Intracellular autoregulation of the Mycobacterium tuberculosis PrrA response regulator

Fanny Ewann, Camille Locht and Philip Supply

INSERM U447, Institut Pasteur de Lille, 1 rue du Professeur Calmette, F-59019 Lille Cedex, France

Correspondence
Philip Supply
philip.supply{at}pasteur-lille.fr


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Two-component systems are major regulatory systems for bacterial adaptation to environmental changes. During the infectious cycle of Mycobacterium tuberculosis, adaptation to an intracellular environment is critical for multiplication and survival of the micro-organism within the host. The M. tuberculosis prrA gene, encoding the regulator of the two-component system PrrA–PrrB, has been shown to be induced upon macrophage phagocytosis and to be transiently required for the early stages of macrophage infection. In order to study the mechanisms of regulation of the PrrA–PrrB two-component system, PrrA and the cytoplasmic part of the PrrB histidine kinase were produced and purified as hexahistidine-tagged recombinant proteins. Electrophoretic mobility shift assays indicated that PrrA specifically binds to the promoter of its own operon, with increased affinity upon phosphorylation. Moreover, induction of fluorescence was observed after phagocytosis of a wild-type M. tuberculosis strain containing the gfp reporter gene under the control of the prrAprrB promoter, while this induction was not seen in a prrA/B mutant strain containing the same construct. These results indicate that the early intracellular induction of prrA depends on the autoregulation of this two-component system.


Abbreviations: His6, hexahistidine


   INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Two-component systems are major elements in bacterial adaptation to environmental changes. These systems are implicated in a large variety of adaptive responses, such as quorum sensing, chemotaxis and metabolic changes [for reviews, see Hoch (2000) and Stock et al. (2000)]. In many pathogenic bacteria, two-component systems are central regulatory elements for the production of virulence factors. Two-component systems basically involve a histidine kinase and a response regulator, which communicate through a phosphorylation cascade. In addition to a so-called receiver domain, which includes a conserved aspartyl residue as the final target of the phosphorelay, most response regulators contain an effector domain able to bind specific DNA sequences, and to thereby act as transcriptional regulators.

Mycobacterium tuberculosis possesses a complex infectious cycle, which includes intra- and extra-cellular phases, both within and outside the lungs, as well as a latency phase, suggesting that the expression of many of its genes must be subjected to regulation. However, the regulatory mechanisms governing the adaptive responses of M. tuberculosis, especially during phagocytosis, are still poorly understood. The M. tuberculosis genome contains 11 pairs of genes encoding two-component systems, in addition to a few isolated genes encoding orphan histidine kinases or response regulators (Cole et al., 1998). Several of these genes have been characterized at least partially (Dasgupta et al., 2000; Ewann et al., 2002; Graham & Clark-Curtiss, 1999; Haydel et al., 1999; Himpens et al., 2000; Perez et al., 2001; Sherman et al., 2001; Supply et al., 1997; Via et al., 1996; Zahrt & Deretic, 2000, 2001). For example, an M. tuberculosis strain with a mutation in the phoP gene was found to be impaired in intra-cellular growth within macrophages, one of the major target cells of M. tuberculosis, and its virulence was found to be attenuated in mice (Perez et al., 2001). MprA–MprB is required for persistence in murine infection (Zahrt & Deretic, 2001) and DevR–DevS is induced in response to hypoxia and required to survive it (Boon & Dick, 2002; Park et al., 2003; Sherman et al., 2001). MtrA–MtrB was found to be essential for survival, as so far it has not been possible to obtain mtrA knockout strains of M. tuberculosis (Zahrt & Deretic, 2000). In addition, mtrA has been shown to be upregulated upon phagocytosis in Mycobacterium bovis BCG but not in M. tuberculosis (Via et al., 1996). The upregulating mechanism in M. bovis BCG has not yet been identified. The PrrA–PrrB system has been found to be induced after macrophage phagocytosis and to be transiently required during the early stages of the macrophage infection (Ewann et al., 2002; Graham & Clark-Curtiss, 1999). The PrrA–PrrB system belongs to a wide subfamily of two-component systems, of which OmpR–EnvZ is the prototype. Many members of this subfamily have been demonstrated to be autoregulated. However, autoregulation is not a general rule, as illustrated by the hilA gene in Salmonella typhimurium (Bajaj et al., 1996; Lucas et al., 2000). Here, we investigated the possible autoregulation of prrA and its role in the induction of the expression of this gene upon phagocytosis of M. tuberculosis.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bacterial strains and plasmids.
The bacterial strains and plasmids used in this study are listed in Table 1. All cloning steps were carried out in Escherichia coli XL-1 Blue (Stratagene). Recombinant hexahistidine (His6)–PrrA and His6–PrrB were produced in E. coli M15 (Qiagen) and SG13009 (Qiagen), respectively. All DNA fragments were amplified by PCR from chromosomal DNA of the M. tuberculosis clinical isolate Mt103 (Jackson et al., 1999). The M. tuberculosis Mt21D3 mutant derivative, which contains a transposon inserted five nucleotides upstream of the predicted prrA start codon, has been described previously. This insertion presumably prevents transcription of the prrAprrB operon, as a transcriptional terminator is present at the 3' end of the aph gene in the transposon and results in an impairment of the intracellular multiplication capacity during the first days of murine macrophage infection (Ewann et al., 2002). This strain will be subsequently referred to as the prrA/B mutant.


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Table 1. Strains and plasmids used in this study

 
pFluo1Sm was constructed by inserting a streptomycin-resistance cassette into pFluo1 (Ewann et al., 2002) digested with EcoRV. This cassette was obtained by digesting pHP45{Omega} (Prentki & Krisch, 1984) with HindIII and Klenow fragment (Roche Diagnostic).

PrrA and the cytoplasmic domain of PrrB were produced as His6-tagged recombinant proteins using pQE-30 (Qiagen). The prrA-coding sequence was amplified by PCR using the oligonucleotide pair 5'-AAAAGATCTATGGGCGGCATGGACACTGGTGTGA-3' and 5'-AAAAAGCTTATTCTGCATACGCAGCACGAATCCGACT-3'; these primers include a BglII and a HindIII restriction site, respectively (underlined). The cytoplasmic domain of PrrB was identified by alignment against the EnvZ and SenX3 sequences (Forst et al., 1989; Himpens et al., 2000) as the C-terminal part of the protein starting from amino acid 204. The corresponding DNA sequence was amplified by PCR using oligonucleotides 5'-GCGGATCCATCGAGATCGCCGAGGC-3' and 5'-TTTAAGCTTCTAACTGGGTCCGGGAAGGC-3', containing a BamHI and a HindIII restriction site, respectively (underlined). The amplified fragments were digested with BamHI and HindIII and inserted into pQE30 restricted by the same enzymes to yield pQE30-PrrA and pQE30-PrrB, respectively.

Purification of the His6-tagged recombinant proteins under native conditions.
The recombinant E. coli strains containing pQE-PrrA and pQE-PrrB were grown in 1 l LB medium containing 100 µg ampicillin ml-1 and 25 µg kanamycin ml-1. When the OD600 value reached 1·2–1·4, expression of the genes encoding the recombinant proteins was induced with 1mM IPTG for 3 h. The cells were then harvested by centrifugation, resuspended in 5 ml lysis buffer (300 mM NaCl, 50 mM Na2HPO4, 10 mM imidazole, pH 8) per gram of fresh weight. The cells were then lysed using a French Press under a pressure of 1000 p.s.i. (6·9 MPa). The lysates were clarified by centrifugation at 10 000 g for 20 min. The supernatants were filtered using a 0·45 µm filter before loading onto a 1·5 ml Ni-NTA column (Qiagen) equilibrated in lysis buffer. The column was washed first with lysis buffer until the OD280 value reached less than 0·01 and then with 5 ml lysis buffer containing 50 mM imidazole. The proteins were eluted with 7 ml lysis buffer containing 250 mM imidazole, and fractions of 1·5 ml were collected. The fractions were analysed by SDS-PAGE, using a 12·5 % polyacrylamide gel, and Coomassie blue staining. After dialysis overnight against PBS (137 mM NaCl, 2·7 mM KCl, 10 mM Na2HPO4, 2 mM KHPO4) containing 10 % (v/v) glycerol, the protein concentrations were measured using BCA kit (Pierce); the proteins were dispensed into aliquots and then stored at -20 °C.

Phosphorylation assays.
Phosphorylation assays were performed using 2 µg of His6–PrrB incubated for 20 min at 37 °C in the presence of 10 µCi of [{gamma}-32P]ATP (3000 Ci mmol-1, 111 TBq mmol-1; Amersham Biosciences) in 20 µl of phosphorylation buffer containing 100 mM Tris/HCl (pH 8·0), 50 mM KCl and 5 mM MnCl2. For phosphotransfer assays, 20 µl of phosphorylation buffer containing 10 µg of His6–PrrA were subsequently added. The reactions were stopped by the addition of 5 µl of 0·5 M EDTA (pH 8·0), and the incubation mixtures were subjected to SDS-PAGE using a 12·5 % polyacrylamide gel. After electrophoresis and Coomassie blue staining, the gel was dried and exposed for autoradiography to an X-ray film (Biomax; Kodak).

Electrophoretic mobility shift assays.
The promoter region of the prrAprrB operon, PprrA/B, was amplified by PCR using primers prrA/PF (5'-tcggggattgtcgacaccatc-3') and prrA/PR (5'-ccatttgcctgattaccgtc-3'). The amplified fragment containing the entire intergenic region separating prrA from its flanking gene (Rv0904) was sequenced and then labelled by T4 kinase (Roche diagnostics) using 10 µCi of [{gamma}-32P]ATP (3000 Ci mmol-1; Amersham Biosciences). One nanogram of the labelled PCR fragment was incubated for 20 min at room temperature with 0–2 µg of His6–PrrA, in 10 µl binding buffer containing 2 mM Tris/HCl (pH 8·0), 0·4 mM MgCl2, 10 mM KCl, 200 µM DTT, 10 % (v/v) glycerol and 0·01 % Nonidet P40. The reaction mixtures were loaded onto a 12 % (w/v) polyacrylamide/45 mM Tris-borate/1 mM EDTA (pH 8·0) native gel and subjected to electrophoresis. After drying, the gel was exposed to an X-ray film. When the effect of phosphorylation on binding was tested, His6–PrrA was phosphorylated prior to the assay in the same phosphorylation buffer as above containing 0·5 mM ATP instead of radiolabelled ATP.

Bone-marrow macrophage infection and flow cytometry analysis.
Murine bone-marrow-derived macrophages were prepared and grown as described previously (Ewann et al., 2002). The infection assays were performed with 2x105 cells per well in 24-well cell culture clusters (Techno Plastic products). After removing the culture medium, 1 ml of a suspension of the M. tuberculosis Mt103 wild-type or of the Mt21D3 prrA/B mutant strain (Ewann et al., 2002), each containing pFluo1Sm, or of BCG containing pFluo1Sm was added to obtain an m.o.i. of 10 : 1. Control wells containing non-infected macrophages were filled with 1 ml fresh culture medium. After 4 h incubation at 37 °C, the cells were washed three times in PBS to remove extracellular bacteria. The cells were then scraped and resuspended in 300 µl PBS; the fluorescence was analysed using a FACSVantage apparatus (BD Bioscience). The non-infected macrophage suspension was used as a reference to define the macrophage cell population, to exclude free bacilli and to eliminate the effects of macrophage autofluorescence.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Production of the recombinant proteins under non-denaturing conditions
The PrrA regulator and the cytoplasmic domain of the PrrB sensor were produced as recombinant His6-tagged proteins and purified under non-denaturing conditions to preserve enzymic activity. Since the amino-terminal transmembrane regions are dispensable for in vitro activity of most histidine kinases, this region of PrrB was not included to avoid solubility problems. A major protein with an apparent molecular mass of 30 kDa was detected in the soluble fraction of IPTG-induced E. coli M15(pQE-PrrA) upon SDS-PAGE and Coomassie blue staining (Fig. 1a, lane 2). This protein was then purified by Ni-NTA affinity chromatography (lane 3). In the soluble fraction of IPTG-induced E. coli SG13009(pQE-PrrB), no major band could be detected in the range of the expected size for His6–PrrB (Fig. 1b). Nevertheless, a protein with an apparent molecular mass of 31 kDa could be purified after Ni-NTA chromatography from this fraction. The observed molecular masses of the two recombinant proteins are slightly higher than expected for His6–PrrA and His6–PrrB (27·0 and 27·3 kDa, respectively), as frequently observed for His6-tagged proteins. The identity of the purified His6-tagged proteins was confirmed by immunoblotting using anti-His tag antibodies (not shown).



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Fig. 1. Production of recombinant His6–PrrA and His6–PrrB. His6–PrrA (a) and His6–PrrB (b) were overproduced in E. coli SG13009(pQE30-PrrA) and E. coli SG13009(pQE30-PrrB), respectively, and purified. The non-induced (lanes 1), soluble (lanes 2) and purified (lanes 3) fractions were analysed by SDS-PAGE and Coomassie blue staining. The arrows indicate the positions of His6–PrrA and His6–PrrB. Molecular mass markers (in kDa) are shown on the left-hand side of the images.

 
Phosphotransfer between His6–PrrB and His6–PrrA
The His6–PrrB and His6–PrrA phosphorylation assays were performed in the presence of Mn2+ ions, which have been demonstrated to be more efficient than Mg2+ ions for the mycobacterial TrcS histidine kinase activity (Haydel et al., 1999). As shown in Fig. 2, His6–PrrB was able to autophosphorylate (lane 1). When His6–PrrA was added to the His6–PrrB phosphorylation mixture, phosphotransfer to His6–PrrA was observed (lane 2). In the absence of His6–PrrB, no detectable His6–PrrA phosphorylation occurred (lane 3).



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Fig. 2. Autophosphorylation of His6–PrrB and phosphotransfer on His6–PrrA. Autophosphorylation of His6–PrrB (lane 1) and phosphotransfer to His6–PrrA (lane 2) were performed as described in Methods. The reaction mixtures were subjected to SDS-PAGE and autoradiography. Lane 3 contains His6–PrrA incubated in the absence of His6–PrrB.

 
His6–PrrA DNA binding activity
The ability of His6–PrrA to bind to a 317 bp PCR fragment containing the prrAprrB promoter region was assessed by electrophoretic mobility shift assays. Fig. 3 shows that His6–PrrA was able to bind to this region. Binding to the labelled target was inhibited by the presence of an excess of unlabelled specific competitor (lane 4) but not by an excess of non-specific DNA (lane 5). These results indicate that His6–PrrA specifically binds to the prrAprrB promoter region and suggest that the genes are autoregulated.



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Fig. 3. DNA binding of His6–PrrA. Electrophoretic mobility shift assays were performed in the absence of His6–PrrA (lane 1), or in the presence of 1 µg (lane 2) or 2 µg (lanes 3–5) unphosphorylated His6–PrrA or of 0·5 (lane 6), 1 µg (lane 7) or 2 µg (lanes 8–10) of phosphorylated His6–PrrA. A 50-fold excess of specific (lanes 4 and 9) or a 100-fold excess of non-specific (lanes 5 and 10) competitor DNA was added to the same reaction mixtures.

 
To test the effect of phosphorylation on the His6–PrrA binding activity, mobility shift assays were carried out using the same DNA target and phosphorylated His6–PrrA. The electrophoretic mobility shifts were increased with phosphorylated His6–PrrA in comparison to unphosphorylated His6–PrrA, when the same amounts of His6–PrrA and His6–PrrA~P were compared with each other (compare lanes 2 and 3 with lanes 7 and 8, respectively).

Autoregulation of the prrAprrB operon
To confirm autoregulation of the prrAprrB operon, we introduced pFluo1Sm, a plasmid containing the gfp reporter gene under control of the prrAprrB promoter region, into the M. tuberculosis Mt103 wild-type strain or the Mt21D3 prrA/B mutant derivative (Ewann et al., 2002). Since prrAprrB is not expressed in axenic culture conditions, and since the expression of this operon is induced early after macrophage phagocytosis (Graham & Clark-Curtiss, 1999; Ewann et al., 2002), murine bone-marrow-derived macrophages were infected with the recombinant strains. A BCG strain containing the same construct and a BCG strain containing pJFX4 and constitutively expressing gfp were used as controls. The fluorescence of the infected macrophages was measured by flow cytometry. Induction of fluorescence was readily observed with macrophages infected by BCG or M. tuberculosis Mt103 containing the prrA : : gfp construct (Fig. 4b, c). In contrast, no fluorescence was detected when the macrophages were infected with the M. tuberculosis prrA/B mutant containing the prrA : : gfp construct (Fig. 4d). Although effects of different copy numbers of the prrA : : gfp plasmid in the wild-type and in the prrA/B mutant can not be totally ruled out, these results, taken together with the results of the mobility shift assays, indicate that the prrAprrB operon is autoregulated in M. tuberculosis.



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Fig. 4. Autoinduction of prrA after macrophage infection. Mean fluorescence intensities of murine bone-marrow-derived macrophages were analysed after infection by recombinant BCG (b), M. tuberculosis Mt103 (c) or the prrA/B-deficient strain Mt21D3 (d) containing prrA : : gfp, as described in Methods. Non-infected macrophages (a) were used as a negative control.

 

   DISCUSSION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Adaptive responses upon macrophage infection are probably critical in the infectious cycle of the intracellular pathogen M. tuberculosis. The genes of at least three two-component systems have been shown to be induced in M. tuberculosis or in M. bovis BCG during growth within macrophages (Ewann et al., 2002; Graham & Clark-Curtiss, 1999; Zahrt & Deretic, 2001). Among these systems, the PrrA–PrrB system has been shown to be transiently required for early intracellular multiplication of M. tuberculosis (Ewann et al., 2002). Electrophoretic mobility shift assays indicate that PrrA specifically binds to its own promoter region, suggesting that, like for several other response regulators belonging to the OmpR family, the prrA gene expression is autoregulated. The absence of detectable prrA expression in culture growth conditions hindered a more detailed mapping of the prrA transcriptional start site using primer extension experiments. Recent studies using electrophoretic mobility shift assays and reporter gene expression in heterologous E. coli or M. smegmatis systems have suggested that the RegX3 and TrcR mycobacterial response regulators are also autoregulated (Haydel et al., 2002; Himpens et al., 2000). However, the signals involved in this autogenous control have not been identified yet. Here, autoregulation of the PrrA–PrrB system was indicated by the fact that intracellular expression of prrA : : gfp depends on the presence of the PrrA–PrrB system in M. tuberculosis itself, as it is abolished in a M. tuberculosis prrA/B mutant. Conversely, this abolition shows that this autoregulatory loop controls the intracellular activation of prrAprrB.

Increased binding to the prrAprrB promoter region was observed upon phosphorylation of PrrA, which is consistent with the fact that the intracellular activation of the prrAprrB operon actually depends on both the presence and activity of the PrrA–PrrB two-component system itself. The in vitro transphosphorylation was relatively inefficient under the conditions used in this study, as only a minor fraction of PrrA could be phosphorylated via PrrB. Therefore, the observed effect of PrrA phosphorylation on DNA binding was probably not optimal. Modifications of the Mn2+ concentrations or the replacement of Mn2+ by other bi-valent ions did not significantly improve the enzymic activities (data not shown). It may be possible that optimal phosphotransfer requires additional factors yet to be identified or requires PrrB sequences that are absent from His6–PrrB.

Several other M. tuberculosis genes have been shown to be induced concomitantly to prrA during the first days of the macrophage infection (Graham & Clark-Curtiss, 1999). These genes encode proteins with various functions, such as sigma factors and cation transporters, as well as proteins involved in lipid and cell-wall metabolism or in intracellular invasion or persistence in mice (Chitale et al., 2001; Cole et al., 1998; Kolattukudy et al., 1997; Manganelli et al., 1999; McKinney et al., 2000). The potential control of intracellular induction of these genes by the PrrA–PrrB system can now be investigated using a strategy similar to that described here and used to demonstrate prrA autoregulation. This approach may perhaps be complemented by a non-targeted, albeit more delicate, proteomic and transcriptomic analysis of the available prrA/B mutant grown intracellularly.


   ACKNOWLEDGEMENTS
 
Jean-Louis Herrmann and Ludovic Tailleux are gratefully acknowledged for their help in flow cytometry analysis. The work was supported by INSERM, Institut Pasteur de Lille, a grant from the Ministère de l'Education Nationale, de la Recherche et de la Technologie. F. E. holds a fellowship of the Région Nord-Pas-de-Calais and the Fondation pour la Recherche Médicale. P. S. is a Chercheur du Centre National de Recherche Scientifique.


   REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
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Received 27 May 2003; revised 17 September 2003; accepted 19 September 2003.