MINIREVIEW This paper is available online at www.jbc.org THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 281, NO. 3, pp. 1305–1308, January 20, 2006 © 2006 by The American Society for Biochemistry and Molecular Biology, Inc. Printed in the U.S.A.

(typically five to seven), and a B value that is comparable with that of neighbor A Structural Perspective on atoms. Additional criteria involve proper O–Mϩ distances, on the average 2.4 Å for Naϩ–O and 2.8 Å for Kϩ–O pairs (4), and valence values close to unity Enzymes Activated by (5). Anomalous scattering often provides unequivocal evidence of Kϩ. * Table 1 presents a classification of Mϩ-activated enzymes based on the Monovalent Cations mechanism of activation identified from structural data and on the Mϩ Published, JBC Papers in Press, November 2, 2005, DOI 10.1074/jbc.R500023200 ϩ 1 requirement established from kinetic studies. In Type I enzymes the M func- Enrico Di Cera tions as a cofactor and the requirement is absolute. In Type II enzymes the Mϩ From the Department of Biochemistry and Molecular Biophysics, functions as an allosteric effector and the requirement is not absolute. Washington University School of Medicine, St. Louis, Missouri 63110 K؉-activated Type I Enzymes Enzymes activated by monovalent cations are abundantly represented Diol and glycerol dehydratases provide the simplest example of Type I ϩ ”in plants and the animal world. They have evolved to exploit Na؉ and K؉, enzymes. K is coordinated by five ligands from the protein and acts as a “bait readily available in biological environments, as major driving forces for for the two hydroxyl oxygens from substrate (6, 7). Enzymes involved in phos- substrate binding and catalysis. Recent progress in the structural biology phoryl transfer reactions were long recognized to be the dominant group ϩ of such enzymes has answered long standing questions about the molecu- among M -activated enzymes (2, 3), and this early observation is confirmed by ϩ lar mechanism of activation and the origin of monovalent cation selectiv- the entries in Table 1. In addition to K , these enzymes have an absolute 2ϩ ity. That enables a simple classification of these functionally diverse requirement for a divalent cation, typically Mg (1, 8–11). The mechanism of ϩ 2ϩ enzymes and reveals unanticipated connections with ion transporters. activation involves K and Mg acting in tandem to provide optimal docking for the phosphate moiety of substrate into the protein active site to enable nucleophilic attack on the P␥ or transfer of phosphate groups. The most revealing example of this strategy comes from the ATP-driven folding machine Over 60 years ago, Boyer et al. (1) reported in the pages of this Journal that ϩ ϭ ␮ GroEL, which has the highest K affinity (Kd 80 M) ever reported for a pyruvate kinase would express appreciable catalytic activity only in the pres- ϩ ϩ M -activated enzyme (9). The crystal structure of GroEL bound to ATP ence of K . A similar effect was soon discovered in other systems, and just a 2ϩ ϩ ϩ reveals Mg and K acting in tandem to assist binding of ATP to the protein few decades later the field of enzymes requiring a monovalent cation (M ) for (Fig. 1). Kϩ is coordinated by one P␣ oxygen, the backbone oxygens of Thr-30 optimal activity encompassed hundreds of examples from plants and the ani- and Lys-51, and four water molecules (12). Nucleophilic attack on the P␥ of mal world (2, 3). Since the beginning, this rapidly expanding field had to ϩ ϩ ATP is mediated by Asp-52. K fixes both groups in place through some of its address two basic questions, namely the molecular mechanism of M activa- 2ϩ ϩ ligating water molecules and is assisted by Mg , which anchors all three P tion and the structural basis of M selectivity. Because kinetic investigation groups of ATP. Branched chain ␣-ketoacid dehydrogenase kinase (13) and would only provide indirect answers, further progress in the field had to await ϩ 2ϩ ϩ pyruvate dehydrogenase kinase (14) use a similar K -Mg tandem, whereas high resolution crystal structures of M -activated enzymes, which have the molecular chaperone Hsc70 (15) and the Rad51 recombinase homolog become available only over the last decade. ϩ 2ϩ ϩ from Methanococcus voltae (16) utilize two K in tandem with Mg . Other A classification of M -activated enzymes can be based on the selectivity of variations are the Kϩ-Zn2ϩ tandem in pyridoxal kinase (17) and Kϩ coupled the effect, as established by kinetic studies, and the mechanism of activation, as 2ϩ ϩ with two Mg in fructose-1,6-bisphosphatase (18), S-adenosylmethionine shown from structural analysis. The effect has exquisite specificity, with Na ϩ ϩ ϩ synthase (19), and pyruvate kinase (20). or K being the preferred M . In general, enzymes requiring K such as ϩ ϩ ؉ kinases and molecular chaperones are also activated by NH4 and Rb but are K -activated Type II Enzymes ϩ ϩ not activated as well or at all by the larger cation Cs or the smaller cations Na ϩ ϩ ϩ Some kinases belong to Type II because K does not contact ATP directly. and Li . Enzymes requiring Na such as ␤-galactosidase and clotting pro- ϩ In this case, K exerts its influence indirectly by perturbing the conformation teases are not activated as well by Liϩ or the larger cations Kϩ,Rbϩ, and Csϩ. ϩ ϩ ϩ of active site residues. Ribokinase (21) and aminoimidazole riboside kinase (22) Because the concentration of Na and K is tightly controlled in vivo,M sdo ϩ ϩ ϩ break the K -Mg2 tandem by embracing the M in a ␤-turn adjacent to the not function as regulators of enzyme activity. Rather, they provide a driving ϩ active site. In MutL the K coordination resembles that of branched chain force for substrate binding and catalysis by lowering energy barriers in the ␣-ketoacid dehydrogenase kinase and pyruvate dehydrogenase kinase except ground and/or transition states. Enzymes activated by Mϩs evolved to take for the fact that the contact with the P␤ oxygen of ATP is replaced by a water advantage of the large availability of Naϩ outside the cell and Kϩ inside the cell molecule (23). MutL represents an intriguing intermediate between typical to optimize their catalytic function. Indeed, a strong correlation exists between ϩ ϩ ϩ Type I kinases and ribokinase that evolved a complete separation of K from the preference for K or Na and the intracellular or extracellular localization ϩ ϩ ATP. Here, one can almost envision a transition between enzymes utilizing K of such enzymes. The mechanism of M activation can be established from ϩ ϩ as a cofactor into enzymes utilizing K as an allosteric effector. crystal structures as cofactor-like or allosteric. In the former case, the M ϩ Two K binding sites have been identified in branched chain ␣-ketoacid dehy- anchors the substrate to the active site of the enzyme, often acting in tandem drogenase, one controlling the binding of thiamine diphosphate and the other, also with a divalent cation like Mg2ϩ. In such a mechanism of activation, the Mϩ is ϩ found in pyruvate dehydrogenase (24), stabilizing the quaternary structure (25). absolutely required for catalysis. In the latter, the M enhances enzyme activ- ϩ Dialkylglycine decarboxylase is a beautiful example of M -dependent molecular ity through conformational transitions triggered upon binding to a site where ϩ switch. In this pyridoxal phosphate (PLP)2-dependent enzyme K binds in a loop the Mϩ makes no direct contact with substrate. In this case, the Mϩ is not near the active site (26, 27). Substrate binding requires Tyr-301 to be in the con- absolutely required for catalysis. ϩ ϩ ϩ formation seen in the K -bound form (Fig. 1). When K is replaced by Na , the Crystallographic assignment of Naϩ or Kϩ is non-trivial, even for high res- activity of the enzyme drops 95% and the change in coordination within the loop olution structures. Naϩ has a small ionic radius (0.97 Å) and the same number causes the O␥ of Ser-80 to swing away and clash with the phenyl ring of Tyr-301 of electrons as a water molecule. Kϩ has a higher electron density but an ionic that reorients and compromises substrate binding. Likewise, in other PLP-de- radius (1.33 Å) almost identical to that of a water molecule. Hence, correct ϩ ϩ pendent enzymes like Ser hydratase (28), tryptophanase (29), and tyrosinase (30), assignment of Na and K must be based on several criteria: the presence of a ϩ the K binding site makes no contact with substrate but helps organize the archi- spherical electron density peak at a ␴ level in the F Ϫ F map considerably o c tecture of the active site. above that of all other water molecules, the number of surrounding oxygens Na؉-activated Type I Enzymes The strategy used by Kϩ-activated Type I kinases to anchor substrate to the * This minireview will be reprinted in the 2006 Minireview Compendium, which will be available in January, 2007. This work was supported in part by National Institutes of active site is also exploited by fructose-1,6-biphosphate aldolase (31) and tag- Health Research Grants HL49413, HL58141, and HL73813. 1 To whom correspondence should be addressed. Tel.: 314-362-4185; Fax: 314-747-5354; 2 The abbreviations used are: PLP, pyridoxal phosphate; CPK, Corey-Pauling-Koltun E-mail: [email protected]. model.

JANUARY 20, 2006•VOLUME 281•NUMBER 3 JOURNAL OF BIOLOGICAL CHEMISTRY 1305 MINIREVIEW: Enzymes Activated by Mϩ

TABLE 1 Classification of M؉-activated enzymes Enzyme Protein Data Bank entrya Ligandsb Ref. c K؉-activated Type I Branched chain ␣-ketoacid dehydrogenase 1GJV, 1GKZ 5 (4-0-1) 13 kinase Diol dehydratase 1DIO, 1EEX, 1EGM, EGV, 1IWB, 1UC4, 1UC5 7 (5-0-2) 6 Fructose-1,6-bisphosphatase 1FPI 4 (3-0-1) 18 Glycerol dehydratase 1IWP, 1MMF 7 (5-0-2) 7 GroEL 1KP8, 1PCQ, 1SVT, 1SX3 7 (2-4-1) 12 Hsc70 1BUP, 1HPM, 1KAX, 1KAY, 1KAZ, 1QQM, 1QQN, 1QQO; 8 (2-3-3) 15, 43 Naϩ bound: 1BA0, 1BA1, 1UD0, 3HSC 7 (5-1-1) MvRadA 1XU4 4 (2-1-1) 16 Pyridoxal kinase 1LHR, 1RFT 6 (4-1-1) 17 Pyruvate dehydrogenase kinase 1Y8N, 1Y8O, 1Y8P 5 (4-0-1) 14 Pyruvate kinase 1A3W, 1A3X, 1A49, 1AQF, 1F3W, 1F3X, 1LIU, 1LIW, 1LIX, 6 (4-1-1) 20 1LIY, 1PKN, 1T5A; Naϩ bound: 1A5U S-Adenosylmethionine synthase 1MXA, 1MXB, 1MXC, 1O90, 1O92, 1O93, 1O9T, 1P7L, 4 (3-0-1) 19 1QM4, 1RG9, 1XRA, 1XRB, 1XRC K؉-activated Type II Branched chain ␣-ketoacid dehydrogenase 1DTW, 1OLS, 1OLU, 1OLX, 1U5B, 1V11, 1V16, 1V1M, 1V1R, 5 (5-0-0) 25 1X7W, 1X7X, 1X7Y, 1X7Z, 1X80 Dialkylglycine decarboxylase 1D7R, 1D7S, 1D7U, 1D7V, 1DKA, 1M0N, 1M0O, 1M0P, 6 (5-1-0) 26, 27 1M0Q; Naϩ bound: 1D7R, 1D7S, 1D7U, 1D7V, 1DGD, 1DKA, 1M0N, 1M0O, 1M0P, 1M0Q, 2DKB MutL 1NHI 5 (4-1-0) 23 Ribokinase 1GQT, 1TYY, 1TZ3, 1TZ6 6 (6-0-0) 21, 22 Ser dehydratase 1PWH 6 (6-0-0) 28 Tryptophanase 1AX4 7 (4-3-0) 29 Tyrosinase 1TPL, 2TPL 7 (4-3-0) 30 Na؉-activated Type I ␤-Galactosidase 1DP0, 1JYN, 1JYV, 1JYW, 1JYX, 1JYY, 1JYZ, 1JZ0, 1JZ1, 1JZ2, 5 (3-1-1) 33, 35 1JZ3, 1JZ4, 1JZ5, 1JZ6, 1JZ7, 1JZ8, 1PX3, 1PX4, 1TG7, 1XC6 Fructose-1,6-bisphosphate aldolase 1B57, 1RV8, 1RVG 6 (4-1-1) 31 Tagatose-1,6-bisphosphate aldolase 1GVF 6 (5-0-1) 32 Na؉-activated Type II Factor Xa 1P0S, 2BOK 6 (4-2-0) 41 Thrombin 1A2C, 1A46, 1A4W, 1A5G, 1A61, 1AD8, 1B5G, 1BB0, 1C1U, 6 (2-4-0) 40, 42 1C1V, 1C1W, 1C4U, 1C4V, 1C5L, 1C5N, 1C5O, 1CA8, 1D3D, 1D3P, 1D3Q, 1D3T, 1D4P, 1D6W, 1D9I, 1DE7, 1DOJ, 1DX5, 1GHV, 1GHW, 1GHX, 1GHY, 1GJ4, 1GJ5, 1JM0, 1JOU, 1K21, 1K22, 1O2G, 1O5G, 1OYT, 1SB1, 1SFQ, 1SG8, 1TBZ, 1VZQ, 1XMN, 1Z8I, 1Z8J, 2THF, 7KME, 8KME; Kϩ bound: 2A0Q Trp synthase 1A50, 1A5S, 1BKS, 1C29, 1C8V, 1C9D, 1CW2, 1CX9, 1FUY, 4 (3-1-0) 38, 39 1K3U, 1K7E, 1K7X, 1K8X, 1K8Y, 1K8Z, 1KFC, 1KFE, 1KFJ, 1KFK, 1QOP, 1UBS, 1V8Z, 2TRS, 2TSY, 2TYS, 2WSY; Kϩ bound: 1A5A, 1A5B, 1BEU, 1TTQ a From a total of 1508 structures containing Naϩ or Kϩ, as of September 8, 2005. Many Mϩs reported in the PDB have no functional role. In other cases, e.g. lysozyme (53), rhodanese (54), uridine phosphorylase (55), transformylase/cyclohydrolase enzyme (56), homoserine dehydrogenase (57), carbonic anhydrase (58), formyltransferase (59), inosine-monophosphate dehydrogenase (60), ␣-amylase (61), and aldehyde dehydrogenase (62), kinetic evidence of specific activation is controversial or not significant. The Kϩ binding sites in Kex2 do not explain the kinetics of activation (63). b The format is N (p-w-s), where N is the sum of ligands from the protein (p), water (w), and substrate (s). c Only the most relevant references are listed. atose-1,6-bisphosphate aldolase (32), where the tandem Naϩ-Zn2ϩ replaces enabling the indole intermediate to be shuttled to the active site for complex- ϩ ϩ K -Mg2 . Tagatose 1,6-bisphosphate is unique in that it replaces a water ation with L-Ser (38). The tunnel is partially blocked by residues Phe-280 and molecule in the Naϩ coordination shell with a cation-␲ interaction. ␤-Galac- Tyr-279 in the Naϩ form and is more open in the Kϩ form. Long range allos- tosidase from Escherichia coli deserves special mention. As the gene product of teric communication in Trp synthase is further demonstrated by the fact that the lacZ operon, it occupies a most prominent place in the history of molecular binding to the active site in the ␣ subunit can displace Naϩ from its site in the biology (33). The activating effect of Naϩ and Mg2ϩ was discovered by Monod ␤ subunit (39). in 1951 (34). In this enzyme, the interplay between Naϩ and Mg2ϩ is quite In the clotting protease thrombin the allosteric effect of Naϩ affects a basic different from the partnership seen in kinases, with Mg2ϩ binding away from mechanism of substrate recognition. Naϩ binds close to the primary specificity substrate and Naϩ being in contact with the galactosyl 6-hydroxyl (Fig. 1) (33, pocket and orients Asp-189 for correct engagement of the Arg side chain of 35). Naϩ is coordinated by three protein atoms and two water molecules in the substrate at the P1 position (Fig. 1), enabling the enzyme to accomplish its free enzyme, and lactose binding replaces one of the waters in the coordination procoagulant role in the blood (40). Long range effects induced by Naϩ binding shell. This change in Naϩ coordination triggers a rearrangement of Phe-601, propagate through a network of buried water molecules up to the catalytic one of the Naϩ ligands, to promote substrate binding. Ser-195 located 15 Å away (40). A similar architecture of Naϩ recognition is observed in clotting factor Xa (41). Na؉-activated Type II Enzymes ؉ Among the enzymes involved in PLP catalysis, Trp synthase has been stud- M Selectivity ied extensively both structurally and kinetically (36). The enzyme is peculiar in Several Mϩ-activated enzymes have been crystallized free or in the presence that it binds Naϩ with only slightly higher affinity than Kϩ (37). The crystal of Naϩ,Kϩ, or other Mϩ, and the resulting information has broadened our structures of Trp synthase bound to Naϩ or Kϩ show that the Mϩ makes no understanding of Mϩ selectivity. In the case of Trp synthase, the changes contact with substrate or PLP and binds to the ␤ subunit near the tunnel between the Naϩ-bound and Kϩ-bound structures are significant (38) but are

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FIGURE 1. Examples of the four classes of M؉-activated enzymes (see also Table 1). Shown are substrate (CPK, C in yellow), relevant residues (CPK, C in cyan), Kϩ or Naϩ (yellow ball), and Mg2ϩ (green ball). not matched by differences in the kinetics of activation (36). In pyruvate kinase the replacement of Kϩ with Naϩ results in no structural changes (20) although the enzyme is practically inactive without Kϩ (1). Future structural studies will hopefully clarify this lack of correlation with function. In thrombin, however, changes in coordination between Naϩ and Kϩ propagate to the oxyanion hole and explain the differences in the kinetics of activation (40, 42). In the case of Dialkylglycine dehydrogenase (26, 27) and Hsc70 (15, 43), replacement of the essential Kϩ with Naϩ changes drastically the geometry of coordination and perturbs residues that control binding of PLP or ATP. These enzymes have evolved Kϩ selectivity by imposing geometric constraints on the coordination shell that cannot be obeyed by the smaller ionic radius of Naϩ. The linkage with enzyme activation is ensured by the functional connection of these constraints with the optimal orientation of catalytic residues. Rigidity of the coordination shell guarantees selectivity by increasing the entropic cost of any reorganiza- tion meant to accommodate a Mϩ of different size. Interestingly, an analogous strategy has been exploited successfully in the synthesis of selective chelators (44, 45). Other examples are provided by ion transporters for which Mϩ selectivity is absolute. The V-type Naϩ-ATPase (46), the F-type Naϩ-ATPase (47), and the bacterial Naϩ/ClϪ-dependent neurotransmitter homologue (48) cage Naϩ in ϩ ϩ FIGURE 2. Overlay of the signature sequence GYG (residues 77–79) in the selectivity rigid environments, practically inaccessible to K . In the KcsA K channel, the ؉ ؉ ϩ filter of the KcsA K channel (CPK, K yellow balls, 1BL8) with the GYG sequence ,backbone oxygens lining the channel maintain distances suitable only for K (residues 325–327) of pyruvate dehydrogenase kinase (magenta,K؉ magenta ball ϩ coordination and provide an exact replica of the coordination shell of K in 1Y8P), the GFG sequence (residues 337–339) of branched chain ␣-ketoacid dehy- drogenase kinase (green,K؉ green ball, 1GJV), and the KYG sequence (residues solution (49). One striking feature of the channel is the signature sequence ؉ ϩ GYG (residues 77–79) whose backbone oxygens shape part of the selectivity 224–226) of thrombin (wheat,K wheat ball, 2A0Q). K sits in equivalent positions relative to the sequence in all cases. filter (Fig. 2). Remarkably, the conformation of this sequence relative to the bound Kϩ in the channel is very similar to the GYG sequence (residues 325– 327) near the Kϩ binding site of pyruvate dehydrogenase kinase (14), the GFG near the Naϩ binding site of thrombin (42). Furthermore, mutation of Tyr in sequence (residues 337–339) near the Kϩ binding site of branched chain ␣-ke- this sequence has very similar functional consequences in the Kϩ channel (50) toacid dehydrogenase kinase (13), and the KYG sequence (residues 224–226) and thrombin (51). This unexpected connection is a testimony to the basic

JANUARY 20, 2006•VOLUME 281•NUMBER 3 JOURNAL OF BIOLOGICAL CHEMISTRY 1307 MINIREVIEW: Enzymes Activated by Mϩ similarity in the mechanism of Mϩ recognition that evolution has bestowed on 28. Yamada, T., Komoto, J., Takata, Y., Ogawa, H., Pitot, H. C., and Takusagawa, F. (2003) proteins with widely different functions. Biochemistry 42, 12854–12865 29. Isupov, M. N., Antson, A. A., Dodson, E. J., Dodson, G. G., Dementieva, I. S., Zakomir- Concluding Remarks dina, L. N., Wilson, K. S., Dauter, Z., Lebedev, A. A., and Harutyunyan, E. H. (1998) J. ϩ Mol. Biol. 276, 603–623 The information that emerged from crystal structures of M -activated 30. Sundararaju, B., Antson, A. A., Phillips, R. S., Demidkina, T. V., Barbolina, M. V., enzymes rationalizes decades of kinetic studies and unravels details on the Gollnick, P., Dodson, G. G., and Wilson, K. S. (1997) Biochemistry 36, 6502–6510 ϩ molecular origin of M activation and selectivity. A simple classification of 31. Hall, D. R., Leonard, G. A., Reed, C. D., Watt, C. I., Berry, A., and Hunter, W. N. (1999) these enzymes is now made possible by merging structural and functional data J. Mol. Biol. 287, 383–394 bases. Future studies should explore the intriguing connections between 32. Hall, D. R., Bond, C. S., Leonard, G. A., Watt, C. I., Berry, A., and Hunter, W. N. (2002) Mϩ-activated enzymes and ion transporters. The enormous amount of knowl- J. Biol. Chem. 277, 22018–22024 33. Juers, D. H., Jacobson, R. H., Wigley, D., Zhang, X. J., Huber, R. E., Tronrud, D. E., and edge gathered from kinetic and structural studies should facilitate the redesign ϩ Matthews, B. W. (2000) Protein Sci. 9, 1685–1699 of M specificity and activation (23, 52). A most exciting task would be to 34. Cohn, M., and Monod, J. (1951) Biochim. Biophys. Acta 7, 153–174 ϩ ϩ introduce M activation into proteins devoid of M binding, which could 35. Juers, D. H., Heightman, T. D., Vasella, A., McCarter, J. D., Mackenzie, L., Withers, benefit many areas of biotechnology and medicine. S. G., and Matthews, B. W. (2001) Biochemistry 40, 14781–14794 36. Woehl, E. U., and Dunn, M. F. (1995) Coord. Chem. Rev. 144, 147–197 37. Peracchi, A., Mozzarelli, A., and Rossi, G. L. (1995) Biochemistry 34, 9459–9465 Acknowledgments—I am grateful to Drs. David Davies, Thomas Hurley, Yu 38. Rhee, S., Parris, K. D., Ahmed, S. A., Miles, E. W., and Davies, D. R. (1996) Biochem- Luo, Mischa Machius, Barry Stoddard, Michael Toney, Jimin Wang, and Wei istry 35, 4211–4221 Yang for helpful exchanges of information. 39. Weyand, M., and Schlichting, I. (1999) Biochemistry 38, 16469–16480 40. Pineda, A. O., Carrell, C. J., Bush, L. A., Prasad, S., Caccia, S., Chen, Z.-W., Mathews, F. S., and Di Cera, E. (2004) J. Biol. Chem. 279, 31842–31853 41. Scharer, K., Morgenthaler, M., Paulini, R., Obst-Sander, U., Banner, D. W., Schlatter, REFERENCES D., Benz, J., Stihle, M., and Diederich, F. (2005) Angew. Chem. Int. Ed. Engl. 44, 1. Boyer, P. D., Lardy, H. A., and Phillips, P. H. (1942) J. Biol. Chem. 146, 673–681 4400–4404 2. Suelter, C. H. (1970) Science 168, 789–795 42. Papaconstantinou, M. E., Carrell, C. J., Pineda, A. O., Bobofchak, K. M., Mathews, 3. Evans, H. J., and Sorger, G. J. (1966) Annu. Rev. Plant Physiol. 17, 47–76 F. S., Flordellis, C. S., Maragoudakis, M. E., Tsopanoglou, N. E., and Di Cera, E. (2005) 4. Harding, M. M. (2002) Acta Crystallogr. Sect. D Biol. Crystallogr. 58, 872–874 J. Biol. Chem. 280, 29393–29396 5. Nayal, M., and Di Cera, E. (1996) J. Mol. Biol. 256, 228–234 43. Flaherty, K. M., DeLuca-Flaherty, C., and McKay, D. B. (1990) Nature 346, 623–628 6. Shibata, N., Masuda, J., Tobimatsu, T., Toraya, T., Suto, K., Morimoto, Y., and Ya- 44. Minta, A., and Tsien, R. Y. (1989) J. Biol. Chem. 264, 19449–19457 suoka, N. (1999) Structure 7, 997–1008 45. Cram, D. J. (1988) Science 240, 760–767 7. Liao, D. I., Dotson, G., Turner, I., Jr., Reiss, L., and Emptage, M. (2003) J. Inorg. 46. Murata, T., Yamato, I., Kakinuma, Y., Leslie, A. G., and Walker, J. E. (2005) Science Biochem. 93, 84–91 308, 654–659 8. O’Brien, M. C., and McKay, D. B. (1995) J. Biol. Chem. 270, 2247–2250 47. Meier, T., Polzer, P., Diederichs, K., Welte, W., and Dimroth, P. (2005) Science 308, 9. Viitanen, P. V., Lubben, T. H., Reed, J., Goloubinoff, P., O’Keefe, D. P., and Lorimer, 659–662 G. H. (1990) Biochemistry 29, 5665–5671 48. Yamashita, A., Singh, S. K., Kawate, T., Jin, Y., and Gouaux, E. (2005) Nature 437, 10. Mudd, S. H., and Cantoni, G. L. (1958) J. Biol. Chem. 231, 481–492 215–223 11. Wu, Y., Qian, X., He, Y., Moya, I. A., and Luo, Y. (2005) J. Biol. Chem. 280, 722–728 49. Doyle, D. A., Morais Cabral, J., Pfuetzner, R. A., Kuo, A., Gulbis, J. M., Cohen, S. L., 12. Wang, J., and Boisvert, D. C. (2003) J. Mol. Biol. 327, 843–855 Chait, B. T., and MacKinnon, R. (1998) Science 280, 69–77 13. Machius, M., Chuang, J. L., Wynn, R. M., Tomchick, D. R., and Chuang, D. T. (2001) 50. Nakamura, R. L., Anderson, J. A., and Gaber, R. F. (1997) J. Biol. Chem. 272, 98, Proc. Natl. Acad. Sci. U. S. A. 11218–11223 1011–1018 14. Kato, M., Chuang, J. L., Tso, S. C., Wynn, R. M., and Chuang, D. T. (2005) EMBO J. 24, 51. Guinto, E. R., Caccia, S., Rose, T., Futterer, K., Waksman, G., and Di Cera, E. (1999) 1763–1774 Proc. Natl. Acad. Sci. U. S. A. 96, 1852–1857 15. Wilbanks, S. M., and McKay, D. B. (1995) J. Biol. Chem. 270, 2251–2257 52. Prasad, S., Wright, K. J., Roy, D. B., Bush, L. A., Cantwell, A. M., and Di Cera, E. (2003) 16. Wu, Y., He, Y., Moya, I. A., Qian, X., and Luo, Y. (2004) Mol. Cell 15, 423–435 Proc. Natl. Acad. Sci. U. S. A. 100, 13785–13790 17. Li, M. H., Kwok, F., Chang, W. R., Lau, C. K., Zhang, J. P., Lo, S. C., Jiang, T., and Liang, 53. Funahashi, J., Takano, K., Yamagata, Y., and Yutani, K. (2002) J. Biol. Chem. 277, D. C. (2002) J. Biol. Chem. 277, 46385–46390 18. Villeret, V., Huang, S., Fromm, H. J., and Lipscomb, W. N. (1995) Proc. Natl. Acad. Sci. 21792–21800 U. S. A. 92, 8916–8920 54. Lijk, L. J., Torfs, C. A., Kalk, K. H., De Maeyer, M. C., and Hol, W. G. (1984) Eur. 19. Komoto, J., Yamada, T., Takata, Y., Markham, G. D., and Takusagawa, F. (2004) J. Biochem. 142, 399–408 Biochemistry 43, 1821–1831 55. Caradoc-Davies, T. T., Cutfield, S. M., Lamont, I. L., and Cutfield, J. F. (2004) J. Mol. 20. Larsen, T. M., Benning, M. M., Rayment, I., and Reed, G. H. (1998) Biochemistry 37, Biol. 337, 337–354 6247–6255 56. Greasley, S. E., Horton, P., Ramcharan, J., Beardsley, G. P., Benkovic, S. J., and Wilson, 21. Andersson, C. E., and Mowbray, S. L. (2002) J. Mol. Biol. 315, 409–419 I. A. (2001) Nat. Struct. Biol. 8, 402–406 22. Zhang, Y., Dougherty, M., Downs, D. M., and Ealick, S. E. (2004) Structure (Camb.) 57. DeLaBarre, B., Thompson, P. R., Wright, G. D., and Berghuis, A. M. (2000) Nat. 12, 1809–1821 Struct. Biol. 7, 238–244 23. Hu, X., Machius, M., and Yang, W. (2003) FEBS Lett. 544, 268–273 58. Premkumar, L., Greenblatt, H. M., Bageshwar, U. K., Savchenko, T., Gokhman, I., 24. Ciszak, E. M., Korotchkina, L. G., Dominiak, P. M., Sidhu, S., and Patel, M. S. (2003) Sussman, J. L., and Zamir, A. (2005) Proc. Natl. Acad. Sci. U. S. A. 102, 7493–7498 J. Biol. Chem. 278, 21240–21246 59. Mamat, B., Roth, A., Grimm, C., Ermler, U., Tziatzios, C., Schubert, D., Thauer, R. K., 25. Evarsson, A., Chuang, J. L., Wynn, R. M., Turley, S., Chuang, D. T., and Hol, W. G. and Shima, S. (2002) Protein Sci. 11, 2168–2178 (2000) Structure (Camb.) 8, 277–291 60. Gan, L., Petsko, G. A., and Hedstrom, L. (2002) Biochemistry 41, 13309–13317 26. Toney, M. D., Hohenester, E., Cowan, S. W., and Jansonius, J. N. (1993) Science 261, 61. Machius, M., Declerck, N., Huber, R., and Wiegand, G. (1998) Structure 6, 281–292 756–759 62. Perez-Miller, S. J., and Hurley, T. D. (2003) Biochemistry 42, 7100–7109 27. Toney, M. D., Hohenester, E., Keller, J. W., and Jansonius, J. N. (1995) J. Mol. Biol. 245, 63. Holyoak, T., Kettner, C. A., Petsko, G. A., Fuller, R. S., and Ringe, D. (2004) Biochem- 151–179 istry 43, 2412–2421

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