Observing biological dynamics at atomic resolution using NMR

Observing biological dynamics atatomic resolution using NMRAnthony K. Mittermaier1 and Lewis E. Kay2

1 Department of Chemistry, McGill University, 801 Sherbrooke St. W., #322, Montreal, Quebec, Canada, H3A 2K62 Departments of Chemistry, Biochemistry, and Molecular Genetics, University of Toronto, 1 King’s College Circle, Toronto,

Ontario, Canada, M5S 1A8

Review

Biological macromolecules are highly flexible and con-tinually undergo conformational fluctuations on a broadspectrum of timescales. It has long been recognized thatdynamics have an important role in the action of thesemolecules. However, the relationship between molecu-lar function and motion is extremely challenging todelineate, because the conformational space availableto macromolecules is vast and the relevant excursionscan be infrequent and short-lived. Recent advances insolution nuclear magnetic resonance (NMR) spec-troscopy permit biomolecular dynamics to be observedwith unprecedented detail. Applications of these newNMR techniques to the study of fundamental processessuch as binding and catalysis have provided newinsights into how living systems operate at an atomiclevel.

Molecular dynamics by NMRNuclear magnetic resonance (NMR) spectroscopy isuniquely suited to characterizing biomolecular dynamics[1–4,5]. A typical experiment simultaneously detects sig-nals from many reporters (NMR active nuclei) within amacromolecule, providing a comprehensive description ofinternal motions with atomic resolution; indeed, there areliterally hundreds of nuclei within a biological macromol-ecule that can act as hidden observers of the conformation-al fluctuations that accompany biological activity.Techniques have been developed to quantify motions thattake place over a wide range of timescales, from picose-conds to hours or even days. Samples are largely unaf-fected by the strong magnetic fields and radiofrequency(RF) irradiation applied in theNMRexperiment and can bestudied under physiological conditions, in some cases inliving cells [6].

NMR dynamics experiments were initially performedon proteins nearly forty years ago, providing some of thefirst indications that stable, folded proteins can undergolarge-amplitude motions [7–10]. Since then, the array ofNMR dynamics methods has been enormously expandedand improved. The past few years have produced remark-able NMR studies, driven by simultaneous advancementsin a number of areas, which have increased our under-standing of how macromolecules function. Especially sig-nificant has been the ability to label specific positions inbiomolecules with NMR active nuclei [11] and the conco-mitant development of methodologies that exploit these

Corresponding author: Mittermaier, A.K. ([email protected]).

0968-0004/$ – see front matter � 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.tibs.2009.

novel labeling patterns to significantly extend the sizelimitations of molecules amenable to dynamics studies[12,13]. Developments in NMR hardware that have ledto significant increases in sensitivity of the acquired datahave also been crucial. Finally, computational strategiesfor analyzing and interpreting NMR relaxation data haveemerged [14–18].

In this review, we discuss an array of NMR dynamicstechniques that are used to characterize protein motionson timescales spanning many orders of magnitude(Figure 1). The sensitivity of NMR parameters to molecu-lar dynamics is well illustrated by the scenario shown inFigure 1 in which a probe (nucleus) attached to a moleculereports on conformational exchange between a pair ofstates. Provided that the NMR resonance frequency ofthe probe in each of the two states is different, then largespectral changes are observed as the exchange rate varies.At low exchange frequencies, a pair of peaks derived fromthe individual states is observed in NMR spectra, whereasthe signals broaden and merge into a single resonance athigher rates. This is but one example of the unique sen-sitivity of NMR parameters to motion; different ways ofexploiting this sensitivity are discussed below.

Magnetization exchange spectroscopyThe magnetization exchange spectroscopy (EXSY) class ofexperiment is used to study slow conformational dynamics,where each probe in each conformer produces a distinctpeak in NMR spectra (Box 1). Initial applications of thisexperiment, developed in the late 1970s, focused on slowdynamics in small molecules [19]. Incorporation of thebasic EXSY sequence into more sophisticated pulseschemes has extended its applicability to larger and morecomplex systems [20–23]. This is demonstrated in a recentapplication to ClpP, an oligomeric protease comprising 14subunits with a total molecular mass of 300 kDa [24]. ClpPis a member of the family of cylindrical, self-compartmen-talizing proteases that includes eukaryotic and archeal20S proteasomes. [25–28]. The active sites of the enzymeare sequestered from the cytosol within the lumen of thecomplex. Proteins targeted for degradation are recognizedby the chaperones ClpA and ClpX, which unfold theirtargets in an ATP-dependent manner and deliver themto the interior of ClpP through axial pores (Box 1, FigureIIa). Although the mechanism by which proteolytic degra-dation products exit the protease had been less well under-stood, important insight was gained via an NMR dynamicsstudy.

07.004 Available online 19 October 2009 601

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Figure 1. Overview of NMR dynamics experiments. The sensitivity of NMR to

molecular motions on a broad range of timescales is well illustrated by the classic

example of two-site chemical exchange [82]. The figure shows simulated one-

dimensional NMR spectra for nuclei exchanging between two distinct chemical

environments, populated in the ratio 3:1 and associated with chemical shifts vA

and vB, respectively (vB � vA = 100 Hz). The indicated times on the right-hand side

correspond to (kAB + kBA)�1. (a) When the rate of exchange is much slower than the

difference in chemical shift (expressed in frequency units), separate peaks are

observed for the two states, and EXSY experiments (Box 1) can be used to

determine the kinetics of exchange. The stochastic variations in chemical shift

produced by exchange can cause the NMR signal to decay more rapidly, leading to

broader, weaker signals. (b) This effect is most pronounced when the exchange

rate is roughly equal to the difference in chemical shifts between exchanging

probes in the two states. Exchange broadening resulting from processes with

(kAB + kBA) < � 2 kHz can be analyzed by CPMG experiments (purple oval, left-hand

Review Trends in Biochemical Sciences Vol.34 No.12

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Using a strategy of perdeuteration, selective 1H/13Cisoleucine d-methyl isotopic labeling, and methyl trans-verse relaxation optimized spectroscopy (TROSY) exper-iments [29], Sprangers et al. obtained high-quality ClpPspectra even at 0.5 8C, where the rotational correlationtime of the molecule is estimated to be greater than 0.4 ms(by comparison, proteins with correlation times on theorder of 30 ns or less are now routinely studied using‘conventional’ NMR spectroscopy). The spectrum wasassigned using site-directed mutagenesis, and it wasobserved that the d-methyl groups of Ile149 and Ile151were each associated with two peaks, one broad and onenarrow, designated F and S for fast and slow relaxing,respectively (Box 1, Figure IIb). Using a methyl TROSY-based EXSY pulse sequence, Sprangers et al establishedthat the ‘doubled’ peaks are due to conformationalexchange between the F and S states (exchange rate of�40 s�1 at 0.58C). Forward and reverse rate constantsextracted from fits of the intensities of the auto- andcross-peaks are the same for the two residues, suggestingthat they sense the same protein motions. TROSY-basedCarr–Purcell–Meiboom–Gill (CPMG) measurements (Box2) [30] are in excellent agreement with the EXSY data,providing validation of themethodology. Ile149 and Ile151,which are present in each of the 14 ClpP monomers, arelocalized in a band around the circumference of the pro-tease at the interface between the heptameric rings (Box 1,Figure IIa). Dynamics in this region could open transientpores and allow product release. This hypothesis wastested by the introduction of a cysteine residue at position153 through site-directed mutagenesis. The cysteinesinserted in this manner can be oxidized to form disulfidebridges between adjacent monomers in the complex, whichare likely to quench the exchange process observed byNMR. A series of biochemical experiments showed thatpeptides can exit rapidly from the reduced form of theenzyme, but are retained within the cavity of the disulfide-linked, oxidized form of the enzyme for >10 h, stronglysuggesting that the protein motions measured by NMR arerelated to clearance of proteolysis products fromClpP. Thisstudy illustrates the potential of combining NMR withbiochemical approaches to produce detailed pictures offunctional dynamics in large oligomeric assemblies.

side; Box 2), whereas processes up to approximately 20 kHz can be studied by R1r

relaxation dispersion methods (purple dots, left-hand side), yielding the kinetic

parameters of interconversion [82]. (c) When exchange is far more rapid than the

difference in chemical shift, a single sharp peak is obtained at the population-

weighted average chemical shift. Under these conditions, the decay of the NMR

signal is dictated primarily by the rotational diffusion of the molecule in solution

and nano- to picosecond timescale internal motions. Analyses of rates at which the

NMR signal returns to its equilibrium value after perturbation (NMR spin

relaxation) provide a measure of the amplitudes and frequencies of fast time-

scale dynamics [83] (Box 4). In addition, structure-based measurements, such as

residual dipolar couplings (RDCs) [2] (Box 3), paramagnetic relaxation

enhancements (PREs) [74] (Box 4), scalar couplings [84], and nuclear Overhauser

enhancements (NOEs) [85] provide data that can be interpreted in terms of

dynamics. These approaches are relatively insensitive to the timescale of

dynamics, requiring only that exchange among conformations is rapid enough

to ensure that experimental measurements are averages across all well-populated

states. Finally, extremely infrequent and transient conformational excursions can

be quantified by measurements of 1H/2H exchange [3]. Thus, NMR can be used to

measure macromolecular dynamics on timescales ranging from picoseconds to

hours or days.

Box 1. Magnetization exchange spectroscopy

The EXSY technique can be applied to molecules in slow exchange

between well-populated conformational states. Here, we consider a

system comprising two interconverting states, A and B. EXSY

experiments are essentially simple variants of multi-dimensional

correlation spectra. For example, in a standard 1H/15N correlation

experiment (directed at 1H/15N pairs such as those of backbone

amide positions), magnetization is transferred from 1H to 15N, the 15N

chemical shift is recorded, the magnetization is returned to 1H, and

the 1H chemical shift is detected. In the corresponding EXSY

experiment, a delay of length T is inserted between the 15N and 1H

chemical shift detection periods (Figure Ia) [22]. When T = 0, two

peaks corresponding to states A and B are obtained (Figure Ib). When

T >0, some of the nuclei initially in state A convert to state B during

the delay, giving rise to a cross-peak at the 15N chemical shift of A and

the 1H chemical shift of B, and vice versa. The intensities of the cross-

peaks (AB and BA) initially increase as a function of T, as the

molecules are given more opportunity to exchange (Figure Ic). For

longer delays, the intensities of all peaks decrease, due to spin

relaxation. A series of NMR spectra are recorded with T varying from

0 to a maximum value that is limited by the longitudinal spin

relaxation rate (Box 4), typically about 1 s. The intensities of both

cross- and auto-peaks are then fitted to extract the forward and

reverse rate constants, as well as the longitudinal spin relaxation

rates [82]. This technique can be applied to systems with rate

constants ranging from about 0.5 s-1 to over 50 s-1, provided that the

peaks are not broadened beyond detection (Figure 1) (Figure II).

Figure I. Exchange spectroscopy (EXSY). (a) A schematic representation of an EXSY NMR pulse scheme [22] for a 1H/15N spin pair exchanging between two states, A and

B. (b) Simulated EXSY spectra for a 1H/15N spin pair undergoing slow two-site exchange. Red and blue peaks correspond to the auto-peaks of states A and B,

respectively, and green cross-peaks are produced by conformational exchange during the delay. (c) Dependence of peak intensities on the delay T.

Figure II. Slow conformational exchange in the protease ClpP. (a) Surface representation of ClpP with two monomers shown as yellow and blue ribbons [86]. Locations

of dynamic isoleucine residues are identified by green and red circles. Substrate entry pores are indicated with blue arrows. (b) The 1H/13C methyl TROSY correlation

spectrum collected for a uniformly [15N, 2H], Ile d1 [13C,1H] labeled ClpP sample. I149 and I151 are each associated with two d1 methyl peaks, designated F and S,

reflecting slow exchange between two distinct, functionally important, conformations. The figure is from Ref. [24]. Copyright (2005) National Academy of Sciences,

U.S.A.


CPMG relaxation dispersionChemical shift fluctuations due to molecular motions on amilli- to microsecond timescale cause peak broadening inNMR spectra and elevate relaxation rates (Figure 1; Box

2). Studies in the 1950s provided the first evidence that thiscontribution to relaxation can be suppressed by applyingRF pulses, and that the effect can be exploited to quantifyconformational exchange processes in small molecules

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Box 2. Carr–Purcell–Meiboom–Gill (CPMG) measurements

The CPMG technique can be applied to systems in which conforma-

tional exchange causes chemical shifts to fluctuate on timescales in

the range 100 s-1 � 2000 s-1 (Box 1). In special cases, the exchange

limits can be expanded to slower time regimes [87], whereas R1r-

based relaxation dispersion methods can be applied to more rapidly

exchanging systems (Figure 1, dotted extensions) [88]. In a typical

series of experiments, variable numbers of refocusing pulses are

applied to magnetization as it evolves under the influence of a

chemical shift that varies stochastically due to the exchange process.

The effect of pulses is illustrated in Figure I, where the signal phase is

Figure I. CPMG relaxation dispersion experiments. (a–c) Simulated signal

trajectories for an ensemble of nuclei exchanging stochastically between two

environments with different chemical shifts during a 20 ms relaxation delay.

Vertical bars at the top of each panel represent refocusing pulses applied

during the relaxation delay. (d) Simulated peak that increases in intensity at

higher pulse repetition rates. (e) Calculated transverse relaxation rates (R2)

plotted as a function of nCPMG = 1/(2t), where t is the delay between successive

refocusing pulses.

Figure II. Millisecond dynamics in DHFR changing in response to substrates

and products. (a) Catalytic cycle of DHFR. (b–f) Locations of exchange

broadening for DHFR in complex with (b) NADPH, (c) NADP+–folate, (d)

NADP+–THF, (e) THF, and (f) NADPH–THF. Green, red, and blue spheres

correspond to 1H nuclei in the NADPH binding site, active site loop, and DHF

binding site, respectively, for which CPMG relaxation dispersion data were fit

to obtain kinetic and chemical shift parameters. Yellow spheres indicate 1H

nuclei with broadened signals for which CPMG traces were of insufficient

quality to extract exchange parameters. (From Ref. [47]. Reprinted with

permission from AAAS).

plotted as a function of time, the slope represents the chemical shift,

and each refocusing pulse reverses the effective direction of signal

evolution. The sets of colored traces correspond to signals from sets

of nuclei undergoing exchange between two different environments

associated with two different chemical shifts. Due to the stochastic

fluctuations in chemical shift (slope), each nucleus follows a slightly

different trajectory, leading to dephasing of the magnetization

(Figure Ia), weak peaks in NMR spectra, and a large transverse

relaxation rate, R2. As the number of refocusing pulses is increased,

the trajectories are prevented from deviating greatly from the initial

position (Figure Ib,c), which suppresses dephasing, increases peak

intensity (Figure Id), and leads to smaller values of R2. CPMG data are

often presented as plots of R2 versus pulse repetition rate, nCPMG,

which is defined as 1/(2t), where t is the time between adjacent

refocusing pulses (Figure Ie). Assuming two-site exchange, the

shapes of dispersion profiles (Figure Ie) are governed by the

populations of the two states, the rate of exchange, and the

difference in chemical shift between the states. Therefore, fits of

CPMG data provide thermodynamic and kinetic information for the

exchange reaction, as well as structural information for the ex-

changing states, even in cases where only the dominant conformer is

‘visible’ in NMR spectra (Figure II).


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[31–33]. Notably, this approach can be used to characterizeconformers that are populated to less than 1% and thatwould otherwise be invisible to conventional NMRmethods. A decade ago, an ingenious modification to thebasic CPMG experiment [34] greatly extended the appli-cability of this technique so that multi-site studies ofbiomolecules are now possible. Since then, relaxation dis-persion experiments have been the focus of intensivemethodological development [35–40], and their use hasbecome widespread in studies of protein and nucleic aciddynamics. Many biological processes, including proteinfolding, binding, and enzyme catalysis, take place on themilli- to microsecond timescale; therefore, they are ideallysuited to characterization by relaxation dispersion tech-niques [18,41–46].

The great potential of thismethodologywas realized in alandmark study of the enzyme dihydrofolate reductase(DHFR) by Wright and co-workers [47]. DHFR uses thecofactor NAPH to reduce dihydrofolate (DHF) to tetrahy-drofolate (THF) via a catalytic cycle that involves fivemajor intermediate forms (Box 2, Figure IIa) [48]. NMRspectra of the enzyme–substrate and enzyme–product com-plexes exhibit line-broadening characteristic ofmillisecondtimescale motions. Amide 15N and 1H CPMG dynamicsdata weremeasured for each of the enzyme complexes (Box2, Figure II). In all cases, the data agreed with a simpleanalytical model in which a low-energy ground stateexchanges with a high-energy excited state. Differencesin 15N and 1H chemical shifts between the ground andexcited states were extracted from the fits, providing spec-troscopic fingerprints of the various excited states. A com-parison of CPMG-derived chemical shift differences andpeak positions indicated that the conformation of eachexcited state resembles the ground state structure of theenzyme complex that immediately precedes and/or followsin the catalytic cycle. These results implicate proteinmotions as being critical to the mechanism of enzymefunction, because the enzyme transiently samples confor-mations similar to the following ground state at manystages in the catalytic cycle, This conclusion is supportedby the kinetic parameters extracted from the CPMG fits.The THF dissociation rate matches the rate at which theTHF-bound protein (E–NADPH–THF) transientlyaccesses an excited state resembling the dissociated form(E–NADPH). The rate of hydride transfer matches the rateat which an excited state resembling the Michaelis com-plex (E–NADPH–DHF) returns to the product-bound (E–

NADP+–THF) ground state. This strongly suggests thatprotein conformational transitions are rate limiting atthese stages of the reaction. In the emerging view ofcatalysis, the energy landscape of protein conformationsis modulated by interactions with substrates and productssuch that the enzyme is ‘funneled through its catalyticallycompetent conformations along a kinetically preferredpath’ [47].

Relaxation dispersionmeasurements are most powerfulwhen the structures of excited states can be established. Inthe previous example, this was made possible by the factthat the chemical shifts of the excited states correspond tothose of the preceding and following ground states, whosestructures have been solved by X-ray crystallography.

However, not all excited states resemble stable forms thatare amenable to standard structure determinationmethods. Vallurupalli and co-workers have developednew experiments that permit the de novo structuralcharacterization of conformational states that are popu-lated to only a small percentage. Utilizing samples dis-solved in liquid crystalline alignment media andspecialized CPMG experiments, they quantified a varietyof different magnetic interactions [36,49] that can be usedto characterize weakly populated (�5%) protein states(Box 3). These measurements provide information on theorientation of inter-nuclear vectors and elements of localstructure within the excited state. Employing the peptide-binding reaction of a Src homology 3 (SH3) domain as amodel system, they demonstrated that these alignmentdata, togetherwith backbone chemical shifts also extractedfrom CPMG experiments, allow accurate, high-resolutionstructures to be determined for weakly populated proteinstates that are invisible using conventional techniques ofstructural biology [50].

Dynamical analyses of residual dipolar couplingsResidual dipolar couplings (RDCs) contain a wealth ofstructural information and they are sensitive to dynamicsranging from the pico- and nano- to the millisecond time-scales (Box 3). Although they are routinely used in thedetermination of static macromolecular structures [51], itremains quite challenging to interpret RDCs in terms ofmolecular motions [2]. RDCs derive from magnetic inter-actions between pairs of proximal NMR-active spins(nuclei); the magnitude of a given RDC is governed bythe orientation of the vector connecting the pair of inter-acting nuclei in a macromolecular frame as well as thealignment of the molecule as a whole (Box 3). Wheninternal dynamics cause the nuclei to reorient, the exper-imentally observed dipolar coupling is the population-weighted average value over all vector orientations. Con-sequently, both structural and dynamic information for alllow-energy macromolecular conformations is encodedwithin an RDC dataset. Extracting dynamical informationfrom RDCs is difficult, as the structure, internal motions,and overall molecular alignment must be fit to the datasimultaneously. The challenge is typically addressed bycollecting RDC data inmultiple different alignmentmedia,with the assumption that the structure and dynamics ofthe macromolecule are identical in each case. RDC data-sets can be fit with explicit geometric models describinginternal motions [52,53]. Alternatively, ensembles of mol-ecular structures can be generated, such that the averagesof dipolar couplings across all members of the ensemblematch the experimental values [54–57].

In a detailed study of the small protein ubiquitin, DeG-root and co-workers assembled an extensive dataset com-prising amide 1H/15N, 1H/13C0, 15N/13C0 and methyl 1H/13CRDCs obtained for 36, 6, 6, and 11 different alignmentconditions, respectively [56]. They generated an ensembleof protein structures that collectively satisfied the exper-imental RDC values (Box 3, Figure II), providing a detailedglimpse into the motions of the protein free in solution.Ubiquitin is a key player in many cellular processes andforms tight, specific interactions with a large number of

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Box 3. Residual dipolar couplings

Each NMR-active nucleus produces a weak magnetic field that interacts

with the magnetic dipoles of nearby nuclei (Figure Ia). The sign and

strength of a dipolar coupling interaction depends on the relative

orientation of the twonuclei. In this example, themagnetic field linesdue

to nucleus (i) point upwards at (ii) and downwards at (iii), leading to

dipolar interaction energies that are of opposite sign for the (i,ii) and (i,iii)

pairs. The strengths of dipolar couplings carry structural information

owing to their orientation dependence. However, all orientations of

inter-nuclearvectorsare equally likely inan isotropic solution.Rotational

diffusion of the molecules rapidly averages the dipolar couplings to zero

and the structural information is lost. This information can be recovered

if macromolecules are dissolved in liquid crystalline media (Figure Ib)

[51], comprising, for example, an ordered suspension of rod-like

particles, such as bacteriophage [89]. Under these conditions, macro-

molecules become weakly aligned, magnetic dipolar interactions no

longer average to exactly zero, and residual dipolar coupling constants

(RDCs) can be measured between proximal pairs of nuclei. RDC values

depend on both the orientation of inter-nuclear vectors within a

molecular reference frame and the overall alignment of the macro-

molecule itself, which varies depending on the type of alignment

medium employed (Figure Ic). A unique feature of RDC data is that inter-

nuclear vectors are defined relative to a common molecular reference

frame, thereby providing global information on the relative orientations

of distal portions of the molecule. Similar information can be gained

from an analysis of peak positions in weakly aligned samples, as the

chemical shifts of many nuclei are orientation dependent (Figure II)

Figure I. Residual dipolar couplings (RDCs). (a) Magnetic field lines emanating from a magnetic dipole at position (i) oriented parallel with the static magnetic field of the

NMR spectrometer (B0). A nucleus at position (ii) senses a local field parallel with B0, whereas a nucleus at (iii) senses a local field anti-parallel to B0. (b) Schematic

representation of protein molecules (dark red) dissolved in a liquid crystalline medium (vertical cylinders). (c) The alignment of an inter-nuclear vector (dotted line)

depends on its orientation within the molecular frame of reference (x, y, z) and the alignment of the entire protein within the laboratory frame of reference (x0, y0, z0).

Figure II. Structural ensemble of ubiquitin calculated from residual dipolar couplings. A total of 40 structures randomly selected from a 116 member ensemble are

shown. Bonds are colored according to the magnitude of motions occurring on a timescale slower than molecular tumbling, with blue, green, yellow, orange, and red

indicating increasing flexibility. (From Ref. [56]. Reprinted with permission from AAAS.)


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different proteins [58–60]. The structures of many ubiqui-tin complexes have been solved by NMR and X-ray crystal-lography, with the conformations adopted by ubiquitinvarying considerably among the complexes. These findingsraise the question of how structural changes might influ-ence the affinity and specificity of binding. Insight into therelationship between binding and flexibility was obtainedthrough a comparison of the RDC ensemble and 46 X-raycrystal structures of ubiquitin complexes. The analysisshowed a high degree of overlap between the RDC (free)and X-ray (bound) ensembles, implying that the differentstructures adopted by ubiquitin in the complexes reflectthe intrinsic motions of the free protein. In other words,binding occurs via conformational selection and involveslow-energy configurations of ubiquitin that are populatedeven in the absence of binding partners. A principle com-ponent analysis of the RDC ensemble further establishedthat the range of ubiquitin structures found in the com-plexes can be accounted for by a small number of collectivemotions in the free protein. This finding suggests that thereduction in protein conformational entropy upon complexformation is not large, even though the magnitude ofmotions in the free protein is substantial. The authorsconclude that ubiquitin has ‘evolved to be as rigid aspossible while remaining as flexible as necessary to engagein different interfaces’ [56] Notably, the motions respon-sible for conformational selection are too rapid to bedetected by relaxation dispersion techniques (Box 2) andtoo slow to be characterized by spin relaxation (Box 4).Thus, dynamical analysis of RDCs promises to open awindow on functional dynamics that have been hiddenfrom view.

Spin relaxationThe rate at which nuclear spin magnetization relaxes toequilibrium after being perturbed by RF pulses is governedby the amplitudes and frequencies of bond vector reorien-tation. Experimentally determined spin relaxation rateshave been used to characterize rapid internal dynamics ofmacromolecules since the 1970s [10]. In the most com-monly applied approach, referred to as model-free [61,62],bond vector motions are separated into contributions frominternal dynamics that occur on the picosecond timescaleand the overall rotational diffusion of the macromolecule,which is typically in the nanosecond regime (Box 4). Stu-dies focusing on picosecond internal dynamics have pro-vided many insights into protein and nucleic acid function[63–65], whereas reports quantitatively linking spin relax-ation parameters to conformational entropy promise toimprove our understanding of biothermodynamics [15–

17,66–69].When macromolecules comprise multiple rigid domains

separated by flexible linkers, the bending and twisting ofthe domains with respect to each other can also contributeto spin relaxation. However, for many of the biomoleculesthat are currently studied by NMR, inter-domain motionsgenerally occur on the same timescale as rotational diffu-sion and it is difficult to separate the two contributions[70]. Al-Hashimi and co-workers have developed a simpleand powerful solution to this problem [71]. Applying theirmethod to the transactivation responsive (TAR) regulatory

RNA from the HIV-1, they demonstrate that this moleculeundergoes functional inter-domain dynamics on a nanose-cond timescale.

The TAR RNA studied by NMR comprises two A-formhelices, termed domains I and II, which are separated by aflexible three nucleotide bulge (Box 4, Figure IIa). The TARsequence is present in viral mRNAs and helps to regulatetranscription by interacting with the viral protein Tat [72]Structures of TAR, both free in solution and in complexwith Tat-derived peptides and a variety of small molecules,show that the bulge region can act as a pivot point andallow large-amplitude bending and twisting of the twohelical domains with respect to each other. This findingraises the question of whether the structural heterogeneityreflects intrinsic motion of the RNA, or whether TAR has asingle preferred conformation free in solution that isdeformed through interactions with ligands.

The amplitude and timescale of inter-helical motionwasquantified by studying a TAR construct in which domain Iwas extended by 22 basepairs (Box 4, Figure IIa). Thissimple modification makes the extended domain I helixmuch larger than the domain II stem–loop, and has twoimportant consequences: Firstly, rotational diffusion of theentire TAR molecule is dominated by the hydrodynamicproperties of the extended helix, and can be treated inde-pendently of dynamics at the pivot point. Secondly, diffu-sive motions of the small domain II stem–loop at the pivotpoint occur on a much faster timescale than overall tum-bling, allowing them to be detected more easily. Using thisapproach, they found that the distribution of fast-timescaledynamics present in the free RNA correlates strongly withthe range of conformations observed in TAR complexes(Box 4, Figure IIb). Analysis of 15N spin relaxation dataindicated that the overall tumbling of the molecule occurswith a time constant of about 20 ns, whereas bending at thepivot point occurs tenfold more rapidly, closely matchingthe diffusion limit. Binding of a small molecule ligand toTAR entirely quenches the bending motions. Thus, TARfreely samples a wide range of conformations in solutionthat allow it to structurally adapt to a variety of bindingpartners. Notably, the helix-extension technique was usedin a dynamical analysis of RDCs, yielding a more precisepicture of the conformations sampled in solution [73] andfurther establishing that the free RNA undergoes motionsthat carry it through a range of structures that resemblevarious ligand-bound states. In this regard, the functionaldynamics of RNA are similar to those described for ubi-quitin. The RDC/spin-relaxation studies of TAR RNA areamong the most complete characterizations of macromol-ecular motions to date.

Paramagnetic relaxation enhancementUnpaired electrons can produce dramatic enhancements ofspin relaxation rates for nearby nuclei. Recent work hasestablished that paramagnetic relaxation enhancements(PREs) are exquisitely sensitive to the existence of low-lying excited states, provided that the kinetics of exchangeare rapid and that the unpaired electron and nucleus aresignificantly closer in the weakly populated conformerthan in the ground state (Box 4) [74]. This effect has beenexploited to study weak self-association [75], transient

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chain collapse in unfolded proteins [76], encounter com-plexes in binding reactions [77], and non-specific sliding ofproteins along DNA [78]. In an elegant application of thisapproach, Clore and co-workers have resolved long-stand-ing questions regarding the bacterial maltodextrin sensor,the maltose-binding protein (MBP) [79]. MBP comprisestwo domains separated by a short, three-stranded linker.According to X-ray crystal structures of the free and boundforms of the protein, the interaction withmaltose induces alarge rigid-body domain reorientation that closes the bind-ing cleft around the ligand (Box 4, Figure III). Small-angle

Box 4. Spin relaxation experiments

At equilibrium, the nuclear magnetization of an NMR sample is

aligned with the static magnetic field of the NMR spectrometer.

Following perturbation by radiofrequency pulses, magnetization

returns to equilibrium. The rate at which this occurs is governed by

molecular dynamics that cause the energies of nuclear interactions to

fluctuate on a nano- to picosecond timescale. Thus, information on

fast-timescale motions can be extracted from measurements of

nuclear spin relaxation rates. Common experiments focus on the R1

and R2 rate constants, which describe the build-up of longitudinal

(parallel with the static magnetic field) and the decay of transverse

(perpendicular to the field) magnetization, respectively, as the system

returns to equilibrium (Figure Ia). Typically, a series of two-dimen-

sional correlation spectra is collected with a variable relaxation delay,

and peak intensities are fit to extract the relaxation rate of interest

(Figure Ib,c). In the case of protein amide and nucleic acid imino15N/1H pairs, 15N relaxation rates are governed almost entirely by

motions of the NH bond vector, which modulate both the 15N

chemical shift and the strength of the 15N/1H dipole-dipole interaction

(Box 3). Analysis of 15N relaxation rates usually separates NH bond-

vector motions into independent contributions from picosecond-

Figure I. Spin relaxation. (a) Return of magnetization to equilibrium following perturb

buildup of longitudinal magnetization parallel with B0 occur with rate constants R2 and

in NMR relaxation experiments. (d) Processes contributing to the dynamic reorientati

rapid association and dissociation with a paramagnetic probe (red circle). Protein ato

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X-ray scattering and NMR RDC [80] data for the free formof the protein largely agree with the ‘‘open’’ X-ray crystalstructure. However, neither of these techniques is sensi-tive to small populations of excited states. The mechanisticrelationship between ligand binding and domain reorien-tation remained unresolved: does maltose bind the openstructure and induce closure, or do transient excursions tothe closed form allow binding? A detailed analysis of PREdata shows that the actual mechanism is likely morecomplicated than either of these scenarios.

timescale internal dynamics and nanosecond-timescale rotational

diffusion of the molecule as a whole (Figure Id) [83].

Nuclear relaxation rates are enhanced enormously by nearby

unpaired electrons, due to dipole–dipole interactions involving the

large electron magnetic moment [74]. Paramagnetic relaxation

enhancements (PREs) decrease as the sixth power of the electron/

nucleus distance; therefore, they are extremely sensitive to

transient close encounters between nuclei and paramagnetic labels.

When exchange is very rapid (greater than several thousand per

second), the experimental PREs are the population-weighted

averages over all conformational sub-states. Thus, even if the

paramagnetic label is in close proximity only a small fraction of the

time, the PREs for this state still make sizeable contributions to

experimental relaxation rates. In the simple scenario shown in

Figure Ie, the PRE measurement for each nucleus is related to its

distance from the unpaired electron (red sphere) in state B, as well

as the relative populations of the two states. In this way, PRE

datasets can provide qualitative thermodynamic information as well

as structural details relating to weakly populated excited states

(Figures II and III).

ation by RF pulses. The decay of magnetization in the transverse (x, y) plane and

R1, respectively. (b,c) Decreasing peak intensities obtained for increasing delays

on of internuclear vectors. (e) Schematic representation of a protein undergoing

ms close to the probe in state B exhibit large PREs, i.e. rapid relaxation.

Figure III. Domain reorientation in maltose-binding protein (MBP). X-ray

crystal structures of the maltose-free (blue, 1OMP [90]) and maltose-bound

(red, 3MBP [91]) forms of MBP, superimposed using the N-terminal domain

(gray) as a reference. Maltose is represented in ball and stick format. Locations

of the engineered cysteine residues used for conjugation of TEMPO spin labels

are indicated with green arrows. (From Ref. [79]. Reprinted by permission from

Macmillan Publishers Ltd.)

Figure II. Functional nano- to picosecond timescale dynamics in the TAR RNA. (a) Sequence and secondary structure of the extended TAR RNA studied by NMR spin

relaxation experiments. In order to eliminate spectral crowding from the Extension, two samples were used in the experiments: one with 13C/15N A,U and unlabeled G,C

where R = G and Y = C, and one with 13C/15N G,C and unlabeled A,U where R = A and Y = U. The naturally occurring six-nucleotide loop in domain II was replaced with a

more stable UUCG tetraloop. (b) Correlation between fast timescale dynamics and heterogeneity in structures of TAR complexes. Higher normalized peak intensities

qualitatively indicate larger amounts of motion on a nano- to picosecond timescale. <Du> is the mean angular deviation in CH bond orientations among eight TAR

structures that have been superimposed using domain I as a reference. Data for domains I, II, and the three nucleotide bulge are colored red, green, and yellow,

respectively. Plotted values are for sugar C10H10 (diamonds) and base C2H2 (squares), C5H5 (circles), C6H6 (triangles), and C8H8 (inverted triangles) bond vectors. (From

Ref. [71]. Reprinted with permission from AAAS.)


Naturally occurring proteins do not contain unpairedelectrons in the absence of metal ligands [81]; therefore,external paramagnetic probes must be employed for PREanalyses. Clore and co-workers used two conjugationschemes in which a stable nitroxide radical (2,2,6,6-tetra-methyl-1-piperidinyloxyl; TEMPO) was covalently linkedto either the N-terminal or C-terminal domain via engin-eered cysteine residues (Box 4, Figure III). In both cases,domain closure substantially shortens the distance be-tween the TEMPO label and the opposite domain, leadingto large PRE increases. PRE data collected for MBP in thepresence of maltose are in excellent agreement with thebound X-ray crystal structure. By contrast, many PREvalues for MBP in the absence of maltose are much largerthan expected. This is not simply due to differences in theground state conformation for the solution and crystalenvironments, as NMR RDC data for the free TEMPO-linked form match the X-ray structure. Therefore, in the

absence of ligands, MBP must experience transientdomain closure that is invisible to population-weightedmeasurements such as RDCs. The PRE value for each1H nucleus in the protein is related to its distance fromthe TEMPO spin label in the transiently closed form, whichprovides detailed structural information on this minorspecies. The analysis reveals that the transiently closedform is populated to a level of about 5% and differs struc-turally from the bound form of MBP in an importantrespect: whereas the binding cleft is occluded in the boundstructure, it is easily accessible to solvent in the transientlyclosed form. Therefore, ligands have easy access to thebinding site of the minor species, which might be poised toreorient into the fully closed conformation, thereby facil-itating maltose binding. The domain motions of free MBPare not detected by relaxation dispersion experiments,indicating that they occur on a timescale faster than about50 ms. They are readily characterized by PREs, illustratingthe power of this method to obtain high-resolution struc-tural information on protein motions that are inaccessibleto other techniques.

Concluding remarksModern NMR spectroscopy offers a rich assortment oftechniques for characterizing biomolecular motions. Thesemethods are helping to fill the gaps between the structuralsnapshots provided by conventional X-ray crystallographicand NMR analyses. It is becoming clear that instead ofadopting a single rigid structure, macromolecules oftenexplore conformational space extensively, allowing themto adapt to different binding partners or to progress effi-ciently through catalytic cycles. Motions can occur over abroad range of timescales and can take many forms, fromdiffusive transitions within a single energy well to acti-vated excursions between ground and excited states. Chan-ging conditions or interactions with ligands can shift thebalance of populations, converting an excited state to aground state and vice versa. This diversity in dynamicsraises many questions. For example, how is flexibilityencoded by the primary amino acid or nucleotide sequence?When do functional motions actively promote a kineticpathway, and when are they a passive consequence ofobligatory flexibility? What is the relationship between

609


dynamics, entropy, and free energy? The ever-increasingability of NMR spectroscopy to observe biological dynamicswith atomic resolution will be central to answering thesequestions and to explaining how living systems function ata fundamental level.

AcknowledgementsA.K.M and L.E.K acknowledge support in the form of grants from theNatural Science and Engineering Research Council of Canada and theCanadian Institutes of Health Research (L.E.K.). L.E.K. holds a CanadaResearch Chair in Biochemistry.

References1 Mittermaier, A. and Kay, L.E. (2006) Review - new tools provide new

insights in NMR studies of protein dynamics. Science 312, 224–2282 Tolman, J.R. and Ruan, K. (2006) NMR residual dipolar couplings as

probes of biomolecular dynamics. Chem. Rev. 106, 1720–17363 Bai, Y.W. (2006) Protein folding pathways studied by pulsed-and

native-state hydrogen exchange. Chem. Rev. 106, 1757–17684 Palmer, A.G. (2004) NMR characterization of the dynamics of

biomacromolecules. Chem. Rev. 104, 3623–36405 Ishima, R. and Torchia, D.A. (2000) Protein dynamics from NMR. Nat.

Struct. Biol. 7, 740–7436 Serber, Z. et al. (2006) Investigating macromolecules inside cultured

and injected cells by in-cell NMR spectroscopy. Nat. Protocols 1, 2701–

27097 Wuthrich, K. and Wagner, G. (1978) Internal Motions in Globular

Proteins. Trends in Biochemical Sciences 3, 227–2308 Snyder, G.H. et al. (1975) Complete tyrosine assignments in high-field

H-1 nuclear magnetic resonance spectrum of bovine pancreatic trypsininhibitor. Biochemistry 14, 3765–3777

9 Campbell, I.D. et al. (1975) Proton magnetic resonance studies oftyrosine residues of hen lysozyme – assignment and etection ofconformational mobility. Proc. Roy. Soc. B 189, 503–509

10 Allerhand, A. et al. (1971) Conformation and segmental motion ofnative and denatured ribonuclease A in solution. Application ofnatural-abundance carbon-13 partially relaxed fourier transformnuclear magnetic resonance. J. Am. Chem. Soc. 93, 544–545

11 Tugarinov, V. et al. (2006) Isotope labeling strategies for the study ofhigh-molecular-weight proteins by solution NMR spectroscopy. Nat.Protocols 1, 749–754

12 Palmer, A.G. et al. (2005) Solution NMR spin relaxation methods forcharacterizing chemical exchange in high-molecular-weight systems.In Nuclear Magnetic Resonance of Biological Macromolecules, Part C,Methods in Enzymology (Vol. 394) (James, T.L., ed.), In pp. 430–465,Academic Press

13 Tugarinov, V. et al. (2004) Nuclear magnetic resonance spectroscopy ofhigh-molecular-weight proteins. Ann. Rev. Biochem. 73, 107–146

14 Zhang, Q. and Al-Hashimi, H.M. (2008) Extending the NMR spatialresolution limit for RNA by motional couplings. Nat. Methods 5, 243–

24515 Frederick, K.K. et al. (2007) Conformational entropy in molecular

recognition by proteins. Nature 448, 325–32916 Trbovic, N. et al. (2009) Protein side-chain dynamics and residual

conformational entropy. J. Am. Chem. Soc. 131, 615–62217 Li, D.W. andBruschweiler, R. (2009) A dictionary for protein side-chain

entropies from NMR order parameters. J. Am. Chem. Soc. 131, 7226–

722718 Sugase, K. et al. (2007) Mechanism of coupled folding and binding of an

intrinsically disordered protein. Nature 447, 1021–102519 Jeener, J. et al. (1979) Investigation of exchange processes by 2-

dimensional NMR spectroscopy. J. Chem. Phys.71 4546–455320 Montelione, G.T. and Wagner, G. (1989) 2D chemical exchange NMR

spectroscopy by proton-detected heteronuclear correlation. J. Am.Chem. Soc. 111, 3096–3098

21 Wider, G. et al. (1991) Studies of slow conformational equilibria inmacromolecules by exchange of heteronuclear longitudinal 2-spin-order in a 2D difference correlation experiment. J. Biomol. NMR 1,93–98

22 Farrow, N.A. et al. (1994) A heteronuclear correlation experiment forsimultaneous determination of 15N longitudinal decay and chemical

610

exchange rates of systems in slow equilibrium. J. Biomol. NMR 4, 727–

73423 Sahu, D. et al. (2007) TROSY-based z-exchange spectroscopy:

application to the determination of the activation energy forintermolecular protein translocation between specific sites ondifferent DNA molecules. J. Am. Chem. Soc. 129, 13232–13237

24 Sprangers, R. et al. (2005) Quantitative NMR spectroscopy ofsupramolecular complexes: dynamic side pores in ClpP areimportant for product release. Proc. Natl Acad. Sci. USA 102,16678–16683

25 Pickart, C.M. and Cohen, R.E. (2004) Proteasomes and their kin:Proteases in the machine age. Nat. Rev. Mol. Cell Biol. 5, 177–187

26 Wickner, S. et al. (1999) Posttranslational quality control: folding,refolding, and degrading proteins. Science 286, 1888–1893

27 Lowe, J. et al. (1995) Crystal structure of the 20S proteasome from thearchaeon T-acidophilum at 3.4 angstrom resolution. Science 268, 533–

53928 Groll, M. et al. (1997) Structure of 20S proteasome from yeast at 2.4

angstrom resolution. Nature 386, 463–47129 Tugarinov, V. et al. (2003) Cross-correlated relaxation enhancedH-1-C-

13 NMR spectroscopy of methyl groups in very high molecular weightproteins and protein complexes. J. Am. Chem. Soc. 125, 10420–10428

30 Korzhnev, D.M. et al. (2004) Probing slow dynamics in high molecularweight proteins by methyl-TROSY NMR spectroscopy: application to a723-residue enzyme. J. Am. Chem. Soc. 126, 3964–3973

31 Carr, H.Y. and Purcell, E.M. (1954) Effects of diffusion on freeprecession in nuclear magnetic resonance experiments. Phys. Rev.94, 630–638

32 Meiboom, S. and Gill, D. (1958) Modified spin-exho method formeasuring nuclear relaxation times. Rev. Sci. Instrum. 29, 688–691

33 Luz, Z. and Meiboom, S. (1963) Nuclear magnetic resonance study ofprotolysis of trimethylammonium ion in aqueous solution - order ofreaction with respect to solvent. J. Chem. Phys. 39, 366–370

34 Loria, P.J. et al. (1999) A relaxation-compensated Carr-Purcell-Meiboom-Gill sequence for characterizing chemical exchange byNMR spectroscopy. J. Am. Chem. Soc. 121, 2331–2332

35 Ishima, R. and Torchia, D.A. (2003) Extending the range of amideproton relaxation dispersion experiments in proteins using a constant-time relaxation-compensated CPMG approach. J. Biomol. NMR 25,243–248

36 Vallurupalli, P. et al. (2007) Measurement of bond vector orientationsin invisible excited states of proteins. Proc. Natl Acad. Sci. USA 104,18473–18477

37 Korzhnev, D.M. et al. (2004) Multiple-quantum relaxation dispersionNMR spectroscopy probing millisecond time-scale dynamics inproteins: theory and application. J. Am. Chem. Soc. 126, 7320–7329

38 Orekhov, V.Y. et al. (2004) Double- and zero-quantum NMR relaxationdispersion experiments sampling millisecond time scale dynamics inproteins. J. Am. Chem. Soc. 126, 1886–1891

39 Long, D. et al. (2008) Accurately probing slow motions on millisecondtimescales with a robust NMR relaxation experiment. J. Am. Chem.Soc. 130, 2432–2433

40 Millet, O. et al. (2000) The static magnetic field dependence of chemicalexchange linebroadening defines the NMR chemcial shift time scale. J.Am. Chem. Soc. 122, 2867–2877

41 Wolf-Watz,M. et al. (2004) Linkage between dynamics and catalysis in athermophilic-mesophilic enzymepair.Nat.Struct.Mol.Biol.11, 945–949

42 Eisenmesser, E.Z. et al. (2005) Intrinsic dynamics of an enzymeunderlies catalysis. Nature 438, 117–121

43 Li, P.L. et al. (2008) Internal dynamics control activation and activity ofthe autoinhibited Vav DH domain. Nat. Struct. Mol. Biol. 15, 613–618

44 Korzhnev, D.M. et al. (2004) Low-populated folding intermediates ofFyn SH3 characterized by relaxation dispersion NMR. Nature 430,586–590

45 Demers, J.P. andMittermaier, A. (2009) Bindingmechanism of an SH3domain studied by NMR and ITC. J. Am. Chem. Soc. 131, 4355–4367

46 Watt, E.D. et al. (2007) The mechanism of rate-limiting motions inenzyme function. Proc. Natl Acad. Sci. USA 104, 11981–11986

47 Boehr, D.D. et al. (2006) The dynamic energy landscape ofdihydrofolate reductase catalysis. Science 313, 1638–1642

48 Schnell, J.R. et al. (2004) Structure, dynamics, and catalytic function ofdihydrofolate reductase. Ann. Rev. Biophys. and Biomol. Struct. 33,119–140


49 Vallurupalli, P. et al. (2008) Probing structure in invisible proteinstates with anisotropic NMR chemical shifts. J. Am. Chem. Soc.130, 2734–2735

50 Vallurupalli, P. et al. (2008) Structures of invisible, excited proteinstates by relaxation dispersion NMR spectroscopy. Proc. Natl Acad.Sci. USA 105, 11766–11771

51 Bax, A. and Grishaev, A. (2005) Weak alignment NMR: a hawk-eyedview of biomolecular structure. Curr. Opin. Struct. Biol. 15, 563–570

52 Bouvignies, G. et al. (2005) Identification of slow correlated motions inproteins using residual dipolar and hydrogen-bond scalar couplings.Proc. Natl Acad. Sci. USA 102, 13885–13890

53 Bouvignies, G. et al. (2008) Characterization of protein dynamics fromresidual dipolar couplings using the three dimensional Gaussian axialfluctuation model. Proteins: Struct. Funct. Bioinformatics 71, 353–363

54 Jensen, M.R. et al. (2008) Quantitative conformational analysis ofpartially folded proteins from residual dipolar couplings: applicationto the molecular recognition element of Sendai virus nucleoprotein. J.Am. Chem. Soc. 130, 8055–8061

55 Clore, G.M. and Schwieters, C.D. (2006) Concordance of residualdipolar couplings, backbone order parameters and crystallographicB-factors for a small alpha/beta protein: a unified picture of highprobability, fast atomic motions in proteins. J. Mol. Biol. 355, 879–886

56 Lange, O.F. et al. (2008) Recognition dynamics up to microsecondsrevealed from an RDC-derived ubiquitin ensemble in solution. Science320, 1471–1475

57 De Simone, A. et al. (2009) Toward an accurate determination of freeenergy landscapes in solution states of proteins. J. Am. Chem. Soc. 131,3810–3811

58 Hicke, L. et al. (2005) Ubiquitin-binding domains. Nat. Rev. Mol. CellBiol. 6, 610–621

59 Brzovic, P.S. and Klevit, R.E. (2006) Ubiquitin transfer from the E2perspective - why is UbcH5 so promiscuous? Cell Cycle 5, 2867–2873

60 Harper, J.W. and Schulman, B.A. (2006) Structural complexity inubiquitin recognition. Cell 124, 1133–1136

61 Lipari, G. and Szabo, A. (1982) Model-free approach to theinterpretation of nuclear magnetic relaxation in macromolecules: 2.Analysis of experimental results. J. Am. Chem. Soc. 104, 4559–4570

62 Lipari, G. and Szabo, A. (1982) Model-free approach to theinterpretation of nuclear magnetic relaxation in macromolecules: 1.Theory and range of validity. J. Am. Chem. Soc. 104, 4546–4559

63 Henzler-Wildman, K.A. et al. (2007) A hierarchy of timescales inprotein dynamics is linked to enzyme catalysis. Nature 450, 913–916

64 Shajani, Z. et al. (2007) Binding of U1A protein changes RNA dynamicsas observed by C-13 NMR relaxation studies. Biochemistry 46, 5875–

588365 Igumenova, T.I. et al. (2006) Characterization of the fast dynamics of

protein amino acid side chains using NMR relaxation in solution.Chem. Rev. 106, 1672–1699

66 Lee, A.L. et al. (2000) Redistribution and loss of side chain entropyupon formation of a calmodulin-peptide complex. Nat. Struct. Biol. 7,72–77

67 Yang, D. and Kay, L.E. (1996) Contributions to conformational entropyarising from bond vector fluctuations measured from NMR-derivedorder parameters: application to protein folding. J. Mol. Biol. 263,369–382

68 Akke, M. et al. (1993) NMR order parameters and free energy: ananalytic approach and application to cooperative calcium binding bycalbindin D9k. J. Am. Chem. Soc. 115, 9832–9833

69 Li, Z. et al. (1996) Insights into the local residual entropy of proteinsprovided by NMR relaxation. Protein Sci. 5, 2647–2650

70 Chen, K. and Tjandra, N. (2008) Extended model free approach toanalyze correlation functions of multidomain proteins in the presenceof motional coupling. J. Am. Chem. Soc. 130, 12745–12751

71 Zhang, Q. et al. (2006) Resolving the motional modes that code for RNAadaptation. Science 311, 653–656

72 Jones, K.A. and Peterlin, B.M. (1994) Control of RNA initiation andelongation at the HIV-1 promoter. Ann. Rev. Biochem. 63, 717–743

73 Zhang, Q. et al. (2007) Visualizing spatially correlated dynamics thatdirects RNA conformational transitions. Nature 450, 1263–1267

74 Clore, G.M. et al. (2007) Elucidating transient macromolecularinteractions using paramagnetic relaxation enhancement. Curr.Opin. Struct. Biol. 17, 603–616

75 Tang, C. et al. (2008) Visualization of transient ultra-weak protein self-association in solution using paramagnetic relaxation enhancement. J.Am. Chem. Soc. 130, 4048–4056

76 Felitsky, D.J. et al. (2008) Modeling transient collapsed states of anunfolded protein to provide insights into early folding events. Proc.Natl Acad. Sci. U. S. A. 105, 6278–6283

77 Iwahara, J. and Clore, G.M. (2006) Detecting transient intermediatesin macromolecular binding by paramagnetic NMR. Nature 440, 1227–

123078 Iwahara, J. et al. (2006) NMR structural and kinetic characterization of

a homeodomain diffusing and hopping on nonspecific DNA. Proc. NatlAcad. Sci. U. S. A. 103, 15062–15067

79 Tang, C. et al. (2007) Open-to-closed transition in apo maltose-bindingprotein observed by paramagnetic NMR. Nature 449, 1078–1082

80 Evenas, J. et al. (2001) Ligand-induced structural changes tomaltodextrin-binding protein as studied by solution NMRspectroscopy. J. Mol. Biol. 309, 961–974

81 Bertini, I. et al. (2001) Paramagnetic probes in metalloproteins. Nucl.Magn. Reson. Biol. Macromol. B 339, 314–340

82 Palmer, A.G. et al. (2001) Nuclear magnetic resonance methods forquantifying microsecond-to-millisecond motions in biologicalmacromolecules. Nucl. Magn. Reson. Biol. Macromol. B 339, 204–

23883 Jarymowycz, V.A. and Stone, M.J. (2006) Fast time scale dynamics of

protein backbones: NMR relaxation methods, applications, andfunctional consequences. Chem. Rev. 106, 1624–1671

84 Lindorff-Larsen, K. et al. (2005) Interpreting dynamically-averagedscalar couplings in proteins. J. Biomol. NMR 32, 273–280

85 Lindorff-Larsen, K. et al. (2005) Simultaneous determination of proteinstructure and dynamics. Nature 433, 128–132

86 Wang, J.M. et al. (1997) The structure of ClpP at 2.3 angstromresolution suggests a model for ATP-dependent proteolysis. Cell 91,447–456

87 Tollinger, M. et al. (2001) Slow dynamics in folded and unfolded statesof an SH3 domain. J. Am. Chem. Soc. 123, 11341–11352

88 Ishima, R. and Torchia, D.A. (1999) Estimating the time scale ofchemical exchange of proteins from measurements of transverserelaxation rates in solution. J. Biomol. NMR 14, 369–372

89 Hansen, M.R. et al. (1998) Tunable alignment of macromolecules byfilamentous phage yields dipolar coupling interactions. Nat. Struct.Biol. 5, 1065–1074

90 Sharff, A.J. et al. (1992) Crystallographic evidence of a large ligand-induced hinge-twist motion between the 2 domains of the maltodextrinbinding protein involved in active transport and chemotaxis.Biochemistry 31, 10657–10663

91 Quiocho, F.A. et al. (1997) Extensive features of tight oligosaccharidebinding revealed in high-resolution structures of the maltodextrintransport chemosensory receptor. Structure 5, 997–1015

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Observing biological dynamics at atomic resolution using NMR