Enzymes: An integrated view of structure, dynamics and function
© Agarwal; licensee BioMed Central Ltd. 2006
Received: 08 November 2005
Accepted: 12 January 2006
Published: 12 January 2006
Microbes utilize enzymes to perform a variety of functions. Enzymes are biocatalysts working as highly efficient machines at the molecular level. In the past, enzymes have been viewed as static entities and their function has been explained on the basis of direct structural interactions between the enzyme and the substrate. A variety of experimental and computational techniques, however, continue to reveal that proteins are dynamically active machines, with various parts exhibiting internal motions at a wide range of time-scales. Increasing evidence also indicates that these internal protein motions play a role in promoting protein function such as enzyme catalysis. Moreover, the thermodynamical fluctuations of the solvent, surrounding the protein, have an impact on internal protein motions and, therefore, on enzyme function. In this review, we describe recent biochemical and theoretical investigations of internal protein dynamics linked to enzyme catalysis. In the enzyme cyclophilin A, investigations have lead to the discovery of a network of protein vibrations promoting catalysis. Cyclophilin A catalyzes peptidyl-prolyl cis/trans isomerization in a variety of peptide and protein substrates. Recent studies of cyclophilin A are discussed in detail and other enzymes (dihydrofolate reductase and liver alcohol dehydrogenase) where similar discoveries have been reported are also briefly discussed. The detailed characterization of the discovered networks indicates that protein dynamics plays a role in rate-enhancement achieved by enzymes. An integrated view of enzyme structure, dynamics and function have wide implications in understanding allosteric and co-operative effects, as well as protein engineering of more efficient enzymes and novel drug design.
An integrated view of protein structure, dynamics, and function is emerging, where proteins are considered as dynamically active machines and internal protein motions are closely linked to function such as enzyme catalysis. Currently there is wide interest, both from experimental and computational groups, in investigating this interconnection. A number of investigations have provided fascinating details about the movement of protein parts and their involvement in enzyme function. Techniques such as X-ray crystallography and small-angle scattering [13, 14], NMR studies [15–17], hydrogen-deuterium exchange , neutron scattering , biochemical and mutational analysis [7, 20, 21] have provided vital clues at individual time-scales; however, the detailed understanding of protein dynamics requires information over a broad range of time-scales. Moreover, the hydration-shell and bulk solvent fluctuations have been suggested to impact protein dynamics, and therefore, protein function [22, 23]. Theoretical studies and computational modeling are playing a vital role in investigating the link between protein dynamics, solvent fluctuations and enzyme catalysis at multiple time-scales [8, 10–12].
In this review, we describe recent biochemical and theoretical/computational studies that have investigated the link between protein dynamics and enzyme catalysis. In particular, we describe the recent investigations of the peptidyl-prolyl cis/trans isomerization activity of the enzyme cyclophilin A, followed by a discussion on similar evidence from other enzyme reactions, namely the hydride transfer reactions catalyzed by dihydrofolate reductase and by liver alcohol dehydrogenase. There are wide implications of understanding the interconnection between protein structure, dynamics and function such as enzyme catalysis. It is known that enzymes catalyzing the same reactions belong to a protein "fold" family, where the overall characteristic shape of the protein is similar. Also, enzymes catalyzing mechanistically similar reactions often belong to the same super-family of protein fold. The benefits of better understanding of enzyme "folds" and dynamics include the possibility of improving the efficiency of microbial factories by engineering of enzymes, as well as designing new enzymes with novel functionalities. Further, there are medical implications of allosteric and cooperative effects for enzyme activity in novel drug design.
A number of factors make CypA an attractive system for investigating the link between internal protein dynamics and enzymatic activity; it is a small protein and does not require metal ions or cofactors for PPIase activity and it catalyzes peptide bond isomerization in a wide variety of substrates. Further, there is also biomedical interest in CypA; cyclophilins are of interest as drug targets because of their likely involvement in the broad spectrum, anti-infective activities of cyclosporin A and non-immunosuppressive derivatives thereof [32, 33]. In addition, Gag-encoded capsid protein (CA) from human immunodeficiency virus type 1 (HIV-1), is a naturally occurring biologically relevant substrate for CypA . The protein-protein complex between CypA and CA has been the subject of many experimental studies [16, 34–37]. There is medical interest in CypA-CA complex, as incorporation of CypA is required for infectious activity of HIV-1 [38, 39].
NMR studies of CypA performed by Kern and coworkers, have suggested a link between internal protein dynamics and substrate isomerization step [15, 16]. The studies were based on 15N spin relaxation investigations of small peptide substrate as well as two-dimensional (2D) 1H-15N heteronuclear exchange studies of the N-terminal of capsid protein (CAN) from HIV-1. In these studies, conformational fluctuations within the active-site of CypA were detected on the time-scale of the reaction (hundreds of microseconds) and the rates of conformational dynamics were found to be strongly correlated with the substrate isomerization step. Several active-site and surface loop regions showed motions only in the presence of substrate, these regions included the residues: Arg55, Lys82, Leu98, Ser99, Ala101, Asn102, Ala103, and Gly109. Based on these studies, the authors proposed a reaction mechanism for PPIase activity of CypA, where the isomerization step takes place with a rate constant of about 9000 s-1, and motions of the protein coincide with the rate of substrate turnover step. CypA residue Arg55 is a major contributor to catalysis , for which the observed changes in backbone conformation are likely to be coupled with motions of the catalytically essential side chain.
Structural analysis of the active-site along the course of reaction indicates the role of important hydrophilic (Arg55 and Asn102) and hydrophobic (Phe60, Phe113, Leu122 and His126) residues of CypA in stabilization of the substrate peptide. The target proline from substrate remains essentially fixed in the hydrophobic pocket formed by CypA residues, while the carbonyl oxygen atom of the preceding substrate residue rotates 180°. Quantum mechanical modeling of the active-site indicates single bond character for the peptide bond near the TS. The results from theoretical modeling were found to be in agreement with the reaction mechanism proposed on the basis of crystallographic studies . This mechanism requires minimum deviation from the ground state crystal structure and displays single bond character for the peptide bond near TS. Dynamical fluctuations of the enzyme backbone in certain regions (CypA 101–104) were found to impact the nature of interactions between the enzyme and substrate, therefore, alter the nature of peptide bond during the course of reaction mechanism.
Computational modeling has identified a variety of internal protein dynamical events linked to CypA enzyme activity, ranging from femtosecond (10-15 s) to microsecond and longer (> 10-6 s) time-scales. On one side of this range there are fast motions, occurring at femtosecond-nanosecond time-scales, consisting of harmonic movements of bonds, angles and a few atoms. These motions are commonly referred to as vibrations. On the other side of this range there are concerted conformational fluctuations occurring on the microsecond (and longer) time-scale. These slower motions or conformational fluctuations, which have been previously referred to as breathing motions, span a large part of the protein. Normal mode analysis is a computational technique that has been commonly used to obtain information regarding the dynamical motions in proteins. This technique provides information about dynamics at several time-scales for a particular protein conformation (present at a local minimum). Normal mode analysis is not suitable for obtaining the slow protein motions occurring at the time-scale of the reaction due to the large changes in protein conformations involved. Another computational technique, known as quasi-harmonic analysis, can be used to calculate vibrational modes from a collection of conformations or system snapshots . Quasi-harmonic analysis of CypA-substrate conformations along the entire reaction pathway provided protein vibrational modes representing conformational fluctuations at the time-scale of the reaction (microsecond-millisecond time-scale). These computed slow protein vibrational modes show concerted motions over a large region of the protein, the backbone in several regions of the protein and side-chains of the many residues (especially on the surface) show large displacements. In CypA, a subset of these modes was found to be coupled to the reaction; 3 protein vibrational modes with the largest coupling to the catalytic step show displacements in several conserved residues in the active-site as well as in other parts of the enzyme structure. Note, these conserved vibrational modes are different from random thermal fluctuations observed in the biomolecules.
Conservation of network hydrogen bonds in cyclophilin structures from various species. 3-dimensional structures were aligned using secondary structure elements and equivalent hydrogen bonds were selected based on sequence and structural similarities. Hydrogen bond lengths are in given Å and PDB codes are given in parenthesis . Reprinted with permission from Agarwal et al., Biochemistry (2004) 43, 10605–10618. Copyright (2004) American Chemical Society.
CypA (1AWQ/2CYH/1RMH average)
Human Cyclophilin B
Cyclophilin 3 (1DYW)
Cyclophilin 5 (1H0P)
Other enzymes: dihydrofolate reductase and liver alcohol dehydrogenase
Experimental and computational investigations have revealed the impact of protein dynamics on catalysis in other enzyme systems. Experimental and computational studies of the enzyme dihydrofolate reductase (DHFR) have indicated a link between protein dynamical events and the substrate turnover step of hydride transfer. X-ray crystallography has demonstrated changes in orientation of surface loops along different sub-states along the reaction pathway . Similarly, the surface loop conformations have been linked to the catalytic step by NMR studies . Theoretical and computational studies using hybrid quantum-mechanical and molecular mechanics (QM/MM) methodology have discovered a network of coupled promoting motions [8, 52, 53]. Similar to the network of protein vibrations in CypA described above, the network in DHFR is also formed by interconnection of residues and crucial interactions ranging from surface regions all the way to active-site. Changes in hydrogen-bonds and crucial interactions along the reaction profile have been observed similar to those present during catalysis by CypA. An important discovery by the computational methods was the identification of the residue Ile14 as a dynamical contributor to catalysis. Recently, the importance of this residue in the catalytic step has been confirmed by NMR studies . The presence of this DHFR network has been confirmed by investigations from several research groups [55, 56]. Investigations of DHFR have provided evidence that changing the enzyme structure leads to changed dynamics and, therefore, change in function [21, 53, 57]. Mutation of a single surface residue, more than 12 Å away from active site, changes the dynamics and leads to a rate reduction by a factor of 163.
Liver alcohol dehydrogenase (LADH) is another enzyme where dynamical motions of the protein residues have been linked to the catalytic step. Detailed biochemical and computational studies have identified conserved active-site residue Val203, whose motion are a key player in altering the active-site chemical environment to promote the reaction [58–63].
In this review, we have presented recent developments that continue to support an emerging integrated view of protein structure, dynamics and function such as enzyme catalysis. The success of microbial cell factories depends on optimal performance of molecular machines inside the cell. Enzymes perform their function with remarkable efficiency, as they increase the reaction rate by many orders of magnitude. Until recently, enzymes (and proteins in general) were considered static assemblies; however, recent investigations continue to provide evidence which indicate that enzymes are dynamically active assemblies. Detailed experimental and theoretical/computational investigations of enzyme CypA suggest that the internal protein motions are a designed part of the protein structure and contribute to its function of catalyzing peptidyl-prolyl cis/trans isomerization. Supporting evidence from other systems (DHFR and LADH) indicates that the interconnection between structure, dynamics and function is present in other enzymes as well.
The integrated view is supported by evidence from investigations of many other proteins and enzymes as well [64–66]. Sequence analysis with thermodynamic mapping have indicated long range energetic coupling in proteins ; slow conformational fluctuations could possibly be the mechanism of energy transfer over long ranges in protein structure and, therefore, provide insights into understanding allosteric effects. Simulations have already revealed that energy can be transferred between specific vibrational modes in a protein [68, 69]. It is also interesting to note that designing active-site mimics of the enzymes is difficult and change in enzyme structure far away from the active-site leads to slow or inactive enzymes. The integrated view offers a possible explanation, as the distal regions of the enzyme contribute to catalysis through dynamical coupling with the solvent and by transferring the required energy to the active-site. Therefore, this integrated view has wide implications in enzyme chemistry, protein engineering and drug design. Manipulation of enzyme catalyzed reactions may be possible; for example, laser pulse has already been used to initiate an enzyme reaction involving thermally excited protein dynamics (molecular motions on picosecond time-scale) . On the basis of better understanding of enzyme structure, dynamics and function it may be possible to design more efficient enzymes or enzymes with novel functionalities. Further, the understanding of allosteric and cooperative effects could help in designing better and novel drugs.
List of abbreviations
activation energy barrier (energy difference between reactant and the activated state)
capsid protein from HIV-1
N-terminal of capsid protein
human immunodeficiency virus type 1
liver alcohol dehydrogenase
nuclear magnetic resonance
peptidyl-prolyl cis/trans isomerase
transition state theory
PKA would like to thank Dr. Brahma Ghosh for feedback on the manuscript.
- Neet KE: Enzyme catalytic power minireview series. J Biol Chem. 1998, 273: 25527-25528. 10.1074/jbc.273.40.25527.View ArticleGoogle Scholar
- Radzicka A, Wolfenden R: A Proficient Enzyme. Science. 1995, 267: 90-93.View ArticleGoogle Scholar
- Kraut J: How Do Enzymes Work. Science. 1988, 242: 533-540.View ArticleGoogle Scholar
- Knowles JR: Enzyme Catalysis - Not Different, Just Better. Nature. 1991, 350: 121-124. 10.1038/350121a0.View ArticleGoogle Scholar
- Fischer E: Ber Dtsch Chem Ges. 1894, 27: 3189-View ArticleGoogle Scholar
- Haldane JBS: Enzymes. 1930, London, Longmans, GreenGoogle Scholar
- Cannon WR, Benkovic SJ: Solvation, reorganization energy, and biological catalysis. J Biol Chem. 1998, 273: 26257-26260. 10.1074/jbc.273.41.26257.View ArticleGoogle Scholar
- Agarwal PK, Billeter SR, Rajagopalan PTR, Benkovic SJ, Hammes-Schiffer S: Network of coupled promoting motions in enzyme catalysis. Proc Natl Acad Sci U S A. 2002, 99: 2794-2799. 10.1073/pnas.052005999.View ArticleGoogle Scholar
- Benkovic SJ, Hammes-Schiffer S: A perspective on enzyme catalysis. Science. 2003, 301: 1196-1202. 10.1126/science.1085515.View ArticleGoogle Scholar
- Agarwal PK, Geist A, Gorin A: Protein dynamics and enzymatic catalysis: investigating the peptidyl-prolyl cis-trans isomerization activity of cyclophilin A. Biochemistry. 2004, 43: 10605-10618. 10.1021/bi0495228.View ArticleGoogle Scholar
- Agarwal PK: Cis/trans isomerization in HIV-1 capsid protein catalyzed by cyclophilin A: insights from computational and theoretical studies. Proteins: Struct Func Bioinform. 2004, 56: 449-463. 10.1002/prot.20135.View ArticleGoogle Scholar
- Agarwal PK: Role of protein dynamics in reaction rate enhancement by enzymes. J Am Chem Soc. 2005, 127: 15248-15256. 10.1021/ja055251s.View ArticleGoogle Scholar
- Schramm VL, Shi WX: Atomic motion in enzymatic reaction coordinates. Curr Opin Struct Biol. 2001, 11: 657-665. 10.1016/S0959-440X(01)00269-X.View ArticleGoogle Scholar
- Heller WT: Influence of multiple well defined conformations on small-angle scattering of proteins in solution. Acta Crystallogr D. 2005, 61: 33-44. 10.1107/S0907444904025855.View ArticleGoogle Scholar
- Eisenmesser EZ, Bosco DA, Akke M, Kern D: Enzyme dynamics during catalysis. Science. 2002, 295: 1520-1523. 10.1126/science.1066176.View ArticleGoogle Scholar
- Bosco DA, Eisenmesser EZ, Pochapsky S, Sundquist WI, Kern D: Catalysis of cis/trans isomerization in native HIV-1 capsid by human cyclophilin A. Proc Natl Acad Sci U S A. 2002, 99: 5247-5252. 10.1073/pnas.082100499.View ArticleGoogle Scholar
- Wand AJ: Dynamic activation of protein function: A view emerging from NMR spectroscopy. Nat Struct Biol. 2001, 8: 926-931. 10.1038/nsb1101-926.View ArticleGoogle Scholar
- Zavodszky P, Kardos J, Svingor A, Petsko GA: Adjustment of conformational flexibility is a key event in the thermal adaptation of proteins. Proc Natl Acad Sci U S A. 1998, 95: 7406-7411. 10.1073/pnas.95.13.7406.View ArticleGoogle Scholar
- Zaccai G: Biochemistry - How soft is a protein? A protein dynamics force constant measured by neutron scattering. Science. 2000, 288: 1604-1607. 10.1126/science.288.5471.1604.View ArticleGoogle Scholar
- Osborne MJ, Schnell J, Benkovic SJ, Dyson HJ, Wright PE: Backbone dynamics in dihydrofolate reductase complexes: Role of loop flexibility in the catalytic mechanism. Biochemistry. 2001, 40: 9846-9859. 10.1021/bi010621k.View ArticleGoogle Scholar
- Cameron CE, Benkovic SJ: Evidence for a functional role of the dynamics of glycine-121 of Escherichia coli dihydrofolate reductase obtained from kinetic analysis of a site-directed mutant. Biochemistry. 1997, 36: 15792-15800. 10.1021/bi9716231.View ArticleGoogle Scholar
- Fenimore PW, Frauenfelder H, McMahon BH, Young RD: Bulk Solvent and hydration-shell fluctuations, similar to a- and b-fluctuations in glasses, control protein motions and functions. P Natl Acad Sci USA. 2004, 101: 14408-14413. 10.1073/pnas.0405573101.View ArticleGoogle Scholar
- Frauenfelder H, Fenimore PW, McMahon BH: Hydration, slaving and protein function. Biophys Chem. 2002, 98: 35-48. 10.1016/S0301-4622(02)00083-2.View ArticleGoogle Scholar
- Handschumacher RE, Harding MW, Rice J, Drugge RJ: Cyclophilin - a Specific Cytosolic Binding-Protein for Cyclosporin-A. Science. 1984, 226: 544-547.View ArticleGoogle Scholar
- Takahashi N, Hayano T, Suzuki M: Peptidyl-Prolyl Cis-Trans Isomerase Is the Cyclosporin-a-Binding Protein Cyclophilin. Nature. 1989, 337: 473-475. 10.1038/337473a0.View ArticleGoogle Scholar
- Gothel SF, Marahiel MA: Peptidyl-prolyl cis-trans isomerases, a superfamily of ubiquitous folding catalysts. Cell Mol Life Sci. 1999, 55: 423-436. 10.1007/s000180050299.View ArticleGoogle Scholar
- Rovira P, Mascarell L, Truffa-Bachi P: The impact of immunosuppressive drugs on the analysis of T-cell activation. Curr Med Chem. 2000, 7: 673-692.View ArticleGoogle Scholar
- Fischer G: Chemical aspects of peptide bond isomerisation. Chem Soc Rev. 2000, 29: 119-127. 10.1039/a803742f.View ArticleGoogle Scholar
- Zhao YD, Ke HM: Crystal structure implies that cyclophilin predominantly catalyzes the trans to cis isomerization. Biochemistry. 1996, 35: 7356-7361. 10.1021/bi9602775.View ArticleGoogle Scholar
- Zhao YD, Ke HM: Mechanistic implication of crystal structures of the cyclophilin-dipeptide complexes. Biochemistry. 1996, 35: 7362-7368. 10.1021/bi960278x.View ArticleGoogle Scholar
- Vajdos FE, Yoo SH, Houseweart M, Sundquist WI, Hill CP: Crystal structure of cyclophilin A complexed with a binding site peptide from the HIV-1 capsid protein. Protein Sci. 1997, 6: 2297-2307.View ArticleGoogle Scholar
- Page AP, Kumar S, Carlow CKS: Parasite Cyclophilins and Antiparasite Activity of Cyclosporine-A. Parasitology Today. 1995, 11: 385-388. 10.1016/0169-4758(95)80007-7.View ArticleGoogle Scholar
- Chappell LH, Wastling JM: Cyclosporine-a - Antiparasite Drug, Modulator of the Host-Parasite Relationship and Immunosuppressant. Parasitology. 1992, 105: S25-S40.View ArticleGoogle Scholar
- Gamble TR, Vajdos FF, Yoo SH, Worthylake DK, Houseweart M, Sundquist WI, Hill CP: Crystal structure of human cyclophilin A bound to the amino- terminal domain of HIV-1 capsid. Cell. 1996, 87: 1285-1294. 10.1016/S0092-8674(00)81823-1.View ArticleGoogle Scholar
- Yoo SH, Myszka DG, Yeh CY, McMurray M, Hill CP, Sundquist WI: Molecular recognition in the HIV-1 capsid/cyclophilin a complex. J Mol Biol. 1997, 269: 780-795. 10.1006/jmbi.1997.1051.View ArticleGoogle Scholar
- Saphire ACS, Bobardt MD, Gallay PA: trans-complementation rescue of cyclophilin A-deficient viruses reveals that the requirement for cyclophilin A in human immunodeficiency virus type 1 replication is independent of its isomerase activity. J Virol. 2002, 76: 2255-2262. 10.1128/jvi.76.5.2255-2262.2002.View ArticleGoogle Scholar
- Howard BR, Vajdos FF, Li S, Sundquist WI, Hill CP: Structural insights into the catalytic mechanism of cyclophilin A. Nat Struct Biol. 2003, 10: 475-481. 10.1038/nsb927.View ArticleGoogle Scholar
- Braaten D, Franke EK, Luban J: Cyclophilin A is required for an early step in the life cycle of human immunodeficiency virus type 1 before the initiation of reverse transcription. J Virol. 1996, 70: 3551-3560.Google Scholar
- Wiegers K, Krausslich HG: Differential dependence of the infectivity of HIV-1 group O isolates on the cellular protein cyclophilin A. Virology. 2002, 294: 289-295. 10.1006/viro.2001.1347.View ArticleGoogle Scholar
- Li G, Cui Q: What is so special about Arg 55 in the catalysis of cyclophilin A? insights from hybrid QM/MM simulations. J Am Chem Soc. 2003, 125: 15028-15038. 10.1021/ja0367851.View ArticleGoogle Scholar
- Garcia-Viloca M, Gao J, Karplus M, Truhlar DG: How enzymes work: Analysis by modern rate theory and computer simulations. Science. 2004, 303: 186-195. 10.1126/science.1088172.View ArticleGoogle Scholar
- Torrie GM, Valleau JP: Non-Physical Sampling Distributions in Monte-Carlo Free-Energy Estimation - Umbrella Sampling. J Comput Phys. 1977, 23: 187-199. 10.1016/0021-9991(77)90121-8.View ArticleGoogle Scholar
- Kumar S, Bouzida D, Swendsen RH, Kollman PA, Rosenberg JM: The Weighted Histogram Analysis Method for Free-Energy Calculations on Biomolecules .1. The Method. J Comput Chem. 1992, 13: 1011-1021. 10.1002/jcc.540130812.View ArticleGoogle Scholar
- Levy RM, Karplus M, Kushick J, Perahia D: Evaluation of the Configurational Entropy for Proteins - Application to Molecular-Dynamics Simulations of an Alpha-Helix. Macromolecules. 1984, 17: 1370-1374. 10.1021/ma00137a013.View ArticleGoogle Scholar
- Eisenmesser EZ, Millet O, Labeikovsky W, Korzhnev DM, Wolf-Watz M, Bosco DA, Skalicky JJ, Kay LE, Kern D: Intrinsic dynamics of an enzyme underlies catalysis. Nature. 2005, 438: 117-121. 10.1038/nature04105.View ArticleGoogle Scholar
- Bouvignies G, Bernado P, Meier S, Cho K, Grzesiek S, Bruschweiler R, Blackledge M: Identification of slow correlated motions in proteins using residual dipolar and hydrogen-bond scalar couplings. P Natl Acad Sci USA. 2005, 102: 13885-13890. 10.1073/pnas.0505129102.View ArticleGoogle Scholar
- Tournier AL, Xu JC, Smith JC: Translational hydration water dynamics drives the protein glass transition. Biophys J. 2003, 85: 1871-1875.View ArticleGoogle Scholar
- Tournier AL, Xu JC, Smith JC: Solvent caging of internal motions in myoglobin at low temperatures. Physchemcomm. 2003, 6-8. 10.1039/b209839c.Google Scholar
- Gerlt JA, Babbitt PC: Mechanistically diverse enzyme superfamilies: the importance of chemistry in the evolution of catalysis. Curr Opin Chem Biol. 1998, 2: 607-612. 10.1016/S1367-5931(98)80091-4.View ArticleGoogle Scholar
- Babbitt PC, Gerlt JA: Understanding enzyme superfamilies - Chemistry as the fundamental determinant in the evolution of new catalytic activities. J Biol Chem. 1997, 272: 30591-30594. 10.1074/jbc.272.49.30591.View ArticleGoogle Scholar
- Sawaya MR, Kraut J: Loop and subdomain movements in the mechanism of Escherichia coli dihydrofolate reductase: Crystallographic evidence. Biochemistry. 1997, 36: 586-603. 10.1021/bi962337c.View ArticleGoogle Scholar
- Agarwal PK, Billeter SR, Hammes-Schiffer S: Nuclear quantum effects and enzyme dynamics in dihydrofolate reductase catalysis. J Phys Chem B. 2002, 106: 3283-3293. 10.1021/jp020190v.View ArticleGoogle Scholar
- Watney JB, Agarwal PK, Hammes-Schiffer S: Effect of mutation on enzyme motion in dihydrofolate reductase. J Am Chem Soc. 2003, 125: 3745-3750. 10.1021/ja028487u.View ArticleGoogle Scholar
- Schnell JR, Dyson HJ, Wright PE: Effect of cofactor binding and loop conformation on side chain methyl dynamics in dihydrofolate reductase. Biochemistry. 2004, 43: 374-383. 10.1021/bi035464z.View ArticleGoogle Scholar
- Garcia-Viloca M, Truhlar DG, Gao JL: Reaction-path energetics and kinetics of the hydride transfer reaction catalyzed by dihydrofolate reductase. Biochemistry. 2003, 42: 13558-13575. 10.1021/bi034824f.View ArticleGoogle Scholar
- Thorpe IF, Brooks CL: The coupling of structural fluctuations to hydride transfer in dihydrofolate reductase. Proteins: Struct Func Bioform. 2004, 57: 444-457. 10.1002/prot.20219.View ArticleGoogle Scholar
- Rajagopalan PTR, Lutz S, Benkovic SJ: Coupling interactions of distal residues enhance dihydrofolate reductase catalysis: Mutational effects on hydride transfer rates. Biochemistry. 2002, 41: 12618-12628. 10.1021/bi026369d.View ArticleGoogle Scholar
- Bahnson BJ, Colby TD, Chin JK, Goldstein BM, Klinman JP: A link between protein structure and enzyme catalyzed hydrogen tunneling. Proc Natl Acad Sci U S A. 1997, 94: 12797-12802. 10.1073/pnas.94.24.12797.View ArticleGoogle Scholar
- Colby TD, Bahnson BJ, Chin JK, Klinman JP, Goldstein BM: Active site modifications in a double mutant of liver alcohol dehydrogenase: Structural studies of two enzyme-ligand complexes. Biochemistry. 1998, 37: 9295-9304. 10.1021/bi973184b.View ArticleGoogle Scholar
- Agarwal PK, Webb SP, Hammes-Schiffer S: Computational studies of the mechanism for proton and hydride transfer in liver alcohol dehydrogenase. J Am Chem Soc. 2000, 122: 4803-4812. 10.1021/ja994456w.View ArticleGoogle Scholar
- Webb SP, Agarwal PK, Hammes-Schiffer S: Combining electronic structure methods with the calculation of hydrogen vibrational wavefunctions: Application to hydride transfer in liver alcohol dehydrogenase. J Phys Chem B. 2000, 104: 8884-8894. 10.1021/jp001635n.View ArticleGoogle Scholar
- Billeter SR, Webb SP, Agarwal PK, Iordanov T, Hammes-Schiffer S: Hydride transfer in liver alcohol dehydrogenase: Quantum dynamics, kinetic isotope effects, and role of enzyme motion. J Am Chem Soc. 2001, 123: 11262-11272. 10.1021/ja011384b.View ArticleGoogle Scholar
- Billeter SR, Webb SP, Iordanov T, Agarwal PK, Hammes-Schiffer S: Hybrid approach for including electronic and nuclear quantum effects in molecular dynamics simulations of hydrogen transfer reactions in enzymes. J Chem Phys. 2001, 114: 6925-6936. 10.1063/1.1356441.View ArticleGoogle Scholar
- Hammes GG: Multiple conformational changes in enzyme catalysis. Biochemistry. 2002, 41: 8221-8228. 10.1021/bi0260839.View ArticleGoogle Scholar
- Kohen A: Kinetic isotope effects as probes for hydrogen tunneling, coupled motion and dynamics contributions to enzyme catalysis. Prog React Kinet Mec. 2003, 28: 119-156.View ArticleGoogle Scholar
- Tousignant A, Pelletier JN: Protein motions promote catalysis. Chem Biol. 2004, 11: 1037-1042. 10.1016/j.chembiol.2004.06.007.View ArticleGoogle Scholar
- Lockless SW, Ranganathan R: Evolutionarily conserved pathways of energetic connectivity in protein families. Science. 1999, 286: 295-299. 10.1126/science.286.5438.295.View ArticleGoogle Scholar
- Moritsugu K, Miyashita O, Kidera A: Vibrational energy transfer in a protein molecule. Phys Rev Lett. 2000, 85: 3970-3973. 10.1103/PhysRevLett.85.3970.View ArticleGoogle Scholar
- Moritsugu K, Miyashita O, Kidera A: Temperature dependence of vibrational energy transfer in a protein molecule. J Phys Chem B. 2003, 107: 3309-3317. 10.1021/jp027823q.View ArticleGoogle Scholar
- Heyes DJ, Hunter CN, van Stokkum IHM, van Grondelle R, Groot ML: Ultrafast enzymatic reaction dynamics in protochlorophyllide oxidoreductase. Nat Struct Biol. 2003, 10: 491-492. 10.1038/nsb929.View ArticleGoogle Scholar
This article is published under license to BioMed Central Ltd. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.