SGM Journals
Research Article

Autoregulation of hpdR and its effect on CDA biosynthesis in Streptomyces coelicolor

Haihua Yang, Yang An, Linqi Wang, Shuli Zhang, Yue Zhang, Yuqing Tian, Gang Liu, Huarong Tan

  • 1State Key Laboratory of Microbial Resources, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, PR China
  • 2Key Laboratory of Systematic Mycology and Lichenology, Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, PR China
  • Correspondence
    Huarong Tan
    tanhr{at}sun.im.ac.cn

    • Microbiology 2010; 156(9):2641–2648 · https://doi.org/10.1099/mic.0.038604-0

    View at publisher PubMed

    Abstract

    HpdR, an IclR-family regulator in Streptomyces coelicolor, is a substrate-dependent repressor for the tyrosine catabolic gene hppD. In this study, S1 nuclease protection assays revealed that hpdR is subject to a negative autoregulation. Purified HpdR showed specific DNA-binding activity for the promoter region of hpdR, indicating that the autoregulation of hpdR is performed directly. The disruption of hpdR led to reduced production of CDA by S. coelicolor J1501, suggesting a positive effect of hpdR on CDA biosynthesis. Electrophoretic mobility shift assays showed that HpdR specifically bound to the promoter region of hmaS (SCO3229 in the CDA gene cluster), encoding 4-hydroxymandelic acid synthase. Disruption of hmaS in J1501 abolished CDA production. It is possible that hpdR regulates CDA biosynthesis by controlling the transcription of hmaS.

    Edited by: J.-H. Roe

    INTRODUCTION

    Streptomycetes are high-G+C Gram-positive, filamentous soil bacteria that have become a major focus for the study of microbial development and differentiation, antibiotic biosynthesis, and the mechanism of transcriptional regulation (Chater, 1993; Chater & Chandra, 2008). The genetically well-studied strain Streptomyces coelicolor A3(2) produces at least four chemically distinct classes of antibiotics: the blue-pigmented polyketide actinorhodin, the red-pigmented undecylprodigiosins, the lipopeptide calcium-dependent antibiotic (CDA) and the cyclopentanone antibiotic methylenomycin. The triggering of antibiotic biosynthesis in Streptomyces species is generally tightly linked with the initiation of morphological differentiation, and both processes are subject to genetic modulation via a hierarchical regulatory network, integrating various physiological and environmental signals (Li et al., 2006).

    Transcriptional regulators have been found to play important roles in development and differentiation in Streptomyces species (Liu et al., 2005; Tian et al., 2007; Wang et al., 2009; Li et al., 2009). IclR-family regulators contain a helix–turn–helix (HTH) motif in their N-terminal domain and bind their target promoters as dimer or tetramers. They play significant roles in diverse biological processes in Streptomyces species, such as regulation of drug resistance, degradation of aromatic compounds, oxidative catabolism of tyrosine, and the inactivation of quorum-sensing signals (Yang et al., 2007, 2009, Santamarta et al., 2007). ssgR activates the transcription of ssgA in S. coelicolor, and is developmentally regulated at the transcriptional level (Traag et al., 2004). NadR plays a global regulatory role in amino acid metabolism, quorum sensing, morphological changes, antibiotic production and expression of chaperones in S. coelicolor (Yang et al., 2009). AreB from Streptomyces clavuligerus modulates leucine biosynthesis and cephamycin C and clavulanic acid production (Santamarta et al., 2007). In our previous work, HpdR, an IclR-type regulatory protein in S. coelicolor, was shown to be a negative regulator of the expression of hppD in tyrosine catabolism, in a substrate-dependent manner (Yang et al., 2007). It bound a 90 bp region spanning the promoter of hppD, which also overlapped the promoter of the adjacent hpdA, by which HpdR negatively regulated the transcription of hppD and hpdA. These studies were performed in S. coelicolor M145. When the development and differentiation of S. coelicolor strain J1501 was studied, we found some phenomena different from those of strain M145. In this study, we describe the autoregulation of hpdR and the effect of hpdR disruption on the transcription of hmaS, involved in CDA biosynthesis in S. coelicolor J1501.

    METHODS

    Bacterial strains, plasmids and growth conditions.

    Streptomyces coelicolor J1501 (hisA1 uraA1 strA1 pgl-1 SCP1−, SCP2−; differentiation phenotype as wild-type strain) (Chater et al., 1982) was used as host strain for gene propagation and gene disruption. Other S. coelicolor strains used in this work were all derived from J1501 (Table 1⇓). Escherichia coli JM109 was used as standard host for plasmid replication. The non-methylating E. coli ET12567 (dam dcm hsds; MacNeil et al., 1992) was used to propagate DNA when it was to be introduced into S. coelicolor derivatives.

    Table 1.

    S. coelicolor strains used in this study

    pKC1132 and pSET152 are bifunctional plasmids with autonomous replication in E. coli and capable of integrating into the Streptomyces chromosome, and harbour the apramycin-resistance gene aac(3)IV (Bierman et al., 1992). The Streptomyces/E. coli shuttle vector pKC1132 was used to construct the disruption derivatives of hpdR and hmaS. pSET152 (Bierman et al., 1992), which integrates into the S. coelicolor chromosome by site-specific recombination at the bacteriophage φC31 attachment site, attB (Kuhstoss & Rao, 1991), was used to introduce single copies of genes into the S. coelicolor chromosome. pGEX-4T-3 (Pharmacia Biotech, 27-4583-01), containing an ampicillin-resistance gene, was used for overexpression of hpdR in E. coli. Media, culture conditions, protoplast transformations and chromosomal DNA isolation for S. coelicolor strains were carried out as described by Kieser et al. (2000).

    When necessary, antibiotics were used in media as follows, at the concentrations given in parentheses: for E. coli, ampicillin (100 μg ml−1), kanamycin (100 μg ml−1), apramycin (50 μg ml−1) or hygromycin (50 μg ml−1) in LB; for Streptomyces, apramycin (100 μg ml−1), kanamycin (100 μg ml−1) or hygromycin (40 μg ml−1) in solid R2YE medium, or kanamycin (10 μg ml−1) or hygromycin (10 μg ml−1) in liquid YEME medium. The stock concentrations of apramycin, kanamycin and hygromycin stored at −25 °C were 100, 50 and 50 mg ml−1, respectively.

    DNA manipulation and sequencing anaylsis.

    Isolation of plasmid and chromosomal DNA, procedures for Southern blotting, and transformation of Streptomyces protoplasts were as described by Liu et al. (2005). All other DNA techniques, including transformation of competent E. coli cells, and blunt-ending of DNA by Klenow fragment and T4 polymerase, were as described by Sambrook et al. (1989), except that hpdR and hmaS probes used in Southern blot analysis were labelled non-radioactively (digoxigenin-11-UTP kit; Roche). For database searches, the programs tfasta (Lipman & Pearson, 1985) and blast (Altschul et al., 1990) were employed. Sequence analysis and homologue comparisons were performed with the programs of NCBI blastx.

    Construction of an hpdR null mutant.

    The hpdR disruption mutant, hpdRDMJ, was constructed as described previously (Yang et al., 2007), except that the host strain was J1501. In brief, the upstream region and partial 5′-end sequence encoding the α-helix–turn-α–helix domain of hpdR was replaced by the kanamycin-resistance gene (aphII) in J1501. To study the transcriptional level of the hpdR promoter without the influence of HpdR protein, plasmid pSET152 : : PhpdR, containing the 0.4 kb KpnI–HincII fragment of the hpdR promoter region, was introduced into strain hpdRDMJ to generate strain hpdRDMJ-P. The hpdR disruption mutant (hpdRDMJ) was selected by its KanR AprS phenotype and confirmed by Southern blot hybridization.

    Complementation of the hpdR mutant.

    To complement hpdRDMJ, a 1.4 kb KpnI fragment containing the complete promoter region and the coding region of hpdR was purified, filled in by T4 polymerase, and then subcloned into the site-directed recombinant plasmid pSET152 digested with EcoRV. The resulting plasmid pSET152 : : hpdR was introduced into hpdRDMJ, from which complemented colonies were selected by their KanR AprR phenotype. The complemented strain, hpdRCS, was further confirmed by Southern hybridization.

    Construction of an hmaS null mutant in J1501.

    An 11 kb KpnI/EcoRI-digested fragment containing hmaS and its flanking sequence from SupercosE63 (provided by Professor Keith Chater, John Innes Centre, Norwich, UK) was inserted into the same sites of pIJ2925. The resulting plasmid pIJ2925 : : hmaS was digested with EcoRI and NotI, from which a 3.36 kb fragment containing hmaS was isolated and inserted into the same sites of pBluescript M13− to generate pM13− : : hmaS, which was then further digested with BglII and ligated with the 1.7 kb BglII DNA fragment, from pIJ963, containing the hygromycin-resistance gene (hyg) to give pM13 : : hmaS : : hyg. This plasmid was subsequently digested with PstI and HindIII, from which a 3.2 kb DNA fragment containing the 1.5 kb left flanking arm and the 1.7 kb hyg (ArmL : : hygR) was isolated, and end-blunted with mung bean nuclease (Takara). pM13− : : hmaS was also digested with HincII, from which a 4.0 kb DNA fragment consisting of M13− and 1 kb right flanking sequence of hmaS was isolated. This fragment was ligated with the end-blunted 3.2 kb fragment (ArmL : : hygR) to generate plasmid pM13− : : ArmL : : HygR : : ArmR, which was further digested with XhoI and BstXI to give a 4.2 kb fragment containing ArmL : : HygR : : ArmR; this fragment was blunted by mung bean nuclease and then ligated with pKC1132 digested with EcoRV. Finally, the BglII–HincII fragment in hmaS was replaced by a 1.7 kb hygromycin-resistance gene to give pKC1132 : : ΔhmaS : : hyg, which was subsequently passed through E. coli ET12567 to remove possible Dam/Dcm methylation and then introduced into J1501 protoplasts using the PEG-mediated transformation method (Kieser et al., 2000). Colonies which were potential disruptants of hmaS were selected by their HygR AprS phenotype and identified by Southern blot hybridization. For Southern blotting assay, genomic DNA isolated from strains J1501 and hmaSDM was digested with NcoI and BglII, and the DIG-labelled 0.4 kb fragment upstream of the non-coding region was used as a probe.

    Overexpression and purification of GST–HpdR.

    In order to produce HpdR in E. coli, the complete coding region of the hpdR gene was amplified by PCR using chromosomal DNA as template and oligonucleotides corresponding to the start and stop codons as forward and reverse primers: hpdR-primer1, 5′-ATGGATCCGTGACCGCGGAGACCT-3′ (BamHI-recognized site underlined), and hpdR-primer2, 5′-ATACTCGAGCTACCGCAGCGCCTCCGCG-3′ (XhoI-recognized site underlined). The hpdR PCR product digested by BamHI and XhoI was inserted into pGEX4T-3 at the same restriction sites. The resulting pGEX-4T-3 : : hpdR was transformed into E. coli BL21(DE3) and the strain was cultured in 50 ml LB with 100 μg ampicillin ml−1 at 37 °C. This culture was grown to mid-exponential phase (OD600 0.6), then IPTG was added to a final concentration of 0.1 mM and incubation continued for an additional 2 h at 30 °C. Cells were harvested by centrifugation at 7700 g, 4 °C for 10 min, washed twice with 50 ml buffer (140 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.3) and then resuspended in 4 ml of the same buffer. The cell suspension was broken by sonification on ice. Triton X-100 was added to the cell lysate at a final concentration of 1 %, and the mixture was gently shaken at 4 °C for 30 min to get more soluble protein. After centrifugation at 12 000 g for 10 min at 4 °C, the supernatant was mixed with 0.4 ml glutathione Sepharose 4B solution (Amersham Pharmacia, 17-0756-01) and gently shaken in an ice bath for another 30 min, and then the mixture was loaded onto a mini-column (USB). After washing with 5 ml 1× PBS for three times, the GST–HpdR protein was specifically eluted with 1 ml elution buffer (10 mM reduced glutathione in 50 mM Tris/HCl, pH 8.0) and concentrated to about 3 μg μl−1 by ultrafiltration (Millipore membrane, 10 kDa cut-off). As a control for protein purification and protein–DNA binding assay, GST was purified by the same strategy and used at the same concentration in the EMSA (see below).

    Preparation of DNA probes for electrophoretic mobility shift assays (EMSAs).

    A 20 pmol sample of the downstream primer corresponding to the hpdR promoter, hpdR-primer4 (5′-CGTATCGGCCAGCAGTTTGAGGA-3′), was end-labelled with [γ-32P]ATP and 10 U T4 polynucleotide kinase (Promega) as described previously (Yang et al., 2007). It was used with the unlabelled hpdR-primer3 (5′-CGCCTACGTCCACCAGAGCG-3′) to generate the 518 bp fragment containing the hpdR promoter region by PCR. The labelled DNA probe containing the hpdR promoter region was purified using the PCR Purification kit Uniq10 spin columns from Shangon (Shanghai Sangon Biological Engineering & Technology Co.). Similarly, the probe containing the promoter region of hmaS was prepared using primers hmaS-up (5′-ACCTGCTCCTCACAATGC-3′) and hmaS-reverse (5′-GCGATGTCACTGGGCGGCAT-3′).

    EMSAs.

    For EMSAs of HpdR binding to target DNA, the reaction (20 μl) was carried out with about 0.2 pmol radiolabelled probe in buffer [125 mM HEPES pH 7.5, 2 mM DTT, 200 mM KCl, 10 mM MgCl2, 0.16 μg calf BSA μl−1, 10 % glycerol and 1 μg poly(dI-dC)]. Poly(dI-dC) from Sigma was included in each reaction to eliminate non-specific protein binding. The mixture was incubated with purified GST–HpdR or GST at 30 °C for 20 min. For competition gel shift assays, a 30-fold excess of unlabelled probe (specific competitor) was added to the reaction system with 900 ng GST–HpdR and incubated for 20 min at 30 °C, after which the corresponding labelled probe was added and incubated for a further 20 min at 30 °C. DNA–protein complexes were separated from free probe on a 5 % glycerol-containing non-denaturing 4.5 % acrylamide gel in 1× TBE buffer. The gels were dried and exposed to Kodak Biomax radiographic film.

    S1 nuclease protection assays of hpdR transcription.

    RNAs for S1 nuclease mapping were isolated from S. coelicolor J1501 and hpdRDMJ-P after incubation for different periods on R2YE medium covered with cellophane (Yang et al., 2007). S1 protection assays were performed using the hrdB gene (AY628703) as a control as before (Yang et al., 2007). The probe used for detecting hpdR transcript was generated by PCR using the unlabelled hpdR-primer4 and radiolabelled hpdR-primer 3. High-resolution S1 nuclease mapping was carried out to identify the transcription start point and gene expression pattern of hpdR in the wild-type and mutant strains. S1 nuclease mapping was performed as described previously (Kieser et al., 2000; Liu et al., 2005). When examining the expression level of hpdR in the wide-type and mutant strains, the probes for hpdR and hrdB (positive control for RNA integrity and loading) were added together to each RNA sample. The sequence ladder was made using an fmol DNA cycle sequencing kit (Promega) with the same labelled primer.

    qRT-PCR assays of hmaS.

    Total RNA samples from J1501 and hpdRDMJ-P, isolated as mentioned above, were treated with Genomic DNA cleaning reagent (Applygen), and then with DNase (Promega) to which RiboLock RNase Inhibitor (Fermentas) was added. The DNA-free total RNA sample was used to synthesize cDNA first. Pre-assays were performed using serial dilutions of cDNA as PCR template to examine the efficiency of amplification. To determine the level of hmaS transcription, real-time RT-PCR was performed using primers hmaS-primer1 (5′-TCCTTTCCCCTTCCTTCATTG-3′) and hmaS-primer2 (5′-GGGACGAAACCGAGGCTGT-3′) with the SYBR PrimeScript RT-PCR kit II (Takara). The relative level of amplified mRNA was normalized to mRNA expression of the housekeeping gene S. coelicolor hrdB, which was amplified as an internal control using primers hrdB-primer3 (5′-CGCACCATCCGTATCCC-3′) and hrdB-primer4 (5′- TCACCGAACTCGCTGTCG-3′).

    Well-based bioassay of CDA.

    An inoculum of 108 spores of S. coelicolor was plated on Oxoid Nutrient Agar (25 ml) and incubated at 30 °C for 2 to 7 days. The plates were frozen at −70 °C for 10 h and then slowly thawed at room temperature. The cells and medium were removed by centrifugation (12 857 g, 10 min) and the resulting supernatant was collected for the assay of CDA production. The CDA-sensitive Staphylococcus aureus was used as indicator strain. A 50 μl aliquot of an overnight culture of Staph. aureus was added into 25 ml melted soft nutrient agar (per litre: 13 g nutrient broth, 7 g Bactoagar; temperature about 45 °C) in the presence of added Ca(NO3)2 (16 mM) and poured into a plate. Agar plugs were cut from the dried plates, creating wells into which the collected supernatant was added. Calcium-dependent bioactivity was observed as a zone of inhibition after overnight incubation at 37 °C. The calcium-dependent bioactivity was assayed three times for each strain. The diameter of the inhibition zone was measured and used to evaluate the quantity of CDA production. Inhibition zones did not appear on control plates lacking Ca(NO3)2.

    RESULTS

    Identification of the hpdR promoter region

    For precise localization of the transcription start point of hpdR, high-resolution S1 mapping was performed using RNA isolated from S. coelicolor J1501 (see Methods). A single transcription start point (tsp) was identified as a G residue around 210 nt upstream of the translational start of hpdR (Fig. 1⇓). This tsp is preceded by the typical −35 and −10 hexamers, TTGATA and ATTCTG, in the upstream region.

    Figure image not available in archive
    Fig. 1.

    Determination of the transcription start point (tsp) of hpdR. (a) High-resolution S1 nuclease mapping of hpdR in S. coelicolor J1501. The tsp is indicated by an asterisk and an arrow. Lanes G, A T, and C are a dideoxy sequencing ladder generated using the same oligonucleotide as was used to make the S1 mapping probe. (b) Sequence of the hpdR promoter region. The −35 and −10 regions are indicated by bold and underlined letters, the tsp is indicated by bent arrow, the potential Shine–Dalgarno sequence (RBS) is underlined, and the potential translational initiation codon (GTG) is indicated by bold letters.

    Transcription of hpdR is developmentally regulated

    To study the transcription of hpdR, high-resolution S1 mapping was performed using RNA isolated from S. coelicolor J1501 grown on R2YE medium for different times. The transcription of hpdR could be readily detected throughout the life cycle, but with different levels at different time points. hpdR was transcribed at a higher level when the strain was growing as substrate mycelia and forming the primary aerial hyphae (at the 24 and 48 h time points, respectively), and then transcription decreased and remained at a relatively low level during the subsequent growth of aerial hyphae and sporulation (Fig. 2a⇓). These results indicated that hpdR was transcribed in an attenuated pattern during the life cycle in J1501. Therefore, its transcription may be developmentally regulated.

    Figure image not available in archive
    Fig. 2.

    S1 nuclease mapping of hpdR in the parent strain J1501 (a) and hpdRDMJ-P (b). The strains were incubated on solid R2YE medium. The incubation time points are indicated above each panel. hrdB, the major sigma factor of S. coelicolor, was used as a positive internal control.

    hpdR is subject to negative autoregulation

    To study whether hpdR is subject to autoregulation, like other members of the IclR family (Gui et al., 1996), the transcription analysis was applied to the hpdR promoter region in an hpdR disruption mutant (hpdRDMJ-P), which contains the intact promoter region of hpdR but lacks the complete coding region for HpdR. S1 mapping assay showed that the transcription of the hpdR promoter in hpdRDMJ-P was at nearly the same level as that of J1501 at 24 h, but it remained at the same level from 24 h to 120 h, whereas the transcription of hpdR in J1501 decreased after 48 h (Fig. 2⇑). These results indicated that the transcription of hpdR seems to be subtly autoregulated during the life cycle in J1501.

    HpdR specifically binds to the promoter region of its own gene

    To assess whether the autoregulation of hpdR was direct or not, an EMSA was performed. A 518 bp DNA fragment containing the promoter of hpdR was labelled and used as a probe. The EMSA showed that HpdR specifically bound to the radiolabelled fragment and formed different DNA–HpdR complexes at low or high concentrations of protein (Fig. 3⇓). The retardation was inhibited by the addition of a 30-fold excess of unlabelled hpdR promoter DNA fragment; additionally, an equal amount of GST protein had no binding activity to the same DNA fragment (data not shown). These results suggested that HpdR in S. coelicolor can specifically bind to its own promoter region.

    Figure image not available in archive
    Fig. 3.

    EMSA of HpdR binding to the hpdR promoter region (PhpdR). Radioactively labelled PhpdR DNA (5 nM in each lane) was incubated with increasing amounts of GST–HpdR (see Methods). Lane 1, free labelled DNA probe (518 bp) in the binding buffer. Lanes 2–7, labelled probe was incubated with 0.1, 0.2, 0.3, 0.4, 0.5 and 0.6 μM GST–HpdR, respectively. Lane 8, labelled probe was incubated with 0.6 μM GST–HpdR and a 30-fold excess of unlabelled probe.

    To determine the shortest fragment bound by HpdR, several DNA fragments were used in the EMSA experiments. A 159 bp probe (−125 to +34 relative to the tsp), containing the −35, −10 and tsp regions of the hpdR promoter, was bound by HpdR and exhibited a similar profile (data not shown). This result indicated that the binding region of HpdR was located in the 159 bp DNA fragment.

    hpdR disruption leads to decreased CDA production

    hpdR was disrupted in S. coelicolor J1501 via double-crossover (Fig. 4a, b⇓) and the resulting disruption mutant was designated hpdRDMJ. This mutant produced 10–30 % of the amount of CDA produced by J1501 (Fig. 4c⇓), suggesting that hpdR may positively regulate CDA biosynthesis. The phenotype of hpdRDMJ could be complemented by introducing a wild-type copy of hpdR on the integrating vector pSET152, and therefore was attributed to functional inactivation of hpdR rather than polar effects on the transcription of downstream genes or mutations in other loci.

    Figure image not available in archive
    Fig. 4.

    CDA production by hpdRDMJ. (a) Construction of hpdRDMJ and hpdRCS. The DNA fragment containing the promoter and HTH coding region of hpdR was replaced by a kanamycin-resistance gene (aphII) in J1501 to generate the hpdR disruption mutant hpdRDMJ. A 1.4 kb KpnI fragment containing the promoter region and coding sequence of hpdR was blunted and inserted into the EcoRV site of pSET152 to generate pSET152 : : hpdR, which was used to complement hpdRDMJ. (b) Southern hybridization analysis of KpnI-digested chromosomal DNA from wild-type (lane 1), the disruption mutant (lanes 2–3; duplicates) and the complemented strain (lanes 4–5; duplicates) to confirm the integration via homologous recombination. The DIG-labelled 0.3 kb DNA fragment containing the internal region of hpdR was used as a probe. The 1.4 kb signal (I) in the wild-type strain (J1501) is from a KpnI fragment corresponding to the complete hpdR gene. Strain hpdRDM shows a 2.0 kb signal (II) corresponding to a fragment with part of the hpdR gene and the 1.0 kb kanamycin-resistance gene. The complemented strain contains the 1.4 kb complete hpdR that was inserted into the chromosomal φC31 attB site of hpdRDM by pSET152 mediation. (c) Assay for CDA production by J1501 (A), hpdRDMJ (B) and hpdRCS (C) in the presence or absence of Ca(NO3)2.

    HpdR positively controls CDA biosynthesis via directly targeting hmaS

    The positive effect of hpdR on CDA production led us to speculate that hpdR may control CDA biosynthesis via directly regulating the biosynthetic gene(s) involved. To find the potential targets regulated by HpdR in the CDA biosynthetic gene cluster, the genes and their upstream regulatory regions in this cluster were analysed. The sequence alignment showed that the upstream regulatory region of hmaS, which encodes an essential enzyme, HmaS (SCO3229), involved in CDA biosynthesis (Hojati et al., 2002), has 55 % identity with the HpdR-bound region upstream of hppD (Fig. 5a⇓). Both HmaS and HppD are non-haem, α-keto-acid-dependent iron oxygenases; they catalyse the conversion of 4-hydroxyphenylpyruvate (4HPP) to different products, 4-hydroxymandelate and homogentisate, respectively (Denoya, et al., 1994; Hojati et al., 2002; Gunsior et al., 2004). Therefore, hmaS may be a potential target of HpdR.

    Figure image not available in archive
    Fig. 5.

    hmaS is a potential target of HpdR. (a) Comparison of the upstream regulatory regions of hmaS and hppD in S. coelicolor. The upstream region of hppD (PhppD) contains the complete HpdR-binding site, indicated by the overhead line. The boxed sequences indicate the −35 and −10 regions of the hppD promoter. (b) Specific binding of HpdR to the promoter region of hmaS. Lane 1, free labelled PhmaS DNA (328 bp) in the binding buffer. Lanes 2–4, the labelled PhmaS fragment was assayed in the presence of 0.1, 0.3 and 0.9 μM purified HpdR–GST. Lane 5, labelled PhmaS was assayed in the presence of 0.9 μM HpdR–GST and 30-fold excess of unlabelled PhmaS (as competitor DNA). Lane 6, labelled PhmaS DNA was assayed in the presence of 0.9 μM GST. (c) Transcription assay of hmaS in J1501 and hpdRDM at 72 h (left), and bioassay of CDA production by J1501, hmaSDM and hpdRDM (right). The qRT-PCR assays were performed as described in Methods; transcription is shown in arbitrary units.

    To investigate this possibility, an EMSA was first performed. A 328 bp DNA fragment containing the intergenic region between hmaS and its upstream gene was used as a probe. HpdR showed specific binding with this probe (Fig. 5b⇑), suggesting that hmaS may be regulated by HpdR in a direct manner. To further identify the role of HpdR in regulating hmaS, the transcriptional level of hmaS in both wild-type J1501 and hpdRDMJ was studied by real-time PCR. The results showed that the transcription of hmaS in J1501 was activated by HpdR at 72 h (Fig. 5c⇑), which is consistent with the production of CDA; no CDA production was detectable at 24 or 48 h (data not shown). The transcriptional level of hmaS in hpdRDM was decreased compared to that in J1501 (Fig. 5c⇑), which is also consistent with the effects on CDA production. These results imply that the transcription of hmaS is partially controlled by hpdR.

    When hmaS was disrupted in J1501 via double crossover (Fig. 6a⇓), the production of CDA was completely abolished in the resulting disruption mutant, hmaSDM (Fig. 6b⇓); this was similar to the phenotype of an hmaS-disrupted mutant of S. coelicolor MT1110 (Hojati et al., 2002). All the above results (Figs 5⇑ and 6⇓) showed that hmaS, an essential gene in CDA biosynthesis, is a novel and direct target of HpdR.

    Figure image not available in archive
    Fig. 6.

    Disruption of hmaS abolishes CDA biosynthesis in J1501. (a) Southern hybridization analysis of NcoI/BglII-digested chromosomal DNA from wild-type strain J1501 (lane 1) and the hmaS disruption strain hmaSDM (lanes 2–5; samples from four colonies) to confirm correct integration by homologous recombination. A DIG-labelled 0.4 kb DNA fragment containing the upstream region of hmaS was used as a probe. The 1.38 kb band in the wild-type strain represents the NcoI–BglII fragment. The hmaS disruption mutant (hmaSDM) shows a 2.2 kb signal corresponding to the remaining partial hmaS and the inserted 1.7 kb hygromycin-resistance gene. (b) CDA production assay of J1501and hmaSDM after growth at 28 °C for 5 days on Nutrient Agar. The CDA-sensitive Staph. aureus was used as indicator strain.

    DISCUSSION

    In S. coelicolor J1501, the transcription level of hpdR was different at the early and late developmental stages (Fig. 2⇑), indicating that its transcription may be related to the developmental and differentiational regulation of S. coelicolor J1501. This strain showed substrate mycelium growth and growth of primary aerial hyphae at 24 h and 48 h, respectively, and showed abundant aerial hyphae formation and sporulation at the later time points. Our previous study showed that the disruption of hpdR resulted in a white phenotype in J1501, suggesting an important role of hpdR in the morphological differentiation of this strain (Tan et al., 2002). The target genes regulated by HpdR and involved in the morphological differentiation of J1501 will be determined in future work.

    Our results showed that hpdR is subject to a fine-tuning autoregulation. The transcription level of the hpdR promoter was almost the same at 24–48 h in either the presence or absence of HpdR protein, but it was greatly increased after 48 h without HpdR. Therefore, it is deduced that hpdR may start the autoregulation from 48 h in response to some developmental signal(s). Our previous study revealed that 4HPP is a specific signal from tyrosine catabolism, which can affect the binding of HpdR to the target promoter region of hppD (Yang et al., 2007), but whether it also serves as the signal to regulate the autoregulation of HpdR remains unclear.

    4HPP has been proved to be an intermediate in both the catabolic pathway of tyrosine and the biosynthetic pathway of CDA. In the main catabolic pathway of tyrosine, HppD catalyses the conversion of 4HPP to homogentisate (Brownlee et al., 2004; Johnson-Winters et al., 2003). In the CDA biosynthetic pathway, HmaS catalyses the conversion of 4HPP to l-4-hydroxymandelic acid (Hojati et al., 2002). HpdR negatively regulates the transcription of hppD and positively regulates the transcription of hmaS, implying that this protein plays a significant regulatory role in both tyrosine catabolism and CDA biosynthesis. The repression of hppD by HpdR keeps HppD at a constant low level, which prevents the overconversion of 4HPP to homogentisate. HpdR also activates the transcription of hmaS, thus accelerating the conversion of 4HPP to l-4-hydroxymandelic acid and the final steps of CDA biosynthesis. Therefore, HpdR possibly co-regulates the transcription of genes involved in a primary catabolic pathway (tyrosine catabolism) and a secondary metabolic pathway (CDA biosynthesis). This co-regulation would lead to the inhibition of 4HPP being converted to homogentisate, and enhanced CDA biosynthesis.

    Our results have shown that HpdR exhibits different regulatory functions in S. coelicolor. It might act as a repressor for hppD and hpdA (Yang et al., 2007) or an activator for hmaS. hpdR appears to play a significant role in a hierarchical regulatory network. The HpdR-bound DNA sequence upstream of hpdR has comparatively low identity to the upstream regulatory regions of hppD and hmaS, suggesting that HpdR-bound motifs are not very well conserved. This also implies that HpdR may bind to other regulatory regions and regulate the transcription of the corresponding genes. Another IclR-type regulator, NadR, was proved to be involved in amino-acid-dependent growth, quorum sensing and antibiotic production in S. coelicolor (Yang et al., 2009). Other studies have shown that IclR-type regulators usually play multiple regulatory roles in different biological pathways (see Introduction). Further work is needed to determine whether HpdR controls other target genes in S. coelicolor or hpdR is regulated by other unknown regulators.

    Acknowledgments

    This work was supported by grants from the National Natural Science Foundation of China (Grant no. 30871346), the Ministry of Science and Technology of China (Grant nos 31030242 and 2009CB118905) and the Chinese Academy of Sciences. We thank Professor Keith Chater (John Innes Centre, Norwich, UK) for kindly providing E. coli ET12567, and plasmids pKC1132 and pSET152. We thank Dr Brenda Leskiw (University of Alberta, Canada) for the gift of apramycin.

    References

    1. Altschul, S. F., Gish, W., Myers, E. W. & Lipman, D. J. (1990). Basic local alignment search tool. J Mol Biol 215, 403–410.
    2. Bierman, M., Logan, R., O'Brien, K., Seno, E. T., Rao, R. N. & Schoner, B. E. (1992). Plasmid cloning vectors for the conjugal transfer of DNA from Escherichia coli to Streptomyces spp. Gene 116, 43–49.
    3. Brownlee, J. M., Johnson-Winters, K., Harrison, D. H. & Moran, G. R. (2004). Structure of the ferrous form of (4-hydroxyphenol) pyruvate dioxygenase from Streptomyces avermitilis in complex with the therapeutic herbicide, NTBC. Biochemistry 43, 6370–6377.
    4. Chater, K. F. (1993). Genetics of differentiation in Streptomyces. Annu Rev Microbiol 47, 685–713.
    5. Chater, K. F. & Chandra, G. (2008). The use of the rare UUA codon to define “expression space” for genes involved in secondary metabolism, development and environmental adaptation in streptomyces. J Microbiol 46, 1–11.
    6. Chater, K. F., Bruton, C. J., King, A. A. & Suarez, J. E. (1982). The expression of Streptomyces and Escherichia coli drug resistance determinants cloned into the Streptomyces phage PC31. Gene 19, 21–32.
    7. Denoya, C. D., Skinner, D. D. & Morgenstern, M. R. (1994). A Streptomyces avermitilis gene encoding a 4-hydroxyphenylpyruvic acid dioxygenase-like protein that directs the production of homogentistic acid and ochronotic pigment in Escherichia coli. J Bacteriol 176, 5312–5319.
    8. Gui, L., Sunnarborg, A., Pan, B. & LaPorte, D. (1996). Autoregulation of iclR, the gene encoding the repressor of the glyoxylate bypass operon. J Bacteriol 178, 321–324.
    9. Gunsior, M., Ravel, J., Challis, G. L. & Townsend, C. A. (2004). Engineering 4-hydroxyphenolpyruvate dioxygenase to a p-hydroxymandelate synthesase and evidence for the proposed benzene oxide intermediate in homogentisate formation. Biochemistry 43, 663–674.
    10. Hojati, Z., Milne, C., Harvey, B., Gordon, L., Borg, M., Flett, F., Wilkinson, B., Sidebottom, P. J., Rudd, B. A. M. & other authors (2002). Structure, biosynthetic origin, and engineered biosynthesis of calcium-dependent antibiotics from Streptomyces coelicolor. Chem Biol 9, 1175–1187.
    11. Johnson-Winters, K., Purpero, V. M., Kavana, M., Nelson, T. & Moran, G. R. (2003). (4-Hydroxyphenol) pyruvate dioxygenase from Streptomyces avermitilis: the basis for ordered substrate addition. Biochemistry 42, 2072–2080.
    12. Kieser, T., Bibb, M. J., Buttner, M. J., Chater, K. F. & Hopwood, D. A. (2000). Practical Streptomyces Genetics. Norwich. : John Innes Foundation.
    13. Kuhstoss, S. & Rao, R. N. (1991). Analysis of the integration function of the streptomycete bacteriophage φC31. J Mol Biol 222, 897–908.
    14. Li, W., Ying, X., Guo, Y., Yu, Z., Zhou, X., Deng, Z., Kieser, H., Chater, K. F. & Tao, M. (2006). Identification of a gene negatively affecting antibiotic production and morphological differentiation in Streptomyces coelicolor A3(2). J Bacteriol 188, 8368–8375.
    15. Li, R., Xie, Z., Tian, Y., Yang, H., Chen, W., You, D., Liu, G., Deng, Z. & Tan, H. (2009). polR, a pathway-specific transcriptional regulatory gene, positively controls polyoxin biosynthesis in Streptomyces cacaoi var. asoensis. Microbiology 155, 1819–1831.
    16. Lipman, D. J. & Pearson, W. R. (1985). Rapid and sensitive protein similarity searches. Science 227, 1435–1441.
    17. Liu, G., Tian, Y., Yang, H. & Tan, H. (2005). A pathway-specific transcription regulational regulatory gene for nikkomycin biosynthesis in Streptomyces ansochromogenes that also influences colony development. Mol Microbiol 55, 1855–1866.
    18. MacNeil, D. J., Gewain, K. M., Ruby, C. L., Dezeny, G., Gibbons, P. H. & MacNeil, T. (1992). Analysis of Streptomyces avermitilis genes required for avermectin biosynthesis utilising a novel integrating vector. Gene 111, 61–68.
    19. Sambrook, J., Fritsch, E. F. & Maniatis, T. (1989). Molecular Cloning: a Laboratory Manual, 3rd edn. Cold Spring Harbor, NY. : Cold Spring Harbor Labortory.
    20. Santamarta, I., López-García, M. T., Pérez-Redondo, R., Koekman, B., Martín, J. F. & Liras, P. (2007). Connecting primary and secondary metabolism: AreB, an IclR-like protein, binds the ARE (ccaR) sequence of S. clavuligerus and modulates leucine biosynthesis and cephamycin C and clavulanic acid production. Mol Microbiol 66, 511–524.
    21. Tan, H., Tian, Y., Yang, H., Liu, G. & Nie, L. (2002). A novel Streptomyces gene, samR, with different effects on differentiation of Streptomyces ansochromogenes and Streptomyces coelicolor. Arch Microbiol 177, 274–278.
    22. Tian, Y., Fowler, K., Findlay, K., Tan, H. & Chater, K. (2007). An unusual response regulator influences sporulation at early and late stages in Streptomyces coelicolor. J Bacteriol 189, 2873–2885.
    23. Traag, B. A., Kelemen, G. H. & van Wezel, G. P. (2004). Transcription of the sporulation gene ssgA is activated by the IclR-type regulator SsgR in a whi-independent manner in Streptomyces coelicolor A3(2). Mol Microbiol 53, 985–1000.
    24. Wang, L., Tian, X., Wang, J., Yang, H., Fan, K., Xu, G., Yang, K. & Tan, H. (2009). Autoregulation of antibiotic biosynthesis by binding of the end product to an atypical response regulator. Proc Natl Acad Sci U S A 106, 8617–8622.
    25. Yang, H., Wang, L., Xie, Z., Tian, Y., Liu, G. & Tan, H. (2007). The tyrosine degradation gene hppD is transcriptionally activated by HpdA and repressed by HpdR in Streptomyces coelicolor, while hpdA is negatively autoregulated and repressed by HpdR. Mol Microbiol 65, 1064–1077.
    26. Yang, Y. H., Song, E., Kim, E. J., Lee, K., Kim, W. S., Park, S. S., Hahn, J. S. & Kim, B. G. (2009). NadR, an IclR-like regulator involved in amino-acid-dependent growth, quorum sensing, and antibiotic production in Streptomyces coelicolor. Appl Microbiol Biotechnol 82, 501–511.