NMR supplement

nature structural biology • NMR supplement • july 1998 517

The traditional NMR approach to structuredetermination of biomolecules is based oninterpretation of rates of magnetizationtransfer between pairs of protons in termsof distance constraints. The approachrequires not only the measurement of mag-netization transfer (NOEs or nuclearOverhauser effects), but the resolution andassignment of NMR signals to specific pro-tons, of specific residues, in a known pro-tein sequence. Assignment of resonancesand measurement of adequate numbers ofNOEs have always been obstacles that madestructure determination time consumingand limited to relatively small proteins(<10,000 Mr for early homonuclear stud-ies). The limitations have been pushed backover the years with additional structuralinformation from scalar coupling constantsand chemical shifts. Assignment strategiesbased on the use of through bond connec-tivites between 13C and 15N sites in isotopi-

cally enriched proteins have also made itpossible to assign resonances in increasinglylarger proteins. However, full structuredeterminatons have still remained confinedto reasonably compact systems of molecularweights less than 30,000–40,000 Mr

1,2.There have been, within the last two

years, experiments reported that could dra-matically change the range of applicabilityof NMR structural methods. Interestingly,they share an origin in anisotropic magneticinteractions that are not normally observ-able in high resolution NMR spectra. Oneimportant class of experiments yields struc-tural constraints that are orientational,rather than distance based. The experi-ments rely on the measurement of residualdipolar couplings, and, in some cases,chemical shift anisotropy (CSA)3–5. Themeasurements can be made with great effi-ciency, and when combined with otherrecent discoveries that take advantage of

interference between the same dipole-dipole and CSA interactions6,7,8, it appearsthat NMR may be poised to take anotherlarge step forward in applicability to larger,more complex systems.

Residual dipolar interactionsThe dipole-dipole interaction, the lead-ing term of which is described in equa-tion (1), is actually the basis of the NOEeffect:

Dij = -ξ ij

(3 cos2 θ - 1) IziIzj (1)2

The interaction constant, ξij, contains fac-tors that describe the magnitudes of mag-netic moments for a pair of nuclei i and j,and the internuclear distance depen-dence that shows up in NOE measure-ments. The spin operator, IziIzj, has thesame form as a first order through-bondspin-spin coupling interaction suggest-

New techniques in structural NMR —anisotropic interactionsJ.H. Prestegard

Structure determination of biomolecules by NMR has traditionally been based on nuclear Overhauser effects(NOEs). Now there are additional sources of information that can complement NOEs in cases where positioningof remote parts of molecules is important, and where extension to larger and more complex systems is desired.

. 1. Palmer, A. Curr. Opin. Biotech. 4, 385–391 (1993).2. Kay, L.E., Torchia, D.A. & Bax, A. Biochemistry 28,

8972–8979 (1989).3. Pascal, S.M., Yamazaki, T., Singer, A.U., Kay, L.E. &

Forman-Kay, J.D. Biochemistry 34, 11353–11362(1995).

4. Tjandra, N., Feller, S.E., Pastor, R.W. & Bax, A. J. Am.Chem. Soc. 117, 12562–12566 (1995).

5. Bruschweiler, R., Liao, X. & Wright, P.E. Science268, 886–889 (1995).

6. Zheng, Z., Czaplicki, J. & Jardetzky, O. Biochemistry15, 5212–5223 (1995).

7. Lee, L.K., Rance, M., Chazin, W.J. & Palmer, A.G. J.Biomol. NMR 9, 287–298 (1997).

8. Lipari, G. & Szabo, A. J. Am. Chem. Soc. 104,4546–4559 (1982).

9. Yamazaki, T., Muhandiram, R. & Kay, L.E. J. Am.Chem. Soc. 116, 8266–8278 (1994).

10. Engelke, J. & Ruterjans, H. J. Biomol. NMR 5,173–182 (1995).

11. Cordier, F., Brutscher, B. & Marion, D. J. Biomol.NMR 7, 163–168 (1996).

12. Dayie, K.T. & Wagner, G. J. Am. Chem. Soc. 119,7797–7806 (1997).

13. Fischer, M.W.F. et al. J. Am. Chem. Soc. 119,12629–12642 (1997).

14. Zeng, L., Fischer, M.W.F. & Zuiderweg, E.R.P. J.Biomol. NMR 7, 157–162 (1996).

15. LeMaster, D.M. & Kushlan, D.M. J. Am. Chem. Soc.118, 9255–9264 (1996).

16. Muhandiram, D.R., Yamazaki, T., Sykes, B.D. & Kay,L.E. J. Am. Chem. Soc. 117, 11536–11544 (1995).

17. Yang, D., Mittermaier, A., Mok, Y.K. & Kay, L.E. J.Mol. Biol. 276, 939–954 (1998).

18. Werbelow, L.G. & Grant, D.M. Adv. Magn. Reson. 9,189–299 (1977).

19. Vold, R.L. & Vold, R.R. Progress in NMRSpectroscopy 12, 79–133 (1978).

20. Daragan, V.A. & Mayo, K.H. Biochemistry 32,11488–11499 (1993).

21. Daragan, V.A. & Mayo, K.H. J. Magn. Reson. Ser. B107, 274–278 (1995).

22. Akke, M. & Palmer, A.G. J. Am. Chem. Soc. 118,911–912 (1996).

23. Akke, M., Liu, J., Cavanagh, J., Erickson, H.P. &Palmer, A.G. Nature Struct. Biol. 5, 55–59 (1998).

24. Denisov, V.P. & Hale, B. J. Mol. Biol. 245, 682–697(1995).

25. Denisov, V.P., Peters, J., Horlein, H.D. & Halle, B.Nature Struct. Biol. 3, 505–509 (1996).

26. Tolman, J.R., Flanagan, J.M., Kennedy, M.A. &Prestegard, J.H. Nature Struct. Biol. 4, 292–297 (1997).

27. Steinbach, P.J. et al. Biochemistry 30, 3988–4001(1991).

28. Feher, V.A., Baldwin, E.P. & Dahlquist, F.W. NatureStruct. Biol. 3, 516–521 (1996).

29. Beeser, S.A., Goldenberg, D.P. & Oas, T.G. J. Mol.Biol. 269, 154–164 (1997).

30. Housset, D., Kim, K.S., Fuchs, J., Woodward, C. &Wlodawer, A. J. Mol. Biol. 220, 757–770 (1991).

31. Nicholson, L.K. et al. Nature Struct. Biol. 2, 274–280(1995).

32. Wlodawer, A. & Erickson, J.W. A. Rev. Biochem. 62,543–585 (1993).

33. Sommerville, R. Prog Nucleic Acid Res 42, 1–38(1992).

34. Gryk, M.R., Jardetzky, O., Klig, L.S. & Yanofsky, C.Protein Science 5, 1195–1197 (1996).

35. Ogata, K. et al. Nature Struct. Biol. 3, 178–187 (1996).36. Ogata, K. et al. Nature. Struct. Biol. 2, 309–320

(1995).37. Pawson, T. Nature, 573–580 (1995).38. Kuriyan, J. & Cowburn, D. Curr. Opin. Str. Biol. 3,

828–837 (1993).39. Pascal, S.M., et al. Cell 77, 461–472 (1994).40. Lee, C.H., et al. Structure 2, 423–438 (1994).41. Kay, L.E., Muhandiram, D.R., Wolf, G., Shoelson, S.E.

& Forman-Kay, J.D. Nature Struct. Biol. 5, 156–163(1998).

42. Akke, M., Bruschweiler, R. & Palmer, A. J. Am.Chem. Soc. 115, 9832–9833 (1993).

43. Yang, D. & Kay, L.E. J. Mol. Biol. 263, 369–382(1996).

44. Philippopoulos, M. & Lim, C. J. Mol. Biol. 254,771–792 (1995).

45. Farrow, N.A., Zhang, O., Forman-Kay, J.D. & Kay,L.E. Biochemistry 34, 868–878 (1995).

46. Li, Z., Raychaudhuri, S. & Wand, A.J. Prot. Sci. 5,2647–2650 (1996).

47. Wand, A.J., Urbauer, J.L., McEvoy, R.P. & Bieber, R.J.Biochemistry 35, 6116–6125 (1996).

48. Mandel, A.M., Akke, M. & Palmer, A.G.Biochemistry 35, 16009–16023 (1996).

49. Yang, D., Mok, Y.-K., Forman-Kay, J.D., Farrow, N.A.& Kay, L.E. J. Mol. Biol. 272, 790–804 (1997).

50. Alexandrescu, A.T. et al. Prot. Sci. in the press(1998).

51. Stivers, J.T., Abeygunawardana, C., Mildvan, A.S. &Whitman, C.P. Biochemistry 35, 16036–16047(1996).

52. Gagne, S.M., Tsuda, S., Spyracopoulos, L., Kay, L.E. &Sykes, B.D. J. Mol. Biol. in the press (1998).

NMR supplement

518 nature structural biology • NMR supplement • july 1998

ing that the dipole interaction can inprinciple add to the multiplet splittingsnormally seen in high resolution NMRspectra. If it were this simple, measure-ment would be extraordinarily easy.

The feature of most interest for our dis-cussion is the dependence on the anglebetween the vector connecting the inter-acting nuclei and the applied magnetic

field of the spectrometer, θ. As can beimagined from a look at Fig. 1, knowing θfor a bonded pair of nuclei, such as a back-bone amide 15N and 1H in a protein, couldbe very useful in defining a molecularstructure. The bar over the θ dependentterm, however, denotes a time average.Normally in solution we assume that thetime average results from a moleculartumbling that uniformly samples direc-tions in space (isotropic sampling). The(3cos2 θ − 1) term in equation (1) thenaverages to zero, and no static contribu-tions to spin interaction energies remain.This is why we are often relegated to mea-suring the dipole-dipole interaction indi-rectly through spin relaxation basedphenomena such as the NOE.

Sampling of orientational space, how-ever, need not be isotropic. If moleculeshave preferred orientations in the pres-ence of a magnetic field, a non-uniformBoltzmann distribution will result, andthere will be secular contributions. Thesewill appear as new multiplet splittings ofresonances in the case of non-bondednuclei, or as changes in splittings of reso-nances in the case of bonded nuclei9.

The utility of residual dipolar couplingsin molecular structure determination ofsoluble molecules was anticipated yearsago in works by Bothner-By andMcLean10. The reason for preferred mole-cular orientations in these early works wasthe interaction between the applied mag-netic field and an orientation dependent

induced magnetic moment in the mole-cule under study. However, departuresfrom isotropic averaging in solution, dueto these interactions are normally small(less than a part in a thousand), and mea-surement with precision was difficult.

About two years ago an example ofmeasurement at a useful level of precisionfor a protein in solution appeared3. Therewere several reasons for reemergence ofthis measurement. One was simply that aparamagnetic protein with a particularlyhigh magnetic susceptibility anisotropywas used (cyanometmyoglobin). A secondwas that the 15N dimension in the het-eronuclear single quantum coherence(HSQC) experiment used on the 15Nenriched protein offered particularly highresolution. And a third was that new, veryhigh field magnets, became available (ori-entation due to field induced interactionsrises approximately as the square of themagnetic field).

The utility of residual dipolar couplingsin structural analysis had also been recog-nized earlier in the liquid crystal commu-nity11. Liquid crystals orient in thepresence of a magnetic field for the samereasons isolated magnetically anisotropicmolecules do, but here molecules formlarge cooperative domains. The net orien-tational effect is large, and residual dipolarcouplings are large. In fact, couplings areusually so large that spectra suffer fromsecond order effects that make analysis ofspectra from even small molecules dis-solved in liquid crystals difficult.

A few years ago applications of liquidcrystal NMR technology reemerged in thestructural biology community, particular-ly the segment involved in membrane pro-tein structure determination. Fragmentsof lipid bilayers, a well accepted buildingblock of biological membranes, werefound to form liquid crystal arrays whenprepared at 20–30 weight % lipid to aque-ous buffer using mixtures of long chainphospholipids such as dimyristoylphos-phatidylcholine (DMPC) and lipids withdetergent-like properties such as dihexa-noylphosphatidylcholine (DHPC)12. Thebilayer fragments appear to be small dis-coidal particles a few hundred Å in diame-ter, and have become known as ‘bicelles’13.At 30–40 ºC they orient with the normalsof the bilayer surfaces perpendicular to themagnetic field. Proteins and peptides thatassociate with lipid bilayers attach to theoriented discoidal surfaces and themselvesbecome oriented. Applications share someof the difficulties associated with strongdipolar couplings in other liquid crystalmedia. However, use of isotopically

Fig. 2 Induced pro-tein orientation bydilute phospholipidbicelles. The proteintumbles rapidly, butanisotropically, inlarge aqueous inter-bicelle spaces.

Fig. 1 Dipolar coupled 15N-1H spin pair in anamide bond. The bond length, r, is assumedfixed and the primary variable is the angle, θ,between the magnetic field, B0, and the inter-nuclear vector.

NMR supplement

nature structural biology • NMR supplement • july 1998 519

labeled peptides, (15N, 13C), again allowsavoidance of some of these problems14,and the physical properties of bicelles con-tinue to be improved15. The general area ofmembrane protein structure determina-tion by solid state NMR on samplesincluding bicelles was highlighted in thelast supplement series16.

An important event for the present dis-cussion is that the areas of field inducedorientation of single molecules in solutionand liquid crystal induced orientation ofbicelle associated molecules have mergedto provide a new and very promising routeto utilization of residual dipolar cou-plings. As noted above, one area suffersfrom splittings that are too small and theother from splittings that are too large. Itmay seem obvious that a suitable compro-mise should exist, but one cannot easilyincrease further the alignment of isolatedoriented molecules, and one cannot ingeneral decrease domain sizes or degreesof order in liquid crystals without produc-ing heterogeneous mixtures of differentphases. It turns out that at least at certaintemperatures, and with certain lipid mix-tures, one can dilute the bicelle medium

used for membrane protein studies by afactor of five or six to produce a coopera-tively oriented homogeneous mediumwith aqueous spaces between bicelles largeenough to accommodate soluble proteins(Fig. 2)4.

Proteins in aqueous spaces in the dilutebicelle preparations adopt some of theorder of the surrounding bicelles, and thelevel of order can be ‘tuned’ to yield a veryfavorable ±10 Hz residual dipolar splittingby adjusting the concentration of thebicelles in the mixture. At this point anunderstanding of the mechanism ofordering may not be complete, but in theinitial application the level of order is con-sistent with a simple weak collisionalmodel that depends only on the physicalbarriers provided by the bicelles and onthe non-spherical shape of the dissolvedproteins. If this mechanism holds, therewill be minimal effects on molecular tum-bling rates, and hence, line widths. Moreimportantly, structures determined willbe truly representative of molecules inbulk aqueous solution. Reports currentlyemerging include applications to not onlyproteins, but nucleic acids and carbohy-

drates. It is likely that any variation inmechanistic understanding will have aminor effect on applications to structuredetermination.

An example of the type of data attain-able in a bicelle medium is shown in Fig. 3.This is an HSQC spectrum of a twodomain fragment of a carbohydrate bind-ing protein from barley17, that has beenordered in a 5% 3:1 DMPC/DHPC bicelledispersion. Decoupling of protons during15N evolution has been omitted from thenormal HSQC sequence so that a doubletappears for each correlated amide pro-ton–amide nitrogen pair. A region of thespectrum that shows two doublets thatincrease and decrease splittings respec-tively, upon orientation is shown. Thebicelle preparations have a convenientproperty in that an isotropic phase con-taining the same lipids, presumably inmore symmetric micelles, is obtained bylowering the temperature slightly. Spectraunder these conditions (left of Fig. 3)show only scalar through-bond couplingsof 93 ±2 Hz. Comparing isotropic and ori-ented spectra, Lys 53 shows a negative 5Hz residual dipolar contribution and Lys95 shows a positive 2 Hz contribution.These are indicative of an average N-Hvector orientation perpendicular and par-allel to the magnetic field respectively — auseful structural constraint.

In the above illustration it is clear thatmeasurements can be made directly fromthe frequency domain spectrum. Thesemeasurements are straightforward and veryefficient compared to NOE measurements.

Fig. 4 Chemical shift anisotropy of an amide car-bonyl carbon. Chemical shift tensor elements, δ11,δ22, and δ33, are taken to be 223, 79 and 55 p.p.m.respectively. Shifts observed in oriented samplesare functions of the angle between the magneticfield, B0, and the principle shift tensor axes.

Fig. 3 Segments from a proton coupled, nitrogen decoupled, 15N-1H HSQC spectrum of a 0.4 mMsolution of a barley lectin fragment in a 5% DMPC/DHPC 3:1 bicelle (doped with a positivelycharged amphiphile). Left is an isotropic spectrum at 25 oC, right is an oriented spectrum at 35 oC.Both increases and decreases in couplings are observed.

NMR supplement

520 nature structural biology • NMR supplement • july 1998

Spectra such as the one shown can beacquired in as little as one hour on 1 mMprotein samples. There are also a number ofoptions for making these measurementsmore precise18–20. In addition, measure-ments are not confined to 15N-1H pairs, butextend to 13C-1H pairs21–22 and even non-bonded 1H-1H pairs4.

Chemical shift anisotropyResidual dipolar coupling is not the onlyanisotropic spin interaction that can pro-vide useful structural information.Chemical shifts are also anisotropic. Nucleiwhich are part of various molecu-lar functional groups resonate atdifferent frequencies depending onshielding by the local electronicenvironment. Electronic environ-ments are seldom isotropic, andhence, shielding is different for dif-ferent orientations of functionalgroups. For the case of a moleculewith an axially symmetric chemi-cal shift tensor, the contribution tospin energy levels, which leads toan offset in resonance positionfrom that seen with isotropic aver-aging, is given in equation (2):

Ci = ∆δ (3 cos2 θ - 1) γiB0Izi (2)2

The coefficient ∆δ is the differ-ence in chemical shift in direc-tions parallel and perpendicularto the symmetry axis, and γiB0Izi

is the Zeeman interaction opera-tor. It is significant that an angu-lar dependence analogous to thatseen for the residual dipole inter-action occurs.

An illustration of anisotropicchemical shift effects is given forthe 13C of the carbonyl group ofthe peptide in Fig. 4; if the pep-tide preferred an orientationwith the field perpendicular tothe plane, the carbonyl carbon would res-onate at a higher field (or lower frequen-cy) than in a case where isotropicsampling of orientations occurred.Measured chemical shift offsets are clear-ly of structural value.

Chemical shift offsets are easily mea-sured in the well oriented cases repre-sented by membrane associated peptides,and both 15N and 13C chemical shift off-sets have been used to help deduce struc-tural properties12,15,23. In weakly orientedsolution systems, and in dilute bicellesystems, measurements are more diffi-cult. Small changes in resonance positiondue to temperature and other factors

must be considered. Nevertheless, usefulmeasurements have been made even invery weakly oriented solution samples5.

Structure determinationHow can structure be determined from allthis new orientational information?Published examples utilizing orientationalinformation from simple solution anddilute bicelle studies have been confinedto refinement of existing protein struc-tures. It is possible to add orientationalpenalty functions to force fields used insimulated annealing protocols, and in this

way combine orientational constraintswith the distance constraints used in con-ventional NMR structure determinations.A good example is the work on a solutionof a DNA binding domain from the tran-scription factor GATA-1 bound to a 16base pair double helical DNA24. A conven-tional NMR structure determination hadpreviously been completed. Adding orien-tation constraints improved consistencywith accepted values of backbone torsionangles in several cases and did significant-ly change the structure of a loop that hadbeen poorly defined by the distance con-straint data. It was also pointed out that itwas possible to deduce the orientation of

the DNA grove binding α-helix from thesmall negative values of the couplings forN–H bonds in this protein segment.

Currently there are few examples ofdetermining structures using orienta-tional data from solution or bicelle stud-ies as a primary structure determinant.There are some fundamental problems inthat error functions used with orienta-tional data are more complex than thoseused with distance data because theinverse of the basic angular function ofequations (1) and (2) is multivalued.However, there are viable alternative

approaches that can be gleanedfrom work occurring on moreordered membrane samples12,23.One can in principle determine,within a degeneracy of two, theorientation of any rigid substruc-ture of a molecule from five inde-pendent orientationalmeasurements. Such a substruc-ture can be a peptide bond if 15N-1H, 13C-1H, and 13C-13C dipolarvectors are combined with chem-ical shift anisotropy effects.Then, peptide bonds can be ori-ented one at a time. Such a proce-dure has recently been used tosuggest a structure for a mem-brane bound myristoylated pep-tide from the ARF-1 protein12.

Substructures can also be sin-gle domains within multidomainproteins. If the structures of indi-vidual domains are known fromconventional NMR or X-raystudies, the entire domain can betreated as a rigid substructure.Even if only N-H dipolar interac-tion vectors are used, there isplenty of information to deter-mine orientation relative to aprinciple orientational frame.

For a well structured molecule,domains should see a common

principle orientational frame. Thisrequirement is illustrated in Fig. 5 for atwo domain transcription factor boundto a DNA double helix (taken from the X-ray structure of Fairall et al.25). If thedomains were positioned properly, theprinciple orientational frames, deter-mined independently from orientationalNMR data in solution, would differ onlyby translation. If they were not, domainscould be reoriented to achieve a commonframe. The positioning of the domains isentirely independent of how far apartthey may be, thus illustrating the fact thatresidual dipolar data and CSA data can beideal complements to short range dis-

Fig. 5 Properly oriented domains in a two zinc-finger–DNA com-plex have common orientations of independently determined prin-ciple orientation tensors. The structure shown is modeled on acrystal structure of the TramTrack–DNA complex by Fairall et al.25

NMR supplement

nature structural biology • NMR supplement • july 1998 521

tance constraints from NOEs. A combi-nation of NOE and residual dipolar datahas recently been used to determine therelative orientation of two zinc fingerdomains from the transcription factorSp1 as they bind to a 14 base pair DNAdouble helix (V.A. Narayan & J.P.Caradonna, pers. comm.).

Larger molecules and the futureWhy is there so much excitement aboutthe prospects for using residual dipolarcoupling and other data from anisotropicinteractions to determine structures ofbiomolecules? Some certainly arises fromthe improvements in precision that mightarise in refinement of existing NMR struc-tures24. But, we believe that more will arisefrom the unique contributions that can bemade in cases where NOEs are hard toobtain. The example involving position-ing of remote domains in DNA bindingtranscription factors is a good illustrationof what we mean. But, another ariseswhen biomolecules are simply large.

In conventional NMR structure deter-minations of proteins, side chain–sidechain NOEs play a particularly importantrole in determining a three dimensionalfold. Frequently only side chain protonsof sequentially remote residues comeclose enough to one another to yield anNOE. Yet, assignments of side chain pro-ton resonances, or those of their attachedcarbons, are hard to establish becausethey require multiple transfers throughshort lived 13C magnetization to get fromthe more easily assigned peptide back-bone resonances to those of the side chainextremities.

Assignment of backbone resonancescan often be accomplished with muchlarger proteins than assignment of sidechain resonances. In fact, resolution andassignment of backbone amide protonand nitrogen resonances extends to par-ticularly large systems when perdeutera-tion of non-exchangeable protons isemployed26. While relatively few NOEsare observed that connect backboneamide protons to sequentially remoteparts, observed short and intermediaterange NOEs do define secondary struc-ture elements. Under these circumstancesthe orientational constraints for N-Hvectors in secondary structure elementsmay be used to supplant some sidechain–side chain NOEs.

Interference ofanisotropic interactionsThere are some reasons to believe thatfurther improvement in an ability to

resolve and assign amide nitrogen andproton resonances are at hand6. In fact,with suitable advances in magnetic fieldstrengths it may be possible to considerNMR based structure determinations ofproteins several times the size of thosestudied today. One reason is an expecteddramatic narrowing of resonances innewly devised experiments. The expectedadvance is actually based on interferenceof the same two interactions that we havebeen discussing above, the dipole-dipoleinteraction and the dipole-CSA interac-tion.

In addition to shifts in resonance posi-tion and changes in doublet splitting,dipole-dipole and CSA-dipole interac-tions contribute to line broadening. Thefact that the two contributions interfereis well known and can be seen in the sim-ple proton coupled 15N-1H cross peaks ofFig. 4. The cross peaks belonging to a 1Hcoupled doublet are not of the samewidth. The downfield doublet compo-nent is actually narrower; in fact, it isnarrower than the single cross peak thatappears when protons are decoupled andthe two components collapse to a singlepeak. Differences are small here becausethey are obscured by a relatively shortacquisition time and unresolved longrange couplings to protons that could bereplaced with deuterons. Nevertheless,the differences are real.

The reason that differential line broad-ening occurs is that the nitrogen spinrelaxes in response to the square of thetotal fluctuation in the local magneticfield as opposed to the sum of the squaresof the individual fluctuations27. Whenlocal field fluctuations come from twosources such as the magnetic dipole ofthe attached proton and the chemicalshift anisotropy of the nitrogen, it is thevector sum that counts. For the two linesof a nitrogen doublet, the magneticdipoles of the protons project in oppositedirections. So, the dipolar and CSA fieldsadd for one line and subtract for theother. Hence, one line relaxes efficientlyand the other less efficiently.

At fields corresponding to 500 MHz forprotons the CSA contribution in anamide 15N-1H pair is smaller than thedipolar contribution, but large enough toproduce lines of observably differentwidth. However, the CSA field is itselffield dependent. At higher fields the sizesof the CSA and dipolar terms becomemore comparable and eventually becomeequal. If the CSA tensor were axially sym-metric and colinear with the dipolar vec-tor, exact cancellation would occur at alittle over 1 GHz, and the line width ofthe narrower line would be determinedpredominantly by remote spin interac-tions. The line widths of the two lines as afunction of spectrometer operating fre-

Fig. 6 Line widths for the two lines of a proton coupled 15N doublet as a function of spectrometeroperating frequency. A correlation time of 20 ns and a nitrogen CSA of -160 p.p.m. have beenassumed. Note the minimum line width predicted at a frequency slightly above 1 GHz.

NMR supplement

522 nature structural biology • NMR supplement • july 1998

quency, or equivalently magnetic field,are plotted in Fig. 6. Clearly there wouldbe tremendous advantage in observingonly the narrow peaks in coupled HSCQspectra.

Wüthrich and coworkers have devisedpulse sequences for the selective acquisi-tion of the narrower peaks in two dimen-sional HSQC spectra and illustrated thefirst applications of this new methodolo-gy6. Improvement in resolution (andimprovement in sensitivity that can comewith it) is key to doing NMR based struc-ture determination of larger molecules.Increasing the molecular weight of a pro-tein by a factor of two approximatelydoubles the number of peaks, and dou-bles the line width of each peak.Decreasing the line width by a factor oftwo to four would largely compensate forthe degradation in resolution and sensi-tivity on doubling the size of a protein.Coupling these improved HSQC experi-ments with some of the assignment andNOE experiments traditionally used inprotein structure determination will cer-tainly be possible. It may also be possibleto couple them with experiments used todetermine residual dipolar couplings andCSA offsets.

We should mention that the above isnot the only case where interferencebetween dipole-dipole and CSA interac-tions promises to improve structure deter-mination of biomolecules in solution.There is an inherent geometry depen-dence of this interference. In the aboveexample, it is important that the approxi-mate symmetry axis of the nitrogen CSAtensor is nearly parallel to the 15N-1Hdipolar vector. When it is not, effects vary.Some authors have used these effects toadvantage in determining torsion anglesalong protein backbones7,8. The informa-tion returned is similar to that obtainedthrough three bond scalar coupling, butapplications to larger molecules will bepossible. Other authors have used inter-ference in spin relaxation to get extrainformation needed to deal withanisotropic motion in proteins28.

A caveat about motionAll of the above discussion about structurepresumes an ability to separate motionaleffects from structural effects. In manycases, analysis of observables is predicatedon the assumption of a rigid structuralmodel. Proteins and other biomoleculesare not rigid. We can, to a certain extent,ignore this fact in NOE based determina-tions because models that include rapiduncorrelated centrosymmetric motionsare degenerate with rigid models. This isnot the case with residual dipolar andother anisotropic data. Motions almostalways reduce the magnitude of theireffects. Moreover, all motions from thoseon picosecond time scales to those on mil-lisecond time scales can contribute. Inearly work on residual dipolar couplingsin a paramagnetic form of myoglobin,deviations from rigid model predictionswere observed and found consistent withaveraging by internal motions29. At thecurrent time it appears that the effects ofthese motions on structures determinedcan be minimized by using a combinationof spin relaxation and amide exchangerate experiments to identify residueswhich may be most susceptible to motion-al averaging and eliminate data from theseresidues when attempting to producestructural models. Nevertheless, theremay be reason for caution in interpreta-tion, particularly for applications thatrefine to high resolution. There are ofcourse good as well as bad aspects to theexistence of motional effects. A new probeof motional properties that is sensitive to abroad range of motions is interesting initself. These more interesting aspects ofmotional effects are discussed in anotherarticle in this supplement30.

AcknowledgmentsSupport from the NIH and NSF is gratefullyacknowledged. We also thank J. Losonczi, A. Fowler,M. Fischer and H. Al-Hashimi for their help inpreparing the figures.

J.H. Prestegard is at the ComplexCarbohydrate Research Center, University

of Georgia, Athens, Georgia 30602-4712,USA.

Correspondence should be addressed toJ.H.P. email: jpresteg@ccrc.uga.edu

1. Clore, G.M. & Gronenborn, A.M., Nature Struct.Biol. 4, 849–853 (1997).

2. Wagner, G., Nature Struct. Biol. 4, 841–844 (1997).3. Tolman, J.R., Flanagan, J.M., Kennedy, M.A., &

Prestegard, J.H. Proc. Natl. Acad. Sci. USA 92,9279–9283 (1995).

4. Tjandra, N. & Bax, A. Science 278, 1111–1114(1997).

5. Ottiger, M., Tjandra, N. & Bax, A. J. Am. Chem. Soc.119, 9825–9830 (1997).

6. Pervushin, K., Riek, R., Wider, G. & Wüthrich, K.Proc. Natl. Acad. Sci. USA 94, 12366–12371 (1997).

7. Reif, B., Hennig, M. & Griesinger, C. Science, 276,1230–1233 (1997).

8. Brutscher, B., Skrynnikov, N.R., Bremi, T.,Brueschweiler, R. & Ernst, R.R. J. Magn. Reson.130, 346–351 (1998).

9. Prestegard, J.H., Tolman, J.R., Al-Hashimi, H.M. &Andrec, M. In Modern techniques in protein NMR(N.R. Krishna & L.J. Berliner, eds) (Plenum, NewYork, 1998) in the press.

10. Bastiaan, E.W., Maclean, C., Van Zijl, P.C.M. &Bothner-By, A.A. Ann. Rep. NMR Spect. 19, 35–77(1987).

11. Emsley, J.N. & Lindon, J.C. (Pergamon Press,Oxford, 1975).

12. Sanders II, C.R., Hare, B.J. Howard, K.P. &Prestegard, J.H. Prog. Nucl. Magn. Resonan. 26,421–444 (1994).

13. Sanders II, C.R. & Landis, G.C. Biochemistry, 34,4030–4040 (1995).

14. Losonczi, J.A. & Prestegard, J.H. Biochemistry 37,706–716 (1998).

15. Prosser, R.S.,Hunt, S.A, DiNatale, J.A. & Vold, R.R. J.Am. Chem. Soc. 118, 269–270 (1996).

16. Opella, S. J. Nature Struct. Biol. 4, 845–848 (1997).17. Weaver, J.L. & Prestegard, J.H. Biochemistry. 37,

116–128 (1998).18. Ottiger, M., Delaglio, F. & Bax, A. J. Magn. Reson.

131, 373–378 (1998).19. Tjandra, N., Grzesiek, S. & Bax, A. J. Am. Chem.

Soc. 118, 6264–6272 (1996).20. Tolman, J.R. & Prestegard, J.H. J. Magn. Reson.

112 245–252 (1996).21. Bax, A. & Tjandra, N. J. Biomol. NMR. 10, 289–292

(1997).22. Yang, D., Tolman, J.R., Goto, N.K. & Kay, L.E. J.

Biolmol. NMR 11, (1998) in the press.23. Kovacs, F.A. & Cross, T.A. Biophys. J. 73, 2511–2517

(1997).24. Tjandra, N., Omichinski, J.G., Gronenborn, A.M.,

Clore, G.M. & Bax, A. Nature Struct. Biol. 4,732–738 (1997).

25. Fairall, L., Schwabe, J.W., Chapman, L., Finch, J.T. &Rhodes, D. Nature. 366, 483–487 (1993).

26. Venters, R.A., Metzler, W.J., Spicer, L.D., Mueller,L., & Farmer III, B.T. J. Am. Chem. Soc. 117,9592–9593 (1995).

27. Werbelow, L.G. & Grant, D.M. Adv. Magn. Reson.9, 189–299 (1977).

28. Fischer, M.W., et. al. J. Am. Chem. Soc. 119,12629–12642 (1997).

29. Tolman, J.R., Flanagan, J.M., Kennedy M. A. &Prestegard, J.H. Nature Struct. Biol. 4, 292–296(1997).

30. Kay, L.E. Nature Struct. Biol. 5, 514–518 (1998).