Molecular Dynamics Simulations of Substrate Dephosphorylation by

Chapter 29

Molecular Dynamics Simulations of Substrate Dephosphorylation by Low Molecular Weight Protein Tyrosine Phosphatase Karin Kolmodin, Tomas Hansson, Jonas Danielsson, and Johan Åqvist

Downloaded by UNIV MASSACHUSETTS AMHERST on August 7, 2012 | http://pubs.acs.org Publication Date: April 8, 1999 | doi: 10.1021/bk-1999-0721.ch029

Department of Molecular Biology, Uppsala University, Biomedical Center, Box 590, S-751 24 Uppsala, Sweden

Dephosphorylation of phosphotyrosine by the low molecular weight protein tyrosine phosphatase proceeds via nucleophilic substitution at the phosphorous atom yielding a covalent enzyme-substrate intermediate that is subsequently hydrolyzed. The reactive nucleophile is the thiolate anion of Cys12. Here we calculate the free energy profiles of putative reaction mechanisms by molecular dynamics and free energy perturbation simulations, utilizing the empirical valence bond method to describe the reaction potential surface. Binding calculations addressing the protonation state of the enzyme-substrate complex are also performed. The calculations give a consistent picture of the catalytic mechanism that is compatible with experimental data.

Phosphorylation and dephosphorylation of tyrosine residues in proteins are important regulatory mechanisms involved in cellular processes such as cell growth, proliferation and differentiation (7-7). Protein tyrosine kinases are responsible for the phosphorylation of tyrosine residues, resulting in either activation or deactivation of the substrate protein. The kinases are counteracted by protein tyrosine phosphatases (PTPases) which hydrolyze the tyrosylphosphates. Considerable progress has been made in recent years towards elucidating the catalytic machinery of PTPases and several enzymological studies as well as crystal structures have been reported (8-17). Despite these advances there are, however, a number of fundamental questions regarding the catalytic reaction mechanism and pathway that remain unanswered. In this work we employ the empirical valence bond (EVB) method (18-20) together with molecular dynamics (MD) simulations and free energy perturbation (FEP) to investigate the mechanism of the low molecular weight (low M ) PTPases. r

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© 1999 American Chemical Society

In Transition State Modeling for Catalysis; Truhlar, D., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1999.

371 The low M protein tyrosine phosphatases are small cytosolic enzymes (18,000 Da) without sequence homology to other proteins in the protein tyrosine phosphatase family, such as FTP IB and Yersinia PTPase. However, they possess the active site signature motif C-(X) -R-(S/T) common to all PTPases, where the eight amino acid residues comprise the phosphate binding loop or P-loop. The fact that kinetic properties such as formation of an E-S intermediate, pH-rate profiles etc. are similar for different types of PTPases (21-24) indicates that they employ a common mechanism for catalysis. The physiological substrates of the low M PTPases are yet unknown, but they readily hydrolyze both phosphotyrosyl proteins and peptides as well as small aryl phosphates, and are themselves regulated by tyrosine phosphorylation at position Y131 and Y132 (25, 26). The catalytic reaction has been shown to proceed via a double displacement mechanism (27) involving a phophoenzyme intermediate where the phosphate group becomes covalently bound to the cysteine residue in the active site motif (Cysl2 in low M PTPase) (28-33). The formation of this intermediate is accomplished by a substitution reaction where Cysl2 attacks the phosphorous atom and the leaving group is protonated by an appropriately positioned acid (Asp 129 in low M PTPase) (34, 35). This aspartate residue is thought to subsequently activate a water molecule which hydrolyzes the phosphorylated cysteine in the next step (Figure 1). It is still unclear whether the catalytic cysteine is in its thiol or anionic form in the free enzyme. Zhang et al. (36) measured the pK value of Cysl2 by alkylation with iodoactetate and iodoacetamide. These results suggest a pAT of 6.75-7.52 depending on the reagent used, whereas Evans et al. (24) estimate the pK of the same residue to be below 4.0 from pH-rate profiles. Earlier calculations by us on the low M PTPase have shown that the enzyme environment significantly stabilizes proton transfer from the cysteine residue to one of the substrate (phenylphosphate) oxygens thereby enabling activation of the nucleophile for attack on the phosphate group. It is thus clear that the p^ of Cysl2 is lowered relative to the normal value of cysteine in solution (p/C ~8.3). However, if Cysl2 is already ionized in the free enzyme it seems likely that the substrate then would bind as the monoanion. Otherwise, a total negative charge of three on the reacting groups is expected to lead to strong electrostatic repulsion between the substrate and the nucleophile, thereby displacing the substrate from its optimal binding position. Crystal structures of the Cys->Ser mutant in Yersinia PTPase in complex with S0 " (8) as well as the corresponding mutant in PTP1B in complex with small peptides containing phosphorylated tyrosine (12, 13) also show more or less identical binding conformations compared to the native low M PTPase in complex with S04 ~. Since the serine residue in the two mutants is undoubtedly protonated this indicates that these different complexes all have one proton and bear the same overall charge of -2 on the relevant groups. In the present study we investigate the energetics of the first half of the catalytic reaction in the low M PTPase, i.e. formation of the phosphoenzyme intermediate, using the EVB approach. We address the activation of the reactive cysteine, the nucleophilic substitution at the phosphorous atom and the degree of concertedness of this substitution with leaving group protonation by Asp 129. Calculations on the mutation of Asp 129 to alanine are also reported and suggest an T

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In Transition State Modeling for Catalysis; Truhlar, D., et al.; ACS Symposium Series; American Chemical Society: Washington, DC, 1999. r

Figure 1. The reaction mechanism catalyzed by lowM PTPase.


373 alternative mechanism for this mutant. The protonation state of the reaction complex is examined by binding energy calculations. The results of these different simulations give a detailed picture of the mechanism of catalysis that is consistent with experimental data.


Computational Models and Methods The reaction potential energy surface is represented by the EVB model that has been described in detail elsewhere (18-20). The valence bond structures used in the present calculations are shown in Figure 2 and the reaction is thus modeled in terms of conversions between these different states. For example, nucleophilic activation is described by the process (i->0) and the nucleophilic substitution is represented, e.g., by (2->3->4)- Since the phosphate oxygens in the enzyme are not equivalent, due to restricted rotation as seen in Figure 3, it is necessary to consider three separate VB structures for each state with a singly protonated phosphate group. This leads to a total of 14 different VB states all of which have been considered here. In addition, we also examine some cases with a total charge of -3 on the reacting fragments, but these are not depicted in Figure 2 since it was found that substrate binding does not seem possible in this case (see below). The EVB hamiltonian was calibrated against relevant solution reactions utilizing experimental energetics data as well as semi-empirical (AM1/SM2 and PM3/SM3) and ab initio (HF/6-31G*) calculations of geometries. As described elsewhere (18, 20) this involves determining gas-phase energy difference Aodj as well as off-diagonal matrix elements H between pairs of VB states so that the EVB potential surface reproduces experimental reaction free energies and barrier heights of relevant reference reactions in solution. This calibration procedure thus involves simulations of uncatalyzed reaction steps with the reacting fragments in water and fitting the above parameters so that calculated and observed free energies coincide. In the case of proton transfer steps, such as (Oi->02), (04->Os) and (O4-X&6), the pXa difference between donor and acceptor together with linear free energy relationships (LFERs) for proton transfer states were used as described in (37, 38) for calibration of the relevant EVB parameters. Calibration of nucleophilic displacement steps, such as 2a,b,c) show that there is no significant discrimination of the acceptor. Proton transfer is feasible from the cysteine to any of the three oxygens, with a slight preference for the two which are hydrogen bonded to Argl8 (O3 and O4 in the crystal structure). The position of the proton does not have any major effect on the catalysis of the approach to the transition state (02a,b,c"^ I3a,b,c)- On the other hand, it appears that stabilization of the phosphoenzyme intermediate resulting from leaving group departure is more sensitive to the nature of phosphate protonation. When the proton is bound to oxygen 03, which accepts a hydrogen bond from N of Argl8, it can be engaged in hydrogen bonding to the negatively charged Asp 129. When the proton is bound to 0 the distance to Asp 129 becomes too large to allow such hydrogen bonding. For the third case with the proton bound to O4 we observe a stabilization of the phosphoenzyme intermediate that is somewhere in between the other two cases. 2

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Simulations of the P-0 bond cleavage and leaving group departure clearly indicate that bond cleavage at the bridging oxygen has to be concerted with protonation of the leaving group in order to depress a charge separation in the active site. This is also in agreement with interpretations of experimental data. The bond cleavage was first simulated along the stepwise pathway, 0 3 - ^ 0 4 followed by 4->5, which is predicted to be energetically unfavorable in the enzyme, yielding a barrier of -22 kcal/mol. A developing negative charge on the leaving group oxygen apparently cannot be stabilized by the proton environment and, since the binding cavity is very narrow, solvating water molecules are excluded from the active site. The concerted pathway, (5) is strongly facilitated by the enzyme and the resulting negative charge on Asp 129 is, unlike the phenolate ion, accessible to solvent. The apparent need for concerted leaving group protonation may indicate that the nucleophilic attack in the second half of the overall reaction (Figure 1), where the phosphoenzyme is hydrolyzed by a water molecule, also has to be concerted with proton abstraction by Asp 129 from the water nucleophile. This issue is, however, left for future work. 3

An alternative mechanism for the protonation of the leaving group has also been investigated. Here the phosphate group itself was used as the acid for leaving group protonation ( 0 3 a . b . c " ^ 6 ) resulting in a double negative charge on the cysteinyl phosphate. It appears that also this reaction is significantly catalyzed by the enzyme and it is therefore possible that such a reaction mechanism may be utilized in mutants lacking the general acid/base Asp 129. In order to examine this issue we calculated the free energy profile for substrate dephosphorylation also for the D129A mutant assuming leaving group protonation by the phosphate monoanion. The resulting free



379 energy profile for the D129A mutant (Figure 4) shows that it is considerably less efficient than the wild type enzyme but still retains catalytic power. The difference in activation energy between WT and D129A at the rate limiting transition state for substrate dephosphorylation is approximately 5 kcal/mol, which corresponds to a rate decrease by a factor of 4000 for the mutant. This appears to be consistent with experimental data which show that the D129A mutant is about 2000-3000 fold slower than the wild type (34, 35). The calculated activation energies of both the wild type enzyme and the D129A mutant are thus consistent with kinetic rate experiments, but as noted above, the dephosphorylation of the protein may be the rate limiting step for the wild type enzyme (33, 49). The protonation state of the reaction complex is one of the main issues in elucidating the catalytic mechanism of PTPases. The rates of reaction of substrates such as phenyl phosphate in solution are usually much faster for the monoanion than for the dianion (39). It is also noteworthy that PTPases generally display optimal activity in the acidic pH range. Furthermore, as mentioned above the crystal structures of various inhibitor complexes with WT and catalytic Cys->Ser mutated enzymes generally show van der Waals contact between Cys/Ser and the ligand oxygens. Since some of these complexes, such as the Cys->Ser mutants with various ligands (8, 12, 13) and WT low M PTPase in complex with HEPES (17), clearly have a total charge of -2 on the nucleophile-substrate moiety, it seems reasonable to expect that the corresponding complex of the active enzyme has the same protonation state. The most straightforward way to examine this issue is to try to evaluate the difference in substrate affinity for different protonation states. We performed FEP calculations where the phenylphosphate substrate was transformed from monoanion to dianion in aqueous solution and in the solvated protein with Cysl2 in its anionic form. The calculated difference in binding free energy was -15 kcal/mol for the monoanion to dianion perturbation, indicating that there is no affinity for the substrate dianion with Cysl2 ionized. It can also be seen from the MD structures that the doubly charged substrate is significantly displaced from its initial binding site due to electrostatic repulsion with the nucleophile. With Cysl2 ionized and the substrate in its monoanionic form, on the other hand, the structural agreement between MD and the crystal complexes are excellent. We also performed calculations of binding free energies using the linear response approach described in (57, 52), which confirmed the hypothesis that a doubly negative substrate does not have affinity for low M PTPase with Cysl2 in its anionic form. T

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Discussion In this work we have used MD/FEP/EVB simulations to investigate details of the substrate dephosphorylation mechanism in low M PTPase. The reaction mechanism with leaving group departure concerted with protonation by Asp 129 is found to be fully compatible with experimentally observed reaction rates. It appears that the enzyme is designed to stabilize negative charge more or less precisely in the plane of the so called P-loop where the equatorial phosphate oxygens are positioned during nucleophilic substitution. For a stepwise mechanism where the leaving group departs in its anionic form, and thus moves out of the 'focus' of the P-loop, the calculations T



380 predict very unfavourable energetics due to insufficient solvation of the phenol anion. On the other hand, the concerted protonation by Asp 129 gives significant catalysis and the resulting negative charge on the aspartate becomes more solvent accessible. Simulations of the D129A mutant reaction also give highly unfavourable energetics with phenolate anion on the effective leaving group and suggest that there would be no enzymatic activity of D129A if this were the reaction pathway. However, our calculations predict that for this mutant a mechanism where the protonated phosphate group itself is involved in concerted protonation may be employed. The calculated increase in activation energy (relative to 0 0 for this mechanism is 5 kcal/mol or about a factor 4000 slower catalysis. Hence, it appears that this type of mechanism can explain why the D129A mutant still retains considerable activity. We have found that with a singly protonated Cysl2-phosphate complex the calculated energetics is consistent with experimental data. However, it has been proposed that both the cysteine and the substrate are fully ionized in the reactive complex (23, 55), which would give a total charge of -3 rather than -2 on the reactants. Although such a situation would seem very unfavorable in view of the resulting electrostatic repulsion (the corresponding reaction is not observed at all in solution (59)), one cannot exclude the possibility that the enzyme might stabilize the -3 complex significantly. To address this problem we carried out calculations of the different binding affinity of phenylphosphate mono- and di-anion for the thiolate form of the enzyme. The results, however, clearly predict that the dianion does not bind when Cysl2 is ionized and the cysteine-phosphate distance increases significantly with respect to the typical distances seen in various enzyme-ligand complexes. The fact that pH-rate profiles for ^ t show a descending slope of -1 on the basic side of the pHoptimum with an apparent pK that is substrate dependent and agrees exactly with the p £ of the substrate phosphate group (21, 54), also suggests that the ES complex is singly protonated. In this context it is also interesting to note the recently reported crystal structure by Zhang et al. (17) of the low M PTPase in complex with vanadate. This structure shows a typical trigonal bipyramidal conformation with the vanadate covalently linked to Cysl2. Since vanadate at the given concentration exists as H V04~ in solution between pH 4 and 8.3 (55) it is most likely that the observed complex with low M PTPase actually corresponds to (Cysl2)S-V04H ~. This structure would thus correspond closely to our high-energy intermediate (O3). Furthermore, the reported binding of inorganic phosphate is three times stronger at pH 5 than at pH 7.5 (17) which agrees well with the relative monoanion concentration (33%) at the higher pH value. Regarding the actual pK of Cysl2 the situation is somewhat unclear. While reaction experiments with iodoacetate have indicated that its p £ is as high as 7.5 (36), pH-rate profiles show a typical ionization around pH 4-5 that is usually attributed to the catalytic cysteine. At any rate it is clear that its p £ is lowered in the free enzyme from the normal value of -8.3, which can be explained by the efficient stabilization of negative charge in the active site. Exactly how the two ionizations of Cysl2 and the substrate phosphate groups are shifted upon binding is less clear. However, the characteristic k&x vs. pH rate profile with optimum around 5-6 flanked by an acidic limb with slope +1 and a basic limb with slope -1 at least appears consistent with two possible ionizations of the Cys-phosphate moiety. a

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381 As noted above, the characteristic geometry of the active site with backbone amide groups and the Argl8 side chain focused towards the center of the P-loop appears ideally adapted for stabilizing negative charge in the plane where the equatorial oxygens are positioned in the transition state(s). This situation resembles that encountered, e.g. in the Ras-RasGAP and transducin a structures, where GTP is proposed to be hydrolyzed by analogous mechanisms (56-58). In summary, we have shown that the enzymatic environment catalyzes the reaction pathway where a singly protonated reaction complex of total charge -2 forms a phosphoenzyme intermediate, by almost eight orders of magnitude. The resulting reaction barrier is compatible with experiments and the calculations indicate that the present model of the reaction pathway is a good representation of the actual catalytic process in the low M PTPase. r

Acknowledgments Support from the EC Biotechnology Program DGXII and the NFR is gratefully acknowledged. Literature Cited 1. 2. 3. 4.

5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

Hunter, T. Cell, 1989, 58, 1013-1016. Ramponi, G.; Manao, G.; Camici, G.; Cappugi, G.; Ruggiero, M.; Bottaro, D. P. FEBS letters, 1989, 250, 469-473. Fischer, E. H.; Charbonneau, H.; Tonks, N. K. Science, 1991, 253, 401-406. Ruggiero, M.; Pazzagli,C.;Rigacci, S.; Magnelli, L.; Raugei, G.; Berti, A.; Chiarugi, V. P.; Pierce, J. H.; Camici, G.; Ramponi, G. FEBS letters, 1993, 326, 294-298. Mondesert, O.; Moreno, S.; Russel, P. J. Biol. Chem. 1994, 269, 27996-27999. Barford, D.; Jia, Z.; Tonks, N. K. Nature Struct. Biol. 1995, 2, 1043-1053. Stone, R. L; Dixon, J. E. J. Biol. Chem. 1994, 269, 31323-31326. Stuckey, J. A.; Schubert, H.L.; Fauman, E. B.; Zhang, Z.-Y.; Dixon, J. E.; Saper, M. A. Nature, 1994, 370, 571-575. Su, X.-D.; Taddei, N.; Stefani, M.; Ramponi, G.; Nordlund, P. Nature, 1994, 370, 575-578. Logan, T. M.; Zhou, M. M.; Nettesheim, D. G.; Meadows, R. P.; Van Etten, R. L.; Fersik, S. W. Biochemistry, 1994, 33, 11087-11096. Zhang, M.; Van Etten, R. L.; Stauffacher, C. V. Biochemistry, 1994, 33, 1109711105. Barford, D.; Flint, A. J.; Tonks, N. K. Science, 1994, 263, 1397-1404. Jia, Z.; Barford, D.; Flint, A. J., Tonks, N. K. Science, 1995, 268, 1754-1758. Yuvaniyama, J.; Denu, J. M.; Dixon, J. E.; Saper, M. A. Science, 1996, 272, 1328-1331. Bilwes, A. M.; den Hertog, J; Hunter, T.; Noel, J. P. Nature, 1996, 382, 555559. Fauman, E. B.; Yuvaniyama, C.; Schubert, H. L.; Stuckey, J. A.; Saper, M. A. J. Biol. Chem. 1996, 271, 18780-18788.


382 17. 18.


19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44.

45. 46. 47.

Zhang, M.; Zhou, M.; Van Etten, R.L.; Stauffacher, C.V. Biochemistry, 1997, 36,15-23. Warshel, A. Computer Modeling of Chemical Reactions in Enzymes and Solutions; Wiley, New York, NY, 1991. Warshel, A.; Sussman, F.; Hwang, J.-K. J. Mol. Biol. 1988, 201,139. Åqvist, J.; Warshel, A. Chem. Rev. 1993, 93, 2523-2544. Zhang, Z.-Y.; Malachowski, W. P.; Van Etten, R. L.; Dixon, J. E. J. Biol. Chem. 1994, 269, 8140-8145. Zhang, Z.-Y. J. Biol. Chem. 1995, 270, 11199-11204. Denu, J. M.; Zhou, G.; Guo, Y.; Dixon, J. E. Biochemistry, 1995, 34, 33963403. Evans, B.; Tishmack, P. A.; Pokalsky, C.; Zhang, M.; Van Etten, R. L. Biochemistry, 1996, 35, 13609-13617. Rigacci, S.; Degl'Inocenti, D.; Bucciantini, M.; Cirri, P.; Berti, A.; Ramponi, G. J. Biol. Chem. 1996, 271, 1278-1281. Tailor, P.; Gilman, J.; Williams, S.; Couture,C.;Mustelin, T. J. Biol. Chem. 1997, 272, 5371-5374. Saini, M. S.; Buchwald, S. L.; Van Etten, R. L.; Knowles, J. R. J. Biol. Chem. 1981, 256, 10453-10455. Chiarugi, P.; Marzocchini, R.; Raugei, G.; Pazzagli, C.; Berti, A.; Camici, G.; Manao, G.; Cappugi, G.; Ramponi, G. FEBS letters, 1992, 310, 9-12. Cirri, P.; Chiarugi, P.; Camici, G.; Manao, G.; Raugei, G.; Cappugi, G.; Ramponi, G. Eur. J. Biochem. 1993, 214, 647-657. Guan, K.-L.; Dixon, J. E. Proteins: Struct. Funct. Genet. 1991, 9, 99-107. Zhang, Z.-Y.; Dixon, J. E. Advan. Enzymol. 1994, 68, 1-36. Zhang, Z.-Y.; Wang, Y.; Dixon, J. E. Proc. Natl. Acad. Sci. USA, 1994, 91, 1624-1627. Davis, J. P.; Zhou, M.-M.; Van Etten, R. L. J. Biol. Chem. 1994, 269, 87348740. Zhang, Z.; Harms, E.; Van Etten, R. L. J. Biol. Chem. 1994, 269, 25947-25950. Taddei, N.; Chiarugi, P.; Cirri, P.; Fiaschi, T.; Stefani, M.; Camici, G.; Raugei, G.; Ramponi, G. FEBS letters, 1994, 350, 328-332. Zhang, Z.-Y.; Davis, J. P.; Van Etten, R. L. Biochemistry, 1992, 31, 1701-1711. Åqvist, J.; Fothergill, M. J. Biol. Chem. 1996, 271, 10010-10016. Hansson, T.; Nordlund, P.; Åqvist, J. J. Mol. Biol. 1996, 265, 118-127. Kirby, A. J.; Younas, M. J. Chem. Soc. 1970, 1165-1172. Åkerfeldt, S. Acta Chem. Scand. 1963, 17, 319-328. Bourne, N.; Williams, A. J. Org. Chem. 1984, 49, 1200-1204. Guthrie, J. P. J. Am. Chem. Soc. 1977, 99, 3991-4001. Cramer, C. J.; Truhlar, D. G. Science, 1992, 256, 213-217. van Gunsteren, W. F.; Berendsen, H. J. C. Groningen Molecular Simulation (GROMOS) Library Manual, Biomos B. V., Nijenborgh 16, Groningen, The Netherlands, 1987. Åqvist, J. Q, Version 2.1; Uppsala University, 1996. King, G.; Warshel, A. J. Chem. Phys. 1989, 91,3647-3661. Essex, J.W.; Jorgensen, W. L. J. Comput. Chem. 1995, 16, 951-972.


383 48. 49. 50. 51. 52.


53. 54. 55. 56. 57. 58.

Lee, F. S.; Warshel, A. J. Chem. Phys. 1992, 97, 3100-3107. Zhang, Z.-Y.; Van Etten, R. L. Biochemistry, 1991, 30, 8954-8959. Zhang, Z.-Y.; Van Etten, R. L. J. Biol. Chem. 1991, 266, 1516-1525. Åqvist, J.; Medina,C.;Samuelsson, J.-E. Protein Eng. 1994, 7, 385-391. Marelius, J.; Graffner-Nordberg, M.; Hansson, T.; Hallberg, A.; Åqvist, J. J. Comput.-Aided Mol. Design, 1998, 12, 000-000. Hengge, A. C.; Zhao, Y.; Wu, L.; Zhang, Z.-Y. Biochemistry, 1997, 36, 79287936. Zhang, Z.-Y.; Van Etten, R. L. Arch. Biochem. Biophys. 1990, 282, 39-49. Pope, M. T.; Dale, B. W. Q. Rev. Chem Soc. 1968, 22, 527-548. Scheffzel, K.; Ahmadian, M. R.; Kabsch, W.; Weismüller, L.; Lautwein, A.; Schmitz, F.; Wittinghofer, A. Science, 1997, 277, 333-338. Sondek, J.; Lambright, D. G.; Noel, J. P.; Hamm, H. E.; Sigler, P. B. Nature, 1994, 372, 276-279. Langen, R.; Schweins, T.; Warshel, A. Biochemistry, 1992, 31, 8691-8696.


Molecular Dynamics Simulations of Substrate Dephosphorylation by

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