Kinetic Basis for Activation of CDK2/Cyclin A by Phosphorylation*

Jonathan C. HagopianDagger , Matthew P. KirtleyDagger §, Lisa M. Stevenson§, Roxanne M. GergisDagger , Alicia A. Russo||, Nikola P. Pavletich||, Stanley M. Parsons**, and John LewDagger DaggerDagger

From the Dagger  Department of Molecular, Cellular and Developmental Biology, and the ** Department of Chemistry and Biochemistry and the Interdepartmental Program in Biochemistry and Molecular Biology, University of California, Santa Barbara, California 93106, || Howard Hughes Medical Institute, Cellular Biochemistry and Biophysics Program, Memorial Sloan-Kettering Cancer Center, New York, New York 10021, and the  Interdepartmental Graduate Program in Biochemistry and Molecular Biology, University of California, Santa Barbara, California 93106

Received for publication, August 11, 2000, and in revised form, September 29, 2000



    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The activation of most protein kinases requires phosphorylation at a conserved site within a structurally defined segment termed the activation loop. A classic example is the regulation of the cell cycle control enzyme, CDK2/cyclin A, in which catalytic activation depends on phosphorylation at Thr160 in CDK2. The structural consequences of phosphorylation have been revealed by x-ray crystallographic studies on CDK2/cyclin A and include changes in conformation, mainly of the activation loop. Here, we describe the kinetic basis for activation by phosphorylation in CDK2/cyclin A. Phosphorylation results in a 100,000-fold increase in catalytic efficiency and an approximate 1,000-fold increase in the overall turnover rate. The effects of phosphorylation on the individual steps in the catalytic reaction pathway were determined using solvent viscosometric techniques. It was found that the increase in catalytic power arises mainly from a 3,000-fold increase in the rate of the phosphoryl group transfer step with a more moderate increase in substrate binding affinity. In contrast, the rate of phosphoryl group transfer in the ATPase pathway was unaffected by phosphorylation, demonstrating that phosphorylation at Thr160 does not serve to stabilize ATP in the ATPase reaction. Thus, we hypothesize that the role of phosphorylation in the kinase reaction may be to specifically stabilize the peptide phosphoacceptor group.



    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cellular proliferation is controlled by a family of protein kinases in which the catalytic subunits are members of the cyclin-dependent kinase (CDK)1 family and the regulatory subunits are cyclins. To date, nine distinct CDKs in addition to eight different cyclins have been identified, in which different CDK/cyclin combinations serve to regulate distinct points in the mammalian cell division cycle. Although Cdc2 (CDK1)/cyclin B controls the transition of cells from the G2 to M-phase, the activities of CDK2/cyclin E and CDK2/cyclin A are critical for G1/S-phase transition and progression through S-phase, respectively (1). Since the critical role of the CDKs in cell cycle control has been well established, understanding the details of their regulation is now of fundamental importance.

The three-dimensional structures of several forms of CDK2 have been solved by x-ray crystallography. Like all protein kinases, CDK2 displays a globular fold consisting of two lobes, a smaller N-terminal lobe that is principally beta -sheet and a larger C-terminal lobe that is principally alpha -helix. The bilobal interface constitutes the active site cleft into which the adenine ring of substrate ATP is deeply buried. The ATP gamma -phosphate is directed toward the mouth of the active site where peptide and protein substrates bind and where phosphoryl group transfer occurs (for a review see Ref. 2). Located near the mouth of the active site is a conserved loop structure termed the activation loop (residues 146-166). This loop structure is present in all protein kinases (3), and phosphorylation at a conserved site within the activation loop is necessary for full activation of most protein kinases. In CDK2, this site is Thr160, phosphorylation of which is catalyzed by a heterologous kinase, CAK (cdk-activating kinase) (4, 5).

Activation of CDK2 requires binding to its regulatory subunit, cyclin, in addition to phosphorylation at Thr160 (5). The structural consequences of cyclin binding to CDK2 and phosphorylation at Thr160 in CDK2 have been revealed by x-ray crystallography. Interaction with cyclin A results in the repositioning of an active site helix and the consequent alignment of key catalytic residues in CDK2, including the invariant residues Lys33 and Glu51, which function to stabilize the alpha - and beta -phosphates of ATP, and Asp145, which chelates an essential Mg2+ ion also serving to stabilize ATP. Furthermore, the active site cavity of CDK2 is exposed upon cyclin binding by repositioning of the activation loop by over 15 Å(6, 7). Subsequent phosphorylation of the CDK2/cyclin A complex at Thr160 (in CDK2) is associated with less dramatic changes in structure that are localized primarily to the activation loop (8). Nonetheless, the latter modification is associated with a dramatic increase in catalytic power.

Although abundant structural information regarding the activation of CDK2 is available, it is not known how the associated alterations in structure correlate with increased catalytic rate. In particular, it is not known which steps along the catalytic reaction pathway are altered in response to Thr160 phosphorylation to achieve kinase activation. Thus, it has not been possible to make a correlation between CDK structure and regulation. In this study, we describe the catalytic reaction pathway for both the unphosphorylated and Thr160-phosphorylated CDK2/cyclin A complexes and report the kinetic basis for activation by phosphorylation. Our results show that phosphorylation affects mainly the rate of chemistry, doing so by specific stabilization of the protein phosphoacceptor group.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Protein Expression and Purification

Unphosphorylated CDK2/cyclin A ((non-p)CDK2/Cyclin A)-- Cdk2 was expressed as the full-length human CDK2 N-terminally fused to GST in pGEX-2T. Cyclin A was expressed as a truncated fragment of bovine cyclin A3, (corresponding to human cyclin A residues Val171 to the end) fused to a C-terminal hexahistidine tag in pET21d (9). Escherichia coli BL21 (DE3) transformed with either GST-CDK2 or cyclin A-His6 were grown at 37 °C in LB to an A600 of 0.6-0.8. Expression of GST-CDK2 was then induced with IPTG (0.4 mM) at room temperature for 18 h, whereas expression of cyclin A-His6 was induced with IPTG (0.2 mM) at room temperature for 3 h. Cell pellets from 1 liter of each culture were separately resuspended in 10 ml of lysis buffer (20 mM MOPS, pH 7.4 50 mM NaCl, 1 mM EDTA), mixed, and colysed in the presence of 1 mM dithiothreitol, 1 µg/ml each of leupeptin, pepstatin, aprotinin, 0.5 mM phenylmethylsulfonyl fluoride, 100 mM MgCl2, 1 mM benzamidine, and 0.2 mg/ml lysozyme for 20 min on ice, sonicated briefly (five 20-s bursts), and then centrifuged at 15,000 rpm (Sorvall SS-34) for 30 min. The supernatant was batch-incubated with 0.5 ml glutathione-agarose (Sigma)/liter of GST-CDK2 culture for 1 h at 4 °C. The resin was washed with buffer A (20 mM MOPS, 0.1 EDTA, 25 mM NaCl, 1 mM dithiothreitol, pH 7.4), and bound protein was eluted in a column using buffer A containing 15 mM glutathione (Sigma). The eluted protein (2-5 mg/ml) was cleaved with thrombin (Sigma, 100 units/ml) in a total volume of 2 ml at room temperature overnight. The cleaved material was digested with Dnase I (Sigma, 0.2 µg/ml) for 15 min at room temperature and then loaded onto a 1-ml Uno Q column (Bio-Rad Biologics). Protein was eluted by a gradient of increasing ionic strength (0.02-0.3 M NaCl/40 ml). The elution profile was monitored by absorbance at A280, peak fractions were analyzed by SDS-PAGE, and those containing protein subunits corresponding to CDK2 and cyclin A were pooled, dialyzed against buffer A, and chromatographed again on Uno Q anion exchange. Fractions corresponding to the major peak at A280 were analyzed for purity by SDS-PAGE followed by assays for ATPase activity and kinase activity toward Peptide 1.

CAK1-- The cDNA clone encoding the Saccharomyces cerevisae CAK1 protein fused to an N-terminal hexahistidine tag was kindly provided by D. Morgan (University of California, San Francisco). CAK1 was subcloned into pGEX-2T, and overexpression of GST-CAK1 protein was carried out in E. coli BL21 (DE3) according to Kaldis et al. (10). Ten liters of bacterial culture were grown to an A600 of 0.6 and induced with IPTG (0.4 mM) at room temperature for 20 h, the cells were harvested, and an extract was prepared as described for (non-p)CDK2/cyclin A with the exception that MgCl2 was present at 5 mM. GST-CAK1 was enriched by glutathione-agarose chromatography (0.5 ml packed resin/liter of culture) as described for (non-p)CDK2/cyclin A.

[T160-P]CDK2/Cyclin A-- GST-CDK2 and cyclin A-His6 were separately purified. GST-CDK2 was purified by glutathione-agarose chromatography and subjected to thrombin cleavage. Cyclin A-His6 was purified by chromatography over Ni2+-NTA-resin according to Brown et al. (9) except that 100 mM MgCl2 was included during lysis as well as throughout purification. Cdk2 (0.8 mg) was phosphorylated at Thr160 by incubation with GST-CAK1, ATP (1 mM), and MgCl2 (10 mM) overnight at room temperature based on a protocol used by Brown et al. (11). Pyruvate kinase (7.5 units/ml) and phosphoenolpyruvate (1 mM) were included to remove product ADP and to regenerate ATP. After phosphorylation, [p-Thr160]CDK2 was combined with cyclin A (4 mg), incubated on ice for 30 min, purified on Superose 12, and then twice subjected to chromatography over Uno Q as described for (non-p)CDK2/cyclin A. From 0.8 mg of CDK2, ~100 µg of highly purified [p-Thr160]CDK2/cyclin A was obtained. [p-Thr160]CDK2/cyclin A expressed in Sf9 cells was purified according to Russo (12).

Activation of (non-p)CDK2/Cyclin A-- (non-p)CDK2/cyclin A was tested for its ability to undergo activation by CAK. (non-p)CDK2/cyclin A (0.5 µM) was incubated with a concentrated extract of glutathione-agarose-enriched GST-CAK1 in the presence of 1 mM ATP, 10 mM MgCl2, 20 mM MOPS, 1 mM phosphoenolpyruvate, 7.5 units/ml pyruvate kinase, 50 mM KCl, 1 mM dithiothreitol, and 0.1 mM EDTA, pH 7.4, (100 µl total volume) for 14 h at room temperature. Ten µl of the reaction mixture was then diluted into 90 µl of buffer containing 1 mM ATP and coupling reagents (15 units/ml lactate dehydrogenase, 7.5 units/ml pyruvate kinase, 1 mM phosphoenolpyruvate, and 130 µM NADH), and peptide kinase activity was measured after the addition of 100 µM Peptide 1 in a coupled spectrophotometric assay (see below). The kinase activity was taken as the change in absorbance at A340 after correction for ATPase activity, which was determined by measurement in the absence of peptide.

Kinetic Assays and Data Analysis-- The phosphorylation of Peptide 1 was monitored by a radioisotope assay in which the direct incorporation of 32P from [gamma -32P]ATP into the peptide was measured. Reactions were carried out in phosphorylation buffer (20 mM MOPS, 50 mM KCl, 10 mM MgCl2(free),1 mM dithiothreitol, 0.1 mM EDTA, pH 7.4) containing enzyme and varying amounts of one substrate with the other held fixed. Assays on [p-Thr160]CDK2/cyclin A contained 0.5 mg/ml bovine serum albumin. In the case of (non-p)CDK2/cyclin A, an ATP regeneration system consisting of phosphoenolpyruvate (1 mM) and pyruvate kinase (7.5 units/ml) was employed. The decay in [gamma -32P]ATP specific radioactivity over time follows first-order kinetics with rate constant k = [rate of ADP formation]/[ATP].2 It was assumed that the rate of ADP formation was effectively equal to the rate of ATP hydrolysis and that ADP generated from the kinase reaction was insignificant. The rate of ADP formation at a given concentration of ATP and enzyme was therefore calculated by solving for the Michaelis-Menten equation using Km and kcat values of 90 µM and 10 min-1, respectively, for the ATPase reaction, which were determined in a spectrophotometric assay (see below; data not shown). The calculated rate constant corresponding to each ATP concentration was used to simulate a decay profile using the equation, y = yo × e-kt, where y is the ATP-specific activity at any time, t, yo is the original ATP-specific activity, and k is the rate constant for decay, as defined above. The average ATP-specific radioactivity at any time, t, was obtained by determining the ATP-specific activity time integral for the entire reaction time course, dividing by the total time. Calculations were performed using the program Scientist (Micromath Inc., Salt Lake City, UT). Reactions were initiated by the addition of [gamma -32P]ATP (300-500 cpm/pmol) and allowed to proceed at 23 °C for 6 to 30 min, at which time reactions were terminated with 25% acetic acid. Reaction samples (20 µl) were spotted and subjected to ascending chromatography on phosphocellulose paper (Whatman P81) in 20 mM H3PO4, pH 2. In this system, ATP and Pi migrate with the solvent front; the phosphorylated peptide product remains at the origin. Papers were dried, and radioactivity corresponding to the phosphopeptide spots were excised and quantified by Cerenkov counting.

Steady-state kinetic parameters for the phosphorylation of Peptide 1 were determined from initial velocity data obtained from a matrix of several fixed ATP concentrations at varied peptide substrate concentrations. Equation 1 was globally fit to the data using the program Scientist (Micromath Inc.).
v=V · A · B/((K<SUB>iA</SUB> · K<SUB>mB</SUB>)+(K<SUB>mB</SUB> · A)+(K<SUB>mA</SUB> · B)+(A · B)) (Eq. 1)
where v is the initial velocity, V is the maximal initial velocity, A and B are the concentrations of the fixed and varied substrates, respectively, Km is the Michaelis constant, and KiA is the dissociation constant for A. kcat was determined by dividing the maximal initial velocity by the enzyme concentration.

ATPase activity was monitored using a coupled spectrophotometric assay in which the regeneration of ATP from ADP and phosphoenolpyruvate catalyzed by pyruvate kinase is coupled to the reduction of pyruvate by NADH to form lactate and NAD+, the latter of which is catalyzed by lactate dehydrogenase (13). The concentrations of the coupling reagents were as follows: 15 units/ml lactate dehydrogenase, 7.5 units/ml pyruvate kinase, 1 mM phosphoenolpyruvate, and 130 µM NADH. Reactions were performed in phosphorylation buffer with coupling reagents in a total volume of 100 µl at 23 °C. Progress of the reaction over time was monitored as a linear decrease in absorbance at 340 nm in a Shimadzu UV1601 spectrophotometer. The micromolar change in product concentration was calculated based on an extinction coefficient for NADH of 6220 cm-1 M-1 at 340 nm. Steady-state kinetic parameters were determined by nonlinear least-squares analysis using the Michaelis-Menten equation, which was fit to the velocity data.

Solvent Viscosity Studies-- Steady-state assays were carried out as described above in buffer containing varied sucrose or fructose. Relative solvent viscosity (eta rel) was determined from the following equation: eta rel = t/to · rho /rho o, where eta rel is the solvent viscosity relative to buffer containing no viscosogen, t is the solvent transit time measured by an Ostwald capillary viscometer, and rho  is the solvent specific gravity. The superscript "o" denotes the absence of viscosogen.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Purification and Steady-state Kinetic Analysis-- [p-Thr160]CDK2/cyclin A was produced by overexpression in either E. coli or Sf9 insect cells. Steady-state kinetic parameters for the phosphorylation of a peptide substrate displaying the canonical recognition motif XT/SPXK/R (Peptide 1; PKTPKKAKKL) were determined by Michaelis-Menten analysis using a two-substrate model for sequential binding (see "Materials and Methods"). The data are shown in Fig. 1 in double reciprocal form. The regression yields a turnover number (kcat) of 7 ± 0.7 s-1 and Km values for ATP and peptide equal to 55 ± 5 µM and 8 ± 0.7 µM, respectively. The optimized steady-state parameter values yield a catalytic efficiency for peptide that is among the highest reported for protein kinases (kcat/Km(peptide) = 0.9 µM-1 s-1). The kinetic properties of [p-Thr160]CDK2/cyclin A produced in Sf9 cells (12) were also determined and were found to be similar (kcat = 4.5 ± 1 s-1, kcat/Km(peptide) = 0.4 µM-1 s-1).



View larger version (18K):
[in this window]
[in a new window]
 
Fig. 1.   Steady-state kinetic analysis of phosphorylation by [p-Thr160]CDK2/cyclin A. Initial velocities for the phosphorylation of Peptide 1 were determined as described under "Materials and Methods." A model describing two-substrate sequential binding was fit to the untransformed data using global nonlinear regression analysis. The experimental data and the fit were then transformed to the double reciprocal form for display. The concentration of enzyme was 2 nM. ATP concentrations were fixed at 20, 50, 100, 200, and 500 µM (from top to bottom). Kinetic parameters are reported in Table I.

(non-p)CDK2/cyclin A was expressed and purified from E. coli. SDS-PAGE analysis of fractions corresponding to the major protein peak eluting from the final chromatographic step is shown in Fig. 2. Each fraction was assayed for ATPase activity, which was found to correspond exactly to the profile of CDK2/cyclin A protein. However, analysis of the same fractions for peptide kinase activity revealed that, while the early fractions of the protein peak displayed low activity, the later fractions exhibited substantially higher kinase activity (Fig. 2, B and C). The basis for the high kinase activity in these later fractions is not known. For all studies on [un-P]CDK2/cyclin A, fractions free of the contaminating high kinase activity species were used (fractions 31-34, Fig. 2). In these fractions, peptide kinase activity corresponded exactly to the levels of CDK2/cyclin A protein and steady-state kinetic parameters, measured for both fraction 31 and fractions 31-34 pooled, were identical (not shown).



View larger version (20K):
[in this window]
[in a new window]
 
Fig. 2.   Anion exchange chromatography of (non-p)CDK2/cyclin A. (non-p)CDK2/cyclin A was produce by overexpression in E. coli and purified as described under "Materials and Methods." A, A280 of eluted protein and buffer conductivity from Uno Q anion exchange chromatography are shown. B, SDS-PAGE analysis of protein-containing fractions. (non-p)CDK2 and cyclin A migrate with the same relative mobility. Molecular mass markers are 97, 66, 55, 42/40, 31, 21.5, and 14.4 kDa from top to bottom. C, peptide kinase activity. Five µl of each fraction was assayed in the presence of 100 µM peptide and 1 mM [gamma -32P]ATP (~300 dpm/pmol) for 10 min at 23 °C, as described under "Materials and Methods." Although two peaks of CDK2/cyclin A are observed in panel A, the steady-state kinetic parameters measured for both fraction 31 and the pool of fractions 31-34 were identical (not shown). The pool was thus used for all studies.

(non-p)CDK2/cyclin A in these fractions displayed low but measurable kinase activity. The kinase activity of (non-p)CDK2/cyclin A] was linear for up to 2 h, demonstrating that autoactivation does not occur within this time frame. To test the possibility that the low level of activity may be attributable to partial denaturation, (non-p)CDK2/cyclin A was tested for activation by phosphorylation (see "Materials and Methods"). Incubation of (non-p)CDK2/cyclin A with MgATP and CAK resulted in a turnover rate (~9 s-1) corresponding to that of [p-Thr160]CDK2/cyclin A obtained after extensive purification (data not shown). These data indicate that all of the (non-p)CDK2/cyclin A was activable and was therefore native. In addition, the maximal ATPase rates of (non-p)CDK2/cyclin A produced in E. coli (10 min-1) compared with insect cells (~14 min-1) were similar, as were their peptide kinase activities.

It was found that the maximal ATPase rate (10 min-1) was ~20-fold higher than the maximal rate of the peptide kinase reaction (0.5 min-1). This finding presented two problems for kinetic analysis. First, (non-p)CDK2/cyclin A binds ADP with a Kd value of ~1 µM (14). Thus, under initial velocity conditions with respect to the kinase reaction, significant product inhibition would be expected to occur. Second, the fraction of ATP depleted at low initial ATP concentrations would be large even at small fractions of peptide phosphorylation, precluding kinetic measurement under initial velocity conditions. To circumvent these difficulties, an ATP-regenerating system was employed. In this system, ADP is efficiently removed from the reaction mix and the concentration of ATP remains constant. However, under these conditions, the [gamma -32P]ATP-specific radioactivity decays over the reaction time course as ATP is regenerated. This decay was accounted for in all analyses of peptide phosphorylation (see "Materials and Methods"). The corrected initial velocity data for the phosphorylation of Peptide 1 by (non-p)CDK2/cyclin A are shown in Fig. 3, and the optimized steady-state kinetic parameter values are reported in Table I. The turnover rate for (non-P)CDK2/cyclin A is 843-fold lower than that of [p-Thr160]CDK2/cyclin A, whereas the Km(peptide) is 137-fold higher. Thus, phosphorylation of CDK2/cyclin A results in an ~100,000-fold increase in catalytic efficiency.



View larger version (20K):
[in this window]
[in a new window]
 
Fig. 3.   Steady-state kinetic analysis of phosphorylation by (non-p)CDK2/cyclin A. Initial velocities for the phosphorylation of Peptide 1 were determined as described under "Materials and Methods," and kinetic parameters were obtained as described in the legend to Fig. 1. The concentration of enzyme was 1 µM. Peptide concentrations were fixed at 0.5, 1, 2, 4, and 6 mM (from top to bottom), and ATP was varied. Kinetic parameters are reported in Table I.


                              
View this table:
[in this window]
[in a new window]
 
Table I
Kinetic and thermodynamic constants for (non-p)CDK2/cyclin A and [p-Thr160]CDK2/cyclin A

Solvent Viscosity Studies-- The Michaelis-Menten parameters described above are composed of microscopic rate constants combined in a manner dependent upon the order of substrate addition. The steady-state data for both [p-Thr160]CDK2/cyclin A and (non-p)CDK2/cyclin A are consistent with both random and compulsorily ordered mechanisms. For both enzymes, however, if the kinetic mechanism is ordered, it is necessarily ordered with ATP binding first; this is true because the crystal structures of both [p-Thr160]CDK2/cyclin A and (non-p)CDK2/cyclin A were obtained with bound ATP alone. Furthermore, both enzymes display substantial ATPase activity in the absence of peptide or protein substrate.

Under saturating conditions of ATP, the catalytic mechanism of CDK2/cyclin A can therefore be described by Scheme 1. In this model, the catalytic efficiency is given by (kcat/Km(peptide) k2 × k3/(k-2 + k3)], while the turnover rate is given by [kcat = k3 × k4/(k3 + k4)]. To solve for the microscopic constants, k2, k-2, k3, and k4, we employed steady-state solvent viscosometric techniques, which allow separation of the diffusive (k2, k-2, k4) from non-diffusive steps (k3) (15-17). Initial velocity data for peptide phosphorylation by [p-Thr160]CDK2/cyclin A were obtained as a function of peptide substrate concentration at several concentrations of sucrose (Fig. 4A). At these sucrose concentrations, the maximum rate3 of ATP hydrolysis (12 min-1) in the absence of peptide was constant, indicating that sucrose does not perturb the structure of the active site. Equation 2,
v=E·(S/K<SUB>d</SUB>·k<SUB>3</SUB>·k<SUB>4</SUB>/&eegr;)/(k<SUB>4</SUB>/&eegr;+k<SUB>4</SUB>·k<SUB>3</SUB>/k<SUB>−2</SUB>+ (Eq. 2)

S/K<SUB>d</SUB>·(k<SUB>3</SUB>+k<SUB>4</SUB>/&eegr;))
which describes the effect of relative solvent viscosity on initial rate, was fit to the data, where E is the concentration of enzyme, S is the concentration of Peptide 1, eta  is the relative solvent viscosity, and Kd is the equilibrium dissociation constant for Peptide 1 binding to the E·ATP complex (k-2/k2). All other constants are defined in Scheme 1. The derived values for the kinetic constants are reported in Table I. Solvent viscosity studies performed on [p-Thr160]CDK2/cyclin A produced in insect cells revealed similar kinetic parameters (Kd(peptide) = 20 µM, k3 = 8 s-1, k4 = 11 s-1).



View larger version (4K):
[in this window]
[in a new window]
 
Scheme 1.  



View larger version (17K):
[in this window]
[in a new window]
 
Fig. 4.   Effect of relative solvent viscosity on Michaelis-Menten parameters. A, [p-Thr160]CDK2/cyclin A. Equation 2, corresponding to Scheme 1, was fit to a matrix of initial velocity data obtained with varied peptide substrate concentrations at several fixed concentrations of sucrose (giving relative solvent viscosities of 1, 2, 2.5, and 4.2, from top to bottom). The optimized microscopic rate constants are given in Table I. The concentration of ATP was fixed at 1 mM. Identical results were obtained at 2 mM ATP. The concentration of enzyme was 2 nM. B, (non-p)CDK2/cyclin A. Relative changes in initial rates were determined as a function of increasing sucrose concentration at three different subsaturating concentrations of peptide substrate (0.4 mM, solid line, open circles; 2 mM, long dashes, open squares; 4 mM, short dashes, open diamonds). The concentration of ATP was 1 mM. Identical results were obtained at 2 mM ATP. The concentration of enzyme was 1 µM. The "viscosity effect" on the reaction rate (kobseta ) is defined as the slope of the line at a given substrate concentration and can vary between the theoretical limits of 0 and 1. The viscosity effects on catalytic efficiency (kcat/Kmeta ) and turnover rate (kcateta ) are obtained by extrapolation to zero and infinite substrate concentration, respectively. kcat/Kmeta and kcateta relate to the individual rate constants in Scheme 1 as follows: kcat/Kmeta  = k3/(k-2 + k3); kcateta  = k3/(k3 + k4)(25). A value for kcat/Kmeta and kcateta equal to zero implies that k-2 k3 and k3 k4, respectively.

A similar solvent viscosometric analysis was conducted on (non-p)CDK2/cyclin A. Although saturation with Peptide 1 could not be achieved because of the high Km value for this substrate, subsaturating substrate concentrations equal to 0.36, 1.8, and 3.6 times the Km(peptide) revealed no viscosity effect on initial rates (Fig. 4B). Extrapolation to zero and infinite peptide substrate concentrations showed that neither the catalytic efficiency nor turnover rate, respectively, was sensitive to solvent viscosity. The release rates for both substrate peptide (k-2) and product (k4) can therefore be assumed to exceed the rate of chemistry (k3) by at least 10-fold (Table I). Thus, phosphoryl group transfer in (non-p)CDK2/cyclin A is rate determining. A viscosometric analysis was also carried out on the ATPase reaction. No viscosity effect on the turnover rate was observed, suggesting that phosphoryl group transfer in the ATPase reaction is also rate determining (Table I). The lack of a viscosity effect on the steady-state kinetic parameters for both the kinase and ATPase reactions indicates that sucrose perturbs only the rates of diffusion.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The majority of protein kinases require phosphorylation at a conserved site within their activation loops for full catalytic activation (3). This site in CDK2/cyclin A is Thr160. Its phosphorylation by CAK is of interest for at least two reasons: 1) it has been established that phosphorylation at this site is a critical regulatory mechanism for control of CDK2/cyclin A activity and progression through the cell cycle (5); 2) the structural basis for activation of CDK2 by both cyclin binding and phosphorylation is known (6-8), and the structure of [p-Thr160]CDK2/cyclin A bound to a synthetic peptide substrate has been solved (18). Given the abundant structural data, the consequences of phosphorylation at Thr160 on the specific steps for substrate binding and catalytic turnover have not been investigated. Thus, a correlation between atomic structure and the mechanism of regulation is not known. In this study, we have characterized the kinetic reaction pathway for the phosphorylation of a model peptide substrate by unphosphorylated CDK2/cyclin A as well as CDK2/cyclin A phosphorylated at Thr160 by CAK. The information reveals the kinetic basis for activation by phosphorylation in terms of the individual reaction steps.

We have employed a synthetic peptide (Peptide 1) as a model substrate in which the primary structure (PKTPKKAKKL) is patterned after a region of similar sequence found in histone H1 protein (19). The catalytic efficiency of Peptide 1 for [p-Thr160]CDK2/cyclin A is one of the highest reported for protein kinases (kcat/Km sime  1 µM-1 s-1) and is similar to that of histone H1 protein (not shown). The lack of a viscosity effect on kcat/Km(peptide) for both [un-P]CDK2/cyclin A and [p-Thr160]CDK2/cyclin A demonstrates that during steady-state turnover Peptide 1 equilibrates rapidly with the active site of both enzymes. Thus, catalytic efficiency for the kinase activity in both cases is a function of only the affinity of peptide binding and the rate of phosphoryl group transfer (kcat/Km sime  k3/Kd). Phosphorylation at Thr160 increases catalytic efficiency by ~100,000-fold, while the turnover rate is enhanced nearly 1000-fold.

The dramatic increase in catalytic efficiency is in part attributable to a moderate but significant increase in peptide binding affinity. The crystal structure of [p-Thr160]CDK2/cyclin A bound to AMPPNP and a peptide substrate (HHASPRK) (18) reveals the structural basis for interaction with peptide and protein substrates. Phosphorylation of CDK2/cyclin A at Thr160 results in displacement of the activation loop by 5.3-7.1 Å and rotation of the carbonyl oxygen of Val163 out of the so called "P+1 binding pocket" (7, 8). This movement is necessary to accommodate binding of the essential proline residue found in the P+1 position (P0 is the phosphorylation site) in all CDK2/cyclin substrates (18). In addition, the substrate lysine residue at position P+3 forms a hydrogen bond with the phosphate group in [p-Thr160]CDK2/cyclin A (18). Overall, phosphorylation of CDK2/cyclin A at Thr160 increases the binding affinity of Peptide 1 by ~50-fold.

Nonetheless, the large increase in both catalytic efficiency and turnover rate with respect to the kinase pathway is attributable mostly to an approximate 3000-fold increase in the rate of the phosphoryl group transfer step. In contrast, phosphorylation has no effect on the rate of ATP hydrolysis. Solvent viscosometric studies on the ATPase reaction of both [p-Thr160]CDK2/cyclin A and (non-p)CDK2/cyclin A show that phosphorylation affects neither the overall turnover rate nor the rate of the phosphoryl group transfer step (Table I).

The large rate enhancement of chemical reactions afforded by enzymes is classically accounted for by specific stabilization of the transition state with respect to the ground state Michaelis complex (20). Thus, the 3000-fold increase in the rate of phosphoryl group transfer may be attributed to optimization of the alignment of either ATP or peptide, or both, in response to phosphorylation. Phosphorylation does not, however, optimize the alignment of ATP for phosphotransfer in the ATPase reaction, because no rate enhancement of this reaction step was observed; this is consistent with the observation that phosphorylation does not affect the positioning of the invariant Lys33, Glu51, and Asp145 catalytic triad, which serves to stabilize the phosphates of ATP. Instead, the alignment of these residues is achieved by cyclin binding (2).

These results raise the possibility that phosphorylation may serve specifically to align the peptide or protein portion of the transition-state structure for phosphotransfer in the kinase reaction. The dramatically slower rate of phosphotransfer to peptide in (non-p)CDK2/cyclin A compared with [p-Thr160]CDK2/cyclin A may be explained by an unfavorable orientation of the peptide substrate hydroxyl group as a consequence of the steric interference from the carbonyl oxygen of Val163, which is relieved by phosphorylation at Thr160. We are currently conducting studies to test our hypothesis that the alignment of ATP versus that of the peptide/protein substrate may be separately controlled by cyclin binding versus phosphorylation at Thr160, respectively.

Substrate turnover by [p-Thr160]CDK2/cyclin A is partially (65%) limited by product release (k4 = 11 s-1). However, our studies do not discern whether the slow step in this event is the release of ADP or phosphopeptide. If the dissociation rate of the phosphopeptide product is similar to, or greater than, that of the peptide substrate (k-2 >=  220 s-1), then it is the release of ADP that is slow. In addition, we cannot assess the effect that phosphorylation at Thr160 has on the product dissociation rates, as the lack of a viscosity effect on kcat in (non-p)CDK2/cyclin A permits only a lower limit value on the net rate of product release to be estimated (k4 >=  0.083 s-1).

Studies addressing the role of autophosphorylation in the activation of cAMP-dependent protein kinase (PKA) have been carried out employing a nonphosphorylatable mutant as a model of the unphosphorylated enzyme (21). Although the site of phosphorylation in PKA (Thr197) is homologous to residue Thr160 in CDK2/cyclin A, clear differences exist between these enzymes in their kinetic mechanisms of activation. In particular, phosphorylated wild-type PKA, in comparison with PKA(T197A), displays no change in substrate binding affinity, a small increase in the overall rate of substrate turnover (<10-fold), and a moderate increase in the rate of the phosphoryl group transfer step (<200-fold) (21) (cf. Table I). The differences in activation of PKA versus CDK2/cyclin A may relate to their different physiological mechanisms of regulation. For example, reversible phosphorylation of the CDKs by CAK is critical to their function in cell cycle control (5), whereas the role of autophosphorylation in the regulation of PKA remains unclear (22, 23). Instead, the regulation of PKA is achieved mainly by association with either the RI or RII regulatory subunits. Comparison of CDK2/cyclin A with PKA (21) and the v-fps tyrosine kinase (24) does, however, reveal an apparent common theme, that phosphorylation of the activation loop in all cases serves mainly to enhance the rate of phosphotransfer with less influence on the affinity of substrate binding.


    FOOTNOTES

* 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.

§ These authors made equal contributions to this work.

Dagger Dagger To whom correspondence should be addressed. Tel.: 805-893-5336; Fax: 805-893-4724; E-mail: lew@lifesci.ucsb.edu.

Published, JBC Papers in Press, October 11, 2000, DOI 10.1074/jbc.M007337200

2 -dATP32/dt = V  ·  ATP32/(ATP32 + Km  ·  (1 + ATP/Km)) = k  ·  ATP32 and -dATP/dt = V  ·  ATP/(ATP + Km  ·  (1 + ATP32/Km)) = k  ·  ATP, where k approx  V/(ATP + Km) and therefore: k = (-dATP/dt)/ATP = (dADP/dt)/ATP.

3 No change in rate was observed at 1 versus 2 mM ATP.


    ABBREVIATIONS

The abbreviations used are: CDK, cyclin-dependent kinase; (non-P)CDK2/cyclin A, unphosphorylated CDK2/cyclin A; [p-Thr160]CDK2/cyclin A, CDK2/cyclin A phosphorylated at Thr160 in CDK2; PKA, cAMP-dependent protein kinase; IPTG, isopropyl-1-thio-beta -D-galactopyranoside; MOPS, 4-morpholinepropanesulfonic acid; PAGE, polyacrylamide gel electrophoresis; CAK, CDK-activating kinase; GST, glutathione S-transferase.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES


1. Morgan, D. O. (1997) Annu. Rev. Cell Dev. Biol. 13, 261-291[CrossRef][Medline] [Order article via Infotrieve]
2. Pavletich, N. P. (1999) J. Mol. Biol. 287, 821-828[CrossRef][Medline] [Order article via Infotrieve]
3. Johnson, L. N., Noble, M. E., and Owen, D. J. (1996) Cell 85, 149-158[Medline] [Order article via Infotrieve]
4. Kaldis, P. (1999) Cell. Mol. Life Sci. 55, 284-296[CrossRef][Medline] [Order article via Infotrieve]
5. Morgan, D. O. (1995) Nature 374, 131-134[CrossRef][Medline] [Order article via Infotrieve]
6. De Bondt, H. L., Rosenblatt, J., Jancarik, J., Jones, H. D., Morgan, D. O., and Kim, S. H. (1993) Nature 363, 595-602[CrossRef][Medline] [Order article via Infotrieve]
7. Jeffrey, P. D., Russo, A. A., Polyak, K., Gibbs, E., Hurwitz, J., Massague, J., and Pavletich, N. P. (1995) Nature 376, 313-320[CrossRef][Medline] [Order article via Infotrieve]
8. Russo, A. A., Jeffrey, P. D., and Pavletich, N. P. (1996) Nat. Struct. Biol. 3, 696-700[Medline] [Order article via Infotrieve]
9. Brown, N. R., Noble, M. E., Endicott, J. A., Garman, E. F., Wakatsuki, S., Mitchell, E., Rasmussen, B., Hunt, T., and Johnson, L. N. (1995) Structure 3, 1235-1247[Medline] [Order article via Infotrieve]
10. Kaldis, P., Sutton, A., and Solomon, M. J. (1996) Cell 86, 553-564[Medline] [Order article via Infotrieve]
11. Brown, N. R., Noble, M. E., Lawrie, A. M., Morris, M. C., Tunnah, P., Divita, G., Johnson, L. N., and Endicott, J. A. (1999) J. Biol. Chem. 274, 8746-8756[Abstract/Free Full Text]
12. Russo, A. A. (1997) Methods Enzymol. 283, 3-12[CrossRef][Medline] [Order article via Infotrieve]
13. Cook, P. F., Neville, M. E., Jr., Vrana, K. E., Hartl, F. T., and Roskoski, R., Jr. (1982) Biochemistry 21, 5794-5799[Medline] [Order article via Infotrieve]
14. Heitz, F., Morris, M. C., Fesquet, D., Cavadore, J. C., Doree, M., and Divita, G. (1997) Biochemistry 36, 4995-5003[CrossRef][Medline] [Order article via Infotrieve]
15. Brouwer, A. C., and Kirsch, J. F. (1982) Biochemistry 21, 1302-1307[Medline] [Order article via Infotrieve]
16. Blacklow, S. C., Raines, R. T., Lim, W. A., Zamore, P. D., and Knowles, J. R. (1988) Biochemistry 27, 683-733
17. Adams, J. A., and Taylor, S. S. (1992) Biochemistry 31, 8516-8522[Medline] [Order article via Infotrieve]
18. Brown, N. R., Noble, M. E. M., Endicott, J. A., and Johnson, L. N. (1999) Nat. Cell Biol. 1, 438-443[CrossRef][Medline] [Order article via Infotrieve]
19. Felix, M. A., Labbe, J. C., Doree, M., Hunt, T., and Karsenti, E. (1990) Nature 346, 379-382[CrossRef][Medline] [Order article via Infotrieve]
20. Hackney, D. D. (1990) The Enzymes , Vol. 19 , pp. 1-36, Academic Press, New York
21. Adams, J. A., McGlone, M. L., Gibson, R., and Taylor, S. S. (1995) Biochemistry 34, 2447-2454[Medline] [Order article via Infotrieve]
22. Cauthron, R. D., Carter, K. B., Liauw, S., and Steinberg, R. A. (1998) Mol. Cell. Biol. 18, 1416-1423[Abstract/Free Full Text]
23. Cheng, X., Ma, Y., Moore, M., Hemmings, B. A., and Taylor, S. S. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 9849-9854[Abstract/Free Full Text]
24. Saylor, P., Hanna, E., and Adams, J. A. (1998) Biochemistry 37, 17875-17881[CrossRef][Medline] [Order article via Infotrieve]
25. Nakatani, H., and Dunford, H. B. (1979) J. Am. Chem. Soc. 83, 2662-2665


Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.