©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
The Purification and Characterization of a Human Dual-specific Protein Tyrosine Phosphatase (*)

(Received for publication, October 18, 1994)

John M. Denu (1)(§) Gaochao Zhou (1)(¶) Li Wu (1) Rong Zhao (1) Jirundon Yuvaniyama (1) (2)(**) Mark A. Saper (1) (2)(§§) Jack E. Dixon (1)(¶¶)

From the  (1)Department of Biological Chemistry and (2)Biophysics Research Division, The University of Michigan, Ann Arbor, Michigan 48109-0606

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

An expression and purification method was developed to obtain the recombinant human dual-specific protein tyrosine phosphatase (PTPase) VHR in quantities suitable for both kinetic studies and crystallization. Physical characterization of the homogeneous recombinant protein verified the mass to be 20,500 ± 100 by matrix-assisted laser desorption mass spectrometry, confirmed the anticipated NH(2)-terminal amino acid sequence and demonstrated that the protein exists as a monomer. Conditions were developed to obtain crystals which were suitable for x-ray structure determination. Using synthetic diphosphorylated peptides corresponding to MAP (mitogen-activated protein) kinase (DHTGFLpTEpYVATR), an assay was devised which permitted the determination of the rate constants for dephosphorylation of the diphosphorylated peptide on threonine and tyrosine residues. The diphosphorylated peptides are preferred over the singly phosphorylated on tyrosine by 3-8-fold. The apparent second-order rate constant k/K for dephosphorlyation of phosphotyrosine on DHTGFLpTEpYVATR was 32,000 M s while dephosphorylation of phosphothreonine was 14 M s (pH 6). The reaction of DHTGFLpTEpYVATR with VHR is ordered, with rapid dephosphorylation on tyrosine occurring first followed by slow dephosphorylation on threonine. Similar results were obtained with F(NLe)(NLe)pTPpYVVTR, a peptide corresponding to a MAP kinase-like protein (JNK1) which is involved in the stress response signaling pathway.


INTRODUCTION

A class of protein tyrosine phosphatases (PTPases), (^1)referred to as ``dual-specific PTPases'', is emerging as important regulators of cell cycle control and mitogenic signal transduction. The first dual-specific PTPase identified corresponded to the H1 open reading frame in Vaccinia virus. This phosphatase was called VH1 for Vaccinia open reading frame H1 and was capable of hydrolyzing phosphate monoesters from both phosphotyrosine and phosphoserine containing peptides (Guan et al., 1991), distinguishing this catalyst from the tyrosine-specific PTPases. The VH1-like phosphatases and the PTPases share the active site sequence motif HCxxGxxR, but show limited sequence identity beyond this region. Following the discovery of VH1, it was shown that p80 (a protein necessary for the passage through mitosis) shared sequence identity to VH1. Several laboratories demonstrated that p80 had intrinsic tyrosine phosphatase activity and that it would dephosphorylate the cyclin-dependent protein kinase p34 on Thr-14 and Tyr-15. The dephosphorylation then led to the activation of p34 and subsequent entry into mitosis (Dunphy and Kumagai, 1991; Galaktionov and Beach 1991; Gautier et al., 1991; Kumagai and Dunphy, 1991; Millar et al., 1991; Strausfeld et al., 1991).

Ishibashi et al.,(1992) identified a dual-specific PTPase, VHR (for VH1-Related) using an expression cloning strategy (Fig. 1). The human fibroblast cDNA clone encoded a protein of 185 amino acids. The bacterially expressed glutathione S-transferase/VHR fusion protein was shown to be a protein phosphatase with dual specificity based upon its ability to hydrolyze phosphoserine from casein as well as phosphotyrosine from a number of tyrosine-phosphorylated growth factor receptors (Ishibashi et al., 1992).


Figure 1: A schematic diagram of the protein structure of the dual-specific protein tyrosine phosphatases.



Two laboratories have independently identified an identical dual-specific phosphatase (Cdi1 or KAP) (Gyuris et al., 1993; Hannon et al., 1994). Both laboratories demonstrated that Cdi1 (KAP) interacts with the cyclin-dependent kinases CDK2 and cdc2, suggesting that these PTPases may be directly involved in cell cycle control. It is important to note that activation of these cyclin-dependent kinases appears to be dependent upon the phosphorylation state of adjacent tyrosine and threonine residues. The list of dual-specific phosphatases (Table 1) includes the mammalian proteins PAC-1 and MKP-1 (Fig. 1). These phosphatases have been shown to dephosphorylate and inactivate threonine- and tyrosine-phosphorylated MAP (mitogen-activated protein) kinase in vivo (Ward et al., 1994; Sun et al., 1993). MAP kinase must be phosphorylated on adjacent tyrosine 185 and threonine 183 residues for activation (Her et al., 1993). Both MKP-1 and PAC-1 appear to be immediate-early gene products. Dual-specific phosphatases are also present in yeast with recent reports suggesting involvement of MSG5 in the mating pathway (Doi et al., 1994) and of YVH1 in nitrogen regulation (Guan et al., 1992), (Table 1, Fig. 1). Based upon the number of recently reported sequences of dual-specific phosphatases, it would appear that these proteins constitute a large family of catalysts.



In the few cases that have been investigated, these phosphatases appear to display a marked preference for protein kinases which can be phosphorylated on tyrosine and threonine in close proximity (Table 1). Despite the growing number of dual-specific PTPases identified, little is known about the biochemical mechanisms employed by this family of enzymes. The current investigation begins to explore the catalytic properties of the dual-specific phosphatases. Our strategy was to select a prototype of this class of phosphatases, express and purify large quantities of the recombinant enzyme, characterize the protein biochemically and crystallize the enzyme for x-ray structure determination. Since human VHR contains the catalytic domain which is common to all members of this class (Fig. 1), this enzyme was an ideal choice for analysis. Here, we describe the overexpression, purification, and biochemical and kinetic characterization of VHR.


MATERIALS AND METHODS

Overexpression and Purification

The coding sequence for VHR was amplified using PCR (polymerase chain reaction) from a human fetal brain cDNA library (Stratagene). The 5` PCR primer contained an adapter sequence for the DNA restriction enzyme NdeI, and the 3` primer contained an EcoRI site downstream of the native stop codon (Zhou et al., 1994). The PCR product was ligated into the PCRII plasmid (Invitrogen) and sequenced. The coding region was removed by NdeI and EcoRI digestion and ligated into the pT7-7 plasmid to generate pT7-7-VHR. The pT7-7-VHR plasmid was used to transform competent BL21/DE3 bacteria. The transformed bacteria were grown on 2xYT plates containing 100 µg/ml ampicillin. Overnight 10-ml cultures originating from isolated colonies were used to inoculate 1 liter of 2xYT containing 140 mg/liter ampicillin. When the growth reached an optical density of 0.8 at 600 nm, isopropyl-beta-D-thiogalactopyranoside (IPTG) was added at 100 mg/liter, and the bacteria were grown for an additional 8 h. The cells were harvested by centrifugation at 5000 times g, resuspended in 15 ml/liter culture of 50 mM Tris, 1 mM EDTA, 1 mM dithiothreitol (pH 7.4), and lysed by pressure (1200 pounds/square inch) using a French Pressure Cell Press (American Instrument Co. Inc.). The cell debris was removed by centrifugation at 27,000 times g (20 min), and 0.5% polyethylene imine was added to the soluble extract to precipitate polynucleotides and their associated proteins. After 15 min of stirring at 4 °C, the precipitate were removed by centrifugation at 27,000 times g. Ammonium sulfate was added at 35% of saturation, stirred for 15 min, and centrifuged at 27,000 times g for 20 min. To the supernatant, ammonium sulfate was added up to 65%, stirred, and centrifuged as before. The resulting pellet was dissolved in a minimum of 20 mM MES, 1 mM EDTA, 1 mM dithiothreitol (pH 6) and dialyzed against 500 ml of the same buffer for 90 min. The dialysate was loaded onto a 50-ml S-Sepharose (Sigma) cation-exchange column pre-equilibrated with MES buffer. The column was then washed with 500 ml of the MES buffer, and the enzyme was eluted with a 0-0.6 M linear NaCl gradient of 400 ml. VHR elutes at 300 mM NaCl. Fractions containing high phosphatase activity with para-nitrophenylphosphate (pNPP) were pooled and concentrated to between 5-10 ml by filtration (Centricon-3 or 10 Concentrator by Amicon). The concentrated solution was loaded onto a Sephadex G-75 gel filtration column (2.5 times 100 cm) and eluted with 270 ml of 50 mM Tris, 1 mM EDTA (pH 7.4). Fractions exhibiting phosphatase activity were analyzed by SDS-polyacrylamide gel electrophoresis to assess the purity. All fractions which contained only a single band corresponding to VHR were pooled and concentrated by filtration to a final concentration of 0.2-0.5 mM. The enzyme was stored at -20 °C in 50 mM Tris, 1 mM EDTA (pH 7.4), until use.

Protein Characterization: Mass Spectroscopy, Amino Acid Analysis, Amino-terminal Sequencing and Size Exclusion Chromatography

Matrix-assisted laser desorption mass spectrometry (MALDI) of the purified protein was performed on a VESTEC-2000 instrument. Amino acid analysis of the purified enzyme was performed on an Applied Biosystems (ABI) instrument which was composed of three integrated modules: the Applied Biosystems model 420H hydrolyzer/derivatizer, the Applied Biosystems model 130A Separation System, and the Applied Biosystems model 610 Data System. NH(2)-terminal protein sequencing was performed on an Applied Biosystems 473A sequenator. The phenylthiohydantoin-amino acids were identified and quantitated by HPLC and the Applied Biosystems model 610a.

Size exclusion chromatography was performed on a Pharmacia fast protein liquid chromatography system using a Superose 12 column and a mobile phase buffer consisting of 50 mM Tris, 150 mM NaCl, 1 mM EDTA (pH 7.4), flowing at a rate of 0.25 ml/min.

Peptide Synthesis

Synthesis of peptides followed typical FMOC/t-butyl protecting strategies and standard cycles on an Applied Biosystems/Perkin Elmer model-431 peptide synthesizer using HBTU and 1-hydroxybenzotriazole for activation. FMOC amino acids were from Synthetec, tetrazole from Glen Research, di-benzyl-N`,N`-diisopropyl-phosphoramidite was from Novabiochem. Other reagents were from Applied Biosystems/Perkin Elmer and solvents were peptide synthesis grade from Fisher. Purification of the phosphorylated peptides was on a Rainin Dynamax reversed-phase HPLC column using an acetonitrile/water gradient containing 0.1% trifluoroacetic acid. The purity of all peptides was verified by analytical reversed-phase HPLC and the structure was verified by amino acid analysis and electrospray or laser desorption mass spectrometry.

The general approach to phosphopeptide synthesis used in this study, global phosphorylation, involved post-synthetic phosphorylation while the peptide was still attached to the solid-phase support with all of its protecting groups intact except those on selected hydroxyl groups (Kitas et al., 1991; Andrews et al., 1991). The hydroxyamino acids were introduced during synthesis as the FMOC-amino acids with no side chain protection. Tyr, Thr, and Ser residues which were not to be phosphorylated contained t-butyl ether protecting groups. The NH(2)-terminal amino acid was incorporated as the t-Boc-protected amino acid. The resin was dried for 24 h on a vacuum system at room temperature before swelling in anhydrous DMF. Reaction with a 20-fold molar excess of the phosphoramidite and 50-fold molar excess of tetrazole for 1-2 h was similar to previous reports (Kitas et al., 1991; Andrews et al., 1991). The resin was washed thoroughly with dry DMF and then subjected to oxidation with 0.5 Mt-butyl peroxide in DMF for times ranging from 0.5 to 4 h at room temperature. The phosphorylated peptide resin was then washed thoroughly with DMF, methylene chloride, dried, and subjected to cleavage and deprotection in 90% trifluoroacetic acid and 5% thioanisole, 3% ethanedithiol, 2% anisole. The cleaved peptide was then precipitated by addition of 15 volumes cold diethylether.

One of the synthetic peptides (DHTGFLpTEpYVATR) exhibited only low levels of phosphorylation. The use of alternative solvents or longer reaction times did not significantly improve the extent of phosphorylation. This was attributed to steric effects of the amino-terminal segment since shorter forms of this peptide were quite efficiently phosphorylated. The approach used was to synthesize the peptide by normal procedures except that the synthesis was stopped after 8 residues, the FMOC protecting group left on the Leu residue, and the peptide resin subjected to the normal phosphorylation protocol. After the oxidation step, the peptide was thoroughly washed and placed back on the synthesizer, and the last 5 amino acid residues coupled. A significant amount of dephosphorylation occurred at the Thr residue during the piperidine deprotection step in the subsequent cycles. This was reduced by shortening the piperidine deprotection step in each cycle and incorporating a t-Boc-protected amino acid in the amino-terminal position.

Assays

Dephosphorylation of phosphorylated MAP kinase by VHR was determined using the method of Zheng and Guan(1993).

Three different types of quantitative enzymatic assays were employed in these studies. A three-component buffer consisting of 0.05 M Tris, 0.05 M Bis-Tris, and 0.1 M acetate was used in all the kinetic analyses. This buffer maintains constant ionic strength over its useful pH range (Ellis and Morrison, 1982). Since preliminary studies (data not shown) indicated that enzymatic activity was sensitive to ionic strength, maintaining constant ionic strength was critical. This buffer exhibited no inhibitory effects on enzymatic activity. All assays were performed at 30 °C. Using pNPP as a substrate, the phosphatase reaction was followed as an increase in absorbance at 405 nm of the product para-nitrophenolate (Zhang and Van Etten, 1991). Initial rates were determined from the change in absorbance upon addition of 1 N NaOH. Rates were determined over the linear region of the reaction using the molar extinction coefficient of 18,000 M cm for the product para-nitrophenolate. Inital rates at various initial substrate concentrations were then fit directly to the Michaelis-Menten using the nonlinear least-squares program Kinetasyst for the Macintosh (IntelliKinetics, State College, PA).

A continuous spectrophotometric assay described previously by Zhang et al. (1993) was employed to follow the dephosphorylation of various phosphotyrosine-containing peptides. This assay takes advantage of the difference in absorbance at 282 nm between phosphotyrosine and tyrosine and can be utilized to follow the complete time course of the enzyme catalyzed hydrolysis of phosphotyrosine containing peptides. Initial concentrations of peptide ranged from 0.3 to 1.8 mM. The complete time course of the reaction can be fitted to the integrated form of the Michaelis-Menten using a nonlinear least-squares algorithm (Yamaoka et al., 1981). This method has been successfully used in the determination of the kinetic parameters for the reaction between phosphotyrosine peptides and RatPTP1 and the Yersinia PTPase (Zhang et al., 1993). This progress curve analysis yielded the identical values to those determined by typical initial rate analysis. However, because the build up of phosphate can begin to inhibit the reaction as more phosphopeptide is converted to product, was modified to to account for the slight inhibition by phosphate. Since dephosphorylated peptides do not bind to the PTPases (Zhang et al., 1993; Cho et al., 1991), no correction for inhibition by this product was necessary. In , E(0) is the enzyme concentration, p is the product concentration upon complete reaction, p is the product concentrationa t time t, and K(i) is the inhibition constant for phosphate. The peptide dephosphorylation data were fit to and kinetic parameters k, K(m) and k/K(m) were obtained.

The third method involved reverse phase HPLC separation and quantitation of the substrates and products from the reaction of VHR with DHTGFLpTEpYVATR and F(Nle)(Nle)pTPpYVVTR. The DHTGFLpTEpYVATR phosphopeptide corresponded to the putative activation sites on MAP (mitogen-activated protein) kinase, whereas the F(Nle)(Nle)pTPpYVVTR phosphopeptide corresponded to the activation sites of JNK1 (c-Jun NH(2)-terminal kinase). With this method, the rate of enzyme-catalyzed hydrolysis at phosphothreonine of both peptides was determined. A Vydac C18 (0.21 times 25 cm) column was attached to an Applied Biosystems 130A Separation System HPLC instrument with UV detection set at 220 nm. The peptides were eluted (150 µl/min) over 60 min using a linear gradient of 15-35% B (80% acetonitrile, 20% water, 0.1% trifluoroacetic acid). Solution A was 0.1% trifluoroacetic acid in water. The retention times of DHTGFLpTEpYVATR in the various phosphorylation states -pTEpY-, -TEpY-, -TEY-, and -pTEY- were 30.6, 31.6, 43.8, and 47.3 min, respectively. The peptide corresponding to JNK1 kinase (F(Nle)(Nle)pTPpYVVTR) was analyzed in an identical fashion except for the following changes. The peptides were eluted over 60 min using a linear gradient of 20-60% B. The retention times of F(Nle)(Nle)pTPpYVVTR in the various phosphorylation states -pTPpY-, -pTPY-, and -TPY- were 22.8, 27.1, and 28.3 min, respectively. In the F(Nle)(Nle)pTPpYVVTR peptide, Nor-leucine was substituted for methionine (found in the native protein) because of the technical problem of oxidation of the thiol during synthesis. Nor-leucine is similar in size and has the same stereochemistry as methionine. All peaks were resolved and were verified by MALDI mass spectral analysis and by coelution with the corresponding authentic peptide. To quench the reaction, an equal volume of 1.5 M acetic acid was added to the aliquot before injection of 15-20 µl of this sample for HPLC separation. Peaks of the dephosphorylated peptide were collected, and the amount of peptide was determined by amino acid analysis. The concentration of dephosphopeptide was then determined as a function of time and fitted to the integrated form of the Michaelis-Menten . Control experiments with no added enzyme indicated that background hydrolysis was negligible.

Inhibition of VHR

A variety of oxyanions were tested as inhibitors of VHR. The inhibition constant K(i) was determined in the following manner. At various fixed concentrations of inhibitor, the initial velocity at different pNPP concentrations was measured as described above. For all inhibitors analyzed, the inhibition was competitive with respect to substrate. The data were fit to (Kinetasyst) to yield the inhibition constant K(i).


RESULTS

Purification and Physical Characterization

The dual-specific PTPase, VHR, was purified as outlined under ``Materials and Methods.'' The coding sequence of VHR was placed behind the bacteriophage T7 promoter in the pT7-7 plasmid. Induction of the transformed bacterial cultures with IPTG gave high levels of expression of the protein. As indicated in Table 2, approximately 30-40 mg of pure enzyme (30% yield) was routinely obtained from 3 liters of bacterial culture. After elution of VHR from the S-Sepharose cation-exchange column, the enzyme was judged to be at least 95% pure based on SDS-polyacrylamide gel electrophoresis (lane 3 in Fig. 2). Further purification on the Sephadex G-75 column resulted in a homogeneous preparation of enzyme (lane 4 in Fig. 2). This preparation was used in all subsequent studies. There was no loss of enzymatic activity after 10 months of storage at -20 °C. Above pH 6, the enzyme retained 100% of its activity after incubation at 30 °C for 3 days. Below pH 4.5, the enzyme rapidly denatures (data not shown). No significant drop in activity was observed after six cycles of freezing (-20 °C) and thawing (30 °C).




Figure 2: SDS-polyacrylamide gel electrophoresis of the purification steps for VHR. Lane 1, polyethyleneimine (0.5%) precipitation, supernatant; lane 2, 35-65% ammonium sulfate precipitation; lane 3, S-Sepharose cation-exchange column; lane 4, Sephadex G-75 gel filtration column. Amount of protein in lanes 3 and 4 are 6 and 10 µg, respectively.



To establish the oligomeric state of the enzyme, VHR was subjected to size exclusion chromatography as outlined under ``Materials and Methods'' (Fig. 3). The purified enzyme was injected onto the column at a concentration of 10 mg/ml. The elution time corresponded to a protein with an apparent molecular weight of 20,600, which was in good agreement with the mass of the monomer predicted by amino acid sequence composition (20,500 for 185 amino acids).


Figure 3: Size exclusion chromatography of VHR. Enzyme was injected at a concentration of 10 mg/ml and eluted in 50 mM Tris, 1 mM EDTA, 150 mM NaCl (pH 7.4) using a Superose 12 gel filtration column as described under ``Materials and Methods.'' Molecular size standards were bovine serum albumin (66,000), egg albumin (44,000), carbonic anhydrase (29,000), and cytochrome (12,400).



To establish the integrity of the purified enzyme, VHR was subjected to mass spectral, amino acid, and amino-terminal sequence analysis. The first six cycles of amino-terminal sequencing established that the protein begins with Ser-Gly-Ser-Phe-Glu-Leu. This is identical to the predicted sequence except that the initiator Met was removed. Amino acid analysis and the amino acid content predicted by the cDNA sequence were in excellent agreement (data not shown). Matrix-assisted laser desorption mass spectrometry analysis yielded an apparent mass of 20,500 ± 100, in good agreement with the predicted molecular weight. Consistent with these results, the purified enzyme contains 184 amino acids with a molecular weight of 20,400. Based upon amino acid analysis and UV absorbance measurements of purified VHR, the molar extinction coefficient at 280 nm was 11,500 M cm. All subsequent concentrations of VHR were determined by using this molar extinction coefficient.

Inhibition by Oxyanions

The inhibition constant K(i) was determined for tungstate, vanadate, and phosphate at pH 6. The K(i) for arsenate inhibition was determined previously (Zhou et al., 1994). All displayed competitive inhibition with respect to substrate (data not shown). The K(i) values for arsenate, tungstate, vanadate, and phosphate were 20.9 ± 0.7, 689 ± 108, 32.3 ± 8.1, and 969 ± 120 mM, respectively.

Crystallization

Crystallization conditions for VHR (10 mg/ml in 1 mM EDTA, 1 mM dithiothreitol, 50 mM Tris (pH 7.2) were screened using the sparse matrix sampling method (Jancarik and Kim, 1991; Crystal Screen I, Hampton Research) and hanging-drop vapor diffusion techniques. The initial condition (0.2 M Li(2)SO(4), 30% polyethylene glycol 4000, 0.1 M Tris ((pH 8.5) screening solution 17)) was optimized to grow single crystals at 22 °C (Fig. 4). Repeated macroseeding yielded VHR crystals (typical size: 0.3 times 0.2 times 0.2 mm^3) that diffract to at least 2 Å resolution on an SDMS multiwire area detector (Rigaku RU200, 50 kV, 100 mA). The crystal was of space group P2(1) with cell dimensions a = 61.08 Å, b = 60.14 Å, c = 51.94 Å, alpha = = 90°, and beta = 98.35°. We estimated the crystal to be 47% solvent for two 20.4-kDa molecules/asymmetric unit. However, crystal density determination and self-rotation functions will be necessary to conclusively rule out the possibility of one molecule/asymmetric unit (74% solvent). A complete native data set to 2.5 Å has been collected, and data collection from potential heavy atom derivatives is in progress.


Figure 4: Crystal of VHR. Crystal was grown by repeated macroseeding as described under ``Results.'' This typical crystal (0.38 times 0.2 times 0.1 mm^3) diffracted to 2.3 Å resolution.



Dephosphorylation of MAP Kinase, MAP Kinase Peptide, JNK Kinase Peptide and Neu Peptide

There was in vivo evidence that at least two of the dual-specific phosphatases catalyze the dephosphorylation of activated MAP kinase (Ward et al., 1994; Sun et al., 1993). Ward et al.(1994) have pointed out that there are numerous MAP kinase-like proteins, all of which appear to require phosphorylation on a similar -TxY- sequence. We initially examined MAP kinase as a substrate for VHR. The enzyme dephosphorylated MAP kinase on both threonine and tyrosine residues (data not shown). The difficulty in obtaining stoichiometrically diphosphorylated MAP kinase which incorporates 1 mol of phosphate at threonine 183 as well as 1 mol of phosphate at tyrosine 185 makes detailed kinetic analysis of partially phosphorylated MAP kinase problematic. For this reason, we devised an in vitro assay using synthetic peptides corresponding to the activation sites of MAP kinase as well as other kinases having the -TxY- motif.

To explore the dual specificity and the substrate specificity of VHR, the kinetic parameters k, K(m), and k/K(m) were determined for a variety of synthetic peptides including diphosphorylated MAP (DHTGFLpTEpYVATR) kinase and JNK1 kinase (F(Nle)(Nle)pTPpYVVTR), (Table 3). Phosphotyrosine dephosphorylation can be followed continuously using the spectrophotometric methods (Fig. 5) described under ``Materials and Methods.'' The k/K(m) parameter is the apparent second-order rate constant for the reaction between free substrate and free enzyme and involves both the steps in binding of substrate and catalysis. The k/K(m) value is therefore an excellent indicator of the quality of a particular substrate. On the other hand, the kinetic parameter k involves only the steps in catalysis and product release. The diphosphorylated peptides corresponding to MAP and JNK kinases (Table 3) yielded the highest k/K(m) values of all substrates, 32,300 and 26,600 M s, respectively. For comparison, the Neu (DAEEpYLVPQQG) peptide (Zhang et al., 1993), monophosphorylated on tyrosine, yielded a k/K(m) value of 4080 M s that was only slightly higher than pNPP (3240 M s). For VHR, the tyrosine-monophosphorylated MAP kinase peptide DHTGFLTEpYVATR yielded a k/K(m) value of 11,200 M s.




Figure 5: VHR- catalyzed hydrolysis of phosphotyrosine on JNK1 kinase. Initial concentration of F(NLe)(NLe)pTPpYVVTR was 1.8 mM, and enzyme concentration was 5 µM. Conditions: 30 °C and pH 6. Solid line is a fit of the data to , and the circles represent 200 data points.



Because the continuous spectrophotometric assay for phosphotyrosine-specific hydrolysis was invisible to the hydrolysis at phosphothreonine, we developed an HPLC-based assay that would allow us to monitor all possible reactions within a single experiment. This method can provide information on the order of hydrolysis at each site as well as the relative rates of hydrolysis. The apparent second-order rate constant k/K(m) for dephosphorylation of phosphothreonine on the diphosphorylated DHTGFLpTEpYVATR (MAP kinase) and F(NLe)(NLe)pTPpYVVTR (JNK1 kinase) peptides was determined by utilizing HPLC to separate and quantitate all four possible phosphorylation states, -TxY-, -pTxY-, -TxpY-, and -pTxpY-, as described under ``Materials and Methods.'' All peaks in the HPLC elution profiles were identified by a combination of mass spectral analysis and by comparison with the retention times of the authentic peptides. Fig. 6shows the elution profile at different reaction times when VHR was reacted with diphosphorylated MAP kinase peptide DHTGFLpTEpYVATR. Within the first 6 min, the DHTGFLpTEpYVATR peptide was rapidly (32,000 M s) and completely dephosphorylated on tyrosine, thus generating phosphothreonine peptide DHTGFLpTEYVATR which was then slowly hydrolyzed by VHR, yielding completely dephosphorylated peptide DHTGFLTEYVATR. No detectable amount of monophosphorylated phosphotyrosine peptide was observed in these experiments with either MAP kinase or JNK1 kinase. The amount of unlabeled peptide was calculated by amino acid analysis of the peak and was correlated with peak area to determine the concentration of peptide at each time point. Alternatively, the amount of peptide formed was determined by correlating the peak area to the amount of starting peptide. Fig. 7shows the data from the time course of reaction for hydrolysis at phosphothreonine with diphosphorylated MAP kinase peptide (DHTGFLpTEpYVATR) and the fit to the integrated Michaelis-Menten . The fit yielded a k/K(m) value of 13.5 ± 0.3 M s (pH 6) which was about 200-fold lower than the rate obtained with pNPP and about 2000-fold slower than phosphotyrosine hydrolysis on the same diphosphorylated peptide. Because the phosphothreonine peptide substrate DHTGFLpTEYVATR was not present at saturating levels (i.e.K(m) 1 mM), the parameter k could not be determined with good accuracy. At pH 7, the k/K(m) value decreased to 3.5 ± 0.3 M s, consistent with the decrease in values seen with both the phosphotyrosine peptides and pNPP as substrates. These results suggest that the pH optimum is close to pH 6, with activity decreasing at higher pH. Using the same HPLC-based analysis, a similar k/K(m) value of 16.8 ± 0.3 M s for phosphothreonine hydrolysis of diphosphorylated JNK1 kinase peptide (F(NLe)(NLe)pTPpYVVTR) was determined.


Figure 6: HPLC elution profile of the products of diphosphorylated MAP kinase peptide (DHTGFLpTEpYVATR)-catalyzed hydrolysis by VHR. Results represent reactions done at 30 °C and pH 6.0. Initial peptide concentration was 300 µM, and the VHR concentration was 1.65 µM. Details are given under ``Materials and Methods.''




Figure 7: Phosphothreonine hydrolysis of diphosporylated MAP kinase peptide (DHTGFLpTEpYVATR) catalyzed by VHR. Data points represent the quantitation of the results shown in Fig. 6as described under ``Materials and Methods.'' Solid line is a fit of the data to .




DISCUSSION

We have described a simple and convenient purification strategy for obtaining large amounts of the native human dual-specific PTPase VHR. Because results obtained from kinetic and biochemical analyses can be altered by engineered proteins, VHR was overexpressed as the native enzyme coded for by its cDNA sequence and not as a fusion protein nor as a histidine-tagged protein. This has enabled us to crystallize the enzyme for structure determination, physically characterize the enzyme and investigate its substrate specificity. Other than demonstrating the general phosphatase activity of this growing class of PTPases, there has been no report on the details of this reaction, the relative rates of dephosphorylation on the putative Thr-183 and Tyr-185 of MAP kinase and the substrate requirements of this important class of enzymes.

After VHR was purified to homogeneity, numerous physical methods were performed to ensure the integrity of the enzyme. Results from amino acid analysis, MALDI mass spectral analysis, and size-exclusion chromatography are consistent with the predicted mass of 20.4 kDa. With NH(2)-terminal amino acid sequencing revealing that the initiator Met was cleaved, the protein contains 184 amino acids.

Once the integrity of the enzyme was confirmed, the substrate specificity was determined. VHR will effectively dephosphorylate purified recombinant MAP kinase; however, because phosphorylation of MAP kinase by MAP kinase kinase was never stoichiometric on both tyrosine 185 and threonine 183, an alternative assay was developed to unambiguously determine the rates of dephosphorylation. Using stoichiometrically diphosphorylated MAP kinase (DHTGFLpTEpYVATR) and JNK1 kinase (F(Nle)(Nle)pTPpYVVTR) peptides in the continuous spectrophotometric assay and the HPLC method described here, the rates of the VHR-catalyzed hydrolysis at each site were determined. The diphosphorylated peptides are clearly the best substrates in terms of phosphotyrosine hydrolysis, with K(m) values between 100 and 200 µM and k/K(m) values of 30,000 M s. In comparison, peptides monophosphorylated on tyrosine have K(m) values above 1 mM. When the diphosphorylated peptides are reacted with VHR, no monophosphotyrosine peptide was ever detected, indicating that the dephosphorylation is ordered. The phosphotyrosine residue is rapidly hydrolyzed first at a rate of 30,000 M s followed by the slow (15 M s) rate of hydrolysis at the threonine position (Fig. 8).


Figure 8: The ordered dephosphorylation of diphosphorylated MAP kinase peptide (DHTGFLpTEpYVATR) and JNK1 kinase F(NLe)(NLe)pTPpYVVTR by VHR. The numbers refer to the relative rates of dephosphorylation at tyrosine (2000) and threonine(1) .



Both the dual-specific PTPases and the PTPases contain an essential cysteine residue that is thought to be the nucleophile which attacks the phosphorus atom, forming a thiol-phosphate intermediate along the catalytic pathway. We have previously established the importance of Cys-124 in the VHR-catalyzed reaction toward both phosphotyrosine and phosphoserine/threonine containing peptides (Zhou et al., 1994). When this cysteine is replace with serine, the enzyme is functionally inactive toward both types of substrates, suggesting a similar mechanism and the same active site with both types of substrates. The proposed intermediate was trapped and characterized when VHR was mixed with [P]phosphotyrosine-labeled Raytide. However, attempts to trap the intermediate with phosphoserine/threonine-containing peptides failed. This apparent inconsistency can be explained with the results observed in the current study. The amount of intermediate formed in the reaction is directly proportional to the ratio of the net rate of breakdown to the net rate of formation of the intermediate. Given that the k/K(m) value for phosphothreonine peptide hydrolysis is about three orders of magnitude slower than hydrolysis of phosphotyrosine containing peptide, the amount of intermediate would also be expected to drop by the same magnitude. The k/K(m) parameter involves the steps of binding through the release of the first product, the alcohol. Thus, this rate constant measures the net rate of formation of the intermediate. Since a common thiol-phosphate intermediate would be formed with both phosphothreonine and phosphotyrosine substrates, the rate of breakdown of the intermediate is expected to be identical. As a result, the levels of trapped intermediate with phosphoserine/threonine substrates would be negligible and therefore undetectable by the described trapping method.


FOOTNOTES

*
This work was supported by grants from the Walther Cancer Institute and National Institutes of Health NIDDKD Grants 18849 and 18024. Partial funding for crystallographic work (to J. Y. and M. A. S.) was provided by University of Michigan Multipurpose Arthritis Center Grant NIH P560 AR20557. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Recipient of NIH Postdoctoral Fellowship DK07245-17.

Postdoctoral fellow of the American Diabetes Association.

**
Development and Promotion of Science and Technology Talents Project scholar from Thailand.

§§
Pew Scholar in the Biomedical Sciences.

¶¶
To whom correspondence should be addressed: Dept. of Biological Chemistry, University of Michigan Medical School, Rm. 5416 Medical Science I, Ann Arbor, Michigan 48109-0606. Tel.: 313-764-8192; Fax: 313-763-4581.

(^1)
The abbreviations used are: PTPase, protein tyrosine phosphatase; VHR, vaccinia H1 related; IPTG, isopropyl-beta-D-thiogalactopyranoside; Bis-Tris, (bis[2-Hydroxyethyl]imino-tris[hydroxymethyl]methane; pNPP, para-nitrophenylphosphate; MES, (2-[N-morpholino]ethanesulfonic acid); DMF, dimethylformamide; t-Boc, N-tertiary-butoxy-carbonyl; MAP, mitogen-activated protein; PCR, polymerase chain reaction; HPLC, high performance liquid chromatography; MALDI, matrix-assisted laser desorption mass spectrometry.


ACKNOWLEDGEMENTS

We thank Dr. Phil Andrews and the Protein Core Facility at the Medical School, University of Michigan, for performing amino acid analysis, NH(2)-terminal sequencing, MALDI mass spectrometry, and synthesis of the peptides. We also thank Drs. Kun-Liang Guan and Elizabeth Butch for the MAP kinase and MAP kinase kinase.


REFERENCES

  1. Andrews, D. M., Kitchin, J., and Seale, P. W. (1991) Int. J. Peptide Protein Res. 38, 469-475 [Medline] [Order article via Infotrieve]
  2. Cho, H. J., Ramer, S. E., Itoh, M., Winkler, D. G., Kitas, E., Bannwarth, W., Burn, P., Saito, H., and Walsh, C. T. (1991) Biochemistry 30, 6210-6216 [Medline] [Order article via Infotrieve]
  3. Derijard, B., Hibi, M., I., W., Barrett, T., Su, B., Deng, T., Karin, M., and Davis, R. (1994) Cell 76, 1025-1037 [Medline] [Order article via Infotrieve]
  4. Doi, K., Gartner, A., Ammerer, G., Errede, B., Shinkawa, H., Sugimoto, K., and Matsumoto, K. (1994) EMBO J. 13, 61-70 [Abstract]
  5. Dunphy, W. G., and Kumagai, A. (1991) Cell 67, 189-196 [Medline] [Order article via Infotrieve]
  6. Ellis, K. J., and Morrison, J. F. (1982) Methods Enzymol. 87, 405-426 [Medline] [Order article via Infotrieve]
  7. Galaktionov, K., and Beach, D. (1991) Cell 67, 1181-1194 [Medline] [Order article via Infotrieve]
  8. Gautier, J., Solomon, M. J., Booher, R. N., Bazan, J. F., and Kirschner, M. W. (1991) Cell 67, 197-211 [Medline] [Order article via Infotrieve]
  9. Guan, K., Hakes, D. J., Wang, Y., Park, H.-D., Cooper, T. G., and Dixon, J. E. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 12175-12179 [Abstract]
  10. Guan, K. L., Broyles, S. S., and Dixon, J. E. (1991) Nature 350, 359-362 [CrossRef][Medline] [Order article via Infotrieve]
  11. Gyuris, J., Golemis, E., Chertkov, H., and Brent, R. (1993) Cell 75, 791-803 [Medline] [Order article via Infotrieve]
  12. Hannon, G. J., Casso, D., and Beach, D. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 1731-1735 [Abstract]
  13. Her, J., Lakhani, S., Zu, K., Vila, J., Dent, P., Sturgill, T., and Weber, M. (1993) Biochem. J. 296, 25-31 [Medline] [Order article via Infotrieve]
  14. Ishibashi, T., Bottaro, D. P., Chan, A., Miki, T., and Aaronson, S. A. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 12170-12174 [Abstract]
  15. Jancarik, J., and Kim, S. H. (1991) J. Appl. Crystallogr. 24, 409-411 [CrossRef]
  16. Kitas, E. A., Knorr, R., and Bannwarth, W. (1991) Twelfth American Peptide Symposium , June 16-21, 1991, Boston
  17. Kumagai, A., and Dunphy, W. G. (1991) Cell 64, 903-914 [Medline] [Order article via Infotrieve]
  18. Millar, J. B. A., McGowan, C. H., Lenaers, G., Jones, R., and Russell, P. (1991) EMBO J. 10, 4302-4309
  19. Russell, P., and Nurse, P. (1986) Cell 45, 145-153 [Medline] [Order article via Infotrieve]
  20. Strausfeld, U., Labbe, J. C., Fesquet, D., Cavadore, J. C., Picard, A., Sadhu, K., Russell, P., and Doree, M. (1991) Nature 351, 242-245 [CrossRef][Medline] [Order article via Infotrieve]
  21. Sun, H., Charles, C. H., Lau, L. F., and Tonks, N. K. (1993) Cell 75, 487-493 [Medline] [Order article via Infotrieve]
  22. Ward, Y., Gupta, S., Jensen, P., Wartmann, M., Davis, R. J., and Kelly, K. (1994) Nature 367, 651-654 [CrossRef][Medline] [Order article via Infotrieve]
  23. Yamaoka, K., Tanigawara, Y., Nakagawa, T., and Uno, T. (1981) J. Pharmacobio-Dyn. 4, 879-885 [Medline] [Order article via Infotrieve]
  24. Zhang, Z.-Y., and Van Etten, R. L. (1991) J. Biol. Chem. 266, 1516-1525
  25. Zhang, Z.-Y., Maclean, D., Thieme-Sefler, A. M., Roeske, R., and Dixon, J. E. (1993) Anal. Biochem. 211, 7-15 [CrossRef][Medline] [Order article via Infotrieve]
  26. Zheng, C.-F., and Guan, K.-L. (1993) J. Biol. Chem. 268, 16116-16119 [Abstract/Free Full Text]
  27. Zhou, G. Z., Denu, J. M., Wu, L., and Dixon, J. E. (1994) J. Biol. Chem. 269, 28084-28090 [Abstract/Free Full Text]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.