Skip to main content
Antimicrobial Agents and Chemotherapy logoLink to Antimicrobial Agents and Chemotherapy
. 2000 Jul;44(7):1809–1817. doi: 10.1128/aac.44.7.1809-1817.2000

Genetic Localization and Molecular Characterization of the nonS Gene Required for Macrotetrolide Biosynthesis in Streptomyces griseus DSM40695

Wyatt C Smith 1, Longkuan Xiang 1, Ben Shen 1,*
PMCID: PMC89966  PMID: 10858335

Abstract

The macrotetrolides are a family of cyclic polyethers derived from tetramerization, in a stereospecific fashion, of the enantiomeric nonactic acid (NA) and its homologs. Isotope labeling experiments established that NA is of polyketide origin, and biochemical investigations demonstrated that 2-methyl-6,8-dihydroxynon-2E-enoic acid can be converted into NA by a cell-free preparation from Streptomyces lividans that expresses nonS. These results lead to the hypothesis that macrotetrolide biosynthesis involves a pair of enantiospecific polyketide pathways. In this work, a 55-kb contiguous DNA region was cloned from Streptomyces griseus DSM40695, a 6.3-kb fragment of which was sequenced to reveal five open reading frames, including the previously reported nonR and nonS genes. Inactivation of nonS in vivo completely abolished macrotetrolide production. Complementation of the nonS mutant by the expression of nonS in trans fully restored its macrotetrolide production ability, with a distribution of individual macrotetrolides similar to that for the wild-type producer. In contrast, fermentation of the nonS mutant in the presence of exogenous (±)-NA resulted in the production of nonactin, monactin, and dinactin but not in the production of trinactin and tetranactin. These results prove the direct involvement of nonS in macrotetrolide biosynthesis. The difference in macrotetrolide production between in vivo complementation of the nonS mutant by the plasmid-borne nonS gene and fermentation of the nonS mutant in the presence of exogenously added (±)-NA suggests that NonS catalyzes the formation of (−)-NA and its homologs, supporting the existence of a pair of enantiospecific polyketide pathways for macrotetrolide biosynthesis in S. griseus. The latter should provide a model that can be used to study the mechanism by which polyketide synthase controls stereochemistry during polyketide biosynthesis.


The macrotetrolides are a family of cyclic polyethers produced by a number of Streptomyces species (9, 16, 18). Nonactin (NON), the smallest homolog and a symmetric member of the family, was first isolated in 1955 (14). Its structure was initially deduced from spectroscopic analysis and was later confirmed by X-ray crystallography (15, 27), revealing that the intriguing molecular topology of NON consists of the (+)(−)(+)(−)-ester linkage of the enantiomeric nonactic acid (NA) building blocks. The other members of this family are homologs of NON. They contain homononactic acid (HNA) and/or bishomononactic acid (BNA) as building blocks, and they all have the same relative configuration (Fig. 1). NA-type building blocks have also been identified in several macrodiolides (24), including the pamamycins (34, 35).

FIG. 1.

FIG. 1

Structures of macrotetrolides (A) and their monomeric building blocks (B).

The macrotetrolides exert a broad spectrum of biological activities (56), ranging from antibacterial, antifungal, antitumor (11), antiprotozoan, antiparasitic, and insecticidal activities to immunosuppressive activities (12, 4749, 56). In fact, comparative studies on the immunosuppressive activities of tetranactin and cyclosporin, the latter being the most widely used immunosuppressant agent, showed that these two compounds were approximately equally effective and that tetranactin has the advantage of low toxicity (49). The biological activities of the macrotetrolides are generally traced to their ionophoric properties (30, 38, 46), and the potencies of these activities appear to parallel the size of the alkyl substituents of the macrotetrolides: tetranactin is often the most potent member of the family, while NON is generally inactive.

The biosynthesis of NON has been extensively studied by in vivo feeding experiments with 13C-, 2H-, and 18O-labeled precursors and biosynthetic intermediates (25, 13, 45) and by isolation of both enantiomers of NA and the dimeric NA (17). These results established unambiguously the polyketide origin of NON, the assembly of which from one molecule each of propionate and succinate and two molecules of acetate must have invoked (i) the rare use of succinate as an intact four-carbon fragment (C-3 to C-6) and (ii) the derivation of a three-carbon unit (C-7 to C-9) from two molecules of acetate (one of which is activated in the form of malonate) (Fig. 2). Feeding experiments with 13C- and 18O-doubly-labeled precursors indicated that the C-3-O bond is formed during closure of the tetrahydrofuran ring, presumably by an intramolecular Michael addition of the 6-hydroxy group onto the enone moiety of 2-methyl-6,8-dihydroxynon-2E-enoic acid (NEA) (3, 4). The involvement of the latter step in NON biosynthesis was further substantiated by the efficient and enantiospecific incorporation of both (6R,8R)-NEA into NON (45) and by the drastic reduction of NON production upon the addition of an NEA analog into the fermentation medium, which presumably acts as a suicide inhibitor for this enzymatic step (39). Sequential and stereospecific oligomerization of (+)- and (−)-NA into NON was supported by the isolation of both enantiomers of NA and the dimeric NA (17), which could be viewed as derailed biosynthetic intermediates from the enzyme complex. In vitro synthesis of macrotetrolides from NA and its homologs was also demonstrated in the presence of an enzyme preparation that appeared to be a heteromeric complex with an apparent molecular weight of 350,000 (2, 36).

FIG. 2.

FIG. 2

(A) A pair of enantiospecific polyketide pathways proposed for macrotetrolide biosynthesis in S. griseus. Arrows with broken lines indicate intermediates that have been either isolated or confirmed by feeding experiments. (B) Monomeric building block compositions of individual macrotetrolides.

Genetic analysis of NON biosynthesis was initiated by cloning the nonR resistance gene from Streptomyces griseus. By screening for tetranactin resistance in Streptomyces lividans TK64, Plater and Robinson (37) isolated several overlapping clones that harbored a total of 9 kb of S. griseus genomic DNA and determined the DNA sequence of a 3.3-kb fragment to reveal three complete open reading frames (ORFs), including nonR, and an incomplete ORF, orfX. Priestley and coworkers (53) subsequently completed the cloning and DNA sequence determination of orfX—naming it nonS—and characterized NonS as the nonactate synthase that catalyzes the formation of NA from NEA via an intramolecular Michael addition. However, the latter study fell short of shedding light on the stereochemistry of NA biosynthesis because the racemic N-octylcysteamine thioester of NEA was used as a substrate for NonS and the stereochemistry of the resulting NA was not determined. The direct involvement of nonS in NON biosynthesis remains to be established.

We view NON biosynthesis as an excellent model with which to study the mechanism by which polyketide synthase (PKS) controls the stereochemistry during polyketide chain assembly, based on the hypothesis that NON is derived from tetramerization, in a stereospecific fashion, of (+)- and (−)-NA. An intriguing corollary of the latter hypothesis is the stereospecific synthesis of two enantiomeric polyketide intermediates, such as NEA or its homologs [R = CH2CH3, CH(CH3)2; Fig. 2A], presumably by a pair of enantiospecific pathways. We report here on the inactivation of nonS in S. griseus DSM40695 to generate macrotetrolide-nonproducing mutants, in vivo complementation of the nonS mutants by the expression of nonS in trans, and fermentation of the nonS mutant in the presence of exogenous (±)-NA to restore macrotetrolide production. Our results prove the direct involvement of nonS in macrotetrolide biosynthesis, suggest that NonS catalyzes the formation of (−)-NA and its homologs, and support the existence of a pair of enantiospecific pathways that lead to (+)- and (−)-NA and their homologs, respectively, for macrotetrolide biosynthesis in S. griseus (Fig. 2A).

MATERIALS AND METHODS

Bacterial strains, plasmids, and biochemicals.

Escherichia coli DH5α (42), E. coli XL 1-Blue MR (Stratagene, La Jolla, Calif.), E. coli ET12567 (29), S. griseus DSM40695 (Deutsche Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany) (also known as Streptomyces subsp. griseus ETHA7796), and Micrococcus luteus ATCC 9431 (American Type Culture Collection, Rockville, Md.) were used in this work. pDH5 (20), pOJ446 (8), pWHM3 (50), and pWHM79 (43) were described previously. pUO9090 was a gift from M. C. Martin (Universidad de Oviedo, Oviedo, Spain), and other plasmids were from standard commercial sources. Ampicillin, apramycin, and NON were from Sigma (St. Louis, Mo.), and thiostrepton was from S. J. Lucania (Bristol-Myers Squibb, Princeton, N.J.). Racemic methyl nonactate was a gift from P. Metz (32).

Media and culture conditions.

E. coli strains carrying plasmids were grown in Luria-Bertani (LB) medium and were selected with appropriate antibiotics (42). S. griseus strains were grown on GMP (55) at 28°C for sporulation and in Trypticase soy broth (23) supplemented with 5 mM MgCl2 and 0.5% glycine at 28°C and 300 rpm (Incubator Shaker series 25, New Brunswick Scientific Co., Inc., Edison, N.J.) for isolation of genomic DNA. For protoplast preparation, S. griseus strains were grown in SGGP (54) supplemented with 0.5% glycine at 28°C and 300 rpm for 25 h; the resulting mycelia were washed with 0.3 M sucrose, resuspended in P buffer (23), and digested with lysozyme (1 mg/ml) at 37°C for 45 min. For regeneration, protoplasts were plated on R1M (55) and incubated at 28°C. For macrotetrolide production, S. griseus strains were cultured by a three-stage fermentation procedure described by Ashworth and Robinson (4).

DNA isolation and manipulation.

Plasmid preparation and DNA extraction were carried out by using commercial kits (Qiagen, Santa Clarita, Calif.). Genomic S. griseus DNA was isolated by standard protocols (23, 40). Restriction enzymes and other molecular biology reagents were from commercial sources, and digestion and ligation were done by standard methods (42). For Southern analysis, digoxigenin labeling of DNA probes, hybridization, and detection were performed by the protocols provided by the manufacturer (Boehringer Mannheim Biochemicals, Indianapolis, Ind.).

PCR primers were synthesized at the Protein Structure Laboratory, University of California, Davis. PCR was carried out on a Gene Amp PCR System 2400 (Perkin-Elmer/ABI, Foster City, Calif.) with Taq polymerase and buffer from Promega (Madison, Wis.). A typical PCR mixture consisted of 5 ng of S. griseus genomic or plasmid DNA as template, 25 pmol of each primer, 25 μM deoxynucleoside triphosphates, 5% dimethyl sulfoxide, 2 U of Taq polymerase, and 1× buffer in a final volume of 50 μl. The PCR temperature program was as follows: initial denaturation at 94°C for 5 min, 24 to 36 cycles of 45 s at 94°C, 1 min at 60°C, and 2 min at 72°C, followed by an additional 7 min at 72°C. To prepare the nonR probe by PCR, the following pair of primers were used: 5′-CACTGCCGACCGCTTCGG-3′ (forward primer) and 5′-CGTCGCTGAACCCGTCGG-3′ (reverse primer) (37). A distinctive product with the predicted size of 1.3 kb was amplified from S. griseus DSM40695 and was cloned into pGEM-T to yield pBS2001.

Automated DNA sequencing was carried out on an ABI Prism 377 DNA Sequencer (Perkin-Elmer/ABI), and this service was provided by either the DBS Automated DNA Sequencing Facility, University of California, Davis, or Davis Sequencing Inc. (Davis, Calif.). Data were analyzed by the ABI Prism Sequencing 2.1.1 software and a Genetics Computer Group (Madison, Wis.) program.

Genomic library construction and screening.

S. griseus genomic DNA was partially digested with MboI to yield a smear at about 60 kb, as monitored by electrophoresis on a 0.3% agarose gel (40). This sample was dephosphorylated upon treatment with shrimp alkaline phosphatase and was ligated into the E. coli-Streptomyces shuttle vector pOJ446 (8) that was prepared by digestion with HpaI, treatment with shrimp alkaline phosphatase, and additional digestion with BamHI. The resulting ligation mixture was packaged with the Gigapack II XL two-component packaging extract (Stratagene). The package mixture was transduced into E. coli XL 1-Blue MR. The transduced cells were spread onto LB plates that contained apramycin (100 μg/ml), and the plates were incubated at 37°C overnight. The titer of the primary library was approximately 4,500 CFU per μg of DNA. Restriction enzyme analysis of 12 randomly selected cosmids confirmed that the average size of the inserts was about 35 to 45 kb (40).

To screen the genomic library, colonies from four LB plates that contained apramycin (100 μg/ml; with approximately 600 colonies per plate) were transferred to nylon transfer membranes (Micro Separations, Inc., Westborough, Mass.) and were screened by colony hybridization with the PCR-amplified 1.3-kb nonR fragment from pBS2001 as a probe. The resulting cosmid clones were confirmed to be positive by Southern hybridization (42). Further restriction enzyme mapping of and chromosomal walking from these overlapping cosmids led to the genetic localization of a 55-kb contiguous DNA region, as represented by pBS2002, pBS2003, and pBS2004 (see Fig. 3A). A 6.3-kb BamHI fragment from pBS2003 that hybridized to the nonR probe was cloned into the same site of pGEM-3zf to yield pBS2005, the DNA sequence of which was subsequently determined (see Fig. 3B).

FIG. 3.

FIG. 3

(A) Restriction map of a 55-kb contiguous DNA region from S. griseus as represented by three overlapping cosmid clones. (B) Genetic organization of the nonDLSRB genes within the non cluster. nonR probe, the 1.3-kb fragment from pBS2001; B, BamHI.

Construction of nonS mutants by gene disruption and replacement.

For gene disruption, a 0.7-kb internal fragment of nonS was amplified by PCR from pBS2005 with primers 5′-TCTAGACCTGACGCTCTCCCGC-3′ (forward primer) and 5′-GAATTCTCCGTGAACGCGGCC-3′ (reverse primer) and was cloned into pGEM-T to yield pBS2006. The latter plasmid was digested with EcoRI and XbaI, and the resulting 0.7-kb EcoRI-XbaI fragment was cloned into the same sites of pDH5 (20) to yield pBS2007 (see Fig. 4A).

FIG. 4.

FIG. 4

Inactivation of nonS by gene disruption via single-crossover homologous recombination. (A) Construction of the nonS disruption mutant and restriction maps of the S. griseus DSM40695 wild-type and SB2001 mutant strains showing predicted fragment sizes upon BamHI digestion. Southern analysis of wild-type (lane 1) and SB2001 (lane 2) genomic DNA, digested with BamHI, by using the pDH5 vector (B) or the 0.7-kb nonS fragment from pBS2007 (C) as a probe. B, BamHI.

For gene replacement, a 2.2-kb HincII fragment that contained nonS was subcloned from pBS2005 into the same site of pGEM-3zf to yield pBS2008. The latter plasmid was digested with EcoRI and HindIII, and the resulting 2.2-kb EcoRI-HindIII fragment with the desired orientation was cloned into the same sites of pDH5 to yield pSB2009. The aac(3)IV apramycin resistance gene was rescued from pUO9090 as a 1.5-kb BglII-BamHI fragment and was inserted into the unique BglII site of pBS2009 to yield pBS2010, in which the nonS gene was insertionally inactivated by aac(3)IV transcribed in the opposite direction in reference to the direction of transcription for nonS (see Fig. 5A).

FIG. 5.

FIG. 5

Inactivation of nonS by gene replacement via double-crossover homologous recombination. (A) Construction of the nonS replacement mutant and restriction maps of the S. griseus DSM40695 wild-type and SB2003 mutant strains showing predicted fragment sizes upon BamHI digestion. Southern analysis of wild-type (lane 2) and SB2003 (lane 3) genomic DNA, digested with BamHI, by using the aac(3)IV gene (B) or the 0.7-kb nonS fragment from pBS2007 (C) as a probe. Lane 1, bacteriophage λ DNA-HindIII digest marker. B, BamHI; Bg, BglII; H, HincII.

To introduce pBS2007 and pBS2010 into S. griseus by polyethylene glycol (PEG)-mediated protoplast transformation (23), these plasmids were propagated in E. coli ET12567 (29), and the resulting double-stranded plasmid DNA was denatured by alkaline treatment (21). The latter DNA (10 μl) and 25% PEG 1000 in P buffer (200 μl) (23) were sequentially added to 100 μl of S. griseus protoplasts (109) in P buffer. The resulting solution was mixed immediately and was spread onto R1M plates. After incubation at 28°C for 16 to 20 h, the plates were selected with thiostrepton (final concentration, 25 μg/ml) for pBS2007 and apramycin (final concentration, 50 μg/ml) for pBS2010; incubation continued until colonies appeared (in 3 to 5 days). Those transformants that were derived from pBS2007 and that were resistant to thiostrepton were identified as S. griseus SB2001 mutants; those transformants that were derived from pBS2010 and that were resistant to both apramycin and thiostrepton were identified as S. griseus SB2002 mutants.

S. griseus SB2002 was further grown sequentially in liquid SGGP medium for two rounds without any antibiotic and for one round with apramycin (final concentration, 50 μg/ml), each of which was at 28°C and 300 rpm for 20 h, and was then plated onto solid SGGP medium with apramycin (final concentration, 50 μg/ml) and incubated at 28°C for 2 days. The resulting colonies were selected on SGGP plates with apramycin (final concentration, 50 μg/ml) and thiostrepton (final concentration, 25 μg/ml), respectively. Those colonies that were thiostrepton sensitive and apramycin resistant were identified as S. griseus SB2003 mutants.

Construction of nonS expression plasmids.

pBS2011 was constructed by cloning a 2.2-kb EcoRI-HindIII fragment that contained the intact nonS gene from pBS2008 into the same sites of pWHM3. To clone the ermE* promoter, pWHM79 (43) was digested with EcoRI-BamHI, and the resulting 0.45-kb EcoRI-BamHI fragment was inserted into the same sites of pBS2011 to yield pBS2012, in which the expression of nonS is under the control of the ermE* promoter (7).

Analysis of macrotetrolide production.

Macrotetrolides were isolated by the procedure described by Ashworth and Robinson (4) and were analyzed by either thin-layer chromatography (TLC), high-performance liquid chromatography (HPLC), or bioassay. For TLC, Keiselgel 60 F254 glass plates (gel thickness, 0.25 mm; Merck, Rahway, N.J.) were used and developed with CHCl3-ethyl acetate (1:2; vol/vol). Individual macrotetrolides were purified by preparative TLC and were subjected to electron-spray mass spectral analysis. The latter was performed on a VG BioQ spectrometer at the Facility for Advanced Instrumentation of the University of California, Davis. For HPLC analysis, the method of Herlt (19) was used, by which a Microsorb-MV C18 column (4.6 by 250 mm; 5 μm; pore size 100 Å; Rainin, Walnut Creek, Calif.), isocratic elution with tetrahydrofuran-H2O (60:40; vol/vol) at flow rate of 0.8 ml/min on a Dynamax gradient HPLC system (Rainin), and an evaporative light-scattering detector (Alltech, Deerfield, Ill.) were used. Macrotetrolide distribution was determined from the average of several fermentations. For the bioassay, either the fermentation supernatant (200 μl) or the acetone extract (10 μl) was added to stainless steel cylinders or filter paper disks, respectively, both of which were placed on LB plates preseeded with an overnight M. luteus culture (0.01%; vol/vol). The plates were incubated at 37°C for 24 h, and macrotetrolide production was estimated by measuring the sizes of the inhibition zones.

Feeding of (±)-nonactic acid to nonS mutants.

To prepare (±)-NA from racemic methyl nonactate, 10 mg of the methyl nonactate was dissolved in methanol (200 μl), and this solution was mixed with 2 N NaOH (200 μl). The resulting mixture was shaken at room temperature for 3 h, and hydrolysis was quantitative as judged by TLC and development with CHCl3-ethyl acetate (1:2; vol/vol). The mixture was then neutralized by addition of concentrated HCl (33 μl) and was evaporated to dryness. The residue was dissolved in H2O (6.0 ml; pH 8.5), sterilized by filtration (pore size, 0.22 μm; Millipore, Bedford, Mass.), and stored at 4°C for feeding experiments.

To 25 ml of a production culture of S. griseus DSM40695, SB2001, or SB2003, a total of 8.2 mg of (±)-NA was administrated in three equal portions on days 2, 3, and 4, respectively. Fermentation continued for 2 additional days, and macrotetrolides were isolated and analyzed by the procedures described above.

Nucleotide sequence accession number.

The nucleotide sequence reported here has been deposited in the GenBank database under accession numbers AF263011 and AF263012.

RESULTS

Cloning of the non gene cluster from S. griseus DSM40695 and sequence analysis of the nonR locus.

Plater and Robinson (37) cloned the nonR gene from S. griseus and demonstrated that NonR confers macrotetrolide resistance on S. lividans in the presence of K+ ion. Given the fact that antibiotic production genes have been found in nearly all cases to be clustered in one region of the bacterial chromosome that consists of structural, resistance, and regulatory genes (22), we proceeded to isolate and sequence the DNA flanking the nonR gene in order to identify the non gene cluster. We amplified a 1.3-kb nonR fragment (pBS2001) by PCR and used it as a probe to screen an S. griseus genomic library that was constructed in the E. coli-Streptomyces shuttle vector pOJ446 (8). Of the 2,400 colonies screened by colony hybridization, 54 positive clones were identified, and 12 of these were selected for further analysis. Restriction enzyme mapping and Southern analysis showed that 11 of the 12 clones contained a single 6.3-kb BamHI fragment that hybridizes to the nonR probe. Additional chromosomal walking from this locus eventually led to the localization of a 55-kb contiguous DNA region covered by 30 overlapping cosmids, as exemplified by pBS2002, pBS2003, and pBS2004 (Fig. 3A).

The 6.3-kb BamHI fragment was subcloned (pBS2005), and its nucleotide sequence was determined. (During our work, this region was also sequenced [GenBank accession number AF074603] and, very recently, was reported by Priestly and coworkers [51].) Five complete ORFs, transcribed in the same direction, were identified, including nonR and orfB, (also called nonB) described previously by Plater and Robinson (37) and nonS reported very recently by Priestley and coworkers (53) (Fig. 3B). Immediately upstream of nonS are two additional ORFs, nonL and nonD, which encode a 555-amino-acid protein with a molecular weight of 59,595 and an isoelectric point of 5.83 and a 569-amino-acid protein with a molecular weight of 61,566 and an isoelectric point of 4.98, respectively. NonL is homologous to a family of coenzyme A (CoA) ligases or ligase-like domains, including the loading domains of the Rif PKS from Amycolatopsis mediterranei S699 (33% identity and 43% similarity) (6), the FK506 PKS from Streptomyces sp. strain MA6548 (32% identity and 42% similarity) (33), and the second type I PKS from the rapamycin-producing strain Streptomyces hygroscopicus ATCC 29253 (28% identity and 40% similarity) (41). These CoA ligase-like domains of type I PKS catalyze the ATP-dependent activation of free carboxylic acids into the corresponding acyl CoAs before they can be loaded onto the PKS complex for polyketide biosynthesis. An equivalent role could be proposed for NonL, although the substrate for NonL is yet to be determined. The fact that the addition of ATP has no effect on the in vitro synthesis of macrotetrolides from NA and its homologs argues against the notion that the latter acids are activated into acyl CoAs that involve a CoA ligase like NonL before incorporation into macrotetrolides (2, 36). NonD shows low but significant end-to-end similarity to several glutaryl 7-amino cephalosporanic acid (7-ACA) acylases, including the ones from Bacillus subtilis (28.4% identity and 40.8% similarity) (1) and from Zymomonas mobilis (19% identity and 27.3% similarity) (GenBank accession number AF124757), as well as to several proteins of unknown function, such as the hypothetical protein Rv2800 from Mycobacterium tuberculosis (31% identity and 40% similarity; GenBank accession number Z81331) and the hypothetical protein from a Synechocystis sp. (26% identity and 39% similarity; GenBank accession number D90903). Since glutaryl 7-ACA acylase catalyzes the amide hydrolysis of glutaryl 7-ACA to yield 7-ACA, it is not clear how NonD could play a role in macrotetrolide biosynthesis.

Inactivation of the nonS gene in S. griseus DSM40695 by gene disruption and replacement.

To examine if the cloned gene cluster encodes macrotetrolide biosynthesis, nonS was inactivated by both insertional disruption and replacement with a mutant copy to generate macrotetrolide-nonproducing mutant strains. To effect gene disruption, pBS2007—a pDH5 derivative that contains a 0.7-kb internal fragment of nonS—was introduced into S. griseus via PEG-mediated protoplast transformation. One of the thiostrepton-resistant transformants was selected and was named S. griseus SB2001. Since pBS2007 is derived from the Streptomyces-nonreplicating plasmid pDH5, S. griseus SB2001 must have resulted from integration of pBS2007 into the S. griseus chromosome by insert-directed homologous recombination (Fig. 4A).

To confirm that targeted nonS disruption has occurred by a single-crossover homologous recombination event, Southern analysis of genomic DNA from S. griseus SB2001 was performed with either pDH5 or the 0.7-kb fragment of nonS as a probe. As shown in Fig. 4B, a distinctive band of the predicted size of 11 kb was detected with pDH5 as a probe in S. griseus SB2001 (lane 2); this band was absent from the wild-type strain S. griseus DSM40695 (Fig. 4B, lane 1). Complementarily, when the 0.7-kb fragment of nonS was used as a probe, the 6.3-kb band in the wild-type strain (Fig. 4C, lane 1) was shifted to a 11-kb fragment in the SB2001 mutant strain (Fig. 4C, lane 2), as would be expected for disruption of nonS by a single-crossover homologous recombination event (Fig. 4A).

To effect gene replacement, pBS2010—a pDH5 derivative that contains a mutant copy of nonS generated by inserting the aac(3)IV apramycin resistance gene from pUO9090 into the unique BglII site of nonS—was similarly introduced into S. griseus via PEG-mediated protoplast transformation. Transformants resistant to both thiostrepton and apramycin (five colonies) were isolated and identified as S. griseus SB2002, which had resulted from a single-crossover homologous recombination event as confirmed by Southern analysis (data not shown). Direct screening for colonies resistant to apramycin only was unsuccessful, suggesting that the double-crossover homologous recombination event occurs at a very low frequency. To encourage the double-crossover event, the SB2002 strain was cultured sequentially in a liquid medium for several rounds without any added antibiotic and for one round in the presence of apramycin. The resulting culture was then selected on solid medium containing thiostrepton or apramycin. Colonies that are thiostrepton sensitive and apramycin resistant were considered to have resulted from the double-crossover homologous recombination event (Fig. 5A), and under the conditions examined, the latter occurred at frequencies as high as 50%. The apramycin-resistant S. griseus SB2003 strains were identified and selected as representatives of nonS replacement mutants.

To confirm that targeted nonS replacement has occurred by a double-crossover homologous recombination event, Southern analysis of genomic DNA from S. griseus SB2003 was performed with either acc(3)IV or nonS as a probe. As shown in Fig. 5B, a distinctive band of the predicted size of 7.8 kb was detected with acc(3)IV as a probe in S. griseus SB2003 (Fig. 5B, lane 3); this band was absent from the wild-type strain S. griseus DSM40695 (Fig. 5B, lane 2). Complementarily, when nonS was used as a probe, the 6.3-kb band in the wild-type strain (Fig. 5C, lane 2) was shifted to a 7.8-kb fragment in the SB2003 mutant strain (Fig. 5C, lane 3), as would be expected for replacement of nonS with the mutant copy by a double-crossover homologous recombination event (Fig. 5A).

Characterization of the nonS mutants as macrotetralide nonproducers.

S. griseus DSM40695 produces a mixture of macrotetrolides (NON, monactin, dinactin, trinactin, and tetranactin at 9.3:23:100:81:18), as judged by HPLC analysis of acetone extracts of the freeze-dried mycelia with an evaporative light-scattering detector (Fig. 6A) (19). While no apparent difference in growth characteristics and morphologies between the wild-type and the mutant strains was observed, macrotetrolide production is completely abolished in either the SB2001 nonS disruption mutant strain or the SB2003 nonS replacement mutant strain (Fig. 6B and D, respectively). Macrotetrolide production was also determined by assaying the antibacterial activity of the fermentation broth or the acetone extracts against M. luteus ATCC 9431. Consistent with the HPLC analysis, acetone extracts or aliquots of fermentation broth from either the SB2001 or SB2003 strains showed a complete lack of activity, while a clear inhibition zone was observed from that of the wild-type organism as well as from that of the positive control with an authentic sample of NON (data not shown).

FIG. 6.

FIG. 6

HPLC analysis with an evaporative light-scattering detector of macrotetrolides isolated from a wild-type strain (A), a nonS disruption mutant strain of SB2001 (B), an SB2001 strain supplemented with (±)-NA (C), a nonS replacement mutant strain of SB2003 (D), an SB2003 strain supplemented with (±)-NA (E), an SB2003 strain complemented with nonS (F), and an SB2003 strain complemented with nonS (G), the expression of which is ensured by the ermE* promoter.

In vivo complementation of the nonS mutants by expression of nonS in trans.

The ability of the wild-type nonS gene to complement the inactivated nonS gene in vivo was tested in the S. griseus SB2003 mutant strain. Two nonS expression constructs were made in pWHM3 (50), pBS2011 and pBS2012, the latter of which uses the constitutive ermE* promoter (7) to ensure efficient expression of nonS. Both the pBS2011 and pBS2012 constructs, as well as the pWHM3 vector as a control, were introduced via PEG-mediated protoplast transformation into S. griseus SB2003. Acetone extracts from fermentations of three transformants of each variant were analyzed for macrotetrolide production by HPLC. While no adverse effect on the growth characteristics of S. griseus SB2003 was observed upon introduction of pWHM3, pBS2011 clearly restored macrotetrolide production to the SB2003 strain, albeit the yields of these metabolites were low in comparison with the yields from the wild-type strain (Fig. 6A versus F). The latter phenotype could be caused by the poor expression of nonS in pBS2011. This belief was indeed supported by the observation that introduction of pBS2012 into the SB2003 mutant resulted in production of the macrotetrolides to a comparable level, with the distribution of individual macrotetrolides (NON, monactin, dinactin, triactin, and tetranactin at 15:37:100:54:11) being very similar to that for the wild-type strain (Fig. 6A versus G). These data confirm that the abolishment of macrotetrolide production in the SB2003 strain is due to the specific inactivation of nonS.

Fermentation of the nonS mutants in the presence of exogenously added (±)-NA.

To investigate if nonS governs one of the enantiospecific pathways that leads to the biosynthesis of (+)- or (−)-NA and its homologs, S. griseus SB2001 and SB2003, as well as the wild-type strain as a control, were cultured in medium supplemented with exogenous (±)-NA. Acetone extracts from fermentations of all three strains were analyzed for macrotetrolide production by HPLC. No significant differences in either the total production yield or the distribution of individual macrotetrolides were observed when the wild-type strain was cultured in the presence of exogenous (±)-NA. In contrast, additions of (±)-NA to the fermentation medium only partially restored macrotetrolide production to SB2001 (Fig. 6C) and SB2003 (Fig. 6E); only NON, monactin, and dinactin were produced, but trinactin and tetranactin were not found. In contrast to the wild-type strain that produces dianactin as the major macrotetrolide, SB2001 and SB2003, in the presence of exogenous (±)-NA, synthesize NON as the predominant product (NON, monactin, and dinactin at 100:20:1.6). The latter was, in fact, expected, since, unlike the formation of NON, which can be synthesized from the exogenously added (±)-NA, the formation of either monactin or dinactin requires one or two molar equivalents of endogenous (+)-HNA (Fig. 2B), respectively, the concentration of which is apparently very low in comparison with the large excess of (±)-NA added exogenously. Both NON and monactin were isolated and subjected to electron-spray mass spectral analysis with authentic samples of NON and monactin as references. The isolated NON and monactin yielded the characteristic ions as the authentic standards with molecular weights of 737.4 (M + H+), 754.6 (M + NH4+), 759.7 (M + Na+), and 775.6 (M + K+) for NON and of 751.4 (M + H+), 768.5 (M + NH4+), 773.6 (M + Na+), and 789.6 (M + K+) for monactin. Close examination of the macrotetrolide structures reveals that, in addition to the (±)-NA monomer, the production of monactin and dinactin requires only (+)-HNA, and the production of trinactin and tetranactin requires both (+)- and (−)-HNA. The synthesis of either (+)- or (−)-HNA, if abolished as the result of nonS inactivation, cannot be circumvented by exogenous supplementation of (±)-NA (Fig. 2B).

DISCUSSION

Polyketides are structurally classified into four major groups: aromatic polyketides, macrolides, polyenes, and polyethers. Recent advances in genetic and biochemical studies of polyketide biosynthesis have provided a mechanistic explanation of how PKSs achieve the vast structural diversity by varying the similar sets of biosynthetic reactions, leading to the successful cloning and characterization of multiple sets of gene clusters that encode aromatic polyketide, macrolide, or polyene biosynthesis (22). However, genetic analysis of polyether biosynthesis has heretofore met with little success, despite considerable effort. The only published reports were the cloning from Streptomyces longisporoflavus of tnrB1 and tnrB2, which confer tetronasin resistance, by Leadlay and coworkers (28); the cloning from S. griseus of nonR, which confers tetranactin resistance, by Plater and Robinson (37); and the cloning from S. griseus of nonS, which exhibits nonactate synthase activity upon heterologous expression, by Priestley and Earle (39). Very recently, Priestley and coworkers (51) also reported on the DNA sequence analysis of 15.5-kb flanking the nonRS genes. While those studies certainly have set an excellent stage for further investigation, they fell short of proving the direct involvement of the cloned genes in either tetronasin or macrotetrolide biosynthesis. Thus, our report here on the genetic localization of a 55-kb contiguous DNA region, inactivation of nonS in S. griseus for the generation of macrotetrolide-nonproducing mutants, the in vivo complementation of nonS mutants by expression of the nonS gene in trans, and the fermentation of the nonS mutants in the presence of exogenous (±)-NA for the restoration of macrotetrolide production established unambiguously the successful cloning of the non biosynthesis gene cluster. As it has been demonstrated for genetic characterizations of nonS, we envisage that the genetic system developed here for S. griseus should enable us to analyze in detail the entire non gene cluster involved in macrotetrolide biosynthesis.

The intriguing molecular topology of the macrotetrolides, which consists of the (+)(−)(+)(−)-ester linkage of the enantiomeric NA and its homologs, has inspired us and others (45, 53) to propose that the biosynthesis of these enantiomeric polyketide intermediates results from a pair of enantiospecific pathways (Fig. 2A). This hypothesis is consistent with the efficient and enantiospecific incorporations of the N-caprylcysteamine thioesters of both (6R,8R)-NEA and (6S,8S)-NEA into NON (45) and with the in vitro transformation of the racemic N-octylcysteamine thioester of NEA into NA by a cell-free preparation made from either an S. griseus or an S. lividans strain that heterologously expresses the nonS gene (53). It is imagined that a pair of enantiospecific PKS enzymes catalyzes the assembly of the two enantiomers of NEA from one molecule each of acetate, propionate, and succinate and that, on the basis of the paradigm of PKSs, the occurrence of NEA homologs—consequently, HNA and BNA—resulted from a less-than-strict starter unit specificity exhibited by the PKS enzymes. To test this hypothesis, we set out to inactivate nonS by targeted gene disruption (SB2001 mutant strain) or gene replacement (SB2003 mutant strain). We reasoned that inactivation of nonS should abolish the biosynthesis of at least one set of the enantiomeric monomers, consequently rendering the nonS mutants macrotetrolide nonproducers, as depicted in Fig. 2B. Examination of SB2001 and SB2003 confirmed that macrotetrolide production has been completely abolished in both strains (Fig. 6B and D). Complementarily, the nonS mutation can be fully complemented in trans by a plasmid-borne copy of nonS. Thus, introduction of pBS1012—in which the expression of nonS is under the control of the ermE* promoter—into SB2003 not only restored macrotetrolide production to the mutant strain to the wild-type level but also restored the distribution of individual macrotetrolides so that it was similar to that in the wild type (Fig. 6A versus G). These results not only confirm that the abolishment of macrotetrolide production in SB2003 results from the specific inactivation of nonS but also indicate that NonS alone is responsible for the synthesis of at least one enantiomeric set of NA and its homologs.

To probe if nonS governs only one or both enantiospecific pathways that lead to the synthesis of (−)- or (+)-NA and its homologs, we cultured SB2001 and SB2003 in the presence of exogenous (±)-NA. We reasoned, as summarized in Fig. 2B, that if nonS governs both enantiospecific pathways, addition of (±)-NA to the nonS mutants should result in the production of NON only because the synthesis of the other macotetrolides requires endogenous (−)- or (+)-HNA, or both. The biosynthesis of the latter has been abolished as the result of nonS inactivation. The same phenotype could also occur, however, if nonS is responsible only for the synthesis of (+)-NA and its homologs, since NON is the only macrotetrolide whose biosynthesis does not require (+)-HNA. In contrast, if nonS is responsible only for the synthesis of (−)-NA and its homologs, addition of (±)-NA to the nonS mutants should result in the production of NON, monactin, and dinactin; trinactin and tetranactin should not be found due to the lack of endogenous (−)-HNA. While no significant difference in either the total production yield or the distribution of individual macrotetrolides was observed when the wild-type strain was cultured in the presence of exogenous (±)-NA, only NON, monactin, and dinactin were detected from SB2001 (Fig. 6C) and SB2003 (Fig. 6E) upon fermentation in the presence of exogenous (±)-NA; the identities of these macrotetrolides were confirmed by both HPLC and electron-spray mass spectral analysis in comparison with authentic standards (dinactin was not analyzed by mass spectral analysis due to its trace quantity). These data agree well with the hypothesis that nonS is responsible only for the synthesis of (−)-NA and its homologs, providing strong support that macrotetrolide biosynthesis in S. griseus involves a pair of enantiospecific pathways. Indeed, sequence analysis of the non cluster has revealed an ORF 18.2 kb upstream of nonS. The deduced amino acid sequence of this ORF shows significant end-to-end similarity to NonS (21% identity and 29% similarity) as well as to the well-characterized mitochondrial 2-enoyl-CoA hydratase-1 (23% identity and 38% similarity) (31). (NonS shows almost an identical level of homology to mitochondrial 2-enoyl-CoA hydratase-1 [25% identity and 37% similarity]). Experiments are in progress to investigate if the latter enzyme, complementary to NonS, is responsible for the synthesis of (+)-NA and its homologs.

It remains obscure how modular PKS enzymes control the stereochemistry during the polyketide chain assembly at either the alkyl-branched center, such as C-2 of NA, or the oxygenated center (derived from the β-keto group of the polyketide intermediates), such as C-8 and C-6 of NA (Fig. 1B). Progress has recently been made in this direction by studying the 6-deoxyerythronolide B synthase in vivo or in vitro, leading to the finding that the stereochemistry of the alkyl-branched center or the oxygenated center is controlled by the ketoacyl synthase (KS) (10, 31, 52) or the ketoreductase (KR) (10, 25), respectively, in the specific cases examined. However, these studies fall short of providing a satisfactory explanation at the molecular level of how a given KS or KR exhibits its stereospecificity; sequence comparison among various KS or KR domains has also failed to reveal any conserved motifs that could potentially be ascribed to their opposite stereospecificities. We envisaged that PKS enzymes from a pair of enantiospecific polyketide pathways should provide an ideal model that could be used to study the stereochemistry of polyketide biosynthesis. Since these enzymes must recognize and process a set of enantiomeric substrates and intermediates, any difference in their primary amino acid sequences and secondary protein structures shall provide an understanding of the factors by which PKS enzyme controls stereochemistry during polyketide biosynthesis. The current work indicates the existence of such a pair of enantiospecific polyketide pathways for macrotetrolide biosynthesis in S. griseus, setting the stage for in-depth investigations. Furthermore, enantiospecific pairs of enzymes are relatively rare in nature, and only in a few cases have such enzymes been isolated and characterized comparatively (44). Studies with individual enzymes for macrotetrolide biosynthesis, therefore, shall shed light on the formation of enantiomeric natural products in general. The cloning of the non cluster and the establishment of a genetic system in S. griseus have made such studies possible.

ACKNOWLEDGMENTS

We thank C. Richard Hutchinson, University of Wisconsin, Madison, for pWHM3 and pWHM79; Douglas J. MacNeil, Merck Research Laboratories, Rahway, N.J., for a culture of E. coli ET12567; M. C. Martin, Universidad de Oviedo, Oviedo, Spain, for pUO9090; Peter Metz, Technische Universität Dresden, Dresden, Germany, for a gift of the (±)-methyl nonactate; Y. Wang, Institute of Medicinal Biotechnology, Beijing, China, for a culture of M. luteus ATCC 9431; Liangcheng Du, Cesar Sanchez, and Scott Standage for critical reading of the manuscript; and Kara Ishii for editorial assistance. Mass spectral analysis was performed at the Facility for Advanced Instrumentation of the University of California, Davis.

This work was supported in part by a CAREER award (award MCB9733938) from the National Science Foundation and the Searle Scholars Program/The Chicago Community Trust.

W. C. Smith and L. Xiang contributed equally to this work.

REFERENCES

  • 1.Aramori I, Fukagawa M, Tsumura M, Iwami M, Ono H, Kojo H, Kohsaka M, Ueda Y, Imanaka H. Cloning and nucleotide sequencing of a novel 7β-(4-carboxybutanamido)cephalosporanic acid acylase gene of Bacillus laterosporus and its expression in Escherichia coli and Bacillus subtilis. J Bacteriol. 1991;173:7848–7855. doi: 10.1128/jb.173.24.7848-7855.1991. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2.Ashworth D M, Clark C A, Robinson J A. On the biosynthetic origins of the hydrogen atoms in the macrotetrolide antibiotics: their mode of assembly catalyzed by a nonactin polyketide synthase. J Chem Soc Perkin Trans I. 1989;1989:1461–1467. [Google Scholar]
  • 3.Ashworth D M, Robinson J A. Biosynthesis of the macrotetralide antibiotics: an investigation using carbon-13 and oxygen-18 labeled acetate and propionate. J Chem Soc Chem Commun. 1983;1983:1327–1329. [Google Scholar]
  • 4.Ashworth D M, Robinson J A. Biosynthesis of the macrotetrolide antibiotics: the incorporation of carbon-13 and oxygen-18 labeled acetate, propionate, and succinate. J Chem Soc Perkin Trans I. 1988;1988:1719–1727. [Google Scholar]
  • 5.Ashworth D M, Robinson J A, Turner D L. Biosynthesis of nonactin from acetate, propionate, and succinate: the assignment of its carbon-13 NMR spectrum by two-dimensional correlation spectroscopy. J Chem Soc Chem Commun. 1982;1982:491–493. [Google Scholar]
  • 6.August P R, Tang L, Yoon Y J, Ning S, Müller R, Yu T-W, Taylor M, Hoffmann D, Kim C-G, Zhang X, Hutchinson C R, Floss H Z. Biosynthesis of the ansamycin antibiotic rifamycin: deductions from the molecular analysis of the rif biosynthetic gene cluster of Amycolatopsis mediterranei S699. Chem Biol. 1998;5:69–79. doi: 10.1016/s1074-5521(98)90141-7. [DOI] [PubMed] [Google Scholar]
  • 7.Bibb M J, White J, Ward J M, Janssen G R. The mRNA for the 23S rRNA methylase encoded by the ermE gene of Saccharopolyspora erythraea is translated in the absence of a conventional ribosome-binding site. Mol Microbiol. 1994;14:533–545. doi: 10.1111/j.1365-2958.1994.tb02187.x. [DOI] [PubMed] [Google Scholar]
  • 8.Bierman M, Logan R, O'Brien K, Seno E T, Rao R N, Schoner B E. Plasmid cloning vectors for the conjugal transfer of DNA from Escherichia coli to Streptomyces spp. Gene. 1992;116:43–69. doi: 10.1016/0378-1119(92)90627-2. [DOI] [PubMed] [Google Scholar]
  • 9.Birch A W, Robinson J A. Polyethers. In: Vining L C, Stuttard C, editors. Genetics and biochemistry of antibiotic production. Boston, Mass: Butterworth-Heinemann; 1995. pp. 443–476. [Google Scholar]
  • 10.Böhm I, Holzbaur I E, Hanefeld U, Cortes J, Staunton J, Leadlay P F. Engineering of a minimal modular polyketide synthase, and targeted alteration of the stereospecificity of polyketide chain extension. Chem Biol. 1998;5:407–412. doi: 10.1016/s1074-5521(98)90157-0. [DOI] [PubMed] [Google Scholar]
  • 11.Borrel M N, Pereira E, Fiallo M, Garnier-Suillerot A. Mobile ionophores are a novel class of P-glycoprotein inhibitors. The effects of ionophores on 4′-O-tetrahydropyranyl-adriamycin incorporation in K562 drug-resistant cells. Eur J Biochem. 1994;223:125–133. doi: 10.1111/j.1432-1033.1994.tb18973.x. [DOI] [PubMed] [Google Scholar]
  • 12.Callewaert D M, Radcliff G, Tanouchi Y, Shichi H. Tetranactin, a macrotetrolide antibiotic, suppresses in vitro proliferation of human lymphocytes and generation of cytotoxicity. Immunopharmacology. 1988;16:25–32. doi: 10.1016/0162-3109(88)90047-1. [DOI] [PubMed] [Google Scholar]
  • 13.Clark C A, Robinson J A. Biosynthesis of nonactin. The role of acetoacetyl-CoA in the formation of nonactic acid. J Chem Soc Chem Commun. 1985;1985:1568–1569. [Google Scholar]
  • 14.Corbaz R, Ettlinger L, Gäumann E, Keller-Schierlein W, Kradolfer F, Neipp L, Prelog V, Zöhner H. Metabolic products of actinomycetes. III. Nonactin. Helv Chim Acta. 1955;38:1445–1448. [Google Scholar]
  • 15.Dobler M. The crystal structure of nonactin. Helv Chim Acta. 1972;55:1371–1384. doi: 10.1002/hlca.19720550504. [DOI] [PubMed] [Google Scholar]
  • 16.Dutton C J, Banks B J, Cooper C B. Polyether ionophores. Nat Prod Rep. 1995;12:165–181. doi: 10.1039/np9951200165. [DOI] [PubMed] [Google Scholar]
  • 17.Fleck W F, Ritzau M, Heinze S, Gräfe U. Isolation of dimeric nonactic acid from the nonactin-producing Streptomyces spec. JA 5909-1. J Basic Microbiol. 1996;36:235–238. [Google Scholar]
  • 18.Fleming I, Ghosh S K. The synthesis of nonactic acid. Its derivatives and nonactin itself. In: Rahman A, editor. Studies in natural products chemistry. Vol. 18. New York, N.Y: Elsevier Science; 1996. pp. 229–268. [Google Scholar]
  • 19.Herlt T. A facile separation of nonactin and its homologues. J Liq Chromatogr Rel Technol. 1997;20:1295–1300. [Google Scholar]
  • 20.Hillenann D, Pühler A, Wohlleben W. Gene disruption and gene replacement in Streptomyces via single stranded DNA transformation of integration vectors. Nucleic Acids Res. 1991;19:727–731. doi: 10.1093/nar/19.4.727. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Ho S-H, Chater K F. Denaturation of circular or linear DNA facilitates targeted integrative transformation of Streptomyces coelicolor A3(2): possible relevance to other organisms. J Bacteriol. 1997;179:122–127. doi: 10.1128/jb.179.1.122-127.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Hopwood D A. Genetic contributions to understanding polyketide synthases. Chem Rev. 1997;97:2465–2497. doi: 10.1021/cr960034i. [DOI] [PubMed] [Google Scholar]
  • 23.Hopwood D A, Bibb M J, Chater K F, Kieser T, Bruton C J, Kieser H M, Lydiate D J, Smith C P, Schrempf H. Genetic manipulation of Streptomyces: a laboratory manual. Norwich, United Kingdom: The John Innes Foundation; 1985. [Google Scholar]
  • 24.Jois H R Y, Sarkar A, Gurusiddaiah S. Antifungal macrodiolide from Streptomyces sp. Antimicrob Agent Chemother. 1986;30:458–464. doi: 10.1128/aac.30.3.458. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Kao C M, McPherson M, McDaniel R N, Fu H, Cane D E, Khosla C. Alcohol stereochemistry in polyketide backbones is controlled by the β-ketoreductase domains of modular polyketide synthase. J Am Chem Soc. 1998;120:2478–2479. [Google Scholar]
  • 26.Kiema T-R, Engel C K, Schmitz W, Filppula S A, Wierenga R K, Hiltunen J K. Mutagenic and enzymological studies of the hydratase and isomerase activities of 2-enoyl-CoA hydratase-1. Biochemistry. 1999;38:2991–2999. doi: 10.1021/bi981646v. [DOI] [PubMed] [Google Scholar]
  • 27.Kilbourn B T, Dunitz J D, Pioda L A R, Simon W. Structure of the K+ complex with nonactin, a macrotetrolide antibiotic possessing highly specific K+ transport properties. J Mol Biol. 1967;30:559–563. doi: 10.1016/0022-2836(67)90370-1. [DOI] [PubMed] [Google Scholar]
  • 28.Linton K J, Cooper H N, Hunter I S, Leadlay P F. An ABC-transporter from Streptomyces longisporoflavus confers resistance to the polyether-ionophore antibiotic tetronasin. Mol Microbiol. 1994;11:777–785. doi: 10.1111/j.1365-2958.1994.tb00355.x. [DOI] [PubMed] [Google Scholar]
  • 29.MacNeil D J, Gewain K M, Ruby C L, Dezney G, Gibbons P H, MacNeil T. Analysis of Streptomyces avermitilis genes required for avermectin biosynthesis utilizing a novel integration vector. Gene. 1992;111:61–68. doi: 10.1016/0378-1119(92)90603-m. [DOI] [PubMed] [Google Scholar]
  • 30.Marrone T J, Merz K M., Jr Molecular recognition of potassium ion by the naturally occurring antibiotic ionophore nonactin. Biochemistry. 1992;114:7542–7549. [Google Scholar]
  • 31.Marsden A F A, Caffrey P, Aparicio J F, Loughran M S, Staunton J, Leadlay P F. Stereospecific acyl transfers on the erythromycin-producing polyketide synthase. Science. 1994;263:378–380. doi: 10.1126/science.8278811. [DOI] [PubMed] [Google Scholar]
  • 32.Meiners U, Cramer E, Fröhlich R, Wibbeling B, Metz P. A general route to the monomeric subunits of the macrotetrolides—a short synthesis of methyl nonactate. Eur J Org Chem. 1998;1998:2073–2078. [Google Scholar]
  • 33.Motamedi H, Cai S-J, Shafiee A, Elliston K O. Structural organization of a multifunctional polyketide synthase involved in the biosynthesis of the macrolide immunosuppressant FK506. Eur J Biochem. 1997;244:74–80. doi: 10.1111/j.1432-1033.1997.00074.x. [DOI] [PubMed] [Google Scholar]
  • 34.Natsume M, Kondo S, Marumo S. The absolute stereochemistry of pamamycin-607, an aerial mycelium-inducing substance of Streptomyces alboniger. J Chem Soc Chem Commun. 1989;1989:1991–1993. [Google Scholar]
  • 35.Natsume M, Yasui K, Kondo S, Marumo S. The structures of four new pamamycin homologues isolated from Streptomyces alboniger. Tetrahedron Lett. 1991;32:3087–3090. [Google Scholar]
  • 36.Nefelova M V, Karelina I Y, Sverdlova A N, Egorow N S. Isolation and properties of macrotetrolide synthase from the mycelium of the actinomycete producer. Biochemistry (Biokhimiya) 1990;54:1531–1538. [PubMed] [Google Scholar]
  • 37.Plater R, Robinson J A. Cloning and sequencing of a gene encoding macrotetrolide antibiotic resistance from Streptomyces griseus. Gene. 1992;112:117–122. doi: 10.1016/0378-1119(92)90312-d. [DOI] [PubMed] [Google Scholar]
  • 38.Pressman B C. Biological applications of ionophores Annu. Rev Biochem. 1976;45:501–530. doi: 10.1146/annurev.bi.45.070176.002441. [DOI] [PubMed] [Google Scholar]
  • 39.Priestley N D, Earle M J. Synthesis and evaluation of a designed inhibitor for nonactin biosynthesis in S. griseus ETH A7796. Tetrahedron Lett. 1997;7:2187–2192. [Google Scholar]
  • 40.Rao R N, Richardson M A, Kuhstoss S. Cosmid shuttle vectors for cloning and analysis of Streptomyces DNA. Methods Enzymol. 1987;153:166–198. doi: 10.1016/0076-6879(87)53053-1. [DOI] [PubMed] [Google Scholar]
  • 41.Ruan X, Stassi D, Lax S, Katz L. A second type-I PKS gene cluster isolated from Streptomyces hygroscopicus ATCC29253, a rapamycin-producing strain. Gene. 1997;203:1–9. doi: 10.1016/s0378-1119(97)00450-2. [DOI] [PubMed] [Google Scholar]
  • 42.Sambrook J, Fritsch E F, Maniatis T. Molecular cloning: a laboratory manual. 2nd ed. Cold Spring Harbor, N.Y: Cold Spring Harbor Laboratory Press; 1989. [Google Scholar]
  • 43.Shen B, Hutchinson C R. Deciphering the mechanism for the assembly of aromatic polyketides by a bacterial polyketide synthase. Proc Natl Acad Sci USA. 1996;93:6600–6604. doi: 10.1073/pnas.93.13.6600. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Shen B, Gould S J. Opposite facial specificity for two hydroquinone epoxidases: (3-si, 4-re)-2,5-dihydroxyacetanilide epoxidase from Streptomyces LL- C10037 and (3-re, 4-si)-2,5-dihydroxyacetanilide epoxidase from Streptomyces MPP 3051. Biochemistry. 1991;30:8937–8944. doi: 10.1021/bi00101a004. [DOI] [PubMed] [Google Scholar]
  • 45.Spavold Z M, Robinson J A. Nonactin biosynthesis: on the role of (6R,8R)- and (6S,8S)-2-methyl-6,8-dihydroxynon-2E-enoic acids in the formation of nonactic acid J. Chem Soc Chem Commun. 1988;1988:4–6. [Google Scholar]
  • 46.Stererding D, Kadenbach B. The K+-ionophores nonactin and valinomycin interact differently with the protein of reconstituted cytochrome c oxidase. J Bioenerg Biomembr. 1990;22:197–205. doi: 10.1007/BF00762946. [DOI] [PubMed] [Google Scholar]
  • 47.Tanouchi Y, Shichi H. Immunosuppressive effects of polynactins (tetranactin, trinactin and dinactin) on experimental autoimmune uveoretinitis in rats. Jpn J Ophthalmol. 1987;31:218–229. [PubMed] [Google Scholar]
  • 48.Tanouchi Y, Shichi H. Immunosuppressive and anti-proliferative effects of a macrotetrolide antibiotic, tetranactin. Immunology. 1988;63:471–475. [PMC free article] [PubMed] [Google Scholar]
  • 49.Teunissen M B M, Pistoor F H M, Rongen H A H, Kapsenberg M L, Bos J D. A comparison of the inhibitory effects of immunosuppressive agents cyclosporine, tetranactin, and didemnin B on human T cell responses in vitro. Transplantation. 1992;53:875–881. doi: 10.1097/00007890-199204000-00031. [DOI] [PubMed] [Google Scholar]
  • 50.Vara J, Lewandowska-Sharbek M, Wang Y-G, Donadio S, Hutchinson C R. Cloning of genes governing the deoxysugar portion of the erythromycin biosynthesis pathway in Saccharopolyspora erythraea (Streptomyces erythreus) J Bacteriol. 1989;171:5872–5881. doi: 10.1128/jb.171.11.5872-5881.1989. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 51.Walczak R J, Woo A J, Strohl W R, Priestley N D. Nonactin biosynthesis: the potential nonactin biosynthesis gene cluster contains type II polyketide synthase-like genes. FEMS Microbiol Lett. 2000;183:171–175. doi: 10.1111/j.1574-6968.2000.tb08953.x. [DOI] [PubMed] [Google Scholar]
  • 52.Weissman K J, Timoney M, Bycroft M, Grice P, Hanefeld U, Staunton J, Leadlay P F. The molecular basis of Celmer's rules: the stereochemistry of the condensation step in chain extension on the erythromycin polyketide synthase. Biochemistry. 1997;36:13849–13855. doi: 10.1021/bi971566b. [DOI] [PubMed] [Google Scholar]
  • 53.Woo A J, Strohl W R, Priestley N D. Nonactin biosynthesis: the product of nonS catalyzes the formation of the furan ring of nonactic acid. Antimicrob Agents Chemother. 1999;43:1662–1668. doi: 10.1128/aac.43.7.1662. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 54.Yamamoto H, Maurer K H, Hutchinson C R. Transformation of Streptomyces erythraeus. J Antibiot. 1986;39:1304–1313. doi: 10.7164/antibiotics.39.1304. [DOI] [PubMed] [Google Scholar]
  • 55.Zhang H, Shinkawa H, Ishikawa J, Kinashi H, Nimi O. Improvement of transformation system in Streptomyces using a modified regeneration medium. J Ferment Bioeng. 1997;83:217–221. [Google Scholar]
  • 56.Zizka Z. Biological effects of macrotetrolide antibiotics and nonactic acids. Folia Microbiol (Prague) 1998;43:7–14. doi: 10.1007/BF02815533. [DOI] [PubMed] [Google Scholar]

Articles from Antimicrobial Agents and Chemotherapy are provided here courtesy of American Society for Microbiology (ASM)

RESOURCES