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© 2015 Nature America, Inc. All rights reserved.
news and views an interesting question. First, uridine is characteristic for RNA and is most divergent from the four DNA-building molecules. Second, it is statistically enriched in RNA loop sections, owing to its ability to form only two hydrogen bonds with its complementing adenine base. Loop sections are considered the first targets of endoribonucleases to degrade RNA elements. Therefore, the probability that U-rich degradation products are made is high. Finally, it is also conceivable that the presence of higher concentrations of Us could serve as a pathogenic pattern during infections because various microbe-associated RNAs are particularly U rich. For example, most positive-strand RNA viruses use negative-strand RNAs with 5′-poly(U) tracts as replication templates. Therefore, abundant amounts of Us are generated during de novo synthesis of polyadenylated progeny RNA in the course of viral replication15. Furthermore, bacterial small noncoding RNAs carry poly(U) tails and have important regulatory functions within bacteria; therefore, they could represent another potential substrate for the generation of single Us during infections16. Given the importance of single-nucleoside recognition for TLR8 activation, it is interesting
that TLRs 7 and 8 were first deorphanized with the discovery of various small-molecule agonists that triggered their activation17. The available crystal structures of TLR8 (refs. 4,5) now provide compelling mechanistic evidence for how small molecules such as the agonistic imidazoquinolines and uridine in synergy with short oligonucleotides can promote protein-protein interaction and reorganization of the dimer into the active conformation. Currently, imidazoquinoline compounds are approved for the topical treatment of actinic keratosis, basal cell carcinoma and certain virus infections. The demonstration that the natural ligand U interacts at the same site as the small-molecule activators provides a strong rationale as well as a valuable template for the development of small-molecule antagonists. Such compounds could target TLR8 and by extension TLR7, which also responds to U-rich RNA and imidazoquinoline agonists. Pharmacologic approaches to inhibit TLRs 7 or 8 could be promising for the treatment of certain autoimmune diseases in which the recognition of host nucleic acids contributes to autoimmunity and devastating inflammation18.
ACKNOWLEDGMENTS The authors are supported by the Deutsche Forschungsgemeinschaft–funded Excellence Cluster ImmunoSensation. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests. 1. Matzinger, P. Science 296, 301–305 (2002). 2. Akira, S. & Takeda, K. Nat. Rev. Immunol. 4, 499–511 (2004). 3. Heil, F. et al. Science 303, 1526–1529 (2004). 4. Tanji, H. et al. Nat. Struct. Mol. Biol. 22, 109–115 (2015). 5. Tanji, H., Ohto, U., Shibata, T., Miyake, K. & Shimizu, T. Science 339, 1426–1429 (2013). 6. Ohto, U., Tanji, H. & Shimizu, T. Microbes Infect. 16, 273–282 (2014). 7. Gorden, K.K.B. et al. J. Immunol. 177, 8164–8170 (2006). 8. Jurk, M. et al. Eur. J. Immunol. 36, 1815–1826 (2006). 9. Ligoxygakis, P., Pelte, N., Hoffmann, J.A. & Reichhart, J.M. Science 297, 114–116 (2002). 10. Tesar, B.M. et al. Am. J. Transplant. 6, 2622–2635 (2006). 11. Chan, M.P. et al. Nat. Commun. (in the press). 12. Pawaria, S. et al. J. Immunol. (in the press). 13. Horiuchi, T., Horiuchi, S. & Mizuno, D. Nature 183, 1529–1530 (1959). 14. Hausser, E. & Stroun, J. Rev. Med. Suisse Romande 83, 577–586 (1963). 15. White, K.A., Enjuanes, L. & Berkhout, B. RNA Biol. 8, 182–183 (2011). 16. Vogel, J. & Wagner, E.G. Curr. Opin. Microbiol. 10, 262–270 (2007). 17. Hemmi, H. et al. Nat. Immunol. 3, 196–200 (2002). 18. Christensen, S.R. & Shlomchik, M.J. Semin. Immunol. 19, 11–23 (2007).
An enzyme cofactor with a split personality Anthony Mittermaier Little is currently known about the molecular determinants of energy barriers along enzyme catalytic pathways. Kern and co-workers have studied this question in adenylate kinase (Adk) and now reveal that a single Mg 2+ ion can accelerate two distinct steps, thus uncovering an unexpected dual role for this ubiquitous cofactor. Many decades of research have been devoted to understanding how enzymes lower the energy barriers for chemical reactions and accelerate rates by many orders of magnitude. Enzyme catalysis often proceeds through multiple discreet steps, each associated with its own energy barrier1 (Fig. 1), and it has become possible to relate the individual steps of catalytic pathways to specific conformational changes occurring in the enzymes themselves by using NMR techniques2–6. Nevertheless, many questions remain to be answered, including those regarding the factors controlling the heights of the barriers. As reported in this issue, Kern and co-workers7 have studied the role of a magnesium (Mg2+) Anthony Mittermaier is at the Department of Chemistry, McGill University, Montreal, Quebec, Canada. e-mail: [email protected]
cofactor in the Adk cycle, which catalyzes the reversible phosphorylation of AMP by ATP. Using a broad combination of biophysical techniques including NMR, they showed that Mg2+ has a surprising dual role: it dramatically accelerates the on-enzyme rate of phosphoryl transfer and also promotes a lid-opening conformational change that is essential for product release. Interestingly, the two effects depend on different aspects of the cofactor’s ‘personality’. The authors found that Mg2+ enhances lid opening primarily by shielding electrostatic interactions, whereas it enhances the chemical step by coordinating the nucleotide phosphate groups and by acting as a pivot during phosphoryl transfer. Some of the first knowledge of the individual steps in enzyme catalytic pathways came from pre-steady-state kinetic assays 8. In this approach, reactions are performed with relatively high concentrations of enzymes,
nature structural & molecular biology volume 22 number 2 FEbruary 2015
such that the buildup of intermediates and/ or products within the active site can be measured before their release from the enzyme. For example, when protein kinase A (PKA) is added to its ATP and peptide substrates, one enzyme equivalent of the phosphopeptide product is formed very rapidly, at a rate far greater than that of steady-state turnover9. This implies that phosphoryl transfer is fast, whereas product release is rate limiting. X-ray crystal structures provided a possible explanation for the slow rate of product release. The enzyme active site adopts both ‘open’ conformations, which are solvent accessible, and ‘closed’ conformations, in which the ATP γ-phosphate is positioned for transfer10. This suggested that product release is predicated on active site opening. However, it is not possible to ascertain whether the conformational transition itself is rate limiting solely on the basis of enzyme kinetic experiments. 101
news and views Figure 1 Schematic representation of enzymatic catalysis. (a) Enzymes (E) increase the rate of conversion of substrates (S) to products (P) by lowering the energy of the transition state (TS) separating the two forms. (b) In many cases, catalytic mechanisms involve a series of discrete steps, each associated with its own energy barrier, for instance, enzyme conformational rearrangements (TS1 and TS3) and chemical transformation (TS2).
a TS
S
P TS E
E
Energy
b S
P E
TS1
S
P
E
E
TS2
TS3
E
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© 2015 Nature America, Inc. All rights reserved.
Energy
Similarly to that of PKA, the Adk active site adopts both open and closed configurations, characterized by ‘lid’ domains that fold over the active site when ligands are bound. The central role of lid dynamics in Adk catalysis was elegantly demonstrated in earlier work by the Kern laboratory3. Because the reaction catalyzed by the enzyme (ATP + AMP ↔ 2 ADP) is reversible, it was possible to perform high-resolution protein NMR experiments on samples actively performing catalysis on substrates and products exchanging in a dynamic equilibrium. The authors found that NMR signals from protein nuclei in the lid regions of Adk are broadened in the presence of saturating concentrations of ligands (Fig. 2a,b), a signature of exchange between different conformations occurring on millisecond-to-microsecond timescales. They characterized the broadening by using recently developed NMR relaxation dispersion experiments11, which yielded both the exchange rates and differences in chemical shifts between the exchanging states. The chemical-shift differences matched those of the lid-opening process, and the rate of opening matched the kcat of the enzyme, thus clearly showing that the turnover rate is governed by the conformational change of the lids and product release. In this issue, Kern and co-workers7 report pre-steady-state kinetic data for Mg2+-free and Mg2+-bound Adk. In the absence of cofactor, the rates of both phosphoryl transfer and product release are slowed dramatically, by factors of about 105 and 103 respectively, compared to when Mg2+ is present. Intriguingly, these perturbations are orthogonal in the sense that replacing Mg2+ with Ca2+ or Co2+ also substantially slows phosphoryl transfer with product release being unaffected. This prompts 102
the question of how the cofactor simultaneously accelerates such different steps in the catalytic pathway. In order to explain the deceleration of product release, the authors again used relaxation dispersion NMR spectroscopy. NMR signals for residues in the lid region were broadened in the presence of Mg2+ and Ca2+, with exchange rates that match kcat. In contrast, no broadening was observed for the protein without cofactor (although a slight broadening was observed at elevated temperatures). This implies that lid motions are essentially quenched in the absence of cofactors, and the lid remains closed over the products, impeding their release. Electrostatic calculations showed that the presence of Mg 2+ weakens electrostatic interactions between the nucleotide products and basic amino acid residues in the lid, results consistent with its mediating acceleration of the opening rate and product release.
To explain the effects on phosphoryl transfer, the authors determined, and subsequently used, X-ray crystal structures of Adk bound to two molecules of ADP, with or without Mg2+, as starting points for molecular dynamics simulations. These showed that the active site and nucleotide phosphates are far more mobile on the nanosecond-to-picosecond timescale in the absence of Mg2+ than in its presence (Fig. 2c,d). The role of protein dynamics in the chemical step of enzyme catalysis is controversial12, and one theory has suggested that motions conduct proteins into and out of catalytically highly active conformational substates13. This model was invoked to explain recent observations in which a mutant of dihydrofolate reductase with suppressed active site dynamics exhibited decreased activity, presumably because motions in the wild-type enzyme provide transient access to catalytically competent states14. In the case of Adk, interestingly, the opposite argument applies. Increased active site dynamics correlates with decreased activity, probably because the increased conformational sampling perturbs the optimum geometry for phosphoryl transfer. By using a combination of biophysical techniques, it is now possible to map the low-lying excited conformational states of enzymes and to relate them to the catalytic mechanisms. The work from Kern and co-workers7 goes beyond mapping exchange among local minima in
a
b
c
d
Figure 2 Dynamics in the adenylate kinase (Adk) active site. (a,b) Lid domains close over the substrates to protect ATP from hydrolysis by water and open to release reaction products. Regions of the protein that are flexible (red and yellow) and rigid (blue) on the millisecond timescale according to NMR are color coded on the structures of Adk bound to the inhibitor Ap5A15 (green, a) and in the apo state16 (b). Image adapted from ref. 3, Nature Publishing Group. (c,d) The magnesium cofactor prearranges ADP phosphates for efficient phosphoryl transfer. Shown are overlays of representative structures from MD simulations performed with Mg2+ present (yellow, c) and absent (d).
volume 22 number 2 FEbruary 2015 nature structural & molecular biology
news and views the energy landscape to address the question of what controls the heights of the intervening barriers. Their finding that Mg2+ accelerates both chemical and conformational steps of the catalytic cycle is unexpected and highlights the delicate interplay of organization and flexibility within enzyme active sites that makes catalysis possible. COMPETING FINANCIAL INTERESTS The author declares no competing financial interests.
1. Fersht, A.R. Structure and Mechanism in Protein Science: a Guide to Enzyme Catalysis and Protein Folding (W.H. Freeman, 1999). 2. Boehr, D.D., McElheny, D., Dyson, H.J. & Wright, P.E. Science 313, 1638–1642 (2006). 3. Wolf-Watz, M. et al. Nat. Struct. Mol. Biol. 11, 945–949 (2004). 4. Eisenmesser, E.Z. et al. Nature 438, 117–121 (2005). 5. Sprangers, R., Gribun, A., Hwang, P.M., Houry, W.A. & Kay, L.E. Proc. Natl. Acad. Sci. USA 102, 16678–16683 (2005). 6. Watt, E.D., Shimada, H., Kovrigin, E.L. & Loria, J.P. Proc. Natl. Acad. Sci. USA 104, 11981–11986 (2007). 7. Kerns, S.J. et al. Nat. Struct. Mol. Biol. 22, 124–131 (2015). 8. Bender, M.L. & Kezdy, F.J. Annu. Rev. Biochem. 34, 49–76 (1965).
9. Grant, B.D. & Adams, J.A. Biochemistry 35, 2022–2029 (1996). 10. Johnson, D.A., Akamine, P. & Radzio-Andzelm, E. Chem. Rev. 101, 2243–2270 (2001). 11. Palmer, A.G., Kroenke, C.D. & Loria, J.P. Methods Enzymol. 339, 204–238 (2001). 12. Kamerlin, S.C. & Warshel, A. Proteins 78, 1339–1375 (2010). 13. Hammes-Schiffer, S. & Benkovic, S.J. Annu. Rev. Biochem. 75, 519–541 (2006). 14. Bhabha, G. et al. Science 332, 234–238 (2011). 15. Müller, C.W. & Schulz, G.E. J. Mol. Biol. 224, 159–177 (1992). 16. Müller, C.W., Schlauderer, G.J., Reinstein, J. & Schulz, G.E. Structure 4, 147–156 (1996).
Designs on a curve
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© 2015 Nature America, Inc. All rights reserved.
J Fernando Bazan & Andrey V Kajava The structural rules governing the curving folds of solenoid proteins, as distilled down to the level of the underlying sequence repeats, provide designers with the tools to reliably fashion new variants with tunable geometries. Bespoke leucine-rich repeat (LRR) scaffolds, as recognition proteins, can now be tailored to better fit their targets. The frequent glimpses of symmetry in protein structures and their assemblies hint at the evolutionary processes that underlie protein modular architectures and optimize the energetics of folding and association1. Stretching far beyond sequence relationships, symmetries are accentuated by a repertoire of shared domain folds, and deeper links can be divined between smaller structured modules that may represent the primitive building blocks of globular proteins2. Nowhere is this witches’ cauldron of molecular evolution more clearly evident than in repetitive proteins, which are composed of structural motifs that are stable only as part of periodic arrays, thus creating runs of striking fold symmetry with only faint sequence watermarks3. A common way to describe repetitive protein structures is by the solenoidal winding of their polypeptide chains, along with their α-helical, β-structural or mixed fold composition4 (Box 1). Elongated, curved and/or twisted α-helical solenoids include armadillo (ARM), tetratricopeptide (TPR), HEAT and ankyrin (ANK) repeats. β-solenoids can form straight and frequently twisted structures, whereas the mixed α/β repeats of LRR solenoids form J. Fernando Bazan is at Bio-Techne Corporation, Minneapolis, Minnesota, USA, and the Department of Pharmacology, University of Minnesota School of Medicine, Minneapolis, Minnesota, USA. Andrey V. Kajava is at the Centre de Recherche de Biochimie Macromoléculaire, CNRS, University of Montpellier, Montpellier, France. e-mail: [email protected] or [email protected]
sinuous arcs of varying complexity. In addition, circular arrays of β-strand blade repeats fold as β-propellers3. Given the key roles of solenoid proteins in cells as structural scaffolds and binding platforms, their curved shapes have proven to be a vibrant testing ground for protein engineers and designers, who seek to harness them for new uses2. The trend in the field has been to focus on the sparse sequence signatures drawn from clusters of solenoid families as the critical design elements of the respective repeat folds, because they map to
the conserved cores of reference structures. However, this approach does not explore the full structural or functional impact of the tremendous repeat-sequence diversity, nor does it inform the design process. Now the André5 and Baker6 laboratories depart from this search for the uniform features of solenoid folds and chart an intriguing new course by pursuing the question of what makes repeats different, instead of what keeps them the same. The ragged multiplicity of divergent, or possibly convergent, repeat motifs within
Box 1 Basics of solenoid geometry The basic parameters of solenoid structures can be drawn on this simple fold schematic: curvature (purple) and twist (green). Curvature is described by the radius (R) of the superhelical axis: the smaller the radius, the larger the curvature. Twist is determined by following analogous reference points in consecutive structural units (green dots); when connected, these points form a virtual helix (green dashed line). The magnitude of the twist is determined by the size of a dihedral angle between the vectors (red arrows) connecting the superhelical axis with the reference points in two consecutive structural units: the larger the angle, the larger the twist. Twist handedness is that of this helix as it winds around the superhelical axis. The solenoid in the figure has a right-handed twist. Adapted from ref. 4, with permission from Elsevier.
nature structural & molecular biology volume 22 number 2 FEbruary 2015
R
103
© 2015 Nature America, Inc. All rights reserved.
news and views an interesting question. First, uridine is characteristic for RNA and is most divergent from the four DNA-building molecules. Second, it is statistically enriched in RNA loop sections, owing to its ability to form only two hydrogen bonds with its complementing adenine base. Loop sections are considered the first targets of endoribonucleases to degrade RNA elements. Therefore, the probability that U-rich degradation products are made is high. Finally, it is also conceivable that the presence of higher concentrations of Us could serve as a pathogenic pattern during infections because various microbe-associated RNAs are particularly U rich. For example, most positive-strand RNA viruses use negative-strand RNAs with 5′-poly(U) tracts as replication templates. Therefore, abundant amounts of Us are generated during de novo synthesis of polyadenylated progeny RNA in the course of viral replication15. Furthermore, bacterial small noncoding RNAs carry poly(U) tails and have important regulatory functions within bacteria; therefore, they could represent another potential substrate for the generation of single Us during infections16. Given the importance of single-nucleoside recognition for TLR8 activation, it is interesting
that TLRs 7 and 8 were first deorphanized with the discovery of various small-molecule agonists that triggered their activation17. The available crystal structures of TLR8 (refs. 4,5) now provide compelling mechanistic evidence for how small molecules such as the agonistic imidazoquinolines and uridine in synergy with short oligonucleotides can promote protein-protein interaction and reorganization of the dimer into the active conformation. Currently, imidazoquinoline compounds are approved for the topical treatment of actinic keratosis, basal cell carcinoma and certain virus infections. The demonstration that the natural ligand U interacts at the same site as the small-molecule activators provides a strong rationale as well as a valuable template for the development of small-molecule antagonists. Such compounds could target TLR8 and by extension TLR7, which also responds to U-rich RNA and imidazoquinoline agonists. Pharmacologic approaches to inhibit TLRs 7 or 8 could be promising for the treatment of certain autoimmune diseases in which the recognition of host nucleic acids contributes to autoimmunity and devastating inflammation18.
ACKNOWLEDGMENTS The authors are supported by the Deutsche Forschungsgemeinschaft–funded Excellence Cluster ImmunoSensation. COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests. 1. Matzinger, P. Science 296, 301–305 (2002). 2. Akira, S. & Takeda, K. Nat. Rev. Immunol. 4, 499–511 (2004). 3. Heil, F. et al. Science 303, 1526–1529 (2004). 4. Tanji, H. et al. Nat. Struct. Mol. Biol. 22, 109–115 (2015). 5. Tanji, H., Ohto, U., Shibata, T., Miyake, K. & Shimizu, T. Science 339, 1426–1429 (2013). 6. Ohto, U., Tanji, H. & Shimizu, T. Microbes Infect. 16, 273–282 (2014). 7. Gorden, K.K.B. et al. J. Immunol. 177, 8164–8170 (2006). 8. Jurk, M. et al. Eur. J. Immunol. 36, 1815–1826 (2006). 9. Ligoxygakis, P., Pelte, N., Hoffmann, J.A. & Reichhart, J.M. Science 297, 114–116 (2002). 10. Tesar, B.M. et al. Am. J. Transplant. 6, 2622–2635 (2006). 11. Chan, M.P. et al. Nat. Commun. (in the press). 12. Pawaria, S. et al. J. Immunol. (in the press). 13. Horiuchi, T., Horiuchi, S. & Mizuno, D. Nature 183, 1529–1530 (1959). 14. Hausser, E. & Stroun, J. Rev. Med. Suisse Romande 83, 577–586 (1963). 15. White, K.A., Enjuanes, L. & Berkhout, B. RNA Biol. 8, 182–183 (2011). 16. Vogel, J. & Wagner, E.G. Curr. Opin. Microbiol. 10, 262–270 (2007). 17. Hemmi, H. et al. Nat. Immunol. 3, 196–200 (2002). 18. Christensen, S.R. & Shlomchik, M.J. Semin. Immunol. 19, 11–23 (2007).
An enzyme cofactor with a split personality Anthony Mittermaier Little is currently known about the molecular determinants of energy barriers along enzyme catalytic pathways. Kern and co-workers have studied this question in adenylate kinase (Adk) and now reveal that a single Mg 2+ ion can accelerate two distinct steps, thus uncovering an unexpected dual role for this ubiquitous cofactor. Many decades of research have been devoted to understanding how enzymes lower the energy barriers for chemical reactions and accelerate rates by many orders of magnitude. Enzyme catalysis often proceeds through multiple discreet steps, each associated with its own energy barrier1 (Fig. 1), and it has become possible to relate the individual steps of catalytic pathways to specific conformational changes occurring in the enzymes themselves by using NMR techniques2–6. Nevertheless, many questions remain to be answered, including those regarding the factors controlling the heights of the barriers. As reported in this issue, Kern and co-workers7 have studied the role of a magnesium (Mg2+) Anthony Mittermaier is at the Department of Chemistry, McGill University, Montreal, Quebec, Canada. e-mail: [email protected]
cofactor in the Adk cycle, which catalyzes the reversible phosphorylation of AMP by ATP. Using a broad combination of biophysical techniques including NMR, they showed that Mg2+ has a surprising dual role: it dramatically accelerates the on-enzyme rate of phosphoryl transfer and also promotes a lid-opening conformational change that is essential for product release. Interestingly, the two effects depend on different aspects of the cofactor’s ‘personality’. The authors found that Mg2+ enhances lid opening primarily by shielding electrostatic interactions, whereas it enhances the chemical step by coordinating the nucleotide phosphate groups and by acting as a pivot during phosphoryl transfer. Some of the first knowledge of the individual steps in enzyme catalytic pathways came from pre-steady-state kinetic assays 8. In this approach, reactions are performed with relatively high concentrations of enzymes,
nature structural & molecular biology volume 22 number 2 FEbruary 2015
such that the buildup of intermediates and/ or products within the active site can be measured before their release from the enzyme. For example, when protein kinase A (PKA) is added to its ATP and peptide substrates, one enzyme equivalent of the phosphopeptide product is formed very rapidly, at a rate far greater than that of steady-state turnover9. This implies that phosphoryl transfer is fast, whereas product release is rate limiting. X-ray crystal structures provided a possible explanation for the slow rate of product release. The enzyme active site adopts both ‘open’ conformations, which are solvent accessible, and ‘closed’ conformations, in which the ATP γ-phosphate is positioned for transfer10. This suggested that product release is predicated on active site opening. However, it is not possible to ascertain whether the conformational transition itself is rate limiting solely on the basis of enzyme kinetic experiments. 101
news and views Figure 1 Schematic representation of enzymatic catalysis. (a) Enzymes (E) increase the rate of conversion of substrates (S) to products (P) by lowering the energy of the transition state (TS) separating the two forms. (b) In many cases, catalytic mechanisms involve a series of discrete steps, each associated with its own energy barrier, for instance, enzyme conformational rearrangements (TS1 and TS3) and chemical transformation (TS2).
a TS
S
P TS E
E
Energy
b S
P E
TS1
S
P
E
E
TS2
TS3
E
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© 2015 Nature America, Inc. All rights reserved.
Energy
Similarly to that of PKA, the Adk active site adopts both open and closed configurations, characterized by ‘lid’ domains that fold over the active site when ligands are bound. The central role of lid dynamics in Adk catalysis was elegantly demonstrated in earlier work by the Kern laboratory3. Because the reaction catalyzed by the enzyme (ATP + AMP ↔ 2 ADP) is reversible, it was possible to perform high-resolution protein NMR experiments on samples actively performing catalysis on substrates and products exchanging in a dynamic equilibrium. The authors found that NMR signals from protein nuclei in the lid regions of Adk are broadened in the presence of saturating concentrations of ligands (Fig. 2a,b), a signature of exchange between different conformations occurring on millisecond-to-microsecond timescales. They characterized the broadening by using recently developed NMR relaxation dispersion experiments11, which yielded both the exchange rates and differences in chemical shifts between the exchanging states. The chemical-shift differences matched those of the lid-opening process, and the rate of opening matched the kcat of the enzyme, thus clearly showing that the turnover rate is governed by the conformational change of the lids and product release. In this issue, Kern and co-workers7 report pre-steady-state kinetic data for Mg2+-free and Mg2+-bound Adk. In the absence of cofactor, the rates of both phosphoryl transfer and product release are slowed dramatically, by factors of about 105 and 103 respectively, compared to when Mg2+ is present. Intriguingly, these perturbations are orthogonal in the sense that replacing Mg2+ with Ca2+ or Co2+ also substantially slows phosphoryl transfer with product release being unaffected. This prompts 102
the question of how the cofactor simultaneously accelerates such different steps in the catalytic pathway. In order to explain the deceleration of product release, the authors again used relaxation dispersion NMR spectroscopy. NMR signals for residues in the lid region were broadened in the presence of Mg2+ and Ca2+, with exchange rates that match kcat. In contrast, no broadening was observed for the protein without cofactor (although a slight broadening was observed at elevated temperatures). This implies that lid motions are essentially quenched in the absence of cofactors, and the lid remains closed over the products, impeding their release. Electrostatic calculations showed that the presence of Mg 2+ weakens electrostatic interactions between the nucleotide products and basic amino acid residues in the lid, results consistent with its mediating acceleration of the opening rate and product release.
To explain the effects on phosphoryl transfer, the authors determined, and subsequently used, X-ray crystal structures of Adk bound to two molecules of ADP, with or without Mg2+, as starting points for molecular dynamics simulations. These showed that the active site and nucleotide phosphates are far more mobile on the nanosecond-to-picosecond timescale in the absence of Mg2+ than in its presence (Fig. 2c,d). The role of protein dynamics in the chemical step of enzyme catalysis is controversial12, and one theory has suggested that motions conduct proteins into and out of catalytically highly active conformational substates13. This model was invoked to explain recent observations in which a mutant of dihydrofolate reductase with suppressed active site dynamics exhibited decreased activity, presumably because motions in the wild-type enzyme provide transient access to catalytically competent states14. In the case of Adk, interestingly, the opposite argument applies. Increased active site dynamics correlates with decreased activity, probably because the increased conformational sampling perturbs the optimum geometry for phosphoryl transfer. By using a combination of biophysical techniques, it is now possible to map the low-lying excited conformational states of enzymes and to relate them to the catalytic mechanisms. The work from Kern and co-workers7 goes beyond mapping exchange among local minima in
a
b
c
d
Figure 2 Dynamics in the adenylate kinase (Adk) active site. (a,b) Lid domains close over the substrates to protect ATP from hydrolysis by water and open to release reaction products. Regions of the protein that are flexible (red and yellow) and rigid (blue) on the millisecond timescale according to NMR are color coded on the structures of Adk bound to the inhibitor Ap5A15 (green, a) and in the apo state16 (b). Image adapted from ref. 3, Nature Publishing Group. (c,d) The magnesium cofactor prearranges ADP phosphates for efficient phosphoryl transfer. Shown are overlays of representative structures from MD simulations performed with Mg2+ present (yellow, c) and absent (d).
volume 22 number 2 FEbruary 2015 nature structural & molecular biology
news and views the energy landscape to address the question of what controls the heights of the intervening barriers. Their finding that Mg2+ accelerates both chemical and conformational steps of the catalytic cycle is unexpected and highlights the delicate interplay of organization and flexibility within enzyme active sites that makes catalysis possible. COMPETING FINANCIAL INTERESTS The author declares no competing financial interests.
1. Fersht, A.R. Structure and Mechanism in Protein Science: a Guide to Enzyme Catalysis and Protein Folding (W.H. Freeman, 1999). 2. Boehr, D.D., McElheny, D., Dyson, H.J. & Wright, P.E. Science 313, 1638–1642 (2006). 3. Wolf-Watz, M. et al. Nat. Struct. Mol. Biol. 11, 945–949 (2004). 4. Eisenmesser, E.Z. et al. Nature 438, 117–121 (2005). 5. Sprangers, R., Gribun, A., Hwang, P.M., Houry, W.A. & Kay, L.E. Proc. Natl. Acad. Sci. USA 102, 16678–16683 (2005). 6. Watt, E.D., Shimada, H., Kovrigin, E.L. & Loria, J.P. Proc. Natl. Acad. Sci. USA 104, 11981–11986 (2007). 7. Kerns, S.J. et al. Nat. Struct. Mol. Biol. 22, 124–131 (2015). 8. Bender, M.L. & Kezdy, F.J. Annu. Rev. Biochem. 34, 49–76 (1965).
9. Grant, B.D. & Adams, J.A. Biochemistry 35, 2022–2029 (1996). 10. Johnson, D.A., Akamine, P. & Radzio-Andzelm, E. Chem. Rev. 101, 2243–2270 (2001). 11. Palmer, A.G., Kroenke, C.D. & Loria, J.P. Methods Enzymol. 339, 204–238 (2001). 12. Kamerlin, S.C. & Warshel, A. Proteins 78, 1339–1375 (2010). 13. Hammes-Schiffer, S. & Benkovic, S.J. Annu. Rev. Biochem. 75, 519–541 (2006). 14. Bhabha, G. et al. Science 332, 234–238 (2011). 15. Müller, C.W. & Schulz, G.E. J. Mol. Biol. 224, 159–177 (1992). 16. Müller, C.W., Schlauderer, G.J., Reinstein, J. & Schulz, G.E. Structure 4, 147–156 (1996).
Designs on a curve
npg
© 2015 Nature America, Inc. All rights reserved.
J Fernando Bazan & Andrey V Kajava The structural rules governing the curving folds of solenoid proteins, as distilled down to the level of the underlying sequence repeats, provide designers with the tools to reliably fashion new variants with tunable geometries. Bespoke leucine-rich repeat (LRR) scaffolds, as recognition proteins, can now be tailored to better fit their targets. The frequent glimpses of symmetry in protein structures and their assemblies hint at the evolutionary processes that underlie protein modular architectures and optimize the energetics of folding and association1. Stretching far beyond sequence relationships, symmetries are accentuated by a repertoire of shared domain folds, and deeper links can be divined between smaller structured modules that may represent the primitive building blocks of globular proteins2. Nowhere is this witches’ cauldron of molecular evolution more clearly evident than in repetitive proteins, which are composed of structural motifs that are stable only as part of periodic arrays, thus creating runs of striking fold symmetry with only faint sequence watermarks3. A common way to describe repetitive protein structures is by the solenoidal winding of their polypeptide chains, along with their α-helical, β-structural or mixed fold composition4 (Box 1). Elongated, curved and/or twisted α-helical solenoids include armadillo (ARM), tetratricopeptide (TPR), HEAT and ankyrin (ANK) repeats. β-solenoids can form straight and frequently twisted structures, whereas the mixed α/β repeats of LRR solenoids form J. Fernando Bazan is at Bio-Techne Corporation, Minneapolis, Minnesota, USA, and the Department of Pharmacology, University of Minnesota School of Medicine, Minneapolis, Minnesota, USA. Andrey V. Kajava is at the Centre de Recherche de Biochimie Macromoléculaire, CNRS, University of Montpellier, Montpellier, France. e-mail: [email protected] or [email protected]
sinuous arcs of varying complexity. In addition, circular arrays of β-strand blade repeats fold as β-propellers3. Given the key roles of solenoid proteins in cells as structural scaffolds and binding platforms, their curved shapes have proven to be a vibrant testing ground for protein engineers and designers, who seek to harness them for new uses2. The trend in the field has been to focus on the sparse sequence signatures drawn from clusters of solenoid families as the critical design elements of the respective repeat folds, because they map to
the conserved cores of reference structures. However, this approach does not explore the full structural or functional impact of the tremendous repeat-sequence diversity, nor does it inform the design process. Now the André5 and Baker6 laboratories depart from this search for the uniform features of solenoid folds and chart an intriguing new course by pursuing the question of what makes repeats different, instead of what keeps them the same. The ragged multiplicity of divergent, or possibly convergent, repeat motifs within
Box 1 Basics of solenoid geometry The basic parameters of solenoid structures can be drawn on this simple fold schematic: curvature (purple) and twist (green). Curvature is described by the radius (R) of the superhelical axis: the smaller the radius, the larger the curvature. Twist is determined by following analogous reference points in consecutive structural units (green dots); when connected, these points form a virtual helix (green dashed line). The magnitude of the twist is determined by the size of a dihedral angle between the vectors (red arrows) connecting the superhelical axis with the reference points in two consecutive structural units: the larger the angle, the larger the twist. Twist handedness is that of this helix as it winds around the superhelical axis. The solenoid in the figure has a right-handed twist. Adapted from ref. 4, with permission from Elsevier.
nature structural & molecular biology volume 22 number 2 FEbruary 2015
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