Defining the Active Site of Schizosaccharomyces pombe C-terminal Domain Phosphatase Fcp1*

Stéphane Hausmann and Stewart ShumanDagger

From the Molecular Biology Program, Sloan-Kettering Institute, New York, New York 10021

Received for publication, December 26, 2002, and in revised form, January 24, 2003

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Fcp1 is an essential protein serine phosphatase that dephosphorylates the C-terminal domain (CTD) of RNA polymerase II. By testing the effects of serial N- and C-terminal deletions of the 723-amino acid Schizosaccharomyces pombe Fcp1, we defined a minimal phosphatase domain spanning amino acids 156-580. We employed site-directed mutagenesis (introducing 24 mutations at 14 conserved positions) to locate candidate catalytic residues. We found that alanine substitutions for Arg223, Asp258, Lys280, Asp297, and Asp298 abrogated the phosphatase activity with either p-nitrophenyl phosphate or CTD-PO4 as substrates. Structure-activity relationships were determined by introducing conservative substitutions at each essential position. Our results, together with previous mutational studies, highlight a constellation of seven amino acids (Asp170, Asp172, Arg223, Asp258, Lys280, Asp297, and Asp298) that are conserved in all Fcp1 orthologs and likely comprise the active site. Five of these residues (Asp170, Asp172, Lys280, Asp297, and Asp298) are conserved at the active site of T4 polynucleotide 3'-phosphatase, suggesting that Fcp1 and T4 phosphatase are structurally and mechanistically related members of the DXD phosphotransferase superfamily.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The C-terminal domain (CTD)1 of the largest subunit of RNA polymerase II (pol II) is composed of a tandemly repeated heptapeptide motif (consensus sequence YSPTSPS). The CTD undergoes a cycle of extensive phosphorylation and dephosphorylation at positions Ser5 and Ser2, which is coordinated with the transcription cycle and is responsive to stress and developmental cues. A general picture of CTD phosphorylation dynamics has emerged in which dephosphorylated pol II is recruited to the preinitiation complex and CTD phosphorylation ensues shortly after the elongation complex is established (1). The phosphorylation state of the CTD is apparently remodeled during its transit along the gene, the purpose of which may be to control the ingress and egress of the mRNA processing assemblies that modify the nascent transcript (1-3). Wholesale dephosphorylation of the CTD may be required to recycle pol II for a new round of initiation.

The enzyme Fcp1 is the major protein serine phosphatase responsible for removing phosphates from the CTD (4-11). Fcp1 orthologs are present in all known eukaryal proteomes, and the enzyme is essential for cell viability in budding and fission yeast (7,10). Fcp1 catalyzes the metal-dependent hydrolysis of phosphoserine from the CTD in the context of the pol II elongation complex or isolated pol II and, in the case of Schizosaccharomyces pombe Fcp1, from synthetic CTD phosphopeptide substrates. Fcp1 also hydrolyzes the nonspecific substrate p-nitrophenyl phosphate. The mechanism of the Fcp1 reaction is not known.

We are undertaking via site-directed mutagenesis and biochemical methods to dissect the domain structure of S. pombe Fcp1 and localize the essential constituents of the active site. We initially characterized catalytically active deletion mutants Fcp1(140-723) and Fcp1(1-580) and identified two acidic side chains, Asp170 and Asp172, as essential for phosphatase activity (11). Here we delineate a minimal phosphatase catalytic domain, Fcp1(156-580). To map the active site, we introduced alanine and conservative mutations at 14 residues of S. pombe Fcp1 that are invariant in Fcp1 orthologs from other eukarya (see Fig. 1). We thereby identified five new residues that are essential for catalysis and likely comprise the phosphatase active site. Our results consolidate the hypothesis that Fcp1 belongs to the DXDXT family of phosphotransferases (9, 11, 12). A plausible catalytic mechanism for Fcp1 is suggested based on the concordance of mutational data for Fcp1 and the polynucleotide 3'-phosphatase domain of bacteriophage T4 polynucleotide kinase.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Fcp1 Mutants-- Amino acid substitution mutations (and diagnostic restriction sites) were introduced into the fcp1+ cDNA by the two-stage overlap extension method as described previously (11). pET-Fcp1 was used as the template for the first-stage amplification. The mutated full-length cDNAs generated in the second-stage amplification were digested with NcoI and BamHI and then inserted into pET16m. The inserts of the resulting pET-Fcp1* plasmids were sequenced completely to confirm the desired mutations and exclude the acquisition of unwanted changes during amplification or cloning. Deletion mutants Fcp1(140-580), Fcp1(148-580), Fcp1(156-580), and Fcp1(162-580) were constructed by PCR amplification with sense-strand primers that introduced an NcoI restriction site and a methionine codon in lieu of the codons for Gly139, Glu147, Asn155, or Gln161 and an antisense-strand primer that introduced a stop codon in lieu of the codon for Pro581 and a BamHI site 3' of the new stop codon. Additional C-terminal deletions were constructed by PCR amplification with antisense primers that introduced stop codons in place of the codons for Val505 or Trp561 and a BamHI site 3' of the new stop codon. The PCR products were digested with NcoI and BamHI and then inserted into pET16m. The pET-Fcp1* and pET-Fcp1Delta plasmids were introduced into Escherichia coli BL21(DE3)-RIL, and the mutant Fcp1 proteins were purified from soluble bacterial lysates nickel-nitrilotriacetic acid-agarose affinity chromatography as described previously for wild-type Fcp1 (11). Protein concentrations were determined by using the BioRad dye reagent with bovine serum albumin as the standard.

Phosphatase Assay-- Reaction mixtures (100 µl) containing 50 mM Tris acetate (pH 5.5), 10 mM MgCl2, 10 mM p-nitrophenyl phosphate (pNØP), and Fcp1 as specified were incubated for 30 min at 37 °C. The reactions were quenched by adding 900 µl of 1 M sodium carbonate. Release of p-nitrophenol (pNØ) was determined by measuring A410 and interpolating the value to a pNØ standard curve.

CTD Phosphatase Assay-- Reaction mixtures (25 µl) containing 50 mM Tris acetate (pH 5.5), 10 mM MgCl2, 25 µM CTD-PO4 peptide (an N-terminal biotinylated 28-aa CTD phosphopeptide composed of four tandem YSPTSPS repeats containing phosphoserines at positions 2 plus 5), and Fcp1 as specified were incubated for 60 min at 37 °C. The reactions were quenched by adding 1 ml of malachite green reagent (BIOMOL GREEN reagent, purchased from BIOMOL Research Laboratories, Plymouth Meeting, PA). Phosphate release was determined by measuring A620 and interpolating the value to a phosphate standard curve.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Alanine Scanning Mutagenesis of Fcp1-- A goal of the present study was to extend the map of the active site of Fcp1 by identifying individual amino acid functional groups required for the phosphatase reaction. To do this, we introduced single alanine changes at 14 amino acids of the S. pombe Fcp1 polypeptide that are invariant in the putative phosphatase catalytic domains of other Fcp1 orthologs (Fig. 1). We focused in particular on basic and acid residues, which we viewed as potential ligands for the phosphoserine substrate or the divalent cation cofactor. We also mutated the two invariant histidines within the phosphatase domain, His177 and His240, which we considered as possible general acid catalysts. The pH optimum of 5.5 for the phosphatase activity of S. pombe Fcp1 and the sharp decrement in activity seen at >= pH 7.5 had suggested the existence of a protonated functional group with a pKa near neutrality (11).


View larger version (92K):
[in this window]
[in a new window]
 
Fig. 1.   Conservation of Fcp1 primary structure. The amino acid sequence of S. pombe Fcp1 from residues 140-326 is aligned to the sequences of Fcp1 orthologs from Saccharomyces cerevisiae (Sce), Leptosphaeria maculans (Lma), Neurospora crassa (Ncr), Aspergillus nidulans (Ani), Candida albicans (Cal), Xenopus laevis (Xla), Drosophila melanogaster (Dme), Anopheles gambiae (Aga), Homo sapiens (Hsa), Caenorhabditis elegans (Cel), Dictyostelium discoideum (Ddi), and Encephalitozoon cuniculi (Ecu). Gaps in the alignment are indicated by dashes. The essential Asp170 and Asp172 residues of the DXDXT phosphatase motif of S. pombe Fcp1 are shaded. The 14 amino acids targeted for mutational analysis in the present study are indicated by dots (). Newly defined essential residues of S. pombe Fcp1 are shaded. Included at the bottom of the alignment are two conserved motifs that comprise the 3'-phosphatase active site of bacteriophage T4 Pnk.

Full-length wild-type Fcp1 and the 14 Fcp1-Ala mutants were produced in bacteria as His10-tagged fusions and purified from soluble bacterial extracts by nickel-agarose chromatography. SDS-PAGE analysis of the imidazole eluate fractions showed that the preparations were highly enriched with respect to the His-Fcp1 polypeptide (Fig. 2A).


View larger version (55K):
[in this window]
[in a new window]
 
Fig. 2.   Purification of Fcp1-Ala mutants and effects on phosphatase activity. A, aliquots (7 µg) of the nickel-agarose preparations of wild-type (WT) Fcp1 and the indicated Fcp1-Ala mutants were analyzed by SDS-PAGE. Polypeptides were visualized by staining the gels with Coomassie Brilliant Blue dye. A scan of the stained gels is shown. The positions and sizes (in kDa) of marker proteins are indicated on the left. B, reaction mixtures containing 50 mM Tris acetate (pH 5.5), 10 mM MgCl2, 10 mM pNØP, and WT or mutant Fcp1 proteins as specified were incubated for 30 min at 37 °C. pNØ release is plotted as a function of input protein.

Alanine Mutational Effects on Phosphatase Activity-- Initial characterization of the phosphatase activity of the purified recombinant Fcp1-Ala proteins was performed using 10 mM pNØP as a substrate. The pNØ reaction product was detected via its absorbance at 410 nM. The extent of conversion of pNØP to pNØ was directly proportional to the concentration of the recombinant wild-type protein, which released 22 nmol of pNØ per µg of protein in 30 min (Fig. 2B). We defined a significant mutational effect as one that elicits at least a 10-fold decrement in specific activity compared with wild-type Fcp1. The specific activities of the R223A, K280A, D297A, and D298A mutants were <0.5% of the activity of wild-type Fcp1, whereas the D258A mutant was 3% as active as wild-type Fcp1 (Fig. 2B and Table I). We conclude that the Arg223, Asp258, Lys280, Asp297, and Asp298 side chains are essential for Fcp1 activity. Other Fcp1-Ala proteins either retained near wild-type phosphatase activity (H177A, H240A, R271A, R299A, and D301A) or displayed modest reductions in activity (E238A, R267A, and D272A) that did not meet our threshold of significance (Fig. 2B and Table I). We surmise that the His177, Glu238, His240, Arg267, Arg271, Asp272, Arg299, and Asp301 side chains do not contribute significantly to catalysis of the phosphatase reaction. A notable finding was that the D323A mutation resulted in a 4-fold increase in phosphatase activity over wild-type Fcp1 (Fig. 2B and Table I). This stimulation was reproducible for multiple independent preparations of the D323A protein (data not shown). Determination of the steady state kinetic parameters for wild-type Fcp1 (Km = 18 mM pNØP; kcat = 3.7 s-1) and the D323A mutant (Km = 7.1 mM pNØP; kcat = 8.3 s-1) showed that the removal of the Asp323 side chain elicited ~2-fold increases in the Km of Fcp1 for pNØP and the turnover number for pNØP hydrolysis. The kcat/Km ratio for the D323A mutant was 5.6-fold greater than that of wild-type Fcp1.


                              
View this table:
[in this window]
[in a new window]
 
Table I
Summary of mutational effects on Fcp1 phosphatase activity
The specific activity shown for each Fcp1 mutant in dephosphorylating pNempty P is the average of two independent titration experiments. The activities are normalized to that of wild-type Fcp1 (defined as 100%).

Mutational Effects on CTD Phosphatase Activity-- CTD phosphatase activity of the Fcp1-Ala mutants was measured by the release of Pi from a synthetic 28-aa phosphopeptide consisting of four tandem repeats of the CTD heptad sequence (YSPTSPS) in which all Ser2 and Ser5 residues are Ser-PO4 (11). The enzyme preparations were assayed in parallel; the reaction mixtures contained 25 µM CTD phosphopeptide and 2.5 µg of input Fcp1, an amount sufficient for saturating levels of Pi release by wild-type Fcp1 (see Fig. 3B). Conducting the screening assays in this fashion highlighted the most severe mutational effects on CTD phosphatase activity (Fig. 3A). The four Fcp1 mutants that were grossly defective in CTD dephosphorylation, R223A, K280A, D297A, and D298A, are the same mutants that displayed <0.5% of wild-type activity in hydrolyzing pNØP. D258A also displayed reduced CTD phosphatase activity in the screening assay (Fig. 3A). A titration experiment showed that the specific activity of D258A in dephosphorylating the CTD phosphopeptide was 3% of the specific activity of wild-type Fcp1 (Fig. 3B). This value is in accord with the effects of the D258A mutation on hydrolysis of the nonspecific substrate pNØP (Table I). The nine Fcp1-Ala proteins that displayed near wild-type CTD phosphatase activity in the screening assay were the same as those that retained activity in hydrolyzing pNØP. We infer that: (i) a single active site is responsible for catalyzing the hydrolysis of pNØP and the phospho-CTD and (ii) Arg223, Asp258, Lys280, Asp297, and Asp298 are likely to be involved directly in catalysis.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 3.   CTD phosphatase activity. A, reaction mixtures containing 25 µM CTD-PO4 peptide and 2.5 µg of the indicated Fcp1 protein were incubated for 60 min at 37 °C. B, reaction mixtures containing 25 µM CTD-PO4 peptide and wild-type or mutant Fcp1 proteins as specified were incubated for 60 min at 37 °C. Pi release is plotted as a function of input enzyme.

The specific activity of wild-type Fcp1 on the CTD phosphopeptide substrate (8.2 nmol of Pi released in 60 min per µg of protein; Fig. 3B) translates into a turnover number of 0.19 s-1. Parallel titration of the D323A mutant showed that it was 74% as active as wild-type Fcp1 with the CTD phosphopeptide substrate (Fig. 3B). The nearly wild-type CTD phosphatase activity of D323A contrasts with its 410% activity with pNØP (Fig. 2B). We surmise that Asp323 is not a genuine modulator of Fcp1 activity but rather that the carboxylate is a specific impediment to the binding and hydrolysis of pNØP, which is a nonphysiological substrate with low affinity for the Fcp1 active site.

Structure-Activity Relationships at Essential Residues-- To further evaluate the contributions of Arg223, Asp258, Lys280, Asp297, and Asp298 to the phosphatase reaction, we tested the effects of conservative substitutions. Aspartate was replaced by asparagine and glutamate, arginine by lysine and glutamine, and lysine by arginine and glutamine. The R223K, R223Q, D258E, D258N, K280R, K280Q, D297E, D297N, D298E, and D298N proteins were purified from soluble bacterial extracts by nickel-agarose chromatography (Fig. 4A).


View larger version (42K):
[in this window]
[in a new window]
 
Fig. 4.   Effects of conservative substitutions on phosphatase activity. A, aliquots (7 µg) of the indicated nickel-agarose Fcp1 preparations were analyzed by SDS-PAGE. A scan of the Coomassie Blue-stained gel is shown. The positions and sizes (in kDa) of marker proteins are indicated on the left. B, reaction mixtures containing 50 mM Tris acetate (pH 5.5), 10 mM MgCl2, 10 mM pNØP, and WT or mutant Fcp1 proteins as specified were incubated for 30 min at 37 °C. pNØ release is plotted as a function of input protein.

Hydrolysis of pNØP was measured as a function of enzyme concentration for wild-type Fcp1 and the 10 conservative mutants (Fig. 4B). We found that R223K and R223Q were just as defective as the alanine mutant, with <1% of wild-type phosphatase activity. Thus, arginine is specifically required at position 223. K280R and K280Q were as defective as K280A (<0.5% activity), indicating a stringent requirement for lysine at position 280. Conservative replacement of Asp297 with Asn or Glu elicited a severe catalytic defect comparable to that seen with D297A (<1% of wild-type activity). These data establish a requirement for a carboxylate residue at position 297, and they indicate a steric constraint on the distance from the main chain to the carboxylate, whereby the additional methylene group of glutamate is detrimental. Replacing Asp298 with Asn resulted in a modest gain of function (to 5% of wild-type activity) compared with the D298A mutant (<0.5%), but substitution by Glu had no salutary effect. These results indicate the importance of the carboxylate at position 298 and a steric constraint on the main chain to carboxylate distance. Glutamate substitution for Asp258 revived phosphatase activity to above our threshold of significant mutational effect (to 18% activity, compared with 3% for D258A), whereas the D258N mutant was still significantly compromised, with 7% of wild-type activity. Thus, the carboxylate at position 258 is essential, and the enzyme is flexible in its accommodation of the glutamate side chain.

Defining a Minimal Fcp1 Phosphatase Domain-- We showed previously that deletion mutants Fcp1(140-723) and Fcp1(1-580) retained full phosphatase activity in vitro (11). To delineate a minimal catalytic domain, we combined the N- and C-terminal deletions to generate the doubly truncated derivative Fcp1(140-580). Purified recombinant Fcp1(140-580) displayed full activity in hydrolyzing pNØP (Fig. 5B) and in dephosphorylating the synthetic CTD phosphopeptide (Fig. 3B). To finely delineate the N-terminal margin of the catalytic domain, we produced and purified a new set of three incrementally truncated derivatives: Fcp1(148-580), Fcp1(156-580), and Fcp1(162-580). SDS-PAGE analysis revealed the expected serial increases in their electrophoretic mobility (Fig. 5A). The recombinant proteins were assayed for phosphatase activity with pNØP (Fig. 5B). We found that deletion of amino acids 140-147 reduced phosphatase-specific activity to 72% that of Fcp1(140-580) (Fig. 5B). Further removal of amino acids 148-155 reduced the phosphatase activity to 28% of Fcp1(140-580). Because these truncations did not meet our 10-fold threshold, we surmise that the segment from 140 to 155 is not strictly essential for catalysis. However, the next incremental deletion mutant, Fcp1(162-580), was only 1.7% as active as Fcp1(140-580), indicating that the segment from 156 to 161 is essential. These results delineate a requirement for ~15 amino acids upstream of the first essential aspartate (Asp170) of the conserved DXDXT motif.


View larger version (27K):
[in this window]
[in a new window]
 
Fig. 5.   Deletion analysis defines a minimal Fcp1 phosphatase domain. A, aliquots (7 µg) of the indicated Fcp1 deletion mutants were analyzed by SDS-PAGE. A Coomassie Blue-stained gel is shown. The positions and sizes (in kDa) of marker proteins are indicated on the left. B, reaction mixtures containing 50 mM Tris acetate (pH 5.5), 10 mM MgCl2, 10 mM pNØP, and either WT or mutant proteins as specified were incubated for 30 min at 37 °C. pNØ release is plotted as a function of input protein.

We showed previously that the C-terminal deletion mutant Fcp1(1-486) was catalytically inert (11). To better delineate a distal margin for the catalytic domain of Fcp1, we introduced new C-terminal deletions into the NDelta 139 Fcp1 derivative. We found that purified recombinant Fcp1(140-505) had no detectable phosphatase activity (Fig. 5). A longer derivative, Fcp1(140-560), was also catalytically inert (not shown). We surmise that the protein segment from amino acids 560-580 is essential.

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Here we employed site-directed mutagenesis to define a minimal catalytic domain and identify five new essential amino acids of the S. pombe CTD phosphatase Fcp1. As discussed in detail below, our results suggest structural and mechanistic similarities between Fcp1 and the 3'-phosphatase domain of T4 polynucleotide kinase (13-16), and they consolidate the suggestion (9, 11, 12) that Fcp1 is a member of a phosphatase superfamily defined by the DXDXT motif.

Deletion analysis showed that large segments could be deleted simultaneously from the N and C termini of S. pombe Fcp1 without loss of phosphatase activity. The apparent minimal catalytic domain is Fcp1(156-580), which consists of an Fcp1 homology domain (shown in Fig. 1) linked to a downstream BRCT domain. An earlier in vivo deletion analysis of S. cerevisiae Fcp1 showed that removal of the N-terminal region upstream of the Fcp1 homology domain did not affect cell viability nor did removal of the C-terminal segment downstream of the BRCT domain. However, simultaneous deletion of both N- and C-terminal segments was lethal in vivo (17). The present biochemical analysis of S. pombe Fcp1 shows clearly that the flanking N- and C-terminal domains are not essential for phosphatase activity, either with pNØP or CTD-PO4. It is therefore likely that the seemingly redundant in vivo roles of the N- and C-terminal domains of yeast Fcp1 have more to do with ancillary functions, such as protein-protein interactions (17, 18) and the pol II elongation-stimulation activity of Fcp1 (9, 19), than with chemical catalysis or CTD-PO4 recognition.

Fcp1 orthologs from diverse species display extensive sequence similarity in the region spanning S. pombe Fcp1 residues 140-326 (Fig. 1). A short conserved peptide motif (167LIVDLDQTII176 in S. pombe Fcp1) located near the N-terminal margin of the minimal catalytic domain corresponds to the signature sequence of the DXDXT family of metal-dependent phosphohydrolases and phosphotransferases (20-22). Several DXDXT family members have been shown to act via an acyl-phosphoenzyme intermediate in which the phosphate is linked to the first aspartate in the DXDXT motif (20, 23-25). Given that the two aspartates in the DXDXT element of Fcp1 were found previously to be essential for Fcp1 phosphatase activity (11, 12), it was suggested that Fcp1 is a member of the DXDXT superfamily. However, a DXD motif is also a defining feature of the "Toprim" domains found in another class of phosphoryl transfer enzymes (including DNA topoisomerases IA, and II and DNA primases), which do not form acyl-phosphate intermediates (26-28). In the Toprim enzymes, the DXD motif coordinates a divalent cation.

In the absence of an atomic structure for Fcp1, we rely on primary structure and mutational analysis for clues to which family Fcp1 might belong. However, amino acid sequence searches reveal little similarity, exclusive of the DXD motif itself, between Fcp1 and the several DXDXT family members for which atomic structures are available. Nonetheless, instructive clues emerged from a comparison of the mutational results for Fcp1 to those obtained for the polynucleotide 3'-phosphatase domain of T4 polynucleotide kinase (Pnk), which also contains a signature DXDXT motif (Fig. 1). The active site of the Pnk phosphatase domain has been mapped by extensive site-directed mutagenesis. The 10 side chains essential for catalysis include six aspartates, two arginines, one lysine, and one serine (13-15).2 The amino acid composition of the Pnk active site is generally similar to the constellation of essential residues of S. pombe Fcp1, which consists, to date, of five aspartates, one arginine, and one lysine. The most upstream essential residues in the Pnk and Fcp1 sequences are the DXDXT motif aspartates (165DVDGT169 in Pnk; 170DLDQT174 in S. pombe Fcp1). Indeed, the N termini of Pnk and Fcp1 can be deleted to within ~15 amino acids of the first catalytic aspartate without significant loss of phosphatase activity (15). The distal ends of the Pnk and Fcp1 active sites map to a conserved motif defined by two vicinal essential aspartates, which are preceded by a hydrophobic cluster and then followed by an conserved arginine and a second cluster of hydrophobic residues (Fig. 1). The sequence of this "DD" motif in Pnk is 274LAIDDRTQVVEM285, and its putative equivalent in Fcp1 is 294VVIDDRGDVWDW305 (Fig. 1). Note that the length of the polypeptide segment separating the DXD and DD motifs is quite similar in Pnk and Fcp1. The occurrence of two signature motifs with similar order and spacing plus the concordant mutational findings for the two aspartates of the DD motif of Fcp1 and Pnk (14)2 leads us to posit that the Fcp1 and Pnk phosphatase domains are structurally and mechanistically related. The crystal structure of Pnk shows that the fold of the Pnk1 phosphatase domain resembles that of the exemplary DXDXT family member, Methanococcus jannaschii phosphoserine phosphatase (PSP) (16).3 Thus, we infer that Fcp1 is also a bona fide member of the DXDXT superfamily.

Elegant crystallographic snapshots have been obtained for PSP at sequential intermediate states of the catalytic cycle, which reveal the role of the DXD motif in catalysis (24, 29). The proximal Asp plays two key functions: Odelta 1 attacks the phosphorus of the substrate to form the acyl-phosphate intermediate, while Odelta 2 is part of the octahedral coordination complex of the essential divalent cation cofactor. The distal aspartate of the DXD motif of PSP acts as a general acid-base catalyst: it donates a proton to the serine leaving group during the first phosphoryl transfer step, and it abstracts a proton from the attacking water during the second hydrolytic step. We infer that Fcp1 employs a similar mechanism of acid-base catalysis via an acyl-phosphate intermediate. We had inferred previously from the pH profile of S. pombe Fcp1 the existence of at least two different functional groups at which the protonation state has a strong impact on phosphatase activity (11). We would now attribute the falloff in catalysis between pH 5.5/5.0 and pH 4.5 to protonation of the proximal Asp nucleophile of the DXD active site motif. To explain the decline in activity as the pH was increased above neutrality, we had invoked a requirement for protonated functional group on Fcp1 enzyme and discussed a potential histidine general acid with a presumptive pKa of ~6.5. The present mutational analysis vitiates the idea that either of the two conserved Fcp1 histidines might play such a role. Rather, the PSP structure and the sum of the mutational data focus attention on Asp172 of the Fcp1 DXD motif as a potential source of the activity decrement of S. pombe Fcp1 at alkaline pH, i.e. via deprotonation of the Asp general acid.

The PSP crystal structure highlights a lysine side chain that interacts with the phosphate in the ground state and stabilizes the pentacoordinate phosphorane transition state (24, 29). The equivalent essential lysine in the phosphatase domain of Pnk (Lys258) is located 19 aa upstream of the DD motif of Pnk. As shown here, Fcp1 also has an essential lysine (Lys280) situated 17 aa upstream of its DD motif. The structure-activity relationships at Pnk Lys258 and Fcp1 Lys280 are concordant with each other (i.e. neither arginine nor glutamine could replace the essential lysine in Pnk or Fcp1), and the functional data are consistent with the monovalent phosphate contact seen for the catalytic lysine in the PSP crystal.

It is notable that PSP and most other proteins classified as DXDXT phosphatases have no precise counterpart in their primary structures of the DD motif present in Pnk and Fcp1. Rather, the structural superposition of PSP and Pnk shows that two aspartate side chains of PSP within a 167DXXXD171 motif occupy positions at the active site that overlay with the two aspartates of the DD motif of Pnk (16). Note that the proximal Asp277 of the 277DD278 dipeptide of Pnk superimposes on the distal Asp171 of the PSP 167DXXXD171 motif, while the distal Asp288 of Pnk DD overlies the proximal Asp167 of the PSP 167DXXXD171 element. The DXXXD motif was recognized as a signature feature, along with the upstream DXDXT element, of the proteins that were initially felt to comprise the phosphatase superfamily (21, 22). Yet, the structural and mutational data for Pnk and Fcp1 suggest that they belong to a different subgroup of phosphatases within the DXDXT superfamily, which is defined by their DD motifs. The PSP structure shows that the two aspartates of the DXXXD motif comprise part of the divalent cation coordination complex in the active site via direct and water-mediated contacts between the carboxylate oxygens and the metal (24, 29). Thus, we infer that the essential 297DD298 dipeptide of Fcp1 is also concerned with metal binding.

In summary, we have defined seven essential hydrophilic amino acids of S. pombe Fcp1 that are conserved in all known Fcp1 orthologs and likely comprise the phosphatase active site. We identify an essential DD motif that is characteristic of Fcp1 and T4 Pnk. Plausible mechanistic roles are proposed for five of the seven essential side chains of Fcp1, either in nucleophilic attack on Ser-PO4 (Asp170), general acid-base catalysis (Asp172), transition state stabilization (Lys280), or metal coordination (Asp297, Asp298). The functions of the other two essential Fcp1 side chains (Arg223 and Asp258) cannot be surmised with confidence, but, by analogy with Pnk, they may be involved in positioning one or more of the other catalytic side chains at the active site via a network of hydrogen-bonding interactions (14,16). Definitive evaluation of the mechanism proposed here will hinge on crystallization of Fcp1.

    FOOTNOTES

* This work was supported by National Institutes of Health Grant GM52470.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger To whom correspondence should be addressed. Tel.: 212-639-7145; Fax: 212-717-3623; E-mail: s-shuman@ski.mskcc.org.

Published, JBC Papers in Press, January 28, 2003, DOI 10.1074/jbc.M213191200

2 H. Zhu and S. Shuman, unpublished data.

3 L. Wang, S. Shuman, and C. Lima, unpublished data.

    ABBREVIATIONS

The abbreviations used are: CTD, C-terminal domain; pol II, polymerase II; pNØP, p-nitrophenyl phosphate; pNØ, p-nitrophenol; aa, amino acid; Pnk, polynucleotide kinase; PSP, phosphoserine phosphatase; WT, wild type.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

1. Kobor, M. S., and Greenblatt, J. (2002) Biochim. Biophys. Acta 1577, 261-275[Medline] [Order article via Infotrieve]
2. Hirose, Y., and Manley, J. L. (2000) Genes Dev. 14, 1415-1429[Free Full Text]
3. Bentley, D. (2002) Curr. Opin. Cell Biol. 14, 336-342[CrossRef][Medline] [Order article via Infotrieve]
4. Lin, P. S., Marshall, N. F., and Dahmus, M. E. (2002) Prog. Nucleic Acids Res. Mol. Biol. 72, 333-365[Medline] [Order article via Infotrieve]
5. Chambers, R. S., and Dahmus, M. E. (1994) J. Biol. Chem. 269, 26243-26248[Abstract/Free Full Text]
6. Chambers, R. S., and Kane, C. M. (1996) J. Biol. Chem. 271, 24498-24504[Abstract/Free Full Text]
7. Archambault, J., Chambers, R. S., Kobor, M. S., Ho, Y., Cartier, M., Bolotin, D., Andrews, B., Kane, C. M., and Greenblatt, J. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 14300-14305[Abstract/Free Full Text]
8. Archambault, J., Pan, G., Dahmus, G. K., Cartier, M., Marshall, N., Zhang, S., Dahmus, M. E., and Greenblatt, J. (1998) J. Biol. Chem. 273, 27593-27601[Abstract/Free Full Text]
9. Cho, H., Kim, T., Mancebo, H., Lane, W. S., Flores, O., and Reinberg, D. (1999) Genes Dev. 13, 1540-1552[Abstract/Free Full Text]
10. Kimura, M., Suzuki, H., and Ishihama, A. (2002) Mol. Cell. Biol. 22, 1577-1588[Abstract/Free Full Text]
11. Hausmann, S., and Shuman, S. (2002) J. Biol. Chem. 277, 21213-21220[Abstract/Free Full Text]
12. Kobor, M. S., Archambault, J., Lester, W., Holstege, F. C. P., Gileadi, O., Lansma, D. B., Jennings, E. G., Kouyoumdjian, F., Davidson, A. R., Young, R. A., and Greenblatt, J. (1999) Mol. Cell. 4, 55-62[Medline] [Order article via Infotrieve]
13. Wang, L. K., and Shuman, S. (2001) J. Biol. Chem. 276, 26868-26874[Abstract/Free Full Text]
14. Wang, L. K., and Shuman, S. (2002) Nucleic Acids Res. 30, 1073-1080[Abstract/Free Full Text]
15. Wang, L. K., Lima, C. D., and Shuman, S. (2002) EMBO J. 21, 3873-3880[Abstract/Free Full Text]
16. Galburt, E. A., Pelletier, J., Wilson, G., and Stoddard, B. L. (2002) Structure 10, 1249-1260[CrossRef][Medline] [Order article via Infotrieve]
17. Kobor, M. S., Simon, L. D., Omichinski, J., Zhong, G., Archambault, J., and Greenblatt, J. (2000) Mol. Cell. Biol. 20, 7438-7449[Abstract/Free Full Text]
18. Chambers, R. S., Wang, B. Q., Burton, Z. F., and Dahmus, M. E. (1995) J. Biol. Chem. 270, 14962-14969[Abstract/Free Full Text]
19. Mandal, S. S., Cho, H., Kim, S., Cabane, K., and Reinberg, D. (2002) Mol. Cell. Biol. 22, 7543-7552[Abstract/Free Full Text]
20. Collet, J. F., Stroobant, V., Pirard, M., Delpierre, G., and Van Schaftingen, E. (1998) J. Biol. Chem. 273, 14107-14112[Abstract/Free Full Text]
21. Thaller, M. C., Schippa, S., and Rossolini, G. M. (1998) Protein Sci. 7, 1647-1652[Abstract/Free Full Text]
22. Aravind, A., Galperin, M. Y., and Koonin, E. V. (1998) Trends Biochem. Sci. 23, 127-129[CrossRef][Medline] [Order article via Infotrieve]
23. Collet, J. F., Stroobant, V., and Van Schaftingen, E. (1999) J. Biol. Chem. 274, 33985-33990[Abstract/Free Full Text]
24. Cho, H. S., Wang, W., Kim, R., Yokota, H., Damo, S., Kim, S. H., Wemmer, D. E., Kustu, S., and Yan, D. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 8525-8530[Abstract/Free Full Text]
25. Allegrini, S., Scaloni, A., Ferrara, L., Pesi, R., Pinna, P., Sgarella, F., Camici, M., Eriksson, S., and Tozzi, M. G. (2002) J. Biol. Chem. 276, 33526-33532[Medline] [Order article via Infotrieve]
26. Aravind, L., Leipe, D. D., and Koonin, E. V. (1998) Nucleic Acids Res. 26, 4205-4213[Abstract/Free Full Text]
27. Nichols, M. D., DeAngeli, K., Keck, J. L., and Berger, J. M. (1999) EMBO J. 18, 6177-6188[Abstract/Free Full Text]
28. Keck, J. L., Roche, D. D., Lynch, S., and Berger, J. M. (2000) Science 287, 2482-2486[Abstract/Free Full Text]
29. Wang, W., Cho, H. S., Kim, R., Jancarik, J., Yokota, H., Nguyen, H. H., Grigoriev, I. V., Wemmer, D. E., and Kim, S. H. (2002) J. Mol. Biol. 319, 421-431[CrossRef][Medline] [Order article via Infotrieve]


Copyright © 2003 by The American Society for Biochemistry and Molecular Biology, Inc.