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J Bacteriol. Sep 2003; 185(18): 5452–5464.
PMCID: PMC193762

TodK, a Putative Histidine Protein Kinase, Regulates Timing of Fruiting Body Morphogenesis in Myxococcus xanthus

Abstract

In response to starvation, Myxococcus xanthus initiates a developmental program that results in the formation of spore-filled multicellular fruiting bodies. Fruiting body formation depends on the temporal and spatial coordination of aggregation and sporulation. These two processes are induced by the cell surface-associated C signal, with aggregation being induced after 6 h and sporulation being induced once cells have completed the aggregation process. We report the identification of TodK, a putative histidine protein kinase of two-component regulatory systems that is important for the correct timing of aggregation and sporulation. Loss of TodK function results in early aggregation and early, as well as increased levels of, sporulation. Transcription of todK decreases 10-fold in response to starvation independently of the stringent response. Loss of TodK function specifically results in increased expression of a subset of C-signal-dependent genes. Accelerated development in a todK mutant depends on the known components in the C-signal transduction pathway. TodK is not important for synthesis of the C signal. From these results we suggest that TodK is part of a signal transduction system which converges on the C-signal transduction pathway to negatively regulate aggregation, sporulation, and the expression of a subset of C-signal-dependent genes. TodK and the SdeK histidine protein kinase, which is part of a signal transduction system that converges on the C-signal transduction pathway to stimulate aggregation, sporulation, and C-signal-dependent gene expression, act in independent genetic pathways. We suggest that the signal transduction pathways defined by TodK and SdeK act in concert with the C-signal transduction pathway to control the timing of aggregation and sporulation.

Myxococcus xanthus has adopted a complex survival strategy to manage starvation, which ultimately results in the formation of spore-filled fruiting bodies. Fruiting body morphogenesis depends on the temporal and spatial coordination of aggregation and sporulation, the two morphogenetic processes underlying fruiting body formation, and sporulation is not initiated until the aggregation process is complete. In the presence of nutrients the motile, rod-shaped cells grow and divide. M. xanthus cells move by gliding (60), and if cells are present on a solid surface, they form cooperatively feeding swarms. In response to starvation of cells at a high density on a solid surface, cells initiate the developmental program that results in the formation of the spore-filled fruiting bodies (10). The first morphological manifestations of fruiting body formation are evident at 6 h with changes in cell behavior and the formation of small aggregation foci. As more cells aggregate in these foci, they increase in size and become mound shaped. Eventually a mound holds 105 densely packed cells. Inside the mounds the rod-shaped cells differentiate to spores, resulting in mature fruiting bodies. While mound formation is complete after 24 h, spore maturation is completed 72 to 120 h after the onset of starvation. Fruiting body formation is accompanied by temporal changes in gene expression in which genes are turned on or off at specific time points during development (21, 22, 34). Developmental gene expression, in turn, also drives the progression of fruiting body formation (33). Here we have investigated how the timing of aggregation and sporulation is controlled.

The starvation- and RelA-dependent accumulation of guanosine penta- and tetraphosphate [(p)ppGpp] initiates the developmental program that results in fruiting body formation (17, 56). Fruiting body formation also depends on at least five intercellular signals (A to E) (50). Mutants deficient in any of these signaling systems are deficient in aggregation and sporulation and display abnormal developmental gene expression. The B, A, D, and E signals become important for development within the first 5 h. The C signal is a cell surface-associated morphogen (29, 55) which is used repeatedly during development to induce first aggregation at 6 h and later sporulation. Tied to the induction of these processes, the C signal induces full expression of most genes turned on after 6 h (28, 35, 39).

Synthesis of the C signal depends on the csgA gene (53). The CsgA protein exists in two forms (35), the full-length 25-kDa protein, which is homologous to short-chain alcohol dehydrogenases (37), and a 17-kDa protein, which is similar in size to the C-factor protein (29). Exogenous full-length CsgA protein (37) and exogenous C-factor protein rescue development of csgA cells (29). The molecular difference between the full-length CsgA protein and the C-factor protein has yet to be elucidated. Thus, it remains to be resolved whether one of the csgA proteins acts as an enzyme to produce the C signal or whether one of the csgA proteins is the actual C signal.

C-signal transmission occurs by a contact-dependent mechanism that involves direct contact between cells (27, 29). The implementation of the C-signal-dependent responses depends on a branched signal transduction pathway (see Fig. Fig.7).7). One branch in this pathway results in increased transcription of the csgA gene (28). In addition, the RelA-dependent accumulation of (p)ppGpp stimulates csgA transcription (7). A second branch in the pathway results in the activation, conceivably by phosphorylation, of the DNA binding response regulator protein FruA (11). Downstream from phosphorylated FruA the C-signal response pathway contains an additional branch point. One branch leads to aggregation; in this branch, C signal acts as a FruA-dependent input signal to the cytoplasmic Frz chemosensory signal transduction system (57). The second branch leads to C-signal-dependent gene expression and sporulation (11, 21, 58). In addition, the histidine protein kinase SdeK is required for aggregation, sporulation, and the expression of C-signal-dependent genes (12, 46), suggesting that SdeK either is a component in the pathway or positively modulates the activity of the pathway. In the present model for the C-signal-dependent coordination of aggregation and sporulation, an ordered increase in the level of C signaling in combination with specific thresholds at which aggregation and sporulation are induced ensures the correct temporal order of these two events (28, 35, 39).

FIG. 7.
Model of the C-signal transduction pathway. The signaling event in the pathway is the interaction between C signal exposed on the surface of one cell and the hypothetical C-signal sensor on a second cell. Starvation results in a RelA- and (p)ppGpp-dependent ...

To further the understanding of the regulatory circuits that control the onset of aggregation and sporulation, we have characterized an M. xanthus mutant that displays early aggregation and early sporulation. Here, we report the identification of a putative histidine protein kinase, TodK, which is required for normal timing of these two events. A TodK mutant displays early aggregation and early sporulation, suggesting that TodK normally acts to inhibit these processes. Moreover, loss of TodK function specifically results in increased expression of a subset of C-signal-dependent genes. From these results we suggest that TodK acts together with the C signal to ensure the timely initiation of morphogenesis.

MATERIALS AND METHODS

Growth, development, measurements of β-galactosidase activity, and motility assays.

Escherichia coli strains were grown in Luria-Bertani broth in the presence of relevant antibiotics (48). M. xanthus cells were grown in CTT medium in liquid cultures or on CTT agar plates (18). Kanamycin or oxytetracycline was used for selective growth at concentrations of 40 and 10 μg/ml, respectively. Aggregation was monitored on CF agar (54) or on TPM agar as described previously (58). Briefly, cells were grown to a density of 5 × 108 cells/ml in CTT, harvested, and resuspended in TPM buffer (10 mM Tris-HCl [pH 7.6], 1 mM KH2PO4, 8 mM MgSO4) at a calculated density of 5 × 109 cells/ml. Twenty-microliter aliquots of concentrated cells were spotted on CF agar or TPM agar and incubated at 32°C. Aggregation was monitored visually using a Leica MZ8 stereomicroscope. Cells were photographed using a Sony 3CCD color video camera. Levels of sporulation were determined after development for 48, 72, 96, and 120 h on CF agar. Spore titers were determined as the number of sonication- and heat-resistant CFU (58). To measure specific activities of β-galactosidase during development, cells were induced to develop on TPM agar as described above. Cells were harvested at the indicated time points as described previously (58). Quantification of specific activities of β-galactosidase was performed as described previously (34). Strains to be tested for motility were grown in CTT to a density of 5 × 108 cells/ml, harvested, and resuspended in TPM to a calculated density of 5 × 109 cells/ml. A 5-μl cell suspension was spotted on a thin layer of 0.5 or 1.5% agar supplemented with 0.5%·CTT prepared on a sterile microscope slide. The swarming edge morphology was inspected after 24 h (20).

Bacterial strains.

M. xanthus strains used in this work are listed in Table Table1.1. SA1634 was constructed by generalized transduction with Mx8 clp2 ts3 (63) propagated on DK1622-Ω8846 to infect DK1622. SA1636, SA1638, SA1640, SA1642, SA1644, SA1646, SA1648, SA1652, SA1656, SA1658, SA1660, and SA1679 were constructed by generalized transduction with Mx8 clp2 ts3 propagated on SA1634 to infect LS269, DK1217, DK1300, DK5279, DK9007, DK11063, DK9711, DK4499, DK4293, DK5204, SA1704, and DK4300, respectively. DK9007 was constructed by generalized transduction with Mx8 clp2 ts3 propagated on DK4324 to infect DK1622. All strains constructed by generalized transduction were tested by either Southern blot analyses (48) or PCR analyses with primers that anneal to mini-Tn5(tet) or to M. xanthus DNA flanking the mini-Tn5(tet) insertion. SA1686 and SA1688 were constructed by integration of plasmids pAAR136 and pSWU19 (S. Wu, personal communication), respectively, into the Mx8 attB site after electroporation (26). Integration was verified by PCR. For pAAR136 construction, an approximately 2.2-kb HindIII-BamHI fragment from pAAR134 (see below) including the entire todK gene was cloned in pSWU19. SA1681 and SA1682 are markerless deletion strains generated by integration of pAAR138 and pAAR140, respectively, followed by excision of the wild-type gene as described previously (23). For pAAR138 construction, a wild-type todK allele was generated by PCR with chromosomal DNA from DK1622 as template. The PCR product was cloned in pUC18 (70), generating pAAR134. The region between two EagI sites in todK was excised, generating an in-frame deletion of 780 bp in the central part of todK. The ΔtodK allele was cloned in pBJ113 (23), generating pAAR138. For pAAR140 construction, pAAR132 contains dotR on a 1,368-bp SacI-XhoI fragment cloned in pBluescript II SK(−) (Stratagene). An in-frame deletion in dotR was constructed by PCR with pAAR132 as a template and primers that anneal at the very ends of dotR. The PCR product-containing vector sequences and the ends of dotR were digested with EcoRI, ligated, and transformed into TOP10 [F mcrA Δ(mrr-hsdRMS-mcrBC) [var phi]80lacZΔM15 ΔlacX74 deoR recA1 araD139 Δ(ara-leu)7679 galU galK rpsL endA1 nupG] (Invitrogen). Subsequently, a 1,009-bp KpnI-SacI fragment from pAAR134 containing the ΔdotR allele was cloned into pBJ113, generating pAAR139. pAAR139 was digested with SacI, and a SacI fragment from pAAR134, which contains the todK sequence downstream of dotR, was cloned in the correct orientation downstream of dotR, giving rise to pAAR140. SA1704 was constructed by integration of the plasmid pJM200 by homologous recombination in the csgA gene. pJM200 carries a csgA-lacZ transcriptional fusion in the plasmid pEE25. SA1704 contains two intact copies of the csgA gene. For construction of pJM200, pTK100 is a pBluescript II SK(−) derivative, which carries a 3.3-kb SalI-PstI fragment containing csgA and its promoter (35). A BamHI site was introduced immediately downstream of the stop codon in csgA with the primer 5′-GCGCGGATCCCGGCTACCAGGGCACTTC (BamHI site underlined; stop codon in csgA in boldface). Subsequently, a 3,100-bp SalI-BamHI fragment containing csgA and the entire csgA promoter was cloned upstream of a promoterless lacZ gene in the lacZ transcriptional fusion plasmid pEE25. pEE25 is a derivative of pBGS18 (62), which carries the promoterless lacZ gene from pTL25 (40; E. Ellehauge, personal communication).

TABLE 1.
M. xanthus strains

Mutagenesis.

Mini-Tn5(tet) was introduced into wild-type DK1622 on the P4 phage (B. Julien, personal communication). P4 phages were prepared as described previously (24) and introduced into DK1622 (B. Julien, personal communication). Transductants were selected on the basis of their resistance to oxytetracycline and were each transferred to a different well in a 96-well microtiter dish containing 150 μl of CTT agar and 10 μg of oxytetracycline/ml. After 5 days cells were transferred to a plate containing modified CF agar (0.1 instead of 0.015% Casitone). Transductants were allowed to grow and develop. The developmental phenotype was checked after 4 and 5 days. A total of 10,800 transductants from 400 individual infections were generated and screened for developmental defects on the modified CF agar. One hundred ten mutants that displayed developmental defects were selected. These mutants were grown in CTT and spotted on TPM agar and CF agar to verify the phenotype and on 0.5% CTT to analyze whether they displayed wild-type swarming behavior. Fifty-five mutants with developmental defects and normal motility behavior were selected for arbitrary PCR as described previously (44) (primers: Arb1 myxo, 5′ GGCCACGCGTCGACTAGTACNNNNNNNNNNGCGAGC; Arb2, 5′ GCTCTAGAGGCCACGCGTCGACTAGTAC; mini-Tn5 LEXT, 5′ GAACGTTACCATGTTAGGAGGTC; mini-Tn5 LINT, 5′ CGGAATTCCGGGAAAGGTTCCGTTCAGGACGC; mini-Tn5 REXT, 5′ AGCCCTGCAAAGTAAACTGGATG; mini-Tn5 RINT, 5′ CGGAATTCTTGCCGCCAAGGATCTGATGGCGC). Subsequently, PCR products were sequenced, and the sequences were Blast examined against the GenBank database at the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/blast) to identify mutants carrying mini-Tn5(tet) in genes not previously characterized. Of the 55 mutants 35 carried mini-Tn5(tet) in genes not previously characterized.

DNA sequencing of region around mini-Tn5(tet) Ω8846.

Mini-Tn5(tet) Ω8846 insertion was used as a selectable marker to clone todK and dotR. Because mini-Tn5(tet) does not have SacI or XhoI sites, SA1634 genomic DNA was digested with SacI and XhoI and cloned in pBluescript II SK(−), generating pAAR120, which contains M. xanthus genomic DNA extending from approximately position −7800 to +1400, and pAAR121, which contains M. xanthus genomic DNA from approximately position +600 to +2700, respectively (all coordinates are relative to the transcriptional start site of todK). pAAR122 was constructed by cloning a HincII fragment extending from approximately position −1300 to a HincII site at position 160 in mini-Tn5(tet) from pAAR120 into pBluescript II SK(−). pAAR124 was constructed by deleting a HincII fragment extending from position +600 to position 2981 in mini-Tn5(tet) from pAAR121. pAAR125 was constructed by deleting a PstI fragment extending from position +600 to position +1700 including mini-Tn5(tet) from pAAR121. To sequence the region around the mini-Tn5(tet) Ω8846 insertion, pAAR122, pAAR124, and pAAR125 were opened with a restriction enzyme with a unique restriction site and exposed to ExoSizeIII digestion (New England Biolabs). The plasmids were religated and sequenced with primers that anneal to the multiple cloning site in pBluescript II SK(−). Sequencing with specific primers closed any gaps in the sequence.

Immunoblot analysis, primer extension, and quantitative RT-PCR.

Semiquantitative immunoblotting was carried out as described previously (3, 35). Primer extension analyses were carried out essentially as described previously (39). To perform quantitative reverse transcription-PCR (RT-PCR), cells were developed on TPM agar for the indicated periods of time and harvested. RNA was extracted by the hot-phenol method (48). The RNA was DNase I treated and reextracted by phenol-chloroform extraction. The RNA was reverse transcribed using TaqMan reverse transcription reagents with the supplied hexamers according to the protocol recommended by the supplier (Applied Biosystems). cDNA was purified using the High Pure PCR product purification kit (Roche). SYBR Green PCR Master Mix was added to cDNA from the reverse transcription of 100 ng of RNA together with 500 nM concentrations of each of the two primers (primers otodK1, 5′ CTCCCGGACGCCTTCTTC, and otodK2, 5′ GGCCATGTCTGGATTCACGTA). The primers hybridize to the 5′ end of todK and give rise to a DNA fragment with a size of 100 bp. The RT-PCR was performed on an ABI PRISM 7700 sequence detection system with the standard setup. Primers were designed using PrimerExpress as recommended by the 7700 sequence detection system supplier (Applied Biosystems).

Nucleotide sequence accession number.

The sequence reported in this paper has been deposited in the GenBank database (accession no. AY231150).

RESULTS

A new genetic locus required for timely morphogenesis.

To isolate M. xanthus mutants that display abnormal aggregation, the fully motile strain DK1622, which served as the wild-type strain in this study, was exposed to mutagenesis with a mini-Tn5(tet) transposon (Materials and Methods). The DK1622-Ω8846 derivative containing the mini-Tn5(tet) Ω8846 insertion displayed early aggregation with the formation of many small fruiting bodies and increased levels of sporulation on CF starvation medium (data not shown). To ensure that the developmental phenotype in this DK1622 derivative was caused by the mini-Tn5(tet) Ω8846 insertion, the insertion was crossed back into the wild-type strain DK1622 to generate SA1634 by Mx8-dependent generalized transduction (Materials and Methods). The developmental phenotype of SA1634 was tested on CF starvation medium (Materials and Methods). Like its parent strain, SA1634 displayed early aggregation (Fig. (Fig.1).1). Translucent mounds were evident in SA1634 at 12 h of starvation whereas translucent mounds were observed in DK1622 at 24 h. After 72 h, the mounds had darkened to become mature fruiting bodies in both strains. The fruiting bodies formed by SA1634 were smaller than those formed by DK1622, and SA1634 formed approximately twice as many fruiting bodies as observed in DK1622 (Fig. (Fig.1).1). The effect of the mini-Tn5(tet) Ω8846 insertion on sporulation was assessed after starvation of DK1622 and SA1634 on CF medium (Table (Table2).2). At 48 h, both strains sporulated at the same low level. At 72 h, however, SA1634 sporulated at a level 20-fold higher than that of DK1622; after 96 h, the sporulation level was sixfold higher in SA1634 than in DK1622, and after 120 h, the level of sporulation in SA1634 was threefold higher than in DK1622. Spores were detected only inside fruiting bodies (data not shown). As the phenotype of SA1634 is similar to that of DK1622-Ω8846, these data suggest that the mini-Tn5(tet) Ω8846 insertion is responsible for the mutant phenotype.

FIG. 1.
Developmental phenotypes of todK and dotR mutants. The indicated strains were exposed to starvation on CF agar for the indicated periods of time and viewed in a Leica MZ8 stereomicroscope. The genotypes of the strains are indicated below their names. ...
TABLE 2.
Sporulation frequencies of wild-type and mutant M. xanthus strains

Cells of SA1634 appeared indistinguishable from wild-type cells during vegetative growth with respect to growth rate in CTT medium, colony morphology on CTT agar, and pigmentation (data not shown). Moreover, when tested for initiation of development at different nutrient concentrations, SA1634 was found to initiate development at the same nutrient concentration (0.25% Casitone) as DK1622 whereas SA1634 did not initiate development at higher concentrations of Casitone (data not shown). These data suggest that the mini-Tn5(tet) Ω8846 mutation does not interfere with starvation recognition.

Two genetic systems control gliding motility in M. xanthus (60). The social (S) motility system controls gliding of groups of cells, and the adventurous (A) motility system controls gliding of single cells. To test if the developmental phenotype caused by the mini-Tn5(tet) Ω8846 insertion was secondary to an effect on gliding motility, the mini-Tn5(tet) Ω8846 insertion was crossed into representative S (DK1300 sglG1) or A (DK1217 aglB1) backgrounds by Mx8-dependent generalized transduction (Materials and Methods), and the effect of the mini-Tn5(tet) Ω8846 insertion on gliding motility was assessed (Materials and Methods). Strains that carry both an A mutation and an S mutation are nonswarming and grow as small, smooth-edged colonies (19, 20). The mini-Tn5(tet) Ω8846 insertion did not generate a nonswarming (A S) phenotype when crossed into either an A or an S background (data not shown). Thus, the mini-Tn5(tet) Ω8846 insertion interferes with neither the A nor the S motility system, suggesting that the gene carrying the mini-Tn5(tet) Ω8846 insertion is not a constituent of either the A or the S motility system. Together these data provide evidence that the mini-Tn5(tet) Ω8846 insertion causes early aggregation and early, as well as increased levels of, sporulation.

Cloning of the tod locus.

The region containing the mini-Tn5(tet) Ω8846 insertion was cloned into the pBluescript II SK(−) vector based on the tetracycline resistance gene that the mini-Tn5(tet) Ω8846 carries. The DNA sequence surrounding the insertion revealed the presence of three open reading frames (ORFs). These ORFs were generally identified on the basis of a high GC content in the third positions of codons typical of GC-rich organisms (2, 51). The mini-Tn5(tet) Ω8846 insertion was located in an ORF designated todK (timing of development kinase) (Fig. (Fig.2).2). Immediately downstream from todK, an ORF designated dotR (downstream of todK response regulator), which is transcribed in a direction opposite that of todK, was identified. The stop codon in todK overlaps partially with the stop codon in dotR (Fig. (Fig.2).2). Upstream from todK the kefC gene, which encodes a protein with homology to K+-efflux pumps, was identified (GenBank accession no. U37008). kefC is transcribed in the same direction as is todK. In a region covering approximately 700 bp upstream from dotR no ORFs were identified (Fig. (Fig.22).

FIG. 2.
Physical map of the todK-dotR region. ORFs are indicated by open arrows. Arrows indicate the directions of transcription. Coordinates are relative to +1, the transcriptional start site of todK. Below the physical map, the overlap between the ...

The deduced todK gene product, TodK, contains 501 amino acids. Mini-Tn5(tet) Ω8846 is inserted after codon 271 in the todK reading frame (Fig. (Fig.2).2). Sequence analyses indicate that the C-terminal part of TodK (amino acids 260 to 501) contains a histidine protein kinase domain (Fig. (Fig.3A).3A). This part of TodK is 48, 41, and 26% identical to the kinase domains of AsgD of M. xanthus (4), SdeK of M. xanthus (12), and ResE of Bacillus subtilis (59), respectively (Fig. (Fig.3B).3B). Histidine protein kinases function as sensors in two-component signal transduction systems (45). The kinase domain of TodK contains a conserved H box, which contains the potential autophosphorylation site H275. The kinase domain in TodK also contains the highly conserved N, D/F, and G boxes found in histidine protein kinases (Fig. (Fig.3A3A and B) (45). The deduced amino acid sequence in the N-terminal part of TodK yielded no significant overall similarities to protein sequences in the databases in a BLAST search (1). This lack of conservation is consistent with the proposed function of the N-terminal part as sensor domains that change the activity of the protein in response to a specific signal. More detailed analyses of the N-terminal part of TodK with a SMART procedure (49) revealed that this region contains two PAS domains (Fig. (Fig.3A)3A) (64). Further analysis with the program TMHMM2 (31) of the deduced amino acid sequence of TodK revealed no potential membrane-spanning region, suggesting that TodK may localize to the cytoplasmic compartment.

FIG. 3.
Sequence analyses of TodK and DotR. (A) Domain organization of TodK. In the sensor domain the two PAS domains are indicated; in the kinase domain, the conserved H, N, D/F, and G boxes are indicated. The asterisk over the H box indicates the conserved ...

The deduced dotR gene product, DotR, contains 135 amino acids. Sequence analysis revealed that DotR is 40, 40, and 32% identical to the receiver domains of the response regulators PhoP of B. subtilis (38), PhoB of Caulobacter crescentus (13), and PilG of Pseudomonas aeruginosa (9), respectively (Fig. (Fig.3C).3C). Response regulators function as effectors in two-component signal transduction systems and typically consist of a conserved receiver domain and an output domain (45). DotR belongs to the group of response regulators which consist only of a receiver domain. The activity of response regulators is regulated by phosphorylation of a conserved aspartate residue in the receiver domain. The phosphoryl group is transferred to the aspartate from the cognate histidine protein kinase. The putative phosphorylation site D67 is conserved in DotR. In addition to the phosphorylated aspartate residue, DotR contains all conserved amino acid residues normally found in receiver domains including D23, D24, S97, and K119 (Fig. (Fig.3C)3C) (67).

Identification of the gene in the tod locus required for timely aggregation and sporulation.

To determine whether the defects caused by the mini-Tn5(tet) Ω8846 insertion were due to an inactivation of the todK gene, we constructed a strain that carries an in-frame deletion mutation in todK by a markerless deletion method (Materials and Methods). SA1681 contains a 780-bp in-frame deletion in todKtodK) corresponding to codons 159 to 418 in TodK. This deletion removes from TodK the H box containing the H275 residue, which is predicted to be the site of autophosphorylation, and the conserved N and D/F boxes. To examine the developmental defects caused by the ΔtodK mutation, SA1681 was exposed to starvation on CF starvation medium in parallel with the wild-type strain DK1622 and SA1634, which carries mini-Tn5(tet) Ω8846. SA1681 displayed a phenotype similar to that of SA1634, which is characterized by early aggregation, formation of approximately twice as many fruiting bodies as observed in DK1622, and smaller fruiting bodies than those in DK1622 (Fig. (Fig.1).1). Likewise, SA1681 displayed early sporulation and reached a final sporulation level approximately threefold higher than in DK1622 at 120 h (Table (Table2).2). Thus, the developmental defects caused by the ΔtodK mutation are similar to that caused by the mini-Tn5(tet) Ω8846 insertion.

To demonstrate that the loss of todK function was responsible for the developmental defects in SA1634, a genetic complementation experiment was performed. The plasmid pAAR136 carries the wild-type todK gene including 566 bp upstream from the putative start codon (Fig. (Fig.2).2). In addition, pAAR136 carries the Mx8 attP site for integration at the chromosomal Mx8 phage attachment site attB. pAAR136 was introduced into SA1634 by electroporation (Materials and Methods) to give the strain SA1688. As a control the vector pSWU19 used to generate pAAR136 was also introduced into SA1634, giving rise to the strain SA1686. SA1688 and SA1686 cells were assayed for development on CF starvation medium in parallel with cells of DK1622 and SA1634. As shown in Fig. Fig.11 and Table Table2,2, the developmental defects caused by the mini-Tn5(tet) Ω8846 insertion were corrected by pAAR136 whereas pSWU19 did not correct the developmental defects in SA1634. The observation that the accelerated development caused by the mini-Tn5(tet) Ω8846 insertion can be corrected by the introduction of a wild-type copy of the todK gene, together with the observation that the ΔtodK mutation gives rise to the same phenotype as does the mini-Tn5(tet) Ω8846 insertion in todK, provides evidence that the developmental defects in SA1634 and SA1681 are due to an inactivation of todK, thus leading to a loss of TodK activity.

Frequently the structural genes for the histidine protein kinase and response regulator in a two-component regulatory system are located adjacent to each other. To analyze whether the DotR response regulator encoded by the dotR gene acts in the same genetic pathway as todK, a strain that carries an in-frame deletion mutation in dotR was constructed (Materials and Methods). SA1682 contains a 375-bp in-frame deletion in dotRdotR) corresponding to codons 6 to 129 in the 135-amino-acid DotR protein (Fig. (Fig.2).2). Cells of SA1682 were indistinguishable from wild-type cells during vegetative growth with respect to growth rate in CTT medium, colony morphology on CTT agar, and pigmentation (data not shown). Moreover, when tested for initiation of development at different nutrient concentrations, SA1682 was found to initiate development at the same nutrient concentration (0.25% Casitone) as that for DK1622 whereas SA1682 did not initiate development at higher concentrations of Casitone (data not shown). These data suggest that DotR is not involved in starvation recognition. To examine the possible developmental defects caused by the ΔdotR mutation, SA1682 was exposed to starvation on CF starvation medium in parallel with the wild-type strain DK1622. SA1682 displayed a developmental phenotype indistinguishable from that of the wild-type strain DK1622 (Fig. (Fig.11 and Table Table2).2). Likewise, when SA1682 was developed on other starvation media such as TPM medium or in submerged culture, the phenotype of SA1682 was indistinguishable from that of DK1622 (data not shown). The observation that the ΔdotR mutation does not cause developmental defects indicates that TodK and DotR may not act in the same genetic pathway.

Expression of the todK gene during development.

To study the expression of todK, primer extension experiments were carried out (Materials and Methods). Total RNA was isolated from exponentially growing vegetative wild-type DK1622 cells and from cells that had been starved on TPM starvation medium for 3, 6, and 12 h. The 5′ end of the todK mRNA maps to an adenine nucleotide located 25 bp upstream of the putative start codon of todK in vegetative cells as well as in developing cells (Fig. (Fig.4A4A and B). The 5′ end of the todK mRNA was detected at high levels in vegetative wild-type DK1622 cells. At 3 and 6 h of starvation, lower levels of the 5′ end of the todK mRNA were detected, and after 12 h the 5′ end of the todK mRNA was undetectable. To analyze if TodK is involved in the expression of todK, total RNA was isolated from vegetative and developing SA1634 cells, which contain the mini-Tn5(tet) Ω8846 insertion, and primer extension experiments were carried out. The detection profile of the 5′ end of the todK mRNA in SA1634 was indistinguishable from that observed in DK1622 (Fig. (Fig.4A4A).

FIG. 4.
Regulation of transcription of todK. (A) Mapping of the 5′ end of the todK transcript by primer extension analysis. C, T, A, and G show the sequence ladders. Total RNA was isolated from DK1622 and SA1634 that had been starved for the indicated ...

To quantify the decrease in todK mRNA levels during development, quantitative RT-PCR analyses were performed on total RNA isolated from vegetative and starving DK1622 and SA1634 cells (Materials and Methods). As shown in Fig. Fig.4C,4C, the relative levels of todK mRNA in DK1622 and SA1634 were similar both in vegetative cells and during development. The relative level of todK mRNA decreased twofold between 0 and 3 h of starvation in both strains. From 3 to 12 h of starvation, an additional fivefold decrease in the relative level of todK mRNA was detected in both strains. These data show that the amount of todK mRNA decreases 10-fold in response to starvation and that todK expression is not autoregulated.

In response to starvation, wild-type cells initiate a stringent response which is both required and sufficient for initiation of early events in fruiting body formation (17, 56). To test if the decrease in todK mRNA levels during starvation depends on the stringent response, we analyzed accumulation of the todK mRNA in strain MS1000, which carries an in-frame deletion in the relA gene. MS1000 does not accumulate (p)ppGpp in response to starvation, fails to mount the stringent response during starvation, and consequently does not form fruiting bodies (M. Singer, personal communication). MS1000 is a derivative of the DK101 strain which carries a mutation in the pilQ gene, which is involved in synthesis of type IV pili (68). To analyze todK expression in MS1000, total RNA was isolated from vegetative cells of MS1000 and DK101 as well as from cells starved on TPM medium. The relative levels of todK mRNA in the two strains were determined in primer extension experiments. The accumulation profile of the todK mRNA in DK101 was identical to that observed in DK1622. Importantly, the relative levels of todK mRNA in MS1000 and DK101 were indistinguishable in vegetative as well as in starving cells (data not shown). Thus, the stringent response is not required to down-regulate todK mRNA levels during development.

Effect of todK on developmental gene expression.

To further define the role played by TodK in M. xanthus development, the expression of eight developmentally regulated lacZ reporter fusions was studied in the mini-Tn5(tet) Ω8846 mutant background. In wild-type cells each reporter fusion increases its expression at a particular time point during starvation. The eight fusions have different expression times ranging from 0 to 24 h, and their expression depends in different combinations on (p)ppGpp and intercellular signals (Table (Table3).3). Thus, analysis of the expression of these reporter fusions in a todK mutant allows us to monitor the progression of the developmental program and to determine the time of action of TodK. To analyze the effect of the mini-Tn5(tet) Ω8846 insertion on developmental gene expression, the mutation was transduced into the eight strains each containing a lac reporter fusion (Materials and Methods). Subsequently, isogenic TodK+ and TodK strains each carrying one of the lacZ fusions were exposed to starvation on TPM medium and specific activities of β-galactosidase expressed from the fusions were determined (Materials and Methods) (Fig. (Fig.5A).5A). The expression profiles of Tn5lac Ω4408, Tn5lac Ω4521, Tn5lac Ω7540, and the csgA-lacZ fusion (pJM200) were indistinguishable between the two strains. The four fusions Tn5lac Ω4499, Tn5lac Ω4414, Tn5lac Ω4435, and Tn5lac Ω4401 had altered expression profiles in the todK strain. Tn5lac Ω4499, Tn5lac Ω4414, and Tn5lac Ω4435 displayed the same levels of expression of β-galactosidase in the two genetic backgrounds until 12 h. Also, the expression times of these three fusions were identical in the two genetic backgrounds. After 12 h the level of β-galactosidase expressed from Tn5lac Ω4499, Tn5lac Ω4414, and Tn5lac Ω4435 was increased in the todK background compared to the todK+ background. Tn5lac Ω4401 displayed the same level of expression of β-galactosidase in the two genetic backgrounds until 24 h. Also, the expression time of this fusion was identical in the two genetic backgrounds. After 24 h the level of β-galactosidase expressed from Tn5lac Ω4401 was increased in the todK background compared to the todK+ background. For the four fusions Tn5lac Ω4499, Tn5lac Ω4414, Tn5lac Ω4435, and Tn5lac Ω4401, the expression level after 36 h of starvation was approximately twofold higher in the todK background than in the todK+ background. These four lac reporter fusions all depend on the C signal for full levels of expression.

FIG. 5.
Effect of todK mutation on developmental gene expression. (A) Expression of the indicated lacZ fusions in wild-type DK1622 cells and in todK SA1634 cells. Cells were starved on TPM agar, samples were withdrawn at the indicated time points, and specific ...
TABLE 3.
lacZ reporter fusions used to analyze the function of TodK

Full expression of the csgA gene depends on the C signal (28). To verify that the expression of csgA is not affected by a todK mutation (Fig. (Fig.5A),5A), semiquantitative immunoblot analyses with polyclonal anti-CsgA antibodies were carried out on total protein isolated from wild-type cells (DK1622) and todK cells (SA1634) that had been starved on TPM medium (Materials and Methods). The two CsgA proteins displayed similar accumulation profiles in the two strains during starvation (Fig. (Fig.5B).5B). These data support the notion that TodK is not involved in csgA expression. Moreover, they provide evidence that TodK is not involved in synthesis or degradation of the two CsgA proteins. The differential effects of the todK mutation on csgA expression compared with other C-signal-dependent genes suggest that TodK functions only in the regulation of the expression of a subset of C-signal-dependent genes. From the increased levels of expression of certain C-signal-dependent genes in a todK background, it appears that TodK would normally have an inhibitory effect on the expression of these C-signal-dependent genes.

Effect of todK mutation on the accumulation of FruA and FrzCD.

To analyze whether a todK mutation interferes with accumulation of known components in the C-signal transduction pathway, we analyzed the accumulation of the FruA protein and the FrzCD protein. Semiquantitative immunoblot analyses with polyclonal anti-FruA antibodies and polyclonal anti-FrzCD antibodies were carried out on total protein isolated from starving wild-type cells (DK1622) and todK cells (SA1634) (Materials and Methods). Each of the two proteins displayed similar accumulation profiles in the two genetic backgrounds during starvation (data not shown). The finding that FruA accumulates at the same levels in the two strains is consistent with the finding that a todK mutation does not have a significant effect on the expression of the fruA-lacZ fusion in Tn5lac Ω7540 (Fig. (Fig.5A).5A). The FrzCD protein shows modulated methylation during development (41), with the C signal inducing methylation of FrzCD after 6 h of starvation (57). To determine if TodK acts upstream or downstream of the events that lead to FrzCD methylation, the FrzCD methylation pattern was analyzed in wild-type cells (DK1622) and todK cells (SA1634) during starvation (Materials and Methods). The todK mutation in SA1634 did not interfere with FrzCD methylation during development (data not shown).

A todK mutation does not bypass known components in the C-signal transduction pathway.

To analyze whether the early aggregation, early sporulation, and increased expression of a subset of the C-signal-dependent genes caused by a todK mutation bypass the requirement for the known components in the C-signal transduction pathway, the mini-Tn5(tet) Ω8846 insertion was introduced by Mx8-dependent generalized transduction (Materials and Methods) into the following strains: LS269, which carries the csgA::Tn5lac Ω269 mutation; DK11063, which carries the fruA::Tn5lac Ω7540 mutation; DK9711, which carries the frzE::Tn5 Ω226 mutation; and DK5279, which carries the devR::Tn5lac Ω4414 mutation. Subsequently, the developmental phenotype of the double mutants was investigated. In all cases the double mutants displayed the developmental defects characteristic of the strains carrying the csgA, fruA, frzE, or devR single mutation (data not shown). These observations demonstrate that the todK mutation does not bypass the known components in the C-signal transduction pathway.

TodK acts independently of the SdeK histidine protein kinase.

Loss of function of the cytoplasmic SdeK histidine protein kinase has effects on C-signal-dependent activities which are opposite to those of a loss-of-function mutation in todK, i.e., an sdeK mutation results in delayed aggregation, reduced sporulation, and decreased expression of C-signal-dependent genes (12, 46). Moreover, an sdeK mutation—like a todK mutation—does not interfere with the expression of csgA, accumulation of the two CsgA proteins, fruA expression, accumulation of FruA, accumulation of FrzCD, or FrzCD methylation (data not shown). To analyze whether todK and sdeK act in the same genetic pathway to regulate C-signal-dependent activities, genetic epistasis experiments were performed in which the mini-Tn5(tet) Ω8846 insertion was first transduced into strain DK4300, which carries Tn5lac Ω4408 inserted in sdeK, to generate strain SA1679. Subsequently, the developmental phenotypes of the three strains SA1634 [todK::mini-Tn5(tet) Ω8846], DK4300 (sdeK::Tn5lac Ω4408), and SA1679 [todK::mini-Tn5(tet) Ω8846 sdeK::Tn5lac Ω4408] were analyzed on CF starvation medium. As shown in Fig. Fig.6,6, DK4300 displayed delayed aggregation with the formation of few and abnormally shaped fruiting bodies. SA1634 displayed accelerated aggregation with the formation of many small fruiting bodies. Importantly, the double mutant displayed an aggregation phenotype which was different from those of the two strains each carrying a single mutation. After 120 h, the sporulation frequency of the sdeK todK double mutant and the sdeK single mutant was <0.0006%. Taken together, these results suggest that TodK and SdeK act in independent genetic pathways.

FIG. 6.
Epistasis analysis of the relationship between todK and sdeK. The indicated strains were exposed to starvation on CF agar for the indicated time points and viewed in a Leica MZ8 stereomicroscope. The genotypes of the strains are indicated below their ...

DISCUSSION

Aggregation and sporulation are the two major morphogenetic events underlying fruiting body formation in M. xanthus. Aggregation and sporulation are induced by the C signal and are not initiated until 6 and 24 h, respectively, after the onset of starvation. This suggests that M. xanthus has regulatory mechanisms to ensure that these two morphogenetic events are prevented from initiating earlier. Here we report the identification of a gene, todK, which is important for the correct timing of aggregation and sporulation. Loss of TodK function results in early aggregation and early, as well as increased levels of, sporulation. These observations provide evidence that the TodK protein normally acts as an inhibitor of development.

The C-terminal domain of TodK has similarity to the transmitter domain of histidine protein kinases of two-component regulatory systems and contains the conserved sequence motifs normally present in transmitter domains including the conserved histidine residue (H275) which is the site of autophosphorylation in other transmitter domains. The N-terminal 240 amino acids of TodK constitute the sensor domain. This domain does not appear to contain membrane-spanning regions, suggesting that TodK may localize to the cytoplasmic compartment.

Generally, histidine protein kinases are activated in response to an extracellular or an intracellular signal. Activation results in autophosphorylation of the conserved histidine residue; subsequently, the phosphate is transferred to a cognate response regulator, thus resulting in the activation of the response regulator. Frequently partner proteins in two-component regulatory systems are encoded by adjacent structural genes. The dotR gene, which is located downstream from todK and transcribed in a direction opposite that of todK, has similarity to single domain response regulators of two-component regulatory systems. An in-frame deletion of dotR resulted neither in a vegetative phenotype nor in a developmental phenotype. The observation that loss of dotR function does not cause developmental defects provides genetic evidence that TodK and DotR may not constitute a simple two-component regulatory system in which TodK is the cognate kinase of DotR. Alternatively, DotR could be only one of several response regulators which are phosphorylated by TodK. In this model, the function of DotR may be redundant, thus explaining why loss of DotR function does not result in developmental defects.

The todK gene is expressed in vegetative cells, and accumulation of the todK mRNA decreases approximately 10-fold in response to starvation. The DNA sequence upstream from the transcriptional start site for todK has similarity to σA-dependent promoters (Fig. (Fig.4B)4B) (16), suggesting that the todK promoter is recognized by RNA polymerase containing σA. Moreover, accumulation of the todK mRNA is independent of the TodK protein in vegetative as well as in starving cells, suggesting that TodK is not involved in the expression of todK. The RelA-dependent accumulation of (p)ppGpp, in response to starvation, is thought to be the signal that initiates fruiting body formation including the production of the early-acting intercellular A signal (17, 56). Interestingly, the decrease in todK mRNA accumulation during development does not depend on RelA, suggesting that the decrease is independent of the stringent response and the A signal. The observation that todK mRNA accumulation decreases in a RelA-independent manner during starvation suggests that the starvation-dependent signal that results in the decrease in todK mRNA accumulation is neither (p)ppGpp nor the A signal. It remains to be analyzed whether one or more of the B, C, D, and E signals are involved in the regulation of todK mRNA accumulation.

Assuming that the intracellular concentration of TodK protein follows the detection profile of todK mRNA, then TodK is present in vegetative cells and the cellular concentration of TodK decreases during development as expected for an inhibitor of development. Even though TodK is predicted to be present in vegetative cells, loss of TodK function is not evident until 12 h of starvation. The absence of vegetative defects and the relatively late developmental defects may suggest that the signal transduction pathway defined by TodK regulates developmental events initiated after 12 h.

Loss of TodK function does not interfere with starvation recognition. Moreover, loss of TodK function does not interfere with the expression of genes that are induced by the stringent response, the A signal, or the E signal. However, loss of TodK function specifically results in increased expression of a subset of C-signal-dependent genes after 12 h of starvation. Epistasis analyses demonstrated that TodK depends on the known components in the C-signal transduction pathway to induce early aggregation and early sporulation. These data are consistent with the notion that loss of TodK function does not interfere with the stringent response, A signaling, or E signaling. Rather these data suggest that the TodK signal transduction pathway converges on the C-signal transduction pathway to negatively regulate timing of aggregation and sporulation and the expression of a subset of C-signal-dependent genes.

csgA has been implicated in the induction of the stringent response (7). In addition, the stringent response induces csgA transcription (6). The observation that loss of TodK function does not alter transcription of sdeK, csgA, and the Tn5lac Ω4521 fusion, all of which depend on the stringent response for full expression (Table (Table3),3), suggests that the TodK pathway converges on the C-signal transduction pathway downstream of the interaction between this pathway and the stringent response (Fig. (Fig.7).7). Overproduction of the C signal early during development results in a phenotype which is similar to that of a todK mutant (35). However, loss of TodK function does not interfere with the accumulation of the CsgA proteins, thus demonstrating that TodK is not involved in the regulation of C-signal accumulation. The observation that TodK does not interfere with csgA transcription argues that TodK does not act to inhibit the reception of the C signal because reception of the C signal is required for full expression of the csgA gene (28). Likewise, TodK is not involved in the developmentally regulated transcription of fruA or in the accumulation of FruA protein. Taken together, these observations suggest that the TodK pathway converges on the C-signal transduction pathway downstream of the accumulation of the FruA protein (Fig. (Fig.7).7). It was suggested previously that FruA is activated by phosphorylation by a cognate histidine protein kinase, which is activated in a C-signal-dependent manner (11). Downstream from phosphorylated FruA the pathway contains a branch point with one branch leading to aggregation and the other branch leading to C-signal-dependent gene expression and sporulation. The phenotype of a todK mutant is consistent with the idea that TodK may act to inhibit FruA phosphorylation or any other—yet to be identified—component which is common to both branches. In this case, loss of TodK function is predicted to result directly in early aggregation, increased C-signal-dependent gene expression, and, consequently, early and increased sporulation. Alternatively, TodK may act to inhibit the aggregation branch. In this scenario, loss of TodK function is predicted to result in early aggregation, which would then indirectly result in increased C-signal-dependent gene expression and, consequently, early and increased sporulation. Both models would explain early aggregation in a todK mutant. Moreover, in both models the increased expression of C-signal-dependent genes in the sporulation branch would explain early and increased sporulation. We speculate that loss of TodK function does not result in early expression of C-signal-dependent genes because other factors, which are not regulated by TodK, are required for the expression of these genes. From our present understanding of TodK function we suggest that the TodK signal transduction pathway converges on the C-signal transduction pathway downstream from the accumulation of FruA. However, the site of action and the mechanism of TodK remain to be clarified.

The TodK protein does not appear to contain membrane-spanning segments, suggesting that TodK activity is modulated either by a membrane-bound component, which senses an extracellular parameter, or by an intracellular parameter. A clue to the input signal that regulates the activity of the TodK kinase may come from the finding that the sensor domain in TodK contains two PAS domains. PAS domains have been implicated in sensing changes in redox potential, oxygen, light, small ligands, and overall energy level of a cell and in mediating protein-protein interactions (64). In this context it is of interest that the flow of carbon through metabolic pathways in M. xanthus cells changes in response to starvation (69). It therefore remains a possibility that TodK is involved in sensing a cellular parameter related to the energy status of a cell.

The cytoplasmic histidine protein kinase SdeK also contains a PAS domain in the sensor domain (64). An sdeK mutation results in developmental defects opposite to that of a todK mutation, i.e., delayed aggregation, reduced sporulation, and decreased expression of C-signal-dependent genes (46). Moreover, as is the case for TodK, SdeK is not required for accumulation of the csgA proteins, FruA, the Frz proteins, or FrzCD methylation. Therefore, the site at which the SdeK signal transduction pathway converges on the C-signal transduction pathway is also predicted to be downstream of FruA accumulation. Epistasis analyses provided genetic evidence that SdeK and TodK act in independent genetic pathways. PAS domains vary in the parameter that is sensed (64). We propose that TodK and SdeK each monitor a different cellular signal related to the energy status of a cell during starvation. Both signals are predicted to be indicators of continued starvation. Once the continued starvation of cells results in the accumulation of these signals, this would subsequently trigger an alteration in the activity of the TodK and SdeK kinases resulting in the alleviation of the inhibitory effect of TodK and stimulating the activating effect of SdeK on development. In this model, the SdeK and TodK signal transduction pathways converge on the C-signal transduction pathway, and thus the C-signal transduction pathway may be regarded as an integration point at which the intercellular signals needed to coordinate the efforts of thousands of cells during fruiting body formation and the cellular signals reflecting the energy status of individual cells are integrated. This integration would ensure that morphogenesis is not initiated aberrantly but only when the conditions for intercellular signaling between cells as well as the energy status of individual cells are appropriate (Fig. (Fig.77).

Several M. xanthus mutants have been isolated that display early aggregation and/or sporulation (5, 7, 8, 15, 30, 42, 66). The isolation of these mutants together with the isolation of many mutants in which fruiting body formation is deficient suggests that fruiting body morphogenesis is subject to both extensive negative and positive control mechanisms. These control mechanisms may serve to coordinate aggregation and sporulation temporally and spatially. During fruiting body formation a large fraction of cells undergo lysis (47). Therefore, negative control mechanisms may also help to ensure that the decision to initiate fruiting body formation is only taken as the last resort when other survival options have proven insufficient.

Acknowledgments

We thank Mitch Singer for the gift of MS1000, Bryan Julien for invaluable help with the mutagenesis, and Birgitte H. Kallipolitis and Eva Ellehauge for carefully reading the manuscript.

The Danish Natural Science Research Council, the FREJA program, and the Carlsberg Foundation supported this work.

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