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munc-13 and the mint family2. For syn-taxin, critical experiments will focus onunderstanding factors that regulate inter-conversion between closed and openstates of the protein and especially thosethat might work by controlling thearrangement of the helical linker domain.More important than ever will be effortsto monitor these protein conformationsand interactions in functional mem-branes and living cells.

Phyllis I. Hanson is in the Department of CellBiology & Physiology, Washington UniversitySchool of Medicine, 660 S. Euclid, CampusBox 8228, St. Louis, Missouri 63110, USA.email: phanson@cellbio.wustl.edu

1. Rothman, J.E. & Wieland, F.T. Science 272, 227–234(1996).

2. Jahn, R. & Südhof, T.C. Annu. Rev. Biochem. 68,863–911 (1999).

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4. Weber, T. et al. Cell 92, 759–772 (1998).5. Misura, K.M.S., Scheller, R.H. & Weis, W.I. Nature

404, 355–362 (2000).6. Dulubova, I. et al. EMBO J. 18, 4372–4382 (1999).7. Fiebig, K.M., Rice, L.M., Pollock, E. & Brünger, A.T.

Nat. Struct. Biol. 6, 117–123 (1999).8. Fernandez, I. et al. Cell 94, 841–849 (1998).9. Calakos, N., Bennett, M.K., Peterson, K.E. & Scheller,

R.H. Science 263, 1146–1149 (1994).10. Nicholson, K.L. et al. Nat. Struct. Biol. 5, 793–802

(1998)11. Söllner, T., Bennett, M.K., Whiteheart, S.W., Scheller,

R.H. & Rothman, J.E. Cell 75, 409–418 (1993).12. Schekman, R. Curr. Opin. Cell Biol. 4, 587–592 (1992).13. Verhage, M. et al. Science 287, 864–869 (2000).14. Pevsner, J. et al. Neuron 13, 353–361 (1994).15. Carr, C.M., Grote, E., Munson, M., Hughson, F.M. &

Novick, P.J. J. Cell Biol. 146, 333–344 (1999).

Theoretical and experimental work inthe areas of protein folding1 and DNAcondensation2 predict the rapid collapseof extended biopolymers into compactconformations, but little is known abouthow this process might contribute to thefolding of RNA. On pages 362 and 367 ofthis issue of Nature Structural Biology,Buchmueller et al.3 and Russell et al.4

provide the first evidence that group Iribozymes form compact states that aredisordered. Consistent with theoreticalpredictions, they show that collapse ofthe RNA occurs early in the foldingprocess, under conditions in which fewmolecules have reached the native struc-ture. Understanding this collapsed statepromises to reveal much about thenucleation of tertiary structure.

The problem of nucleation and col-lapse has been investigated much moreextensively in proteins than in nucleicacids. Evidence for disordered compactstates come from the molten globuleintermediates of protein folding anddenatured states of proteins such asstaphylcoccal nuclease5,6. The condensa-tion of peptide chains is primarily drivenby hydrophobic interactions, while thecondensation of nucleic acids requiresneutralization of negatively chargedphosphates by cations7.

In RNA, stable tertiary structure isonly observed in the presence of divalentmetal ions or, in some cases, very highconcentrations of monovalent ions7.Both the kinetics and specificity of fold-

ing is expected to be extremely sensitiveto ionic conditions — not only becausebinding of Mg2+ is required to stabilizethe native structure, but also becauseelectrostatic interactions are expected todominate the free energy of nucleationand collapse.

Theoretical modelsTheoretical studies of minimal peptidemodels have provided estimates for thetimescales of folding that agree well withexperimental results1,8,9. These modelsare also useful for predicting the foldingkinetics of RNA10, although the actualtimescales for folding of proteins andRNA may differ. For some small pro-teins, such as chymotrypsin inhibitor 2,specific nucleation of native interactionsleads directly to the native state, aprocess which occurs on timescales of10–100 µs (for a review, see ref. 11) (Fig.1). This is roughly comparable to thetime required to form local secondarystructures in RNA12,13. RNA tertiaryinteractions form more slowly, on theorder of 10–100 ms for tRNA14,15 and thecatalytic domain of the ribozyme from B.subtilis RNase P16.

In general, the initial collapse of thepolypeptide or polynucleotide chain isbelieved to be nonspecific, yielding a col-lection of compact structures that diffu-sively search out low free energyconformations8 (Fig. 1). Many of thesecorrespond to non-native structures thatarise when local energetically favorable

interactions compete successfully withcontacts that define the native topologyof the protein or RNA10. Although theinitial collapse and conformationalsearch in proteins occur on timescales of10–1,000 ms (depending on the length ofthe chain and rate of diffusion)8, thetransition from stable intermediates tothe native structure is slow, because themolecule must at least partially unfold todisrupt the non-native structures17. It isimportant to note that mechanismsinvolving specific and nonspecific col-lapse may occur simultaneously, withsome molecules folding rapidly to thenative structure while others becometrapped in metastable intermediatestates (Fig. 1).

Compact intermediates RNA folding pathways are often domi-nated by partially folded intermediatesthat persist for long times in vitro18. Inthe Tetrahymena ribozyme, the domaincontaining helices P4–P6 folds with atime constant of 1 s, after the formationof tertiary structure is initiated byadding Mg2+ to the RNA19 (Fig. 2).Native interactions within the core of theribozyme form much more slowly (τ ∼ 1 min; refs 19,20), and even longertimes (τ ∼ 10 min) are required to reachthe active state under some condi-tions21,22. Native gel electrophoresisshowed that the ribozyme is distributedamong a collection of intermediateswithin the first 15–30 s after the addition

Compact but disordered states of RNASarah A. Woodson

Large ribozymes collapse to compact structures that are non-native and partially disordered when folding isinduced by magnesium ions. This suggests that the early steps in the folding of these RNAs are non-specific.

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350 nature structural biology • volume 7 number 5 • may 2000

of Mg2+ (ref. 23). Many of these migratednearly as fast as the folded RNA, suggest-ing that they are only slightly moreextended than the native ribozyme.

Statistical mechanical models for fold-ing of macromolecules (Fig. 1) predictthat compact structures will form veryearly in the folding process10. In contrast,sequential assembly of the native statearound stable substructures (such as theP4–P6 domain) predicts that a portion ofthe RNA will remain extended until rela-tively late in the folding process13,20.

Russell et al.4 tested these predictionsby measuring changes in the averageradius of gyration (Rg) of the Tetra-hymena ribozyme using small angle X-ray scattering (SAXS). These experimentstook advantage of the sensitivity andtime resolution offered by bright syn-

chrotron X-ray sources24. As expected,the radius of gyration of the unfoldedRNA (74 Å) was much larger than that ofnative RNA in 10 mM MgCl2 (47 Å)4.More excitingly, they found that theradius of gyration decreased to 51 Åwithin 1 min after adding Mg2+, the deadtime of the experiment (Fig. 2). This is~20-fold faster than formation of thenative structure, and is fully consistentwith rapid collapse to a collection ofcompact, non-native structures4.

Collapsed stateUsing a completely different experimen-tal approach, Buchmueller et al.3 showedthat a protein-dependent ribozymeforms compact structures that appear toentirely lack stable tertiary contacts. ThebI5 group I ribozyme is derived from a

yeast mitochondrial intron that requiresa protein cofactor, CBP2, for splicing invivo25. Binding of CBP2 stabilizes the ter-tiary structure of the riboyzme andrestores catalytic activity under physio-logical conditions26. The bI5 catalyticcore requires 40–50 mM MgCl2 to fold inthe absence of protein (ref. 27). The rateof Mg2+-induced folding is independentof protein concentration, suggesting thatthe protein preferentially recognizes thefolded ribozyme core27 (Fig. 3).

Buchmueller et al.3 used gel filtrationchromatography and native gel elec-trophoresis to measure the hydrodynam-ic radius of the bI5 ribozyme core atvarious Mg2+ concentrations3. As expect-ed, the RNA appeared more compactwith higher amounts of MgCl2, com-pared to a double-stranded RNA control(Fig. 3). Surprisingly, the midpoint ofthe collapse transition (3 mM MgCl2)was much lower than the amount ofMg2+ required to observe tertiary inter-actions in hydroxyl radical cleavageexperiments (Fig. 3)27. Thus, nearly all ofthe RNA is in compact structures underconditions (3–7 mM MgCl2) in whichonly a small fraction of the RNA isnative. By comparison with gel filtrationchromatography standards, the authorsestimated the Stokes radii of the unfold-ed, intermediate and native forms to be57 Å, 48 Å and 44 Å, respectively3.

Native-like intermediates Both groups found that the intermedi-ates are only 10% larger, on average, thanthe native structure. This implies thatthe RNA already has a large amount ofnative structure. In the Tetrahymenaribozyme, there is good evidence fromhydroxyl radical protection experimentsthat many of the native tertiary interac-

Fig. 1 Hypothetical energy folding landscape of RNA. The graph schematically indicates free ener-gy (G, vertical axis) as a function of conformation (horizontal axis). A subset of the extended mol-ecules undergoes specific nucleation and collapse to the native structure (N). The remainingpopulation becomes trapped in a collection of compact, metastable intermediates (I) that corre-spond to local minima in the rough energy landscape. The intermediates contain many native andsome non-native interactions. Transitions from I to N cross significant energy barriers and occurclose to the native structure.

Fig. 2 Folding pathway of the Tetrahymena ribozyme. Progressive formation of tertiary interactions was determined by time-resolved hydroxyl rad-ical footprinting19 and other methods20,36. Radii of gyration (Rg) of different species determined by Russell et al.4 are indicated. The unfolded andintermediate states correspond to multiple conformations. Transitions from misfolded intermediates to the native structure involve at least partialunfolding of the RNA. Green, P4-P6 domain; blue, P3-P9 domain; yellow, non-native helix in the ribozyme core; pink and gray, peripheral helicesP2/P2.1 and P0.1/P9.2. Cartoon of the native structure is based on ref. 28.

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tions are formed within the first 10 s ofthe folding reaction19, including interac-tions with peripheral helices that wraparound the exterior of the core28. Theselong-range contacts are thought to stabi-lize the intermediates.

An important question is how native-like intermediates reach the native state.One possibility is that non-native con-tacts in the ribozyme core can reorganizewithin the confines of a compact struc-ture (Fig. 2). A more likely possibility isthat the intermediates transiently unfoldbefore reaching the native state.Mutations that disrupt peripheral inter-actions increase the rate of folding23,29.This is consistent with the significantactivation barrier for refolding21,30.

In contrast, the collapsed state of bI5RNA contains no detectable tertiarystructure, suggesting that this phase isnonspecific and consists of many confor-mational states3. Buchmueller et al.3

found that disruption of tertiary contactsthat are essential for maintaining thenative fold by site-directed mutagenesishad little effect on the formation of com-pact structures at low Mg2+ concentra-tions3. Although it is hard to imagine howcompact structures would form withoutlong range interactions, bI5 RNA mayfluctuate among marginally stable struc-tures in moderate MgCl2 concentrations.If this is the case, the activation barrier forforming the native structure should below, and the kinetics of RNA foldingshould not be affected by changes in reac-tion temperature. In agreement with thisexpectation, Buchmueller et al.3 foundthat the bI5 folding rate was independentof temperature in 7 mM MgCl2

3. Becausenative bI5 RNA is rapidly bound by CBP2in these experiments, the folding rate in

these experiments was determined bymeasuring the accumulation of catalyti-cally active CBP2–bI5 complex.

Folding of RNA–protein complexesThe differences in the folding intermedi-ates of the Tetrahymena and bI5ribozymes provide a new perspective onthe assembly of RNA–protein complexes.While the intermediates of theTetrahymena ribozyme contain a largepercentage of native structure, the con-formations of the bI5 intermediates arepoorly defined. Self-splicing group Iintrons are invariably stabilized by long-range contacts outside of the conservedcore, although the exact topology ofthese interactions differs among intronsubfamilies28,31. Although these interac-tions greatly stabilize the native struc-ture, they also increase the folding timeby stabilizing non-native structures32. Insome introns, such as yeast bI5, theseRNA–RNA interactions are replaced byRNA-binding proteins27,33. The results ofBuchmueller et al.3 suggest that tertiaryinteractions in bI5 remain relatively fluidin the absence of CBP.

Protein-dependent splicing mecha-nisms are widely considered to haveevolved from autocatalytic RNAs(reviewed in ref. 34). In light of this, onemay ask whether the assembly ofRNA–protein complexes offers anyadvantages over the folding of largeribozymes. Because the probability oflarge RNAs collapsing specifically toform the native state is extremely small,RNA–protein interactions that providegreater opportunities for specific con-tacts13,35 could facilitate the formation ofnative structure. Furthermore, the over-all rate of folding will be greatly

increased by low activation barriersbetween incorrect structures and thenative conformation. These barriers willbe lowest when the RNA interactions areweak. Destabilization of the folded RNAwill be more readily tolerated if proteinspreferentially bind and stabilize thenative RNA conformation.

In summary, the papers byBuchmueller et al.3 and Russell et al.4

now provide good evidence for confor-mationally disordered states in RNA thatare nearly as compact as the native struc-ture, a phase that is reminiscent of the‘molten globule’ states of proteins.However, the forces that induce RNAcollapse are quite different from thosethat drive the folding of polypeptides. Aswith many important observations,these results raise more questions thanthey answer. For example, how is RNAcollapse initiated — is it nucleated bysecondary or tertiary interactions, andwhat is the role of cations in this process?In addition, are the collapsed states aprerequisite for the formation of thenative state — in other words, do somecompact conformations lead directly tonative tertiary structure, or must theRNA chain become completely extendedbefore refolding? Exciting times areahead as the story of RNA folding con-tinues to unfold.

AcknowledgmentsThe author thanks D. Thirumalai and D. Draper forhelpful discussions.

Sarah A. Woodson is in the T. C. JenkinsDepartment of Biophysics, Johns HopkinsUniversity, 3400 N. Charles Street,Baltimore, Maryland 21218-2685, USA.email: swoodson@jhu.edu

Fig. 3 Folding of yeast mitochondrial bI5 ribozyme. The ribozyme core becomes increasingly compact at higher concentrations of MgCl2, as shown byBuchmueller et al.3. CBP2 (yellow) is proposed to selectively bind and stabilize the native structure. The conformations illustrated here are intermediatesin an equilibrium folding pathway, while the structures shown in Fig. 2 are kinetic folding intermediates. The structure of the RNA in the collapsed, non-native phase is unknown, and presumably corresponds to a mixture of conformations.

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1. Chan, H.S. & Dill, K.A. Proteins 30, 2–33 (1998).2. Bloomfield, V.A. Biopolymers 44, 269–282 (1997).3. Buchmueller, K.L., Webb, A.E., Richardson, D.A. &

Weeks, K.M. Nature Struct. Biol. 7, 362–366 (2000).4. Russell, R., Millett, I.S., Doniach, S. & Herschlag, D.

Nature Struct. Biol. 7, 367–370 (2000).5. Flanagan, J.M., Kataoka, M., Shortle, D. &

Engelman, D.M. Proc. Natl. Acad. Sci. USA 89,748–752 (1992).

6. Fink, A.L. Annu. Rev. Biophys. Biomol. Struct. 24,495–522 (1995).

7. Doudna, J.A. & Doherty, E.A. Folding Des. 2, R65–70(1997).

8. Thirumalai, D. Journal de Physique (France) 5,1457–1467 (1995).

9. Wolynes, P.G., Onuchic, J.N. & Thirumalai, D.Science 267, 1619–1620 (1995).

10. Thirumalai, D. & Woodson, S.A. Acct. Chem. Res.29, 433–439 (1996).

11. Eaton, W.A., Thompson, P.A., Chan, C.K., Hage, S.J.& Hofrichter, J. Structure 4, 1133–1139 (1996).

12. Draper, D.E. Parallel worlds. Nature Struct. Biol. 3,397–400 (1996).

13. Brion, P. & Westhof, E. Annu. Rev. Biophys. Biomol.Struct. 26, 113–137 (1997).

14. Cole, P.E. & Crothers, D.M. Biochemistry 11,4368–4374 (1972).

15. Riesner, D., Maass, G., Thiebe, R., Philippsen, P. &Zachau, H.G. Eur. J. Biochem. 36, 76–88 (1973).

16. Fang, X.W., Pan, T. & Sosnick, T.R. Nature Struct.Biol. 6, 1091–1095 (1999).

17. Pan, J., Thirumalai, D. & Woodson, S.A. J. Mol. Biol.273, 7–13 (1997).

18. Treiber, D.K. & Williamson, J.R. Curr. Opin. Struct.Biol. 9, 339–345 (1999).

19. Sclavi, B., Sullivan, M., Chance, M.R., Brenowitz, M.& Woodson, S.A. Science 279, 1940–1943 (1998).

20. Zarrinkar, P.P. & Williamson, J.R. Nature Struct. Biol.3, 432–438 (1996).

21. Rook, M.S., Treiber, D.K. & Williamson, J.R. J. Mol.Biol. 281, 609–620 (1998).

22. Russell, R. & Herschlag, D. J. Mol. Biol. 291,1155–1167 (1999).

23. Pan, J., Deras, M.L. & Woodson, S.A. J. Mol. Biol.296, 133–144 (2000).

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New tricks from an itinerant intronAnna Marie Pyle

A new mechanism for transposition by autocatalytic group II introns provides a working model for introndispersal in eukaryotic genomes.

It is often tempting to think aboutgenomes as stable, predictable archives ofgenetic information, much like a quietlibrary in which changes occur only by themis-shelving of a book (mutation) or by carefully orchestrated inter-libraryexchanges (recombination). But in fact,genomes are more like jungles. Althoughthey have a certain infrastructure, they areelaborate, detritis-strewn ecosystemsfilled with competing species. And despitethe fact that these species share a home(the host genome), many of them havetheir own agenda. Some of the mostunpredictable creatures prowling aroundthe genomic environment are transpos-able elements. These are pieces of nucleicacid that hop from place to place, some-times wreaking havoc and at other timesproviding a selective advantage for thehost organism1–4. The mechanisms of

transposition are very diverse and stillpoorly understood. However, recent workfrom the laboratory of Marlene Belfort5

has considerably advanced our under-standing of the mechanism by which amajor class of transposable elementsmoves to new target sites.

Transposons of various types havepropagated extensively in eukaryoticgenomes, to the extent that 10–40% ofmammalian genomes are comprised ofthis ‘selfish’ or ‘parasitic’ DNA6. Manytransposition events involve the move-ment of DNA fragments that encode orrecruit proteins that facilitate mobility.However, certain types of transpositionevents are catalyzed, at least in part, bynucleic acid molecules1,7. In these cases,the transposing elements are self-splicingor autocatalytic intron RNAs, which havebeen shown to catalyze their own excision

from precursor RNA both in vitro and invivo8.

For some time, it has been known thatthe DNA of self-splicing introns can moveand propagate like other transposable ele-ments7,9. A plausible mechanism for thisprocess was suggested by in vitro studieswhich showed that excised group I andgroup II intron RNAs have the ability to‘reverse splice’, or reincorporate them-selves, directly into RNA or DNA strandscontaining sequences similar to the 5′exons of the original splice sites10–12

25. Gampel, A., Nishikimi, M. & Tzagoloff, A. Mol. CellBiol. 9, 5424–5433 (1989).

26. Weeks, K.M. & Cech, T.R. Science 271, 345–348(1996).

27. Weeks, K.M. & Cech, T.R. Cell 82, 221–230 (1995).28. Lehnert, V., Jaeger, L., Michel, F. & Westhof, E.

Chem. Biol. 3, 993–1009 (1996).29. Treiber, D.K., Rook, M.S., Zarrinkar, P.P. &

Williamson, J.R. Science 279, 1943–1946 (1998).30. Pan, T. & Sosnick, T.R. Nature Struct. Biol. 4, 931–938

(1997).31. Michel, F. & Westhof, E. J. Mol. Biol. 216, 585–610

(1990).32. Pan, J. & Woodson, S.A. J. Mol. Biol. 294, 955–965

(1999).33. Myers, C.A. et al. J. Mol. Biol. 262, 87–104 (1996).34. Lambowitz, A.M. & Perlman, P.S. Trends Biochem.

Sci. 15, 440–44 (1990).35. Steitz, T.A. in The RNA World, Second Edition (eds.

Gesteland, R.F., Cech, T.R. & Atkins, J.F.) 427–450(Cold Spring Harbor Laboratory Press, Cold SpringHarbor, New York; 1999).

36. Downs, W.D. & Cech, T.R. RNA 2, 718–732 (1996).

Fig. 1 Splicing of group II introns is a reversible, two-step process. In Step 1 of splicing, the 2’-OHof an adenosine within the intron (red) is the nucleophile (top). It attacks the 5′-splice site, result-ing in liberation of a 5′-exon RNA and a lariat 3′-exon RNA (middle). In Step 2, the nucleophile isthe 3′-OH of the 5′-exon, which attacks the 3′-splice site, joining the exons and liberating the lari-at intron RNA (bottom). Note that this process can proceed in reverse, resulting in reintegration ofthe intron. In the first step of reverse-splicing (bottom), the 3 ′-OH of the lariat intron attacks thesplice junction (see black dot), resulting in formation of lariat-3′-exon (middle). In the second stepof reverse splicing, the 3′-OH of the free 5′-exon attacks the 2′-3′-5′ linkage at the branch site, lib-erating the 2′-OH as the leaving group and resulting in reincorporation of a linear intronsequence. In group II introns, the 5′-splice site is generally recognized by two sets of base pairs(dotted lines) between the exon binding sites of the intron (EBS1 and EBS2) and the intron bindingsites of the 5′exon (IBS1 and IBS2, green regions). The most important of these for ectopic retro-transposition is the IBS1 site (closest to the splice junction).

Step 1 ofsplicing

Step 2 ofreverse splicing

Step 2 ofsplicing

Step 1 ofreverse splicing

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