes of inoR mutants are correlated with the level of inoA expression

Based on the results in this paper, non-polar disruption of inoR should increase the expression of inoA. We examined this prediction by Western blotting and S1 protection assays on inoRDM, which had an in-frame deletion of a segment encoding the winged helix–turn–helix (wHTH) motif. The inoRDM strain produced small colonies with a relatively thin aerial mycelium, but sporulated earlier than the wild-type (Fig. 10A), and did indeed show de-repressed inoA transcription and increased levels of InoA protein (Fig. 10C and E).

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Figure 10. Effect of inoR disruption on the phenotype and expression of both inoR and inoA. A and B. Phenotype of different inoR disruption mutants on MM mannitol medium in the absence (A) or presence (B) of 0.1 mM inositol for 4 days. C and D. Transcriptional analysis of inoR and inoA in strain inoRDM (C) and strain inoRFS (D); RNAs from M145, strain inoRDM and inoRFS time-courses grown on MM mannitol were hybridized with inoR and inoA probe respectively for S1 mapping, and hrdB was used as a control for RNA quality and loading. E. Western blot to detect the expression level of InoA in strain inoRFS, D3900 and inoRDM compared with M145 during a time-course.

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We also studied two inoR mutants expected to have polar effects. The frameshift mutant inoRFS exhibited delayed development that could be restored to wild-type levels by adding inositol (Fig. 10A and B), and had lower levels of InoA in Western blots than the wild-type M145 (Fig. 10E), and correspondingly low levels of inoA-specific mRNA (Fig. 10D). A probe specific for the inoR sequence upstream of the mutation gave a very high signal, consistent with the loss of the autorepressing activity of InoR (Fig. 10D). As described in our previous work (Zhang et al., 2010), the other polar mutant (D3900, with neo inserted in inoR), had a bald phenotype on inositol-free MM, and complementation experiments indicated that this was due to polar effects on the downstream part of the transcription unit. Consistent with this, InoA protein was barely detectable by Western blotting (Fig. 10E), and the morphological phenotype could be restored to wild-type by adding inositol (Fig. 10A and B).

Taken together, the phenotypes of the three inoR mutants could all be explained by a combination of the InoR autorepression circuitry and the level of readthrough transcription into inoA, which in turn affected the level of InoA protein and hence the biosynthesis of inositol-1-P and inositol.

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

SCO3899 encodes the major route for inositol biosynthesis in S. coelicolor

Based on the close resemblance of the SCO3899 protein to the myo-inositol-1-phosphate synthase (mIPS) of M. tuberculosis, and the inositol dependence of a SCO3899 mutant, we concluded that the SCO3899 gene product is mIPS, and that it is part of the main pathway of inositol biosynthesis. The gene was therefore named inoA. The further conversion of inositol-1-P to inositol may be carried out by the predicted inositol monophosphatase (IMP) encoded by SCO5860.

Possible biological roles of inositol related to S. coelicolor differentiation

As indicated in Introduction, the known roles of inositol in bacteria are as a precursor of mycothiol (MSH), phosphatidylinositol (PI) and di-myo-inositol-phosphate (DIP). Since DIP has not been reported in actinobacteria, and the S. coelicolor genome contains no obvious determinants for DIP biosynthesis, we do not consider DIP further here.

The predicted MSH deficiency of inoA mutants may well contribute to their developmental defect, but it is unlikely to be the full explanation of the phenotypic consequences of inositol limitation, since the phenotypes of mshA (SCO4204) and mshC (SCO1663) disruption mutants (slower differentiation of mutants than the wild-type: Park et al., 2006, and our unpublished results) are less severe than that of strain inoADM: we found that an mshA mutant could produce mature spores after incubation on both MM and R2YE medium for 7 days (our unpublished results), unlike the terminal white phenotype of strain inoADM. Moreover, a sigR mutant, lacking σR, which controls the production of MSH by directly regulating the expression of mshA (Park and Roe, 2008), showed fourfold less MSH production than the wild-type, but was unaffected in growth and morphological differentiation (Paget et al., 1998).

We therefore suggest that a limitation for phosphatidylinositol (PI) may also contribute to the inoADM phenotype. Most studies on the biosynthesis by actinomycetes of PI, which is made by the condensation of inositol with CDP-diacylglycerol, and of PI derivatives, have been carried out in mycobacteria, since lipomannan (LM) and lipoarabinomannans (LAMs), which contain a PI anchor (Hunter and Brennan, 1990), are major mycobacterial antigens and potential immunomodulators in tuberculosis and leprosy (Besra et al., 1997; Nigou et al., 1997; Dinadayala et al., 2006). The PI synthase of M. tuberculosis has been identified and its corresponding gene, pgsA, has been cloned (Salman et al., 1999; Jackson et al., 2000). The further biosynthetic sequence is thought to be PI[RIGHTWARDS ARROW]PIM (phosphatidylinositol mannoside)[RIGHTWARDS ARROW]LM[RIGHTWARDS ARROW]LAM (Besra et al., 1997). In a database search, we found a likely orthologue of pgsA in the S. coelicolor genome (Bentley et al., 2002): the SCO1527 gene product (241 aa) has 43% identity with PgsA (217 aa) in a nearly whole-length amino-acid comparison. SCO1527 is the first of three genes of a probable transcription unit consisting of SCO1525-1527. The SCO1525 product (387 aa) showed 54% identity with PimA (406 aa), which catalyses the transfer of a mannose residue to the myo-inositol ring of PI in M. smegmatis and M. tuberculosis (Kordulákováet al., 2002) to form phosphatidylinositol monomannoside (PIM1) and phosphatidylinositol dimannoside (PIM2); and the SCO1526 product (311 aa) had 49% identity with the MSMEG_2934-encoded acyltransferase (304 aa) of M. smegmatis mc2 155, which catalyses the transfer of palmitate to the mannose residue of PIM1 or PIM2 to form Ac1PIM1 and Ac1PIM2 respectively (Kordulákováet al., 2003). This implies that PI and its derivatives are probably also made in S. coelicolor. Indeed, in the very closely related Streptomyces lividans, PI makes up about 10–15% of the total phospholipid (Chouayekh et al., 2007), and phosphatidylinositol mannosides and dilyso-cardiolipin-phosphatidylinositol (DLCL-PI) have been found as a membrane component in S. hygroscopicus (Hoischen et al., 1997), so PI synthesis is probably a general attribute of the genus (these genes are also conserved in many other actinomycete genomes). Studies of SCO1527 are in progress.

Clearly, these considerations indicate that inositol is involved in the synthesis of PI and its derivatives in S. coelicolor, and we suggest that these phospholipids may be required in particularly large amounts during sporulation septation of aerial hyphae. During this process, double membranes grow in to form sporulation septa between each copy of the genome, synchronously along the complete length of the long apical compartment, and probably in no more than a few minutes (Chater, 1993). For each genome in the apical compartment, this implies the rapid synthesis of c. 1.6 µM2 of membrane per genome (twice the cross-sectional area of the apical compartment, diameter c. 1 µM). However, the growing apical compartments of the thinner vegetative hyphae (c. 0.5 µM diameter) extend by about 20 µM h−1 (Jyothikumar et al., 2008), a length accommodating about 10 copies of the genome, with only rare septation. This corresponds to about 3.14 µM2 of new membrane per genome per hour. Thus, at the time of sporulation septation, the demand for membrane synthesis is transiently perhaps sixfold higher (c. 19.6 µM2 per genome per hour, assuming 5 min for the formation of sporulation septa) than during normal growth (c. 3.14 µM2 per genome per hour). If the proportion of phospholipid in lateral membranes and septa is the same, this would explain the increased inositol consumption during sporulation.

Inositol polyphosphates and PIs are important signalling molecules in eukaryotes, and evidence has been presented to suggest that PIs might fulfil a similar role in streptomycetes (Chouayekh et al., 2007). We cannot exclude the possibility that some aspects of the mutant phenotypes that we describe may reflect such a signalling role.

Regulation of inositol biosynthesis in S. coelicolor

Based on our results, we can propose a model for the regulation of de novo biosynthesis of inositol and the relationship between inositol and differentiation of S. coelicolor (Fig. 11). The inoRA operon organization ensures that, since inoR itself is always expressed at least at a low level governed by its autorepression, the cotranscribed inoA is also always expressed, providing a steady supply of inositol during most stages of growth and development. This basic circuitry may simply depend on the concentration of InoR, or also be influenced by a sensitivity of InoR to some other signal. Analysis of other actinobacterial genomes indicates that inoRA is widespread among them, and that a sequence closely similar to the inverted repeat sequence to which InoR binds is usually present upstream of inoR. No similar sequence was found anywhere else in any of the genomes, indicating that the regulatory effect of InoR is confined to the operon.

image

Figure 11. Model for the regulation of inositol levels during growth and development. The autoregulatory circuitry controlled by InoR determines a basal level of inoA expression and thus of inositol biosynthesis, except during sporulation, when an unknown signal (yellow arrow) causes WhiI to change from a form that is needed for sporulation septation to a form that directly interferes with InoR repression. The resulting overexpression of inoRA leads to increased inositol biosynthesis, which is needed for sporulation septation to be completed. Then, the increased InoR should able to re-establish repression, perhaps forming a particularly effective binding complex to displace WhiI. In this model, the effect of glucose 6-phosphate on binding of InoR to PinoR (indicated in brackets) is not considered to be of developmental significance.

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In yeast, mIPS is feedback repressed by inositol (Hirsch and Henry, 1986), but in S. coelicolor we could find no evidence that inositol or inositol-1-P affected the interaction of InoR with the inoRA promoter. Instead, we found that glucose 6-P, the substrate of mIPS, could interfere with binding. However, this effect was seen only at concentrations somewhat higher than those reported for total hexose phosphate pools in exponentially growing E. coli (8.8 mM) (Bennett et al., 2009), suggesting that it may function mainly to avoid draining the pool of this key glycolytic intermediate, which is at the hub of several biochemical pathways (interestingly, glucose 6-P is closely related to glucosamine 6-P, an important N-acetylglucosamine-derived global signalling molecule for the onset of development and secondary metabolism in S. coelicolor: Rigali et al., 2006; 2008). It is worth noting that the inoR mutant strain inoRDM, which overproduced InoA, exhibited small colonies with reduced aerial growth but earlier sporulation, possibly caused by a nutrient stress response elicited by the excessive conversion of glucose 6-P into inositol. The high concentration of glucose 6-P and glucose led to the release of InoR from its target. It may also suggest that the certain ligand of InoR in vivo is probably some other metabolite that hasn't been tested in our experiments so far. There is also a possible trivial explanation of the glucose 6-P effects: they might conceivably be due to non-enzymatic modification reactions of some reducing sugars (such as glucose 6-P, glucose and fructose 6-P) with protein (Monnier and Cerami, 1981), though similar effects on DNA (Bucala et al., 1984) seem to be ruled out by the DNase I footprinting results (Fig. 9D).

To accommodate the requirement for additional inositol during sporulation, the InoR autorepression circuit is altered through the direct action of WhiI, probably in a form modified as a response to the progression of development: a mutant form of WhiI (D27A) thought to be unable to attain this modified state did not show the sporulation-associated surge in inoRA expression (Fig. 4B). The only form of WhiI available experimentally, a His-tagged WhiI C-terminal domain, was able to bind specifically to the inoRA promoter region (Fig. 7A and B). The simplest model for the role of WhiI in inoRA expression is that WhiI binds cooperatively to site I and site II (an unusual binding position for an activator), displacing InoR from its overlapping binding site, and making the promoter available to RNA polymerase. Repressor displacement is known in eukaryotic systems (e.g. Kukimoto and Kanda, 2001). We could find no example in bacterial systems, though the opposite situation – displacement of an activator by a repressor – has been demonstrated (Squire et al., 2009).

The substantial increase in InoR resulting from the interaction of WhiI with the inoRA promoter should then be able to re-establish repression, perhaps forming a particularly effective binding complex (taking into account the large number of complexes revealed in EMSA experiments in Fig. 5, and the long segment protected against DNase in Fig. 6). This may be responsible for the sharp decline of inoA expression at the last time point (Fig. 4B), ensuring that carbon and energy resources are not diverted unnecessarily into inositol biosynthesis. We found no evidence of any other promoters driving inoA expression, except the secondary mRNA 5′ ends detected in S1 analysis, which were most probably generated by cleavage of the primary transcript, either in vivo or during the S1 nuclease treatment.

There are several examples of vegetatively expressed functions that are also required at high levels at specific stages of development in S. coelicolor. This end is achieved in the case of ftsZ by the use of an additional transcription start site subject to specific developmental activation (Flärdh et al., 2000; Kwak et al., 2001). In the case of glycogen/trehalose metabolism via the glgE pathway (Kalscheuer et al., 2010), there are two almost identical copies of a complex operon, one expressed only in older parts of the substrate mycelium, the other only in sporulating aerial hyphae (Schneider et al., 2000). The dual control of a single primary metabolism promoter by a pathway-specific repressor (possibly subject to metabolic regulation) and a developmental regulator (possibly subject to checkpoint control) described here is a distinct strategy, and further studies are in progress to understand the molecular interactions taking place at the inoRA promoter.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Bacterial strains, plasmids and growth conditions

Media and microbiological methods were as in Kieser et al. (2000) unless stated otherwise. Bacterial strains and plasmids used are summarized in Table 1. S. coelicolor M145 and its derivatives were grown at 30°C on solid medium MS (soya flour with mannitol), and minimal medium with 0.5% mannitol or 1% glucose (supplemented with 0.1 mM inositol if necessary), or R2YE, for phenotypic and morphological observations. For DNA extraction, Streptomyces strains were incubated in YEME (yeast extract-malt extract) liquid medium. For the phenotypic study of an inoA defective mutant, the strain was grown on MS solid medium containing 1 M inositol at 30°C for 7 days to get abundant spores. The spores were harvested and washed twice in distilled water to get rid of any trace of inositol, and then resuspended in 10% glycerol for storage and further inoculation.

Table 1. Strains and plasmids used in this study.
 CharacteristicsReference
S. coelicolor strains (M145 derivatives)
 M145Prototrophic; SCP1-, SCP2- Kieser et al. (2000)
 J2676M145 ΔwhiI Tian et al. (2007)
 J2671M145 whiI (D27A) Tian et al. (2007)
 inoADMM145 ΔinoA::neoThis work
 inoADMC2inoADM with pHK102This work
 inoADMC1inoADM with pHK101This work
 inoADM/pSET152InoADM containing pSET152 as a controlThis work
 D3900disruption mutant of inoR with oppositely oriented neo insertion Zhang et al. (2010)
 inoRFS inoR frameshift mutantThis work
 inoRDMM145 ΔinoRThis work
 inoROEM145/pIJ8600::inoR Zhang et al. (2010)
 M145/pIJ8600M145 containing pIJ8600 as a control Zhang et al. (2010)
E. coli strains  
 DH5αF-, φ80lacZΔM15, Δ(lacZYA-argF) U169, deoR, recA1, endA1, hsdR17 (rk-, mk+), phoA, supE44, λ-, thi-1, gyrA96, relA1Gibco BRL
 ET12567 dam, dcm, hsdS, cat, tet MacNeil et al. (1992)
 BW25113K-12 derivative, ΔaraBAD, ΔrhaBAD Datsenko and Wanner (2000)
 BL21(DE3)F-, ompT, hsdS, gal, dcm (DE3)Novagen
Plasmids  
 pBlueScript II SK (-)routine cloning and subcloning vectorStratagen
 pIJ773 oriT aac(3)IV Gust et al. (2003)
 pKC1132suicide vector for Streptomyces containing oriT RK2 Kieser et al. (2000)
 pSET152 aac(3)IV intφC31 Kieser et al. (2000)
 pIJ8600 tipAp expression vector Kieser et al. (2000)
 pET23bExpression vectorNovagen
 pIJ2925Derivates of pUC18 Kieser et al. (2000)
 pET23b::inoA inoA expression vector carrying His-tag in the C-terminus of InoAThis work
 pET23b::inoR inoR expression vector carrying His-tag in the C-terminus of InoRThis work
 pIJ2925::ΔinoA::neoA 2.7 Kb DNA fragment containing the left and right flanks of inoA and neo was inserted into pIJ2925This work
 pKC1132:: ΔinoA::neoThe 2.7 Kb DNA fragment containing the left and right flanks of inoA and neo was inserted into pKC1132This work
 pHK1011.4 Kb fragment containing inoA and its upstream sequence was inserted into pSET152This work
 pHK1022.2 Kb fragment containing inoR promoter and whole sequence of inoR and inoA was inserted into pSET152This work
 SCH24 S. coelicolor cosmid containing inoR Redenbach et al. (1996)
 pFS1 aac3(IV) flanked by two AflII sites was introduced into inoR of SCH24This work
 pFS2reading frame of inoR was shifted by removing aac3(IV) from pFS1 with AflII digestion and self-ligationThis work
 pFS3cassette of aac3(IV) and oriT replace the neo of pFS2 for conjugal transferThis work
 pDEF1SCH24 derivative containing mutated inoR with the sequence of wHTH domain deleted in-frameThis work
 pDEF2cassette of aac3(IV) and oriT replace the neo of pDEF1 for conjugal transferThis work

PCR and DNA manipulation

PCR reactions were performed under standard conditions in the presence of 5% DMSO. Primers are listed in Table 2. The integrity of all DNA fragments amplified by PCR was confirmed by DNA sequencing. DNA manipulation, purification, ligation, restriction analysis, gel electrophoresis and transformation of E. coli were as in Sambrook and Russell (2001). Isolation of chromosomal DNA and transformation of Streptomyces were as in Kieser et al. (2000). The digoxigenin-11-dUTP labelling and detection kit (Roche Diagnostics GmbH, Mannheim, Germany) was used for the preparation of DNA probe and detection in Southern blot experiments, according to the protocols provided by the manufacturer.

Table 2. Oligonucleotides used in this study.
PrimerSequence (5′ to 3′)
  1. Asterisk indicates the primer was end-labelled with 32P when it was used to generate DNA used in EMSA, S1 mapping or DNaseI footprinting. Restriction sites underlined and the corresponding enzymes were indicated at the end of each primer sequence.

FS-FCCGATGCACGGCTACGAGCTGCTTAAGACGCTCAGTGGAACGAAA (AflII)
FS-RCAGTGACGTGTTGAGCCGTTTACTTAAGCATCTCGTTCTCCGCTCA (AflII)
whiICE1TATACCATGGGCAGCAGCCATCATCATCATCATCACAGCAGCGGCGGCGCCGCCCCCATCGGCG (NcoI)
whiICE2GCAGGATCCTCAGTGGA-TGATGCCCGTGCG (BamHI)
AAC3.4-FAGATCTGATCAAGAGACAGGATGAGGATCGTTTCGCATGATTCCGGGGATCCGTCGACC
AAC3.4-RGTCGCTTGGTCGGTCATTTCGAACCCCAGAGTCCCGCTCATGTAGGCTGGAGCTGCTTC
3900EX-FCTCCGGCTCATATGAGCCGGAGATCCGGCA (NdeI)
3900EX-RCGGCGGCCGCCGTGGCGGTGTCGCCGGG (NotI)
3900OE-RTCACGTGGCGGTGTCGC
3900S1-FGAGGACGAACCGGACTGAC
3900DE-FCCGGCTCATCGTCGCCTC
3900DE-RGAGGCGACGATGAGCCGGGACGCGTACGACGACGAG
3900Bind-F*GCCCGAAGTTCACCTCAAATCA
3900Bind-RCAGCGTCTTGAGGCAGGGATA
3899Bind-F*GGAATGGAGTCCGTGGAGC
3899Bind-RCGGGTCGGCGTCCTTGTAGT
defS1-R*CCGGGTGCGGGCCAGTG
RT-FTCAGCGCTCGACCTCACC
RT-RCTCGATACATCAGCACGGCA
hrdB-FACTCGGGCCACGCGGATTG
hrdB-R*AGCCTTTCCCCGCTCAAT
Q1GTTGAATTCTTGTCGCACGCAGAAGCTC (EcoRI)
Q11ATAGGATCCTGGCTACGCGAACCGAAC (BamHI)
Q2TATGGTACCGCGCCAACGTCGAGAAGT (KpnI)
Q22CGCAAGCTTCCATAGCAGAACCGCATC (HindIII)
Q3CGAGAATTCGGAATGGAGTCCGTGGAGC (EcoRI)
Q33/Q44TATGGATCCGGCACAGCCTCACATACGG (BamHI)
Q4TATGAATTCGGCGGTTTGCGACGGTGCTC (EcoRI)
Q5GAGCTACATATGGGTTCGGTTCGCGTAGC (NdeI)
Q55TATAAGCTTGCGCTCGACCTCACCCGC (HindIII)
3899S1-FGCTTCCGGCTCGTATGTTGT
3899S1-R*GGGTCGGCGTCCTTGTAGTA

Construction of inoA disruption mutant and complementation

PCR-targeted gene disruptions were as in Gust et al. (2003). To construct an inoA disruption mutant, two fragments flanking inoA were prepared by PCR with primer pairs Q1/Q11 and Q2/Q22, and placed on either side of the kanamycin resistance gene (neo) such that 1019 bp of the 1033 bp inoA coding sequence was replaced by neo. The ΔinoA::neo construct was then inserted into the BamHI site of pKC1132 to generate pKC1132::ΔinoA::neo, which was subsequently introduced into S. coelicolor M145 by conjugal transfer via ET12567/pUZ8002. Transformants were selected on MS medium containing kanamycin and inositol and screened for apramycin sensitive candidate double cross-over recombinants. These were confirmed by Southern blot and PCR (data not shown). One such mutant, termed inoADM (inoAdisruption mutant), was selected for further analysis.

Two DNA fragments were used for complementation analysis. A 1.4 kb EcoRI–BamHI DNA fragment carrying the entire coding region of inoA and 235 bp upstream of its start codon was amplified by primers Q3/Q33, and the PCR product was inserted into the same sites of pSET152 to generate pHK101. Similarly, a 2.2 kb EcoRI–BamHI DNA fragment containing inoA, inoR and 379 bp upstream of inoR was amplified by primers Q4/Q44 and subcloned into pSET152 to generate pHK102. Plasmids pHK101 and pHK102 were respectively introduced into strain inoADM by conjugation to give the corresponding inoADM complementation strains (inoADMC1 and inoADMC2).

Construction of frameshift and in-frame deletion mutants of inoR

To construct a frameshift mutation in inoR, the apramycin resistance cassette from pSET152 was amplified using primers FS-F and FS-R, each containing an AflII site, and the resulting PCR product was used to target inoR in cosmid SCH24 in E. coli BW25113/pIJ790. Digestion of the resulting cosmid (pFS1) with AflII, and blunt-end ligation following filling of the overhanging ends with Klenow Fragment, generated pFS2, in which a frameshifting insert of eight nucleotides (TTAATTAA) in inoR was confirmed by DNA sequencing. Using PCR targeting, the neo of the pFS2 backbone was then substituted by a cassette of aac3(IV) and oriTRK2 generated by PCR from pIJ773 using primers aac3.4-F and aac3.4-R, to give inoR frameshift disruption vector pFS3, which was further confirmed by restriction analysis. pFS3 was introduced into strain D3900 (kanR) by conjugation, selecting apramycin-resistant single cross-over recombinants, which were then screened for kanamycin-sensitive double cross-over replacement. One such mutant (inoRFS) was confirmed as an inoRframeshift mutant by DNA sequencing.

A similar strategy was used to construct an inoR in-frame deletion mutant. Two fragments corresponding to the sequence from −435 to +9 and the sequence from +301 to +678 with respect to the translation start codon of inoR were amplified respectively by primers pairs 3900S1-F/3900DE-F and 3900DE-R/3900OE-R. Because primer 3900DE-R had an 18 nt 5′-extension overlapping 3900DE-F, the two amplified sequences could be spliced together using overlap extension PCR with primer 3900S1-F and 3900OE-R. The final 822 bp fragment lacked the part encoding the entire predicted wHTH motif from the fourth to 100th amino acids (Horton et al., 1989). The 822 bp PCR product was co-electroporated into BW25113/pIJ790 along with linearized cosmid pFS1 digested with AflII. Re-circularization of the cosmid was achieved by double cross-over using the PCR product as a bridge, to give pDEF1. Sequencing of inoR in pDEF1 confirmed the in-frame deletion. The neo of the pDEF1 backbone was then substituted by aac3(IV) and oriTRK2 from pIJ773 using PCR targeting to give pDEF2, which was verified by restriction analysis and introduced by conjugation into strain D3900 (kanR). Exconjugants were then plated on antibiotic-free medium to identify double cross-over recombinants sensitive to both kanamycin and apramycin. One such mutant (inoRDM) was confirmed as an inoR in-frame deletion mutant by DNA sequencing.

Expression and purification of InoR and InoA

The inoR coding region was amplified using primers 3900EX-F and 3900EX-R. The amplified fragment digested with NdeI and NotI was inserted into pET23b to generate expression plasmid pET23b::inoR. Similarly, the inoA coding region was obtained by PCR using primers Q5 and Q55. The PCR product was digested with NdeI and HindIII, and inserted into pET23b to give pET23b::inoA. After confirmation by DNA sequencing, pET23b::inoR and pET23b::inoA were respectively introduced into E. coli BL21(DE3) for protein expression. The expression strains were grown at 37°C in 100 ml LB with 100 µg ml−1 ampicillin to exponential growth phase (OD600 of 0.6). IPTG was then added (final concentration 0.1 mM), and the cultures were incubated overnight at 30°C. The cells were harvested by centrifugation (6000 g, 4°C, 3 min), washed twice with binding buffer (20 mM Tris base, 500 mM NaCl, 5 mM imidazole, 5% glycerol, pH 7.9), and then resuspended in 10 ml of the same buffer. The cell suspension was treated by sonication on ice. After centrifugation (14 000 g, 20 min, 4°C), the supernatant was recovered, and the InoR-His6 or InoA-His6 were separated from the whole-cell lysate using Ni-NTA agarose chromatography (Novagen). After extensive washing with buffer (20 mM Tris base, 500 mM NaCl, 100 mM imidazole, 5% glycerol, pH 7.9), the InoR-His6 or InoA-His6 were specifically eluted from the resin with elution buffer (20 mM Tris base, 500 mM NaCl, 150 mM imidazole, 5% glycerol, pH 7.9) and concentrated by ultrafiltration (Millipore membrane, 3 kDa cut-off size) according to the protocol provided by the manufacturer. Protein purity was determined by Coomassie blue staining after SDS-PAGE on 12% polyacrylamide gel, and the protein concentration was quantified by the BCA protein assay Kit (NOVAGEN). The purified proteins were stored in 5% glycerol at −70°C before use for the gel retardation assay or to immunise mice to prepare polyclonal antibody for InoA.

Expression of a His-tagged WhiI C-terminal domain

The fragment which encodes the C-terminal DNA-binding domain of WhiI (WhiI-C) was amplified using the primers whiICE1 (5′-TATACCATGGGCAGCAGCCATCATCATCATCATCACAGCAGCGGCGGCGCCGCCCCCATCGGCG-3′; NcoI site underlined) and whiICE2 (5′-GCAGGATCCTCAGTGGATGATGCCCGTGCG-3′; BamHI site underlined). This fragment was digested with BamHI and NcoI, and subsequently cloned into the corresponding site of pET28a to generate pET28a::whiI-C. Subsequent steps were essentially as described above for InoR and InoA expression, except that the purified protein was stored in the elution buffer at 4°C.

Preparation of anti-WhiI-C antibody

To prepare rabbit anti-WhiI-C serum, 1 mg WhiI-C was used for the primary immunization, and two weeks later, boosting with 0.8 mg WhiI-C was performed weekly. After the fourth boost, the serum was harvested, and the titre and specificity was analysed by Western blot.

Electrophoresis mobility shift assays (EMSAs)

The EMSAs to detect the binding activity of InoR to the promoter region of inoR and inoA were performed as described previously (Yang et al., 2007). The 314 bp probe containing a 158 bp sequence upstream of the start codon of inoR was amplified with primers 3900Bind-R and 3900Bind-F* labelled by [γ-32p]-ATP with T4 polynucleotide kinase (Promage). Similarly, the 326 bp probe containing a 235 bp sequence upstream of the start codon of inoA was amplified using the primers 3899bind-R and 32P-end-labelled 3899bind-F*. During the EMSA, each 32P-labelled DNA probe (1000 c.p.m.) was incubated with different final concentrations of InoR from 0.001 to 12.5 µM at 25°C for 20 min in a buffer containing 1 µg of poly-(dI-dC) (Sigma), 20 mM Tris-HCl (pH 7.5), 1 mM dithiothreitol (DTT), 10 mM MgCl2, 0.8 µg BSA and 5% glycerol in a total volume of 20 µl. After incubation, samples were separated by electrophoresis (10 V cm−1, non-denaturing 4.5% polyacrylamide gels, running buffer containing 45 mM Tris-HCl (pH 8.0), 45 mM boric acid and 1 mM EDTA). Gels were dried and exposed to Biomax radiographic film (Kodak). A DNA fragment containing the promoter of hrdB was also labelled with 32P and used as a negative control to exclude the non-specific binding of InoR with DNA. For competitive gel shift assays, the labelled probe was pre-incubated with InoR-His6 for 20 min at 25°C, followed by the addition of 30- and 100-fold unlabelled probe (specific competitor) and incubation for another 20 min at 25°C. The resulting DNA–protein complexes were then subjected to electrophoresis and autoradiography as described above.

The binding activity of WhiI-C to the promoter region of inoRA was performed similarly. The inoR probe (314 bp) was purified from agarose gel with QIAquick Gel Extraction Kit (QIAGEN). After quantification by NanoVue (GE Healthcare Life Sciences), 1 µg of the probe was end-labelled with [γ-32P]-ATP. The labelled probes were purified with QIAquick Nucleotide Removal Kit (QIAGEN) and eluted in 30 µl ddH2O. One microlitre of 60-fold diluted probe (about 0.5 ng and 1000 c.p.m) was used for EMSAs with 0–600 nM WhiI-C. In some experiments, anti-WhiI-C polyclonal antibody was also added after 30 min incubation of WhiI-C with different probes at 25°C, and incubated for further 30 min at the same temperature before loading onto the EMSA gel, to observe the super-shifted complexes.

To detect the effect of candidate ligand molecules on InoR binding to the inoR promoter, different sugars (glucose-1-P, glucose, fructose-1-P, inositol-1-P and inositol) were added after initial binding described above, and then incubated for further 30 min at the same temperature before loading onto the EMSA gel.

DNase I footprinting

In order to characterize the InoR binding sites on both strands of the inoR promoter, the same probe used in the EMSA was labelled by [γ-32p]-ATP at the 5′ ends of its sense or antisense strands. The footprint reaction mixture contained 30 000 c.p.m. of 32P-labelled DNA probe, 0.025–250 pmol of InoR-His6, 2.5 µg of poly-(dI-dC) (Sigma) and 20 mM Tris-HCl (pH 7.5), 1 mM DTT, 10 mM MgCl2, 2 µg BSA and 5% (v/v) glycerol in a total volume of 50 µl. After incubation of the mixture at 25°C for 20 min, 5.5 µl RQ1 RNase-free DNase I Buffer and 0.5 U DNase I (Promega) were added, and the mixture was incubated for 1 min. The reaction was stopped by adding 50 µl of stop solution (20 mM EGTA, pH 8.0) and extracted with 100 µl of phenol/CH3Cl (1:1, v/v). After precipitation in ethanol, the pellet was washed with 75% (v/v) ethanol. The pellet was resuspended in 5 µl of H2O and 5 µl of formamide/dye mixture, and electrophoresed on a 6% (w/v) polyacrylamide/urea gel.

To characterize the WhiI-C binding sites on both strands of the inoR promoter, the same probe as above was labelled by [γ-32p]-ATP at the 5′ ends of its sense or antisense strands. About 30 ng 32P-labelled DNA probe was used together with the binding mixture, which contains 2.5 µg of poly-(dI-dC) (Sigma) and 20 mM Tris-HCl (pH 7.5), 1 mM DTT, 10 mM MgCl2, 2 µg BSA and 5% (v/v) glycerol, and 140, 210, 420, 840 and 1400 nM WhiI-C protein respectively, in a total volume of 50 µl. After incubation of the mixture at 25°C for 30 min, 5.5 µl RQ1 RNase-free DNase I Buffer and 0.375 U DNase I (Promega) were added, and the mixture was incubated for 70 s. The reaction was stopped and purified as above, and detected by electrophoresis on a 6% (w/v) polyacrylamide/urea gel. To determine the effect of glucose 6-P on the binding activity of InoR on inoR promoter, the same conditions were used, with InoR protein at 750 nM and 1–25 mM glucose 6-P added to the probe mixture.

RNA isolation, RT-PCR and S1 nuclease mapping

RNA corresponding to different stages of Streptomyces development was isolated from cultures on cellophane overlaid on MM mannitol plates as in Tian et al. (2007). The quality and quantity of RNAs were examined by UV spectroscopy and agarose gel electrophoresis.

RNA treated by RNase-free DNase I (Promega) was subjected to reverse transcription by SuperScript III RT (Invitrogen) with specific primer RT-F, as recommended by the manufacturer. Five per cent of the reverse transcription product was used as template for PCR amplification with primers RT-F and RT-R. PCR products were analysed via standard agarose gel electrophoresis. PCR of DNase-treated RNA was performed with the same primer pairs to detect any DNA contamination.

To determine the transcription start point of inoR and temporal expression of inoR and inoA, S1 nuclease mapping was performed using 40 µg of RNA in each assay. Probes were generated by PCR, using primers labelled at their 5′ end with [γ-32P]-ATP by T4 polynucleotide kinase as appropriate. Probes were obtained using the following primer pairs: inoR probe for wild-type used 3900Bind-F and 3900Bind-R* with M145 genomic DNA as template; inoR probe for inoRDM used 3900Bind-F and defS1-R* with inoR in-frame deletion mutant genomic DNA as template; the hybrid probe for inoA used 3899S1-F (annealing to the DNA template of pSET152 vector) and 3899S1-R* with pHK101 as template (asterisks indicate the labelled primers). A probe for hrdB, encoding the principal sigma factor in S. coelicolor, was used as control to assess the RNA quality and loading. The S1 nuclease mapping protocol was as in Kieser et al. (2000). DNA fragments protected by RNA were separated on 6% sequencing gels and visualized by autoradiography. Transcription start points of inoR and DNase I footprints were identified against a dideoxy-sequencing ladder (Promega fmol cycle sequencing kit) produced using the same labelled primer as was used to generate the probes for S1 analysis and DNase I footprinting analysis.

Western blotting assay of InoA expression

Cellophane disc cultures on MM mannitol plates were harvested and treated by sonication, and cell debris was removed by centrifugation. InoA in the supernatants was detected by Western blotting. Ten micrograms of total proteins separated by 12% SDS polyacrylamide gel (Fig. S3) was transferred to PVDF membrane using the Trans-Blot system (Bio-rad) at 50 mA overnight. The membrane was washed twice with PBS for 10 min and blocked in PBS with 7% skimmed milk powder solution (2 h, room temperature). The membrane was then washed twice with PBS and incubated for 1 h with appropriate InoA antiserum (1:2000-1:10 000 diulution) in PBS. Excess antibodies were removed by repeated washing with Tris-buffered saline with Tween-20 (TBST pH 7.4; 1 mM Tris-HCl, 150 mM NaCl, 0.05% Tween-20). After 1 h incubation in TBST containing the secondary antibody (10−4 dilution of goat IgG against mouse) conjugated with alkaline phosphatase, the membrane was washed twice with TBST, and the signals were detected by the BCIP/NBT colorimetric kit.

Scanning electron microscope (SEM) observation

For scanning electron microscopy, colonies were fixed in 2.5% (v/v) glutaraldehyde for 4 h, stained with osmic acid for 2–4 h, and dehydrated with ethanol of different concentrations. Each sample was coated with platinum-gold and examined by Hitachi D-570 scanning microscope.

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

This work was supported by grants from the National Natural Science Foundation of China (grant nos. 31030003 and 30970054) and the Ministry of Science and Technology of China (grant number 2009CB118905). K.F.C. was supported by a John Innes Foundation Emeritus Fellowship. We thank Govind Chandra for some of the bioinformatics analysis.

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  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
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