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Institute of Genetics, University of Bayreuth, D-95440 Bayreuth, Germany
Correspondence
Wolfgang Schumann
wschumann{at}uni-bayreuth.de
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ABSTRACT |
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Supplementary tables of strains and primers are available with the online version of this paper.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Errington, 2003). Bacillus subtilis cells code for an extremely complex and sophisticated decision-making apparatus, which monitors a huge range of internal and external signals. This information is channelled through several separate regulatory systems, of which the most prominent component is the master regulator Spo0A, a DNA-binding protein which directly affects transcription of a total of 121 genes, either negatively or positively (Molle et al., 2003). The synthesis of Spo0A is controlled at the transcriptional level, and the activity of the protein is regulated by phosphorylation through the phosphorelay signal-transduction system (Burbulys et al., 1991; Hoch, 1993). The transfer of the phosphoryl group to Spo0A involves a complex network consisting of several kinases (KinA, KinB, KinC, KinD and KinE), where each probably responds to a different stimulus (Jiang et al., 2000b). Upon autophosphorylation, the phosphoryl group is transferred by two intermediates, Spo0F and Spo0B, and finally to Spo0A (Burbulys et al., 1991). Two of the three phosphoproteins are subject to regulation by phosphatases, and one group, the Rap phosphatases, are regulated by pentapeptides (Perego, 1998). The Rap phosphatases specifically dephosphorylate Spo0F
P, while another three phosphatases (Spo0E, YisI and YnzD) attack Spo0AP.
The ftsH gene, coding for a membrane-anchored metalloprotease, is present in most if not all bacterial species (Schumann, 1999; Ogura & Wilkinson, 2001). The FtsH protein and its biochemical and biological functions have been studied in detail in Escherichia coli. It carries two transmembrane segments close to its N-terminus which anchor the protein into the cytoplasmic membrane in such a way that both its short N- and its long C-terminus are exposed into the cytoplasm (Tomoyasu et al., 1993a). The C-terminal part contains a Walker A and B box, involved in binding and hydrolysis of ATP (Tomoyasu et al., 1993b), and a binding site for Zn2+. The Aquifex aeolicus FtsH protein devoid of its transmembrane segments has been crystallized and shown to form a ring-like hexameric structure (Suno et al., 2006). While the ftsH gene in E. coli is essential (Ogura et al., 1999), a B. subtilis ftsH knockout is viable, but displays a pleiotropic phenotype (Deuerling et al., 1997; Lysenko et al., 1997). Cells with an ftsH null allele are sensitive to heat and osmotic stress, grow largely as filaments and last, but not least, exhibit a significantly reduced sporulation frequency. The reduced sporulation frequency seems to interfere with the synthesis or/and activity of Spo0A since an abrB-lacZ transcriptional fusion fails to be turned off during stage 0 (Deuerling et al., 1997). Based on this result we assume that the FtsH protease has to degrade one or more proteins involved, directly or indirectly, in the production of a sufficient amount of active Spo0A. To this end, we have identified the Spo0E phosphatase as one of the targets of FtsH, and our observations indicate that the C-terminus of Spo0E is necessary for its degradation. Furthermore, we show that FtsH is needed only during stage 0.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Construction of plasmids and recombinant strains.
All transcriptional fusions were constructed using the integration vector pDG1728 (Guérout-Fleury et al., 1996). This vector contains a promoterless lacZ and allows insertion of the operon fusions ectopically at the amyE locus. Three different promoters were fused to lacZ generated by PCR using chromosomal DNA of strain 1012 DNA as template. These promoters are Pskf (amplified by primers ON1 and ON2; see Supplementary Table S2) preceding the skf operon, which is activated by a low amount of active Spo0A (Fujita et al., 2005), and the vegetative- and the stationary-phase-induced promoters Pv (ON3/ON4) and Ps (ON5/ON6), respectively, of the spo0A gene (Chibazakura et al., 1991). Pskf was inserted between the EcoRI and HindIII sites of pDG1728; Pv and Ps were ligated into the EcoRI and BamHI sites.
Inactivation of the four genes rapA, rapB, rapE and spo0E was achieved by the replacement method as follows. First, the two flanking regions of each gene (about 300 bp) were amplified (see Supplementary Table S2 for the primer sequences) and inserted into pBluescript SKII+. Next, the chloramphenicol-resistance cassette was amplified using pDG1662 as template and inserted between the flanking regions of the three rap genes. In the case of the spo0E gene, a phleomycin-resistance marker generated from pBluescript SKII+-phleo was ligated between the two flanking regions. PCR fragments containing the resistance marker and the flanking regions were transformed into B. subtilis 1012 followed by selection on LB plates containing either chloramphenicol or phleomycin. Chromosomal DNA was prepared from several transformants each and checked by Southern blotting for replacement of the wild-type alleles. One knockout mutant each was kept for further studies.
Recombinant vectors allowing overexpression and purification of GST-tagged proteins were prepared using pGEX-2T. The genes ftsH (ON21/22), spo0E (ON23/ON24), yisI (ON25/26) and ynzD (ON29/ON30) were amplified using chromosomal DNA of strain 1012. The mutants spo0E11 and spo0E94 carry stop codons at positions 72 and 60, respectively (Perego & Hoch, 1987). The two truncated versions were generated by amplification of the appropriate coding region (spo0E11, ON23/28; spo0E94, ON23/27) followed by a stop codon. To construct a ynzD-spo0E fusion, the coding region for the last 25 aa of spo0E was fused to ynzD using ON31/ON32. All amplicons were fused in-frame to the coding region of GST.
Expression and purification of GST-tagged proteins.
The recombinant pGEX-2T plasmids were transformed into E. coli strain A8296 (Tatsuta et al., 1998). This strain carries an ftsH knockout to avoid production of two types of FtsH proteins, one encoded by the E. coli and the other by the B. subtilis gene. Expression was induced by adding 1 mM IPTG (final concentration) to the cell cultures at an OD600 of 0.8 followed by transfer of the cultures to 25 °C and further growth overnight. GST-tagged proteins were purified by binding to glutathione (GSH)-agarose beads followed by elution with GSH. A detailed description of the purification procedure has been published (Teff et al., 2000).
Proteolysis experiments.
Degradation reactions were performed as described by Tomoyasu et al. (1995). The complete reaction mixture (30 µl) consisted of the following components: 50 mM Tris/acetate (pH 8.0), 5 mM magnesium acetate, 12.5 µM zinc acetate, 80 mM NaCl, 1.4 mM β-mercaptoethanol, 5 mM ATP, 50 µg BSA ml–1, 100 µg ml–1 of the target GST-tagged protein (or 100 µg ml–1 of β-casein, as a positive control to verify the proteolytic activity of purified GST-FtsH), 50 µg ml–1 of purified B. subtilis GST-FtsH and 1 µl EDTA-free ‘Complete’ inhibitor mix (Roche Diagnostics) solution. The solution was prepared from one inhibitor tablet dissolved in 1 ml water. Reactions were performed at 40 °C for the time points indicated. Aliquots of the reaction mixtures were analysed by 15 % SDS-PAGE followed by staining with Coomassie blue.
β-Galactosidase assay.
Cells were grown in DSM at 37 °C and samples were collected at the indicated time points. β-Galactosidase assays were performed in triplicate on soluble extracts using o-nitrophenyl-β-D-galactoside as substrate (Miller, 1972) and yielded comparable results. The activities of one representative experiment are presented. β-Galactosidase activities are given in units, where one unit is defined as 
A405 min–1xOD578–1x10–3, in which OD578 is the optical density of the growth culture.
Western blots.
Western blotting was carried out as described by Towbin et al. (1979) except that immunoblots were developed by an ECL Western blotting detection kit (Amersham) according to the manufacturer's instructions. Polyclonal anti-Spo0A antibodies were used for the detection of Spo0A (Fujita et al., 2005).
Determination of sporulation frequency.
Sporulation frequencies were determined by the heat-resistance assay (Harwood & Cutting, 1990). Briefly, the strains were inoculated in 50 ml DSM and incubated at 37 °C for 36 h. Cells were serially diluted in potassium phosphate buffer (10 mM potassium phosphate buffer, pH 7.4, supplemented with 50 mM KCl and 1 mM MgSO4), and 100 µl samples of appropriate dilutions were plated on DSM agar to determine the number of vegetative cells. Cells were challenged at 80 °C for 20 min, and then 100 µl samples were plated on DSM agar to determine the number of heat-resistant spores.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). To confirm this assumption, we visualized the amount of Spo0A produced in the presence and absence of ftsH by Western blotting. Cells were grown in DSM and aliquots removed at t0 (corresponding to the end of exponential growth and entry into transition phase) to t3. As can be seen from Fig. 1, Spo0A started to be present from stage 0 on in the wild-type strain and continued to be produced to at least stage 3. In contrast, in the ftsH knockout, Spo0A was present in greatly reduced amounts (Fig. 1).
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P) are sufficient to activate the skf operon (Fujita et al., 2005). When this fusion was analysed in the wild-type background, the β-galactosidase activity started to increase from stage 0 onwards (Fig. 2a). When the same operon fusion was tested in the ftsH knockout, only a very low background activity was measured, without any increase at least up to t3 (Fig. 2a). These results confirm that ftsH does indeed interfere with the synthesis or/and activation of Spo0A, where both regulatory circuits are interwoven (Strauch et al., 1992).
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P, the second component of the phosphorelay (Perego, 1998). First, we asked whether ftsH influences the sporulation frequencies in the presence or absence of one of the three phosphatases. We constructed knockouts in all three genes as described in Methods. Then, these null alleles were each combined with an ftsH knockout, and all six strains were analysed for their sporulation frequencies, for the amount of Spo0A present and for its activity status. While about 59 % of the cells in our wild-type strain were able to form heat-resistant spores (Table 1), the sporulation frequency dropped by five orders of magnitude in the absence of the FtsH protein, confirming earlier data (Deuerling et al., 1997). The sporulation frequencies in all three rap knockouts were higher than that of the wild-type strain and ranged from 67 % to 72 %. A similar observation has been published for RapA and RapE (Jiang et al., 2000a). The sporulation frequencies in the double knockouts were increased by two to three orders of magnitude as compared to a single ftsH null mutant, but remained below 1 % (Table 1). These data clearly indicate an influence of the ftsH allele on all three Rap phosphatases.
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shows the results from a representative assay. The absence of any of the three phosphatases in the otherwise wild-type background exhibited a different outcome. While in the absence of either rapA or rapB the amount of Spo0A was increased at t0 as compared to the wild-type situation, its amount was reduced at both t0 and t1 in the rapE knockout (Fig. 1). When the ftsH null allele was added, the amount of Spo0A dropped, as already observed for the wild-type strain in the absence of ftsH (Fig. 1). Is the Spo0A protein present in the double knockouts active? To answer this question, the Pskf-lacZ fusion was introduced into all six strains followed by measurement of the β-galactosidase activities of the strains grown in sporulation medium. Expression of the Pskf-lacZ fusions was somewhat different among the six strains. While expression started in all strains at t0 and reached its plateau value at t2 in the wild-type and in the rapB strains, it further increased in the rapA and the rapE strains (Fig. 2). In the presence of ftsH, the expression of the operon fusion did not increase over the basal level, with the exception of rapE, where a slight increase to about 20 units was observed (Fig. 2d). These results suggest that both the rapA and the rapE genes could be involved in shutting off Pskf-lacZ at t2.
The FtsH protein interferes with the phosphorylation status of Spo0A through Spo0E
Besides the Rap phosphatases, another set of three phosphatases is involved in the specific dephosphorylation of Spo0AP, designated Spo0E, YisI and YnzD; only the first is active during sporulation (Perego, 2001). While overproduction of Spo0E reduced the sporulation frequency, deletion of spo0E resulted in an increase (Perego & Hoch, 1991). We constructed a spo0E knockout, combined it with the ftsH null allele and measured the sporulation frequencies in both strains. As to be expected the sporulation frequency rose in the absence of the spo0E gene, above the level observed in the wild-type strain (Table 1). If combined with an ftsH knockout, the sporulation frequency was increased 1000-fold over the level measured in the ftsH strain, but was still about 100-fold lower than the wild-type level (Table 1). Next, we analysed the production of Spo0A in both mutant strains. Western-blot analysis revealed that Spo0A was present in large amounts already at t0 in the spo0E strain and remained at that high level for the next 3 h (Fig. 1). In the ftsH strain, the amount of Spo0A was reduced at t0 and further increased to levels comparable to those present in the ftsH+ strain (Fig. 1). This result suggests an interaction between the two proteins, either directly or indirectly, thereby influencing expression of spo0A. When we tested for the activity of Spo0A in both mutant strains, it was found to result in a higher activation of the skf promoter in the ftsH+ strain, but completely failed to activate this promoter in the absence of ftsH (Fig. 2e). In conclusion, the absence of an active spo0E allele in an ftsH knockout leads to expression of the spo0A gene, but the protein remains inactive. These data indicate that ftsH influences production of active Spo0A by either promoting its phosphorylation or preventing its rapid dephosphorylation.
Spo0E is a target protein for FtsH
One possibility to explain the interaction between FtsH and Spo0E is a direct one whereby FtsH degrades Spo0E. To test that possibility, both proteins were purified and incubated under conditions where FtsH is able to degrade β-casein (Kotschwar et al., 2004). FtsH was purified with a GST-tag as reported before; the purification tag keeps the protein soluble in the absence of any detergent (Kotschwar et al., 2004). Since we failed to overproduce Spo0E equipped with a His-tag (unpublished data), we decided to add the GST-tag as well. Next, both proteins were incubated in the presence and absence of ATP. While in the absence of ATP, the GST-Spo0E remained stable during a 4 h incubation, it was largely, but not completely, degraded in the presence of the nucleotide (Fig. 3a). This could be verified by probing some lanes with anti-GST antibody (Fig. 3b). To rule out the possibility that cleavage occurs at or within the GST tag rather than within Spo0E, this tag was purified and incubated with GST-FtsH. GST remained stable for at least 5 h (data not shown).
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). Are these two phosphatases also a substrate of FtsH? While β-casein was completely degraded within 5 h of incubation, both GST-YisI and GST-YnzD remained stable under these conditions (Fig. 4). These data clearly demonstrate that neither YisI nor YnzD is a substrate of FtsH. They further confirm that the GST-tag is not recognized by FtsH.
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P (Perego & Hoch, 1991; Ohlsen et al., 1994). Both mutations resulted in a stop codon, reducing the length of the proteins from 85 to 71 (spo0E11) and 59 aa (spo0E94) (Perego & Hoch, 1987; Ohlsen et al., 1994). To find out whether these two shortened versions of the Spo0E protein are still a target for FtsH, both were tagged with GST, overproduced in E. coli and purified. When the two purified proteins were incubated with FtsH, both remained stable for at least 5 h, while β-casein as a control was degraded under these conditions (Fig. 5). Since the full-length Spo0E protein is unstable when incubated with FtsH, we infer that the C-terminal 25 aa are responsible for this instability.
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). Since Spo0E serves as a target for FtsH, but YisI and YnzD do not, we asked whether the C-terminal extension of Spo0E is responsible for recognition by FtsH. To answer this question, we fused the coding region for the C-terminal 25 aa to ynzD (YnzD-0E). The GST-tagged hybrid protein was overproduced in E. coli, purified by affinity chromatography and incubated with GST-FtsH. As can be seen from Fig. 6 (lane 6), YnzD-0E was largely degraded over time. We conclude from this experiment that indeed the C-terminal region of Spo0E contains the recognition sequence for the FtsH protease.
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). We asked whether there is a crosstalk between these genes concerning their expression level. Is there increased expression of either yisI or/and ynzD in a spo0E knockout? First, we fused the promoters of the two genes to lacZ and integrated both transcriptional fusions at the amyE locus. Next, the spo0E knockout was introduced into both strains. Then, all four strains (AL53–AL56; see Supplementary Table S1) were grown in DSM, samples were taken from t0 up to t3, and the β-galactosidase activity was determined. While no difference was measured for the ynzD promoter in the presence and absence of the spo0E allele (Fig. 7b), there was a slight increase in the transcription of the yisI gene (Fig. 7a) in the absence of spo0E. We conclude that removal of the spo0E gene did not influence expression of the two other genes significantly, excluding a crosstalk at the transcriptional level.
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). The aspartate residue at position 56 of Spo0A acts as the phosphorylation site (Burbulys et al., 1991) and is dispensable in the spo0A-sad67 allele (Ireton et al., 1993). We asked whether an ftsH knockout strain is able to form spores in the presence of the spo0A-sad67D56N allele. We measured the sporulation frequencies in strain SIK190, which carries the spo0A-sad67D56N allele fused to an IPTG-inducible promoter. While a sporulation frequency of 0.06 % was measured in the absence of IPTG (Table 2), induction of the mutant allele at t0 resulted in 37 % heat-resistant cells. Then the ftsH : : tet knockout was introduced into SIK190 (SIK190F) and the sporulation frequency was determined. While the sporulation frequency was low in the absence of IPTG, it was high after IPTG induction and both values were comparable to those measured in the ftsH wild-type strain (Table 2). These results strongly suggest that ftsH is needed only during phase 0 and that the spo0A-sad67D56N allele can be expressed in the absence of ftsH.
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; Kudoh et al., 1985). While Pv is recognized by the housekeeping sigma factor 
A and functions as a low-level promoter to produce a maintenance level of the Spo0A protein during exponential growth (Yamashita et al., 1989), the second promoter, Ps, is recognized by the stationary sigma factor H. This promoter is required for induction of the protein at the end of exponential growth and during stage 0 (Ferrari et al., 1985; Yamashita et al., 1989) and is further activated indirectly by Spo0AP, which represses expression of abrB, a negative regulator of sigH (Perego et al., 1988). Therefore, phosphorylation of Spo0A at the onset of sporulation activates an autoregulatory loop leading to an increase in activated Spo0A. We asked whether ftsH or/and spo0E influences transcription at either promoter. The two promoters were separately fused to lacZ and the transcriptional fusions were ectopically integrated at the amyE locus. Then, either the ftsH or the spo0E knockouts or both were added and the β-galactosidase activities were determined in all eight strains. While the β-galactosidase activity initiated at Pv increased slightly up to t1 followed by a modest decrease in the wild-type strain, its activity was reduced to about 50 % in the ftsH knockout (Fig. 8a). The enzymic activity in the spo0E null mutant was comparable to that measured in the wild-type strain, but addition of the spo0E null allele to that of ftsH resulted in a slight increase in the β-galactosidase activity (Fig. 8a). In summary, the influence of both spo0E and ftsH on the Pv promoter is minor.
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). In the absence of spo0E, it was induced about tenfold, while only a threefold induction was measured in the ftsH knockout, which was not increased in the double knockout (Fig. 8b). We conclude that transcription at Ps is strongly reduced in ftsH, an effect which is not compensated by spo0E. In total, ftsH influences only transcription at Ps, most probably through the strongly reduced level of active Spo0A, which is needed as part of the autoregulatory loop. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). It is also known that never do 100 % of the cells sporulate. The decision to sporulate or not to sporulate is dependent on the amount of active Spo0A at the end of stage 0, which takes about 2 h. Sporulating cells sense a multitude of mostly unknown signals, including the metabolic state, the fate of the chromosomes and the cell density, and integrate and process these signals by a phosphorelay which controls the level of phosphorylated Spo0A. It turned out that, while about 60 % of the cells manage to synthesize active Spo0A above a threshold value [these cells have been called Spo0A-ON (Chung et al., 1994)], the remaining 40 % fail to do so (Spo0A-OFF). The formation of two subpopulations of otherwise isogenic cells is designated bistability (Smits et al., 2006; Dubnau & Losick, 2006). The molecular basis for this bistability remains elusive. But it has to be asked whether cells in the transition phase exhibit multi- rather than bistability based on the observation that so many different genetic programmes are activated. This can be tested by double and triple labelling using transcriptional fusions between programme-specific promoters and gfp and its derivatives (Margolin, 2000).
Several years ago, we and others discovered that the sporulation frequency in an ftsH knockout is reduced by about five orders of magnitude (Deuerling et al., 1997; Lysenko et al., 1997). The ftsH gene codes for a membrane-anchored ATP-dependent metalloprotease which seems to be present in all bacterial species (Schumann, 1999; Ogura & Wilkinson, 2001). The objective of this ongoing research project is to elucidate the role of the FtsH protease during sporulation of B. subtilis. It is based on the assumption that FtsH has to degrade or to regulate the steady-state level of one or more proteins negatively interfering with the synthesis of active Spo0A. Cells carrying an ftsH knockout synthesize a significantly reduced amount of Spo0A which, based on genetic data, is inactive (Figs 1 and 2). This observation explains why ftsH null mutants exhibit a dramatically reduced sporulation frequency. But ftsH could also play a role during subsequent sporulation stages. This possibility was ruled out by introduction of an ftsH null allele into a strain carrying a mutant spo0A allele which is active in the absence of phosphorylation. Upon expression of the spo0A-sad67D56N allele, cells exhibited a normal sporulation frequency (Table 2). Therefore, the role of ftsH is exclusively confined to the synthesis or/and activation of wild-type Spo0A.
The next question to be raised is why the amount of Spo0A is significantly reduced and why this reduced amount is inactive. So far, we have used the candidate approach to identify putative substrate proteins of FtsH. First, we constructed knockouts by the gene replacement method in four different genes coding for phosphatases for which it has already been reported that the sporulation frequencies are slightly, but reproducibly enhanced in the absence of two of them (Jiang et al., 2000a; Perego & Hoch, 1987); these results could be confirmed and extended for an additional phosphatase (RapB). Upon introduction of an ftsH null allele into these four mutant strains, the sporulation frequencies were increased by two to three orders of magnitude, but still remained very low (Table 1). Not surprisingly, no increase in the amount of active Spo0A could be measured. On the basis of this analysis, we conclude that ftsH interferes directly or indirectly with expression or activity of these four phosphatases. In the double knockout, only about 1 % of the cells reach the Spo0A-ON state. This conclusion can be confirmed by fusing the promoter of the skf and the spoIIA operons to gfp and analysing single cells under the fluorescence microscope. Whereas the skf promoter needs a low amount of active Spo0A, the spoIIA promoter requires a high amount to become activated (Fujita et al., 2005).
Based on the results shown in Table 1, we asked whether the phosphatases are a substrate for FtsH. To this end, we overexpressed and purified the Spo0E phosphatase and incubated it with purified FtsH. Indeed, Spo0E is partially degraded by FtsH (Fig. 3); there are at least three possibilities to explain this result. First, our GST-FtsH protease does not exhibit full activity. Second, part of Spo0E is refractory to digestion by FtsH, and third, the GST-tag provides partial protection. We favour the second possibility based on the following observations. In E. coli, two substrate proteins have been identified for which FtsH controls their steady-state level: LpxC and KdtA; both are involved in the LPS biosynthetic pathway (Führer et al., 2006; Katz & Ron, 2008). LpxC is a key enzyme in LPS formation and catalyses the second reaction and the first committed step in the biosynthesis of lipid A (Sorensen et al., 1996). Overproduction of LpxC causes accumulation of abnormal membranes in the periplasm (Ogura et al., 1999), which is toxic for the cell (Sullivan & Donachie, 1984), indicating that the amount of LpxC must be carefully regulated, and this occurs at the protein level via degradation by the protease FtsH (Führer et al., 2006). KdtA is the only essential glycosyltransferase in the core oligosaccharide biosynthesis pathway (Belunis & Raetz, 1992). Here, too, FtsH regulates the concentration of the precursor of the sugar moiety 3-deoxy-D-manno-octulosonate (KDO) via the KdtA enzyme (Katz & Ron, 2008). When both purified enzymes were incubated with purified FtsH protease, no complete degradation was observed, reminiscent of our observation that complete degradation of Spo0E could not be obtained.
Why is FtsH unable to fully degrade these three proteins, at least under in vitro conditions? We assume that these proteins are present in two different conformations, one recognized and the other not recognized by FtsH. An interesting question is whether this ratio can be influenced by a mechanism, e.g. by molecular chaperones.
Perego (2001) identified two homologues of the Spo0E phosphatase, YisI and YnzD. She showed that both phosphatases are able to dephosphorylate Spo0AP in vitro. These two phosphatases are distinguished from Spo0E by two characteristics: first, their genes are expressed during the vegetative growth phase; and second, they lack a C-terminal extension of about 25 aa. She suggested that this C-terminal extension could be recognized by a protease. When GST-tagged YisI or YnzD was incubated with FtsH, these proteins turned out to be stable under the same conditions (Fig. 4). Two truncated versions of Spo0E also turned out to be stable (Fig. 5). Based on these findings, we asked whether the C-terminal region of Spo0E is responsible for its instability. When the C-terminal extension of Spo0E was fused to YnzD, the fusion protein was susceptible to degradation by FtsH (Fig. 6). We conclude that the C-terminal 25 aa of Spo0E contain the residues recognized by FtsH. Since the spo0E11 mutation turned out to be insensitive to proteolysis by FtsH, the recognition site can be narrowed down to the last 14 aa.
Which amino acids are recognized by the FtsH protease? One of the substrate proteins of the E. coli FtsH protease is the LpxC enzyme (Ogura et al., 1999). This enzyme represents the key enzyme in lipopolysaccharide (LPS) formation and controls the ratio between LPS and phospholipids (Sorensen et al., 1996). Since overproduction of LpxC causes accumulation of abnormal membranes in the periplasm (Ogura et al., 1999), leading to cell death (Sullivan & Donachie, 1984), the amount of LpxC must be carefully regulated; this is done by FtsH. Here, too, the C-terminus has been identified as being responsible for recognition and degradation by FtsH (Führer et al., 2006), specifically a stretch of about ten amino acids at the immediate C-terminus, which resembles the SsrA-tag. In the case of Spo0E, there is no similarity to the B. subtilis SsrA-tag (Wiegert & Schumann, 2001). Therefore, the amino acid sequence recognized by the FtsH protease is different from that of the SsrA-tag. We suggest that FtsH regulates the steady-state level of Spo0E in a similar way. This can be analysed by creating a Spo0E-GFP fusion protein followed by single-cell analysis as described above.
The studies reported here strongly suggest that regulation of stability of several proteins involved directly or indirectly in the synthesis of active Spo0A exerts a new level of post-translational regulation through the FtsH protease. Another protease has been identified yielding a comparable phenotype. Inactivation of clpP resulted in cells deficient in sporulation initiation and in competence and in a highly filamentous morphology (Msadek et al., 1998; Gerth et al., 1998). In such a mutant, the expression of spo0A and spo0H, coding for the stationary sigma factor H, was significantly decreased (Nanamiya et al., 2000). Introduction of a mutant spo0E allele into the clpP knockout restored the expression of spo0A, but not sporulation. Based on our results, additional genes influencing the synthesis of active Spo0A need to be identified. This will be accomplished by three different experimental strategies: First, the candidate strategy; second, saturated transposon mutagenesis using pMarA (Le Breton et al., 2006); and third, construction of an ftsH trap mutant (Flynn et al., 2003). Identification of these additional targets should shed light on the molecular mechanism of bi- or even multistability.
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ACKNOWLEDGEMENTS |
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Edited by: M. Hecker
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REFERENCES |
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Received 11 September 2008;
revised 21 November 2008;
accepted 26 November 2008.
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