Journal of Bacteriology, November 2002, p . 6109-6114, Vol . 184, No . 22

The PrpC Serine-Threonine Phosphatase and PrkC Kinase Have Opposing Physiological Roles in Stationary-Phase Bacillus subtilis Cells

Tatiana A . Gaidenko, Tae-Jong Kim, and Chester W . Price^*

Department of Food Science and Technology, University of California, Davis, California 95616

Received 18 December 2001/ Accepted 19 August 2002

Three of these kinases (RsbT, RsbW, and SpoIIAB) and four of the phosphatases (RsbP, RsbU, RsbX, and SpoIIE) act via a partner-switching mechanism to control the activities of the general stress transcription factor ^B and the forespore-specific transcription factor ^F (reviewed in reference 19) . The three partner-switching kinases are unusual in that their sequences reflect more of a kinship with bacterial histidine protein kinases than with typical eukaryotic serine-threonine kinases (9, 15) . The four partner-switching phosphatases all belong to the PPM/PP2C family, but they are also unusual in that they lack conserved domains V_a and V_b that are commonly found in eukaryotic PP2C phosphatases (22) .

In contrast, the PrpC phosphatase and the PrkC kinase more closely resemble their eukaryotic counterparts, but their physiological roles are unknown (18, 22) . Because it is thought that at least one serine-threonine phosphatase in the ^B signal transduction network remains to be discovered (10), we sought to determine the effects of loss-of-function mutations within prpC and prkC . Given the multiple phenotypes elicited by these mutations, we infer that PrpC and PrkC have a significant role in controlling a variety of stationary-phase processes .

ß-Galactosidase accumulation assays. For the experiments shown in Table 1, cells were grown into stationary phase in buffered Luria broth (LB) lacking salt (BLB) (5) . For environmental stress experiments (data not shown), cells were grown to early logarithmic phase in BLB, at which point NaCl was added to a 0.3 M final concentration . For both assays samples were collected and treated as described by Miller (14) . Cells were washed with Z buffer and permeabilized using sodium dodecyl sulfate (SDS) and chloroform . Protein levels were determined using the Bio-Rad protein assay reagent (Bio-Rad Laboratories, Richmond, Calif.) . Activity was defined as A₄₂₀ x 1,000 per minute per milligram of protein .

 
Western blotting experiments. Wild-type and mutant cultures were grown in BLB, harvested in logarithmic phase (50 Klett units in a Klett-Summerson colorimeter, using a number 66 filter), and then resuspended in sonication buffer (50 mM Tris, pH 7.5; 50 mM MgCl₂) . Cells were broken by sonication, and extracts were prepared by centrifugation . Proteins were separated on SDS-polyacrylamide gels and then transferred to polyvinylidene difluoride membranes (Bio-Rad Laboratories) . These blots were blocked and washed in membrane blocking solution (Zymed Laboratories, San Francisco, Calif.) that was compatible with the primary rabbit antibody, either antiphosphothreonine (catalog no . 71-8200; Zymed Laboratories) or antiphosphoserine (catalog no . 61-8100) . Blots were exposed to 1 µg of primary antibody/ml in TBS-T buffer (20 mM Tris, pH 7.6; 137 mM NaCl; 0.1% Tween 20) for 1 h at 25°C and then to the anti-rabbit immunoglobulin G (IgG) peroxidase-conjugated secondary antibody (Sigma, St . Louis, Mo.) . Blots were washed, and the bound antibody was visualized using the ECL Plus Western blotting detection kit (Amersham Pharmacia Biotech, Piscataway, N.J.), according to the manufacturer's instructions .

Immune precipitation experiments. We did two different types of immune precipitation experiments . The first was an immune precipitation from cell extracts, in which cells were grown and extracts made as described for the Western blotting . Here we began with a 150-ml culture, resulting in 4 ml of extract in sonication buffer . To this we added 9 µl of anti-Escherichia coli elongation factor G (EF-G) antibody, the generous gift of Andreas Savelsbergh and Wolfgang Wintermeyer . After incubating for 1 h on ice, we added 300 µl of protein A beads (Sigma) suspended in IP buffer (50 mM Tris, pH 8.0; 150 mM NaCl; 0.1% SDS) and then continued the incubation for 1 h at 4°C with slow shaking . The beads were collected by centrifugation, washed three times with IP buffer, and suspended in 150 µl of Laemmli sample buffer (60 mM Tris, pH 6.8; 100 mM dithiothreitol [DTT]; 2% SDS; 10% glycerol; 0.001% bromphenol blue) . After heating at 85°C for 10 min, samples were loaded onto a polyacrylamide gel for analysis by Coomassie staining and Western blotting (see Fig . 2B) . The second type of immune precipitation was to purify ³²P-labeled EF-G from in vitro kinase reactions . Here, we added 5 µl of anti-E . coli EF-G antibody to 210 µl of kinase reaction mix (see below) . After incubating for 1 h on ice, we added 100 µl of protein A beads suspended in IP buffer and then continued the incubation for 1 h at 4°C with slow shaking . The beads were collected by centrifugation and then washed three times with IP buffer and three times with kinase buffer lacking DTT (see below for kinase buffer) . The beads were resuspended in 300 µl of phosphatase buffer (see below), of which half was used to monitor the immune precipitation (see Fig . 3B) and half was used for the phosphatase release assay .

 
Kinase and phosphatase assays. We constructed overexpression clones which fused the fus (EF-G), prpC, and prkC coding regions to the hexahistidine tag in the pET15b expression vector (Novagen, Madison, Wis.) . Tagged proteins were purified from E . coli BL21(DE3) extracts on nickel affinity columns by using the manufacturer's protocol (Novagen) . The phosphatase and kinase rapidly lost activity when stored in glycerol at -20°C, so all assays were done with freshly purified proteins . For kinase assays, 5 µg of PrkC and 5 µg of EF-G (final concentration, 2 µM each) were incubated at 37°C for 30 min in 30 µl of kinase buffer (50 mM Tris, pH 7.6; 50 mM KCl; 10 mM MgCl₂; 1 mM DTT; 0.1 mM EDTA) together with 1 mM unlabeled ATP and 15 µCi of [-³²P]ATP (Perkin Elmer Life Sciences, Boston, Mass.) . This kinase reaction mixture was also used to prepare labeled substrate for the phosphatase assays . For the assay shown below in Fig . 3A, PrkC and EF-G were labeled in the kinase reaction, separated from unincorporated [-³²P]ATP on a nickel affinity column, resuspended in 30 µl of phosphatase buffer (kinase buffer plus 2 mM MnCl₂), and then incubated at 37°C with 2 µg of purified PrpC (2 µM final concentration) for the times indicated . Reactions were terminated by heating in Laemmli sample buffer, and the proteins were analyzed by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) . The phosphatase assay shown below in Fig . 3C was done essentially as described previously (25) . Immune-precipitated, labeled EF-G in 150 µl of phosphatase buffer was incubated at 37°C together with 5 µg of purified PrpC (1 µM final concentration) . Samples were removed at the times indicated, treated with cold trichloroacetic acid, and centrifuged . The supernatants were counted in a scintillation counter to determine the amount of -³²P released .

In-gel digestion and mass spectrometry. The band containing the protein of interest was excised from a 6% polyacrylamide gel, washed with Milli-Q water (Millipore, Bedford, Mass.), and diced, and the protein was reduced and alkylated (21) . The protein was digested in 50% ammonium bicarbonate containing sequence-grade, modified trypsin (Promega, Madison, Wis.) . After extraction with 0.1% trifluoroacetic acid and with 5% formic acid in 50% acetonitrile, peptide mass mapping was done using a Bruker Biflex III matrix-assisted laser desorption ionization-time-of-flight (TOF) mass spectrometer (Bruken-Franzen Analytik, Bremen, Germany) equipped with a pulsed N₂ laser (337 nm), a delayed extraction ion source, and a reflectron . Mass spectra were acquired in the reflectron mode . Internal mass calibration was performed with two trypsin-autodigested fragments, and then measured monoisotopic masses of the tryptic peptides from the protein of interest were used to search international databases . A mass accuracy of 50 ppm or better was used for each search . De novo sequencing of the tryptic peptides by tandem mass spectrometry (MS/MS) was done using hybrid nanospray-electrospray ionization-Quadrupole-TOF-MS and MS/MS in a QSTAR mass spectrometer (Allied Biosystems, Inc., Foster City, Calif.) . QSTAR instrument calibration was via a standard peptide mixture and routinely gave mass accuracies of 5 ppm or better . De novo sequencing of peptides was done using QSTAR software and confirmed by manual interpretation of MS/MS spectra .

In contrast to the absence of an observable phenotype in logarithmic or early-stationary-phase cells, the prpC and prkC null mutations had large effects on cell density during extended incubation in stationary phase . One such experiment is shown in Fig . 1 . In replicate growth experiments in BLB lacking salt, wild-type cell density decreased to a minimum between 23 and 27 h after the onset of stationary phase (Table 1) . Notably, the density of the prkC null mutant was significantly lower than that of the wild type, whereas the density of the prpC null mutant was significantly higher . Similar results were obtained with cells grown in standard LB, but in this medium wild-type density reached a minimum between 30 and 34 h after the onset of stationary phase (data not shown) .

 
We also did a standard plate count to measure cell viability at the point when wild-type cells reached minimum density (Table 1) . The observed differences between wild-type and mutant cells were reproducible but not significant at the 95% confidence level . However, microscopic examination showed that the prpC mutant formed abundant chains in stationary phase, suggesting that a standard plate count would greatly underestimate the number of viable cells . In contrast, both the wild-type strain and the prkC mutant were found as discrete cells (data not shown) .

Interestingly, the significant differences in cell density between wild-type and mutant strains were reflected in ß-galactosidase accumulation from the ^B-dependent ctc-lacZ fusion they carried . In particular, accumulation in the prkC mutant was significantly less than in the wild type (Table 1) . This distinction was not due to differential lysis, because ß-galactosidase activity was assayed in intact cells, collected by centrifugation . It therefore appears that ^B is less active during late stationary phase in the prkC mutant, or that the reporter fusion is more labile .

Based on the diverse effects the prpC and prkC mutations have on stationary-phase physiology (Table 1), we propose that the PrpC phosphatase and PrkC kinase have important regulatory roles in stationary-phase cells . Moreover, the prkC null allele is epistatic to the prpC null, at least with respect to the three phenotypes we tested . We therefore hypothesize that loss of PrpC phosphatase activity promotes high cell density in stationary phase due to an inability to counter PrkC kinase activity .

Cell extracts of the prpC null mutant have elevated levels of a protein recognized by antiphosphothreonine antibody. In order to identify possible protein substrates of the PrpC phosphatase or the PrkC kinase, we separated extracts of wild-type and mutant cells by SDS-PAGE and probed Western blots of these gels with antibodies that specifically recognize either phosphoserine or phosphothreonine . No significant differences were seen using antibody specific for phosphoserine (data not shown) . However, using antibody specific for phosphothreonine, the extract of the prpC phosphatase mutant displayed a new signal which migrated with an apparent molecular weight of 105,000 (Fig . 2A, lane 3) . Because this signal was absent in extracts from the prkC kinase mutant and the prpC-prkC double mutant, we infer that the PrkC kinase was directly or indirectly required for its appearance in vivo .

Only 56 B . subtilis proteins have calculated molecular weightsexceeding 100,000 (12; http://genolist.pasteur.fr/SubtiList/).We therefore ran SDS-6% polyacrylamide gels to separate the proteins in the region of interest and were able to associate the antiphosphothreonine signal with a single protein band (data not shown) . The band was extracted from the gel and digested with trypsin . The resulting fragments were analyzed by using MS, which identified EF-G with high confidence . A total of 46% (23 of 49) of the theoretical tryptic fragments were identified by mass, and these fragments were distributed over the entire length of the protein . Moreover, the amino acid sequence of these fragments exactly matched the corresponding fragments in EF-G . Therefore, EF-G became a good candidate as a target for the PrpC phosphatase and the PrkC kinase in vivo .

Immune precipitation using anti-E . coli EF-G yields a protein recognized by antiphosphothreonine antibody. The calculated molecular weights of EF-G and the PrkC kinase are 76,360 and 71,687, respectively (12) . However, in our hands hexahistidine-tagged versions of these two proteins both migrated on SDS-PAGE with apparent molecular weights of between 105,000 and 110,000 (see below) . This anomalous mobility was previously observed for native EF-G (4), but the aberrant mobility of PrkC was somewhat surprising . Notably, the PrkC kinase is thought to autophosphorylate on a threonine residue and is also known to be a substrate of the PrpC phosphatase (18) . Given the close comigration of EF-G and PrkC in SDS-PAGE, it remains possible that the antiphosphothreonine signal seen in vivo (Fig . 2A) was in fact PrkC itself, present as a minor component in the EF-G band .

To address this issue, we used anti-E . coli EF-G antibody to perform an immune precipitation from a cell extract of a PrpC phosphatase mutant . This yielded a protein with the expected mobility of B . subtilis EF-G (Fig . 2B, lane 1), and a parallel Western blot using antiphosphothreonine antibody revealed a signal with the same mobility (Fig . 2B, lane 3) . This signal was absent in the wild-type control (Fig . 2B, lane 4) . Because we show below that the anti-E . coli EF-G antibody preferentially precipitated EF-G over PrkC (Fig . 3B) and that the antiphosphothreonine antibody was specific for phosphothreonine (Fig . 3D), these results support the hypothesis that B . subtilis EF-G is phosphorylated on a threonine residue in a mutant lacking PrpC activity .

PrkC adds a phosphate to EF-G in vitro and PrpC removes this phosphate. To determine if EF-G was a substrate for the PrkC kinase and PrpC phosphatase in vitro, we his-tagged all three proteins and purified them on nickel affinity columns . As shown in Fig . 3A, incubation of purified PrkC with [-³²P]ATP alone yielded a labeled protein with the mobility of the PrkC standard (lane 1), whereas incubation of purified EF-G produced no labeled protein (lane 2) . However, when EF-G was incubated together with PrkC, a second labeled protein with the mobility of the EF-G standard appeared (lane 3) . Thus, PrkC is capable of phosphorylating EF-G in vitro .

We then used EF-G phosphorylated by PrkC to determine whether EF-G might be a substrate for the PrpC phosphatase . We first incubated EF-G and PrkC proteins in kinase buffer together with [-³²P]ATP, and then we purified the labeled proteins on a nickel affinity column to separate them from unincorporated [-³²P]ATP . Both labeled proteins were incubated in phosphatase buffer together with the purified PrpC phosphatase . As shown in Fig . 3A (lanes 4 to 7), the presence of PrpC progressively decreased the amount of the labeled PrkC, as expected from the work of Obuchowski et al . (18) . However, the presence of PrpC also progressively decreased the amount of labeled EF-G in the same reaction .

The experiment shown in Fig . 3A did not allow us to distinguish between direct dephosphorylation of EF-G by the PrpC phosphatase or indirect dephosphorylation via a reverse reaction involving the PrkC kinase . To address this issue, we first prepared EF-G phosphate by incubation with PrkC and [-³²P]ATP . We then purified the EF-G phosphate by immune precipitation with antibody specific for E . coli EF-G . This resulted in a preparation in which EF-G phosphate was the primary labeled component (Fig . 3B) . When this EF-G phosphate preparation was incubated with PrpC phosphatase, the ³²P label was efficiently removed in a phosphate release assay (Fig . 3C) . Although we cannot eliminate the possibility that a small amount of PrkC kinase remained in our EF-G phosphate preparation and catalyzed the reverse reaction, we consider it more likely that the PrpC phosphatase directly dephosphorylates EF-G phosphate in vitro .

To determine whether PrkC and EF-G were in fact phosphorylated on a threonine residue in vitro, we performed the Western blotting experiment shown in Fig . 3D . This experiment also served to test antibody specificity . The antiphosphothreonine antibody detected two strong bands at the mobilities expected for phosphorylated PrkC and EF-G (lane 1) . Preincubation of this antibody with phosphothreonine effectively competed both these signals (lane 2), whereas preincubation with phosphoserine had little effect (lane 3) . Because the antiphosphothreonine antibody had no significant avidity to phosphoserine, we conclude that both PrkC and EF-G are phosphorylated on one or more threonine residues .

Activity of EF-2, the EF-G homologue in eukaryotic cells, is known to be dramatically decreased by threonine phosphorylation (reviewed in reference 17) . Moreover, E . coli EF-G is known to be phosphorylated during phage T7 infection (20), and it was suggested earlier that a B . subtilis EF-G homologue becomes phosphorylated during the sporulation process (16) . It is therefore an attractive possibility that the dynamic control of the phosphorylation state of EF-G serves a regulatory role in stationary-phase B . subtilis cells, and that disruption of this control contributes to the observed phenotypes of the prpC and prkC mutants . If threonine phosphorylation of bacterial EF-G serves to decrease its activity, as is the case for eukaryotic EF-2, we can imagine that the resulting slow polypeptide extension would redirect free ribosomal subunits to mRNA species that have low initiation rates, primarily as a consequence of decreased competition with mRNA species that have high initiation rates .

PrkC bears a clear serine-threonine kinase domain resembling those found in eukaryotic kinases, and this domain is widely distributed among prokaryotes . However, true orthologs of the full-length PrkC appear to be confined to the low-GC group of gram-positive bacteria (Table 2) . This reflects the fact that PrkC has two domains—an amino-terminal kinase domain, which is widely shared, and a carboxyl-terminal domain of unknown function, which is of more restricted distribution (24; http://www.ncbi.nlm.nih.gov/COG/) . Among these low-GC gram-positive bacteria, genes encoding PrpC and PrkC orthologs are directly adjacent, as they are in B . subtilis, suggesting that they fulfill equivalent regulatory roles in these organisms .

We thank the two anonymous reviewers for their helpful comments, Young-Moo Lee of the Protein Structure Laboratory for his assistance with the mass spectrometer analysis, and Andreas Savelsbergh and Wolfgang Wintermeyer for providing polyclonal antibody against E . coli EF-G .

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