Tai Lieu Chat Luong Understanding Enzymes This page intentionally left blank Understanding Enzymes Function, Design, Engineering, and Analysis edited by Allan Svendsen CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2016 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S Government works Version Date: 20160419 International Standard Book Number-13: 978-981-4669-33-7 (eBook - PDF) This book contains information obtained from authentic and highly regarded sources Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint Except as permitted under U.S Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers For permission to photocopy or use material electronically from this work, please access www copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400 CCC is a not-for-profit organization that provides licenses and registration for a variety of users For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com March 21, 2016 12:20 PSP Book - 9in x 6in 00-Allan-Svendsen-Prelims Contents xix Introduction PART I ENZYME FUNCTION A Short Practical Guide to the Quantitative Analysis of Engineered Enzymes Christopher D Bayer and Florian Hollfelder 1.1 Introduction 1.2 Quantifying Reaction Progress 1.3 Typical Saturation Plots Give Michaelis–Menten Parameters 1.4 What Can Go Wrong? 1.5 Dealing with Multiphasic and Pre-Steady-State Kinetics 1.6 Evaluating Enzymes Protein Conformational Motions: Enzyme Catalysis Xinyi Huang, C Tony Liu, and Stephen J Benkovic 2.1 Introduction 2.2 Multidimensional Protein Landscape and the Timescales of Motions 2.3 Conformational Changes in Enzyme–Substrate Interactions 2.4 Conformational Changes in Catalysis 2.4.1 Protein Dynamics of DHFR in the Catalytic Cycle 2.4.2 Temporally Overlap: Correlation Does Not Mean Causation 2.4.3 Fast Timescale Conformational Fluctuations 3 12 16 21 21 22 26 28 30 32 34 March 28, 2016 10:38 PSP Book - 9in x 6in 00-Allan-Svendsen-Prelims vi Contents 2.4.4 Effect of Conformational Changes on the Electrostatic Environment 2.5 Conservation of Protein Motions in Evolution 2.6 Designing Protein Dynamics 2.7 Concluding Remarks Enzymology Meets Nanotechnology: Single-Molecule Methods for Observing Enzyme Kinetics in Real Time Kerstin G Blank, Anna A Wasiel, and Alan E Rowan 3.1 Introduction 3.2 Single-Turnover Detection 3.2.1 Fluorescent Reporter Systems 3.2.2 Measurement Setup 3.2.3 Data Analysis 3.3 Single-Enzyme Kinetics 3.3.1 Candida antarctica Lipase B 3.3.2 Thermomyces lanuginosus Lipase 3.3.3 α-Chymotrypsin 3.3.4 Nitrite Reductase 3.3.5 Summary 3.4 New Developments Facilitated by Nanotechnology 3.4.1 Nano-optical Approaches 3.4.2 Nano-electronic Approaches 3.4.3 Nanomechanical Approaches 3.4.4 Summary 3.5 Conclusion Interfacial Enzyme Function Visualized Using Neutron, X-Ray, and Light-Scattering Methods Hanna Wacklin and Tommy Nylander 4.1 Phospholipase A2 : An Interfacially Activated Enzyme 4.1.1 Neutron Reflection 4.1.2 Ellipsometry 4.1.3 Activity of Naja mossambica mossambica PLA2 4.1.4 Fate of the Reaction Products 4.1.5 The Lag Phase and Activation of Pancreatic PLA2 4.1.6 Distribution of Products during the Lag Phase 36 38 39 40 47 48 53 53 56 57 60 63 67 73 78 84 88 89 96 103 108 110 125 126 129 130 130 133 135 138 March 21, 2016 12:20 PSP Book - 9in x 6in 00-Allan-Svendsen-Prelims Contents 4.1.7 Hydrolysis of DPPC by Pancreatic PLA2 4.1.8 Role of the Reaction Products in PLA2 Activation 4.1.9 Effect of pH and Activation by Me-β-cyclodextrin 4.2 Other Lipolytic Enzyme Reactions on Surfaces 4.2.1 Triacylglycerol Lipases and the Role of Lipid Liquid Crystalline Nanostructures 4.3 Cellulase Enzymes 4.4 Conclusion Folding Dynamics and Structural Basis of the Enzyme Mechanism of Ubiquitin C-Terminal Hydroylases Shang-Te Danny Hsu 5.1 Introduction 5.1.1 UCH-L1 5.1.1.1 Genetic association between UCH-L1 and neurodegenerative diseases 5.1.1.2 UCH-L1 in oncogenesis 5.1.2 Molecular Insights into the Pathogenesis Associated with UCH-L1 5.1.3 UCHL3 5.1.4 UCHL5 5.1.5 BAP1 5.2 UCH Structures 5.3 Folding Dynamics and Kinetics 5.4 Substrate Recognition 5.5 Enzyme Mechanism 5.6 Conclusion 139 141 144 150 150 154 158 167 169 171 171 175 175 177 178 179 180 183 184 186 189 Stabilization of Enzymes by Metal Binding: Structures of Two Alkalophilic Bacillus Subtilases and Analysis of the Second Metal-Binding Site of the Subtilase Family 203 Jan Dohnalek, Katherine E McAuley, Andrzej M Brzozowski, Peter R Østergaard, Allan Svendsen, and Keith S Wilson 6.1 Introduction: Subtilases and Metal Binding 203 6.1.1 Calcium-Binding Sites in Bacillus: Proposal for a Standard Nomenclature 209 vii March 21, 2016 12:20 PSP Book - 9in x 6in 00-Allan-Svendsen-Prelims viii Contents 6.1.2 The Weak Metal-Binding Site 6.2 Two New Structures of Subtilases with Altered Calcium Sites 6.2.1 Proteinase SubTY 6.2.1.1 The overall fold 6.2.1.2 The active site 6.2.1.3 SubTY calcium and sodium sites 6.2.1.4 SubTY disulfide bridge 6.2.2 SubHal 6.2.2.1 The unliganded form of SubHal 6.2.2.2 The SubHal:CI2A complex 6.2.2.3 Termini, surface, and pH stability of SubHal 6.2.2.4 The two crystallographically independent SubHal:CI2A complexes 6.2.2.5 The calcium sites in SubHal 6.2.2.6 The active site of SubHal 6.2.3 Enzymatic Activity of SubTY and SubHal 6.2.4 Comparison of SubTY and SubHal with Other Subtilases 6.2.5 The SubHal C-domain Compared to the Eukaryotic PCs, Furin and Kexin 6.2.5.1 Active site comparison 6.2.5.2 The specificity pockets 6.2.5.3 Inhibitor CI2A binding 6.2.6 Activity Profiles 6.2.7 Comparison of Metal Binding at the Strong and Weak Sites in the S8 Family 6.2.8 The Ca-II and Na-II Metal-Binding Sites 6.3 Conclusion: Implications for Structural Studies of Enzymes 6.4 Materials and Methods 6.4.1 SubTY 6.4.1.1 Protein production and purification 6.4.1.2 Purification of the SubTY:CI2A (1:1) complex 6.4.1.3 Crystallization 6.4.1.4 Structure determination 214 216 216 216 216 218 219 220 220 221 221 223 224 226 228 228 232 233 234 234 236 236 237 248 249 249 249 250 250 251 March 21, 2016 12:20 PSP Book - 9in x 6in 00-Allan-Svendsen-Prelims Contents 6.4.2 SubHal 6.4.2.1 Protein production and purification 6.4.2.2 Purification of the SubHal:CI2A (1:1) complex 6.4.2.3 Crystallization 6.4.2.4 Structure determination 6.4.3 Protease Assays 6.4.4 pH Stability 6.4.5 Data Deposition Structure and Functional Roles of Surface Binding Sites in Amylolytic Enzymes Darrell Cockburn and Birte Svensson 7.1 Introduction 7.2 Identification of SBSs: X-Ray Crystallography 7.3 Bioinformatics of SBS Enzymes 7.4 Binding Site Isolation 7.5 Protection of Binding Sites from Chemical Labeling 7.6 Nuclear Magnetic Resonance 7.7 Binding Assays 7.8 Activity Assays 7.9 Future Prospects 7.10 Conclusion Interfacial Enzymes and Their Interactions with Surfaces: Molecular Simulation Studies Nathalie Willems, Mickaăel Lelimousin, Heidi Koldsứ, and Mark S P Sansom 8.1 Introduction 8.2 Enzyme Interactions at Interfaces 8.3 Molecular Dynamic Simulations of Biomolecular Systems 8.4 Lipases 8.4.1 Atomistic MD Studies of Lipase Interactions with Interfaces 8.4.2 The Role of Water in Lipase Catalysis at Interfaces 251 251 252 252 253 256 257 257 267 267 271 273 275 277 277 278 282 283 286 297 297 299 301 303 304 307 ix March 23, 2016 13:17 PSP Book - 9in x 6in 25-Allan-Svendsen-c25 Kinetic Stability Linked to Substantially Unfolded Transition States 845 environment surrounding the protein in vivo) in which irreversible alterations of the unfolded state may readily take place, the rate of irreversible denaturation may be determined by the free-energy barrier for unfolding [12], a scenario that can be experimentally verified in vitro by using proteases to simulate a harsh environment [44], and (ii) the half-life time associated to the crossing of such a barrier (1/k in Eq 25.1) changes in an exponential manner with the barrier height and even a decrease of a few kilojoules per mole in height may cause a large decrease in half-life that can affect organism fitness and be subject to purifying natural selection (Fig 25.3) This interpretation of the stability–/sequence–statistics correlation in terms of kinetic stability was actually supported by exhaustive unfolding kinetics studies on all the 27 variants that demonstrated a good correlation between the mutation effects on the free-energy barrier and the frequency of occurrence of residues in sequence alignments [43] (Fig 25.3) The overall picture that emerges from these studies on thioredoxin variants certainly not only involves natural selection for kinetic stability but also implies a substantially unfolded transition state This picture does provide a molecular-level explanation for kinetic stability in this system, as many stabilizing interactions in the native state are broken up in the transition state, thus contributing to a high free-energy barrier Also, a substantially unfolded transition state is consistent with the correlation between mutation effects on thermodynamic and kinetic stabilities found in this system (Fig 25.3), since many mutations are expected to occur in regions of the structure that become unfolded in the transition state Finally, a substantially unfolded transition state greatly facilitates the rational design of kinetic stability given that modeled effects on thermodynamic stability (unfolding free-energy difference between the native and unfolded states) will be transferred to the free-energy barrier that determines the rate of denaturation As an example, charge reversal mutations were introduced in consensus-stabilized thioredoxin scaffolds to yield variants with both thermodynamic and kinetic stabilities highly tunable by salt [45] Recent work has addressed the stabilization of phytase from Citrobacter braakii (a biotechnologically important enzyme) on the basis of disulfide bridge engineering [46] High-temperature March 23, 2016 13:17 PSP Book - 9in x 6in 846 Kinetic Stability of Variant Enzymes Figure 25.3 Comparing mutational effects on thioredoxin thermodynamic and kinetic stabilities (Left) Dependence of the equilibrium unfolding temperature and the unfolding rate constant with changes in the corresponding free-energy changes (unfolding: G; activation barrier: GN→‡ ) The right axis on the lower plot indicates the rate constant levels corresponding to representative half-lives Note that a free-energy change (kJ/mol) alters the equilibrium unfolding temperature by just a few degrees, while it can have a substantial effect on the half-life time for irreversible denaturation (Right) Correlation between mutational effects on the unfolding free-energy and the activation free-energy barrier (from the analysis of equilibrium and kinetic data for urea-induced unfolding) for the 27 mutations of Fig 25.1 The excellent correlation with slope close to unity supports that the mutation sites are in regions that are substantially unfolded in the transition state Reprinted from Ref [43], Copyright (2006), with permission from Elsevier molecular dynamics simulations were used to determine regions showing large structural displacements at 500 K when compared with the starting structure Design of the disulfide bridges was focused on these regions on the basis of the following assumptions: (i) the targeted regions could plausibly be unfolded or disrupted in the transition state for irreversible denaturation, and therefore, disulfide bridges introduced in those regions may enhance the corresponding free-energy barrier, and (ii) the targeted regions could plausibly be flexible in the native structure, and therefore, introduction of disulfide bridges in those regions is 25-Allan-Svendsen-c25 March 23, 2016 13:17 PSP Book - 9in x 6in 25-Allan-Svendsen-c25 Kinetic Stability Linked to Substantially Unfolded Transition States 847 Figure 25.4 Differential scanning calorimetry of the thermal denaturation of variants of the phytase from Citrobacter braakii (a) Calorimetric profiles for the WT enzyme and variants with one (S1, S2, S3), two (D1, D2), and three (T) engineered disulfide bridges These profiles are partially reversible and show some scan rate dependence; therefore, they only provide a first, rough assessment of the mutational effects on thermodynamic stability (b) Plot of denaturation temperature (as determined from the calorimetric profiles) versus the number of engineered disulfide bridges (c) Reversibility of the thermal denaturation is higher for the phytase variants with two and three engineered disulfide bridges Reprinted from Ref [46] (published under a Creative Commons Attribution license) not likely to distort or strain the native structure Variants with one, two, and three engineered disulfide bridges were prepared and characterized exhaustively on the basis of differential scanning calorimetry (Fig 25.4) and inactivation kinetics at several temperatures (Fig 25.5) The variants showed both enhanced thermodynamic stability and enhanced kinetic stability, supporting that the engineered disulfide bridges were located in regions of the structure that become unfolded in the kinetically relevant transition state Possibly, the most striking result of this study was the strongly non-Arrhenius temperature dependence observed for the inactivation rate of all variants studied (Fig 25.5) In fact, for all the variants the timescale of the irreversible denaturation process reached a minimum at an intermediate temperature within the denaturation transition range [46] This striking behavior was shown to be a signature of the key kinetic role in irreversible aggregation played by a highly unfolded intermediate state/ensemble Such intermediate state would of course be a good model for the kinetically relevant transition state of the irreversible denaturation process March 23, 2016 13:17 PSP Book - 9in x 6in 848 Kinetic Stability of Variant Enzymes Figure 25.5 Kinetics of thermal inactivation of a variant of phytase from Citrobacter braakii with two engineered disulfide bridges (a) compared to the WT protein (b) A/A is the fraction of the initial enzyme activity The variant shows a considerable kinetic stabilization with respect to the WT enzyme Note that in both cases, thermal inactivation is slowest at an intermediate temperature (a strongly non-Arrhenius temperature dependence), a result that suggests a key kinetic role of a substantially unfolded intermediate state (likely a good model for the kinetically relevant transition state) Reprinted from Ref [46] (published under a Creative Commons Attribution license) 25.5 Role of Solvation Barriers in Kinetic Stability: Lipases and Triose Phosphate Isomerases Previous sections would seem to convey a simple structural picture of the transition states for irreversible denaturation processes as partially unfolded or partially disrupted native structures In fact, a transition state belongs to the top of a free-energy barrier and may display rather unusual features when compared with the states (native, unfolded) at free-energy basins Indeed, experimental and computational analyses support that transition states for protein folding/unfolding processes may display a clearly nonnative energetic nonbalance associated to solvation/desolvation of amino acid moieties [47–51] For instance, protein unfolding implies the interaction with water (i.e., solvation) of the amino acid residues buried in the native structure However, the molecule of water has a finite size and buried interacting amino acids must separate a certain distance before water molecules can slip in Such water-unsatisfied separation involves a large increase in energy associated to disrupted internal interactions (van der Waals packing interactions between buried hydrophobic residues, for instance) This situation is often described in the literature as a solvation barrier, and it would be described as a desolvation barrier if the 25-Allan-Svendsen-c25 March 23, 2016 13:17 PSP Book - 9in x 6in 25-Allan-Svendsen-c25 Role of Solvation Barriers in Kinetic Stability 849 Figure 25.6 Cartoon depiction of solvation/desolvation barrier effects Blue color is used for the protein surface that is accessible to the solvent Red color represents broken or disrupted internal interactions that are not satisfied by water molecules A folding–unfolding scenario is shown in this figure, but the type of transition state depicted may also play a role in kinetic stability Reprinted from Ref [51], Copyright (2006), with permission from Elsevier process is seen in the folding direction (i.e., from the unfolded state to the transition state) What is meant by these terms is essentially that there may be a large contribution to the freeenergy barrier for folding–unfolding due to the asynchrony between water penetration/release and disruption/setting up of internal interactions (Fig 25.6) While solvation/desolvation barriers were originally proposed to play a role in folding/unfolding processes, their contribution to protein kinetic stability has been established mainly through the studies on lipases and triose phosphate isomerases summarized below Stabilization of lipase from Thermomyces lanuginosus (an important industrial enzyme) has been addressed on the basis of directed evolution focused to flexible regions detected in hightemperature molecular dynamics simulations [52] The variants thus obtained showed substantially enhanced kinetic stability (Fig 25.7), a result consistent with the general proposal that heated molecular dynamics simulations may plausibly locate regions of March 23, 2016 13:17 PSP Book - 9in x 6in 850 Kinetic Stability of Variant Enzymes Figure 25.7 (Left) Mutational effects on the free-energy barrier for the irreversible denaturation of the lipase from Thermomyces lanuginosus (G‡ values) Free-energy changes have enthalpic and entropic components, and here the G‡ values are plotted versus their enthalpic components, that is, versus the mutational changes in activation enthalpy (H ‡ values) (Right) Activation urea m values (m‡ ) for the several lipase variants studied plotted versus the mutational changes in (a) activation enthalpy and (b) the denaturation temperature The m‡ values shown are substantially smaller than the m values for complete unfolding of lipase (about 17 kJ/[mol·M]), indicating low exposure to the solvent in the transition state Furthermore, the m‡ values not correlate with the corresponding mutational changes in activation enthalpy supporting a transition state akin to that depicted in Fig 25.6 Reprinted from Ref [52], Copyright (2006), with permission from Elsevier 25-Allan-Svendsen-c25 March 23, 2016 13:17 PSP Book - 9in x 6in 25-Allan-Svendsen-c25 Role of Solvation Barriers in Kinetic Stability 851 the structure that become disrupted in the kinetically relevant transition state However, detailed analyses of differential scanning calorimetry thermograms for the irreversible denaturation of the lipase variants led to a more intricate picture Activation enthalpy values can be derived from the fittings of a two-state irreversible denaturation model to the calorimetric profiles Furthermore, calorimetric experiments at different urea concentrations allow the estimation of the denaturant-concentration dependence of the activation free energy, that is, the so-called activation urea m value, which is generally taken as a measure of the exposure to the solvent of the transition state as compared with the native state Surprisingly, for all variants the two values showed a clear mismatch, with the activation enthalpy being very large (on the order hundreds of kilojoules per mole) and the activation m values being comparatively small A high activation enthalpy suggests that a substantial number of interactions are broken or disrupted in the transition state, while a low urea m value indicates the transition state has a low exposure to solvent It follows that disrupted (or partially disrupted) interactions in the transition state are mainly internal and still not satisfied by water molecules; we find, therefore, the structural pattern expected to be associated to a solvation barrier contribution The activation enthalpy versus the urea m value disparity observed with lipase also extends to the corresponding mutation effects on these quantities (Fig 25.7), revealing that the solvation barrier contribution is highly sensitive to mutation [52, 53] Furthermore, studies on the kinetic stability triosephosphate isomerases [53, 54] reproduced the activation-enthalpy/urea m pattern associated to a solvation barrier contribution and also showed such contributions to significantly differ among triose phosphate isomerases from different organisms It emerges, overall, that solvation/desolvation effects may provide an efficient mechanism for the modulation of free-energy barriers The rational exploitation of this mechanism for kinetic stability enhancement is not straightforward at this stage, however, since, among other problems, solvation barrier contributions seem to be characterized by pervasive enthalpy/entropy compensation phenomena [53, 54] that often are difficult to rationalize and predict March 23, 2016 13:17 PSP Book - 9in x 6in 852 Kinetic Stability of Variant Enzymes 25.6 Concluding Remarks We have purposely used in this chapter an approach to protein kinetic stability based exclusively on phenomenological descriptions of the transition states for the kinetically relevant conformational processes Admittedly, this approach has obvious limitations For instance, kinetic stability in a highly proteolytic environment may require not only a high free-energy barrier for unfolding/denaturation [44] but also the suppression of the local structure fluctuations that render the native state susceptible to proteolysis [36] It does not appear that the molecular mechanisms responsible for such suppression can be described in terms of a single transition state associated to extensive conformational changes Also, the longterm kinetic stability at low temperature that is relevant for the shelf life of protein pharmaceuticals may be determined in part by chemical alterations of residues in the native structure (such as deamidation or cyclic imide formation) [14], processes, which, again, may not be linked to large conformational alterations From a more general viewpoint, it must be recognized that mechanisms of irreversible protein denaturation may in some cases be considerably complex [46, 55–57] For instance, protein aggregation may involve steps of nucleation, growth, fragmentation, coalescence, etc Clearly, assuming a single rate-determining step and a single kinetically relevant free-energy barrier may not be realistic in some cases The above caveats notwithstanding, it stems from the examples discussed in this chapter that a description on the basis of a single transition state may be adequate as a first approximation in many cases, and furthermore, it may be useful in the following sense: (i) it may provide clues to the molecular mechanisms behind the natural selection of the kinetic stabilization found in many protein systems; (ii) it may offer suitable guidelines for protein engineering as, for instance, directed evolution procedures aimed at enhancing protein kinetic stability are more likely to be successful if focused to the regions of the protein structure that become unfolded or disrupted in the kinetically relevant transition state; and (iii) it may open up the possibility of using known computational procedures for transition-state prediction in the rational design of kinetic stability, and in this context, it is noteworthy that several studies 25-Allan-Svendsen-c25 March 23, 2016 13:17 PSP Book - 9in x 6in 25-Allan-Svendsen-c25 References 853 (discussed in this chapter) support that high-temperature molecular dynamics simulations can be employed to assess the regions of the structure that are likely unfolded/disrupted in the kinetically relevant transition state Acknowledgments Work in the author’s lab is supported by grants BIO2012-34937 and CSD2000-00088 (Spanish Ministry of Economy and Competitiveness) and Feder Funds References Carpenter, J F., and Manning, M C (2002) Rational Design of Stable Protein Formulations: Theory and Practice (Kluwer Academic/Plenum, New York) Kirk, O., Borchert, T V., and Fuglsang, C C (2002) Industrial enzyme applications, Curr Opin Biotechnol., 13, pp 345–351 Lazar, G.A., Marshall, S A., Plecs, J J., Mayo, S L., and Dejarlais, J R (2003) Designing proteins for therapeutic applications, Curr Opin Struct Biol., 13, pp 513–518 Bloom, J D., Labthavikul, S T., Otey, C R., and Arnold, F H (2006) Protein stability promotes evolvability, Proc Natl Acad Sci U S A, 103, pp 5869 5874 ă Khersonsky, O., Kiss, G., Rothlisberger, D., Dym, O., Albeck, S., Houk, K N., Baker, D., and Tawfik, D S (2012) Bridging the gaps in design methodologies by evolutionary optimization of the stability and proficiency of designed Kemp eliminase KE59, Proc Natl Acad Sci U S A, 109, pp 10358–10363 Becktel W J., and Schellman, J A (1987) Protein stability curves, Biopolymers, 26, pp 1859–1877 Privalov, P L (1990) Cold denaturation of proteins, Crit Rev Biochem Mol Biol., 25, pp 281–305 Sanchez-Ruiz, J M (1995) Chapter 6, Differential scanning calorimetry of proteins In Subcellular Biochemistry, Biswas, B B., and Roy, S., eds (Plenum Press, New York), 24, pp 133–176 March 23, 2016 13:17 PSP Book - 9in x 6in 854 Kinetic Stability of Variant Enzymes Klibanov, A M., and Ahern, T J (1987) Thermal stability of proteins In Protein Engineering, Oxender, D L., and Fox, C F., eds (Alan R Liss, New York), pp 213–218 10 Lumry, R., and Eyring, E (1954) Conformational changes of proteins, J Phys Chem., 58, pp 110–120 11 Freire, E., van Osdol, W W., Mayorga, O L., and Sanchez-Ruiz, J M (1990) Calorimetrically-determined dynamics of complex unfolding transitions in proteins, Annu Rev Biophys Biophys Chem., 19, pp 159– 188 12 Plaza del Pino, I M., Ibarra-Molero, B., and Sanchez-Ruiz, J M (2000) Lower kinetic limit to protein kinetic stability: a proposal regarding protein stability in vivo and its relation with misfolding diseases, Proteins, 40, pp 58–70 13 Sanchez-Ruiz, J M (1992) Theoretical analysis of Lumry-Eyring models in differential scanning calorimetry, Biophys J., 61, pp 921–935 14 Chang, B S., and Hershenson, S (2002) Practical approaches to protein formulation development In Rational Design of Stable Protein Formulations: Theory and Practice, Carpenter, J F., and Manning, M C., eds (Kluwer Academic/Plenum, New York), pp 1–25 15 Sanchez-Ruiz, J M (2010) Protein kinetic stability, Biophys Chem., 148, pp 1–15 16 Pace, C N., Grimsley, G R., Thomson, J A., and Barnett, B J (1988) Conformational stability and activity of ribonucelase T1 with zero, one, and two intact disulfide bonds, J Biol Chem., 263, pp 11820– 11825 17 Zhang, T., Bertelsen, E., and Alber, T (1994) Entropic effects of disulfide bonds on protein stability, Nat Struct Biol., 1, pp 434–438 18 Gribenko, A V., Patel, M M., Liu, J., McCallum S A., Wang, C., and Makhatadze, G I (2009) Rational stabilization of enzymes by computational redesign of surface charge interactions, Proc Natl Acad Sci U S A, 106, pp 2601–2606 19 Ibarra-Molero, B., and Sanchez-Ruiz, J M (2002) Genetic algorithm to design stabilizing surface-charge distributions in proteins, J Phys Chem B, 106, pp 6609–6613 20 Sanchez-Ruiz, J M., and Makhatadze, G I (2001) To charge or not to charge?, Trends Biotechnol., 19, pp 132–135 21 Malakauskas, S M., and Mayo, S L (1998) Design, structure and stability of a hyperthermophilic protein variant, Nat Struct Biol., 5, pp 470–475 25-Allan-Svendsen-c25 March 23, 2016 13:17 PSP Book - 9in x 6in 25-Allan-Svendsen-c25 References 855 22 Potapov, V., Cohen, M., and Schreiber, G (2009) Assessing computational methods for predicting protein stability upon mutation: good on average but not in the details, Protein Eng., 22, pp 553–560 23 Creamer, T P., Srinivasan, R., and Rose, G D (1995) Modeling unfolded states of peptides and proteins, Biochemsitry, 34, pp 16245–16250 24 Fu, H., Grimsley, G., Scholtz, J M., and Pace, C N (2010) Increasing protein stability: importance of DeltaC(p) and the denatured state, Protein Sci., 19, pp 1044–1052 25 Guzman-Casado, M., Parody-Morreale, A., Robic, S., Marqusee, S., and Sanchez-Ruiz, J M (2003) Energetic evidence for the formation of a pH-dependent hydrophobic cluster in the denatured state of Thermus thermophilus ribonuclease H, J Mol Biol., 329, pp 731–743 26 Robic, S., Guzman-Casado, M., Sanchez-Ruiz, J M., and Marqusee, S (2003) Role of residual structure in the unfolded state of a thermophilic protein, Proc Natl Acad Sci U S A, 100, pp 11345–11349 27 Xiao, S., Patsalo, V., Shan, B., Bi, Y., Green, D F., and Raleigh D P (2013) Rational modification of protein stability by targeting surface sites leads to complicated results, Proc Natl Acad Sci U S A, 110, pp 11337– 11342 28 Bryngelson, J D., Onuchic, J N., Socci, N D., and Wolynes, P G (1995) Funnels, pathways, and the energy landscape of protein folding: a synthesis, Proteins, 21, pp 167–195 29 Dill, K A., and Chan, H S (1996) From Levinthal to pathways to funnels, Nat Struct Biol., 4, pp 10–19 30 Sanchez-Ruiz, J M (2011) Probing free-energy surfaces with differential scanning calorimetry, Annu Rev Phys Chem., 62, pp 231–255 ˜ 31 Godoy-Ruiz, R., Henry E R., Kubelka, J., Hofrichter J., Munoz, V., SanchezRuiz, J M., and Eaton, W A (2008) Estimating free-energy barrier heights for an ultrafast folding protein from calorimetric and kinetic data, J Phys Chem B, 112, 5398–5349 ˜ 32 Naganathan, A N., Sanchez-Ruiz, J M., and Munoz, V (2005) Direct measurement of barrier heights in protein folding, J Am Chem Soc., 127, pp 17970–17971 33 Naganathan, A N., Li, P., Perez-Jimenez, R., Sanchez-Ruiz, J M., and ˜ Munoz, V (2010) Navigating the downhill protein folding regime via structural homologues, J Am Chem Soc., 132, pp 11183–11190 34 Fersht, A R., Matouscheck, A., and Serrano, L., (1992) The folding of an enzyme: I Theory of protein engineering analysis of stability and pathway of protein folding, J Mol Biol., 224, pp 771–782 March 23, 2016 13:17 PSP Book - 9in x 6in 856 Kinetic Stability of Variant Enzymes ˜ 35 Naganathan, A N., and Munoz, V (2010) Insights into protein folding mechanisms from large scale analysis of mutational effects, Proc Natl Acad Sci U S A, 107, pp 8611–8616 36 Jaswal, S S., Sohl, J L., and Agard, D A (2002) Energetic landscape of the α-lytic protease optimizes longevity through kinetic stability, Nature, 415, pp 343–346 37 Kelch, B A., and Agard, D A (2007) Mesophile versus thermophile: insights into the structural mechanism of kinetic stability, J Mol Biol., 370, pp 784–795 38 Kelch, B A., Eagen, K P., Erciyas, F P., Hunphris, E L., Thomason, A R., Mitsuiki, S., and Agard, D A (2007) Structural and mechanistic exploration of acid resistance: kinetic stability facilitates evolution of extremophilic behavior, J Mol Biol., 368, pp 870–883 39 Salimi, N L., Ho, B., and Agard, D A (2010) Unfolding simulations reveal the mechanism of extreme unfolding cooperativity in the kinetically stable α-lytic protease, PLOS Comput Biol., 6, p e1000689 40 Kelch, B A., Salimi, N L., and Agard, D A (2012) Functional modulation of a protein folding landscape via side-chain distortion, Proc Natl Acad Sci U S A, 109, pp 9414–9419 41 Godoy-Ruiz, R., Perez-Jimenez, R., Ibarra-Molero, B., and Sanchez-Ruiz, J M (2004) Relation between protein stability, evolution and structure, as probed by carboxylic acid mutations, J Mol Biol., 336, pp 313– 318 42 Godoy-Ruiz, R., Perez-Jimenez, R., Ibarra-Molero, B., and Sanchez-Ruiz, J M (2005) A stability pattern of protein hydrophobic mutations that reflects evolutionary structural information, Biophys J., 89, pp 3320– 3331 43 Godoy-Ruiz, R., Ariza, F., Rodriguez-Larrea, D., Perez-Jimenez, R., IbarraMolero, B., and Sanchez-Ruiz, J M (2006) Natural selection for kinetic stability is a likely origin of correlations between mutational effects on protein energetics and frequencies o amino acid occurrences in sequence alignments, J Mol Biol., 362, pp 966–978 44 Tur-Arlandis, G., Rodriguez-Larrea, D., Ibarra-Molero, B., and SanchezRuiz, J M (2010) Proteolytic scanning calorimetry: a novel methodology that probes the fundamental features of protein kinetic stability, Biophys J., 98, pp L12–L14 45 Pey, A L., Rodriguez-Larrea, D., Bomke, S., Dammers, S., Godoy-Ruiz, R., Garcia-Mira, M M., and Sanchez-Ruiz, J M (2008) Engineering proteins 25-Allan-Svendsen-c25 March 23, 2016 13:17 PSP Book - 9in x 6in 25-Allan-Svendsen-c25 References 857 with tunable thermodynamic and kinetic stabilities, Proteins, 71, pp 165–174 46 Sanchez-Romero, I., Ariza, A., Wilson, K S., Skjøt, M., Vind, J., De Maria, L., Skov, L K., and Sanchez-Ruiz, J M (2013) Mechanism of protein kinetic stabilization by engineered disulfide crosslinks, PLOS ONE, 8, p e70013 47 Chan, H S., Zhang, Z., Wallin, S., and Liu, Z (2011) Cooperativity, localnonlocal coupling, and nonnative interactions: principles of protein folding from coarse-grained models, Annu Rev Phys Chem., 62, pp 301– 326 48 Cheung, S M., Garcia, A E., and Onuchic, J N (2002) Protein folding mediated by solvation: water expulsion and formation of the hydrophobic core occur after structural collapse, Proc Natl Acad Sci U S A, 99, pp 685–690 49 Ferguson, A., Liu, Z., and Chan, H S (2009) Desolvation barrier effects are a likely contributor to the remarkable diversity in the folding rates of small proteins, J Mol Biol., 389, pp 619–636 50 Rank, J A., and Baker, D (1997) A desolvation barrier to hydrophobic cluster formation may contribute to the rate-limiting step in protein folding, Protein Sci., 6, pp 347–354 51 Rodriguez-Larrea, D., Ibarra-Molero, B., and Sanchez-Ruiz, J M (2006) Energetic and structural consequences of desolvation/solvation barriers to protein folding/unfolding assessed from experimental unfolding rates, Biophys J., 91, pp L48–L50 52 Rodriguez-Larrea, D., Minning, S., Borchert, T V., and Sanchez-Ruiz, J M (2006) Role of solvation barriers in protein kinetic stability, J Mol Biol., 360, pp 715–724 53 Costas, M., Rodriguez-Larrea, D., De Maria, L., Borchert, T V., GomezPuyou, A., and Sanchez-Ruiz, J M (2009) Between-species variation in the kinetic stability of TIM proteins linked to solvation-barrier free energies, J Mol Biol., 385, pp 924–927 54 Aguirre, Y., Cabrera, N., Aguirre, B., Perez-Monfort, R., Hernandez´ Santoyo, A., Reyes-Vivas, H., Enriquez-Flores, S., Tuena de Gomez-Puyou, ´ M., Gomez-Puyou, A., Sanchez-Ruiz, J M., and Costas, M (2014) Different contributions of conserved amino acids to the global properties of triosephosphate isomerases, Proteins, 82, pp 323–335 55 Cohen, S I., Vendruscolo, M., Dobson, C M., and Knowles, T P (2012) From macroscopic measurements to microscopic to microscopic mechanisms of protein aggregation, J Mol Biol., 421, pp 160–171 March 23, 2016 13:17 PSP Book - 9in x 6in 858 Kinetic Stability of Variant Enzymes 56 Roberts, C J., Das, T K., and Sahin, E (2011) Predicting solution aggregation rates for therapeutic proteins: approaches and challenges, Int J Pharm., 418, pp 318–333 57 Weiss, W F., Young, T M., and Roberts, C J (2009) Principles, approaches and challenges for predicting protein aggregation rates and shelf life, J Pharm Sci., 98, pp 1246–1277 25-Allan-Svendsen-c25