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Indian Journal of Chemistry Vol. 50A, March-April 2011, pp. 355-362 Advances in Contemporary Research Substrate orientation and the origin of catalytic power in xanthine oxidoreductase Hongnan Cao, James Pauff† & Russ Hille* Department of Biochemistry,The University of California, Riverside, CA 92521, USA Email: [email protected] Received 23 July 2010; accepted 25 August 2010 With the chemical course of the reaction catalyzed by the molybdenum-containing hydroxylase xanthine oxidoreductase now relatively well-understood, efforts in the field have now turned to understanding the catalytic power of the enzyme in the context of its structure. The present minireview is an account of recent efforts, from the authors’ laboratory and elsewhere, towards understanding the role of active site amino acid residues in accelerating reaction rate. On the basis of recent site-directed mutagenesis work, in conjunction with protein X-ray crystallography, it is now possible to attribute the specific extent to which each contributes to transition state stabilization and the means by which this occurs. Keywords: Bioinorganic chemistry, Catalysis, Reaction mechanisms, Molybdenum enzymes, Xanthine oxidoreductase Xanthine oxidoreductase is the eponymous member of a large family of molybdenum-containing enzymes that catalyze the oxidative hydroxylation of aromatic heterocycles and aldehydes. The enzyme has been extensively studied for over a century, but it is only in the past 10-15 years that a clear picture of how the reaction proceeds and the structural context in which this occurs has been achieved. The structure of the bovine enzyme1 shows a homodimer of 295 kDa with the four redox-active centers of each subunit laid out in discretely folding domains, from the N-terminus: a first [2Fe-2S]containing domain with a fold closely resembling spinach ferredoxin; a second [2Fe-2S]-containing domain consisting principally of α-helical rather than β-sheet secondary structure; an FAD-containing domain with a fold resembling that seen in vanillyl oxidase family of flavoproteins; and a C-terminal molybdenum-containing portion that consists of two large, oblong domains lying across one another with the molybdenum center at the domain-domain interface. The dimer interface is in this last portion of the polypeptide. The X-ray structure of the xanthine dehydrogenase from Rhodobacter capsulatus has also been determined2, and found to be very similar in overall protein architecture to the bovine enzyme, and particularly so with regard to the conserved amino —————— † Present affiliation: College of Medicine, The Ohio State University, Columbus, OH 43210, USA. acid residues of the active site. One important difference is that the R. capsulatus protein is an α2β2 heterotetramer rather than an α2 dimer like the bovine enzyme, with the molybdenum center in a separate subunit from the Fe/S- and flavincontaining parts of the protein. This notwithstanding, the corresponding portions of the two proteins have a very high degree of structural homology. The molybdenum center of oxidized xanthine oxidoreductase is best represented as LMoVIOS(OH), where L is a pyranopterin cofactor coordinated to the metal via an enedithiolate side chain that is common to all mononuclear molybdenum- and tungsten-containing enzymes3,4. The coordination geometry of the molybdenum center is that of a distorted square pyramid. Although the initial X-ray crystal structures of xanthine oxidoreductase1 and the related aldehyde oxidoreductase from Desulfovibrio gigas5 placed the sulfido group in the apical position, subsequent magnetic circular dichroism studies of xanthine oxidoreductase6 in conjunction with a comparison with subsequently determined crystal structures of other enzymes of the xanthine oxidoreductase family7,8, have unequivocally demonstrated that it is the oxo rather than sulfido group that occupies the apical position. Furthermore, X-ray absorption spectroscopic studies have shown that the equatorial oxygen is present as a singly protonated hydroxide rather than the doubly protonated water that was originally proposed9. 356 INDIAN J CHEM, SEC A, MARCH-APRIL 2011 Fig. 1—The reaction mechanism of xanthine oxidase. The reaction is initiated by proton abstraction from the Mo-OH group by Glu 1261, followed by nucleophilic attack on the carbon to be hydroxylated and hydride transfer to the Mo=S. This leads to the LMoIVO(SH)(OR) intermediate described in the text, which breaks down by electron transfer out of the molybdenum center and hydroxide displacement of product from the molybdenum coordination sphere. The precise sequence of these last two events depends on the reaction conditions and substrate used. The figure was generated using PyMol (PDB accession no. 1FIQ). The reaction mechanism of xanthine oxidase as currently understood is shown in Fig. 1. It has been known for some time that while water is the ultimate source of the oxygen atom incorporated into the hydroxyl group of product in the course of the reaction10, there is a catalytically labile site on the enzyme that represents the proximal donor of the catalytically introduced hydroxyl group11. This catalytically labile oxygen has been shown to be the equatorial Mo-OH rather than the apical Mo=O12, a suggestion that had been made previously on the basis of a comparison of the properties of a series of inorganic model compounds to the enzyme active site13. The pH dependence of the reaction of xanthine oxidoreductase is bell-shaped, reflecting ionizations with pKas of 6.6 and 7.4; the former has been attributed to an active site base, and the latter to substrate itself (reflecting the fact that enzyme works on the neutral rather than monoionic form of substrate)14. By analogy to the reaction mechanism proposed for the D. gigas aldehyde oxidoreductase, in which it was proposed that a strictly conserved active site glutamate residue functioned as a general base in catalysis15, the reaction of xanthine oxidoreductase is thought to proceed via proton abstraction of the equatorial Mo-OH by the catalytic glutamate residue3,12, with nucleophilic attack of Mo-O- on the carbon to be hydroxylated and concomitant hydride transfer to the Mo=S group to give an initial intermediate that can be formulated as IV LMo O(SH)(OR), where OR represents nowhydroxylated product (uric acid in the case of xanthine as substrate) coordinated to the molybdenum via the catalytically introduced hydroxyl group. It is noteworthy that earlier work with a homologous series of substituted quinazolines had in fact suggested that the reaction mechanism involved nucleophilic attack on substrate16. The LMoIVO(SH)(OR) intermediate breaks down by displacement of product from the molybdenum coordination sphere by hydroxide from solvent, with electron transfer from the molybdenum to the other redox-active centers of the enzyme accompanied by deprotonation of the Mo-SH to return to the Mo=S of oxidized enzyme. The sequence in which these events occur depends on the reaction conditions and the substrate utilized; when partial reoxidation precedes displacement of substrate, an LMoVOS(OR) intermediate forms that gives rise to the so-called “very rapid” EPR signal that has long been recognized as being mechanistically important17,18. Under most conditions, however, product dissociation is rate-limiting, with the rate of the chemical step of the reaction, leading to the LMoIVO(SH)(OR) intermediate, faster by a factor of approximately 75 (ref. 19). It is also important to recognize that formation of this species is an oxidative event from the standpoint of the molybdenum center, and that the initial intermediate is formally a MoIV species20. Although it has been suggested from time to time that the MoV species might be formed directly by direct one-electron transfer from substrate, the lack of correlation between reduction potential and reaction rate in a homologous series of substituted purines indicates that this is not likely to be the case21. With the reaction mechanism thus defined, the question that remains is the origin of the catalytic power of the enzyme, that is, the specific means by which the transition state is stabilized in order to accelerate reaction rate. What follows here is first a description of the active site of the enzyme, followed by a consideration of several protein structures with substrate or substrate analogs bound in the active site that, with complementary site-directed mutagenesis studies, provide clues CAO et al.: SUBSTRATE ORIENTATION & ORIGIN OF CATALYTIC POWER IN XANTHINE OXIDOREDUCTASE 357 as to the specific roles of amino acid residues in transition state stabilization. Structure of the active site The structure of the active site of xanthine oxidoreductase1, showing the several amino acid residues implicated in catalysis, is given in Fig. 2. The substrate binding site is defined by Phe 914 and Phe 1009 (interacting with substrate in a side-on and face-on fashion, respectively), which together constrain the substrate to a well-defined plane relative to the molybdenum center. There are surprisingly few interactions between the polypeptide and the first shell of the molybdenum coordination sphere, with Gln 767 within hydrogen-bonding distance of the apical Mo=O (3.12 Å nitrogen to oxygen) and Glu 1261, the putative active site base, which lies within hydrogen-bonding distance of the equatorial Mo-OH of the molybdenum coordination sphere (3.05 Å oxygen to oxygen). Gln 767 is conserved among all eukaryotic members of the xanthine oxidoreductase family of molybdenum enzymes, but is not strictly conserved in the bacterial aldehydeoxidizing enzymes. Somewhat further removed from the molybdenum center is Glu 802 (with its nearer oxygen 3.6 Å from the apical Mo=O and 4.1 Å from the equatorial Mo-OH oxygen) and Arg 880 (with the nearer guanidinium nitrogen 7.6 Å from the Mo-OH oxygen). Both residues are universally conserved among xanthine-oxidizing enzymes, but not among the closely related ones utilizing aldehyde substrates. Site-directed mutagenesis studies Two heterologous expression systems for functional xanthine oxidoreductase have been developed, that of the rat enzyme in a baculovirus system by Nishino and coworkers22, and the xanthine dehydrogenase from R. capsulatus in E. coli by Leimkühler and coworkers23. (A second expression system in E. coli for the human protein yields soluble protein, but of very low specific activity (<5%) owing to ineffective maturation of the molybdenum center24) Using the rat protein from the baculovirus system, it is possible to make assignments for the two [2Fe-2S] clusters in the enzyme, it having long been known that two distinct EPR signals arise from the enzyme25. Fe/S I which is readily seen at temperatures as warm as 150 K has relatively sharp features and g1,2,3 = 2.022, 1.932 that are typical of [2Fe-2S] clusters. By contrast, Fe/S II has much broader features and g1,2,3 = 2.110, 1.991 and 1.894 (ref. 26) Fig. 2—Structure of the active site of bovine xanthine oxidase. Specific amino acid residues involved in substrate binding and catalysis, as described in the text, are shown. The figure was generated using PyMol (PDB accession no. 1FIQ). and is only observed below 50 K. It has been known for some time that the reduced Fe/S I interacts magnetically with the molybdenum center when the latter is in the Mo(V) oxidation state27, but in the crystal structure of the protein it is the cluster in the unusual α-helical domain rather than the more normal ferredoxin-like domain that lies closer to the molybdenum center. Using the rat xanthine oxidoreductase from the baculovirus system, Nishino and coworkers28 were able to demonstrate that mutation of cysteine residues in the ferredoxin-like domain perturbed the Fe/S II EPR signal, while similar mutations in the α-helical domain perturbed the Fe/S I signal. Thus, counterintuitively, the [2Fe-2S] cluster with the more normal EPR characteristics was shown to reside in the more unusual polypeptide fold. Mutagenesis of amino acid residues in the active site have been largely restricted to work with the R. capsulatus system, where adequate amounts of the recombinant proteins are available for detailed kinetic characterization. Leimkühler et al.29 demonstrated that mutation of Glu 232 (equivalent to Glu 802 in the bovine enzyme) to Ala reduced kcat in steady-state experiments from 108 s-1 to 4.4 s-1, and kred in the anaerobic reduction of enzyme by xanthine from 67 to 5.5 s-1. Very similar results were seen in the steady-state kinetics of an E803V mutant of the E. coli-expressed human xanthine oxidoreductase24. In the rapid reaction work with the R. capsulatus enzyme, the affinity of enzyme for substrate also decreased by a factor of approximately 10, with Kd increasing from 34 to 409 µM; kred/Kd, reflecting the second-order reaction of free enzyme with free substrate through the first irreversible step of 358 INDIAN J CHEM, SEC A, MARCH-APRIL 2011 the reaction (in this case, through to formation of the LMoIVO(SH)(OR) intermediate) decreased by two orders of magnitude from 1.97 × 106 M-1s-1 to 1.34 × 104 M-1s-1. At face value, the effect on kred and Kd is such that the interaction between Glu 232 and substrate provides some 3 kcal/mol of free energy, approximately half of which is utilized to bind substrate (as reflected in the higher Kd seen in the mutant) and half to stabilize the transition state (as reflected in the lower kred exhibited by the mutant). Given that the chemical step of the reaction is likely to become rate-limiting in the mutant, however, and assuming that with wild-type enzyme it is not (and to the same degree as is seen with the vertebrate enzymes, i.e., by a factor of 75), it is likely that the actual free energy available from the interaction is ~5.5 kcal/mol, of which 4 kcal/mol is utilized to stabilize the transition state for the chemical step of the reaction rather than simply bind the substrate. On the basis of a computational study of the relative stabilities of several tautomers of free xanthine and the LMoIVO(SH)(OR) intermediate, it has been suggested that Glu 232 accelerates reaction rate specifically by facilitating a tautomerization in the course of nucleophilic attack that involves proton transfer from N3 to N9 of the purine ring and serves to compensate for the negative charge accumulating on the imidazole subnucleus of the purine in the course of the reaction30. This is illustrated in Fig. 3. Glu 197 (Glu 767 in the bovine enzyme) was also mutated to an alanine. This residue hydrogen-bonds to the apical Mo-O of the molybdenum coordination sphere and does not constitute part of the substrate binding site. Consistent with this, in rapid reaction studies Kd for substrate was unchanged, while the limiting rate constant for reduction at high [xanthine] was reduced by a factor of seven from that seen with wild-type enzyme (15 s-1 versus 108 s-1, respectively). Again, given the likelihood that there has been a Fig. 3—The proposed role of R. capsulatus xanthine dehydrogenase Glu 232 (Glu 802 in the bovine enzyme) in facilitating tautomerization in the course of the reaction. change in rate-limiting step in the overall reaction, from product dissociation to the initial formation of the LMoIVO(SH)(OR) intermediate as discussed above, the actual effect of the mutation on the rate constant for the first step of the reaction may be as great as 500-fold, implying that the Glu 197 hydrogen bond to the Mo=O contributes several kcal/mol to transition state stabilization, presumably by modulating the electronic structure (and hence reactivity) of the molybdenum center. A series of mutants at Glu 730, the putative active site base equivalent to Glu 1261 in the bovine enzyme, were also examined in this study. In no case was discernible activity detected in steady-state assays, nor was any detectable reduction observed in anaerobic reductive half-reaction experiments. Even taking into account the variable amount of functional molybdenum center in the mutants (as low as 33 % in the case of the E730A mutant, as compared with 76 % in the wild-type enzyme), the loss of activity was profound. Whereas 100 µM xanthine reduces the wild-type enzyme to completion in ~100 ms, the E730A was not perceptibly reduced in an overnight incubation; conservatively, this amounts to at least a 107-fold reduction in the limiting rate of enzyme reduction. Clearly, Glu 730 is extremely important catalytically, contributing at least 10 kcal/mol in transition state stabilization. That Glu 730 is working specifically as an active site base is suggested by the fact that the E730A mutant is partially active at pH 10, where the Mo-OH is expected to spontaneously deprotonate (Ibdah & Hille, unpublished data). The final active site residue examined has been is Arg 310 (Arg 880 in the bovine enzyme), which has been mutated to methionine31. This residue lies over 7 Å from the catalytically labile Mo-OH and was expected to be involved primarily in substrate binding rather than transition state stabilization. Surprisingly, the principal effect was a 4000-fold reduction in kred, indicating that Arg 310 contributed significantly to transition state stabilization, by at least 4.5 kcal/mol. Consistent with these results, a homologous R881M mutant in the human enzyme lacked activity in steady-state assays24. Interestingly, in a comparison of a homologous series of purine substrates, all hydroxylated at the 8 position as is xanthine, it was found that the substrates fell into two categories: a first, including xanthine, that were effective substrates for wild-type enzyme and which were profoundly affected by the R310M mutation, and a second consisting of poor substrates for wild-type enzyme but which were minimally affected by the mutation31. CAO et al.: SUBSTRATE ORIENTATION & ORIGIN OF CATALYTIC POWER IN XANTHINE OXIDOREDUCTASE 359 Fig. 4—The proposed role of R. capsulatus xanthine dehydrogenase Arg 310 (Arg 880 in the bovine enzyme) in transition state stabilization in the course of nucleophilic attack. Fig. 5—Proposed good and bad substrate orientations in the active site of xanthine oxidase31. On the basis of these results, it has been suggested that Arg 310 stabilizes the transition state for the chemical step of the reaction by providing charge compensation for the negative charge accumulating on the heterocycle, as illustrated in Fig. 4. Structural studies of substrate binding The above proposals for the catalytic roles of Glu 232/802 and Arg 310/880 depend critically on substrate being positioned in the active site as shown in Figs 3 and 4, with confined to the plane defined by the two phenylalanine residues of the active site. An upside down orientation such as shown in Fig. 5, prevents both residues from functioning effectively, and is the proposed structural basis for the lower activity exhibited by some purines relative to others31. The question is whether this is in fact the case. The first crystallographic evidence as to the orientation of substrate in the active site came from the structure of the R. capsulatus enzyme in the complex with the inhibitor alloxanthine2. Alloxanthine, a pyrazolopyrimidine analog of the purine xanthine, is a mechanism-based inhibitor of xanthine oxidoreductase that has long been known to form a very stable complex with the reduced form of the enzyme32. In the crystal structure, alloxanthine is oriented in the active site as shown in Fig. 6. There is a direct bond between N-2 of the pyrazole subnucleus of the inhibitor and the molybdenum (displacing the Fig. 6—The structure of reduced R. capsulatus xanthine dehydrogenase in complex with the inhibitor alloxanthine. Specific amino acid residues are shown. The figure was generated using PyMol (PDB accession no. 1JRP). catalytically labile Mo-OH from the molybdenum coordination sphere), and its pyrimidine subnucleus is oriented with the “distal” C-6 keto group (equivalent to the C-2 keto group of xanthine, owing to the different numbering conventions for the two heterocycles) pointing toward Arg 310. Interestingly, in a complex of alloxanthine with the oxidized rather than reduced (bovine) enzyme, inhibitor is bound in the opposite orientation (Fig. 5), with the “proximal” C-4 carbonyl (equivalent to C-6 of purine substrates) 360 INDIAN J CHEM, SEC A, MARCH-APRIL 2011 interacting with Arg 880 (Nishino T & Pai E, personal communication). In this structure, as expected, there is no bond between the N-2 nitrogen and molybdenum. On the basis of the above discussion of purine substrates, this second orientation represents the catalytically effective one; in the first (with reduced enzyme), as a result of the closer position required for direct coordination of inhibitor to the molybdenum, the heterocycle binds in an inverted orientation, with the “distal” rather than “proximal” keto group of the pyrimidine subnucleus interacting with the active site arginine. In order to explicitly examine substrate orientation in the active site and test the validity of the proposed catalytic roles of the various active site residues, we have examined the X-ray crystal structures of the bovine enzyme in complex with a variety of substrates and substrate analogs. The first to be examined was 2-hydroxy-6-methylpurine, a slow substrate for the enzyme20. Using this poor substrate, the structure shown in Fig. 7A is observed33 (at a resolution of 2.3 Å), in which bridging electron density between the molybdenum and C-8 of the heterocycle is clearly seen. This can only mean that catalysis has been initiated in the crystal, with the observed structure that of the LMoIVO(SH)(OR) intermediate or its LMoVOS(OR) oxidation product (probably the latter since the observed Mo-S distance of 2.0 ± 0.2 Å suggests a Mo=S rather than MO-SH ligand). The orientation of substrate, and in particular the Mo-C distance of 3.4 Å, is generally consistent with ENDOR studies of the “very rapid” species Fig. 7—The X-ray crystal structures of xanthine oxidase in complex with various substrates. [(a) functional enzyme and 2-hydroxy-6methylpurine33, (PDB 3BNJ); (b) nonfunctional, desulfo enzyme and xanthine35, (PDB 3EUB); (c) functional enzyme and lumazine35 (PDB 3ETR); (d) functional enzyme and hypoxanthine in two different orientations of substrate36 (PDB 3NRZ). All figures were generated using COOT]. 361 CAO et al.: SUBSTRATE ORIENTATION & ORIGIN OF CATALYTIC POWER IN XANTHINE OXIDOREDUCTASE which indicated that this interatomic distance was too large for a direct Mo-C bond and that the product was coordinated to the metal in a simple end-on manner34, as is seen crystallographically. In this structure, the asymmetric electron density seen for the substrate in the active site clearly indicates that the orientation of the coordinated product (2,8-dihydroxypurine) in the active site is such that the “distal” C-2 keto group of the pyrimidine subnucleus is oriented toward Arg880, and is within hydrogen-bonding distance of ~ 3.1 Å. 2-Hydroxy-6-methylpurine is a poor substrate for xanthine oxidase and one that is only modestly affected by the R330M mutation in the R. capsulatus enzyme, the orientation seen here is presumably that of the less effective catalytic orientation, and is in agreement with that expected on the basis of the argument above. The next structures that have examined are those of the nonfunctional desulfo form of the enzyme in complex with xanthine (at 2.6 Å resolution, Fig. 7B), and of the functional form in reaction with the pteridine substrate lumazine (at 2.2 Å resolution, Fig. 7C)35. Both these substrates are “good” substrates in the context of the orientation discussion. In both structures, the orientation of the pyrimidine subnucleus of the heterocycle is found to be opposite to that seen with the poor substrate, 2-hydroxy-6methylpurine, with the “proximal” C-6 rather than C-2 oriented toward the active site arginine, 2.9 Å away in the case of the xanthine structure, and 3.5 Å in the case of the lumazine structure. This orientation is that expected to be catalytically more effective on the basis of the argument above, and the prediction that good substrates bind in a different orientation than poor substrates is borne out by the observed crystal structures. Both these structures have subsequently been resolved to better than 1.8 Å resolution. On the basis of the published crystal structures to date, it is evident that purine substrates that are hydroxylated at the 8 position do indeed bind to xanthine oxidase in two different ways, as proposed on the basis of the kinetic work described above. The more catalytically effective orientation is such that Glu 802 is indeed positioned appropriately to function catalytically in facilitating tautomerization between N-3 and N-9, and Arg 880 by stabilizing negative charge accumulation on the C-6 carbonyl of substrate in the course of nucleophilic attack. Physiologically, xanthine oxidase functions not only to hydroxylate xanthine at C-8 to give uric acid, but also hypoxanthine at C-2 to give xanthine in the immediately preceding step of purine catabolism. Glu 802 O N H H N OH N N O Glu 802 -O Mo O H H H N N O N N O- Fig. 8—Proposed role of Glu 802 in facilitating the tautomerization of hypoxanthine in the course of hydroxylation by xanthine oxidase. To examine the role of amino acid residues in catalysis of C-2 of purine substrates, crystal structures of bovine xanthine oxidase in complex with hypoxanthine (at 1.8 Å resolution) and the chemotherapeutic agent 6-mercaptopurine (at 2.6 Å resolution) have been determined36. For each, different orientations of substrate are observed in the two active sites of the dimeric enzyme (there being one dimmer per asymmetric unit in the crystal). One orientation is appropriate for hydroxylation at C-2 of substrate, leading to xanthine or 6-thioxanthine as product (Fig. 7D). The other has C-8 oriented toward the active site molybdenum center (Fig. 7E), as if to give 6,8-dihydroxypurine, a putative product not previously thought to be formed by the enzyme. Separate kinetics studies demonstrate that hypoxanthine is hydroxylated essentially quantitatively at C-2, indicating that the second crystallographically observed orientation is substantially less catalytically effective than the former, so that even when hypoxanthine binds in an orientation that permits hydroxylation at C-8 the reaction proceeds only very slowly36. The apparent enzyme selectivity for C-2 over C-8 with hypoxanthine appears to be due to both relatively subtle differences in substrate orientation in the active site and to differences in the intrinsic reactivity of the two sites; a computational analysis of hypoxanthine indicates that the C-2 carbon is significantly more positively charged than C-8, making it intrinsically more susceptible to nucleophilic attack. Interestingly, these studies show that, in contrast to the situation with xanthine, the preferred tautomer of hypoxanthine with N-9 of the imidazole subnucleus is (modestly) more stable than that with N-7 protonated36. This being the case, Glu 802 is again positioned to facilitate tautomerization in the course of nucleophilic attack, this time from N-9 to N-3 (the reverse sense as proposed in the hydroxylation of xanthine at C-8), as illustrated in Fig. 8. 362 INDIAN J CHEM, SEC A, MARCH-APRIL 2011 Conclusions A combination of site-directed mutagenesis work and crystallographic studies has substantiated the reaction mechanism of xanthine oxidase as it is presently understood and has provided insight into the specific catalytic roles of active site residues. The majority of the catalytic power of the enzyme can be attributed to the action of (1) Glu 1261 (Glu 730 in the R. capsulatus enzyme) as an active site base, deprotonating the Mo-OH group of the molybdenum coordination sphere and thereby facilitating nucleophilic attack on the carbon center to be hydroxylated; (2) Glu 802 (232) in facilitating tautomerization of substrate, thereby stabilizing negative charge accumulation on the heterocycle in the course of nucleophilic attack; and (3) Arg 880 (310) in formal charge compensation in the course of nucleophilic attack on substrate. Together, these three residues are responsible for at least 17 kcal/mol in transition state stabilization (equivalent to twelve orders of magnitude in rate acceleration). Acknowledgement This work was supported by a grant from the US National Institutes of Health (GM 075036 to RH). References 1 2 3 4 5 6 7 8 9 Enroth C, Eger B T, Okamoto K, Nishino T, Nishino T & Pai E, Proc Nat Acad Sci USA, 97 (2000) 10723. 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