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Journal of Bacteriology, May 2004, p. 2567-2575, Vol. 186, No. 9
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.9.2567-2575.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Identification of PimR as a Positive Regulator of Pimaricin Biosynthesis in Streptomyces natalensis

Nuria Antón,1 Marta V. Mendes,1 Juan F. Martín,1,2 and Jesús F. Aparicio1,2*

Institute of Biotechnology INBIOTEC, Parque Científico de León, 24006 León,1 Area of Microbiology, Faculty of Biology, University of León, 24071 León, Spain2

Received 4 December 2003/ Accepted 15 January 2004


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Sequencing of the DNA region on the left fringe of the pimaricin gene cluster revealed the presence of a 3.6-kb gene, pimR, whose deduced product (1,198 amino acid residues) was found to have amino acid sequence homology with bacterial regulatory proteins. Database comparisons revealed that PimR represents the archetype of a new class of regulators, combining a Streptomyces antibiotic regulatory protein (SARP)-like N-terminal section with a C-terminal half homologous to guanylate cyclases and large ATP-binding regulators of the LuxR family. Gene replacement of pimR from Streptomyces natalensis chromosome results in a complete loss of pimaricin production, suggesting that PimR is a positive regulator of pimaricin biosynthesis. Gene expression analysis by reverse transcriptase PCR (RT-PCR) of the pimaricin gene cluster revealed that S. natalensis {Delta}PimR shows no expression at all of the cholesterol oxidase-encoding gene pimE, and very low level transcription of the remaining genes of the cluster except for the mutant pimR gene, thus demonstrating that this regulator activates the transcription of all the genes belonging to the pimaricin gene cluster but not its own transcription.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Streptomycetes are filamentous soil bacteria that have a complex life cycle that involves differentiation and sporulation. These bacteria have attracted great interest due to their well-known ability to produce a great variety of antibiotics and other secondary metabolites. Production of these compounds is regulated in response to nutritional status alteration and a variety of environmental conditions and, hence, occurs in a growth-phase-dependent manner, at the beginning of the stationary growth phase, and is usually accompanied by morphological differentiation (10, 15).

Control of secondary metabolite production seems to be operating at several levels. The highest levels include genes that exert a pleiotropic control over one or more aspects of secondary metabolism, such as antibiotic production or morphological differentiation (see reference 16 for a review). The lowest level, however, is composed by regulatory genes that only affect a single antibiotic biosynthetic pathway. These pathway-specific regulatory genes are usually found within the respective antibiotic biosynthesis gene cluster, a feature that has greatly facilitated their study.

The first pathway-specific regulatory proteins characterized in actinomycetes belonged to a protein family called SARPs (Streptomyces antibiotic regulatory proteins) (43). Members of this family are characterized by the presence of an OmpR-like DNA-binding domain (31) and include ActII-Orf4 of the actinorhodin pathway (5), DnrI of the daunorubicin gene cluster (28), RedD of the undecylprodigiosin pathway (33), CcaR of the cephamycin and clavulanic acid pathways (36), and TylS and TylT from the tylosin pathway (8), among others.

In recent years, a novel family of transcriptional regulators has been identified (20). This new family, typified by the regulator of the maltose regulon in Escherichia coli, MalT (11), and called large ATP-binding regulators of the LuxR family (LAL) is characterized by the unusual size of its members, the presence of an N-terminal ATP/GTP-binding domain easily identified by the presence of the conserved Walker A motif (42), and a C-terminal LuxR-like DNA-binding domain characterized by a conserved helix-turn-helix motif. Several regulators of the LAL family have been identified in antibiotic gene clusters from actinomycetes, including PikD from the pikromycin pathway in Streptomyces venezuelae (44), RapH from the rapamycin pathway in Streptomyces hygroscopicus (1, 32), and NysRI and NysRIII from the nystatin pathway in Streptomyces noursei (12).

Pimaricin is a 26-member tetraene macrolide antifungal antibiotic produced by Streptomyces natalensis, which is widely used for the treatment of fungal keratitis and also in the food industry to prevent mold contamination of cheese and other nonsterile foods (i.e., cured meat, sausages, and ham, etc.). As a polyene, its antifungal activity lies in its interaction with membrane sterols, thus causing the alteration of membrane structure and leading to the leakage of cellular materials. As other macrocyclic polyketides, pimaricin is synthesized by the action of so-called type I modular polyketide synthases (4). Our laboratory has previously sequenced the pimaricin biosynthetic gene cluster completely and demonstrated its identity by gene disruption experiments (2, 3). The gene cluster encodes 13 polyketide synthase modules within five multifunctional enzymes and 12 additional proteins that presumably govern modification of the polyketide skeleton, export, and regulation of gene expression (Fig. 1) (3, 4).



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  		FIG. 1. Pimaricin biosynthetic gene cluster. The left fringe of the gene cluster is indicated in more detail and includes pimR (in dark grey), the glycosyl transferase-encoding open reading frame (pimK) thought to be involved in the attachment of the mycosamine moiety during pimaricin biosynthesis, and the last two polyketide synthase (PKS) genes (pimS3 and pimS4) involved in the construction of the pimaricinolide (3). Pointed boxes indicate directions of transcription. All NotI (N) sites are shown; the SalI (S) and NruI (Nr) sites involved in vector construction are also indicated.

 
In this paper we describe the cloning, sequencing, and detailed characterization of a novel class of pathway-specific regulators in S. natalensis and demonstrate its role as a transcriptional activator for pimaricin biosynthesis in this bacterium.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Bacterial strains, cloning vectors, and cultivation. S. natalensis ATCC 27448 was routinely grown in YEME medium (25) without sucrose. Sporulation was achieved in TBO medium (2). For pimaricin production, the strain was grown in YEME without sucrose. The same media were supplemented with thiostrepton when used for S. natalensis 6D82 growth and/or metabolite production. E. coli strain XL1-Blue MR (Stratagene) was used as a host for plasmid subcloning in plasmids pBluescript (Stratagene), pUC18, and pUC19. Candida utilis (synonym, Pichia jadinii) CECT 1061 was used for bioassay experiments. Phage KC515 (c+ attP::tsr::vph), a {phi}C31-derived phage (38), was used for gene disruption experiments. Streptomyces lividans JII 1326 (17) served as a host for phage propagation and transfection. Infection with {phi}6D8 (the KC515 recombinant derivative used for gene replacement) was carried out on R5 medium (25). Standard conditions for culture of Streptomyces species and isolation of phages were as described by Kieser et al. (25).

Genetic procedures. Standard genetic techniques with E. coli and in vitro DNA manipulations were as described by Sambrook and Russell (39). Recombinant DNA techniques in Streptomyces species and isolation of Streptomyces total and phage DNA were performed as previously described (25). Southern hybridization was carried out with probes labeled with digoxigenin by using the digoxigenin DNA labeling kit (Roche Biochemicals).

DNA sequencing and analysis. Sequencing templates were obtained by random subcloning of fragments generated by controlled partial HaeIII digestions. DNA sequencing was accomplished by the dideoxynucleotide chain termination method (40) with the Perkin Elmer Amplitaq dye-terminator sequencing system on double-stranded DNA templates with an Applied Biosystems model 310 sequencer (Foster City, Calif.). Each nucleotide was sequenced a minimum of three times on both strands. Alignment of sequence contigs was performed by the DNAStar program Seqman (Madison, Wis.). DNA and protein sequences were analyzed with the EBI worldwide web BLAST and InterProScan servers as well as with the CNRS Prodom Protein Domain Database server.

Construction of a pimR mutant. A 10,636-bp NotI fragment encompassing the entire pimR gene, pimK, and part of the pimS4 gene (Fig. 1) was cloned into a NotI-cut pBluescript vector to yield pNAF0. This plasmid was then digested with SalI and religated to yield pNAF1, which was used as a source of DNA for both sequencing of pimR and for obtaining the DNA fragment used for gene replacement.

The pimR gene was disrupted by KC515 phage-mediated gene replacement as follows. Plasmid pNAF1 was digested with NruI and religated to yield pNAF2. This treatment eliminates a 455-bp NruI fragment that codes for an internal piece of PimR that contains an ATP/GTP-binding site (42) (see below) and results in a mutant pimR gene with a frameshift beyond the new NruI site. A 2-kb SacI-BglII fragment internal to the mutant pimR sequence was cloned into the SacI-BamHI sites of KC515 (38). Transfection of S. lividans protoplasts (25) resulted in a number of phage plaques that were screened by Southern hybridization for the presence of pimR-derived sequences. One of the recombinants, {phi}6D8, was selected and used to infect S. natalensis, thus allowing the selection for lysogen formation. Lysogens were selected by thiostrepton resistance on R5 medium. Gene replacement was sought by repeated rounds of nonselective growth in liquid YEME medium without sucrose, and the loss of the phage was confirmed by genomic Southern hybridization.

Isolation of total RNA. S. natalensis ATCC 27448 and S. natalensis {Delta}pimR were grown for 48 h in YEME medium without sucrose (stationary phase of growth), the cultures were then mixed with 1 volume of 40% glycerol, and mycelia were harvested by centrifugation and immediately frozen by immersion in liquid nitrogen. Frozen mycelium was then broken by shearing in a mortar, and the frozen lysate was added to buffer RLT (Qiagen) in the presence of 1.5% ß-mercaptoethanol. RNeasy mini spin columns were used for RNA isolation according to the manufacturer's instructions. RNA preparations were treated with DNase I (Promega) to eliminate possible chromosomal DNA contamination.

Gene expression analysis by RT-PCR. Transcription was studied by using the SuperScript one-step reverse transcriptase PCR (RT-PCR) system with Platinum Taq DNA polymerase (Invitrogen), with 5 ng of total RNA as the template. Conditions were as follows: first strand cDNA synthesis, 45°C for 40 min followed by 94°C for 2 min; amplification, 28 or 33 cycles of 98°C for 15 s, 60 to 70°C (depending of the set of primers used) for 30 s, and 72°C for 1 min. Primers (18- to 24-mers) (Table 1) were designed to generate PCR products of approximately 500 bp. Negative controls were carried out with each set of primers and Platinum Taq DNA polymerase to confirm the absence of contaminating DNA in the RNA preparations. The identity of each amplified product was corroborated by direct sequencing with one of the primers.


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  		TABLE 1. Primers for RT-PCR

 
Assay of pimaricin production. To assay pimaricin in culture broths, 0.5 ml of culture was extracted with 4 ml of butanol, and the organic phase was diluted in water-saturated butanol to bring the absorbance at 319 nm into the range of 0.1 to 0.4 U. Solutions of pure pimaricin (DSM, Delft, The Netherlands) were used as controls. To confirm the identity of pimaricin, a UV-visible absorption spectrum (absorption peaks at 319, 304, 291, and 281 nm) was routinely determined on a Hitachi U-2001 spectrophotometer. The fungicidal activity of pimaricin was tested by bioassay with C. utilis CECT 1061 as the test organism. Quantitative determination of pimaricin was performed with a Waters 600 high-performance liquid chromatographer (HPLC) with a diode array UV detector set at 304 nm, fitted with a µBondapak RP-C18 column (10 µm; 3.9 by 300 mm). Elution was with a gradient (1 ml/min) of 100% methanol (methanol concentration: 50% from 0 to 3 min, up to 90% from 3 to 12 min, 90% from 12 to 20 min, down to 50% from 20 to 25 min, 50% from 25 to 30 min). The retention time for pimaricin was 14.5 min.

Cholesterol oxidase assay. Cholesterol oxidase activity was assessed by a coupled system with catalase (Roche Biochemicals) in which the cholesterol oxidase catalyzes the oxidation of cholesterol to 4-cholesten-3-one with the reduction of oxygen to hydrogen peroxide, which in the presence of catalase oxidizes methanol to formaldehyde. The latter, in the presence of NH4+, reacts with acetyl acetone to yield a yellow lutidine dye that is measured at 405 nm. The reaction mixture, containing 50 mM ammonium phosphate, 1.4 M methanol, 1.1 U of catalase/µl, 16 mM acetyl acetone, 0.1 µg of cholesterol/µl, and 1.4 U of commercial cholesterol oxidase/µl, was incubated at 37°C for 60 min. To assay cholesterol oxidase extracellular activity, culture broths of S. natalensis wild-ype or {Delta}pimR strains were substituted for the commercial cholesterol oxidase in the above reaction mixture.

Nucleotide sequence accession number. The sequence reported here has been deposited in the GenBank database under the accession number AJ585085.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Cloning of pimR. pimR was identified by genomic walking by using an S. natalensis ATCC 27448 cosmid library (2) and DNA segments from a previously unidentified open reading frame (orfX) upstream from pimK (which encodes a putative mycosamine transferase gene) on the left end of the pimaricin gene cluster (3). The gene was sequenced from plasmid pNAF1 (see Materials and Methods) and turned out to be separated by only 95 bp from the 5' end of pimK, with no obvious terminators between them (Fig. 1). The initiating ATG codon of pimR is preceded by the sequence GAGAGG, which could potentially act as a ribosomal binding site. pimR is 3,597 bp long, with an overall codon usage pattern in good agreement with that of typical Streptomyces genes; however, it contains a number of codons that are rare in such a G+C-rich organism. The presence of two TTA codons could be of particular interest, since their involvement in the regulation of differentiation and secondary metabolism in Streptomyces has been proposed (26).

The pimR gene product is a multidomain protein that partially resembles positive regulators of antibiotic biosynthetic pathways. Computer-assisted analysis of the pimR gene product (1,198 amino acids with an estimated Mr of 130,184) showed a very high sequence identity (86%) with the whole of protein PTER of Streptomyces avermitilis, a putative regulatory protein of 1,096 amino acid residues whose encoding gene was found within the pte gene cluster, which is involved in the biosynthesis of the pentane filipin (23, 34). PimR is some 100 amino acid residues larger than PTER, thus resembling a PTER-like protein with an extra trans-Reg-C domain (InterPro no. IPR001867) in its N terminus (Fig. 2A). This domain (amino acids 34 to 98) resembles the DNA-binding domain at the C terminus of the E. coli activator OmpR (29) and is usually found at the N termini of a family of antibiotic activators known as SARPs (43).



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  		FIG. 2. Domain structure and amino acid alignments of parts of the PimR protein. (A) Predicted domain structure of PimR. TRC, trans-reg-C domain (InterPro no. IPR001867) that resembles the DNA-binding domain of OmpR (see text); BAD, bacterial transcriptional activator domain (ProDom no. PD005851); AAA, domain characteristic of ATPases associated with diverse cellular activities (InterPro no. IPR003593). The asterisk indicates the location of the ATP/GTP-binding motif. Arrows indicate the regions of amino acid identity with proteins of the SARP, LAL, or CYC families (see text). CYC, guanylate cyclase. (B) Alignment of the N-terminal portion of PimR with different proteins of the SARP family. Numbers indicate amino acid residues from the N terminus of the protein. Conserved amino acids are shown highlighted. SAR, Streptomyces ambofaciens transcriptional activator (SWALL Q9KWX4); AknO, SARP of the aclacinomycins pathway in Streptomyces galilaeus (SWALL Q9L4V0); Gra-Orf9, activator from the granaticin pathway in Streptomyces violaceoruber (SWALL Q9ZA48); TylS, positive regulator of tylosin biosynthesis in Streptomyces fradiae (SWALL Q9XCC4); ActII-Orf4, actinorhodin activator in S. coelicolor A3(2) (SWALL P46106). (C) Comparison of the Walker A (left) and B (right) motifs of PimR with those of other proteins. Numbers indicate amino acid residues from the N terminus of the protein. Highly conserved amino acids are shown highlighted. Sc8d9.18, putative LAL regulatory protein from S. coelicolor A3(2) (SWALL Q9Z573); NysRI and NysRIII, LAL regulators of the nystatin pathway in S. noursei (SWALL Q9L4W1 and Q9L4V9); Sma0464 and Sma1591, putative guanylate cyclases encoded by plasmid pSymA of S. meliloti (SWALL Q930F6 and Q92YL0).

 
No end-to-end counterparts were found in the protein databases, however PimR showed significant sequence similarity with several pathway specific regulators in its N-terminal 300 amino acid residues (Fig. 2B). The highest scores were with the S. ambofaciens transcriptional activator SAR (40.3% identity) (18); AknO from the aclacinomycins pathway in S. galilaeus (40.2% identity) (37); Gra-orf9 from the granaticin pathway in S. violaceoruber (39.7% identity) (22); TylT and TylS from the tylosin pathway in S. fradiae (39% and 37.7% identity, respectively) (7), and ActII-orf4 from the actinorhodin pathway in S. coelicolor (36.5% identity) (21). All antibiotic activators described above belong to the family of Streptomyces antibiotic regulatory proteins (SARPs) that appear to turn on the expression of at least some of the genes of their respective clusters, thus controlling antibiotic production. The members of this family (typically between 277 and 666 amino acids) display two conserved amino acid regions (Fig. 2B), one at the N terminus with a proposed function in DNA binding (Trans-Reg-C domain; see above), and the second one at the C-terminal half of the protein. This later hypothetical domain is known as bacterial transcriptional activator domain (BAD; ProDom PD005851), and it is also found in several transcriptional activators of Mycobacterium.

C-terminal of this region, PimR displays an AAA domain (InterPro no. IPR003593) that spans to complete the N-terminal half of the protein (amino acids 439 to 627) (Fig. 2A). This domain is characteristic of a large family of ATPases associated with diverse cellular activities, including cell cycle regulation, protein degradation, and protein transport, among others (35), and contains the ATP/GTP-binding or Walker A motif A/G-X4-G-K-S/T (amino acids 447 to 454) (Fig. 2C) (42).

Comparison of database proteins with a stretch of 450 amino acid residues around the AAA domain (amino acids 350 to 800) yielded significant similarity with several proteins belonging to the LAL family of regulators (20), including various putative transcriptional activators from Streptomyces coelicolor (products of the genes sc5f8.05c, sc8d9.18, and sc2g5.14c) (9) or the NysRIII and NysRI regulators of nystatin biosynthesis in S. noursei (12). The identity is restricted to the N-terminal 400 amino acids of each of these proteins and therefore does not cover the characteristic C-terminal helix-turn-helix motif for DNA binding. Along with LAL regulators, BlastP homology searches identified several guanylate cyclases, including Sma1591, Sma0464, and Sma1789 encoded by the megaplasmid pSymA from Sinorhizobium meliloti (6) or Mll0576 from Mesorhizobium loti (24), although lacking the signature sequence for these proteins (Prosite no. PDOC00425), which is usually located at the N terminus. The identity in this case extends up to the C terminus of PimR (around 25% identity with the whole of the cyclase except for the N-terminal 200 amino acid residues).

Gene replacement of pimR. Since S. natalensis ATCC 27448 has so far proved absolutely resistant to transformation by conventional procedures, we took advantage of the ability of phage KC515, an attP-defective {phi}C31 derivative (38), to infect S. natalensis to introduce DNA into this strain. The recombinant phage used for pimR inactivation, {phi}6D8, was constructed as described in Materials and Methods and used to infect S. natalensis to obtain lysogens. Because phage KC515 and its derivative lack attP, they can only form lysogens by homologous recombination into the chromosome (Fig. 3A).



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  		FIG. 3. Gene replacement of pimR. (A) Predicted restriction enzyme polymorphism caused by gene replacement. The first crossover event is also indicated. The KpnI-PvuII restriction pattern before and after replacement is shown. The probe is indicated by thick lines. The fragment used for gene disruption derives from pNAF2 (see text for details). B, BamHI; K, KpnI; N, NruI; P, PvuII; S, SacI; Sm, SmaI; Z, BglII; wt, wild type. (B) Southern hybridization of the KpnI-PvuII-digested chromosomal DNA of the wild-type (lane 1), 6D82 (lane 2), and {Delta}pimR (lane 3) strains.

 
Five lysogens of S. natalensis were obtained by selection for thiostrepton resistance and tested for the lack of pimaricin production, since phage integration would disrupt the pimR transcript. One of these mutants was randomly selected and named S. natalensis 6D82. The identity of the mutant was confirmed by Southern hybridization (Fig. 3B). This mutant was then used to isolate thiostrepton-sensitive derivatives that had undergone a second recombination event deleting the integrated phage. These thiostrepton-sensitive isolates were obtained after 6 rounds of nonselective growth in YEME medium. Of the nine colonies isolated, six were found to have reverted to the wild type while the other three harbored the desired change.

One of these mutants, where pimR had been replaced by a mutated version of it lacking the ATP-binding site, was randomly selected and named S. natalensis {Delta}pimR. Chromosomal DNAs isolated from S. natalensis ATCC 27448 and mutant {Delta}pimR and digested with both KpnI and PvuII were probed with the 980-bp SmaI fragment used to construct the KC515 derivative utilized for gene replacement (see Materials and Methods). Hybridizing bands of 2.4 and 0.8 kb were found for the wild type as expected (Fig. 3B). However, in the disrupted mutant, a single 2.7-kb band was detected (Fig. 3B), indicating that a double crossover event had occurred. The observed hybridizing bands corresponded exactly to those expected according to the integration process depicted in Fig. 3. Figure 3 also shows the hybridizing bands found for S. natalensis 6D82.

The new strain S. natalensis {Delta}pimR had growth and morphological characteristics identical to those of the S. natalensis wild type when grown on solid or liquid media, indicating that PimR has no role in bacterial growth or differentiation.

Inactivation of pimR blocked pimaricin biosynthesis. The fermentation broth produced by the mutant strain generated by phage-mediated gene replacement, S. natalensis {Delta}pimR, was extracted with butanol and analyzed for the presence of pimaricin. Both the microbiological bioassay against C. utilis (data not shown) and the HPLC assays indicated that no pimaricin was being produced by the mutant strain {Delta}pimR (Fig. 4). This result, together with the significant similarity of PimR in its N-terminal section with well-known transcriptional activators, raised the question of which gene or genes were the potential target of PimR activity.



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  		FIG. 4. (A) Comparison of HPLC analyses of butanol-extracted broths from S. natalensis wild-type (top) and {Delta}pimR (bottom) strains. Detection was carried out at A304. (B) Typical chromophore UV-visible absorption spectrum of pimaricin and its structure.

 
Transcriptional control of pimaricin production. Total RNA was prepared from the S. natalensis wild type and mutant {Delta}pimR after growth for 48 h (when pimaricin is actively produced) (30) and used as a template for gene expression analysis by RT-PCR. Primers for RT-PCR were specific to sequences within pim genes (Table 1) and were designed to produce cDNAs of approximately 500 bp. A primer pair designed to amplify a cDNA of the lysA gene (encoding diaminopimelate decarboxylase) (Table 1) was used as an internal control. Transcripts were analyzed from the 17 genes of the pim cluster, including pimR, after 28 PCR cycles. In the case of pimR, transcripts were analyzed with pairs of primers located before and after the deletion. Whenever 28 cycles did not yield a product, analysis was repeated at 33 cycles. This analysis was carried out at least three times for each primer pair.

All 17 genes were transcribed at 48 h in the S. natalensis wild type; however, when we analyzed the transcription pattern in S. natalensis {Delta}pimR, we found virtually no transcripts for any of the genes of the cluster except for pimR (Fig. 5). No difference was observed between the use of primer pairs located before or after the deletion (Fig. 5A and B, respectively), thus indicating that transcription proceeded unabated across the site of the frameshift deletion in pimR. These results suggest that the gene replacement would have no polar effect on the transcription of pimK, which is located downstream from pimR. The lack of transcripts of pimK in S. natalensis {Delta}pimR must therefore be attributed to the absence of a functional PimR protein in this strain and implies that pimK is transcribed from a monocistronic transcript. Interestingly, the transcription pattern of the lysA gene was comparable in S. natalensis {Delta}pimR and in the parental strain (Fig. 5), thus validating the results described above.



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  		FIG. 5. Gene expression analysis of the pimaricin gene cluster by RT-PCR. Analysis was carried out on S. natalensis wild-type (+) and {Delta}pimR (–) strains as indicated in Materials and Methods. The identity of each amplified product was corroborated by direct sequencing. The absence of contaminating DNA in the RNA samples was assessed by PCR. Due to its small size, transcription of pimF was assessed by using a reverse primer (Table 1) located within the coding sequence of the gene located downstream from it (i.e., pimS0). Transcription of pimR or its mutated version was assessed with two pairs of primers (Table 1), one pair located before the deletion (A) and the second pair located after the deletion (B). Transcription of the lysA gene (encoding diaminopimelate decarboxylase) was also assessed as an internal control. A diagram with the organization of the genes within the pimaricin cluster and their putative transcripts is also included. The top panel shows the amplified products after 28 cycles of PCR, and the bottom panel shows the same analysis after 33 cycles to detect low-level transcripts.

 
Increasing the number of PCR cycles from 28 to 33 allowed the identification of transcripts for the remaining pim genes except for pimE (Fig. 5), thus suggesting that the mutant retains some transcription of the pim genes, albeit at very low levels. In any case, such low-level transcription was not sufficient for pimaricin production given that no pimaricin could be detected in the culture broth of S. natalensis {Delta}pimR.

The strict control of PimR on the transcription of pimE was totally unexpected for several reasons. First, due to the absence of apparent transcriptional terminators in the short intergenic regions between pimA, pimB, pimE, pimC, pimG, pimF, and pimS0, these genes were thought to form an operon resulting in a transcript of more than 13,600 bp (3) which could be controlled coordinately. Second, no obvious function could be predicted for PimE in pimaricin biosynthesis, given that PimE is a functional cholesterol oxidase (M. V. Mendes, N. Antón, J. F. Martín, and J. F. Aparicio, unpublished data). These results now could indicate that the above-indicated genes are actually transcribed from at least three different transcriptional units, namely pimAB, pimE, and pimCGFS0. However, in the absence of evidence indicating that all the transcripts are equally stable, it is also possible that the multicistronic transcript could be processed and subject to different rates of RNA degradation. Opposite to S. natalensis wild-type culture broths, no cholesterol oxidase activity could be detected in S. natalensis {Delta}pimR cultures (data not shown), thus confirming the lack of pimE transcription upon disruption of pimR. The precise role of PimE on pimaricin biosynthesis is intriguing and requires further investigation.


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Sequencing of the left-hand side of the pimaricin gene cluster revealed the presence of a gene, pimR, which could play a role as an activator for pimaricin biosynthesis in S. natalensis. Computer-assisted analysis of PimR revealed that it is a new multidomain protein, with an N-terminal region strikingly similar to transcriptional activators of the SARP family (5, 43) and a C-terminal region with significant similarity with guanylate cyclases and transcriptional regulators of the LAL family (large ATP-binding regulators of the LuxR family) (20, 44), including the ATP/GTP-binding AAA domain present in these protein families but lacking the characteristic signature sequence at the N terminus of guanylate cyclases or the LuxR-type helix-turn-helix motif for DNA binding present at the C terminus of LAL regulators.

The significant similarity of the N-terminal section of PimR with proteins of the SARP family raised two questions: first, whether PimR was actually a transcriptional activator, and second, whether, as an activator, it followed a regulatory behavior comparable to that of SARPs. The absence of pimaricin production upon disruption of the gene provided the first indication of the functional role of PimR as an activator of pimaricin biosynthesis, and the RT-PCR analysis revealed that it controls the expression of all of the pimaricin genes except its own transcription. SARPs activate transcription of their target genes by binding to heptameric repeats found around the –35 regions of cognate promoters. These sequences, corresponding to the consensus 5'-TCGAGCG-3', have been identified in the promoter regions of actinorhodin (43), daunorubicin (41), and mithramycin (27) biosynthetic gene clusters; however, when we examined the DNA sequence of the putative promoter regions of the pimaricin genes, no such sequences were identified, thus suggesting that PimR probably follows a regulatory pattern different from those of SARPs. In fact, the presence of distinct Walker A and B motifs implicated in ATP/GTP binding in PimR suggests that its transcriptional activating activity is dependent on ATP/GTP hydrolysis.

The domain composition of PimR suggests that pimR could have been originated by merging a SARP-like gene with a guanylate cyclase gene that has lost its 5'-end 600 nucleotides. Why would S. natalensis develop such a chimera for the regulation of pimaricin production is unclear, but it is tempting to speculate that the C-terminal region of PimR may act as a sensory transducer for the SARP-like region to bind DNA and activate transcription in an ATP-dependent manner. Both guanylate cyclases and the prototype of LAL regulatory proteins, the regulator of the maltose regulon in E. coli, MalT (11), interact with effectors to modulate their activities (19), and that could be the case for PimR.

The control of pimaricin biosynthesis exerted by PimR takes place through the specific transcriptional activation of all of the key enzyme-encoding genes for pimaricinolide construction. PimR either controls multiple pimaricin biosynthetic promoters or activates another hierarchical regulator(s). In any case, this supposes an important energy savings given that transcript synthesis is energy dependent, a fact that acquires special relevance for the giant mRNAs that are supposed to direct the synthesis of pimaricin (3).

The absence of pimE transcripts on S. natalensis {Delta}pimR compared to those of its surrounding genes was unexpected. The only indication for a functional role of pimE in pimaricin biosynthesis was its chromosomal location just in the middle of the pimaricin gene cluster (2, 3); however, the activity of its gene product as a functional cholesterol oxidase (Mendes et al., unpublished) suggested that such a location could be a matter of chance. Now, these results strongly suggest that PimE is actually involved in pimaricin biosynthesis and raise the question of what is the precise mechanism that drives the cellular role of this enzyme. Interestingly, a cholesterol oxidase-encoding gene, pteG, has been also identified within the gene cluster of the 26-membered polyene filipin produced by S. avermitilis (23, 34). Further experimental analyses (now in progress) will hopefully provide the answer to this question and to the generality of this phenomenon.

Besides that of pimaricin (2, 3), no other biosynthetic gene cluster for a small polyene has been characterized to date. The fact that, as yet, the only orthologue of PimR is PTER, a putative regulator of the pentane filipin produced by S. avermitilis (23, 34), suggests that common regulatory circuits could operate for the expression of these kind of molecules and that they are not shared by other polyenes. The absence of similar regulatory proteins in the recently characterized nystatin (12), amphotericin (13), or candicidin (14) gene clusters supports such a hypothesis. Further characterization of the PimR protein might prove useful for future applications to control and improve the expression of genes involved in the biosynthesis of small polyenes and thus increase their production.


   ACKNOWLEDGMENTS
 
This work was supported by a grant from the Comisión Interministerial de Ciencia y Tecnología (CICYT) to J.F.A. (BIO2001--0040). M.V.M. received a fellowship of the Fundação para a Ciência e a Tecnologia (PRAXIS XXI/BD/15850/98). N.A. was the recipient of an F.P.U. fellowship from Ministerio de Educación, Cultura, y Deporte (AP2002-1446).

We thank M. Driessen (DSM) and E. Recio for helpful discussions and J. Merino and B. Martín for excellent technical assistance.


   FOOTNOTES
 
* Corresponding author. Mailing address: Institute of Biotechnology INBIOTEC, Parque Científico de León, Avda. del Real, no. 1, 24006 León, Spain. Phone: 34-987-210308. Fax: 34-987-210388. E-mail: degjfa{at}unileon.es. Back


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Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Journal of Bacteriology, May 2004, p. 2567-2575, Vol. 186, No. 9
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.9.2567-2575.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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