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Journal of Bacteriology, September 2005, p. 5877-5884, Vol. 187, No. 17
0021-9193/05/$08.00+0     doi:10.1128/JB.187.17.5877-5884.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

The Protein Phosphatases of Synechocystis sp. Strain PCC 6803: Open Reading Frames sll1033 and sll1387 Encode Enzymes That Exhibit both Protein-Serine and Protein-Tyrosine Phosphatase Activity In Vitro

Renhui Li,{dagger} M. Ben Potters, Liang Shi,{ddagger} and Peter J. Kennelly*

Department of Biochemistry and Virginia Institute for Genomics, Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061

Received 20 December 2004/ Accepted 30 May 2005


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ABSTRACT
 
The open reading frames (ORFs) encoding two potential protein-serine/threonine phosphatases from the cyanobacterium Synechocystis sp. strain PCC 6803 were cloned and their protein products expressed in Escherichia coli cells. The product of ORF sll1033, SynPPM3, is a homologue of the PPM family of protein-serine/threonine phosphatases found in all eukaryotes as well as many members of the Bacteria. Surprisingly, the recombinant protein phosphatase dephosphorylated phosphotyrosine- as well as phosphoserine-containing proteins in vitro. While kinetic analyses indicate that the enzyme was more efficient at dephosphorylating the latter, replacement of Asp608 by asparagine enhanced activity toward a phosphotyrosine-containing protein fourfold. The product of ORF sll1387, SynPPP1, is the sole homolog of the PPP family of protein phosphatases encoded by the genome of Synechocystis sp. strain PCC 6803. Like many other bacterial PPPs, the enzyme dephosphorylated phosphoserine- and phosphotyrosine-containing proteins with comparable efficiencies. However, while previously described PPPs from prokaryotic organisms required the addition of exogenous metal ion cofactors, such as Mg2+ or Mn2+, for activity, recombinantly produced SynPPP1 displayed near-maximal activity in the absence of added metals. Inductively coupled plasma mass spectrometry indicated that recombinant SynPPP1 contained significant quantities, 0.32 to 0.44 mol/mole total, of Mg and Mn. In this respect, the cyanobacterial enzyme resembled eukaryotic members of the PPP family, which are metalloproteins. mRNA encoding SynPPP1 or SynPPM3 could be detected in cells grown under many, but not all, environmental conditions.


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INTRODUCTION
 
Examination of the current library of genome sequences indicates that many bacteria possess open reading frames (ORFs) encoding known or potential protein-serine/threonine/tyrosine kinases and phosphatases (30, 46, 49). However, our current body of knowledge concerning the functional properties and physiologic roles of these enzymes remains disappointingly fragmentary (23). The genome of the cyanobacterium Synechocystis sp. strain PCC 6803 (22), for example, encodes 15 potential protein-serine/threonine/tyrosine kinases and 12 potential protein-serine/threonine/tyrosine phosphatases (46, 49). However, the catalytic capabilities of only nine of these deduced intermediaries of signal transduction, one-third of the total, have been experimentally assessed to date. Intriguingly, while all six of the predicted protein kinases characterized thus far displayed the ability to phosphorylate themselves and/or exogenous proteins in vitro (19-21, 26, 45), only two of the three potential protein phosphatases behaved as initially predicted. Specifically, while the PPM family protein phosphatases PphA (16, 41) and IcfG (SynPPM1) (45) exhibited robust protein phosphatase activity, the product of ORF slr0946, initially annotated as a low-molecular-weight protein-tyrosine phosphatase, was discovered to be an arsenate reductase (33).

The predominant source of protein-serine/threonine phosphatase activity in eukaryotic organisms resides in the members of two genetic superfamilies designated PPM and PPP. The M in PPM refers to the universal requirement of these enzymes for the presence of a divalent metal ion cofactor, usually Mg2+, for activity. The family includes pyruvate dehydrogenase phosphatase and the many forms of protein phosphatase 2C in eukaryotes, numerous bacterial representatives, such as the protein phosphatases that modulate sporulation and stress responses in Bacillus subtilis (4, 24), as well as a single deduced archaeal protein (23). The PPP family of protein phosphatases are widely distributed through all domains of life (23). The eukaryotic members of the family include protein phosphatase 1, protein phosphatase 2A, and protein phosphatase 2B, commonly called calcineurin. Eukaryotic PPPs are serine/threonine-specific metalloenzymes (1, 42). Archaeal PPPs also are serine/threonine specific but require the presence of exogenous metal ion cofactors for activity (24). Bacterial PPPs also require exogenous metal ions for activity (24). However, they display extremely broad substrate specificity in vitro that encompasses both protein-bound phosphoserine and phosphothreonine as well as phosphotyrosine and phosphohistidine (24). In addition, the PPP family includes several enzymes that act on nonprotein substrates, such as the bacterial diadenosine tetraphosphatase ApaH (24, 34).

In this paper we describe the basic functional characteristics of two additional protein phosphatases from Synechocystis sp. strain PCC 6803: the products of open reading frames sll1033 and sll1387. While the former, SynPPM3, is one of eight potential PPM family protein phosphatases potentially encoded within the genome of this cyanobacterium, the latter, SynPPP1, is the sole potential representative of the PPP family of protein phosphatases. Both protein phosphatases displayed some unusual features, including the ability to catalyze the dephosphorylation of both phosphoserine- and phosphotyrosine-containing proteins in vitro.


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MATERIALS AND METHODS
 
Materials. Purchased materials included [{gamma}-32P]ATP from NEN Research Products (Boston, MA) and Chelating Sepharose Fast Flo from Pharmacia (Uppsala, Sweden). The pET101/D-TOPO cloning kit, enterokinase EKMax, and 10x reaction buffer were from Invitrogen (Carlsbad, CA). Superscript preamplification system and oligonucleotides were from Life Technologies, Inc. (Frederick, MD). Moloney murine leukemia virus reverse transcriptase, rRNasin inhibitor, and Wizard PCR preps were from Promega (Madison, WI). Chroma Spin-200 diethyl pyrocarbonate-H2O columns were from Clontech (Palo Alto, CA). Nytran membranes and a Spot Blot device were from Schleicher and Schuell (Keene, NH). Sheared salmon sperm DNA was from 5'-3', Inc. (Boulder, CO). Slide-A-Lyzer dialysis cassettes were from Pierce Biotechnology, Inc. (Rockford, IL). All general laboratory reagents and culture media were from Sigma (St. Louis, MO) or Fisher (Pittsburgh, PA), unless otherwise indicated. All absorbance measurements were made using a U-2000 spectrophotometer (Hitachi, Japan).

Standard procedures. Protein concentrations were determined by the method of Bradford (5) using premixed reagent and a standardized solution of bovine serum albumin (BSA), both from Pierce Biotechnology, Inc. (Rockford, IL). Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed as described by Laemmli (28). Gels were stained with Coomassie blue as described by Fairbanks et al. (11). The extraction of inorganic phosphate was performed using a modification of the procedure of Martin and Doty (36). For isolation of genomic DNA, cells were lysed and genomic DNA was isolated as described by Li et al. (32). For analysis of gene expression, cells were lysed and RNA was isolated by the method of Chomczynski and Sacchi (7).

Growth of organism and isolation of mRNA and genomic DNA. Cyanobacterium Synechocystis sp. strain PCC 6803 was grown at 30°C in BG11 medium (40) with continuous lighting (10 µE m2 s–1) and agitation: our control environmental condition. Perturbations in environmental conditions (Table 1) were introduced by taking cells in exponential growth phase (0.3 ≤ optical density at 730 nm [OD730] ≤ 0.4), harvesting by centrifugation, washing, and resuspending in new media. Twenty-four hours after initiation of the environmental perturbation, cells were harvested by centrifugation, washed once with 10 mM Tris-HCl, pH 8.0, containing 1 mM EDTA, and stored at –70°C. For heat shock and its associated control, cells were harvested after only 3 h to minimize morbidity/mortality. For phosphate or sulfate starvation, the period of exposure was extended to 48 h to deplete internal stores.


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  		TABLE 1. Environmental conditions testeda

Detection of mRNA. The levels of mRNA derived from ORFs sll1033 and sll1387 were determined by the array method of Chalifour et al. (6). Briefly, oligonucleotide probes, ~400 to 600 nucleotides in length, were prepared by PCR amplification of genomic DNA. For sll1033, the sequences of the forward and reverse primers were 5'-ATGCTGATCTGTCTCCAGTGC-3' and 5'-AACACATCCCATTCGGAACGG-3', respectively. The corresponding primers for sll1387 were 5'-ATGCCCAATCCTCGACGCA-3' and 5'-TCTAGGCAGTGAGCCAGCC-3'. The homogeneity and size of the resulting probes were verified by agarose gel electrophoresis. Following purification using Wizard PCR preps, the oligonucleotides were denatured by incubating in the presence of 0.3 M NaOH for 1 h at 70°C. After adding 0.4 volumes of 20x salt sodium citrate, duplicate portions (50 ng) of each probe were applied to a 102- by 133-mm Nytran Supercharged membrane in a 96-well Spot Blot apparatus and cross-linked to the membrane using an FB-UVXL-1000 cross-linker (Fisher, Pittsburgh, PA). Sheared salmon sperm DNA (50 ng) was used a negative control. Whole genomic DNA from Synechocystis sp. strain PCC 6803 (50 ng) was used as a positive control. Nonspecific binding sites were then blocked by incubation with sheared salmon sperm DNA.

Portions (2 µg) of cyanobacterial mRNA isolated from cells grown under various conditions were incubated for 60 min at 42°C in a volume of 25 µl containing 200 U of Moloney murine leukemia virus reverse transcriptase, 40 U of rRNasin RNase inhibitor, a mixture of gene-specific primers, 100 nM total, 0.5 mM dTTP, 0.5 mM dCTP, 0.5 mM dGTP, 5 µM dATP, and 0.8 µM [{alpha}-32P]dATP (3,000 Ci/mmol). Following initial denaturation at 94°C for 3 min, each sample was subjected to 25 cycles consisting of denaturation for 1 min at 94°C, annealing at 55°C for 1 min, and extension for 2 min at 72°C. Radiolabeled cDNA was isolated using Chroma Spin-200 diethyl pyrocarbonate-H2O columns according to the manufacturer's protocols. Agarose gel electrophoresis indicated that most cDNAs, where present, ranged from 150 to 400 bp in length. Portions (2 x 106 cpm each) of radiolabeled cDNA were mixed with 200 µg of sheared salmon sperm DNA in 10 ml of 6x salt sodium citrate containing Denhardt's reagent and 0.5% (wt/vol) SDS, applied to the wells of the Spot Blot device, and incubated for 12 h at 68°C. The membranes were removed from the Spot Blot device and washed three times, for 30 min at 68°C, with 200-ml volumes of 2x salt sodium citrate containing 1% (vol/vol) SDS, followed by two washings with 200 ml of 1% (wt/vol) SDS. The quantity of radioactive cDNA binding to each nucleotide probe was determined by electronic autoradiography using an Instantimager (Packard, Meriden, CT). Results were reported as the quantity of radioactive cDNA hybridized to each probe above the negative control, sheared salmon sperm DNA, relative to the positive control, genomic DNA from Synechocystis sp. strain PCC 6803, which was set equal to 1.0.

We determined the threshold of detection for our gene array empirically, specifically by asking how well it predicted the outcome of assays for mRNA expression via an established method, conventional reverse transcription-PCR (RT-PCR) using discrete sets of gene-specific primers. From our matrix of 26 ORFs and the 13 environmental conditions listed in Table 1, roughly 40% (138/338) were checked for the presence of mRNA by RT-PCR using gene-specific primers. Comparison of these data indicated that the highest degree of correlation between the results of gene array and RT-PCR, 126/138 or 91%, was reached when the threshold of detection was set at 0.30 for the gene array (data not shown). The genes encoding two other PPMs from Synechocystis sp. strain PCC 6803 served as important controls. The first, slr2031, has been reported by another laboratory using a completely unrelated methodology, Northern blotting, to be constitutively expressed (15). As predicted, we detected robust expression of slr2031 using our array method. The second, slr1860 or icfG, has been reported elsewhere using reporter constructs to be expressed in media containing glucose or cells deprived of light but not in cells grown under our control conditions or in the presence of added carbonate (15). As predicted, both our gene array and conventional PCR analyses detected expression of slr1860 in cells deprived of light or to which glucose had been added but not in control cells or cells to which carbonate had been added in the absence of glucose (45). Moreover, it should be pointed out that the amount of signal detected under the former conditions, 0.42 to 0.45, was comparable to that observed for sll1387.

Cloning of ORF sll1033 and expression of its recombinant protein product. ORF sll1033 was cloned using the materials provided in the pET101/D-TOPO cloning kit following the manufacturer's protocols. Briefly, ORF sll1033 was amplified by PCR using genomic DNA (20 ng) as template and 10 pmol each of a forward and a reverse oligonucleotide primer. The sequences of the forward and reverse primers were, respectively, 5'-ATTATGCTGATCTGTCTCCA-3' and 5'-AAACTACCACACTCCAAAGG-3'. The resulting PCR products were then ligated into vector PCR T7/NT-TOPO, which added oligonucleotides encoding an N-terminal extension to the full-length sequence of the wild-type protein that contained a hexahistidine sequence and a recognition epitope for the anti-Xpress antibody. The resulting plasmids were used to transform One Shot TOP 10 chemically competent E. coli. The orientation and sequence of the cloned genes were verified by sequencing the isolated plasmids. The plasmids were then used to transform E. coli BL21Star (DE3) One Shot cells. The transformed cells were cultured at 37°C in 200 ml of Luria-Bertani medium containing 0.1 mg/ml ampicillin until the OD600 reached 0.6 to 1.0. Isopropyl-ß-D-thiogalactopyranoside was then added to a final concentration of 1 mM, and the cells were harvested 3 h later.

The cell pellet from cells expressing the recombinant product of ORF sll1033, rSynPPM3 (a recombinantly produced form of SynPPM3 containing an N-terminal histidine tag), harvested as described above, was resuspended in 5 ml of lysis buffer: 50 mM Tris-HCl, pH 7.5, containing 10 mM imidazole, 250 mM NaCl, and 5% (vol/vol) glycerol. Lysozyme (10 mg) was added and the suspension placed on ice for 30 min. The cells then were lysed by sonic disruption and the lysate clarified by centrifugation at 10,000 x g for 20 min. The supernatant liquid was applied to a 0.5- by 5-cm column of Chelating Sepharose Fast Flo that had been charged with NiSO4 and subsequently equilibrated with lysis buffer. The column was then washed in lysis buffer, and adherent proteins were eluted with lysis buffer containing 250 mM imidazole. Protein-containing fractions were pooled, dialyzed versus 20 mM Tris-HCl, pH 7.5, containing 0.5 mM EDTA and 1 mM dithiothreitol, and stored at 4°C.

Cloning of ORF sll1387 and isolation of its recombinant protein product. ORF sll1387 was amplified by PCR from 1 µg of genomic DNA using 50 pmol of a forward primer whose sequence was 5'-CGATCTCGAGCCATGCCCAATCCTCGACGCA-3', 50 pmol of a reverse primer whose sequence was 5'-ATCGGGTACCAACAGGCTCTCAGGCTACCC-3', and 5 U of Pfu DNA polymerase (Stratagene, La Jolla, CA) according to the manufacturer's instructions. The resulting PCR product was ligated into expression vector pRSET-C and sequenced to ensure the fidelity of the DNA amplification and cloning processes. Competent E. coli strain BL21(DE3) pLyS cells were transformed with the isolated plasmid and grown in 200-ml portions of Luria broth containing 50 µg/ml ampicillin until the OD600 reached 0.6 to 1.0. Isopropyl-ß-D-thiogalactopyranoside then was added to a final concentration of 0.4 mM. The culture was then incubated for 4 to 6 h at 25°C and harvested by centrifugation, and the cell pellet was stored at –20°C until needed.

The recombinant product of ORF sll1387, rSynPPP1 (a recombinantly produced form of SynPPP1 containing an N-terminal histidine tag), was isolated from the cell pellets of two 200-ml cultures following the procedure described above for rPPM3 with the following modifications. The concentration of NaCl in the lysis buffer was decreased to 200 mM and glycerol was omitted. The column also was washed with lysis buffer containing 40 mM imidazole prior to elution of adherent proteins.

Assay of protein phosphatase activity. Radiolabeled phosphoprotein substrates were prepared and assayed as previously described (25). Assays of the protein phosphatase activity of rSynPPM3 (40 to 400 ng) were performed at 37°C in 50 µl of 100 mM Tris-HCl, pH 7.5, containing 50 mM NaCl, 2 mM dithiothreitol, 5 mM MnCl2, 0.5 mg/ml BSA, and 1 to 4 µM protein-bound [32P]phosphate. Assays of the protein phosphatase activity of rSynPPP1 were performed in a similar manner, with the following exceptions. The assay temperature was reduced to 30°C, the pH was increased to 8.0, and the BSA concentration was increased to 1 mg/ml. Under the conditions described above, the quantity of product detected was observed to be linearly dependent upon both time of incubation and enzyme concentration. "In gel" assays of protein-serine and protein-tyrosine phosphatase activity were performed on 10-µg portions of rSynPPP1 using 32P-labeled P-Ser casein (casein that has been phosphorylated on serine residues in vitro using the catalytic subunit of the cyclic AMP [cAMP]-dependent protein kinase) or P-Tyr casein (casein that has been phosphorylated on tyrosine residues in vitro using lyn protein-tyrosine kinase) as described by Gates et al. (13).

Assay of activity toward p-nitrophenyl phosphate. The activity of rSynPPP1 and rSynPPM3 toward p-nitrophenyl phosphate (pNPP) was assayed under conditions similar to those described above for protein substrates, with the exception that the quantity of enzyme was increased to 4 µg and the volume of the initial incubation was increased to 400 µl. The concentration of pNPP ranged from 1 to 10 mM as indicated in the individual figure and table legends. For the determination of the reaction product, p-nitrophenol, the incubation was terminated by the addition of 600 µl of 0.5 M EDTA, pH 9.0, to prevent the formation of insoluble Mn(OH)2 (12). p-Nitrophenolate was then measured spectrophotometrically at a wavelength of 420 nm. For the determination of the other reaction product, inorganic phosphate, the Malachite Green method of Lanzetta et al. (29) was utilized. Under the conditions described above, the quantity of product detected was observed to be linearly dependent upon both time of incubation and enzyme concentration.

Metal analysis. In the first (trial) analysis, rSynPPP1, purified as described above, was dialyzed overnight at 4°C against two changes, 100 volumes each, of 25 mM Tris, pH 7.5, containing 0.5 mM EDTA in a Slide-A-Lyzer dialysis cassette whose membrane had a 10-kDa molecular mass cutoff. The final concentration of protein was 0.15 mg/ml. The retentate and the final dialysate were then analyzed for the presence of over 60 metals by West Coast Analytical Service, Inc. (Santa Fe Springs, CA) using inductively coupled plasma mass spectrometry. As the analysis indicated the presence of significant levels of Ni, the metal ion used for the affinity purification step, as well as Mn and Mg, a second set of samples was prepared. An aliquot of the protein (2.5 ml) was diluted to a volume of 3.1 ml with dialysis buffer and dialyzed against three changes, 100 volumes each, of 25 mM Tris, pH 7.5, containing 0.5 mM EDTA. The final concentration of protein was 0.23 mg/ml. The retentate and the final dialysate were then analyzed for the presence of Mg, Mn, and Ni as described above.


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RESULTS
 
mRNAs for SynPPP1and SynPPM3 are expressed under a variety of conditions. Cultures of Synechocystis sp. strain PCC 6803 that had been exposed to a variety of nutritional and environmental conditions (Table 1) were surveyed for the presence of mRNA encoding SynPPP1 or SynPPM3 using an adaptation of the array technique of Chalifour et al. (6). As can be seen in Fig. 1, mRNA encoding SynPPP1 could be detected in cells grown under a variety of conditions. Highest levels were evident in cells that had been exposed to carbon-rich media containing millimolar levels of carbonate, glucose, or glucose plus carbonate or media lacking nitrate, phosphate, or sulfate. Little or no mRNA could be detected in cells that had been deprived of light or cells that had been deprived of fixed carbon. mRNA encoding SynPPM3 was detected under the vast majority (11/13 = 85%) of the conditions surveyed. The sole exceptions were cells that had been deprived of light or cells that had been exposed to high levels of carbonate.



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  		FIG. 1. Detection of mRNA encoding SynPPP1 and SynPPM3 by gene array. The presence and relative levels of mRNA for SynPPP1 (shaded bars) and SynPPM3 (hatched bars) were determined by measuring the levels of radiolabeled cDNA produced by reverse transcription in the presence of [{alpha}-32P]dATP as described in Materials and Methods. All results were corrected for nonspecific binding to salmon sperm DNA and normalized to the levels of radiolabeled cDNA that hybridized to the positive control probe, genomic DNA. Shown are the averages of duplicate determinations ± standard error. The horizontal line indicates the threshold at which it was considered, with 91% confidence, that detectable quantities of the mRNA of interest were present (for further details, see Materials and Methods). For a description of the various culture conditions employed, see Table 1.

As the apparent level of expression of mRNA encoding SynPPP1 was, in many cases, at or near the level of detection of the array technique, selected samples were analyzed by conventional RT-PCR to independently ascertain whether message was present. Four conditions were tested, including BG11 medium with continuous light (control), heat shock, replacement of nitrate by ammonia, and light deprivation. While conventional RT-PCR is only semiquantitative at best, the pattern of expression of the RT-PCR product mirrored that observed by gene array. Specifically, the predicted 620-bp PCR product was visible when mRNA from cells grown under control conditions, subject to heat shock, or exposed to ammonia in the absence of nitrate was used as template but not when mRNA from cells deprived of light was used as template (Fig. 2).



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  		FIG. 2. RT-PCR analysis of expression of mRNA encoding SynPPP1. Shown is an ethidium bromide-stained agarose gel of the RT-PCR products obtained from cells grown under control conditions (lane 5), in the absence of light (lane 4), at 42°C for 3 h (lane 3), or with ammonia as the fixed nitrogen source (lane 2) as described in Table 1. The migration position of the 620-bp PCR product obtained from sll1387 is shown at left. Lane 1 shows the results obtained when reverse transcription was omitted, which permits detection of any contaminating genomic DNA.

Further verification of the gene array method was provided by parallel analyses of two other ORFs encoding PPM family protein phosphatases, slr1860 (2) and slr2031 (15), whose patterns of expression had been reported by other laboratories using independent methods—reporter constructs and Northern blotting, respectively. Expression of slr2031, which reportedly is constitutively expressed in Synechocystis sp. strain PCC 6803 (15), was detected under all 13 conditions listed in Table 1. Expression of slr1860 reportedly was induced by glucose or light deprivation (2). Using both gene array and conventional RT-PCR, we observed expression of this ORF in cells to which glucose had been added to the culture media or which were deprived of light but not in cells grown in light or to which elevated levels of carbonate were added. We also detected expression in cells lacking nitrate or subject to heat shock, but reference data for these conditions were not apparent in the literature. Thus, our gene array analyses were validated by two types of controls, one a constitutively expressed gene and another an inducible gene whose levels of expression, when induced, were comparable (0.42 to 0.45) to those of slr1387 (data not shown).

rSynPPP1 and rSynPPM3 exhibit protein phosphatase activity. ORFs sll1033 and sll1387 encoding SynPPP1 and SynPPM3, respectively, were amplified via PCR and cloned into vectors encoding N-terminal fusion domains containing hexahistidine sequences. The recombinant fusion proteins, designated rSynPPP1 and rSynPPM3, were expressed in E. coli and purified to apparent electrophoretic homogeneity by metal-chelate chromatography. Both recombinant fusion proteins hydrolyzed a common synthetic substrate for phosphohydrolases, p-nitrophenyl phosphate, at measurable rates (Table 2). When challenged with a battery of exogenous phosphoprotein substrates, both enzymes displayed the ability to dephosphorylate protein-bound phosphoserine and phosphotyrosine (Table 2). The radioactive reaction product could be quantitatively extracted into organic solvents as a molybdate complex, verifying that it was indeed inorganic phosphate (data not shown).


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  		TABLE 2. Activity of rSynPPP1 and rSynPPM3 against exogenous substratesa

Unlike previously described PPPs from prokaryotic organisms, which require the addition of divalent metal ion cofactors (24, 44), rSynPPP1 displayed robust catalytic activity in the presence of millimolar concentrations of EDTA. Only when an unusually high level, 50 mM, of chelating agent was added could significant (~40%) inhibition of activity be observed. Conversely, the addition of Mg2+ or Mn2+ only marginally (≤20%) stimulated the protein phosphatase activity of rSynPPP1. rSynPPM3, on the other hand, displayed little on no activity unless divalent metal ions were added. The latter enzyme thus conformed to the consistent pattern displayed by previously described members of the PPM superfamily from both eukaryotic (1) and bacterial (24, 44) organisms. Mn2+ was the most potent activator of rSynPPM3 in vitro, while Ca2+ proved unexpectedly effective. For Mn2+, half-maximal activation was reached at a concentration of ~0.5 mM.

rSynPPP1 and rSynPPM3 displayed maximal activity between pH 7.5 and 8.4 and pH 6.5 and 8.5, respectively. rSynPPP1 and rSynPPM3 each were inhibited by millimolar concentrations of sodium pyrophosphate and moderately inhibited by NaF. Both enzymes were insensitive to tetramisole, a widely used inhibitor of alkaline phosphatases; tartrate, an inhibitor of many acid phosphatases; o-vanadate, a general inhibitor of protein-tyrosine phosphatases; and microcystin-LR or okadaic acid, potent and selective inhibitors of the PPP family protein-serine/threonine phosphatases PP1 and PP2A from eukaryotes (data not shown).

Steady-state kinetic analyses were performed with rSynPPP1 and rSynPPM3 (Table 3) using selected substrates. It was noted that rSynPPP1 exhibited a 10-fold-lower Km toward reduced carboxyamidomethylated and maleylated lysozyme containing 32P-labeled phosphoserine (32P-Ser RCML [RCML that has been phosphorylated on serine residues in vitro using the catalytic subunit of the cAMP-dependent protein kinase]), a small acidic protein, than it did toward 32P-Ser myelin basic protein (32P-Ser MBP [MBP that has been phosphorylated on serine residues in vitro using the catalytic subunit of the cAMP-dependent protein kinase]), a small basic protein. Vmax values for the two substrates were fairly comparable. rSynPPM3 dephosphorylated 32P-Ser casein and 32P-Ser MBP with maximal velocities nearly 8- and 45-fold greater, respectively, than it did a phosphotyrosine-containing protein, 32P-Tyr RCML (RCML that has been phosphorylated on tyrosine residues in vitro using lyn protein-tyrosine kinase). On the other hand, the enzyme's Km for 32P-Tyr RCML was comparable to that for P-Ser casein, another acidic protein, and over 10-fold lower than that toward 32P-Ser MBP.


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  		TABLE 3. Michaelis constants for rSynPPP1 and rSynPPM3a

Verification of the polypeptide sources of protein-serine and protein-tyrosine phosphatase activity. rSynPPP1 was subject to SDS-PAGE in gels in which 32P-Ser casein or 32P-Tyr casein had been copolymerized into the polyacrylamide matrix. After the enzyme was renatured in situ, a prominent band of protein phosphatase activity appeared in each gel whose position coincided with that of the Coomassie blue-stained protein (data not shown). Attempts to similarly renature and assay rSynPPM3 in gel proved unsuccessful. Therefore, site-directed mutagenesis was employed to produce recombinant versions of rSynPPM3 lacking conserved, and therefore presumably functionally essential, amino acid residues as an alternate means for verifying that the recombinant enzyme was the source of both the protein-serine and the protein-tyrosine phosphatase activities detected in the assays described above.

Mutagenic alteration of each of seven conserved acidic residues distributed among subdomains 1, 2, 8, 9, and 11 (Table 4) was observed to perturb catalytic activity. Many of these residues were predicted from the X-ray structure of PP2C{alpha} from humans (8) to participate in binding the two divalent metal ion cofactors essential for catalysis. Substitution of Asp604, which was predicted to coordinate directly to metal ion 1 as well as to a water molecule that interacts with both metal ion 1 and the phosphoryl group of the substrate, with asparagine resulted in the virtual elimination (i.e., <2% of control) of activity towards pNPP as well as two phosphoserine- and one phosphotyrosine-containing proteins (Table 5). Similarly, substitution of either Asp415, which was predicted to coordinate metal ion 2 via a bridging water molecule, or Asp648, which was predicted to directly coordinate metal ion 1, greatly reduced activity toward pNPP as well as 32P-Ser casein and 32P-Tyr RCML. We therefore concluded that rSynPPM3 was the predominant source of both the protein-serine and protein-tyrosine phosphatase activity in our preparations.


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  		TABLE 4. Location of residues mutagenically altered in rSynPPM3a


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  		TABLE 5. Effect of mutagenically produced alterations on the catalytic efficiency of rSynPPM3a

Substitution of Asp415 with alanine instead of asparagine diminished the efficiency with which rSynPPM3 hydrolyzed 32P-Ser MBP and 32P-Tyr RCML by 5- to 10-fold, respectively, but only marginally diminished the enzyme's activity toward 32P-Ser casein. The mutagenic alteration of several other residues elicited similarly complex patterns of behavior. Substitution of Glu414, which was predicted to bind metal ion 2 via a bridging water molecule, with glutamine had no detectable effect on the enzyme's ability to hydrolyze pNPP and decreased the rate at which it hydrolyzed 32P-Ser casein and 32P-Tyr RCML only about twofold. Perhaps this is not surprising, as the presence of an acidic amino acid at this position, while very common, is not absolutely conserved among the members of the PPM family (4). Alteration of Asp444, which was predicted to coordinate both metal ions, to asparagine diminished activity toward 32P-Ser MBP by roughly fivefold but only marginally affected the hydrolysis of 32P-Ser casein or 32P-Tyr RCML. Mutagenic alteration of Glu618, a highly conserved residue not implicated directly in catalysis, to glutamine diminished activity toward 32P-Tyr RCML ca. sixfold while leaving the enzyme's ability to hydrolyze pNPP or 32P-Ser casein largely unimpaired. Most surprising of all, replacement of Asp608 by asparagine resulted in a fourfold increase in activity toward 32P-Tyr RCML, with only minor, i.e., ≤2-fold, decreases in activity toward pNPP or 32P-Ser casein.

rSynPPP1 is a metalloprotein. As the eukaryotic members of the PPP superfamily are metalloproteins containing tightly associated metals (1, 42), we asked whether rSynPPP1 also was a metalloprotein. rSynPPP1 was extensively dialyzed against buffers containing a low (0.5 mM) concentration of EDTA and then analyzed for the presence of over 60 different metals by inductively coupled plasma mass spectrometry. Initial analyses revealed that only Mn, Mg, and Ni were present in the retentate at levels above that detected in the dialysate (Table 6, analysis 1). A more extensive regimen of dialysis dramatically lowered the level of Ni without markedly altering the levels of Mn or Mg (Table 6, analysis 2), suggesting that the former metal had carried over from the Ni2+ affinity column used to purify the protein, while Mg and Mn were associated predominantly with the protein phosphatase.


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  		TABLE 6. Metal analysis of rSynPPP1a


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DISCUSSION
 
Since the PPP family encompasses a broad range of phosphodiesterases, pyrophosphatases, and phosphomonesterases that act on nucleotides, lipids, and other nonprotein biomolecules (34), we asked whether SynPPP1 was in fact a protein phosphatase. The recombinantly expressed form of SynPPP1, rSynPPP1, dephosphorylated pNPP, a synthetic substrate hydrolyzed by a broad range of phosphoesterases, as well as several exogenous phosphoproteins in vitro. The latter included P-Ser casein, which we had observed in preliminary studies (45). Compared to protein phosphatase 1 (PP1), protein phosphatase 2A (PP2A), and protein phosphatase 2B (PP2B), the most prominent members of the PPP family of protein phosphatases in eukaryotes, the Vmax values for the cyanobacterial enzyme toward phosphoserine-containing proteins were 10- to 100-fold lower than those reported for PP2A (14, 37) or PP2B (3, 10, 39) but similar to the only reported value for PP1 (14). When one considers that the kinetic parameters for PP1, PP2A, and PP2B were determined using cognate phosphoprotein substrates, or phosphopeptides modeled thereupon, while the cyanobacterial enzyme was assayed with the physiologically and phylogenetically irrelevant phosphoproteins available, i.e., casein and MBP, the observed differences would not appear to be particularly surprising. Moreover, the apparent affinity of rSynPPP1 for these phosphoproteins, a potentially more incisive indication of the general nature of the enzyme's intended substrate(s), was quite comparable to that of its eukaryotic counterparts (3, 10, 14, 18, 37, 39, 50). It would therefore appear that SynPPP1 is a bona fide protein phosphatase.

Like virtually all of its bacterial counterparts, but in stark contrast to the archaeal and eukaryotic members of the PPP family of protein phosphatases, rSynPPP1 dephosphorylated exogenous phosphotyrosine-containing proteins at rates comparable to phosphoserine-containing ones. Intriguingly, the sole exception to this pattern of dual specificity among bacterial PPPs, the psychrophilic phosphatase I from Shewanella sp. (48), was reported to be exclusively phosphotyrosine specific (47).

The members of the PPP family of phosphatases utilize a pair of active site metal ions to polarize the water molecule responsible for hydrolyzing their phosphoester substrates (1, 34). In the case of eukaryotic PPPs, these metals are incorporated into the protein as stably bound prosthetic groups (9). On the other hand, the catalytic metal ions for all previously examined bacterial and archaeal PPPs behaved as readily dissociated cofactors (24, 44). Hence, assays of the catalytic activity of prokaryotic PPPs required the addition of Mn2+ and/or Mg2+. Surprisingly, rSynPPP1 exhibited near maximum catalytic activity in the absence of added metal ions. Exposure to a sufficiently high level of EDTA reduced activity by nearly half, implying that the catalytically essential metals were already present and tightly bound. This inference was buttressed using inductively coupled plasma mass spectrometry, which revealed the presence of significant quantities of Mg and Mn.

While the quantity of Mn and Mg detected (0.32 to 0.44 mol/mol) fell well short of the predicted stoichiometry of 2.0 (42), it should be noted that the enzyme was subject to prolonged dialysis prior to analysis. It would not be unexpected that bound metals would slowly leach from the protein during this process. Alternatively, the mass of exogenous polypeptide produced via recombinant expression may have overwhelmed the capacity of the E. coli host to insert the catalytic metals into the apoprotein, resulting in the production of a mixture of metal-containing and metal-deficient protein. If so, the addition of metals in vitro to the apoprotein apparently was insufficient to generate catalytically competent holoenzyme. If E. coli cells are capable of properly processing recombinantly produced SynPPP1, this implies that PrpA and PrpB, the PPP family protein phosphatases indigenous to the bacterium, also are metalloproteins. Unfortunately, no information is available on the effect of divalent metals, or the lack thereof, on these enzymes, although Mn2+ and Mg2+ apparently were present in all assays of their catalytic activity (38).

While rSynPPM3 displayed the dependence on divalent metal ion cofactors typical of all previously described members of the PPM family of protein phosphatases (1, 24), it dephosphorylated both of the phosphotyrosine-containing proteins with which it was challenged in vitro. As only one other member of the PPM family has been reported to display a similar dual specific capability, the latter observation was quite unexpected. Intriguingly, the exception to the pattern of absolute specificity for protein-bound phosphoserine and/or phosphothreonine residues, PphA, also is from Synechocystis sp. strain PCC 6803 (41). While the Km values for rSynPPM3 were comparable to those of PphA (41) and generally lower than those reported for PP2C from humans (12, 17) or paramecia (27), the former's Vmax values were noticeably lower as well.

Mutagenic alteration of Asp415, Asp604, or Asp648 to asparagine eliminated or greatly reduced the ability of our recombinant enzyme preparations to dephosphorylate pNPP, P-Ser casein, and P-Tyr RCML, establishing that rSynPPM3 was the predominant source of both the protein-serine and protein-tyrosine phosphatase activity in our preparations. However, the effects of mutagenically altering several other conserved amino acids proved remarkably substrate dependent. As the majority of the amino acids targeted for substitution were predicted to anchor the binuclear metal center, the simplest explanation for the observed mosaic pattern was that each alteration perturbed its geometry or Lewis acid potential in a subtly different manner.

An examination of the handful of published studies on the mutagenic alteration of other members of the PPM family offered few insights regarding this hypothesis, as each laboratory employed a single substrate for the assessment of catalytic activity in vitro. However, hints of complex effects can be gleaned. For example, while Sheen (43) reported that substitution of the methionine-glutamic acid-aspartic acid sequence in motif 1 by isoleucine-glycine-histidine in the PPM family protein phosphatase ABI1 from Arabidopsis thaliana (Table 6) eliminated activity toward 32P-Ser casein in vitro, Jackson et al. (17) observed that substitution of the corresponding aspartic acid residue in this motif, Asp38, with alanine in PP2C{alpha} from humans had no significant impact on the enzyme's ability to hydrolyze pNPP. Moreover, even though the altered form of ABI1 appeared to lack catalytic activity in vitro, it blocked the induction of gene expression by abscisic acid as effectively as the unaltered phosphatase in vivo (43). By contrast, replacement of the aspartic acid-glycine-histidine sequence in motif 2 by lysine-leucine-asparagine in ABI1 eliminated both activity toward 32P-Ser casein and the ability to block abscisic acid-mediated signals. In the final analysis, X-ray crystallographic studies of enzyme-substrate complexes using both the native enzyme and mutagenically altered forms thereof will be needed to deconvolute this complex mosaic.

Might SynPPM3 or other PPM family protein phosphatases act as a protein-tyrosine phosphatase in vivo? Kinetic analyses indicated that the Vmax values of rSynPPM3 toward P-Tyr RCML were roughly 8- and 45-fold higher, respectively, than those toward P-Ser casein and P-Ser MBP. However, the Km value toward the tyrosine phosphorylated protein was comparable to that toward P-Ser casein and much lower than that toward P-Ser MBP. Moreover, substitution of Glu608 with glutamine substantially enhanced the enzyme's activity toward 32P-Tyr RCML, suggesting at least latent potential as a protein-tyrosine phosphatase. Intriguingly, several other members of the PPM family contain asparagine at this position (4, 46), suggesting that some forms of this enzyme may target phosphotyrosine-containing proteins in vivo. What is clear at this point is that bacteria have been much more liberal than their eukaryotic brethren in adapting existing protein phosphatase scaffolds for the hydrolysis of a wide range of different phosphoamino acid residues. Further exploration of the prokaryotic domains undoubtedly will reveal more protein phosphatases that exhibit novel physical and/or functional properties.


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ACKNOWLEDGMENTS
 
These studies were supported by grant GM 55067 from the National Institutes of Health.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biochemistry-0308, Virginia Polytechnic Institute and State University, Blacksburg, VA 24061. Phone: (540) 231-4317. Fax: (540) 231-9070. E-mail: pjkennel{at}vt.edu. Back

{dagger} Present address: Department of Microbiology, University of Tennessee, Knoxville, TN 37996. Back

{ddagger} Present address: Microbiology Group, Pacific Northwest National Laboratory, Richland, WA 99352. Back


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REFERENCES
 
    1
  1. Barford, D. 1996. Molecular mechanisms of the protein serine/threonine phosphatases. Trends Biochem. Sci. 21:407-412.[CrossRef][Medline]
    2. 2
  2. Beuf, L., S. Bedu, M.-C. Durand, and F. Joset. 1994. A protein involved in co-ordinate regulation of inorganic carbon and glucose metabolism in the facultative photoautotrophic cyanobacterium Synechocystis PCC6803. Plant Mol. Biol. 25:855-864.[CrossRef][Medline]
    3. 3
  3. Blumenthal, D. K., K. Takio, R. S. Hansen, and E. G. Krebs. 1986. Dephosphorylation of cAMP-dependent protein kinase regulatory subunit (type II) by calmodulin-dependent protein phosphatase. Determinants of substrate specificity. J. Biol. Chem. 261:8140-8145.[Abstract/Free Full Text]
    4. 4
  4. Bork, P., N. G. Brown, H. Hegyi, and J. Schultz. 1996. The protein phosphatase 2C (PP2C) superfamily: detection of bacterial homologues. Protein Sci. 5:1421-1425.[Medline]
    5. 5
  5. Bradford, M. M. 1976. A rapid and simple method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254.[CrossRef][Medline]
    6. 6
  6. Chalifour, L. E., R. Fahmy, E. L. Holder, E. W. Hutchinson, C. K. Osterland, H. M. Schipper, and E. Wang. 1994. A method for analysis of gene expression patterns. Anal. Biochem. 216:299-304.[CrossRef][Medline]
    7. 7
  7. Chomczynski, P., and N. Sacchi. 1987. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal. Biochem. 162:156-159.[Medline]
    8. 8
  8. Das, A. K., N. R. Helps, P. T. W. Cohen, and D. Barford. 1996. Crystal structure of the protein serine/threonine phosphatase 2C at 2.0 A resolution. EMBO J. 15:6798-6809.[Medline]
    9. 9
  9. Egloff, M.-P., P. T. W. Cohen, P. Reinemer, and D. Barford. 1995. Crystal structure of the catalytic subunit of human protein phosphatase 1 and its complex with tungstate. J. Mol. Biol. 254:942-959.[CrossRef][Medline]
    10. 10
  10. Enz, A., G. Shapiro, A. Chappius, and A. Dattler. 1994. Nonradioactive assay for protein phosphatase 2B (calcineurin) activity using a partial sequence of the subunit of cAMP-dependent protein kinase as substrate. Anal. Biochem. 216:147-153.[CrossRef][Medline]
    11. 11
  11. Fairbanks, G., T. L. Steck, and D. F. H. Wallace. 1971. Electrophoretic analysis of the major polypeptides of the human erythrocyte membrane. Biochemistry 10:2606-2617.[CrossRef][Medline]
    12. 12
  12. Fjeld, C. C., and J. M. Denu. 1999. Kinetic analysis of human serine/threonine protein phosphatase 2C. J. Biol. Chem. 274:20336-20343.[Abstract/Free Full Text]
    13. 13
  13. Gates, R. E., J. L. Miller, and L. E. King, Jr. 1996. Activity and molecular weight of protein tyrosine phosphatases in cell lysates determined by renaturation after gel electrophoresis. Anal. Biochem. 237:208-215.[CrossRef][Medline]
    14. 14
  14. Hemmings, H. C., Jr., A. C. Nairn, J. I. Elliott, and P. Greengard. 1990. Synthetic peptide analogs of DARPP-32 (Mr 32,000 dopamine and cAMP-regulated phosphoprotein), an inhibitor of protein phosphatase-1. J. Biol. Chem. 265:20369-20376.[Abstract/Free Full Text]
    15. 15
  15. Huckauf, J., C. Nomura, K. Forchhammer, and M. Hagemann. 2000. Stress responses of Synechocystis sp. strain PCC6803 mutants impaired in genes encoding putative alternative sigma factors. Microbiology 146:2877-2889.[Abstract/Free Full Text]
    16. 16
  16. Irmler, A., and K. Forchhammer. 2001. A PP2C-type phosphatase dephosphorylates the PII signaling protein in the cyanobacterium Synechocystis PCC6803. Proc. Natl. Acad. Sci. USA 98:12978-12983.[Abstract/Free Full Text]
    17. 17
  17. Jackson, M. D., C. C. Fjeld, and J. M. Denu. 2003. Probing the function of conserved residues in the serine/threonine phosphaase PP2C{alpha}. Biochemistry 42:8513-8521.[CrossRef][Medline]
    18. 18
  18. Jones, J. A., and Y. A. Hannun. 2002. Tight binding inhibition of protein phosphatase-1 by phosphatidic acid. Specificity of inhibition by phospholipid. J. Biol. Chem. 277:15530-15538.[Abstract/Free Full Text]
    19. 19
  19. Kamei, A., S. Yoshihara, T. Yuasa, X. Geng, and M. Ikeuchi. 2003. Biochemical and functional characterization of a eukaryotic-type protein kinase, SpkB, in the cyanobacterium, Synechocystis sp. PCC6803. Curr. Microbiol. 46:296-301.[CrossRef][Medline]
    20. 20
  20. Kamei, A., T. Yuasa, X. Geng, and M. Ikeuchi. 2002. Biochemical examination of the potential eukaryotic-type protein kinase genes in the complete genome of the unicellular cyanobacterium Synechocystis sp. PCC6803. DNA Res. 9:71-78.[Abstract]
    21. 21
  21. Kamei, A., T. Yuasa, K. Orikawa, X. X. Geng, and M. Ikeuchi. 2001. A eukaryotic-type protein kinase, SpkA, is required for normal motility of the unicellular cyanobacterium Synechocystis sp. PCC6803. J. Bacteriol. 183:1505-1510.[Abstract/Free Full Text]
    22. 22
  22. Kaneto, T., S. Sato, H. Kotani, A. Tanaka, E. Asamizu, Y. Nakamura, N. Miyajima, M. Hirosawa, M. Sugiura, S. Sasamoto, T. Kimura, T. Hosouchi, A. Matsuno, A. Muraki, N. Nakazaki, K. Naruo, S. Okumura, S. Shimpo, C. Takeuchi, T. Wada, A. Watanaba, M. Yamada, M. Yasuda, and S. Tabata. 1996. Sequence analysis of the genome of the unicellular cyanobacterium Synechocystis sp strain PCC 6803. II. Sequence determination of the entire genome and assignment of potential protein-coding regions. DNA Res. 30:109-136.
    23. 23
  23. Kennelly, P. J. 2002. Protein kinases and protein phosphatases in prokaryotes: a genomic perspective. FEMS Microbiol. Lett. 206:1-8.[CrossRef][Medline]
    24. 24
  24. Kennelly, P. J. 2001. Protein phosphatases: a phylogenetic perspective. Chem. Rev. 101:2291-2312.[CrossRef][Medline]
    25. 25
  25. Kennelly, P. J., K. A. Oxenrider, J. Leng, J. S. Cantwell, and N. Zhao. 1993. Identification of a serine/threonine-specific protein phosphatase from the Archaebacterium Sulfolobus solfataricus. J. Biol. Chem. 268:6505-6510.[Abstract/Free Full Text]
    26. 26
  26. Kim, Y. H., Y. M. Park, S.-J. Kim, Y.-I. Park, J.-S. Choi, and Y.-H. Chung. 2004. The role of Slr1443 in pilus biogenesis in Synechocystis sp. PCC6803: involvement in post-translational modification of pilins. Biochem. Biophys. Res. Commun. 315:179-186.[CrossRef][Medline]
    27. 27
  27. Klumpp, S., C. Hanke, A. Donella-deana, A. Beyer, R. Keilner, L. A. Pinna, and J. E. Schultz. 1994. A membrane-bound protein phosphatase type 2C from Paramecium tetraurelia. Purification, characterization, and cloning. J. Biol. Chem. 269:32774-32780.[Abstract/Free Full Text]
    28. 28
  28. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685.[CrossRef][Medline]
    29. 29
  29. Lanzetta, P. A., L. J. Alvarez, P. S. Reinach, and O. A. Candia. 1979. An improved assay for nanomole amounts of inorganic phosphate. Anal. Biochem. 100:95-97.[CrossRef][Medline]
    30. 30
  30. Leonard, C. J., L. Aravind, and E. V. Koonin. 1998. Novel families of putative protein kinases in Bacteria and Archaea: evolution of the "eukaryotic" protein kinase superfamily. Genome Res. 8:1038-1047.[Abstract/Free Full Text]
    31. 31
  31. Leung, J., S. Merlot, and J. Giraudat. 1997. The Arabidopsis abscisic acid-insensitive2 (ABI2) and ABI1 genes encode homologous protein phosphatases 2C involved in abscisic acid production. Plant Cell 9:759-771.[Abstract]
    32. 32
  32. Li, R., H. J. Debella, and W. W. Carmichael. 2001. Isolates identifiable as Athrospira maxima and Athrospira fusiformis (Oscillatoriales, Cyanobacteria) appear identical on the basis of a morphological study in culture and 16S rRNA sequences. Phycologia 40:367-371.
    33. 33
  33. Li, R., J. D. Haile, and P. J. Kennelly. 2003. An arsenate reductase from Synechocystis sp. strain PCC6803 exhibits a novel combination of catalytic characteristics. J. Bacteriol. 185:6780-6789.[Abstract/Free Full Text]
    34. 34
  34. Lohse, D. L., J. M. Denu, and J. E. Dixon. 1995. Insights derived from the structures of the ser/thr phosphatases calcineurin and protein phosphatase 1. Curr. Biol. 3:987-990.
    35. 35
  35. Mann, D. J., D. G. Campbell, C. H. McGowan, and P. T. W. Cohen. 1992. Mammalian protein serine/threonine phosphatase 2C: cDNA cloning and comparative analysis of amino acid sequences. Biochim. Biophys. Acta 1130:100-104.[Medline]
    36. 36
  36. Martin, J. B., and D. M. Doty. 1949. Determination of inorganic phosphate. Modification of the isobutyl alcohol procedure. Anal. Chem. 21:965-967.[CrossRef]
    37. 37
  37. Mertz, P., L. Yu, R. Sikkink, and F. Rusnak. 1997. Kinetic and spectroscopic analyses of mutants of a conserved histidine in the metallophosphatases calcineurin and {lambda} protein phosphatase. J. Biol. Chem. 272:21296-21302.[Abstract/Free Full Text]
    38. 38
  38. Missiakis, D., and S. Raina. 1997. Signal transduction pathways in response to protein misfolding in the extracytoplasmic compartments of E. coli: role of two new phosphoprotein phosphatases PrpA and PrpB. EMBO J. 16:1670-1685.[CrossRef][Medline]
    39. 39
  39. Price, N. E., and M. C. Mumby. 2000. Effects of regulatory subunits on the kinetics of protein phosphatase 2A. J. Biol. Chem. 39:11312-11318.
    40. 40
  40. Rippka, R., J. Deruelles, J. B. Waterbury, M. Herdman, and R. Y. Stanier. 1979. Genetic assignments, strain histories, and properties of pure cultures of cyanobacteria. J. Gen. Microbiol. 111:1-61.
    41. 41
  41. Ruppert, U., A. Irmler, N. Kloft, and K. Forchhammer. 2002. The novel protein phosphatase PphA from Synechocystis PCC 6803 controls dephosphorylation of the signalling protein PII. Mol. Microbiol. 44:855-864.[CrossRef][Medline]
    42. 42
  42. Rusnak, F., L. Yu, and P. Mertz. 1996. Metalloenzymes and signal transduction: the protein serine/threonine phosphatases, a novel class of binuclear metal-containing enzymes. J. Biol. Inorg. Chem. 1:388-396.[CrossRef]
    43. 43
  43. Sheen, J. 1998. Mutational analysis of protein phosphatase 2C involved in abscisic acid signal transduction in higher plants. Proc. Natl. Acad. Sci. USA 95:975-980.[Abstract/Free Full Text]
    44. 44
  44. Shi, L. 2004. Manganese-dependent protein O-phosphatases in prokaryotes and their biological functions. Front. Biosci. 9:1382-1397.[Medline]
    45. 45
  45. Shi, L., K. M. Bischoff, and P. J. Kennelly. 1999. The icfG gene cluster of Synechocystis sp. strain PCC 6803 encodes an Rsb/Spo-like protein kinase, protein phosphatase, and two phosphoproteins. J. Bacteriol. 181:4761-4767.[Abstract/Free Full Text]
    46. 46
  46. Shi, L., M. Potts, and P. J. Kennelly. 1998. The serine, threonine, and/or tyrosine-specific protein kinases and protein phosphatases of prokaryotic organisms: a family portrait. FEMS Microbiol. Rev. 22:229-253.[CrossRef][Medline]
    47. 47
  47. Tsuruta, H., and Y. Aizono. 1999. Enzymatical properties of psychrophilic phosphatase I. J. Biochem. (Tokyo) 125:690-695.[Abstract/Free Full Text]
    48. 48
  48. Tsuruta, H., and Y. Aizono. 2000. Cloning of phosphatase I gene from a psychrophile, Shewanella sp., and some properties of the recombinant enzyme. J. Biochem. (Tokyo) 127:143-149.[Abstract/Free Full Text]
    49. 49
  49. Zhang, C.-C., L. Gonzalez, and V. Phalip. 1998. Survey, analysis and genetic organization of genes encoding eukaryotic-like signaling proteins by a cyanobacterial genome. Nucleic Acids Res. 26:3619-3625.[Abstract/Free Full Text]
    50. 50
  50. Zhang, Z., S. Zhao, S. Deans-Zirattu, G. Bai, and E. Y. C. Lee. 1993. Mutagenesis of the catalytic subunit of rabbit muscle protein phosphatase-1. Mol. Cell. Biochem. 127/128:113-119.

Journal of Bacteriology, September 2005, p. 5877-5884, Vol. 187, No. 17
0021-9193/05/$08.00+0     doi:10.1128/JB.187.17.5877-5884.2005
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