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J. Biol. Chem., Vol. 279, Issue 29, 30459-30468, July 16, 2004
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**
From the
Department of Molecular Pharmacology, Albert Einstein College of Medicine, Bronx, New York 10461, ¶Molecular Probes, Inc., Eugene, Oregon 97402, the Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, Texas 75390, and the ||Department of Biochemistry, Purdue University, West Lafayette, Indiana 47907
Received for publication, February 27, 2004 , and in revised form, May 4, 2004.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTS AND DISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTS AND DISCUSSIONREFERENCES |
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Despite their structural similarities, the biological function of Cdc14 phosphatases may vary between species. Thus, in S. cerevisiae, Cdc14 is absolutely required for mitotic exit. In contrast, the Cdc14 homolog Clp1/Flp1 is not required for mitotic exit in Schizosaccharomyces pombe, but rather is mainly involved in regulating cytokinesis and affects the timing of entry into mitosis (10, 11). In addition, CeCdc14 phosphatase is required for cytokinesis in the Caenorhabditis elegans embryo (12). The functional roles for the two human Cdc14 homologs, hCdc14A and hCdc14B, have not been fully defined, though depletion of hCdc14A by RNAi causes a variety of mitotic defects, including cytokinesis failure (13). It is possible that Cdc14 phosphatases may antagonize CDK activity by dephosphorylation of different substrates and regulate distinct cell-cycle transitions in different species, as CDK activity is important for many different cell cycle processes (14).
Although the role of the Cdc14 phosphatase in cell cycle has been studied extensively, relatively little is known about its catalytic mechanism. Given their critical roles in mitotic exit and cytokinesis, the Cdc14 phosphatases have been implicated as potential therapeutic targets for anticancer drug development. Thus it is important to define Cdc14 active site substrate specificity and identify residues that are essential for Cdc14 catalysis. Interestingly, the Cdc14 phosphatases contain the active site sequence CX5R, which is a hallmark for the protein tyrosine phosphatase (PTP) superfamily. The PTPs are a diverse family of enzymes that includes tyrosine-specific, dual specificity, Cdc25, as well as low molecular weight phosphatases (15). Detailed enzymological studies suggest that the PTPs employ a conserved two-step mechanism (16), shown in Scheme 1, in which E is the enzyme, 
, the substrate, 
, the enzyme-substrate Michaelis complex, E-P, the phosphoenzyme intermediate, and ArOH, the leaving group phenol. In this mechanism, the side chain of the active site Cys residue serves as a nucleophile to accept the phosphoryl group from the substrate to form E-P. The active site Arg makes bidentate hydrogen bonds with the phosphoryl group in the substrate through its guanidinium group and plays an important role in both substrate binding and transition state stabilization during catalysis. To facilitate substrate turnover, PTPs also utilize an Asp residue, which acts as a general acid by protonating the ester oxygen of the leaving group, thus greatly enhancing the rate of E-P formation. In the second step, E-P hydrolysis occurs by the attack of a nucleophilic water molecule assisted by the same Asp, functioning as a general base, with subsequent release of the free enzyme and inorganic phosphate. Aside from the conserved active site CX5R motif, there are little sequence similarities between Cdc14 and other members of the PTP superfamily. Consequently, it is not known whether other residues also participate in Cdc14 catalysis. This is in contrast to other PTPs for which a number of conserved, catalytically important residues have been identified and characterized (16). In the following, we describe a kinetic investigation of Cdc14 substrate specificity with both small molecule aryl phosphates and a physiological protein substrate hCdh1. We also describe results from site-directed mutagenesis experiments in combination with steady-state and pre-steady-state kinetic analyses that lead to the identification of several catalytically important residues including the general acid/base in Cdc14 catalysis.
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTS AND DISCUSSIONREFERENCES |
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-naphthyl phosphate, and 3-O-methylfluorescein phosphate (OMFP) were obtained from Sigma. 6,8-Difluoro-4-methylumbelliferyl phosphate (DiFMUP) was purchased from Molecular Probes. 2-Chloro-4-nitrophenyl phosphate, 4-cyanophenyl phosphate, 4-trifluoromethylphenyl phosphate, and 4-chlorophenyl phosphate were synthesized as described (17). 8-Fluoro-4-methylumbelliferyl phosphate (8-FMUP), 6-fluoro-4-methylumbelliferyl phosphate (6-FMUP), and 5-fluoro-4-methylumbelliferyl phosphate (5-FMUP) were prepared as described (18).
Expression and Purification of Cdc14 and Mutant Proteins—The cDNA for budding yeast Cdc14 was subcloned into vector pET-15b. The Cdc14 mutants were obtained using the QuikChange site-directed mutagenesis kit from Stratagene. The sequences of the wild-type and mutant Cdc14s were confirmed by DNA sequencing. Wild-type and mutant Cdc14s were transformed in Escherichia coli BL21(DE3) cells and induced by 0.3 mM isopropyl-1-thio--D-galactopyranoside at room temperature for 20 h. The His6-tagged proteins were purified by Ni-NTA resin based upon the procedure described by Qiagen. Proteins were judged to be at least 90% pure by SDS-PAGE.
Steady-state Kinetics—All assays with small molecule substrates were performed at 30 °C. The following buffers were used: 50 mM succinate (pH 5.0 
6.0), 50 mM 3,3-dimethyl glutarate (pH 6.0 7.2), and 100 mM Tricine (pH 7.2 9.0). In addition, each buffer also contained 1 mM EDTA, 2 mM dithiothreitol and was adjusted to an ionic strength of 150 mM with sodium chloride. To ensure initial rate conditions, the product versus time progress curves were routinely checked for linearity at different substrate and enzyme concentrations. In all cases, the enzyme concentration was at least 100-fold lower than that of the substrate, and the quantity of substrate consumed was less than 10% of the initial value, so that the steady-state assumption was fulfilled. There was no product inhibition under these conditions. For substrates 2-chloro-4-nitrophenyl phosphate and pNPP, assay mixtures of 200 µl in total volume were set up in 1.1-ml polypropylene tubes from Marsh Biomedical. Reactions were started by the addition of an appropriate amount of Cdc14. The reaction mixtures were quenched with 50 µl of 5 M sodium hydroxide, and the absorbance at 405 nm was read using a plate reader. For substrates 5-FMUP, 8-FMUP, 6-FMUP, MUP, -naphthyl phosphate, 4-cyanophenyl phosphate, 4-trifluoromethylphenyl phosphate, 4-chlorophenyl phosphate, phenyl phosphate, the reaction mixtures were quenched with 10% trichloroacetic acid, and the amount of inorganic phosphate released was quantitated based on the method of Black and Jones (19). For the substrates 6,8-DiFMUP, 5-FMUP, 8-FMUP, 6-FMUP, MUP, continuous assays were performed. The reaction (1-ml total volume) was set up in a 1-ml quartz cuvette. The fluorescence of the product was monitored by a PerkinElmer Life Sciences 50B Luminescence spectrometer. When the substrate concentrations were well below the Km, the kcat/Km values were obtained from a linear least square fit of the plot of initial rate versus substrate concentration using Equation 1,
 | (Eq. 1) |
 | (Eq. 2) |
 | (Eq. 3) |
pH Dependence Studies—The kcat and kcat/Km values for the Cdc14-catalyzed hydrolysis of pNPP and DiFMUP were determined as a function of pH at 30 °C. To fit the kcat/Km versus pH data for the hydrolysis of pNPP, Equation 4 was used,
 | (Eq. 4) |
 | (Eq. 5) |
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Pre-Steady-state Kinetic Study—Pre-steady-state kinetic measurements of the wild-type Cdc14 and its D253A mutant-catalyzed hydrolysis of DiFMUP were conducted at pH 7.0 and 30 °C. Experiments were performed by mixing the enzyme and substrate in a temperature-controlled Applied Photophysics SX.18MV stopped-flow spectrofluorimeter. The excitation wavelength was 390 nm, and the emission wavelength was selected using a 420-nm cutoff filter. The reaction was monitored by the increase in fluorescence of the DiFMU product. A standard curve was generated by measuring the fluorescence of solutions of DiFMU that were reconstituted in the assay buffer. Data were collected and fit to Equation 7.
 | (Eq. 7) |
 | (Eq. 8) |
 | (Eq. 9) |
Preparation of Phosphorylated hCdh1 Protein—A procedure similar to that described previously (7, 22) was used for the preparation of phosphorylated hCdh1. Briefly, a recombinant baculovirus encoding hCdh1 with His6 tag fused to its N terminus was constructed using the Bac-to-Bac system (Invitrogen). Sf9 cells were infected with the hCdh1 baculovirus at a multiplicity of infection (MOI) of 5 for 50 h. The Sf9 cells were treated with 0.5 µM of okadaic acid for 4 h prior to harvesting. The cells were lysed with a buffer containing 20 mM Tris (pH 7.7), 150 mM NaCl, and 0.1% Triton X-100. After centrifugation, the supernatant was incubated with Ni2+-NTA beads (Qiagen). After washing, hCdh1 was eluted with a step gradient of imidazole and dialyzed into the appropriate buffer for subsequent assays.
Continuous Spectrophotometric Assay for Cdc14 and Its Mutants Using Phosphorylated hCdh1 as a Substrate—Kinetic parameters for the dephosphorylation of the phosphorylated hCdh1 protein were determined using a continuous spectrophotometric assay. This assay incorporates a coupled enzyme system, which uses purine nucleoside phosphorylase and its chromogenic substrate 7-methyl-6-thioguanosine for the quantification of inorganic phosphate produced in the phosphatase reaction (23, 24). The change in absorbance at 360 nm was due to the conversion of 7-methyl-6-thioguanosine to 7-methyl-6-thioguanine in the presence of inorganic phosphate. Quantitation of the inorganic phosphate produced in the phosphatase reaction was determined using the extinction coefficient of 11,200 M–1 cm–1 at 360 nm and pH 7.0. Experiments of wild-type Cdc14 were carried out at 25 °C in a 1.6-ml reaction mixture containing 100 mM Tris at pH 7.0, 100 mM NaCl, 1 mM EDTA, 0.1 mg/ml purine nucleoside phosphorylase (Sigma), and 50 µM 7-methyl-6-thioguanosine. The spectrophotometric measurements were conducted using a PerkinElmer Lambda 14 spectrophotometer equipped with a magnetic stirrer in the cuvette holder. The kcat/Km values for the Cdc14 catalyzed dephosphorylation of hCdh1 were obtained by nonlinear least square fit to Equation 2. Experiments with Cdc14 mutants were carried out at 25 °C in a 200-µl reaction mixture under the same conditions in 96-well plates. The spectrophotometric measurements were obtained using a Molecular Devices Spectro MAX 340 plate reader. The kcat/Km values were obtained from a linear least square fit of the initial rate versus substrate concentration to Equation 1. In these measurements, the phosphorylated hCdh1 concentration was less than 4 µM, and the concentrations of the Cdc14 and its mutants ranged from 6.4 64 nM.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTS AND DISCUSSIONREFERENCES |
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To begin to probe the active site properties of Cdc14, we explored the ability of Cdc14 to catalyze the hydrolysis of a panel of aromatic phosphates, including 1-, 2-, and multi-ringed substrates. The chemical structures for pNPP, DiFMUP, and OMFP are shown in Fig. 1. All of the experiments described in this study were performed with the full-length recombinant Cdc14 from the budding yeast with an N-terminal His6 tag to enable facile purification. Steady-state parameters for the Cdc14-catalyzed hydrolysis of aryl phosphates were determined at pH 7.0 and 30 °C.
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-naphthyl phosphate is 5.3-fold higher than that for 4-chlorophenyl phosphate even though these substrates also have the same leaving group pKa value. Unlike other dual specific phosphatases, such as Cdc25 phosphatases, mitogen-activated protein kinase phosphatases, and VH1-related (VHR), which display a striking preference (2–3 orders of magnitude) for bulky multi-ringed substrates like OMFP over pNPP (29–34), the kcat/Km for the Cdc14-catalyzed hydrolysis of OMFP is only 8-fold higher than that of pNPP. Interestingly, the tyrosine-specific PTPs exhibit very limited preference for substrates with more than one aromatic ring (20, 25, 31). Collectively, the unique affinity of Cdc14 for 2-ringed aryl phosphate substrates suggest that the active site properties of the Cdc14 phosphatases are distinct from those of other dual specific and tyrosine-specific PTPs. This information regarding the range of functionality that can be readily accommodated by the active site of Cdc14 phosphatases may be useful when designing active site-directed inhibitors for these enzymes.
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The requirement for general acid/base in Cdc14 catalysis has not been established. As shown in Table I, the kcat value for the Cdc14-catalyzed hydrolysis of pNPP is slightly less than that for DiFMUP, but the kcat/Km value for pNPP is 1,820-fold lower than that for DiFMUP. The kcat parameter reflects the overall rate-limiting step of enzyme catalysis, and, in the case of PTPs, previous studies have shown this to be mainly associated with hydrolysis of the E-P intermediate (k3 in Reaction Scheme 1) under acidic to neutral conditions (49). The fact that the Cdc14-catalyzed hydrolysis of pNPP and DiFMUP, two substrates that differ markedly in their leaving group pKa (7.14 for p-nitrophenol and 4.7 for DiFMU), display similar kcat values suggests that the rate-determining step is mostly E-P hydrolysis (also see below).
The kinetic parameter kcat/Km monitors the PTP reaction beginning with binding of the substrate and up to and including the first irreversible step in the kinetic mechanism (Reaction Scheme 1), which is E-P formation accompanied by release of the leaving group. Kinetic isotope effect studies indicate that the chemical step (i.e. phosphoryl transfer from substrate to the enzyme nucleophile) is rate-limiting for the kcat/Km term (36, 41, 43). In general, the PTP-catalyzed E-P formation exhibits little leaving group dependence due to protonation of the phenolic oxygen by the general acid. In contrast, general acid deficient mutant PTPs show large leaving group dependence on kcat/Km (37, 40, 50). For Cdc14-catalyzed hydrolysis of pNPP and DiFMUP, even when one excludes the contribution from preferential binding of 2-ringed substrates over 1-ringed substrates (10-fold), the kcat/Km value for DiFMUP is still 180-fold higher than that of pNPP. It is rare for a wild-type PTP to display such a large leaving group dependence on kcat/Km. Consequently, it is not clear whether the Cdc14-catalyzed reaction involves general acid catalysis.
In order to further define the catalytic mechanism of Cdc14 and to identify additional catalytically important residues, in particular the general acid/base, we performed site-directed mutagenesis in Cdc14 and assessed the mutational effects by kinetic analysis. Because previous studies have shown that the general acid/base functionality in PTP catalysis is associated with acidic residues, we primarily focused our attention on the nine invariant acidic residues in the catalytic domain of Cdc14, namely, Asp50, Asp129, Asp143, Glu168, Glu171, Glu174, Asp177, Asp253, and Glu308 (Fig. 2). We made the most conservative site-directed mutations possible (i.e. Glu to Gln and Asp to Asn) in order to remove the ability of these residues to function effectively in general acid-base catalysis while minimizing structural perturbations in the proteins. All of the mutations were verified by DNA sequencing. Recombinant proteins were expressed in E. coli, and purified to near homogeneity as judged by SDS-PAGE (data not shown).
While this article was in preparation, the x-ray crystal structure of the catalytic domain of human Cdc14B (residues Pro44–His386) was published (51). Unlike other members of the PTP superfamily, the structure of hCdc14B reveals a novel arrangement of two structurally equivalent domains arranged in tandem. The N-terminal domain (domain A, residues 44–198 in hCdc14B) may contribute to Cdc14 substrate specificity and shares no sequence similarity with other dual specific phosphatases. The C-terminal domain (domain B, residues 213–386 in hCdc14B) contains the conserved PTP motif. The active site of hCdc14B is located within a long groove between the interface of the A- and B-domain (Fig. 3). Residues of two surface loops of the A-domain, the extended WPD(A) (residues Thr37–Asn55 in Cdc14) and 
5A/6A (residues Leu117–Thr140 in Cdc14) loops, form one side of the groove. The WPD (residues Leu249–Asp258 in Cdc14) and Q-loops (residues Arg317–Gly323 in Cdc14) of the B-domain create the opposite face of the active site groove, whereas the interdomain linker -helix (residues Leu164–Phe175 in Cdc14) is positioned at the entrance to one end of the channel. This structure provides a framework for the interpretation of our mutational data. All of the residues that were subjected to mutagenesis are highlighted in the hCdc14B structure (Fig. 3).
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5A/6A loop) are located within the active site cavity (Fig. 3), it is possible that these two residues may play more important roles in the hydrolysis of the E-P intermediate. Only moderate reduction in kcat and no change or a slight increase in kcat/Km was observed for E171Q and D177N, suggesting that Glu171 and Asp177 are not essential for the Cdc14-catalyzed pNPP hydrolysis.
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atom of the Ser(P) substrate (Ref. 51 and Fig. 3). Based on the structural data, it seems that Asp253 could function as a general acid/base in the Cdc14-cataltzed reaction. Previous studies have shown that mutations of the general acid in PTPs generally result in two to three orders of magnitude decrease in both kcat/Km and kcat. As shown in Table II, the kcat values for the Cdc14-catalyzed hydrolysis of pNPP and DiFMUP decrease 640 to 1200-fold when Asp253 is replaced by an Ala. Less dramatic reductions in kcat (50–100-fold) are observed for the D253N mutant. Surprisingly, the kcat/Km values for D253A and D253N drop only 7- and 26-fold respectively, with pNPP as a substrate, while the kcat/Km values are virtually the same as those of the wild-type Cdc14 using DiFMUP as a substrate. In order to determine whether Asp253 indeed functions as a general acid/base in Cdc14 catalysis, we carried out additional experiments with both the wild-type and Asp253 mutant Cdc14 enzymes. Characterization of Asp253 as a General Acid/Base in Cdc14 Catalysis—PTPs with functional general acids show bell-shaped pH profiles where the maximum catalytic rate usually occurs around pH 5–6 with aryl phosphates as substrates. PTPs with general acids also show relatively flat leaving group dependences, i.e. the rate of catalysis is affected minimally by the pKa of the leaving group. Removal of the general acid through site-directed mutagenesis results in a flat pH profile with a significant drop in the rate of catalysis, and a significant leaving group dependence where the rate of catalysis drops as the leaving group pKa rises.
Leaving Group Dependence—The leaving group dependence for the hydrolysis of aryl phosphates catalyzed by Cdc14 was investigated at pH 7.0 and 30 °C using a series of substrates with leaving group pKa values ranging from 4.7 to 10. Since we have shown that the size of the aryl substrates can have substantial effects on the rate of Cdc14 catalysis (Table I), we analyzed the leaving group effect with the one-ringed and two-ringed substrates separately, in order to exclude steric effects. Fig. 4 shows the Brønsted plots, which relate the logarithm of kcat/Km to the pKa values of the leaving group. The slope of the Brønsted plot corresponds to the lg value, which reflects the amount of negative charge developed on the phenolic oxygen at the transition state.
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lg value of –1.01 ± 0.01. This value is similar to that of the uncatalyzed aryl phosphate dianion reaction (lg =–1.23, Ref. 54), which indicates a full negative charge on the leaving group oxygen at the transition state and suggests that no general acid catalysis occurs for substrates with leaving group pKa values between 4.7 and 7. Interestingly, the leaving group dependence for two-ringed substrates with leaving group pKa from 7–9.4 is flat, giving a lg value of –0.12 ± 0.02. In addition, the leaving group dependence for one-ringed substrates with leaving group pKa values from 7.14 to 9.99 is also flat, yielding a lg value of –0.06 ± 0.05. These values are similar to those determined for PTPs with functional general acid catalysis (20, 40, 55) and the uncatalyzed monoanion reaction (lg =–0.27, Ref. 54), in which the leaving group is protonated in the transition state.
The results described above suggest that the Cdc14-catalyzed reaction does not involve a general acid for substrates with low leaving group pKa values (4.7–7), whereas the hydrolysis of substrates with high leaving group pKa values (>7) appears to require a general acid. To determine if Asp253 fulfills the function of a general acid, we also quantified the leaving group effect for the Cdc14/D253A-catalyzed reaction. Unfortunately, the activities of Cdc14/D253A-catalyzed hydrolysis of one-ringed substrates with leaving group pKa > 8 are too low to be measurable. However, as shown in Fig. 4, we were able to determine the kinetic parameters for the Cdc14/D253A-catalyzed hydrolysis of 2-ringed substrates with leaving group pKa values 4.7–7.8, which yielded a lg value of –1.19 ± 0.07. This value indicates that no general acid catalysis occurs for substrates with leaving group pKa values between 4.7 and 7.8. That the kcat/Km values for the D253A mutant catalyzed hydrolysis of substrates with leaving group pKa between 4.7 and 7 are similar to those of the wild-type enzyme is consistent with the notion that the dephosphorylation of substrates with leaving group pKa below 7 does not require general acid catalysis. Importantly, there was clearly no break at pKa = 7 in the leaving group profile for the D253A mutant as the reaction rates for substrates with leaving group pKa values higher than 7 continued to drop. The fact that a lg value of –1.19 was obtained for the Cdc14/D253A-catalyzed reaction that extends beyond for substrates with leaving group pKa above 7 and that a further decrease in kcat/Km for substrates with leaving group pKa values >7 suggest that Asp253 may indeed act as a general acid in Cdc14-catalyzed hydrolysis of high leaving group pKa (7.0 9.99) substrates.
pH Dependence—To further substantiate that Asp253 is the general acid/base in Cdc14, we determined the pH-rate profiles for the hydrolysis of pNPP and DiFMUP by both the wild-type and the D253A mutant Cdc14 phosphatases (Fig. 5). Since the leaving group pKa for DiFMUP (4.7) is <7, we predicted that the wild-type Cdc14-catalyzed DiFMUP hydrolysis would not exhibit general acid catalysis for the E-P formation step (kcat/Km) but may require general base catalysis for the E-P hydrolysis step (kcat). As predicted, DiFMUP shows a flat pH-kcat/Km profile between pH 5.5 and 8.0 (Fig. 5A), consistent with a lack of general acid and with the leaving group dependence described above. Since no general acid catalysis is involved, removal of the carboxyl group at position 253 has no effect on kcat/Km, and the D253A mutant also displays a flat pH dependence for DiFMUP with kcat/Km values virtually identical to those of the wild-type enzyme (Fig. 5A).
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0.007 s–1) are more than 3 orders of magnitude slower than (kcat)max of the wild-type enzyme and are insensitive to pH values (Fig. 5B). Since removal of Asp253 in Cdc14 does not affect E-P formation, the large reduction on kcat for D253A appears primarily due to an impairment of E-P hydrolysis. Thus the rate-limiting step for the D253A-catalyzed DiFMUP reaction is also likely E-P hydrolysis (also see below). Consistent with Asp253 serving as a general base in the Cdc14-catalyzed reaction, mutating residue 253 from Asp into Ala eliminates the acid limb of the pH-kcat profile for DiFMUP. Since the leaving group pKa for pNPP is 7.14, we predicted that the wild-type Cdc14-catalyzed pNPP hydrolysis would exhibit characteristics of general acid/base catalysis. As expected, the wild-type Cdc14-catalyzed pNPP hydrolysis displays bell-shaped pH dependences for both kcat and kcat/Km, as observed with other PTPs using pNPP as a substrate (20, 35, 42). As shown in Fig. 5C, pNPP shows a pH optimum at 5.4 and steadily decreasing kcat/Km values at higher pH values, quite unlike the pH dependence of DiFMUP. This behavior is consistent with a Cdc14 residue that must be protonated for optimal catalytic efficiency, possibly a general acid. Because of the instability of Cdc14 at low pH values, the kcat/Km values below pH 5.0 could not be accurately measured. Consequently, the data were fitted to Equation 4 with the second ionization constant of pNPP (pKS2) fixed at the experimentally determined value of 5.1 (29). From this analysis, the pKE1 (ionization constant of the active site Cys residue) was determined to be 5.5 ± 0.5, the pKE2 (ionization constant of the general acid) 5.1 ± 0.4, and the (kcat/Km)max (2.3 ± 1.6) x 104 M–1 s–1. These ionization constants are similar to those obtained for other PTPs. Consistent with the involvement of a general acid in the Cdc14-catalyzed hydrolysis of substrates with leaving group pKa values > 7, the kcat/Km values for Cdc14/D253A are dramatically reduced, especially at the optimal pH, and replacement of Asp253 with an Ala abolishes the basic limb of the pH profile for pNPP (Fig. 5C). The pKE1 value for Cdc14/D253A could not be measured because of its low activity and instability at pH below 6. Therefore, we conclude that D253 is responsible for the basic limb of the pH-kcat/Km profile seen for wild-type Cdc14 with pNPP and that Asp253 must be protonated for optimal catalysis of this substrate.
As shown in Fig. 5D, the pH versus kcat profile for the Cdc14-catalyzed pNPP reaction shows a pH optimum at 6.8, and the (kcat)max,pKES1, and pKES2 values are determined to be 12.9 ± 1.4 s–1, 6.7 ± 0.1, and 6.9 ± 0.1, respectively. It is important to recognize that pKES1 and pKES2 are apparent values and may not represent microscopic ionizations of particular groups (56). It is also important to recognize that there are uncertainties in pKa values calculated from kinetic data when the differences in pKa are less than 0.6 units (57). Nevertheless, the acidic limb of the pH-kcat profile for pNPP is similar to that of DiFMUP, suggesting that the rate-limiting step likely corresponds to E-P hydrolysis in this pH range and that pKES1 likely reflects the ionization of the general base function of Asp253. In accordance with this, removal of Asp253 reduces the kcat by 3 orders of magnitude and eliminates the acidic limb of the pH-kcat profile (Fig. 5D). Interestingly, unlike the DiFMUP reaction, whose kcat values level off at a maximum plateau at pH > 7 (Fig. 5B), the pH-kcat profile for pNPP shows a steady decrease in kcat as the pH is raised (Fig. 5D). This may be caused by the fact that under more basic conditions, Asp253 is deprotonated, so that E-P formation step eventually becomes the rate-limiting step for substrates that require general acid catalysis. Consequently, the apparent pKES2 (6.9) derived from the basic limb of the pH-kcat profile for pNPP may not represent an intrinsic ionization constant but reflect a change in the rate-limiting step, from E-P hydrolysis at lower pH values to E-P formation at higher pH values. Since Asp253 is also involved in E-P formation, its removal also abolishes the basic limb of the pH-kcat profile for pNPP (Fig. 5D).
We also analyzed the pH dependence of the D253N-catalyzed hydrolysis of DiFMUP and pNPP (data not shown). Similar to D253A, the D253N mutant exhibits flat pH-rate profiles for both kcat and kcat/Km. The kcat/Km values for the D253N-catalyzed reaction are only slightly lower than those of the D253A mutant. Interestingly, the kcat for the D253N mutant is 10-fold higher than D253A (Table II), suggesting that the carboxyamide group in D253N is more efficient than the methyl group in D253A in promoting E-P hydrolysis. Collectively, the results from both leaving group and pH dependence of Cdc14-catalyzed reaction are consistent with a general acid independent mechanism for substrates with leaving group pKa < 7 and a general acid-dependent mechanism for substrates with leaving group pKa > 7. Experiments with both the D253A and D253N mutants indicate that Asp253 act as a general acid in the E-P formation step (k2) and as a general base in the E-P hydrolysis step (k3) (Scheme 1).
Pre-Steady-state Burst Kinetics—We next proceeded to carry out pre-steady-state burst kinetic experiments in order to determine the rate of E-P formation and breakdown for the Cdc14-catalyzed reaction and to further define the role of Asp253 in catalysis. Unfortunately, burst kinetics was not feasible for the pNPP reaction because of the large Km value, which makes it difficult to saturate the enzyme. However, we successfully performed a burst kinetic analysis of the Cdc14-catalyzed hydrolysis of DiFMUP at pH 7.0 and 30 °C. If E-P formation is sufficiently rapid relative to its breakdown, we would expect to see a burst of product (DiFMU) upon rapid mixing of the enzyme and substrate. As shown in Fig. 6A, the time course for the Cdc14-catalyzed hydrolysis of DiFMUP is biphasic, characterized by a rapid, exponential burst phase, which is then followed by a slower linear phase. By varying the initial concentration of the enzyme and substrate we found that the amplitude of the burst is proportional to the enzyme concentration. The presence of a pre-steady-state burst upon the hydrolysis of DiFMUP by Cdc14 is consistent with rate-determining E-P hydrolysis for this substrate. This is in full agreement with the conclusion derived from the steady-state kinetic evidence discussed above. The rate constants of the individual chemical steps k2 and k3 as well as the dissociation constant KS can be obtained by plots of kburst (rate constant for the burst phase) versus DiFMUP concentrations (Fig. 6B) as described under "Experimental Procedures" (21). The k2, k3, and KS values are correspondingly 151 ± 29 s–1,20 ± 5s–1, and 209 ± 66 µM. Thus, for the hydrolysis of DiFMUP, k2 is 7.6-fold greater than k3. Since kcat = k2k3/(k2 + k3), and Km = KSk3/(k2 + k3), kcat/Km can be transformed to k2/KS. The k2/KS value (720,000 M–1 s–1) determined by the stopped-flow method is similar to the kcat/Km value (770,000 M–1 s–1) obtained by steady-state experiments.
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Since k3, the rate constant for E-P hydrolysis, must be identical for dephosphorylation of either substrates, we can approximately solve for the kinetic parameter k2 in the Cdc14-catalyzed pNPP reaction based on kcat = k2k3/(k2 + k3). The calculated k2 for pNPP is 5.8 s–1. Therefore, the rate of E-P formation is 26-fold faster with DiFMUP as a substrate than with pNPP. Since k2 and k3 for the pNPP reaction are comparable, both E-P formation and breakdown contributes to the rate-limiting step in pNPP hydrolysis. This also support the conclusion derived from the pH-kcat analysis of the pNPP reaction that the kcat is rate-limited by both k2 and k3 at pH 7. Since kcat/Km = k2/KS, KS for pNPP can be calculated to be 13.8 mM using the estimated k2 value and the experimentally determined kcat/Km for pNPP (420 M–1 s–1). Thus, the dissociation constant (KS) of Cdc14 for DiFMUP (209 µM) is 66-fold lower than that for pNPP, providing further evidence that Cdc14 possesses intrinsically higher affinity for 2-ringed aryl phosphates. This property may relate to the existence of a hydrophobic pocket adjacent to the active site that recognizes the p + 1 Pro residue in Cdc14 substrates (51). The enhanced substrate binding may be partially responsible for the faster rate of E-P formation with DiFMUP than pNPP.
Activity of Cdc14 Toward Its Physiological Substrate hCdh1—The crystal structure of hCdc14B shows that acidic residues Glu168, Glu171, and Asp177 in the interdomain linker -helix (residues Leu164–Phe175 in Cdc14) cluster at the entrance of the catalytic site, suggesting that these residues may confer specificity for basic residues in the substrates (51). However, substitution of these acidic residues with an Ala reduces the kcat/Km for hydrolysis of a peptide substrate, Ala-Ser(P)-Pro-Arg-Arg-Arg, by only 2-fold (51). It is not known whether these acidic residues play a role in recognizing protein substrates.
To gain further insight into the molecular basis of Cdc14 substrate specificity, we chose to analyze the Cdc14-catalyzed dephosphorylation of a physiological protein substrate Cdh1. Cdh1 is a substrate-specific co-activator of the APC (58, 59), which functions as an E3 ubiquitin ligase to degrade the mitotic cyclin Clb2 and other substrates during the G1 phase of the cell cycle. Cdh1 is present throughout the cell cycle, but its binding to APC is blocked by Cdk phosphorylation (22, 60). Cdc14 activates APC by dephosphorylating Cdh1, leading to destruction of Cdc28-Clb2 and mitotic exit (4). In addition, human Cdc14A can also dephosphorylate human Cdh1 (hCdh1) to promote the activation of APCCdh1 in vitro (7).
To follow the Cdc14-catalyzed hCdh1 dephosphorylation, we employed a continuous spectrophotometric enzyme-coupled assay in which the coupling enzyme, purine nucleoside phosphorylase, uses the inorganic phosphate, generated by the action of Cdc14, to convert 7-methyl-6-thioguanosine to 7-methyl-6-thioguanine and ribose-1-phosphate, resulting in an increase in absorbance at 360 nm (23, 24). The reaction was carried out at pH 7 and 25 °C. Fig. 7 shows a typical progress curve of the Cdc14-catalyzed hCdh1 dephosphorylation. To ensure first-order conditions, the reaction was conducted at several hCdh1 concentrations ranging from 0.5 to 4 µM. Identical kcat/Km values were obtained by fitting the progress curves to Equation 2, indicating that the substrate concentrations were << Km. The kcat/Km values for hCdh1 dephosphorylation by the full-length Cdc14 and the catalytic domain (residues 1–374) were determined to be (1.8 ± 0.04) x 105 and (1.3 ± 0.01) x 105 M–1 s–1, respectively. This is consistent with previous observations that C-terminal domain of Cdc14 (residues 375–551) is not required for its phosphatase activity (1). Interestingly, under the same conditions the human Cdc14A catalytic domain (residues 1–379) was able to dephosphorylate hCdh1 with a kcat/Km value of (4.0 ± 0.04) x 105 M–1 s–1, indicating that hCdh1 is an efficient substrate for both the yeast and human enzymes.
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The residues that we identified as important for hCdh1 dephosphorylation are located in structural elements that surround the Cdc14 catalytic site (Fig. 3). Thus, Asp50 and Asp129 reside in the WPD(A) (residues Thr37–Asn55) and 5A/6A (residues Leu117–Thr140) surface loops respectively. Asp253 is located at the WPD loop (residues Leu249–Asp258). And Glu168, Glu171, and Asp177 in the interdomain linker -helix (residues Leu164–Phe175) are positioned at the entrance to one end of the active site groove. Although none of these residues (with the exception of Asp253) are found in direct contact with the phosphopeptide Ala-Ser(P)-Pro in the crystal structure of the hCdc14B complex (51), our kinetic data reveal that Asp50, Asp129, Glu168, Glu171, and Asp177 are important for Cdc14-catalyzed hCdh1 dephosphorylation.
In summary, we have carried out a detailed kinetic and mechanistic study of the Cdc14 phosphatases with both small molecule aryl phosphates and a physiological protein substrate hCdh1. We found that Cdc14 has a unique preference for 2-ringed aromatic substrates, a finding that may have significant implications for inhibitor design. Steady-state kinetics analysis of the Cdc14-catalyzed reaction showed that Cdc14 hydrolyzes low leaving group pKa substrates (pKa < 7), like DiFMUP, without the aid of a general acid, and higher pKa substrates (pKa > 7), like pNPP, with a general acid. The use of both DiFMUP and pNPP in combination with leaving group dependence, pH dependence, and pre-steady-state kinetic techniques enabled the isolation and analysis of both the E-P formation and hydrolysis step. We were able to establish clearly the requirement of general acid catalysis for the E-P formation step in reactions with high leaving group pKa substrates, and the presence of general base catalysis in E-P hydrolysis. Mutational study of invariant acidic residues in Cdc14 unambiguously identified Asp253, which acts as a general acid during E-P formation and as a general base in E-P hydrolysis. Finally, we provided evidence that Asp50, Asp129, Glu168, Glu171, and Asp177 are involved in substrate recognition for the physiological protein substrate hCdh1.
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FOOTNOTES |
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** To whom correspondence should be addressed. Tel.: 718-430-4288; Fax: 718-430-8922. E-mail: zyzhang{at}aecom.yu.edu.
1 The abbreviations used are: CDK, cyclin-dependent kinase; Tricine, N-[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]glycine; PTP, protein-tyrosine phosphatase; pNPP, p-nitrophenyl phosphate; KAP, kinase-associated phosphatase; PRL, phosphatase of regenerating liver; PTEN, phosphatase and tensin homolog deleted on chromosome 10; MUP, 4-methylumbelliferyl phosphate; 8-FMUP, 8-fluoro-4-methylumbelliferyl phosphate; 6-FMUP, 6-fluoro-4-methylumbelliferyl phosphate; 5-FMUP, 5-fluoro-4-methylumbelliferyl phosphate; DiFMUP, 6,8-difluoro-4-methylumbelliferyl phosphate; OMFP, 3-O-methylfluorescein phosphate; GST, glutathione S-transferase; NTA, nitrilotriacetic acid; APC, anaphase-promoting complex.
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTS AND DISCUSSIONREFERENCES |
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