© 2005 Nature Publishing Group Intrinsic dynamics of an enzyme underlies catalysis Elan Z. Eisenmesser 1 , Oscar Millet 2 †, Wladimir Labeikovsky 1 , Dmitry M. Korzhnev 2 , Magnus Wolf-Watz 1 †, Daryl A. Bosco 1 †, Jack J. Skalicky 3 †, Lewis E. Kay 2 & Dorothee Kern 1 A unique feature of chemical catalysis mediated by enzymes is that the catalytically reactive atoms are embedded within a folded protein. Although current understanding of enzyme function has been focused on the chemical reactions and static three- dimensional structures, the dynamic nature of proteins has been proposed to have a function in catalysis 1–5 . The concept of conformational substates has been described 6 ; however, the challenge is to unravel the intimate linkage between protein flexibility and enzymatic function. Here we show that the intrinsic plasticity of the protein is a key characteristic of catalysis. The dynamics of the prolyl cis–trans isomerase cyclophilin A (CypA) in its substrate-free state and during catalysis were characterized with NMR relaxation experiments. The characteristic enzyme motions detected during catalysis are already present in the free enzyme with frequencies corresponding to the catalytic turnover rates. This correlation suggests that the protein motions necessary for catalysis are an intrinsic property of the enzyme and may even limit the overall turnover rate. Motion is localized not only to the active site but also to a wider dynamic network. Whereas coupled networks in proteins have been proposed pre- viously 3,7–10 , we experimentally measured the collective nature of motions with the use of mutant forms of CypA. We propose that the pre-existence of collective dynamics in enzymes before cata- lysis is a common feature of biocatalysts and that proteins have evolved under synergistic pressure between structure and dynamics. The most basic principle of enzyme catalysis is the ability of an enzyme to decrease the transition-state energy, thereby catalysing the chemical reaction. A wealth of information about the kinetics and thermodynamics of enzyme-catalysed reactions has been obtained by monitoring the conversion of substrates into products. However, much less is known about the kinetics and energetics of confor- mational processes in the protein. Because it is the protein com- ponent that alters the transition state energy, enzyme function depends on transitions from ground states to higher-energy states of the enzyme and the reactant. A detailed understanding of the entire trajectory of catalysis is therefore a current challenge. Structures of intermediates provide snapshots of conformational changes, and a set of experimental methods, such as fluorescence, mass spectroscopy and NMR, have been developed to monitor the kinetics of these changes. We have recently provided a proof of principle that enzyme dynamics can be monitored during catalysis at multiple sites by NMR relaxation experiments with the use of cyclophilin A (CypA) 5 . This enzyme belongs to the family of prolyl-isomerases that catalyse the cis–trans isomerization of prolyl peptide bonds. Although CypA is involved in a series of biomedically important processes 11 , its natural biological role and mechanism of action are still in debate. CypA is the target of the immunosuppressive drug cyclosporin A (CsA) and is essential for HIV-1 virulence 12 . Here we use new NMR relaxation dispersion experiments 13 to compare motions in free CypA with those during turnover. We show that protein dynamics associated with catalysis is a built-in property of the enzyme that is also manifested in the free protein. The motions are collective, propagating from the active site to remote sites. The results show that intrinsic plasticity on a basic structural template is a crucial element for catalytic function, and that proteins have there- fore evolved under synergistic pressure between structure and dynamics. Relaxation dispersion experiments probe molecular motions in the microsecond to millisecond timescale quantitatively and with much higher sensitivity than traditional transverse relaxation experi- ments 13 . The additional line-broadening of NMR signals caused by conformational exchange between two states (R ex ) depends on the sum of forward and reverse rates of interconversion (k ex ), the relative populations of the exchanging species (p A and p B ) and the chemical shifts between the exchanging species (Dq), with R ex ¼ p A p B Dq 2 k ex 1 2 4v CPMG k ex tanh k ex 4v CPMG   ð1Þ in the fast exchange limit 14 . The dependence of R ex on the applied external field (v CPMG , in which CPMG stands for Carr–Purcell–Meiboom–Gill) is the key element in relaxation dispersion experiments (Fig. 1a, b). It is noted that an atom with R ex contributions reports on a change in its electronic environment, but this does not necessarily represent physical movement of this atom. We applied this method to 15 N-labelled CypA catalysing the cis–trans isomerization of N-succinyl-Ala-Phe-Pro-Phe-p-nitroanilide (Fig. 1, right). The high sensitivity of these experiments leads to the identification of many more amides in CypA with conformational exchange than originally described 5 . Clear and quantitative analysis of the relaxation data require conditions of two-state exchange. However, CypA interconverts between at least three states during the catalytic cycle: the free enzyme (E) and the two enzyme–substrate (ES) complexes with substrate bound in the cis and trans confor- mations. The system was therefore biochemically ‘tuned’ to two-state exchange: by using an excess of substrate at 10 8C, 95% saturation with substrate could be reached in which the dispersion profiles detect only two-state conformational exchange corresponding to the catalytic step of cis–trans isomerization (scheme (2)). Under these conditions, contributions to R ex from substrate binding and dis- sociation are negligible (see Supplementary Information). Individual fits of dispersion profiles on a collection of 30 amides resulted in similar rate constants, a remarkable result that justifies a LETTERS 1 Department of Biochemistry, Howard Hughes Medical Institute, Brandeis University, Waltham, Massachusetts 02454, USA. 2 Departments of Medical Genetics, Biochemistry and Chemistry, University of Toronto, Toronto, Ontario M5S 1A8, Canada. 3 National High Magnetic Field Laboratory at Florida State University, Tallahassee, Florida 32310, USA. †Present addresses: Plataforma de Biomolecules, Parc Cientific de Barcelona, Josep Samitier 1-5, 08028 Barcelona, Catalonia, Spain (O.M.); The Scripps Research Institute, Department of Chemistry, La Jolla, California 92037, USA (D.A.B.); University of Utah School of Medicine, Department of Biochemistry, Salt Lake City, Utah 84132, USA (J.J.S.); University of Umea ˚, Department of Biochemistry, Umea ˚ SE-901 87, Sweden (M.W.W.). Vol 438|3 November 2005|doi:10.1038/nature04105 117