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1 Section of Microbiology and Center for Genetics and Development, 268 Briggs Hall, University of California, Davis, CA 95616, USA
2 Department of Biological Sciences, University of Warwick, Coventry CV4 7AL, UK
Correspondence
Mitchell Singer
mhsinger{at}ucdavis.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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Present address: Department of Civil and Environmental Engineering, Massachusetts Institute of Technology, Cambridge, MA 02139, USA.
Present address: Department of Molecular Biology, Princeton University, Princeton, NJ 08544, USA.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Shimkets, 1999; Ward & Zusman, 2000).
One way in which bacteria sense and process such environmental signals is through signal transduction pathways known as two-component systems (TCSs). These typically consist of a sensor histidine kinase and a partner effector protein called the ‘response regulator’. For most TCSs, the pathway becomes activated upon detection of a signal by the histidine kinase, which then autophosphorylates a conserved histidine residue in its C-terminal H-box domain. The signal is subsequently transduced by transfer of the phosphoryl group from the H-box histidine to a conserved aspartate residue in the receiver domain of the partner response regulator. In its phosphorylated form, the response regulator effects a variety of physiologic changes in the cell, which usually involve the activation or repression of genes to coordinate an appropriate response to the activating signal (for reviews, see Kenney, 2002; Parkinson & Kofoid, 1992; Stock et al., 2000).
The recently completed genome of M. xanthus reveals the presence of 255 known and putative TCS components, 44 % of which are orphaned or unpartnered (D. E. Whitworth, unpublished data). One of these is a putative response regulator that we have named PhoP4, for phosphate regulation protein 4. It shows homology to PhoB in Escherichia coli and PhoP in Bacillus subtilis, which in those organisms regulate the Pho regulon of phosphate-starvation-inducible genes, including genes for phosphate transport and several phosphatases (Hulett et al., 1994; Makino et al., 1989).
The regulation of phosphate pools is important for development in M. xanthus, as evidenced by the fact that starvation for inorganic phosphate induces fruiting-body formation (Manoil & Kaiser, 1980). Furthermore, phosphatase activities with pH optima at 5.2, 7.2 and 8.5, known as the magnesium-independent (MI) acid, MI neutral and MI alkaline activities, respectively, have been reported that are specifically activated in development at discrete times following fruiting-body formation (Weinberg & Zusman, 1990). The tight temporal regulation of these MI phosphatase activities suggests that they play an integral role in development; however, the number and identities of all the phosphatases contributing to these activity profiles remain to be elucidated.
Previously, the laboratory of Muñoz-Dorado identified and characterized three known Pho TCS operons named phoP1–phoR1, phoR2–phoP2 and phoR3–phoP3 (Carrero-Lerida et al., 2005; Martinez-Canamero et al., 2003; Moraleda-Muñoz et al., 2003). Deletions in any of these systems affect specific developmental phosphatase activity patterns in M. xanthus, while leaving the vegetative phosphatase activities, which are magnesium dependent, unchanged. In addition, none of these mutations appears to affect sporulation.
We report here the identification and characterization of a member of a fourth Pho TCS, PhoP4. Our data show that PhoP4 is required for (I) expression of all three known development-specific phosphatase activities, (II) wild-type-level sporulation, and (III) expression of the predicted assimilatory inorganic phosphate pstSCAB–phoU operon. In addition, our yeast two-hybrid analyses show reciprocal interactions between PhoP4 and the histidine kinase of another M. xanthus Pho TCS, PhoR2. In sum, these data indicate that PhoP4 (and the TCS system it represents) is a very important component of the M. xanthus Pho regulatory circuit, specifically with regard to activities required for development.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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. Plasmids were propagated in E. coli JM109 (Yanisch-Perron et al., 1985). An internal fragment of the phoP4 ORF was amplified by PCR using forward primer PhoP4-F43 and reverse primer PhoP4-R575, which contain engineered PstI and EcoRI restriction sites (underlined, see Table 2), respectively. The resultant amplicon was digested with the two restriction enzymes and inserted directionally into vector pBGS18 to generate plasmid pDV525. Wild-type M. xanthus strain DK1622, which does not maintain E. coli vectors, was transformed by electroporation with pDV525 using a GenePulser Xcell electroporator (Bio-Rad Laboratories), as described elsewhere (Kashefi & Hartzell, 1995). Kanamycin-resistant (KmR) transformants were selected and then screened by Southern blot analysis for the insertion of pDV525 by homologous recombination to generate the phoP4 : : pDV525 insertion strain VP965.
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). Briefly, sequences upstream and downstream of the phoP4 gene were amplified by PCR using the primers listed in Table 2
, then ligated into the multiple-cloning site of plasmid pBJ114 to form plasmid pCS66. Insertion of this plasmid at the phoP4 locus was accomplished as described above. KmR transformants were plated on 2 % galactose CTTYE agar (CTT supplemented with 0.2 % yeast extract) and KmS GalR candidates (which had lost the plasmid by homologous recombination) were screened by Southern blot analysis for the presence of the deletion allele, yielding strain VP963, in which phoP4 was deleted between bases 77 and 541. Plasmids pDV888, pDV890 and pDV892 were generated by cloning the PstSqF1/R1, PstBqF1/R1 and PhoUqF1/R1 PCR amplicons, respectively, into the pCR 2.1-TOPO vector as directed by the TOPO TA Cloning Kit protocol (Invitrogen).
Culture media and conditions.
E. coli strains were grown in Luria–Bertani broth (Sambrook et al., 1989) in a roller drum at 37 °C and 60 r.p.m. Unless otherwise stated, M. xanthus strains were grown on CTTYE media at 33 °C, as previously described (Hodgkin & Kaiser, 1977). When needed, 40 µg kanamycin monosulfate (Sigma-Aldrich) was added per millilitre of medium.
Developmental media and conditions.
Mid-exponential-phase M. xanthus cells were concentrated to a density of 5x109 cells ml–1, and spotted onto TPM agar (10 mM Tris, pH 7.6, 1 mM potassium phosphate, pH 7.8, 8 mM magnesium sulfate; Kroos et al., 1986) at 33 °C for developmental and sporulation assays. Fruiting-body formation was visually evaluated by light microscopy with a Nikon SMZ800 microscope. Myxospore preparations were performed as previously described (Pham et al., 2005).
Yeast two-hybrid assays.
The Saccharomyces cerevisiae strain used was EGY48 (Gyuris et al., 1993). Cultivation and yeast two-hybrid methods were as described previously (Whitworth & Hodgson, 2001). Briefly, DNA encoding the receiver domains and H boxes of pho genes was amplified by PCR according to a protocol described by Whitworth & Hodgson (2001), using the primers listed in Table 2. PCR products were cloned into both pEG202 and pJG4-5, generating LexA and activation domain (AD) fusions, respectively (Table 1), using EcoRI and XhoI restriction sites. Testing the suitability of all LexA fusions for use in the two-hybrid system and subsequent screening for protein–protein interactions were also as described by Whitworth & Hodgson (2001). A positive control was provided by plasmids pDEW002 and pDEW020 (Table 1), encoding a pair of fusion proteins known to interact strongly and specifically (Browning et al., 2003). Due to the semi-quantitative nature of the yeast two-hybrid assay, interactions were assessed visually on medium containing X-Gal, with the degree of indigo colouration scored in the range 1–3, as previously reported (Browning et al., 2003; Higgs et al., 2005; Whitworth & Hodgson, 2001). When tested with ONPG as the substrate, typical ‘1’, ‘2’, and ‘3’ interactions were found to express 
-galactosidase with specific activities of 2.9±0.6, 10.7±1.0 and 2862±1962 nmol (mg protein)–1 min–1, respectively. The lowest specific activities, at 0.7±0.3, corresponded to the absence of interaction, which is indicated with a – in Table 4.
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) from approximately 1010 quick-frozen M. xanthus cells. These cells were harvested either during vegetative growth at a cell density of 5x108 cells ml–1, or 48 and 60 h after the initiation of starvation on TPM agar plates.
Quantitative PCR (QPCR).
Analysis was performed as described previously (Diodati et al., 2006). Expression of the pstS, pstB and phoU genes was calculated from DNA quantities obtained for concurrent standard curve reactions. Typically, a standard curve was generated by reacting a 10-fold dilution series of plasmid DNA (pDV888 for pstS, pDV890 for pstB, and pDV892 for phoU), ranging from 1010 to 101 copies per microlitre, with the appropriate primer set and the DyNAmo HS SYBR Green QPCR Master Mix (Finnzymes).
DNA microarrays.
PCR-generated DNA microarrays containing probes to the 7235 M. xanthus ORFs of the M1 genome (Diodati et al., 2006; Jakobsen et al., 2004) were spotted onto poly-L-lysine coated glass slides by the Stanford Functional Genomics Facility (Stanford, CA). Samples from three independent biological replicates were collected as described above, and DNA microarray analyses were performed as previously described (Diodati et al., 2006). Hierarchical clustering and statistical analyses were performed using Cluster and Java TreeView software (Saldanha, 2004), and SAM (Tusher et al., 2001). All DNA microarray data have been submitted to the Gene Expression Omnibus (GEO) at the National Center for Biotechnology Information (NCBI) (http://www.ncbi.nlm.nih.gov/projects/geo/), and accession numbers are provided in the legend of Fig. 3.
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; Hirani et al., 2001; Wanner, 1996). Phosphatase assays were performed as previously described (Weinberg & Zusman, 1990), with the exception that cells used for the assay were cultured in CTT media without additional phosphate. We previously reported that CTT medium without added phosphate contains 
0.75 mM phosphate (all of it presumably derived from casitone), while fully supplemented CTT medium contains 1.75 mM phosphate (Harris et al., 1998). To perform the phosphatase assay, mid-exponential-phase cells grown on low-phosphate CTT media were concentrated to 5x109 cells ml–1, spotted in triplicate sets onto non-phosphate-supplemented CF agar plates (Hagen et al., 1978), and harvested at various time-points into TM buffer (TPM, buffer without the potassium phosphate) or 20 mM HEPES buffer. Cells were disrupted mechanically by sonication (Branson Sonifier 450, Branson Ultrasonics) with glass beads, and cell and spore breakage was confirmed by observation under light microscopy. Aliquots of cell lysates were added to appropriate buffers containing 0.01 M p-nitrophenol phosphate, incubated at 37 °C, and enzymic activity was assessed colorimetrically at A410. Protein concentration was determined by Bradford assay (Bio-Rad Laboratories). For specific activity calculations, extinction coefficients for each buffer were derived empirically.
Sequence analyses.
Secondary structure predictions were performed using the GCG suite of sequence analysis tools, version 2 (Accelrys). Amino acid sequences were aligned using the CLUSTALW method (Thompson et al., 1994).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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), which in other bacteria regulate genes needed for phosphate scavenging. PhoP4 is predicted to be 229 amino acids in length, and contains a putative receiver domain between residues 5 and 119 (the putative conserved phosphoryl-acceptor aspartate is at position 52), and an OmpR-like DNA-binding/transactivation domain between residues 132 and 223. A putative winged helix–turn–helix structure (Okamura et al., 2000), including the recognition helix between residues 194 and 206, is predicted by the method of Chou & Fasman (1978).
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). Belonging to the ABC (ATP-binding cassette) superfamily of transporter genes, the pstSCAB–phoU operon encodes a periplasmic binding protein (PstS, MXAN4788), integral membrane-channel proteins (PstC, MXAN4789, and PstA, MXAN4790), an ABC family traffic ATPase (PstB, MXAN4791), and a component hypothesized to participate in Pi incorporation into ATP (PhoU, MXAN4792). The gene order is identical to that of the homologous E. coli operon; however, in enteric bacteria, the phoB gene is not located near the pstSCAB–phoU operon (Blattner et al., 1997; Karp et al., 2002).
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); these may serve as binding sites for PhoP4, in a similar manner to E. coli PhoB binding of PHO boxes in the promoter regions of its target genes, including the pstSCAB–phoU operon (Makino et al., 1996; Wanner, 1996). However, it is important to emphasize that the consensus sequence of a PHO box varies from species to species (Fig. 2; Liu & Hulett, 1998; Makino et al., 1996; Wanner, 1996), and future direct DNA binding studies are needed to evaluate whether any of the candidate sites identified can actually be bound by purified PhoP4 protein.
phoP4 mutants are defective for expression of the predicted pstSCAB–phoU operon
DNA microarrays were used to compare the patterns of global gene expression between the 
phoP4 mutant and wild-type cells at 0, 48 and 60 h post initiation of development (Fig. 3). During vegetative growth, i.e. at 0 h, no significant differences were observed between wild-type and the phoP4 mutant cells using DNA microarrays. However, at 48 and 60 h post initiation, the expression of several genes was affected in the phoP4 mutant. The data summarized in Fig. 3 show those genes whose expression either decreased (blue) or increased (yellow) by at least twofold in the phoP4 mutant as compared to wild-type. Many of the genes identified are ‘hypothetical proteins’. However, several are genes whose expression is known or predicted to be developmentally regulated, e.g. myxobacterial haemagglutinin (Romeo et al., 1986) and polyketide synthase. Most interesting was the identification of pstS, the putative M. xanthus homologue of the E. coli inorganic phosphate transporter gene. Further analysis revealed that transcription of other genes in the predicted pstSCAB–phoU operon was reduced by roughly 50 % (Fig. 4a). These data were then confirmed and quantified by QPCR using probes to the pstB gene (Fig. 4b): whereas expression of pstB in the wild-type was more than three times higher at 48 and 60 h into development than during vegetative growth (at 0 h), in the phoP4 mutant, the late development pstB transcript levels remained essentially unchanged compared to the 0 h time-point. This pattern was very similar to those observed for QPCR analyses of pstS and phoU, the two other genes in the predicted pstSCAB–phoU operon that were examined as part of this study (data not shown).
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were not affected in either the insertion or in-frame deletion strains). Hence, it is also possible that the expression defects observed by microarray and QPCR described above could have been caused by the inadvertent mutation of unknown cis elements when the phoP4 ORF was knocked out.
PhoP4 is required for development-specific phosphatase activities under phosphate-limiting conditions
Mutations in the other M. xanthus Pho TCSs partially decrease the MI acid phosphatase activity (AcPA) and/or the MI neutral phosphatase activity (NPA), without impairing the MI alkaline phosphatase activity (AlPA) (Carrero-Lerida et al., 2005; Moraleda-Muñoz et al., 2003). In order to determine whether PhoP4 is similarly involved in their regulation, phosphatase activites over developmental time-courses were assayed for both the phoP4 mutant VP963 and the phoP4 : : pDV525 insertional disruption mutant VP965, along with the parent wild-type strain DK1622. These analyses revealed that both phoP4 mutants were nearly identically defective for all three development-specific phosphatase activites: AcPA and AlPA were down about fivefold compared to wild-type, while NPA was decreased by 10-fold (Fig. 5). Close examination of the activity profiles also indicated that differences between the wild-type and either phoP4 mutant began to become pronounced at around 44 h into development, which coincides with the formation of the first myxospores (Fig. 5). This is significant, because it has previously been demonstrated that most development-specific phosphatase activities originate within spores (Weinberg & Zusman, 1990). In contrast to their effects on the developmental phosphatase activities, the phoP4 mutants, like other M. xanthus pho mutants, caused no defects in the vegetative phosphatase activities (data not shown).
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phoR2–phoP2 cells, unlike the wild-type, fruit and sporulate when grown on media containing concentrations of 0.2 % casitone or less (Moraleda-Muñoz et al., 2003). In the case of the phoP4 mutants, this defect was not observed when either phoP4 : : pDV525 or phoP4 cells were spotted onto media containing 0.1, 0.2, 0.5 or 1 % casitone (data not shown). However, when sporulation was assessed on TPM starvation media, the data revealed that phoP4 mutants produced viable spore levels nearly two orders of magnitude lower than those of wild-type cells (Table 3).
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). Cross-communication among different Pho regulon TCSs is known in other systems (Amemura et al., 1990), and therefore interactions among the M. xanthus Pho TCS proteins were systematically tested using the yeast two-hybrid system.
The results showed strong and specific interactions among some of the Pho protein fusions (Table 4). Strong reciprocal interactions were observed between PhoP1 and PhoR1, as would be expected of true TCS partners, while PhoR1 interacted with itself, suggesting homodimerization. An expected interaction between the protein fusions of the putative TCS partners PhoP2 and PhoR2 was also observed, though this interaction was not reciprocal. The PhoP2 receiver domain was apparently able to interact with itself, which is known to be the case for some response regulators (Lewis et al., 2002; Muller-Dieckmann et al., 1999). No interactions were seen for the Pho3 proteins, except that PhoR3 interacted strongly with PhoP2, which is not surprising given the high similarity of the Pho2 and Pho3 systems (Moraleda-Muñoz et al., 2003). Some other predicted interactions for these two systems were also not observed (or were not reciprocal), suggesting a significant rate of false negatives in the assay results.
Most interestingly, the PhoP4 fusion proteins, regardless of whether assayed as bait or prey, interacted strongly with both PhoR2 fusions. Such reciprocal interactions provide solid evidence of a PhoP4–PhoR2 interactional relationship. While PhoP4 also interacted with both the PhoP1 and PhoP2 fusions, these interactions were not reciprocal, and their significance must therefore be considered more warily; this is also true for the non-reciprocal PhoP2–PhoR3 and PhoP2–PhoR2 interactions.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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The data in this study showed that the function of one Pho TCS, represented here by PhoP4, is tied closely to processes linked to phosphate regulation. Not only did phoP4 mutations decrease spore viability by nearly two orders of magnitude (Table 3), perhaps by disrupting the developmental response to overall phosphate starvation, but they also impaired the magnesium-independent phosphatase activities that are associated with myxospores (Fig. 5). Furthermore, mutating phoP4 also decreased expression of the predicted downstream pstSCAB–phoU operon (Fig. 4), which is homologous to systems responsible for inorganic phosphate assimilation (Wanner, 1996). Interestingly, the PhoP4-dependent Pst system appeared to be active very late in development (at 60 h), when most M. xanthus cells either have lysed or are developing into myxospores (Julien et al., 2000). Possibly, the scavenging of inorganic phosphate by the Pst system at this stage, combined with the co-occurring high phosphatase activities, may be important for spore formation; this possibility is further strengthened by the observed defect in spore viability for the phoP4 mutants. Moraleda-Muñoz et al. (2003) have reported that a phoR2–phoP2 phoR3–phoP3 double mutant, while having no defect in overall spore viability, nevertheless does produce a fraction of myxospores (5–10 %) that do not reshape completely: they are ellipsoid rather than being roughly round like wild-type myxospores. Similar high-resolution microscopic imaging of the few phoP4 myxospores produced may reveal whether structural defects are caused by the pho4 mutations. Any defects found will likely be more severe than the simple misshaping of a fraction of the phoP4 myxospores formed (given the observed deficit in spore viability), and may include deformities in the spore coat, intermediate coat or cortex. Indeed, visual inspection of phoP4 myxospores by phase-contrast microscopy revealed no discernible differences compared to wild-type myxospores in terms of phase brightness, shape or size.
Beyond examining the various phenotypes caused by phoP4 mutants, which are quantitatively the strongest among the M. xanthus pho mutants so far reported, the question regarding the identity of the histidine kinase partner(s) to PhoP4 has yet to be fully answered. We noted that, whereas the mutants of each M. xanthus Pho TCS displayed different complements of phenotypes (Carrero-Lerida et al., 2005; Martinez-Canamero et al., 2003; Moraleda-Muñoz et al., 2003), they all nevertheless affected certain subsets of the as-yet unidentified, developmentally active phosphatases (Weinberg & Zusman, 1990). This suggested to us that the four Pho TCSs may be functionally interlinked, rather than acting entirely separately from one another. Intriguingly, as reported above, a yeast two-hybrid analysis for interactions among the seven known Pho TCS components showed strong, reciprocal interactions between the receiver and H-box domains of PhoP4 and PhoR2, respectively (Table 4). It is possible that inputs from signals that activate the Pho2 and Pho4 systems, if received simultaneously, may lead to cross-communication between them, and amplify the expression of genes under their common control, e.g. those connected with the acid phosphatase activities.
However, besides interacting with PhoR2, we hypothesize that PhoP4 must be partnered with at least one other histidine kinase to regulate sporulation, and with the developmental alkaline and neutral phosphatase activities, since these processes are not affected in the phoR2 mutant. The recently completed genome of M. xanthus may allow for a systematic approach towards identifying this histidine kinase partner of PhoP4. Since PhoP4 is a homologue of E. coli PhoB, it is predicted that its histidine kinase partner would resemble E. coli PhoR, the PhoB partner. The M. xanthus genome has been parsed for TCS components, and they have been classified based on domain architecture, gene order and deduced sequence phylogenies (D. E. Whitworth & D. A. Hodgson, unpublished results). These analyses have revealed a grouping of 13 histidine kinases, termed class II histidine kinases, which share these features with E. coli PhoR. In the next step, yeast two-hybrid analyses similar to the ones described in this study will be used to assess whether any of these components interact with PhoP4. Coupled with expression studies to evaluate the physiological relevance of any interactions, and phenotypic analyses comparing phoP4 mutants to mutants in genes of candidate partners, these future experiments will all help decipher the complexity of the M. xanthus phosphate regulatory circuitry.
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ACKNOWLEDGEMENTS |
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Received 29 December 2005;
revised 23 February 2006;
accepted 26 February 2006.
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) and phoP4 (
) cells were induced to undergo development on CF starvation agar. Threefold replicate samples were harvested at various time-points between 0 and 72 h into development, and phosphatase activities at pH 5.2, 7.2 and 8.5 were separately monitoredby the production of p-nitrophenol (pNP). The phosphataseactivity curves for VP963 (