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Previous Article | Next Article 
Journal of Bacteriology, October 2006, p. 7111-7122, Vol. 188, No. 20
0021-9193/06/$08.00+0 doi:10.1128/JB.00574-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Tomoyasu Nishizawa,
Kosaku Takahashi,
Takeshi Hosaka,
Hiroyuki Aoki,¶ and
Kozo Ochi*
National Food Research Institute, Tsukuba, Ibaraki 305-8642, Japan
Received 24 April 2006/ Accepted 31 July 2006
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ABSTRACT |
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INTRODUCTION |
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Upon binding to RNA polymerase, ppGpp inhibits transcription of one set of genes and stimulates transcription of another. Although inhibition of transcript elongation by ppGpp has been reported, ppGpp acts primarily at the stage of transcription initiation, during the formation of the open promoter complex and/or during the first few rounds of RNA synthesis (7, 60). It is thought, therefore, that the major specificity determinants for regulation by ppGpp are intrinsic to negatively regulated promoters, which are distinguished by two characteristic features: the presence of a guanosine- and cytosine-rich discriminator sequence situated between the –10 promoter region and the transcription start site and the formation of short-lived open complexes (4, 7, 57). Although the dependence of ppGpp control on these features has been experimentally established for some promoters, this dependence is not absolute, and the features themselves are not well conserved. Additional evidence of the role of ppGpp in control of rRNA synthesis comes from genetic studies in which a variety of rpoB (encoding the RNA polymerase ß subunit) mutants that confer rifampin resistance were isolated and analyzed. These rpoB mutations frequently circumvent the ppGpp0 phenotype (i.e., inability to grow in a chemically defined medium or to produce antibiotics), suggesting that the mutant enzymes behave like "stringent" RNA polymerases (3, 20, 31, 34, 65, 68) and that RNA polymerase mutants could be subject to stringent control. Also noteworthy is the recent finding by Jishage et al. (23) that in E. coli, alternative 
factors compete against 70 significantly better in the presence of ppGpp, which is suggestive of a ppGpp-dependent alteration in factor competition for binding to the RNA polymerase core. Recently, we reported that the intracellular ppGpp level is fine-turned by EshA, which is capable of binding cyclic aMP and which is an important protein for triggering antibiotic production in Streptomyces spp. (50a).
Despite much investigation (8, 24, 56), until recently the binding site for ppGpp in RNA polymerase remained undefined, and so the mechanism by which ppGpp selectively regulates the transcription of a large number of genes remained obscure. However, through the collaborative efforts of three laboratories, including ours, new insights into the mechanism of transcriptional regulation by ppGpp have been gained from a structural analysis of the Thermus thermophilus RNA polymerase holoenzyme in complex with ppGpp (2). The results indicate that (i) ppGpp binds to a single site on the RNA polymerase surface adjacent to, but not overlapping, the active center in two alternative orientations and that (ii) base pairing of ppGpp with cytosines in the nontemplate DNA strand might be an essential component of transcriptional control by ppGpp. Because this structural analysis of RNA polymerase was carried out using only thermophilic bacteria (T. thermophilus or Thermus aquaticus), our aim in the present study was to clarify the physiological basis of the stringent response in this thermophilic bacterial group.
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MATERIALS AND METHODS |
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Preparation of thiostrepton-resistant mutants. Spontaneous thiostrepton-resistant mutants were obtained from colonies that grew within 3 days after cells were spread on MTM agar medium containing 0.3 µg/ml or 3 µg/ml thiostrepton. To detect the mutations, a 5' part of the rplK gene was amplified by PCR using the oligonucleotide primers TTL11f (5'-ATGAAGAAAGTTGTTGCGG-3') and TTL11r (5'-CGTGCTTAGGCATCCTTCAC-3'), which were based on the sequence described previously (18), since spontaneous rplK mutations conferring the resistance to thiostrepton (or thiopeptin) in bacteria have always been found in the 5' part of the rplK gene (see Results). The reaction mixture (50 µl) contained GCI buffer (Takara), 20 ng of total DNA, 0.2 mM (each) deoxynucleoside triphosphate, 1 mM (each) primer, and 2.5 U of LA-Taq DNA polymerase (Takara). Amplification was carried out in a GeneAmp PCR system 9700 (Applied Biosystems) using a protocol that entailed initial denaturation at 98°C for 2 min followed by 30 cycles of denaturation at 98°C for 20 s, annealing at 58°C for 10 s, and elongation at 72°C for 30 s. The DNA was then sequenced using a DYEnamic ET terminator cycle sequencing premix kit (Amersham Biosciences) and an ABI PRISM 310 genetic analyzer (Applied Biosystems).
Genetic transformation of T. thermophilus. T. thermophilus wild-type strain HB8 was transformed to be thiostrepton resistant as described by Koyama et al. (28). Strain HB8 was cultured overnight, diluted 100-fold with fresh TM medium, and incubated with shaking at 70°C for 4 h. Then, the culture was mixed with chromosomal DNA (final concentration of 10 µg/ml) prepared from thiostrepton-resistant mutant cells. DNA was isolated by the method of Saito and Miura (50). The mixture was then incubated with shaking at 70°C for 1 h, followed by cooling on ice. Samples were appropriately diluted with TM medium and plated on a TM agar plate containing 1 µg/ml thiostrepton for the detection of thiostrepton-resistant transformants, which developed after 48 h of incubation at 70°C.
Chromosomal DNA isolation and manipulation. Standard methods for DNA isolation and manipulation were used as described by Sambrook et al. (51). Chromosomal DNA extraction for PCR amplification was carried out using an InstaGene Matrix resin (Bio-Rad) according to the instruction manual. DNA fragments were isolated from agarose gels using a QIAEX II gel extraction kit (QIAGEN).
Construction of RSHTt (relA) disruptants. The plasmids used to disrupt the T. thermophilus relA/spoT homolog (RSHTt) through homologous recombination were constructed as follows. Based on the database nucleotide sequence, internal PCR primers for RSHTt and a thermostable kanamycin-resistant marker gene (HTK) were generated. A pBluescript SK(–) plasmid vector (Toyobo) was then used to clone PCR products in which an EcoRV site was introduced by PCR using the following primers (restriction sites are underlined, and modified sequences are in italics): TNP-020 (for the promoter region of RSHTt) (5'-TAATCTAGACGGGTAGGGGTGCTACACTAAAAGTAGTGG-3'), TNP-021 (reverse primer for RSHTt) (5'-CAAGATGGGAGTGGGGCATGGTTAGG-3'), TNP-024 (for the 5' end of HTK) (5'-CCGCTCGAGCGTTGACGGCGGATATGGTA-3'), and TNP-025 (for the 3' end of HTK) (5'-AATCTCGAGCGTAACCAACATGATTAACA-3'). PCR with chromosomal DNA from T. thermophilus HB8 and the pUC18-HTK vector for HTK was carried out with TaKaRa PrimeSTAR HS DNA polymerase (Takara) as instructed by the manufacturer. The reactions were run in a GeneAmp PCR 9700 system (Applied Biosystems). The amplification protocol typically entailed 25 cycles of denaturation at 98°C for 10 s, annealing at 56°C for 5 s, and extension at 72°C for 2.5 min. The PCR fragment containing RSHTt was digested with HindIII-XbaI and cloned into the HindIII-XbaI sites of pBluescript SK(–), yielding pTN1018. The 2.3-kb HindIII-XbaI fragment containing RSHTt was subcloned into the HindIII-XbaI site of pUC119 (Toyobo), yielding pTN1022. To add the restriction site in the Kanr gene cassette (HTK), PCR amplification was performed using primers TNP-024 and TNP-025 based on the pUC118-HTK, and then the 1.0-kb fragment was cloned into the pBluescript SK(–)/EcoRV site, yielding pTN1020. The 1.0-kb XhoI HTK(XhoI) fragment from pTN1020 was inserted into the XhoI sites (in RSHTt), generating pTN1023. The respective plasmids for gene disruption were introduced into T. thermophilus strains HB8 (wild type) and KO-572 (Phe–), after which kanamycin (50 µg/ml)-resistant transformants were selected as described by Hashimoto et al. (17). An overnight culture was diluted 50-fold with one-half-diluted MTM medium (1/2 MTM) and shaken at 70°C until the optical density at 650 nm (OD650) reached 0.45 to 0.55. This culture (400 µl) was mixed with 1.0 µg of gene disruption plasmid DNA, after which the mixture was incubated at 70°C for 2 h and then spread on 1/2 MTM agar medium containing 50 µg/ml kanamycin. The plates were then incubated at 70°C for 24 h. Disruption of RSHTt was confirmed with primers TNP-030 (5'-GGAGGCCCTCAAAGAGTTGG-3') and TNP-031 (5'-GGAGTGGGGCATGGTTAGG-3') by detecting the HTK cassette in the expected region (central region) of RSHTt.
Real-time quantitative PCR (RT-qPCR) analysis. T. thermophilus cultures (100 ml), grown to an OD650 of 0.5 to 0.6, were treated with serine hydroxamate (final concentration, 10 mM) for the appropriate time, placed into ice-cold 50-ml tubes, and then centrifuged at 6,000 x g for 10 min at 4°C. The cell pellets were frozen and kept at –80°C until RNA extraction. At that time, the frozen pellet was incubated at room temperature for 30 min in lysozyme (0.5 mg/ml) solution. Total RNA was extracted using the ISOGEN-LS reagent (Nippon Gene) with chloroform, and the purification steps were carried out following the manufacturer's instructions. The extracted RNA was then purified further using an RNeasy kit with RNase-free DNase I (QIAGEN), after which the concentration of purified RNA was measured by spectroscopy at OD260. cDNA was generated from a 1-µg sample of the purified total RNA by reverse transcription (ReverTra Ace, 100 U per reaction; Toyobo) at 45°C using the reverse transcription primers TNP-035 (for the 23Sa and 23Sb rRNA genes) (5'-CTGAGATGTTTCAGTTCCC-3') and TNP-044 (for rpoD) (5'-GCCTTGTTCTCAATCTGCC-3') and then subjected to RNase H treatment (2 U per reaction). Samples (50 ng per assay [for rpoD] or 0. 5 ng per assay [for 23Sa-23Sb]) were then analyzed using the 7300 real-time PCR system and Power SYBR Green PCR master mix (Applied Biosystems). The PCR primers used were TNP-032 (the forward primer for 23Sa and 23Sb [5'-TGCCCTGAGGGGGGTAGC-3']), TNP-034 (the reverse primer for 23Sa and 23Sb [5'-CTGGCTTATCGCAGGTAGC-3']), TNP-042 (rpoD forward primer [5'-CAACCTCCGGCTCGTGGTCTCC-3']), and TNP-043 (rpoD reverse primer [5'-GGTGGCGTAGGTGGAGAAC-3']). The standard curves for quantification were calculated by serial dilution of plasmid, generating pTN1024 and pTN1027, which contain an amplified 23Sa and 23Sb rRNA nucleotide sequence (ca. 260 bp) and a rpoD nucleotide sequence (ca. 700 bp), respectively. These plasmids were constructed based on pBluescript SK(–). The amplification protocol entailed 1 cycle of 96°C for 1 min, followed by 40 cycles of 96°C for 30 s (step 1), 58°C for 30 s (step 2), and 72°C for 35 s (step 3). Data collection and real-time analysis were done at step 3. After the final cycle, melt curve data were obtained using an additional stage of dissociation, beginning at 56°C for 10 s and then incrementally increasing the temperature until 96°C.
Measurement of RNA synthesis. The RNA synthesis rate in growing cells was determined as previously described (38) by measuring the incorporation of [2-14C]uracil into acid-precipitated material. Cells were grown to mid-exponential phase (OD650 = 0.5) in a chemically defined medium (medium 162), after which [2-14C]uracil (0.1 µCi/ml, 100 µM), with or without serine hydroxamate (10 mM), was added to the culture and incubation continued with shaking. Samples (1 ml) were then removed at the indicated times and immediately added to 1 ml of ice-cold 10% trichloroacetic acid. The resultant precipitates were collected by filtration, washed with 5% trichloroacetic acid containing 10 mM cold uracil, dried, and counted in a liquid scintillation counter using clear-sol (Nakarai).
Assay of nucleotide pools. The intracellular concentrations of nucleotides, including ppGpp and pppGpp, were assayed by high-performance liquid chromatography (HPLC) after extraction with 1 M formic acid as previously described (38) and expressed as pmol per mg dry weight.
Ribosome-dependent in vitro ppGpp synthesis.
Ribosomes were isolated from exponential-phase T. thermophilus (grown in MTM medium) or E. coli cells (grown in L medium) as described previously (30). The resultant pellets of ribosomes were resolved in buffer A (50 mM Tris acetate [pH 8.0], 15 mM magnesium acetate, 60 mM potassium acetate, 30 mM ammonium acetate, 1 mM dithiothreitol, 0.2 mM EDTA, 0.5 mM phenylmethylsulfonylfluoride), and the ribosome concentration was determined using a spectrophotometer (at 260 nm) and expressed in terms of A260 (U/ml). Ribosome-dependent synthesis of ppGpp or pppGpp was carried out according to the method of Martinez-Costa et al. (35). The reaction mixture (50 µl), containing 2 mM ATP, 1.3 mM GTP, 10 µCi/ml [
-32P]GTP (3,000 Ci/mmol; Amersham Biosciences), 18% (vol/vol) methanol, and 60 A260 U/ml ribosome, was incubated for 30 min at the indicated temperature. The reaction was stopped by addition of 2 µl of 88% formic acid, after which the precipitated proteins were removed by centrifugation at 10,000 x g for 5 min. A 5-µl aliquot of the supernatant was subjected to thin-layer chromatography on polyethyleneimine-cellulose (Merck) with 1.5 M KH2PO4. Labeled mononucleotides were detected by autoradiography.
Construction of a plasmid for in vitro transcription assays. The region of the T. thermophilus 23S/5S rRNA operon (designated the 23Sb gene in the NCBI database [accession no. AP008226; previously accession no. M35674]) containing the promoter sequence was amplified by PCR using oligonucleotide primers TTrrn-1 (5'-CAGCAAAGCTTTGCCAGCGAAAAGGCCC-3') and TTrrn-2 (5'-ACATCCAGGGATCCACGCCCGCTAC-3'), which were based on the sequence described previously (16). The PCR product was digested using the corresponding restriction enzymes at the HindIII site generated by TTrrn-1 (underlined sequence) and an internal BamHI site, and the resultant fragment was cloned into the HindIII and BamHI sites of the pSP64-polyA vector (Promega), yielding pTRP, which was then sequenced to ensure the correctness of the nucleotide sequence.
Purification of RNA polymerase. RNA polymerase was purified according to the method of Burgess and Jendrisak (5) or Xue et al. (66) with some modification. All steps were carried out at 4°C. TGED buffer (0.01 M Tris-HCl [pH 7.9], 5% glycerol, 0.1 mM EDTA, and 0.1 mM dithiothreitol) was used as the basal buffer throughout the purification procedures. Polyethylenimine P-70 solution (30% [vol/vol] solution) was purchased from Wako Pure Chemicals, and a 10% (vol/vol) solution titrated with HCl to pH 7.9 was prepared and subjected to filtration through Miracloth (Calbiochem) to remove insoluble materials. Cells (10 g) in 40 ml of grinding buffer (TGED buffer plus 23 µg/ml phenylmethylsulfonylfluoride and 2 mg/ml lysozyme) were then disrupted using a French press at 12,000 lb/in2. After the resultant cell extract was centrifuged at 8,000 x g for 45 min and the supernatant was collected, 10% polyethylenimine P-70 solution (pH 7.9) was slowly added with stirring to a final concentration of 0.5%. The stirring was then continued for an additional 5 min before the mixture was centrifuged at 6,000 x g for 15 min. The resultant pellet was washed by suspending it in 40 ml of TGED buffer containing 0.2 M NaCl and centrifuged again at 6,000 x g for 15 min, after which the RNA polymerase was eluted with 40 ml of TGED buffer containing 1 M NaCl. The eluate was then centrifuged at 6,000 x g for 30 min, the supernatant collected, and solid ammonium sulfate added with stirring to 50% saturation. The mixture was then stirred for an additional 1 h and centrifuged at 8000 x g for 45 min, and the supernatant was discarded. To remove the nucleic acids, the pellet was washed with 40 ml of TGED buffer containing 2 M ammonium sulfate. After centrifugation, the pellet was resuspended in TGED buffer and dialyzed for 24 h against TGED buffer containing 0.15 M NaCl. Thereafter, a calf thymus DNA-cellulose (denatured; Amersham Biosciences) column (5 ml in volume) was equilibrated with 20 ml of TGED buffer containing 0.15 M NaCl. Each sample of dialyzate was pumped onto the column at a flow rate of 0.2 ml/min, washed with 20 ml of the same buffer, and then eluted with 40 ml of a 0.15 to 1 M NaCl linear salt gradient. Peak fractions were pooled and precipitated with 50% saturated ammonium sulfate at 0°C for 15 min with gentle stirring, centrifuged at 8,000 x g for 30 min, and dissolved in 1 ml of TGED buffer containing 0.5 M NaCl. The RNA polymerase sample thus obtained was then further fractionated using a superose 6 HR 10/30 gel filtration column with an AKTA explorer 10XT fast-protein liquid chromatography system (Amersham Biosciences). The sample was put onto the column, equilibrated with TGED buffer containing 0.5 M NaCl, and eluted at a flow rate of 0.1 ml/min. Peak fractions were collected and dialyzed for 24 h with storage buffer (the same as TGED buffer containing 0.1 M NaCl, except that the glycerol concentration was 50% instead of 5%), and the resultant enzyme solution was stored at –20°C until use.
Assay of RNA polymerase activity. RNA polymerase activity was assayed using the method of Barker et al. (4) with some modification. As template DNA, the 23S/5S rRNA operon region (23Sb gene) of plasmid pTRP was amplified by PCR using the oligonucleotide primers pSP64f (5'-ATTTAGGTGACACTATAGAATAC-3') and pSPr (5'-ACAGCTATGACATGATTACGAATTC-3') and then extracted twice with phenol-chloroform, precipitated with ethanol, and resuspended in 10 mM Tris-HCl (pH 8.0). Transcription was initiated by addition of RNA polymerase (5 µg) into 50 µl of reaction mixture containing 40 mM Tris-HCl (pH 8.0), 10 mM MgCl2, 1 mM dithiothreitol, 0.1 µg/µl bovine serum albumin, 200 mM NaCl, 0.02 µg of template DNA, 0.5 mM (each) nucleotide triphosphate (ATP, GTP, CTP, and UTP), and 2 µCi/ml of [3H]UTP. After addition of the indicated amount of ppGpp, pppGpp, or ppApp to the mixture, the reaction was run at 60°C until 10-µl samples of the reaction mixture were collected and mixed with an equal volume of stop solution (1% sodium dodecyl sulfate [SDS] and 50 mM sodium pyrophosphate) to stop the reaction. The samples were then applied to DE81 DEAE filters (Whatman), which were then washed three times with 5% Na2HPO4 and twice with water, dried, and counted in a liquid scintillation counter using clear-sol (Nakarai).
To confirm the specific rRNA transcript, an in vitro transcription assay was also carried out as an alternative method using polyacrylamide gel electrophoresis. Transcription was performed in 20 µl of reaction mixture containing 40 mM Tris-HCl (pH 8.0), 10 mM MgCl2, 1 mM dithiothreitol, 0.1 µg/µl bovine serum albumin, 100 mM NaCl, 1 µg of RNA polymerase, 8 ng of template DNA, 0.5 mM (each) ATP, GTP, CTP, and UTP, and 0.2 mCi/ml of [-32P]UTP. In some cases, ppGpp, ATP, or GTP was added. The reaction was run at 60°C for 20 min and mixed with 2 µl of 20% SDS to stop the reaction. The sample was treated with water-saturated phenol:chloroform:isoamyl alcohol (50:49:1, vol/vol/vol) and precipitated with ethanol, and the pellet was dissolved in 15 µl of water. After addition of the same volume of 2x formamide sample buffer (95% formamide, 18 mM EDTA, 0.025% SDS, 0.1% xylene cyanol, and 0.1% bromophenol blue), the transcripts were separated on 8 M urea-5% polyacrylamide gel. The radioactivity was detected with a BAS-2500 system (Fuji Film) and quantitated with Image Gauge version 3.41 software (Fuji Film).
Assay of IMP dehydrogenase activity. IMP dehydrogenase activity of T. thermophilus strain HB8 (grown to mid-exponential phase in MTM medium) was measured as described previously (37), except that the reaction was carried out at 50°C for 20 min in a reaction mixture with a pH adjusted to 8.5 instead of 8.0.
Reagents. ppGpp, pppGpp, and ppApp were prepared enzymatically in our laboratory using Streptomyces morookaense (44). HPLC analysis showed the purity of these samples to be >98% for ppGpp and pppGpp and 93% for ppApp.
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RESULTS |
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1%, harbored a P21R mutation and displayed a diminished capacity to accumulate ppGpp upon exposure to serine hydroxamate, confirming that the rplK mutation was responsible for the observed phenotype. Construction of relA disruptant. relA mutants, which harbor defects in their relA gene, encoding ppGpp synthetase I, together with relC mutants have been studied in great detail with respect to the stringent response represented by ppGpp accumulation (7). We therefore attempted to construct a T. thermophilus relA disruptant. T. thermophilus HB8 possesses a relA/spoT homologue (RSHTt) encoding a 727-amino-acid protein (KEGG Genes Database [http://www.genome.jp/]). Based on a DBGET search, RSHTt was found to encode a HD His-Arp domain (amino acids 49 to 151), which is characteristic of metal-dependent phosphohydrolases (1), a conserved RelA/SpoT region (248 to 371), a TGS (for "ThrRS, GTPase, and SpoT") domain (409 to 472), which is predicted to possess a predominant ß-sheet structure (52, 63), and an aspartokinase domain (656 to 726), which binds specifically to a particular amino acid, leading to regulation of the linked enzyme (53). We successfully constructed T. thermophilus relA disruptants (KO-571 and KO-652) (see Materials and Methods) and found that they grew as well as the wild-type strain in nutritionally rich media (e.g., MTM medium) but lost completely the ability to accumulate ppGpp, as indicated by the absence of an effect of serine hydroxamate. For that reason, we will hereafter refer to RSHTt as relA.
Phenotypic characterization of relA and relC mutants.(i) Changes in nucleotide pool size. That the ability of the relA disruptant (KO-571) and relC mutant (KO-564) to accumulate ppGpp was severely impaired (100% and 80% reduction, respectively) was confirmed by monitoring the changes in nucleotide levels after serine hydroxamate treatment (Fig. 3A). Importantly, levels of GTP declined markedly (to less than half) in the wild-type strain (and also in the relC mutant), possibly due to potent inhibition of GMP synthesis by ppGpp (41), while its level even increased in the relA disruptant, possibly due to the complete absence of ppGpp synthesis under the growth-limiting (GTP consumption-limiting) condition caused by serine hydroxamate. In fact, an increase in the size of the GTP pool (about twofold) was also detected by treating the cells with a high concentration (25 µg/ml each) of streptomycin or chloramphenicol (data not shown). On the other hand, levels of ATP increased 3.5-fold in the wild-type strain, presumably due to direct and/or indirect inhibition of RNA synthesis (see below) by ppGpp (Fig. 3A). No such dramatic changes in ATP levels were observed in the relA disruptant. The intracellular concentrations of ATP and GTP (just before the addition of serine hydroxamate) and of ppGpp (5 min after the addition) were calculated as being 1.2 mM, 0.17 mM, and 10 µM, respectively, according to the method described previously (37).
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(ii) RNA synthesis. One of the most characteristic components of the bacterial stringent response is suppression of RNA synthesis (7). Thus, RNA synthesis ([14C]uracil incorporation) was substantially reduced in wild-type T. thermophilus following treatment with serine hydroxamate (reduction rate = 60%) (Fig. 4). The same treatment reduced RNA synthesis in the relA disruptant by only 36% and in the relC mutant by an intermediate 52%.
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In vitro ppGpp synthesis using the T. thermophilus ribosome. We next attempted to clarify the intrinsic mechanism of ppGpp synthesis in T. thermophilus by examining ribosome-dependent ppGpp synthesis in vitro (Fig. 6). E. coli W3110 served as a reference strain in these experiments. Ribosomes were isolated from exponentially growing cells, and the reaction mediating ppGpp synthesis was carried out according to the method of Martinez-Costa et al. (35). It is evident from Fig. 6A that ribosomes from wild-type T. thermophilus have the ability to produce ppGpp in addition to much greater amounts of pppGpp when the reaction mixture was incubated at 50°C; less than 10% of the product was ppGpp. Neither ppGpp nor pppGpp was detected in the absence of ribosome. Surprisingly, only trace amounts of ppGpp and pppGpp were produced at 70°C, although that temperature is optimal for growth of T. thermophilus. The observed low productivity was not due to the lability of ppGpp and pppGpp, as only 10% or less of a standard sample in water or buffer solution was degraded after 1 h at 70°C. Ribosomes from E. coli and T. thermophilus were also capable of producing ppGpp and pppGpp at 30°C but at a much lower level than was produced by the latter at 50°C (Fig. 6A). Thus, ribosomes from wild-type T. thermophilus show an enormous capacity to produce pppGpp at 50°C.
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ppGpp inhibits IMP dehydrogenase activity. IMP dehydrogenase is the first enzyme of the pathway leading to GTP from IMP. Like the case for B. subtilis and Streptomyces (37, 41), IMP dehydrogenase from T. thermophilus was a target of (p)ppGpp. The activity of the T. thermophilus IMP dehydrogenase was measured with various concentrations of IMP (0.02 to 2 mM) in the presence or absence of (p)ppGpp (see Materials and Methods). The Ki values determined for ppGpp and pppGpp were 20 µM and 11 µM, respectively. These Ki values are somewhat lower than the previously reported Ki value (50 µM) for ppGpp for Streptomyces griseus (30), implying that ppGpp is capable of functioning more potently for T. thermophilus than for S. griseus. The marked and abrupt decline in the level of GTP in rel+ cells accompanied by ppGpp accumulation (Fig. 3 A and B) can be accounted for in this way.
ppGpp inhibits RNA polymerase activity. To date, a number of biochemical and genetic events have been attributed to the activity of (p)ppGpp, mainly on the basis of results with E. coli (7). Among (p)ppGpp's functions, inhibition of RNA polymerase is the most prominent, leading to severe inhibition of certain genes, including the rrn genes for rRNA synthesis. We therefore undertook an in vitro analysis of the effects of ppGpp and pppGpp (and ppApp) on expression of the T. thermophilus 23Sb gene (driven by its own promoter) coding for 23S/5S rRNA, although the rRNA gene promoters from this organism have not yet been well characterized (16).
We purified RNA polymerase from T. thermophilus cells growing exponentially in MTM medium (see Materials and Methods); an SDS-polyacrylamide gel electrophoresis profile of the enzyme fraction at each purification step is shown in Fig. 7. RNA polymerase activity was determined by measuring [3H]UTP incorporation in the presence and absence of (p)ppGpp or ppApp. We found that RNA polymerase-catalyzed transcription of the 23S/5S rRNA gene was severely inhibited by increasing levels of ppGpp or pppGpp such that the activity was inhibited by 80% in the presence of 1 mM (p)ppGpp (Fig. 8A). It is worth mentioning that relatively low concentrations of (p)ppGpp (e.g., 0.05 mM) also negatively influenced RNA polymerase activity, though the observed inhibition was slight. It is thus concluded that in T. thermophilus both ppGpp and pppGpp could act by exerting an inhibitory effect on rRNA gene transcription. By contrast, ppApp displayed no effect, even at a high (1 mM) concentration. This conclusion was confirmed using an alternative method, the detection of the specific transcripts on the gel, in which the runoff transcripts from in vitro transcription reactions were separated (see Materials and Methods). The gel separation profile indicating the specific transcripts and a quantification of the effects of various concentrations of ppGpp on the level of the runoff transcripts are shown in Fig. 8B and C, respectively.
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DISCUSSION |
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-phosphohydrolase), although there are alternative routes via which pppGpp is degraded (7). The preferential accumulation of pppGpp rather than ppGpp that we observed in vitro (Fig. 6A) may reflect the weak activity of, or the absence of, this enzyme in our reaction system. One aspect of the stringent response that has remained unclear is why there are significant differences (more than 10-fold) in the magnitudes of the effects of ppGpp on rRNA promoter activity in vitro and in vivo (4, 9): the inhibitory effect of ppGpp observed in vitro was always much smaller than that observed in vivo (4). This was also the case in the present study: whereas 0.1 mM ppGpp was required to substantially suppress rRNA synthesis in vitro (Fig. 8A and C), the maximum concentration of ppGpp observed 5 min after addition of serine hydroxamate to whole cells (Fig. 1 and 3) was only 10 to 15 µM, as calculated according to the method described previously (37). This concentration was not sufficient to elicit substantial inhibition in vitro (Fig. 8) but inhibited rRNA synthesis in vivo (Fig. 4 and 5). Gourse and colleagues (45) recently suggested this discrepancy reflects the level of the DksA protein, which amplifies the effects of ppGpp in E. coli cells by binding to RNA polymerase. They found that the concentration of ppGpp required for half-maximal inhibition of rrnB P1 promoter activity was 20 µM in the absence of DksA but only 1 to 2 µM in its presence. In addition, the structural basis for ppGpp-DksA synergism during transcription has been determined from analysis of the three-dimensional structure of E. coli DksA (46). A similar mechanism for amplifying the effect of ppGpp may operate in T. thermophilus cells, though a DksA homologue has not yet been found in that strain. Alternatively, it is possible that the intracellular GTP concentration, rather than ppGpp per se, is a more important component in regulation of rRNA gene expression in T. thermophilus, as recently proposed for B. subtilis by Krasny and Gourse (29). These authors found that in contrast to the case with E. coli, changes in B. subtilis rRNA promoter activity always correlate with changes in the intracellular GTP concentration, thus reaching a conclusion that in contrast to the situation for E. coli, where ppGpp decreases rRNA promoter activity by directly inhibiting RNA polymerase, ppGpp may not inhibit B. subtilis RNA polymerase directly. Rather, an increase in the ppGpp concentration might reduce the available GTP pools (perhaps by inhibiting IMP dehydrogenase activity), thereby modulating rRNA promoter activity indirectly (29). In fact, the present work dealing with T. thermophilus demonstrated that although the maximum concentration of ppGpp observed was only 10 to 15 µM, this value is comparable to the Ki value (20 µM) for ppGpp (see Results), thus showing efficacy of the observed ppGpp level in inhibiting IMP dehydrogenase activity. Despite the lack of examination in vivo for the possible effect of GTP pool size on T. thermophilus rRNA promoter activity due to insusceptibility to drugs known to block GMP synthesis, it is most conceivable that in T. thermophilus, rRNA promoter activity is regulated mainly by changes in the GTP pool size, which is modulated by ppGpp, possibly in cooperation with direct inhibition of RNA polymerase activity by binding of ppGpp. This notion is supported by the fact that activity of the T. thermophilus 23S/5S rRNA promoter was significantly influenced by a wide range of GTP concentration in vitro (Fig. 9). A temporarily increased rRNA transcription during serine hydroxamate treatment for the relA disruptant (Fig. 5) can also be accounted for by the increased GTP pool size (Fig. 3A), which would affect positively the rRNA gene transcription (Fig. 9). In the framework of the above notion, our recent finding that production of an antibiotic bacilysin in B. subtilis is regulated cooperatively by ppGpp and GTP (22) is notable.
One of the characteristic properties of T. thermophilus is its capacity for genetic transformation (28). We recently reported that in B. subtilis, GTP acts as a nutritional signal for competence development (and sporulation) (21) via CodY, which can be thought of as a GTP-sensing transcriptional regulator having a predicted GTP binding pocket (54). Consequently, a sufficient reduction in GTP is required to fully induce genetic competence (and sporulation). Because ppGpp has been shown to potently inhibit IMP dehydrogenase, resulting in a rapid reduction in the level of GTP (37; this study), the stringent response likely contributes to the transformability of T. thermophilus by moderating the level of GTP.
Structural analysis of the RNA polymerase-ppGpp complex suggested that given the flexibility of the architecture of the ppGpp base subsite, ppApp, which is certainly produced in certain Streptomyces species (44), might be accommodated within the same binding site (2). As demonstrated by Travers (58) using E. coli, ppApp acts as a positive effector for transcription of some stringently controlled genes, thus displaying a functionality that opposes the effect of ppGpp. However, unlike ppGpp, ppApp has not yet been detected in E. coli. Although in preliminary experiments we have identified a low level (4 pmol/mg dry weight) of ppApp in T. thermophilus (unpublished data), our present findings clearly demonstrate no substantial effect of ppApp on rRNA gene transcription in T. thermophilus even at a high (1 mM) concentration, irrespective of the absence (Fig. 8A) or presence (not shown) of ppGpp.
The mutations harbored by the KO-564 and KO-565 relC mutants were both situated at Pro21 of the L11 protein, mutation of which was previously shown to confer the RelC phenotype (Fig. 2). It is important to note that all but one relC mutation found so far in Streptomyces spp. were deletion mutations (the exception is the KO-450 S. coelicolor relC mutation; see Fig. 2) and located in this specific region of L11 (27), which is indicative of this region's importance to the cell's ability to synthesize ppGpp. On the basis of the crystal structure of the Thermotoga maritima L11-rRNA complex, Wimberly et al. (62) proposed that the C-terminal domain of L11 binds RNA tightly, while the N-terminal domain makes only limited contact with RNA and functions as a switch that reversibly associates with an adjacent region of RNA. They also found that mutations conferring resistance to thiostrepton are located in a narrow cleft between the RNA and the N-terminal domain, which is suggestive of the mechanism of ribosome-dependent ppGpp synthesis.
In summary, our present findings complement these earlier structural analyses by providing physiological evidence that the stringent response is mediated in T. thermophilus by ribosome-dependent synthesis of (p)ppGpp, which in turn inhibits, directly and/or indirectly, RNA polymerase activity. However, the evidence linking ppGpp to the stringent RNA control response was not sufficient in the present study, due to the fact that T. thermophilus is characterized by a low-level ppGpp accumulation under amino acid starvation conditions. This means that if there is a classical stringent response and these tiny amounts of ppGpp function in T. thermophilus, then the response elements (such as RNA polymerase and IMP dehydrogenase) must be correspondingly more sensitive. Based on the current findings, it is at present most conceivable that in T. thermophilus, rRNA promoter activity is controlled directly by the GTP pool size, which is modulated by ppGpp via inhibition of IMP dehydrogenase activity, a model proposed recently for B. subtilis rRNA transcription by Krasny and Gourse (29). Thus, a more convincing demonstration of ppGpp sensitivity of rRNA gene transcription will require a very careful study in itself, because DksA to amplify the regulatory signal is not present in T. thermophilus, at least from the approach of an amino acid sequence homology search.
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ACKNOWLEDGMENTS |
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We are grateful to Shinichi Etoh and Kazuhiko Kurosawa for preliminary work performed in several of the experiments, Seiki Kuramitsu, Takato Yano, and Yoshinori Koyama for advice on gene engineering, and Yuzuru Tozawa for generous support in proceeding with this study.
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FOOTNOTES |
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Present address: Plant function and their control, CREST, Japan Science Technology Agency, Nihonbashi, Tokyo, Japan.
Present address: Graduate School of Agriculture, Hokkaido University, Sapporo, Japan.
¶ Present address: Best Institute, University of Toronto, Toronto, Canada.
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REFERENCES |
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29-PALG-32 mutation leads to reduced expression of the regulatory genes ccaR and claR and very low transcription of the ceaS2 gene for clavulanic acid biosynthesis in Streptomyces clavuligerus. Mol. Microbiol. 61:758-770.[CrossRef][Medline]
