Institute of Molecular Biology, Slovak Academy of Sciences, Dubravská cesta 21, 842 51 Bratislava, Slovak Republic1
Author for correspondence: J. Kormanec. Tel: +421 7 5941 2432. Fax: +421 7 5477 2316. e-mail: umbijkor{at}savba.sk
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Keywords: glucose induction, differentiation, GAPDH, promoter, S1-nuclease mapping
Abbreviations: GAPDH, glyceraldehyde-3-phosphate dehydrogenase; tsp, transcription start point(s); wt, wild-type
The GenBank/EMBL/DDBJ accession number for the sequence described in this paper is U21191.
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Streptomycetes are mycelial, high-GC Gram-positive bacteria that undergo a complex cycle of morphological differentiation involving the development of spore-bearing aerial hyphae on mycelial colonies. They produce a variety of biologically active secondary metabolites, including the majority of known antibiotics (Chater, 1998 ). Previously, we identified and characterized the GAPDH-encoding gap gene in Streptomyces aureofaciens (Kormanec et al., 1995
). Transcriptional studies of the gap gene suggested monocistronic organization and developmental regulation of the gene. A single promoter, gap-P, was induced by the presence of glucose in the medium, and at the onset of aerial mycelium formation (Kormanec et al., 1997
). Sequence analysis of the region upstream of gap has revealed the 3' end of a gene encoding a protein similar to the AraC/XylS family of bacterial transcriptional regulators (Gallegos et al., 1997
; Kormanec et al., 1995
). Its close proximity to gap suggests a function in the regulation of gap expression, since transcriptional regulators of this family are usually located upstream of the operon they regulate (Gallegos et al., 1997
). To elucidate a possible function of the gene, which we named gapR, in the regulation of gap expression in S. aureofaciens, we disrupted the gene and analysed expression of the gap gene under various conditions in this mutant. The GapR protein, overproduced in an E. coli expression system, was shown to bind upstream of the gap-P promoter region. The results suggest a direct role of GapR in regulation of gap expression in S. aureofaciens.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
DNA manipulations.
DNA manipulations in E. coli were done as described in Ausubel et al. (1987) , and those in Streptomyces were according to Hopwood et al. (1985)
. DNA fragments were isolated from agarose gels by binding to a DEAE-paper as recently described (Kormanec, 2000
). Nucleotide sequencing was performed by the chemical method (Maxam & Gilbert, 1980
). Site-directed mutagenesis was done with a Chameleon mutagenesis kit from Promega.
Disruption of the gapR gene.
The plasmid used for disruption of S. aureofaciens gapR was prepared as follows. Plasmid pRPO7-11C contained the 1170 bp BamHIPstI fragment (Fig. 1a) bearing the full-length S. aureofaciens gap gene (Kormanec et al., 1995
) in pBluescript SK(+). This gap-bearing insert was removed as a 1170 bp XhoIXbaI (blunt-ended) fragment and inserted into XhoI- and SmaI-digested plasmid pTSR1 (Kormanec et al., 1998
) containing the Streptomyces azureus tsr gene conferring resistance to thiostrepton, to create pRPO7-11S. The region upstream of the gapR gene was cloned as a 1250 bp EcoRISmaI fragment (Fig. 1a
) into EcoRI- and SmaI-digested plasmid pAPHII1 (Kormanec et al., 1998
) containing the kanamycin-resistance gene of Tn5, to create pRPO7-11T. The resulting plasmid, pRPO7-11U, was prepared by inserting a 2400 bp XbaIXhoI (blunt-ended) fragment from pRPO7-11S into pRPO7-11T cut with XbaI and SacI (blunt-ended). The plasmid pRPO7-11U was used to transform S. aureofaciens protoplasts as described by Kormanec et al. (1993)
. Thiostrepton-resistant clones were further analysed for thiostrepton resistance and kanamycin sensitivity, which might indicate a double crossover event. Two kanamycin-sensitive clones were identified, and correct integration was confirmed by Southern blot hybridization. Both clones had a similar phenotype, and one clone, S. aureofaciens
gapR, was chosen for further study.
|
Preparation of cell-free extracts.
Liquid-grown S. aureofaciens mycelium was harvested by centrifugation at 12000 g for 10 min, and washed by ice-cold STE buffer (10 mM Tris, 100 mM NaCl, 1 mM EDTA, pH 8). The mycelium was disrupted by sonication on ice (30 min total time, 30 s at amplitude 22 microns and 100 s pause; model Soniprep 150, MSE). Cell debris was then removed by centrifugation for 30 min at 30000 g. The cell-free extracts were stored in aliquots at -70 °C.
Protein analysis.
Protein concentrations were determined according to Bradford (1976) , with BSA as a standard. Denaturing SDS-PAGE of proteins was done as described by Laemmli (1970)
, and gels were stained with Coomassie blue R250. GAPDH was assayed by the arsenolysis procedure; 1 unit (U) is defined as the amount of enzyme which reduces 1 mmol NAD+ min-1 at A340 (Byers, 1982
).
Preparation of radiolabelled DNA fragments for GapR-binding studies.
A 291 bp gap-P promoter DNA fragment (positions -290 to +1 bp, in relation to the tsp of the gap-P promoter; Fig. 1a) was generated by PCR from plasmid pRPO7-11A (Kormanec et al., 1995
). For labelling of a coding strand, a 5'-end-labelled oligonucleotide primer, mut73, internal to gapR (5'-CACCGGGTACACGCCGATGC-3') and an unlabelled oligonucleotide, mut75, from the gap-P promoter region (5'-CCACCAGGTTCCCCCGCTGG-3') were used. For labelling of a noncoding strand, oligonucleotide mut75 was 5'-end-labelled, and mut73 was unlabelled. The oligonucleotides were end-labelled with [
-32P]ATP (Amersham; 111 TBq mmol-1) and T4-polynucleotide kinase (Biolabs) as described in Ausubel et al. (1987)
. The labelled fragments were purified by PAGE as described in Kormanec (2000)
.
Gel mobility-shift assay.
The assays were done as described by Ausubel et al. (1987) . 32P-labelled DNA fragments (0·5 ng, 500010000 c.p.m.) were incubated with cell-free extracts or partially purified GapR protein for 15 min at 30 °C in 15 µl total volume of the binding buffer (12·5 mM Tris pH 7·9, 60 mM KCl, 1 mM EDTA, 1 mM DTT, 12% glycerol) with 2 µg sonicated salmon sperm DNA and 4·5 µg BSA added. After incubation, protein-bound and free DNA were resolved on nondenaturing polyacrylamide gels (4% acrylamide, 0·05% bisacrylamide and 2·5% glycerol), running in a high-ionic-strength buffer (50 mM Tris, 380 mM glycine and 2 mM EDTA, pH 8·5) at 4 °C. The gels were dried and exposed to an X-ray film.
DNaseI footprinting.
Binding reactions were performed under the same conditions as for the gel mobility-shift assays with 4 ng 32P-labelled DNA fragments (1000030000 c.p.m.), in 30 µl binding buffer. After incubation, 3 µl DNase I solution [5 U DNase I ml-1 (Boehringer Mannheim) in 100 mM MgCl2, 100 mM DTT] was added to the binding reaction. The reaction mixture was incubated for 40 s at 37 °C, and stopped with 7·5 µl DNase I stop buffer (3 M ammonium acetate, 0·25 M EDTA, 0·1 mg tRNA ml-1), and extracted with 30 µl alkaline phenol/chloroform. The aqueous phase was precipitated with 3 vols ethanol. The resulting pellet, after washing with 70% ethanol and Speed Vac drying, was suspended in 5 µl Maxam loading buffer (80% formamide; 1 mM EDTA; 10 mM NaOH; 0·05%, w/v, bromphenol blue; 0·05%, w/v, xylene cyanole FF). The DNA fragments were analysed on 6 % DNA sequencing gels together with G+A and T+C sequencing ladders derived from the end-labelled fragments (Maxam & Gilbert, 1980 ). After electrophoresis, the gels were dried and exposed to an X-ray film.
Expression of gapR in E. coli.
The S. aureofaciens gapR gene was mutated to introduce a single NdeI site in the translational start codon using a mutagenic primer, mut79 (5'-GTACGAGGTACATATGTGCGGCGCGG-3'). To produce N-terminal His-tagged fusion GapR protein, the gene was cloned as a 1200 bp NdeISacI fragment in E. coli expression plasmid pET28a (Novagen) cut with the same enzymes, resulting in plasmid pET-gapR1. The DNA sequence of the fusion region was verified. The host strain for the pET series expression plasmids, E. coli BL21(DE3) pLysS, transformed with the plasmid, was grown in LB medium (Ausubel et al., 1987 ) containing 30 µg chloramphenicol ml-1 and 40 µg kanamycin ml-1 at 30 °C until OD600 0·5. Expression was induced with 1 mM IPTG. After 3 h, the cells were harvested by centrifugation at 12000 g for 10 min, and washed with ice-cold STE buffer. The pelleted cells were suspended in the binding buffer, and disrupted by sonication. The cell lysates were centrifuged for 30 min at 30000 g, and the supernatants were stored in aliquots at -70 °C. The purification of His-tagged GapR protein on His-Tag Bind resin (Novagen) under denaturing conditions was carried out as directed by the manufacturer.
Partial purification of GapR.
E. coli cells containing pET-gapR1 (2 g) were suspended in 10 ml buffer A (20 mM Tris/HCl pH 8·6, 1 mM EDTA, 20 mM KCl, 12% glycerol, 5 mM mercaptoethanol), and disrupted by sonication on ice. Following centrifugation at 15000 g for 30 min, the supernatant was applied to a DEAE-cellulose column (25 ml; DE 52 Whatman) equilibrated with buffer A. After the column had been washed with the buffer, proteins were eluted with a linear gradient of KCl from 0 to 1 M in a total volume of 500 ml at a flow rate of 1 ml min-1. Fractions containing DNA-binding activity (24 ml) were pooled, concentrated and dialysed overnight against binding buffer. The sample was stored at -70 °C.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
|
|
Expression of the gap gene in the S. aureofaciens gapR mutant
We have previously shown that a single gap-P promoter directing transcription of gap is induced both by glucose, and at the onset of aerial mycelium formation (Kormanec et al., 1997 ). To investigate whether gapR disruption has an effect on gap transcription, S1-nuclease mapping was performed using RNA isolated from wt and
gapR S. aureofaciens strains during differentiation on solid medium, and grown in liquid minimal medium NMP with mannitol or glucose as a carbon source to different growth phases (Fig. 3
). As shown in Fig. 3(b)
, a single RNA-protected fragment corresponding to the gap-P promoter (Kormanec et al., 1997
) was identified using probe 2 with the RNA isolated from the wt strain. The level of gap mRNA was substantially increased in the wt with glucose in the medium, and at the onset of aerial mycelium formation, as described previously (Kormanec et al., 1997
). However, only a very weak RNA-protected fragment (visible after overexposure of the autoradiogram) was identified with all RNAs from the
gapR mutant strain (Fig. 3b
). Its intensity was comparable irrespective of carbon source used or stage of differentiation. The results indicate that the
gapR mutation dramatically affected transcription from the gap-P promoter. The level of gap mRNA is much lower in the
gapR mutant, irrespective of carbon source or developmental stage. As a control, S1-nuclease mapping was performed with the same RNA samples using a probe fragment specific for the S. aureofaciens hrdB-P2 promoter, which is expressed fairly constantly during differentiation (Kormanec & Farka
ovský, 1993
). RNA-protected fragments corresponding to the hrdB-P2 promoter were identified with all RNA samples (Fig. 3c
). Moreover, a similar pattern of gap-P expression to the wt was identified using RNA isolated from the S. aureofaciens
gapR mutant strain with the gapR gene introduced in trans (data not shown).
These results were in contrast with the phenotype of the gapR mutant strain. GAPDH is an essential glycolytic enzyme for carbon flux in primary metabolism, and such a dramatic decrease of gap expression should strongly affect growth of the mutant strain on glycolytic carbon sources. However, the growth rate of the
gapR mutant was only partially decreased (see above). Therefore, we measured phosphorylating GAPDH activity in cell-free extracts from the wt and
gapR strains grown for 20 h in liquid minimal medium NMP with mannitol or glucose as a carbon source. Similar to results of transcriptional analysis, the specific activity of GAPDH was about 2·5 times higher in the presence of glucose (4·01±0·35 U mg-1) than with mannitol (1·81±0·22 U mg-1) in the wt strain. However, although glucose induction of GAPDH activity was detected in the
gapR strains, the specific activity was decreased (2·6±0·28 U mg-1 with glucose, and 1·17±0·12 U mg-1 with mannitol). The results clearly show that the
gapR mutation also affects GAPDH activity, but S. aureofaciens seems to contain another glucose-inducible GAPDH-encoding gene, gapR-independent expression of which is sufficient to ensure growth. Our previous hybridization analysis suggested the presence of a second gap gene in S. aureofaciens (Kormanec et al., 1995
).
Overproduction of GapR in E. coli, and its binding to the gap promoter region
To investigate whether GapR acts directly on gap-P induction, we overproduced this protein in E. coli and probed its binding in the gap-P promoter region. The S. aureofaciens GapR was overproduced as an N-terminal fusion with the 6xHis tag in E. coli using a T7 RNA polymerase expression system. Total protein extracts of E. coli transformed with the plasmid pET-gapR1 and the cloning plasmid pET28a, before and after induction with IPTG at 30 °C, were examined by SDS-PAGE. A prominent band with increasing intensity (with a maximum after 3 h) was clearly visible after induction with IPTG in the region corresponding to a molecular mass of 32 kDa (Fig. 5a). This value corresponds to the calculated Mr of the 6xHis-tagged GapR protein. However, almost all His-tagged GapR protein was found in the insoluble fraction. The amount of the soluble form did not significantly increase at lower temperature, or after coexpression with groEL, groES or trx genes (data not shown). We also failed to renaturate the 6xHis-tagged GapR protein isolated under denaturating conditions by Ni2+-affinity chromatography, employing various conditions. This insolubility is a typical characteristic of regulators within the AraC/XylS family and has hampered the biochemical analysis of these proteins (Gallegos et al., 1997
). However, when soluble cell-free protein extracts of E. coli transformed with plasmid pET-gapR1 and pET28a, respectively, were used in a gel retardation assay with the 32P-labelled 291 bp gap-P promoter DNA fragment (positions -290 to +1 bp, in relation to the tsp of the gap-P promoter; Fig. 1a
), a retarded band was clearly visible only with the cell-free extracts of E. coli containing plasmid pET-gapR1 (Fig. 5b
). The specificity of the interaction was demonstrated by the competitive binding of the unlabelled fragment (Fig. 5b
, lane 6). These results indicate that a small residual portion of GapR is in a soluble form and is capable of binding to the gap-P promoter region. However, we were unable to purify it by Ni2+-affinity chromatography under native conditions.
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Recent experiments in Gram-negative E. coli and low-GC Gram-positive B. subtilis revealed that the expression of gap is induced by glucose. However, this induction is caused by different mechanisms in these bacteria. In E. coli, glucose induction of both gap genes, gapA and gapB, depends upon a component of the PTS system, the EIIGlc protein. However, this dependence is indirect. Glucose is assumed to function as an external signal modulating the phosphorylation state of EIIGlc, which transmits the signal to the yet unknown regulator (Charpentier et al., 1998 ). In B. subtilis, two gap genes were identified encoding GAPDHs with opposite physiological roles: glycolytic gapA and gluconeogenic gapB. While gapA is induced, gapB is strongly repressed by glucose (Fillinger et al., 2000
). This glucose induction of the glycolytic gapA is indirectly dependent on the catabolite control protein CcpA (Fillinger et al., 2000
; Tobisch et al., 1999
). The glycolytic gapA gene is coexpressed in an operon with the upstream gene cggR, which encodes a repressor of the cggRgapA operon. CggR belongs to the SorC/DeoR family of transcriptional regulators. The activity of this repressor is inhibited by glucose, and it is assumed that the role of CcpA in glucose induction is mediated by CggR (Fillinger et al., 2000
). Likewise, expression of gap is also induced by glucose in high-GC Gram-positive S. aureofaciens (Kormanec et al., 1997
). However, in contrast to B. subtilis, the glycolytic gap gene is monocistronic in S. aureofaciens (Kormanec et al., 1997
), and an activator protein of the AraC/XylS family, GapR, which directly binds to the gap promoter region, regulates its expression. Considering S1-nuclease mapping experiments in S. aureofaciens wt and
gapR mutant strains, transcription of gap is substantially reduced in the
gapR mutant, irrespective of carbon source used for cultivation. Only a very low residual amount of gap mRNA was detected in the mutant strain, visible only after overexposure of the gel. Based on these results, it seems that the GapR protein is absolutely required for gap transcription. The mechanism of glucose induction in S. aureofaciens still remains unclear. However, GapR protein may ensure gap expression at an uninduced level, irrespective of the carbon source used. In the presence of glucose, an unknown signal might be transduced to the GapR protein by mechanisms similar to those previously described for E. coli or B. subtilis (Charpentier et al., 1998
; Fillinger et al., 2000
). Such modified GapR might ensure induced gap expression.
In addition to the glucose induction, the gap-P promoter is induced at the onset of aerial mycelium formation (Kormanec et al., 1997 ), and this induction is also substantially decreased in the
gapR mutant. A possible explanation for this induction might be connected with the phase I glycogen degradation that occurs in the course of Streptomyces differentiation in substrate hyphae that undergo aerial mycelium formation (Homerová et al., 1996
; Plaskitt & Chater, 1995
). Thus, GapR might respond to a common product of glucose and glycogen catabolism that is present in increased amounts either in a high glucose uptake in substrate mycelium grown in the presence of glucose, or after phase I glycogen degradation at the beginning of aerial mycelium formation. It is assumed that glycogen accumulation and degradation may play a role in morphological differentiation (Homerová et al., 1996
; Plaskitt & Chater, 1995
), and this proposed linking of gap-P induction indicates physiological relevance of regulation of glycolysis in connection with differentiation.
Considering the results of the DNaseI footprinting analysis, the proposed binding site of GapR in the gap-P promoter spans a region from -73 to -28 bp (for the coding strand), and -82 to -33 bp (for the noncoding strand) (Fig. 4a). This protected region contains a tandem repeat (Fig. 4b
) that could be the GapR-binding consensus sequence. Moreover, a similar tandem repeat was identified in the proposed binding site of the S. griseus AraC/XylS homologue AdpA, in the strR promoter (Ohnishi et al., 1999
; Vujaklija et al., 1993
) (Fig. 4b
). Based on these preliminary binding studies, it is difficult to suggest a consensus element for the GapR-binding site. However, considering the similarity between these two proposed binding sites for GapR and AdpA, this tandem repeat might constitute a consensus sequence of the GapR-binding site. No similarity to any of the other identified binding sites for AraC/XylS transcriptional activators was found, but binding sites for this family are highly variable and, in general, unique for each homologue (Gallegos et al., 1997
). The GapR-protected region overlaps the -35 hexamer of the gap-P promoter. Thus, surface residues of GapR, like those of many activators, could contact RNA polymerase. This binding is similar to several other activators of AraC/XylS (Gallegos et al., 1997
).
Considering the comparison of GapR with other AraC/XylS homologues (Fig. 2), it is conceivable that this type of gap regulation also exists in other Streptomyces species, since two AraC/XylS-homologous proteins discovered by genome sequencing of S. coelicolor (www.sanger.ac.uk/Projects/S_coelicolor/) are very similar to GapR in the N-terminal domain. This domain is generally responsible for an effector binding, and AraC/XylS homologues that are similar in this domain also have similar functions (Gallegos et al., 1997
). Searching the nucleotide sequence around these two S. coelicolor genes did not reveal any gap gene or other glycolytic gene. However, one of these proteins might activate expression of a gap gene located in a different position in the S. coelicolor genome. Based on the present data of the genome sequencing project, S. coelicolor also contains two gap orthologues. However, we did not find any similarity between the S. aureofaciens gap-P promoter (including the GapR-binding site) and the regions upstream from the S. coelicolor gap orthologues. An intriguing finding was the high sequence similarity between S. aureofaciens GapR and AdpA of S. griseus (Ohnishi et al., 1999
) in this N-terminal region (Fig. 2
). The AdpA protein is an A-factor-responsive transcriptional activator of the AraC/XylS family, which binds to the promoter of the pathway-specific regulatory gene strR responsible for transcription of streptomycin biosynthetic genes in S. griseus. Disruption of adpA affected streptomycin biosynthesis and morphological differentiation in S. griseus (Ohnishi et al., 1999
; Vujaklija et al., 1993
). This phenotype is in contrast with the results of gapR disruption, which did not affect sporulation and production of secondary metabolites in S. aureofaciens. Thus, it seems that these two proteins are not homologous in their function. Moreover, compared to other AraC/XylS homologues, S. griseus AdpA has an extraordinary long C-terminal DNA-binding domain (Fig. 2
). In this respect, it is interesting that the binding sites of both proteins (GapR and AdpA) are similar (Fig. 4b
). This might reflect sequence similarity of these proteins in proposed
-helixturn
-helix DNA-binding domains (Fig. 2
).
An interesting finding was that, though gap transcription was substantially reduced in the gapR mutant, the mutation only partially affected growth of S. aureofaciens, irrespective of carbon source used. Estimation of GAPDH activity actually revealed a decrease of enzyme activity in the
gapR mutant, but the residual activity was still high enough to ensure growth. These results indicate the presence of a second gap gene in S. aureofaciens, as was also suggested by hybridization studies (Kormanec et al., 1995
). Based on the residual GAPDH activity in cell-free extracts in the
gapR mutant, the second gap gene seems to be induced by glucose.
![]() |
ACKNOWLEDGEMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. O., Seidman, J. S., Smith, J. A. & Struhl, K. (1987). Current Protocols in Molecular Biology. New York: Wiley.
Baylis, H. A. & Bibb, M. J. (1987). The nucleotide sequence of a 16S rRNA gene from Streptomyces coelicolor A3(2). Nucleic Acids Res 15, 1716.
Boschi-Muller, S., Azza, S., Pollastro, D., Corbier, C. & Branlant, G. (1997). Comparative enzymatic properties of GapB-encoded erythrose-4-phosphate dehydrogenase of Escherichia coli and phosphorylating glyceraldehyde-3-phosphate dehydrogenase. J Biol Chem 272, 15106-15112.
Bradford, M. M. (1976). A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72, 248-254.[Medline]
Branlant, G., Flesch, G. & Branlant, C. (1983). Molecular cloning of the glyceraldehyde-3-phosphate dehydrogenase genes of Bacillus stearothermophilus and Escherichia coli, and their expression in Escherichia coli. Gene 25, 1-7.[Medline]
Byers, L. D. (1982). Glyceraldehyde-3-phosphate dehydrogenase from yeast. Methods Enzymol 89, 326-335.[Medline]
Charpentier, B. & Branlant, C. (1994). The Escherichia coli gapA gene is transcribed by the vegetative RNA polymerase holoenzyme E70 and by the heat shock RNA polymerase E
32. J Bacteriol 176, 830-839.[Abstract]
Charpentier, B., Bardey, V., Robas, N. & Branlant, C. (1998). The EIIGlc protein is involved in glucose-mediated activation of Escherichia coli gapA and gapB-pgk transcription. J Bacteriol 180, 6476-6483.
Chater, K. F. (1998). Taking a genetic scalpel to the Streptomyces colony. Microbiology 144, 1465-1478.
Della Seta, F., Boshi-Muller, S., Vignais, M. L. & Branlant, G. (1997). Characterization of Escherichia coli strains with gapA and gapB genes deleted. J Bacteriol 179, 5218-5221.[Abstract]
Fillinger, S., Boschi-Muller, S., Azza, S., Dervyn, E., Branlant, G. & Aymerich, S. (2000). Two glyceraldehyde 3-phosphate dehydrogenases with opposite physiological roles in a nonphotosynthetic bacterium. J Biol Chem 275, 14031-14037.
Gallegos, M.-T., Schleif, R., Bairoch, A., Hofmann, K. & Ramos, J. L. (1997). AraC/XylS family of transcriptional regulators. Microbiol Mol Biol Rev 61, 393-410.[Abstract]
Harris, J. I. & Waters, M. (1976). Glyceraldehyde-3-phosphate dehydrogenase. In The Enzymes , pp. 1-49. Edited by P. D. Boyer. New York:Academic Press.
Homerová, D., Benada, O., Kofroová, O.,
e
uchová, B. & Kormanec, J. (1996). Disruption of a glycogen branching enzyme gene, glgB, specifically affects the sporulation-associated phase of glycogen accumulation in Streptomyces aureofaciens. Microbiology 142, 1201-1208.
Hopwood, D. A., Bibb, M. J., Chater, K. F. & 7 other authors (1985). Genetic Manipulation of Streptomyces: a Laboratory Manual. Norwich: The John Innes Foundation.
Horinouchi, S., Hara, O. & Beppu, T. (1983). Cloning of a pleiotropic gene that positively controls biosynthesis of A-factor, actinorhodin, and prodigiosin in Streptomyces coelicolor A3(2) and Streptomyces lividans. J Bacteriol 155, 1238-1248.[Medline]
Kang, J.-G., Hahn, M.-Y., Ishihama, A. & Roe, J.-H. (1997). Identification of sigma factors for growth phase-related promoter selectivity of RNA polymerases from Streptomyces coelicolor A3(2). Nucleic Acids Res 25, 2566-2573.
Kormanec, J. (2000). Analyzing the developmental expression of sigma factors with S1-nuclease mapping. In Nuclease Methods and Protocols. Methods in Molecular Biology , pp. 498-513. Edited by C. H. Chein. Totowa, NJ:Humana Press.
Kormanec, J. & Farkaovský, M. (1993). Differential expression of principal sigma factor homologues of Streptomyces aureofaciens correlates with the developmental stage. Nucleic Acids Res 21, 3647-3652.[Abstract]
Kormanec, J., e
uchová, B. & Farka
ovský, M. (1993). Optimization of Streptomyces aureofaciens transformation and disruption of the hrdA gene encoding a homologue of the principal
factor. J Gen Microbiol 139, 2525-2529.
Kormanec, J., Lempelová, A., Farkaovský, M. & Homerová, D. (1995). Cloning, sequencing and expression in Escherichia coli of a Streptomyces aureofaciens gene encoding glyceraldehyde-3-phosphate dehydrogenase. Gene 165, 77-80.[Medline]
Kormanec, J., Lempelová, A., Nováková, R., e
uchová, B. & Homérová, D. (1997). Expression of the Streptomyces aureofaciens glyceraldehyde-3-phosphate dehydrogenase gene (gap) is developmentally regulated and induced by glucose. Microbiology 143, 3555-3561.[Abstract]
Kormanec, J., ev
íková, B., Spru
anský, O., Benada, O., Kofro
ová, O., Nováková, R.,
e
uchová, B., Potú
ková, L. & Homérová, D. (1998). The Streptomyces aureofaciens homologue of the whiB gene is essential for sporulation and its expression correlates with the developmental stage. Folia Microbiol 43, 605-612.
Laemmli, U. K. (1970). Cleavage of structural proteins during assembly of the head of bacteriophage T4. Nature 227, 680-685.[Medline]
Maxam, A. M. & Gilbert, W. (1980). Sequencing end-labelled DNA with base specific chemical cleavages. Methods Enzymol 65, 449-560.[Medline]
Nováková, R., ev
íková, B. & Kormanec, J. (1998). A method for the identification of promoters recognized by RNA polymerase containing a particular sigma factor: cloning of a developmentally regulated promoter and corresponding gene directed by the Streptomyces aureofaciens sigma factor RpoZ. Gene 208, 43-50.[Medline]
Ohnishi, Y., Kameyama, S., Onaka, H. & Horinouchi, S. (1999). The A-factor regulatory cascade leading to streptomycin biosynthesis in Streptomyces griseus: identification of a target gene of the A-factor receptor. Mol Microbiol 34, 102-111.[Medline]
Plaskitt, K. A. & Chater, K. F. (1995). Influences of developmental genes on localized glycogen deposition in colonies of a mycelial procaryote, Streptomyces coelicolor A3(2): a possible interface between metabolism and morphogenesis. Philos Trans R Soc Lond B 347, 105-121.
Tobisch, S., Zuhlke, D., Bernhardt, J., Stulke, J. & Hecker, M. (1999). Role of CcpA in regulation of the central pathways of carbon catabolism in Bacillus subtilis. J Bacteriol 181, 6996-7004.
Vujaklija, D., Horinouchi, S. & Beppu, T. (1993). Detection of an A-factor-responsive protein that binds to the upstream activation sequence of strR, a regulatory gene for streptomycin biosynthesis in Streptomyces griseus. J Bacteriol 175, 2652-2661.[Abstract]
Wright, F. & Bibb, M. J. (1992). Codon usage in the G+C rich Streptomyces genome. Gene 113, 55-65.[Medline]
Received 4 October 2000;
revised 9 January 2001;
accepted 29 January 2001.
HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
INT J SYST EVOL MICROBIOL | MICROBIOLOGY | J GEN VIROL |
J MED MICROBIOL | ALL SGM JOURNALS |