Synechocystis sp. PCC 6803 glutamine synthetase type I (GS) activity is controlled by direct interaction with two inactivating factors (IF7 and IF17). IF7 and IF17 are homologous polypeptides encoded by the gifA and gifB genes respectively. We investigated the transcriptional regulation of these genes. Expression of both genes is maximum in the presence of ammonium, when GS is inactivated. Nitrogen starvation attenuates the ammonium-mediated induction of gifA and gifB as well as the ammonium-mediated inactivation of GS. Putative binding sites for the transcription factor NtcA were identified at −7.5 and −30.5 bp upstream of gifB and gifA transcription start points respectively. Synechocystis NtcA protein binding to both promoters was demonstrated by gel electrophoresis mobility shift assays. Constitutive high expression levels of both genes were found in a Synechocystis NtcA non-segregated mutant (SNC1), which showed a fourfold reduction in the ntcA expression. These experiments indicate a repressive role for NtcA on the transcription of gifA and gifB genes. Our results demonstrate that NtcA plays a central role in GS regulation in cyanobacteria, stimulating transcription of the glnA gene (GS structural gene) and suppressing transcription of the GS inactivating factor genes gifA and gifB.
Assimilation of ammonium by most microorganisms occurs through the sequential action of glutamine synthetase (GS) and of glutamate synthase (GOGAT), commonly known as the GS-GOGAT cycle (for reviews, see Merrick and Edwards, 1995; Reitzer, 1996). The reaction catalysed by GS involves the ATP-dependent amination of glutamate to yield glutamine. GOGAT then catalyses the transfer of the amide group from glutamine to 2-ketoglutarate to yield two molecules of glutamate. This pathway represents the connecting step between carbon and nitrogen metabolism. Because of this, it is not surprising that both the activity and the synthesis of the first enzyme of the pathway, GS, are finely regulated in many organisms. In most of the studied systems, the control of GS activity responds to carbon and nitrogen signals. In the presence of abundant carbon sources, nitrogen deficiency results in a high level of GS activity. In contrast, when a nitrogen-rich source is present, GS activity is downregulated.
In enteric bacteria, GS activity is regulated by adenylylation (for reviews, see Rhee et al., 1989; Stadtman, 1990; Merrick and Edwards, 1995; Reitzer, 1996). Although the adenylylated form is highly sensitive to feedback inhibition, the deadenylylated form is relatively insensitive to feedback inhibition. Adenylylation involves the transfer of an adenylyl group from ATP to a Tyr residue, each of the 12 subunits of the enzyme being susceptible to adenylylation. The adenylylation state of GS is controlled by a bicyclic cascade involving two bifunctional proteins, the adenylyltransferase (ATase) and the uridylyltransferase (UTase), and the signal transducing protein PII. In addition to controlling GS activity by covalent modification, UTase and PII, together with the two-component regulatory system comprising NtrC and NtrB, modulate the level of transcription of the GS structural gene (glnA) (Merrick and Edwards, 1995; Magasanik, 1996).
Cyanobacterial GS type I (hereafter referred to as GS) activity is not controlled by adenylylation (Fisher et al., 1981). However, an ammonium-dependent GS inactivation was characterized in the unicellular cyanobacterium Synechocystis sp. PCC 6803 (Mérida et al., 1991a, b; Reyes and Florencio, 1995). We have recently reported that the mechanism of Synechocystis GS inactivation involves the binding of two polypeptides (IF7 and IF17) to the GS, yielding an inactive GS–IFs complex (García-Domínguez et al., 1999). IF7 and IF17 are homologous proteins encoded by two unlinked genes, gifA and gifB. ΔgifA and ΔgifB Synechocystis mutant strains are severely impaired in GS inactivation, and the double mutant ΔgifA/ΔgifB is completely deficient in GS inactivation.
In the present work, we studied the transcriptional control of gifA and gifB genes. We demonstrate that transcription of both genes is repressed in the absence of ammonium and that the transcription factor NtcA is responsible for this repression. This is the first time that a role for NtcA in transcriptional repression is demonstrated.
gifA and gifB expression is controlled by the nitrogen availability
As a first step in the characterization of gifA and gifB gene expression, we analysed the steady-state levels of gifA and gifB mRNA under different nitrogen conditions by means of Northern blotting experiments. Levels of gifA and gifB mRNA were maximum in ammonium-grown cells, decreasing to about 35% and 20%, respectively, in nitrate-grown cells. Transcript levels were barely detectable under nitrogen deficiency (Fig. 1). It is worth noting that accumulation of Synechocystis 6803 glnA transcript follows an inverse correlation, high levels under nitrogen deficiency and low levels in the presence of ammonium (Reyes et al., 1997). Addition of ammonium to nitrate-grown cells leads to a fast and transitory increase of the gifA and gifB mRNA levels co-incident with the drop in GS activity (Fig. 2A; García-Domínguez et al., 1999). We have previously reported that nitrogen-starved Synechocystis 6803 cells showed a delay in the ammonium-dependent GS inactivation (Fig. 2C, right graph; Mérida et al., 1991a). Therefore, if expression of gifA and gifB is directly involved in GS inactivation, ammonium-dependent induction of gifA and gifB mRNAs should also be delayed under these conditions. Thus, when ammonium was added to nitrogen-starved cells, gifA and gifB mRNA levels increased slowly, reaching the maximum after 6 and 3 h respectively (Fig. 2B). On the contrary, when ammonium-grown cells were transferred to a medium free of combined nitrogen, expression of both genes was completely repressed in a few minutes (data not shown). Addition of nitrate to nitrogen-starved cells had a much smaller effect on gifA and gifB mRNA levels and on GS activity (Fig. 2B and C). These data confirm that there is a precise inverse correlation between the levels of gifA and gifB mRNA and the levels of GS activity.
gifA and gifB promoters contain NtcA binding sites
To identify the promoter regions of gifA and gifB genes (PgifA and PgifB), the transcription start points (TSP) of both genes were determined by primer extension analysis. The gifA TSP was mapped to nucleotide −51 with respect to the translation start codon (Fig. 3A). Six nucleotides upstream of the TSP, a putative −10 box in the form TATAAA was found. No obvious −35 box was detected. The gifB TSP was mapped to 104 nucleotides upstream of the gifB start codon (Fig. 3A). Putative −10 and −35 promoter boxes were found at the appropriate positions (Fig. 3B). As observed in RNA blotting experiments, primer extension products from both gifA and gifB promoters were more abundant in samples from cells grown in the presence of ammonium than from those grown in the presence of nitrate, and were undetectable using RNA from nitrogen-starved cells. As mentioned previously, the transcriptional regulator NtcA activates transcription of a number of promoters under conditions of ammonium deprivation (Luque et al., 1994). As gifA and gifB are repressed under these conditions, a putative role for NtcA as a repressor of both genes could be hypothesized. To verify this possibility, the presence of NtcA binding sites in the gifA and gifB promoter regions was checked. A consensus NtcA binding site (Luque et al., 1994) centred at position −30.5 upstream of the gifA TSP was found (Fig. 3B). A consensus NtcA binding site was also found overlapping the putative −10 promoter box of the gifB gene (centred at position −7.5; Fig. 3B). The position of the gifA and gifB NtcA binding sites contrasts with the position of the NtcA binding sites of those promoters in which NtcA activates transcription (Fig. 3B).
NtcA binds to gifA and gifB promoters
To confirm that NtcA binds to PgifA and PgifB, electrophoretic mobility shift assays using purified Synechocystis 6803 NtcA protein were performed. Synechocystis NtcA was expressed in Escherichia coli and purified as a GST-fusion protein, as described previously (Muro-Pastor et al., 1996). A 195 bp BstXI–AccI fragment and a 100 bp HincII–BamHI fragment containing PgifA and PgifB, respectively, were retarded by purified NtcA protein (Fig. 4, lanes 5, 6, 10 and 11), but not by purified GST protein (lanes 4 and 9). The NtcA-dependent shift was severely diminished in the presence of a 10-fold excess of the same unlabelled fragment (lanes 7 and 12), and it was unaffected by the presence of an excess of an unrelated DNA fragment (data not shown). A 150 bp PvuII–BamHI fragment from the 3′ region of the gifA gene was used as a non-related probe. This fragment was not retarded by NtcA protein (Fig. 4, lanes 1 and 2). These experiments demonstrate that NtcA binds specifically to the gifA and gifB promoter regions in vitro.
NtcA represses transcription of gifA and gifB genes
To demonstrate that the transcriptional regulator NtcA controls the synthesis of gifA and gifB mRNA, we decided to determine the levels of gifA and gifB transcripts in a NtcA mutant strain. To generate a Synechocystis 6803 NtcA mutant, we constructed a plasmid containing the ntcA gene disrupted by a chloramphenicol (Cm) resistance cassette (Fig. 5A). This plasmid was used to transform Synechocystis 6803. CmR colonies were obtained and subjected to several rounds of segregation. This strain was named SNC1. Synechocystis 6803 is a polyploid bacterium with about 12 chromosomes per cell (Labarre et al., 1989). Southern blot analysis revealed that only partial segregation of the inactivated version of the ntcA gene was achieved. Therefore, SNC1 strain contains both wild type and ntcA mutated chromosomes. The maximum segregation level was obtained after several rounds of culture using ammonium as a nitrogen source and with Cm (80 µg ml−1). Under these conditions, the level of segregation was stable and the ratio of mutated to wild-type chromosomes was about 4 (Fig. 5B). This segregation level was not affected after 6 h under nitrate utilizing conditions (data not shown). To determine the level of expression of the ntcA gene in both the wild type and the SNC1 strain under these conditions, Northern blot experiments were carried out. As shown in Fig. 5C, the amount of ntcA transcript in SNC1 cells was about fourfold less than in wild-type cells.
We previously postulated that Synechocystis 6803 glnA (structural gene for glutamine synthetase type I), glnN (structural gene for glutamine synthetase type III) and glnB (encoding the signal transducing protein PII) gene transcription is controlled positively by NtcA, based on the presence of NtcA binding sites in their respective promoters (García-Domínguez and Florencio, 1997; Reyes et al., 1997). We analysed the steady-state mRNA levels of these genes in the wild type and in the SNC1 strain under three different conditions: nitrate utilization, ammonium utilization and nitrogen starvation. Ammonium-grown SNC1 or wild-type cells were transferred for 6 h to nitrate- or ammonium-containing medium or to nitrogen-free medium and samples were taken for RNA isolation. As previously reported, levels of glnA, glnN and glnB mRNA were upregulated in nitrogen-deficient wild-type cells. However, induction was severely impaired or absent in SNC1 mutant cells (Fig. 6). This experiment demonstrates that NtcA is a transcriptional activator of glnA, glnN and glnB genes in Synechocystis 6803. gifA and gifB transcript levels were high in wild-type ammonium-grown cells and were downregulated in both nitrate-grown cells and nitrogen-starved cells. In contrast, gifA and gifB transcript levels in SNC1 cells remained high under all conditions tested. Furthermore, ammonium-grown SNC1 cells showed approximately twofold higher gifA transcript levels than ammonium-grown wild-type cells. This experiment indicates that NtcA represses both gifA and gifB promoters.
Synechocystis 6803 GS is controlled by a mechanism that differs from the classic adenylylation system present in many prokaryotes. In the presence of ammonium, two proteins, IF7 and IF17, bind to the GS, yielding an inactive complex (García-Domínguez et al., 1999). Our previous results show that recombinant IF7 and IF17 produced in E. coli are able to inactivate GS in vitro, suggesting that both factors can interact with the GS without further modification (García-Domínguez et al., 1999). Therefore, our data indicate that GS inactivation is determined by the intracellular levels of IF7 and IF17. We show in this work that transcription of the gifA and gifB genes, encoding IF7 and IF17 respectively, is upregulated in the presence of ammonium and is repressed in the absence of this nitrogen source, and that the transcription factor NtcA is responsible for this regulation.
In most of the NtcA-dependent promoters characterized to date, the position of the NtcA binding site is centred at about nucleotide −41.5 (see Fig. 3B), similarly to the class II CRP-dependent promoters (Kolb et al., 1993; Busby and Ebright, 1997). In all these promoters, NtcA behaves as an activator of transcription. In class I CRP-dependent promoters, the DNA-binding site for CRP is located upstream of the site for RNA polymerase, at about −61.5 (Ebright, 1993). No NtcA-dependent promoters have been identified with these characteristics. Certain activators of the CRP family of transcription factors can also mediate repression. This has been clearly characterized for several promoters controlled by CRP or FNR (Collado-Vides et al., 1991; Kolb et al., 1993). In these cases, transcription factor binding sites overlap the RNA polymerase-binding sites between −40 and +20. A role for NtcA as a repressor has also been hypothesized based on the presence of NtcA binding sites in typically repressive positions in the gor and the rbcLS promoters (Ramasubramanian et al., 1994; Jiang et al., 1995). However, this repressive role had not been demonstrated. In the gifA and gifB genes, NtcA binding sites are centred at −30.5 and −7.5 bp, respectively, suggesting a repressive role of NtcA at these promoters. The constitutive expression of both genes in the NtcA mutant (SNC1) demonstrates this repressive activity in vivo. The presence of a consensus −35 box in the gifB but not in the gifA promoter, together with the different location of the NtcA binding site, may account for the fact that gifB is more sensitive to the presence of ammonium than gifA (see Fig. 2).
We and others have previously shown that the transcription factor NtcA activates transcription from several cyanobacterial glnA promoters in the absence of ammonium (Frias et al., 1994; Luque et al., 1994; Cohen-Kupiec et al., 1995; Reyes et al., 1997). Data presented in this work demonstrate that NtcA represses transcription of gifA and gifB genes in the absence of ammonium. Therefore, in Synechocystis 6803, NtcA plays a central role in the regulation of glutamine synthetase. Our current model is as follows (Fig. 7). Under conditions of nitrogen excess, NtcA is in an inactive form and is unable to activate or repress transcription of glnA or gif genes respectively. This provokes a derepression of gifA and gifB transcription, which determines the inactivation of GS. This situation is also characterized by a basal level of transcription of the glnA gene. Under nitrogen deficiency, NtcA in its active form activates transcription of the glnA promoter (about fourfold induction) and represses gifA and gifB genes. Our results demonstrate a central role for transcriptional regulation in the mechanism by which GS activity is controlled in cyanobacteria. This is in contrast to the classic adenylylation system of GS regulation, in which most of the concurring elements (uridylyltransferase, adenylyltransferase and PII protein) are constitutively expressed irrespective of the nitrogen status of the cell (van Heeswijk et al., 1993). Two similarities between the enterobacterial and the cyanobacterial systems can be highlighted. Both regulatory systems sense the nitrogen–carbon balance of the cell. In the case of the enterobacterial system, the ratio of glutamine to 2-ketoglutarate determines the modification state of the PII protein (Stadtman, 1990; Reitzer, 1996; Jiang et al., 1997b). PII protein is modified by uridylylation through the action of the uridylyltransferase enzyme, a bifunctional enzyme able to carry out both the uridylyl transfer and the removing activity. A high intracellular concentration of glutamine induces the uridylyl-removing activity, provoking the deuridylylation of PII. The interaction of the unmodified PII with the adenylyltransferase results in adenylylation of GS. A high intracellular concentration of 2-ketoglutarate activates the uridylyltransferase activity, which transfers an UMP group to each one of the three subunits of the PII protein to form PII-UMP. PII-UMP interacts with ATase, which in turn catalyses the removal of AMP, yielding active GS. In the case of Synechocystis 6803, nitrogen starvation attenuates the ammonium-mediated induction of gifA and gifB and, therefore, the ammonium-mediated inactivation of GS. As nitrogen-starved Synechocystis 6803 cells contain sevenfold higher levels of 2-ketoglutarate than nitrate-grown cells (Mérida et al., 1991a), these data suggest that GS regulation is modulated through the internal balance between carbon–nitrogen compounds and carbon compounds. How this balance modulates the activity of NtcA remains to be elucidated. A second point of similarity between the Synechocystis 6803 and the E. coli system is the fact that both control of GS activity and control of GS synthesis (transcriptional regulation of glnA gene) are finely co-ordinated through a common element. In the case of E. coli, the common element is the PII protein. PII together with the two-component regulatory system comprising NRI (NtrC) and NRII (NtrC) modulate the level of transcription of the glnA gene (Merrick and Edwards, 1995; Magasanik, 1996; Reitzer, 1996). In the case of Synechocystis 6803, the common element is NtcA, which controls the synthesis of the GS and the synthesis of its inhibitors. A PII protein has been identified and characterized in cyanobacteria (Harrison et al., 1990; Tsinoremas et al., 1991; Forchhammer and Tandeau de Marsac, 1994; García-Domínguez and Florencio, 1997). Cyanobacterial PII protein is post-transcriptionally modified by phosphorylation at Ser-49 (Forchhammer and Tandeau de Marsac, 1994). The fact that GS inactivation is operative in a Ser-49→Ala Synechocystis PII point mutant indicates that PII protein is not involved in the GS inactivation signal transduction pathway (M. García-Domínguez and F. J. Florencio, unpublished). Furthermore, repression by ammonium and induction in the absence of ammonium of NtcA-dependent genes is not affected in PII null mutants in Synechococcus PCC 7942, indicating that PII is not involved in the control of NtcA (Forchhammer and Tandeau de Marsac, 1995; Lee et al., 1998). These data strongly suggest that PII protein is not directly involved in GS control in cyanobacteria.
Two questions remain in our model for Synechocystis 6803 GS control (Fig. 7). One of them, the mechanism of regulation of NtcA activity, has been already discussed, the second one concerns the GS reactivating mechanism. Ammonium-dependent Synechocystis 6803 GS inactivation is a reversible process (Mérida et al., 1991a, b; Reyes and Florencio, 1995). Thus, GS is fully reactivated within 10 min after removing ammonium from Synechocystis 6803 cultures. Ammonium removal provokes the repression of gifA and gifB expression (data not shown). One possibility is that the simple dilution of IF7 and IF17 may determine the release of both factors from GS and the reactivation of the enzyme. However, the fast reactivation process suggests the involvement of specific mechanisms.
Although it has been actively pursued, complete segregation of a ntcA Synechocystis 6803 mutant has been unsuccessful. A possible explanation for the requirement of ntcA for viability in Synechocystis 6803 can be deduced from the regulatory circuits that we present in this work. Previous data from our group demonstrated that Synechocystis 6803 is unable to grow in the absence of glutamine synthetase activity. Thus, glnA or glnN single mutants were viable, but ΔglnAglnN double mutants were non-viable (Reyes and Florencio, 1994). The partially segregated ntcA mutant (SNC1) showed basal levels of glnA expression and undetectable levels of glnN expression under all tested conditions. In addition, SNC1 cells showed high levels of gifA and gifB expression, suggesting the existence of high intracellular levels of IF7 and IF17. Therefore, low amounts of GS protein and high levels of GS inhibitors would determine a very low GS activity. The inability to reach complete segregation may be related to the absence of GS activity expected in fully segregated Synechocystis 6803 ntcA mutants. In Synechococcus 7942 and Anabaena 7120 strains, knock out ntcA mutants are viable, showing a reduction in the level of GS activity (about 50% of the total GS activity present in the wild-type strain; Vega-Palas et al., 1990). Long-term ammonium-dependent decrease of GS activity (50% decrease) has been observed in these two species (Rowell et al., 1979; data not shown), contrasting with the rapid inactivation of GS in Synechocystis 6803 and suggesting that regulation of GS activity in these strains is not as dramatic as in Synechocystis 6803.
In summary, the present study reveals the dual role of NtcA as an activator and a repressor, which allows a precise co-ordination of GS activity and synthesis. Further experiments are required to determine the early steps in the cyanobacterial GS regulatory pathway, i.e. how changes in nitrogen availability modulate the transcriptional activity of NtcA.
Strains and growth conditions
Synechocystis sp. strain PCC 6803 was grown photoautotrophically at 30°C in BG11 medium (Rippka et al., 1979) supplemented with 1 g l−1 NaHCO3 (BG11c) and bubbled with a continuous stream of 1% (v/v) CO2 in air under continuous fluorescent illumination (50 µmol m−2 s−1 photons from white light). For plate cultures, BG11c liquid medium was supplemented with 1% (w/v) agar. Chloramphenicol was added to a final concentration of 80 µg ml−1 when required. When ammonium was used as nitrogen source, nitrate was replaced by 10 mM NH4Cl and the medium was buffered with 20 mM N-tris(hydroxymethyl)-methyl-2-aminoethane-sulphonic acid (TES) buffer.
Escherichia coli DH5α (Bethesda Research Laboratories) grown in Luria–Bertani broth medium, as described in Sambrook et al. (1989), was used for plasmid construction and replication. E. coli was supplemented with 100 µg ml−1 ampicillin when required.
Insertional mutagenesis of the Synechocystis ntcA gene
The ntcA gene was amplified by standard PCR procedures, using oligonucleotides M1 and M2 (Muro-Pastor et al., 1996). The Synechocystis 6803 SNC1 mutant strain was created by interrupting the ntcA gene with the C.C1 cassette (Elhai and Wolk, 1988) containing a cat gene (chloramphenicol acetyltransferase) that confers resistance to chloramphenicol. C.C1 was isolated as a 1.9 kb HincII DNA fragment and introduced into the unique DraIII site of ntcA. Transformation of Synechocystis 6803 cells was carried out as previously described (Chauvat et al., 1988).
For Southern blotting analysis, total DNA from cyanobacteria was isolated as previously described (Cai and Wolk, 1990). DNA was digested and electrophoresed in 0.7% agarose gels in a tris-borate/EDTA buffer system (Sambrook et al., 1989), then DNA was transferred to nylon Z-probe membranes (Bio-Rad). DNA probes were 32P-labelled with a random-primer kit (Pharmacia) using [α-32P]-dCTP (3000 Ci mmol−1).
RNA isolation and Northern blot hybridization
Total RNA was isolated from 25 ml samples of Synechocystis 6803 cultures at the mid-exponential phase (3–5 µg ml−1 chlorophyll). Extractions were performed by vortexing cells in the presence of phenol–chloroform and acid-washed baked glass beads (0.25–0.3 mm diameter; Braun) as previously described (García-Domínguez and Florencio, 1997).
For Northern blotting analysis, 15 µg of total RNA was loaded per lane and electrophoresed in 1.2% agarose denaturing formaldehyde gels. Transfer to nylon membranes (Hybond N-plus; Amersham), prehybridization, hybridization and washes were performed as described in the Amersham instruction manual. Hybridization was conducted at 42°C in the presence of 50% formamide. The 409 bp DNA fragment obtained by PCR amplification with oligonucleotides sif3 (TACATATGTCTACTCAACAACAGG) and sif4 (GTTAATGGGATCCTAGTTAATATC) and the 637 bp DNA fragment obtained with oligonucleotides lif3 (GCGCCATATGCAATTAAGTTACCG) and lif4 (TTGGATCCTCCGTTATCTGAATAG) were used as the gifA and gifB probes respectively. M1 and M2 oligonucleotides were used to generate the ntcA probe (Muro-Pastor et al., 1996). As a control, in all the cases the filters were reprobed with a Hin dIII–Bam HI 580 bp probe from plasmid pAV1100 that contains the constitutively expressed RNase P RNA gene from Synechocystis 6803 (Vioque, 1992). To determine the counts per minute (c.p.m.) of radioactive areas in Northern blot hybridizations, an InstantImager Electronic Autoradiography apparatus (Packard Instrument Company) was used.
Primer extension analysis
Oligonucleotides sif5 (GCGCGAGCCTGTTGTTGAGTAGAC) and lif5 (AACCACGGTAGCTGCCCGCTAGCC) end-labelled with T4 polynucleotide kinase and [γ-32P]-dATP (3000 Ci mmol−1) following standard procedures (Sambrook et al., 1989) were used for primer extension analysis of gifA and gifB promoters respectively. For annealing, a 10 µl mixture containing 0.15 M KCl, 10 mM tris-HCl, pH 8.0, 1 mM EDTA, 20 µg of total RNA and about 2 pmol of oligonucleotide (106 c.p.m.) was prepared. The annealing mixture was heated for 2 min at 90°C in a water bath and cooled slowly to 50°C. For extension, a 10 µl mixture was prepared with one-half of the annealing mixture, 10 mM DTT, 0.5 mM each dNTP, 2 µg actinomycin D, 50 mM tris-HCl, pH 8.3, 75 mM KCl, 3 mM MgCl2 and 100 U of Superscript II RNase H-Reverse Transcriptase (Gibco BRL). The mixture was incubated for 45 min at 45°C and the reaction was stopped by adding 4 µl of formamide-loading buffer. One-half of the reaction was electrophoresed on a 6% polyacrylamide sequencing gel together with a sequencing reaction of the gifA or gifB gene 5′ regions, using the sif5 or lif5 oligonucleotide respectively.
Gel retardation assays
GST–NtcA fusion protein was expressed and purified as previously described (Muro-Pastor et al., 1996). The indicated fragments were end-labelled with [α-32P]-dCTP using Sequenase version 2.0 enzyme. The binding reactions and electrophoresis were carried out as previously described (Muro-Pastor et al., 1996). The gifA promoter probe was obtained by BstXI–AccI digestion of a PCR-amplified fragment using oligonucleotides sif1 (GCAAACATCCGCCCATGGATCAAC) and sif5. The gifB promoter probe was obtained by Hin cII–Bam HI digestion of a PCR-amplified fragment using oligonucleotides lif1 (TCCAGCACTGGCGCCAGCATTTCC) and lif7 (CAGGAATGGATCCCTACTTCCGTC).
GS activity was determined in situ by using the Mn2+-dependent γ-glutamyl-transferase assay in cells permeabilized with mixed alkyltrimethylammonium bromide (Mérida et al., 1991a). One unit of GS activity corresponds to the amount of enzyme that catalyses the synthesis of 1 µmol min−1 of γ-glutamylhydroxamate.
We thank J. J. Ghislain and A. Vioque for critical reading of the manuscript. M.G.-D. was the recipient of a fellowship from Spanish Ministerio de Educación y Cultura. This study was supported by grant PB97-0732 from DGESIC and by Junta de Andalucía (group CV1-0112).