Enzymatic and Functional Analysis of a Protein Phosphatase, Pph3, from Myxococcus xanthus

Abstract

A protein phosphatase, designated Pph3, from Myxococcus xanthus showed the enzymatic characteristics of PP2C-type serine/threonine protein phosphatases, which are metal ion-dependent, okadaic acid-insensitive protein phosphatases. The pph3 mutant under starvation conditions formed immature fruiting bodies and reduced sporulation.

TEXT

Myxococcus xanthus is a Gram-negative soil bacterium that demonstrates complex social behavior (4, 9). Under starvation conditions, the vegetative cells undergo a developmental cycle involving cell-cell interactions. More than 10⁵ cells migrate to an aggregation center and form a fruiting body, within which cells differentiate into myxospores.

Living organisms sense and respond to environmental conditions using an array of signal transduction systems. Since M. xanthus demonstrates complex social behavior, signal transduction systems in this bacterium have been studied widely. Protein phosphorylation is a key mechanism by which environmental signals are transmitted to cause changes in protein expression or activity in both eukaryotes and prokaryotes. Signal transduction pathways in eukaryotes are typically mediated by various serine/threonine (Ser/Thr) and/or tyrosine (Tyr) protein kinases (21). All members of this well-defined superfamily exhibit extensive sequence similarity. The catalytic subunits of many Ser/Thr/Tyr protein kinases are highly conserved (26). These eukaryotic-like protein kinases are also found in many bacteria, especially species of Myxococcales, Actinomycetales, and Nostocales (22). In addition, these bacteria have eukaryotic-type protein phosphatases that carry out the dephosphorylation of phosphoproteins. Protein phosphatases in eukaryotes are grouped into three families (phosphoprotein phosphatases [PPP], metal ion-dependent protein phosphatases [PPM], and protein tyrosine phosphatases [PTP]) (10). The PPP family covers the PP1, PP2A, and PP2B groups, and the PPM family includes the PP2C group, which acts as a monomer (10). To date, two M. xanthus protein phosphatases, Pph1 and Pph2, which belong to the PPM family and the PPP family, respectively, have been characterized (7, 28). pph1 mutation caused defects in vegetative growth, cell motility, and aggregation and sporulation. The pph2 mutant is defective in aggregation, sporulation yield, and germination efficiency. Moreover, the addition of carbonyl cyanide m-chlorophenyl hydrazone reduces the viability of the pph2 mutant. In this study, we analyzed the enzymatic properties and function of a protein phosphatase of M. xanthus, Pph3, which belongs to the PPM family.

Sequence analysis indicates that Pph3 belongs to the PPM family.

The pph3 (MXAN_4398) gene encodes a protein of 424 amino acids with a calculated molecular mass of 45.6 kDa. Two domains could be defined for Pph3, the N-terminal domain (from residues 8 to 250), which is highly similar to the catalytic domain of PP2C-like protein phosphatases, including all 11 conserved motifs; and the C-terminal domain (from residues 296 to 405), with a cyclic nucleotide-monophosphate (cNMP)-binding domain. PP2C is one of the four major classes of mammalian serine/threonine-specific protein phosphatases, and PP2Cs have no apparent amino acid sequence homology to other types of PPP phosphatases. The amino acid sequences of the catalytic domain of Pph3 showed 87% identity to Stigmatella aurantiaca protein phosphatase, 48% identity to Sorangium cellulosum protein phosphatase, 36% identity to M. xanthus Pph1, and 20% identity to Homo sapiens Ser/Thr protein phosphatase 2Cα (Fig. 1) (18). Eight invariant residues in the conserved motifs of PP2C-like protein phosphatases also exist in Pph3 (24, 27).

Fig. 1.

Amino acid sequence alignment of homologous regions in M. xanthus Pph3 (Pph3), S. aurantiaca protein phosphatase (Sau; STAUR_4761), S. cellulosum protein phosphatase (Sce; sce6484), M. xanthus Pph1 (Pph1), and Homo sapiens PP2Cα (Hsa). Amino acid residues in agreement for more than three residues are indicated by filled boxes. Gray shading indicates degrees of similarity among amino acid residues. Closed circles mark amino acid residues that are invariant residues in the conserved motifs of PP2C-like protein phosphatases. Amino acid residues are numbered to the left of the sequence.

Expression of Pph3 in E. coli.

The pph3 gene was amplified by PCR using the primers 5′-GAAGGTAGGCATATGGGCCTGCGTAACGTGCGGAG-3′ and 5′-GACAAGCTTGAATTCCGGTCGACGAGGGGCCATGAC-3′, and the product was cloned into an expression vector, pCold or pCold-TF (Takara Bio.). The expression plasmids were transferred to E. coli BL21(DE3). The fusion protein (trigger factor [TF] [52 kDa]-Pph3) with an N-terminal hexahistidine tag was expressed only in the soluble fraction. The fusion protein produced in the soluble fraction in E. coli was purified by affinity chromatography on a Talon CellThru column (Clontech), and then the purified protein was further purified using a gel filtration column (TSKgel G3000SW; Tosoh). The purified Pph3 was analyzed by SDS-PAGE, which revealed a single band corresponding to a molecular mass of 107 kDa (data not shown).

Pph3 is an Mn²⁺-dependent phosphatase.

Purified recombinant Pph3 was used to determine phosphatase activity against phosphopeptides. The dephosphorylation activities of Pph3 toward three phosphorylated peptides are shown in Fig. 2 A. Pph3 demonstrated the highest dephosphorylation activity toward phosphothreonyl hexapeptide, RRATpVA, for PP2A, PP2B, and PP2C substrates. The protein phosphatase activity of Pph3 required Mn²⁺ as an essential cofactor (Table 1) and 2-mercaptoethanol as a reducing agent. The optimal Mn²⁺ concentration for Pph3 activity against phosphothreonine peptide was 10 to 15 mM. PPM phosphatases are Mg²⁺- or Mn²⁺-dependent protein phosphatases. M. xanthus Pph1, a member of the PPM family, was activated not only by Mn²⁺ but also by Mg²⁺ (28); however, Pph3 activity was not affected by Mg²⁺. In addition, Pph3 activity was not activated by Ca²⁺, Fe²⁺, and Zn²⁺. The effects of various metal ions on enzyme activity in the presence of 10 mM MnCl₂ were examined. The results are shown in Table 1, indicating that Fe²⁺ or Zn²⁺ strongly inhibited the protein phosphatase activity of Pph3. In the presence of 10 mM Mn²⁺, the addition of 10 mM Mg²⁺ slightly activated Pph3 activity. Pph3 was characterized by an optimum pH and optimum temperature. The optimum pH was identified by measuring at pH 5.0, 6.0, 7.0, 7.5, 8.0, 8.5, and 9.0, and the optimum temperature was found by carrying out enzymatic assays at 20, 25, 30, 35, 40, 50, and 60°C. The enzyme had an optimum pH of 8.0 for the phosphothreonine peptide with 0.2 M Tris-HCl buffer. The enzyme showed less than about 40% optimal activity below pH 7.0. The optimum temperature of Pph3 was 35°C with phosphothreonine peptide. The activity of this enzyme was lost above 50°C.

Fig. 2.

Phosphatase activity of Pph3. (A) Substrate specificity of Pph3. Phosphatase activities were determined using serine phosphopeptide, RKRSpRAE (1); tyrosine phosphopeptide, RRLIEDAEYpAARG (2); and threonine phosphopeptide, RRATpVA (3). Assays were carried out in 100 mM Tris-HCl, pH 8.0, containing 10 mM MnCl₂, 0.05% 2-mercaptoethanol, 0.1 mM substrate, and 2 μg Pph3 at 35°C for 45 min. Phosphate released was detected using a malachite green assay kit (Enzo Life Sciences). The data are the means ± standard errors of triplicate determinations. (B) Effect of inhibitors on protein phosphatase activity. Pph3 activity for threonine phosphopeptide was assayed in the absence (1) or presence (2) of 20 μM okadaic acid, 0.2 mM cyclosporine A (3), or 2 mM sodium orthovanadate (4). (C) Effect of cNMP on protein phosphatase activity. The purified fusion enzyme was cleaved with HRV 3C protease (Novagen) at 4°C for 12 h, and then protein phosphatase activity of Pph3 against threonine phosphopeptide was assayed in the absence (1) or presence (2) of 1 mM cAMP or cGMP (3).

Table 1.

Effect of metal ions on protein phosphatase activities^a

Metal	Concn (mM)	Relative activity (%) with:
		No Mn2+	10 mM Mn2+
None	0	6 ± 1	100 ± 7
MnCl2	0.1	7 ± 6	ND
	0.5	12 ± 2	ND
	2.5	38 ± 0	ND
	10.0	100 ± 5	ND
	15.0	113 ± 2	ND
MgCl2	10.0	4 ± 1	128 ± 1
CaCl2	10.0	2 ± 1	101 ± 8
FeCl2	10.0	9 ± 4	28 ± 2
ZnCl2	10.0	4 ± 0	14 ± 1

^aProtein phosphatase activities were measured in a reaction mixture containing 100 mM Tris-HCl, pH 8.0, 0.1 mM RRATpVA substrate, 0.05% 2-mercaptoethanol, enzyme, and the metal ions shown above at the indicated concentration in a total volume of 50 μl. The mixtures were incubated at 35°C for 60 min. The data represent the means ± standard errors of triplicate determinations. ND, not determined.

In contrast to phosphothreonine peptide, Pph3 hardly dephosphorylated the phosphorylated PKG substrate, RKRSpRAE (Fig. 2A). Since human PP2C has a striking preference for phosphothreonine (RRATpVA) over phosphoserine (RRASpVA) (3), Pph3 also had similar enzymatic properties with PP2C-type phosphatase. Pph3 also showed dephosphorylation activity against tyrosine-phosphorylated peptide, RRLIEDAEYpAARG, for PTP substrate. The specific activity of Pph3 against tyrosine phosphopeptide was about 5-fold lower than that against threonine phosphopeptide. In human PP2Cα, no detectable activity was observed against tyrosine-phosphorylated peptide KIGEGTYpGVVYK (5). On the other hand, a PP2C-like phosphatase, PTC1, from Saccharomyces cerevisiae had weak tyrosine phosphatase activity (17). The kinetic parameter of Pph3 was determined using RRATpVA as a substrate. The K_m value of Pph3 for RRATpVA was 164 ± 13 μM, similar to that of human recombinant PP2Cα (152 μM) (20).

Okadaic acid 20 μM, an inhibitor specific to PP1 and PP2A, and 0.2 mM cyclosporine A, an inhibitor specific to PP2B, did not inhibit Pph3 activity (Fig. 2B). M. xanthus Pph3 also showed the enzymatic characteristics of PPM because PPM phosphatases are known to be insensitive to the broad-spectrum phosphatase inhibitor okadaic acid. Protein phosphatase activity of Pph3 was inhibited by about 50%; however, this was in the presence of 2 mM sodium orthovanadate. Vanadate is a classic protein phosphatase inhibitor that mimics the transition state of phosphoryl-transfer reactions and competes with the substrate for interaction with the nucleophile (2).

Pph3 has an additional C-terminal cNMP-binding domain. The C-terminal cNMP-binding domain of Pph3 shared a high degree of identity with the conserved consensus sequence ([Q/E/K]G[D/E]xG[D/K/E]x[F/M/L][Y/F]xxxxG) (where x is any residue) of the cyclic AMP (cAMP)-binding motif. Unlike the PPP family, phosphatases that belong to the PPM family are monomeric enzymes that are not regulated through regulatory subunits. To determine whether the protein phosphatase activity of Pph3 is regulated by cAMP or cyclic GMP (cGMP), we determined Pph3 activity in the presence of 1 mM cAMP or cGMP. As shown in Fig. 2C, the phosphatase activity of Pph3 was not affected appreciably by the addition of cAMP or cGMP. The second messengers cAMP and cGMP are important regulators of numerous biological functions in bacteria (1, 19). The cellular concentrations of cAMP and cGMP changed during the development in M. xanthus (8, 30). We have reported the roles and enzymatic characterization of two receptor-type adenylyl cyclases and some cNMP phosphodiesterases in M. xanthus (11–15). Other myxobacteria also have protein phosphatase/cyclic nucleotide-binding domain proteins; for example, genes of Haliangium ochraceum, S. cellulosum, Plesiocystis pacifica, and S. aurantiaca encode 4, 3, 2, and 1 protein phosphatase/cyclic nucleotide-binding domain proteins, respectively, although the abilities of these protein phosphatases to respond to cyclic nucleotides have not yet been demonstrated directly. Further work is needed to understand the enzymatic characterization of Pph3 by using various Pph3 mutant proteins, which contain Pph3 lacking the C-terminal cNMP-binding domain, or with point mutations in the cNMP-binding domain.

Pph3 is involved in the formation of fruiting bodies and spores.

To study the biological function of Pph3, the pph3 gene was disrupted by insertion of the kanamycin resistance (Km^r) gene. Oligonucleotide primers, pph3N (5′-CCGAGAAGTCTTCGAGCGGAACCTG-3′) and pph3C (5′-GGAAGTCAGCAGCCGGGCCATGCGG-3′), were used to amplify the DNA fragment containing the pph3 gene from the M. xanthus FB (IFO 13542) genome, and then the PCR product was cloned into a pBluescript SK vector. The Km^r gene was inserted between the NruI sites of the pph3 gene, and the insertion-deletion mutation was moved into the chromosome of M. xanthus FB by electroporation (6). Using PCR and restriction enzyme analyses, we confirmed that the Km^r gene was inserted into the pph3 gene on the chromosome of the mutant (data not shown).

The timing and level of expression of the pph3 gene were determined by quantitative reverse transcriptase PCR (qRT-PCR) analysis. Total RNA was isolated from M. xanthus FB in the exponential growth phase and during development at 24 h, 48 h, and 72 h as described previously (16). cDNA was synthesized with PrimeScript II RTase (Takara Bio.), and PCR analysis was performed with Kapa SYBR Fast qPCR master mix (KAPABiosystems), a 5′ gene-specific primer (5′-AGGACGAGGAGGTCCTTCC-3′), a 3′ gene-specific primer (5′-CTGCGCTTCGCTCGACTCC-3′), and the synthesized cDNA using an ABI 7300 real-time cycler. Standard curves were made at 10³, 10⁴, 10⁵, 10⁶, and 10⁷ pCold-TF-pph3 plasmid copies/μl. qRT-PCR analysis using cycle-to-threshold values showed that the pph3 gene was expressed at different stages of development, with maximal expression during the aggregation stage (24 h), and then decreasing until fruiting body formation (48 h and 72 h). The relative cDNA quantities at 48 h and 72 h were 15% and 6%, respectively, relative to that at 24 h (defined as 100%). The pph3 gene was also expressed during exponential growth, and the relative cDNA quantity was 75%, relative to that at 24 h of development. We also confirmed that a gene (MXAN_4397) located downstream of pph3 was transcribed at similar levels in the wild type and a pph3 Km^r insertion mutant, using qRT-PCR with a primer set (5′-GCAGCTTCGTCCGGAATCC-3′ and 5′-CTCATGGCGCAGCTCAACC-3′) (data not shown), suggesting that the phenotypes of the pph3 mutant were not due to polar effects. These qRT-PCR products were not amplified without reverse transcription.

When cultured in Casitone-yeast extract (CYE) growth medium at 27°C, the wild-type strain and pph3 mutant showed similar growth rates during exponential growth; however, the final cell density (2.7 × 10⁹ cells/ml) of the pph3 mutant was about 83% of that of the wild-type strain. A similar phenotype was observed in the pph1 mutant of M. xanthus (28). In lower eukaryotes, such as S. cerevisiae and Schizosaccharomyces pombe, PP2C-deficient mutants are reported to be temperature and high-osmolarity sensitive for growth (23, 25). The growth rates of the pph3 mutant at a high temperature (37°C) and osmotic pressure (0.2 M NaCl) were similar to those of the wild-type strain (data not shown); however, the final cell densities of the pph3 mutant at a high temperature (5.6 × 10⁸ cells/ml) and osmotic pressure (1.3 × 10⁹ cells/ml) were reduced to 61% and 57%, respectively, compared with those of the wild-type strain (defined as 100%).

On clone fruiting (CF) starvation agar developmental medium, wild-type vegetative cells glided to aggregation centers where they assembled into multicellular structures called mounds at 20 to 24 h of development and formed fruiting bodies at 48 to 72 h. Within the fruiting bodies of the wild type, rod-shaped vegetative cells were converted to spherical myxospores. The pph3 mutant cells formed fruiting bodies about 24 h later than the wild-type strain; however, they formed immature fruiting bodies even after 8 days of incubation (Fig. 3 A). In the mutant strain, many spores were observed around the immature fruiting bodies. Accordingly, the viable spore yield of the pph3 mutant strain at 8 days was approximately 45% of that of the wild-type strain (Fig. 3B). The phenotype of reduced viable spore formation is also shown in M. xanthus pph1 and pph2 mutants (7, 28). In the pph3 mutant as well as in the M. xanthus pph2 mutant, many short rod-shaped spores were observed (data not shown). On the other hand, when vegetative cells were cultured with 0.5 M glycerol in CYE medium, the mutant cells sporulated at the same rate as wild-type cells.

Fig. 3.

Fruiting body and spore formations of M. xanthus wild-type and pph3 mutant strains. (A) Fruiting body development of the pph3 mutant. Cell suspensions were placed on CF agar at 28°C for 8 days. (B) Spore formation of the wild type (open circles) and pph3 mutant strains (closed circles). Cells were developed on CF agar, and undifferentiated cells were killed by sonication for 1 min and by incubation at 60°C for 15 min. Total spores (dashed lines) were counted with a hemocytometer. Viable spores (solid lines) were determined by plating out serial dilutions on Casitone-Tris (CTT) agar plates. Data are expressed as the means of triplicate experiments. Standard deviations are shown by error bars.

M. xanthus fibrils are used to attach to other cells and are required for cell-cell cohesion and social (S) motility (31). S motility is necessary for tight packing of spores within fruiting bodies (29). In agglutination buffer, wild-type cells began to agglutinate after 40 min of incubation and were precipitated within 120 min. In contrast, pph3 mutant cells agglutinated more slowly than wild-type cells and completely precipitated after 160 min of incubation (data not shown). This result indicates that the amount of produced fibrils in the pph3 mutant may be slightly less than that in the wild-type strain. M. xanthus cells move on a solid surface by gliding motility with occasional reversals of direction. In M. xanthus pph1 mutant cells, pph1 mutant cells exhibited reduced cellular reversal frequency compared with that of the wild-type strain (28): however, there was no significant difference in the reversal period between the wild-type strain (5.5 min) and the pph3 mutant strain (7.5 min).

This study and previous studies suggest that a eukaryotic protein phosphatase, Pph3, is involved in dephosphorylation reactions of phosphorylated proteins under starvation conditions, together with Pph1 and Pph2, and dephosphorylation by the reactions of eukaryotic protein phosphatases is required for the formation of fruiting bodies and spores in M. xanthus.