cystatin forms a tetramer through structural rearrangement of domain-swapped dimers prior to...

Cystatin forms a Tetramer through StructuralRearrangement of Domain-swapped Dimers priorto Amyloidogenesis

Anna Sanders, C. Jeremy Craven, Lee D. Higgins, Silva GianniniMatthew J. Conroy, Andrea M. Hounslow, Jonathan P. Waltho andRosemary A. Staniforth*

Department of MolecularBiology and BiotechnologyUniversity of Sheffield, FirthCourt, Western Bank, SheffieldS10 2TN, UK

The cystatins were the first amyloidogenic proteins to be shown to oligo-merize through a 3D domain swapping mechanism. Here we show that,under conditions leading to the formation of amyloid deposits, thedomain-swapped dimer of chicken cystatin further oligomerizes to atetramer, prior to fibrillization. The tetramer has a very similar circulardichroism and fluorescence signature to the folded monomer and dimerstructures, but exhibits some loss of dispersion in the 1H-NMR spectrum.8-Anilino-1-naphthalene sulfonate fluorescence enhancement indicates anincrease in the degree of disorder. While the dimerization reaction isbimolecular and most likely limited by the availability of a predominantlyunfolded form of the monomer, the tetramerization reaction is first-order.The tetramer is formed slowly (t1/2 ¼ six days at 85 8C), dimeric cystatinis the precursor to tetramer formation, and thus the rate is limitedby structural rearrangement within the dimer. Some higher-orderoligomerization events parallel tetramer formation while others followfrom the tetrameric form. Thus, the tetramer is a transient intermediatewithin the pathway of large-scale oligomerization.

q 2004 Elsevier Ltd. All rights reserved.

Keywords: cystatin; amyloid intermediate; CAA; domain-swapping; dimer*Corresponding author

Introduction

The deposition of amyloid is associated with arange of debilitating human disorders includingAlzheimer’s disease, type II diabetes, the spongi-form encephalopathies (CJD, BSE and scrapie),cancer and rheumatoid arthritis.1 – 7 The process ofamyloidogenesis involves the self-assembly ofsoluble protein or peptide into insoluble fibrillarmaterial. Although amyloidogenic proteins haveno sequence or structural homology in the solubleform, amyloid fibers typically maintain a commoncross-b-structure; that of an extended b-sheet

with strands running perpendicular to the fibrilaxis.8 – 14 In vivo amyloidogenesis can be mimickedin vitro, both for amyloidogenic proteins15 – 17 andfor some proteins not associated with amyloiddiseases,18 – 21 using solution conditions thatdestabilize their native states. However, thedetailed structure of amyloid and the mechanismby which it is formed remain poorly understood.Current models of amyloidogenesis21 – 28 are limitedowing to the relatively featureless nature of thetransitions, generally comprising a pre-exponential(lag) phase and an exponential (propagation)phase, and the difficulty in identifying the corre-sponding changes in structure. However, confor-mational transitions within the pre-exponentialphase of fibrillogenesis have been reported forseveral proteins,29 – 34 including the cystatins,35 – 37

but the fate of these species in the hierarchy ofassembly remains largely undefined. In additionto defining the mechanism of fibrillogenesis, theimportance of understanding initial steps is furtherunderlined by the current lack of identification of

0022-2836/$ - see front matter q 2004 Elsevier Ltd. All rights reserved.

E-mail address of the corresponding author:[email protected]

Abbreviations used: hCC, human cystatin C; cC,chicken cystatin; GdnHCl, guanidine hydrochloride; CR,Congo red; ANS, 8-anilino-1-naphthalene sulfonate; M,monomer; D, dimer; T, tetramer; SEC, size-exclusionchromatography; AUC, analytical ultracentrifugation;CD, circular dichroism; pdb, Protein Data Bank.

doi:10.1016/j.jmb.2003.12.011 J. Mol. Biol. (2004) 336, 165–178

the primary cause of clinical symptoms ofamyloid diseases and the consequent targeting oftherapies towards the initial stages of proteinassociation.

The cystatins are a superfamily of structurallyhomologous cysteine proteinase inhibitors38

(Figure 1A) implicated in a number of degenera-tive disorders. Cystatin B (also known as stefin B),a member of the type I cystatin family (termedstefins), is linked to a hereditary progressivedisease that manifests itself as a fatal form of myo-clonus epilepsy.39 Cystatin C, a member of the typeII cystatin family,40 is associated with cerebralamyloid angiopathies (CAA), a disease in whichthe deposition of amyloid in blood vessels leads torecurrent hemorrhagic stroke. This prevalentdisease of the elderly41,42 is commonly observed inAlzheimer’s patients where layers of cystatin Care often co-deposited within Ab plaques.43 – 48 Inaddition, hereditary cystatin CAA (HCCAA) is theresult of a destabilizing mutation (L68Q) withinthe core of cystatin C and leads to prematuredeath (30–40 years of age).49 – 51

All of the cystatins studied in detail to date formstable yet biologically inactive dimers, under con-ditions that destabilize the native structure andthat lead to amyloidosis.35,52 – 54 These dimers are3D domain-swapped (Figure 1B),37,55 i.e. a non-covalent interface within the monomer is brokenand its place is taken by the corresponding regionof the other molecule. This relatively uncommonself-association phenomenon has been widelyproposed as a candidate for some of the steps inamyloid formation,56 – 59 and has subsequentlybeen shown to occur in a second amyloidogenicprotein, human prion protein.60 This mechanismof protein assembly has the attraction of reconcil-ing aggregation events with the rules that govern

the folding of globular proteins: both phenomenacomply with the principle of minimal frustration,sharing common folding nuclei and favoringnative over non-native contacts.61 To date, though,it has not been determined whether domain swap-ping exists in mature amyloid. However, thedimensions of domain-swapped oligomers andthe large, stable, intermolecular contact surfacethat is generated by domain swapping are bothconsistent with the observed properties of amyloidfibers.37 Furthermore, in the case of the cystatins,when fibrillogenesis is triggered, 3D domain-swap-ping dimerization is an inevitable occurrence in thepre-exponential phase.35 In this study, we investi-gate the fate of domain-swapped dimers duringthe pre-exponential phase of cystatin fibrillo-genesis and report the tetramerization of chickencystatin, the chicken homologue of human cystatinC. This protein has the experimental advantageover human cystatin C of a higher thermodynamicstability for the folded state.

Results

Cystatins oligomerize under pre-denaturingconditions

The destabilization of the folded state introducedby the mutation involved in HCCAA (L68Q) is suf-ficient for human cystatin C (hCC) to dimerizethrough 3D domain swapping at a measurablerate under physiological conditions.35 Dimerizationis similarly observed at room temperature for theequivalent mutant (I66Q) of chicken cystatin(cC).37 In the case of wild-type proteins, destabiliz-ing but pre-denaturing conditions of pH,temperature or chemical denaturants (such as

Figure 1. Representative examples of the fold of cystatin (pdb code 1CEW) monomers and 3D domain-swappeddimers. A, The monomer of chicken cystatin. The two disulfide bonds are shown in green. The long mobile loop con-necting strands 3 and 4, colored yellow, is positioned upper left. Loop1, the hinge region where the domain swappingevent associated with dimerization occurs, colored blue, is positioned lower right. B, The dimer of human cystatin C(pdb code 1G96) showing where the individual polypeptide chains, colored red and blue, are positioned followingdomain swapping. Loop 1 now forms a continuous b-strand linking the two cystatin fold units.

166 Tetramers of Cystatin

guanidinium chloride or SDS) are required to bringthe dimerization rate into an experimentally acces-sible time window.37,62 Extrapolating back to phys-iological conditions, dimerization remainsthermodynamically favored at all measuredprotein concentrations of cC (as low as 600 nM).37

Prolonged incubation of chicken cystatin (cC)under destabilizing but pre-denaturing conditionsleads to further self-association, as illustrated byincubation at high temperature (Figure 2A and B).A size-exclusion chromatogram after incubation ofmonomeric cC at 85 8C for 50 hours shows that, inaddition to dimers, a species with a retentionvolume of 8.3 ml is produced. This retentioncorresponds to a species that is consistent with theformation of a tetramer of cC (predictedMr ¼ 54 kDa). Intriguingly, and in contrast withother domain swapping proteins such as CD263 orRNase A,58,64 no trimeric or pentameric species aredetectable, suggesting that in the present case, the

dimer rather than the monomer is likely to be theassembly competent species (see below).

Isolation of a tetrameric species

Using temperature as the means of destabiliza-tion has the advantage of providing a convenientand rapid means of trapping transient kinetic inter-mediates through quenching to ambient tempera-tures. The putative tetramer was prepared withover 95% purity via thermal quenching andisolation using preparative size exclusionchromatography (SEC) at room temperature(Figure 2C). The purity of the tetramer wasconfirmed independently using ion-exchangechromatography. On storage at room temperaturefor greater than ten days, no significant re-equi-libration to other oligomeric states was observed.The molecular mass of the isolated tetramer wasdetermined independently to be 55(^3) kDa by

Figure 2. Resolution and molecular mass of the cC tetramer. A, Size exclusion chromatograms of aliquots of cC(66 mM) incubated at 85 8C for zero hours (dotted line), five hours (continuous line), and 50 hours (broken line). Specieswith elution volumes of approximately 10 ml, 9 ml and 8 ml correspond to monomers, dimers and tetramers,respectively. B, Vertical expansion of the time-course shown in A illustrating the build-up of tetrameric cC. Highermolecular mass species are also evident but are less definable. C, Size-exclusion chromatograms of purified samplesof monomer (continuous line), dimer (broken line) and tetramer (dotted line) samples of cC showing the resolutionof the purification. D, Experimental (continuous line) and fitted (broken line) sedimentation velocity AUC profiles ofisolated cC tetramer (52 mM). The derived sedimentation coefficient of 3.1 S corresponds to four cC monomer units.

Tetramers of Cystatin 167

fitting sedimentation velocity data (Figure 2D)obtained from analytical ultracentrifugation(AUC) to the Svedberg equation, using a trans-lational diffusion coefficient, determined by light-scattering, of 5 £ 1027 cm2 s21. The sedimentationcoefficient as a function of protein concentrationconfirmed that no significant re-equilibration orfurther aggregation occurs after quenching of thereaction to room temperature on the timescale ofthe AUC experiment (three days). The tetramerdoes not contain intermolecular disulfide bonds orother chemical modifications since it reverts to amonomeric form under non-reducing SDS-PAGEand mass spectrometry (data not shown).

The tetramer is structurally analogous to thefolded monomer and dimer states

The 1D 1H-NMR spectrum (Figure 3A) of thetetramer of cC shows loss of resolution, owing toline-broadening, compared with equivalent spectraof the dimer or monomer. This is expected as aresult of the increase in molecular mass, but isfurther exacerbated most likely by non-specificself-association and/or intermediate chemicalexchange. NMR line-broadening by thesephenomena is also observed for monomers anddimers of cC,37,65,66 and hCC,62 which otherwise arehomogeneous species amenable to high-resolutionstructure determination. Some loss of resonancedispersion is also observed for the tetramer com-pared to the dimer, suggesting a degree of disrup-tion of the overall fold, though this loss is not tothe extent observed within spectra of moredisordered states such as molten globules. Amolten globule species has been observed for cCupon reduction of the disulfide bond betweenresidues 95 and 115 (termed R in Figure 3A),where the resonance dispersion is close to that ofits unfolded state.37 Unfortunately, the resonanceline-width and the relative lack of dispersion inthe NMR spectrum preclude further straight-forward structural analysis.

Other spectroscopic measurements confirm thatthe overall cystatin fold, present within monomersand dimers (Figure 1), is preserved in tetramers.Secondary structure analysis using far-UV CD(Figure 3B) shows the monomer, dimer andtetramer states of cC to be indistinguishable. Near-UV CD and tryptophan fluorescence spectra(Figure 3C and D) reveal only marginal differenceswith the folded states of dimers and monomers,indicating that aromatic residues are found inequivalent environments in tetramers. A smallblue-shift in the fluorescence maximum of thetryptophan residue (348 nm ! 343 nm), whichoccurs on dimerization, is preserved in thetetramer lending further support to the hypothesisthat tetramer is made up of dimers. In contrast,8-anilino-1-naphthalene sulfonate (ANS), whichprobes exposed hydrophobic clusters, bindstetramers of cC preferentially (Figure 3E) indicat-ing that there is some increase in solvent exposure

of the protein core. Taken together, the structuraldata support the hypothesis that, although thecystatin fold is maintained in the tetramer of cC,some disruption of the stability of the fold hasoccurred. This disruption may involve only someregions of the protein, or may be transient, i.e. arapid folded state–molten globule state exchangemay be occurring in which both states arepopulated significantly.

The cystatin multimerization reaction

The kinetics of self-association of cC at 85 8C, pH7.0 were monitored using SEC (Figure 4A). At a cCconcentration of 66 mM, 3D domain swappingdimerization occurs with a half-time of 0.6 hours.The dimer population subsequently converts witha half-time of 40 hours partially to tetramers and

Figure 3. Structural characterization of the isolatedtetramer. A, 1H-NMR spectra of the amide (left) andmethyl (right) chemical shift regions of (from top tobottom) the tetramer (T), the reduced molten globule(R), native-like dimer (D) and monomer (M) of cC. B toE, Comparative spectroscopy of isolated monomers (con-tinuous line), dimers (broken line), and tetramers (dottedline) of cC in 50 mM PO4 (pH 7.0), using far-UV CD (B),near-UV CD (C), intrinsic tryptophan fluorescence (D)and extrinsic ANS fluorescence (E).


partially to multimers. The multimer population isnot necessarily a single species; it comprises all ofthe species that exceed the molecular mass limit ofthe SEC column (i.e. Mr . 1 MDa, which corre-sponds to in excess of 70 monomers). The tetramerpopulation also eventually converts to multimers,this process being more clearly observed undermore strongly destabilizing conditions. At 85 8C,pH 7.0 in the presence of 3 M GdnHCl (Figure4B), conditions which are still pre-denaturationalfor cC, the half-time for dimerization falls withinthe dead time of the experiment (0.08 hours),while the dimer-to-tetramer and dimer-to-multimer transitions have half-times of ,2 hours.The half-time of the tetramer to multimertransition under these conditions is ,20 hours.

The transition state to tetramer formation ismore folded than for dimerization

The guanidine dependence of the tetrameriza-tion reaction rate (Figure 4C) highlights differencesbetween the dimerization and the tetramerizationtransitions. This dependence (termed m‡) reportson the degree of solvation (or of denaturation) ofthe transition state ensemble.67 It reveals a far smal-ler effect for tetramerization ðm‡ðD!TÞ ¼ 1 M21Þthan observed for the dimerization reactionðm‡ðM!DÞ ¼ 4:4 M21Þ, and as reported37 for dimeri-zation at 25 8C ðm‡ðM!DÞ ¼ 5 M21Þ: Dimerization ofcC is rate-limited by a bimolecular process, whichindicates one of two experimentally indistinguish-able scenarios. Either the reaction occurs througha poorly populated intermediate state, Mp, in aScheme where 2M ! 2Mp ! D and KMp=M is verysmall (hence 2Mp ! D is rate-limiting) or, alterna-tively, an energetically expensive D0 ! D transitionoccurs after association, within a reaction Schemewhere 2M ! 2Mp ! D0 ! D. In this latter case, theputative D0 state would have to be too unstable tobe significantly populated, since a transient dimerstate is not observed (using a spectroscopic tech-nique such as NMR37) but the transition state

would be dimeric rather than monomeric. Thehigh guanidine hydrochloride dependence of thedimerization rate reflects the degree of denatura-tion of, in the former case, the excited state, Mp,and in the latter case, the transition state betweenD0 and D compared with M. In either case, the reac-tion is rate-limited by a major unfolding event. Incontrast, the degree of denaturation of the tran-sition state to tetramer formation is low, indicatingthat this reaction is rate-limited by a very differentprocess, with limited unfolding.

The tetramer is formed directly from the dimer

In principle, the formation of the tetramer couldbe the result of one of two Schemes, namely:

4M O 2D O T ðScheme 1Þ

2D O 4M O T ðScheme 2Þ

At 85 8C and pH 7.0, the rate of tetramer formation(1.3 £ 1026 s21) is indistinguishable whether thereaction is started from 100% dimer or from 100%monomer. In the latter case, dimerization precedestetramerization by two orders of magnitude orgreater at cC concentrations throughout the rangemeasured (10 mM to 1.8 mM), and no residualmonomer is detectable following dimerization.While this behavior does not preclude eitherScheme, there are a number of observations withwhich a mechanism requiring re-conversion of thedimer-to-monomer (Scheme 2) can be discounted.Firstly, no burst phase of tetramer formation isobserved when starting from 100% monomer,which, if Scheme 2 were correct, would indicatethat M must form D at least an order of magnitudefaster than it can form T. Since at, 10 mM cC, theinitial rate of the M ! D transition is1.4 £ 1024 s21, this provides a limit to the putativedirect M ! T transition inherent within Scheme 2of slower than 1.4 £ 1025 s21. Secondly, a require-ment for a minimum excess of D over M of oneorder of magnitude at 10 mM cC, as no residual M

Figure 4. Kinetics of cystatin assembly. SEC analysis of the formation of higher-order species of cC (monomer (X),dimer (O), tetramer (B), multimer (A)) when incubated at a concentration of 66 mM in 10 mM PO4 (pH 7.0) at 85 8Cin the absence (A) and in the presence (B) of 3 M GdnHCl. In C, the derived GdnHCl dependence of the observedrate constants of dimerization (þ ) and tetramerization (W) are compared.


is detectable following dimerization, further limitsa putative M ! T transition to faster than1.3 £ 1025 s21, since the observed rate of D ! T is1.3 £ 1026 s21. Taken together, the only scenario inwhich Scheme 2 is plausible is where the M ! Ttransition occurs at ,1025 s21. However, thisscenario can be eliminated when comparing theproduction of T as a function of protein concen-tration. As the concentration of cC is increased,the rates of dimer formation and tetramer for-mation both increase. Given the above rate limits,at low cC concentration the production of tetramerpredicted by Scheme 2 in the early stages of thereaction (i.e. d½T�=dt $ 1025½M�) is not observed.In contrast, the prediction from Scheme 1 that therate of production of the tetramer should be relatedto the concentration of dimer (i.e. d½T�=dt / ½D�) isobserved throughout (see the next section).

Tetramer formation is limited by aunimolecular process

In contrast to the monomer-to-dimer transition,the dimer-to-tetramer transition remains pre-dominantly first-order at protein concentrations aslow as were measurable (10 mM [M]initial;Figure 5A–C). A double logarithmic plot ofd½T�=dt versus [D] exhibits a slope of 1.1 ^ 0.2(Figure 5D). Consequently, the rate of tetramer for-mation is limited by a slow conformationalrearrangement (an opening step) within the dimer,rather than by the subsequent step depicted inScheme 3:

D þ D OkD!Dp

kDp!D

Dp þ Dp OkDp!T

kT!Dp

T ðScheme 3Þ

Fitting the data using the differential equationsimplicit in Scheme 3 (see Materials and Methods)and using observed dimer concentrations yieldsan activation (opening) rate constant for the dimerkD!Dp ¼ 1:3 £ 1026 s21 and a value for the ratiokDp!T=ðkDp!DÞ

2 ¼ 2 £ 105 mM21 s. At sufficientlyhigh protein concentration this Scheme wouldlead to a classical uni-molecular reaction. Thecross-over between bi and uni-molecularlimiting regimes occurs when kD!Dp ½D�=g , 1,where g ¼ ðkDp!DÞ

2=4kDp!T (i.e. when½D� , ðkDp!DÞ

2=4kD!DpkDp!T ¼ 1 mM), which isbelow the concentration range measurable here.Some considerations can also be made as tothe values of kDp!D and kDp!T: The lack of anobservable lag phase in the formation oftetramer following the build-up of dimer in anexperiment where ½M�o ¼ 10 mM, places a lowerlimit on kDp!D of ,3 £ 1024 s21. SincekDp!T=ðkDp!DÞ

2 ¼ 2 £ 105 mM21 s, a lower limit forkDp!T becomes ,2 £ 104 M21 s21. An upper limitcan be estimated for kDp!D by assuming that kDp!T

is diffusion limited at 1 £ 109 M21 s21, which corre-sponds to a maximum rate of 7 £ 1022 s21.

Multimerization of dimers and tetramers

The rate of formation of tetramers and the rate ofmultimerization of dimers are comparable(Figure 4A). At longer times (for example, a half-time of ,20 hours in 3 M GdnHCl, Figure 4B), thepopulation of tetramers is diminished in favor ofthe population of multimers and the tetramer istherefore a transient intermediate. Structurally, themultimer ensemble generated by incubation at85 8C and pH 7.0, when examined by negativestain electron microscopy, is heterogeneous, inwhich both fibrous material and more amorphousmaterial are well represented (Figure 6A and B).The fibrous material is of the type referred to

Figure 5. Concentration dependence of tetramerformation. Time courses showing the appearance of thetetrameric form of cC starting with 1000 mM, 100 mMand 10 mM monomeric protein in A, B and C, respect-ively. Error bars represent an estimate of the reproduci-bility of the individual measurements. Continuous linesare fits to the data according to the uni-molecularrelationship d½T�=dt ¼ kD!Dp :[D], where kD!Dp is1 £ 1026 s21. In D, the double logarithmic plot of the rateof tetramer formation as a function of [D] shows thedata collectively, and a linear fit gives a slope of1.1 ^ 0.2, characteristic of a uni-molecular reaction.


previously as “protofibrils” or “immature fibrils”for the cases of an SH3 domain,68 alpha-synuclein69

or beta-2-microglobulin.17 An examination of themultimer ensemble at lower resolution (Figure 6Cand D) reveals that the generated cC multimersstain with Congo red, but only a portion displaythe golden green birefringence characteristic ofamyloid. While this also demonstrates that kineticpartitioning occurs, it precludes a straightforwarddemonstration of the independence of the multi-merization routes based on the observed kinetics.It is also not possible to distinguish whether multi-merization of tetramers occurs directly, or whetherthis proceeds via re-formation of dimers. Underthe experimentally accessible conditions, the rateof re-equilibration of tetramer to smaller speciesðkT!DÞ may be comparable or faster than the rateof multimerization ðkT!MultimerÞ, since only upperlimits on either of these parameters can be deter-mined. Whether the tetramer plays a direct or anindirect role in the formation of fibrous ratherthan amorphous material was also not discernibleby starting multimerization from the purifiedtetramer. No substantial change in the hetero-geneity of the final multimer population isobserved whether purified tetramer, dimer ormonomer is the starting material, which is consist-ent with either re-equilibration or downstreambranching of the pathways for multimerization.

Discussion

Oligomerization in the pre-exponential phase

During the pre-exponential phase of cystatinmultimerization, at all measurable protein concen-trations, the folded monomer (the active proteinaseinhibitor) firstly undergoes a bimolecular transitionto a domain-swapped dimer, via a predominantlyunfolded transition state, followed by a uni-molecular transition to a tetramer via a predomi-nantly folded transition state, that can be describedby the following Scheme:

4M OkM!Mp

kMp!M

4Mp OkMp!D

kD!Mp

2D OkD!Dp

kDp!D

2Dp OkDp!T

kT!Dp

T ðScheme 4Þ

The values or ranges for the component rate con-stants for oligomerization at pH 7.0 and 85 8C aregiven in Table 1. Whether a transition state (orindeed any state) is predominantly folded orunfolded is determined by the relative denaturantconcentration dependence of its free energy, com-monly referred to as its m-value. The relationshipof the m-values of the dimerization (dotted line)and tetramerisation (broken line) transition states,compared with the states involved in the normalfolding and unfolding of monomeric cC, is illus-trated in Figure 7. Monomeric cC refolds througha well-populated intermediate state (I) and a com-mon transition state (TS) regardless of whether theendpoint is the native state ðFoxÞ, in the casewhere both disulfide bonds are intact, or a moltenglobule state ðFredÞ; in the case where both disulfidebonds are reduced.70 The transition state for dimer-ization, namely either an excited state of themonomer, Mp, or a high energy transition within aloosely associated dimer, is more unfolded thaneither the kinetic intermediate state or the refoldingtransition state of monomeric cC. The high freeenergy implied by such a major unfolding event isconsistent with the low population of Mp beingthe source of rate limitation, as illustrated inScheme 4. Assuming this scenario, the free energyof Mp (Figure 7) would be appropriate for itsdenaturant dependence within the normal folding

Figure 6. Characterization of multimers. Negativestain transmission electron microscopy images of fibrous(A) and amorphous (B) aggregates. The short, curlymorphology evident in the right image is typical ofprotofibrils or “immature” amyloid fibers. The bar rep-resents 100 nm in both images. Representative aggre-gates of cC generated by prolonged incubation at 85 8Cand stained with Congo red as viewed under partiallypolarized (C) and under fully cross-polarized light (D).

Table 1. Rate constants for the tetramerization of chickencystatin at 85 8C

kM!D 14 ^ 2 M21 s21

KMp=Ma 1.4 ^ 0.2 £ 1028

kD!Dp 1.0 ^ 0.3 £ 1026 s21

kDp!D 3 £ 1024–7 £ 1022 s21

kDp!T 2 £ 104–1 £ 109 M21 s21

kT!D ,1 £ 1026 s21

Values for the constants describing reaction Scheme 4 arelisted. Since only the ratio kDp!T=ðkDp!DÞ

2 can be determined(2 £ 105 mM21 s), upper and lower limits for the values of kDp!D

and kDp!T are listed.a Theoretical value for the equilibrium constant describing

the stability of Mp compared with M, in the limit where dimeri-zation is rate-limited by the diffusion of activated monomers,i.e. kMp!D ¼ 1 £ 109 M21 s21.


scheme of the protein. In contrast, the transitionstate for tetramerization (which is associated withthe transition D ! Dp) is significantly more foldedthan the kinetic folding intermediate (I) or themajor refolding transition state (TS). The denatur-ant dependence of its free energy is closer to thatof the well-defined and relatively stable equi-librium molten globule state of cC generated ondisulfide reduction ðFredÞ, indicating that rate-limitation of the D ! Dp transition arises from adifficult conversion within the folded stateensemble.

Domain swapping within the tetramer

The slow kinetics of tetramer formation and theconformational data are inconsistent with modelswhere the formation of tetramer is a simple,diffusion-controlled association of dimers. Thelow rate of the uni-molecular step precedingtetramer formation is consistent with more sub-stantial structural rearrangements in the dimerwhere the product returns to a fold highlyanalogous to that found in monomers and dimers.The 3D domain swapping is a highly likely candi-date mechanism to account for these observations,

particularly since domain swapping already occursunder these same conditions during dimerization.It is therefore of value to consider models throughwhich this may occur, in comparison with othermodels recently reported.58 – 60,71 The ability of aprotein to favor domain swapping over intra-molecular folding depends on the free energychange associated with the formation of any newinterface, and the effective concentration ðCeffÞ ofthe two swapping domains: Ceff ¼ KðintraÞ=KðinterÞ,where KðintraÞ and KðinterÞ are the association con-stants for the domains within the monomer andoligomer, respectively. Ceff is generally large inprotein folding, where it is a manifestation of thecooperativity that leads to a high value for KðintraÞ

from linked but individually weak, non-covalentinteractions. A reduction in the effective concen-tration of the domains occurs when the folded con-formation of the macromolecule in the monomericstate is strained in the inter-domain (hinge) region,and also when the inter-domain region is flexible.In the latter case, Ceff typically falls with the lengthof the flexible region.

Although dimerization and tetramerization of cCare rate-limited by different transition states(Figure 7), this does not exclude the possibilitythat the same process is occurring on bothoccasions (Figure 8A), as in the hypotheticalmodel suggested previously for hCC.55 This modelimplies similar properties for Mp and Dp and there-fore similar conformational changes involved inthe M N Mp and D N Dp transitions. Withoutknowledge of the dimerization dissociation con-stant, KD=M, which is below the experimental detec-tion limits (i.e. below the cross-over point to abimolecular regime, at 1 mM ½M�initial), it remainspossible that the change in the rate-limiting stepsimply reflects the increased stability of the dimerover both that of the monomer and that of the tran-sition state for the 3D domain swapping involvedin dimerization. Similarly, limits in sensitivity alsopreclude the evaluation of the solvation propertiesof Dp and thus a direct comparison with theproperties of Mp. Against this model is the diffi-culty in identifying a thermodynamic drivingforce for further domain swapping of this type,since dimerization already relieves the strain inthe loop I region of cystatins (Figure 1).37

Equally likely is domain swapping usingalternative interfaces, of which there are two strongcandidates (Figure 8B and C). First, the extendedloop linking strands 3 and 4 will have a reducedeffective concentration for its two ends owing tothe mobility demonstrated for cC65 and impliedfor hCC,52,62 and therefore would have a loweredconcentration threshold for domain swapping.Secondly, conformational heterogeneity is apparentin the solution behavior of cystatin A at the end ofthe helix and the start of strand 2,72 implicatingthis region in some packing anomalies that mayreduce the effective concentration of parts of thisregion. In the latter event, domain swappingwould exchange strand 1 and the helix between

Figure 7. Relationship between refolding and oligo-merization. The free energy as a function of the degreeof folding (GdnHCl dependency or m-value) of theunfolded state (U), the kinetic intermediate state (I), therefolding transition state (TS) and the folded (F) statefor the oxidized (ox) monomeric form of cC (X), and thereduced (red) monomeric form (A). Superimposed onthis are the corresponding GdnHCl dependencies fordimerization (dotted line) and tetramerization (brokenline). The free energy value illustrated for dimerization(filled grey circle) is that predicted from a model wheredimerization is rate-limited by the diffusion of Mp.


monomer fold units. Whatever the mechanism, theability of the tetramer to bind ANS indicates thatthe rigidity of structure of some of the hydrophobiccore of the monomer and dimer is compromisedwithin the tetramer. This mirrors the observationsfor the fibrillogenesis of yeast prion, where oligo-meric intermediates are less rigidly structuredthan the mature fibers.25 Meanwhile, conditionsthat stabilize cC tetramer or alternative types ofcystatin tetramers are being actively sought so asto distinguish between these different models.

Tetramers in the context of fibers

The considerable population of tetramers duringthe pre-exponential phase makes these strong can-didates for intermediates in the formation offurther oligomers and, potentially, of fibrils. Thehydrodynamic radius of the tetramer ðRh , 46 �AÞ,derived from a combination of light scatteringand AUC, is appropriate for the dimensions ofamyloid fibrils prepared from cystatins (diameter7–13 nm),73,74 and hence the tetramer is a plausiblebuilding block from which fibrils are made.Assembly of tetramers into fibrils may be occur-ring by lateral association or further domain swap-ping. An interesting alternative is that cystatinfibrils are constructed from domain swapping ofdimers, for example as illustrated in Figure 8D,where the observed formation of tetramers are theresult of an off-pathway cyclization reaction, thetrue but lowly populated intermediate to propa-gation being a linear rather than a cyclizedtetramer. The resolution of these multimerizationphases is now underway; screening for conditionsin which a homogeneous fibrillar sample can beobtained will allow us to carry out a more in-depth structural study.

The population of critical oligomeric speciesprior to elongation events is a feature of some cur-rent fibrillization models.21,25 In these cases, theintermediates are populated as the assemblyswitches from bulk to linear growth of aggregatedspecies and appear to be heterogeneous both instoichiometry and structure. These models alsopredict that their final amyloid structure is notstabilized until the later stages in the process (inthe case of b-amyloid formation34 the oligomerpopulation is partially a-helical), i.e. the reaction

Figure 8. Models of 3D domain-swapped tetramersand multimers. Proposed models of symmetric tetramers

derived from 3D domain-swapped dimers generatedthrough further domain swapping using (A) the sameinterface as for the initial swapping event, (B) theextended loop linking strands 3 and 4 and (C) the endof the helix and the start of strand 2. Each color corre-sponds to an original monomeric unit. D, A schematicrepresentation of how domain swapping in cystatinscould propagate into an elongated b-sheet structurebased, in this case, on model C. The repeat unit isshown such that the addition of an infinite number ofunits in a head-to-tail fashion will make up the modelfiber.


is not a simple two-state process where monomersare incorporated onto growing fibers. In contrast,an oligomeric intermediate populated duringfibrillogenesis of prion protein75 has been shownto be off the pathway of amyloid formation. How-ever, off-pathway intermediates have attractedmuch attention from a medical viewpoint. Forexample, discrete oligomeric intermediates ofa-synuclein have been shown to disruptmembranes76 and are stronger candidates thanfibrillar forms for the observed in vitro cytotoxity.

Cystatin assembly represents an example wherediscrete intermediates can be both defined and iso-lated. In the early phases of multimerization, it isshown here that cystatins undergo a hierarchicalassembly process that involves slow and well-defined transitions to dimeric and tetramericspecies in which 3D domain swapping is at leastpartly involved.

Materials and Methods

Production and isolation of tetramericchicken cystatin

Recombinant wild-type chicken cystatin (cC, Mr ¼13,500 Da) was expressed in Escherichia coli (TG1) usingthe pIN-III-OmpA system, and purified from the peri-plasmic extract using papain affinity chromatography77

followed by size-exclusion chromatography (Superdex75, Pharmacia) under conditions which favor dis-sociation of substrate proteinases (0.5 M KCl, 50 mMK3PO4/K2HPO4 (pH 11.5)). Protein concentrations wereevaluated using an extinction coefficient at 280 nm of0.871 AU mg ml21. Tetramerization was achieved byincubating 50 mM–2 mM wild-type cC at 85 8C (10 mMpotassium phosphate (pH 7.0)) for 72 hours. Thetetramer was separated from residual dimeric cC bysize-exclusion chromatography (Superdex 75), in 50 mMpotassium phosphate (pH 7.0). The purity of the tetra-mer was established by ion-exchange chromatography(Shodex IEC QA-825, Japan) using a 0.1–0.4 M NaClgradient in 50 mM Tris (pH 8.0). The tetramer was thendialyzed (spectrapor MWCO of 6–8 kDa) against 2 £ 5 lof 10 mM potassium phosphate, 1 mM sodium azide(pH 7.0), and stored at 4 8C at protein concentrations,50 mM.

Molecular mass determination

The molecular mass and purity of the tetramer wasdetermined by size-exclusion chromatography (SEC)using a BIOSEP-SEC 3000 HPLC column (Phenomenex,CA) in 50 mM potassium phosphate (pH 7.0), previouslycalibrated using a standard set of low molecular massmarkers (Pharmacia). Elution times were determined bymeasuring the absorbance at 280 nm and 224 nm. Peakareas were deconvolved by fitting to a series ofGaussians using Datamax v. 4.14 software (Jobin YvonInc., NJ). In addition, sedimentation velocities weremeasured using analytical ultracentrifugation (AUC) ata range of protein concentrations (0.2–0.7 mg ml21)using a Beckman Optima XL-1, previously equilibratedto 20 8C. Data were fitted to the Svedberg equation: Mr ¼RTs=ðDð1 2 vrÞÞ; where Mr is the molecular mass (Da), s

is the sedimentation coefficient (S), R is the gas constant,T is the temperature (Kelvin), D is the translational diffu-sion coefficient (cm2 s21), v is the partial specific volumeof the solute (ml g21) and r is the solvent density(g ml21). The translational diffusion coefficient, D, wasmeasured by dynamic light scattering (DLS). A 4 mMsolution of tetrameric cC (10 mM potassium phosphate(pH 7.0)) was injected through a 300 A syringe filter intoa Dynapro-801 TC molecular sizing instrument. Thewavelength of the laser was 7830 A and a 908 scatteringangle was measured.

Spectroscopy

One-dimensional 1H-NMR spectra of the different cCstates were acquired at 20 8C and 37 8C on a BrukerDRX-600 spectrometer and were processed using Felix(Accelrys). All protein concentrations were 500 mM interms of monomer equivalents. Circular dichroism (CD)measurements were carried out on a Jasco J-810 spectro-polarimeter (far-UV CD: pathlength ¼ 1 mm; proteinconcentration ¼ 20–30 mM; near-UV CD: pathlength ¼5 mm; protein concentration ¼ 75 mM). Fluorescencemeasurements were carried out on a ShimadzuRF5301PC fluorimeter using a protein concentrationof 5 mM with or without the addition of 50 mM ANS(8-anilino-1-naphtalene sulfonate). Wavelengths usedwere lex ¼ 290 nm, lem ¼ 300– 420 nm for tryptophanfluorescence and lex ¼ 380 nm, lem ¼ 400– 550 nm forANS fluorescence. All protein solutions were in50 mM potassium phosphate (pH 7.0) at 20 8C.

Kinetic measurements

The time dependence of tetramer production was fol-lowed by removing aliquots of the incubation mixture(monomeric cC, 10 mM potassium phosphate (pH 7.0)at 85 8C) at set intervals and by quenching the reactionby dilution with protein-free incubation buffer (20 8C) toa final protein concentration of 25 mM or by rapid cool-ing on ice (for samples in the lower protein concentrationrange). These aliquots were then analyzed using SECHPLC as described above. The rates of the transitionsbetween monomer (M) and dimer (D) were derived asdescribed.37 The rates of the transitions involving thetetramer (T) were analyzed according to the followingminimal Scheme:

D þ D OkD!Dp

kDp!D

Dp þ Dp OkDp!T

kT!Dp

T

where D converts to an excited (open) form (Dp) prior tocombination with a second molecule in the Dp state toform T. Thus, kD!Dp is the uni-molecular opening rate ofthe dimer, kDp!D is the uni-molecular closing rate of Dp,kDp!T is the bi-molecular rate constant by which two Dp

associate to form T, and kT!Dp is the uni-molecular dis-sociation of T. In such a model, the bi-molecular ratekDp!T is limiting at low concentrations of dimer, and theuni-molecular rate kD!Dp becomes limiting at high con-centrations. The accessible experimental window ininitial protein concentration was extensive, rangingfrom 10 mM to 1.8 mM initial monomer units. Rates oftetramer formation were detected throughout the con-centration range of dimer, including during its formationfrom monomer and its depletion to tetramer ormultimers.

The parameters in the model were determined by fit-ting the observed tetramer concentration to the observed


dimer concentration as a function of time at initial mono-mer concentrations of 10 mM, 100 mM and 1000 mM. Thefitting involved using numerical integration of the differ-ential equations implicit in the above Scheme to calculatethe formation of tetramer as a function of time givenexperimentally observed dimer concentrations, coupledwith parameter optimization via a Levenberg Marquadtnon-linear least-squares algorithm. It was observed inthe fitting procedure that whereas the rate constantkD!Dp could be determined, the quality of the fit wasonly sensitive to the ratio ðkDp!DÞ

2=kDp!T, so that the par-ameters kDp!D and kDp!T could not be independentlydetermined. This can be rationalized as follows. Sincewe observe no lag phase in the formation of T, we canmake the steady-state approximation that:

d½Dp�=dt ¼ kD!Dp ½D�2 kDp!D½Dp�2 kDp!T½D

p�2

þ kT!Dp ½T� ¼ 0 ð1Þ

Solving equation (1) for [Dp] and ignoring the back rate,kT!Dp , gives:

d½T�=dt ¼ g{ 2 1 þpð1 þ kD!Dp=g½D�Þ}2 ð2Þ

where g ¼ ðkDp!DÞ2=4kDp!T:

The rate of tetramer formation therefore depends onlyon kD!Dp and on the ratio of ðkDp!DÞ

2=kDp!T, as observed.The values obtained for these parameters were notaffected by inclusion of a back rate kT!Dp in the simu-lations. The discounting of this parameter was furtherjustified by the lack of any significant depletion of [T]over the fitted data ranges, as only initial data pointswere included in the fits to the reactions occurring athigh guanidine hydrochloride concentrations.

Equation (2) yields the expected limiting cases:At high protein concentration:

d½T�=dt ¼ kD!Dp ½D�

and at low concentration:

d½T�=dt ¼ kDp!TðkD!Dp=kDp!DÞ2½D�2

with the crossover between these regimes occurring for:

½D� , ðkDp!DÞ2=ð4kD!DpkDp!TÞ

Transmission electron microscopy

The end product of the reactions was either adsorbeddirectly or, after washing with a solution of 1%(w/v)sodium dodecyl sulfate (SDS) over a 100 kDa cut-offfilter, on glow discharged carbon-coated copper gridsand negative stained with 0.8% (w/v) uranyl formate.Micrographs were recorded at a nominal magnificationof 50,000 £ on a Philips CM100 electron microscopeoperating at 100 kV.

Congo red binding

Both the spectral shift in the absorbance spectrum ofCongo red (CR) in the presence of amyloid in solution78

and the golden green birefringence of stained amyloidaggregates observed under polarizing light were usedto characterize cC multimers formed during the courseof the reaction. Absorbance spectra of solutions of100 mg ml21 protein in the presence of CR at 2 mM,10 mM and 20 mM were recorded and the shift from403 nm (unbound CR) to 541 nm (bound CR) wasevaluated using insulin fibers as a control. These aggre-

gate-CR samples were then placed onto a microscopeslide under a cover slip and screened using a cross-polarizer.

Molecular modelling

Models of potential forms of cystatin tetramers wereconstructed using Insight II and XPLOR (Accelrys, Inc).Dimer units of hCC55 were placed in appropriatepositions according to potential domain-swapping inter-faces. The chain identities were changed as appropriateand the new interfaces optimized manually beforeenergy minimization.

Acknowledgements

We are grateful to Professor Ennes Auerswaldfor providing us with the expression system forcC, the National Centre for MacromolecularThermodynamics, Sutton Bonington, UK forhydrodynamic measurements, and the mass spec-trometry facility at the Department of Chemistry,University of Bristol, UK. We are also indebted toDr Christopher Walters for useful discussions. Wethank Accelrys for their continued support for theprograms FELIX and INSIGHT, and The WellcomeTrust, EPSRC and the Lister Institute for PreventiveMedicine for financial support. R.A.S. is a RoyalSociety University Research Fellow.

References

1. Glenner, G. G. (1980). Amyloid deposits andamyloidosis. New Engl. J. Med. 302, 1283–1292.

2. Prusiner, S. B. (1991). Molecular-biology of priondiseases. Science, 252, 1515–1522.

3. Tan, S. Y. & Pepys, M. B. (1994). Amyloidosis.Histopathology, 25, 403–414.

4. Koo, E. H., Lansbury, P. T. & Kelly, J. W. (1999).Amyloid diseases: abnormal protein aggregation inneurodegeneration. Proc. Natl Acad. Sci. USA, 96,9989–9990.

5. Rochet, J. C. & Lansbury, P. T. (2000). Amyloid fibril-logenesis: themes and variations. Curr. Opin. Struct.Biol. 10, 60–68.

6. Sipe, J. D. & Cohen, A. S. (2000). History of theamyloid fibril. J. Struct. Biol. 130, 88–98.

7. Collinge, J. (2001). Prion diseases of humans and ani-mals: their causes and molecular basis. Annu. Rev.Neurosci. 24, 519–550.

8. Blake, C. & Serpell, L. (1996). Synchrotron X-raystudies suggest that the core of the transthyretinamyloid fibril is a continuous beta-sheet helix.Structure, 4, 989–998.

9. Sunde, M., Serpell, L. C., Bartlam, M., Fraser, P. E.,Pepys, M. B. & Blake, C. C. F. (1997). Common corestructure of amyloid fibrils by synchrotron X-raydiffraction. J. Mol. Biol. 273, 729–739.

10. Jimenez, J. L., Guijarro, J. L., Orlova, E., Zurdo, J.,Dobson, C. M., Sunde, M. & Saibil, H. R. (1999).Cryo-electron microscopy structure of an SH3amyloid fibril and model of the molecular packing.EMBO J. 18, 815–821.


11. Jimenez, J. L., Nettleton, E. J., Bouchard, M.,Robinson, C. V., Dobson, C. M. & Saibil, H. R.(2002). The protofilament structure of insulinamyloid fibrils. Proc. Natl Acad. Sci. USA, 99,9196–9201.

12. Serpell, L. C. & Smith, J. M. (2000). Direct visualiza-tion of the beta-sheet structure of syntheticAlzheimer’s amyloid. J. Mol. Biol. 299, 225–231.

13. Balbach, J. J., Petkova, A. T., Oyler, N. A., Antzutkin,O. N., Gordon, D. J., Meredith, S. C. & Tycko, R.(2002). Supramolecular structure in full-lengthAlzheimer’s beta-amyloid fibrils: evidence for aparallel beta-sheet organization from solid-statenuclear magnetic resonance. Biophys. J. 83,1205–1216.

14. Wetzel, R. (2002). Ideas for amyloid fibril structure.Structure, 10, 1031–1036.

15. Colon, W. & Kelly, J. W. (1992). Partial denaturationof transthyretin is sufficient for amyloid fibril for-mation in vitro. Biochemistry, 31, 8654–8660.

16. Huang, T. H. J., Yang, D. S., Fraser, P. E. &Chakrabartty, A. (2000). Alternate aggregation path-ways of the Alzheimer beta-amyloid peptide—an invitro model of preamyloid. J. Biol. Chem. 275,36436–36440.

17. Kad, N. M., Thomson, N. H., Smith, D. P., Smith,D. A. & Radford, S. E. (2001). beta(2)-microglobulinand its deamidated variant, N17D form amyloidfibrils with a range of morphologies in vitro. J. Mol.Biol. 313, 559–571.

18. Guijarro, J. I., Sunde, M., Jones, J. A., Campbell, I. D.& Dobson, C. M. (1998). Amyloid fibril formation byan SH3 domain. Proc. Natl Acad. Sci. USA, 95,4224–4228.

19. Chiti, F., Webster, P., Taddei, N., Clark, A., Stefani,M., Ramponi, G. & Dobson, C. M. (1999). Designingconditions for in vitro formation of amyloid proto-filaments and fibrils. Proc. Natl Acad. Sci. USA, 96,3590–3594.

20. Morozova-Roche, L. A., Zurdo, J., Spencer, A.,Noppe, W., Receveur, V., Archer, D. B. et al. (2000).Amyloid fibril formation and seeding by wild-typehuman lysozyme and its disease-related mutationalvariants. J. Struct. Biol. 130, 339–351.

21. Modler, A. J., Gast, K., Lutsch, G. & Damaschun, G.(2003). Assembly of amyloid protofibrils via criticaloligomers—a novel pathway of amyloid formation.J. Mol. Biol. 325, 135–148.

22. Griffith, J. S. (1967). Self-replication and Scrapie.Nature, 215, 1043–1044.

23. Prusiner, S. B. (1982). Novel proteinaceous infectiousparticles cause scrapie. Science, 216, 136–144.

24. Jarrett, J. T. & Lansbury, P. T. (1993). Seeding one-dimensional crystallization of amyloid—a patho-genic mechanism in Alzheimer’s disease andscrapie. Cell, 73, 1055–1058.

25. Serio, T. R., Cashikar, A. G., Kowal, A. S., Sawicki,G. J., Moslehi, J. J., Serpell, L. et al. (2000). Nucleatedconformational conversion and the replication ofconformational information by a prion determinant.Science, 289, 1317–1321.

26. Kelly, J. W. (2000). Mechanisms of amyloidogenesis.Nature Struct. Biol. 7, 824–826.

27. Nielsen, L., Khurana, R., Coats, A., Frokjaer, S.,Brange, J., Vyas, S. et al. (2001). Effect of environ-mental factors on the kinetics of insulin fibril for-mation: elucidation of the molecular mechanism.Biochemistry, 40, 6036–6046.

28. Chiti, F., Taddei, N., Baroni, F., Capanni, C., Stefani,

M., Ramponi, G. & Dobson, C. M. (2002). Kinetic par-titioning of protein folding and aggregation. NatureStruct. Biol. 9, 137–143.

29. Lai, Z. H., Colon, W. & Kelly, J. W. (1996). The acid-mediated denaturation pathway of transthyretinyields a conformational intermediate that can self-assemble into amyloid. Biochemistry, 35, 6470–6482.

30. Booth, D. R., Sunde, M., Bellotti, V., Robinson, C. V.,Hutchinson, W. L., Fraser, P. E. et al. (1997).Instability, unfolding and aggregation of humanlysozyme variants underlying amyloid fibrillo-genesis. Nature, 385, 787–793.

31. Kelly, J. W. (1998). The alternative conformations ofamyloidogenic proteins and their multi-stepassembly pathways. Curr. Opin. Struct. Biol. 8,101–106.

32. Jackson, G. S., Hosszu, L. L. P., Power, A., Hill, A. F.,Kenney, J., Saibil, H. et al. (1999). Reversible conver-sion of monomeric human prion protein betweennative and fibrillogenic conformations. Science, 283,1935–1937.

33. McParland, V. J., Kad, N. M., Kalverda, A. P., Brown,A., Kirwin-Jones, P., Hunter, M. G. et al. (2000). Par-tially unfolded states of beta(2)-microglobulin andamyloid formation in vitro. Biochemistry, 39,8735–8746.

34. Kirkitadze, M. D., Condron, M. M. & Teplow, D. B.(2001). Identification and characterization of keykinetic intermediates in amyloid beta-protein fibrillo-genesis. J. Mol. Biol. 312, 1103–1119.

35. Abrahamson, M. & Grubb, A. (1994). Increased bodytemperature accelerates aggregation of the Leu-68 ! Gln mutant cystatin C, the amyloid formingprotein in hereditary cystatin C amyloid Angiopathy.Proc. Natl Acad. Sci. USA, 91, 1416–1420.

36. Gerhartz, B., Ekiel, I. & Abrahamson, M. (1998). Twostable unfolding intermediates of the disease-causingL68Q variant of human cystatin C. Biochemistry, 37,17309–17317.

37. Staniforth, R. A., Giannini, S., Higgins, L. D., Conroy,M. J., Hounslow, A. M., Jerala, R. et al. (2001). Three-dimensional domain swapping in the folded andmolten-globule states of cystatins, an amyloid-form-ing structural superfamily. EMBO J. 20, 4774–4781.

38. Barrett, A. J. (1987). The cystatins—a new class ofpeptidase inhibitors. Trends Biochem. Sci. 12, 193–196.

39. Pennacchio, L. A., Lehesjoki, A. E., Stone, N. E.,Willour, V. L., Virtaneva, K., Miao, J. M. et al. (1996).Mutations in the gene encoding cystatin B in pro-gressive myoclonus epilepsy (EPM1). Science, 271,1731–1734.

40. Barrett, A. J., Davies, M. E. & Grubb, A. (1984). Theplace of human g-trace (cystatin C) amongst thecysteine proteinase inhibitors. Biochem. Biophys. Res.Commun. 120, 631–636.

41. Greenberg, S. M. (1998). Cerebral amyloid angio-pathy. Neurology, 51, 690–694.

42. Labovitz, D. L. & Sacco, R. L. (2001). Intracerebralhemorrhage: update. Curr. Opin. Neurol. 14, 103–108.

43. Bornebroek, M., Haan, J., Maat-Schieman, M. L. C.,Van Duinen, S. G. & Roos, R. A. C. (1996). Hereditarycerebral hemorrhage with amyloidosis-Dutch type(HCHWA-D). 1. A review of clinical, radiologic andgenetic aspects. Brain Pathol. 6, 111–114.

44. Castano, E. M., Prelli, F., Soto, C., Beavis, R.,Matsubara, E., Shoji, M. & Frangione, B. (1996).The length of amyloid-beta in hereditarycerebral hemorrhage with amyloidosis, Dutchtype—implications for the role of amyloid-beta 1-42


in Alzheimer’s disease. J. Biol. Chem. 271,32185–32191.

45. Maat-Schieman, M. L., van Duinen, S. G.,Bornebroek, M., Haan, J. & Roos, R. A. (1996).Hereditary cerebral hemorrhage with amyloidosis-Dutch type (HCHWA-D). 2. A review of histo-pathological aspects. Brain Pathol. 6, 115–120.

46. Wei, L. H., Berman, Y., Castano, E. M., Cadene, M.,Beavis, R. C., Devi, L. & Levy, E. (1998). Instabilityof the amyloidogenic cystatin C variant of hereditarycerebral hemorrhage with amyloidosis, Icelandictype. J. Biol. Chem. 273, 11806–11814.

47. Crawford, F. C., Freeman, M. J., Schinka, J. A.,Abdullah, L. I., Gold, M., Hartman, R. et al. (2000).A polymorphism in the cystatin C gene is a novelrisk factor for late-onset Alzheimer’s disease.Neurology, 55, 763–768.

48. Levy, E., Sastre, M., Kumar, A., Gallo, G., Piccardo, P.,Ghetti, B. & Tagliavini, F. (2001). Codeposition ofcystatin C with amyloid-beta protein in the brain ofAlzheimer disease patients. J. Neuropathol. Exp.Neurol. 60, 94–104.

49. Cohen, D. H., Feiner, H., Jensson, O. & Frangione, B.(1983). Amyloid fibril in hereditary cerebralhemorrhage with amyloidosis (HCHWA) is relatedto the gastroentero-pancreatic, neuroendocrineprotein, g-trace. J. Expt. Med. 158, 623–628.

50. Jensson, O., Gudmundsson, G., Arnason, A.,Blondal, H., Petursdottir, I., Thorsteinsson, L. et al.(1987). Hereditary cystatin C (g-trace) amyloidangiopathy of the CNS causing cerebral hemorrhage.Acta Neurol. Scand. 76, 102–114.

51. Abrahamson, M. (1996). Molecular basis foramyloidosis related to hereditary brain hemorrhage.Scandin. J. Clin. Lab. Invest. 56, 47–56. Suppl. 226.

52. Ekiel, I. & Abrahamson, M. (1996). Folding-relateddimerization of human cystatin C. J. Biol. Chem. 271,1314–1321.

53. Zerovnik, E., Jerala, R., KroonZitko, L., Turk, V. &Lohner, K. (1997). Characterization of the equi-librium intermediates in acid denaturation ofhuman stefin B. Eur. J. Biochem. 245, 364–372.

54. Jerala, R. & Zerovnik, E. (1999). Accessing the globalminimum conformation of stefin A dimer by anneal-ing under partially denaturing conditions. J. Mol.Biol. 291, 1079–1089.

55. Janowski, R., Kozak, M., Jankowska, E., Grzonka, Z.,Grubb, A., Abrahamson, M. & Jaskolski, M. (2001).Human cystatin C, an amyloidogenic protein,dimerizes through three-dimensional domainswapping. Nature Struct. Biol. 8, 316–320.

56. Schlunegger, M. P., Bennett, M. J. & Eisenberg, D.(1997). Oligomer formation by 3D domain swapping:a model for protein assembly and misassembly.Advan. Protein Chem. 50, 61–122.

57. Cohen, F. E. & Prusiner, S. B. (1998). Pathologic con-formations of prion proteins. Annu. Rev. Biochem. 67,793–819.

58. Liu, Y., Gotte, G., Libonati, M. & Eisenberg, D. (2001).A domain-swapped RNase A dimer with impli-cations for amyloid formation. Nature Struct. Biol. 8,211–214.

59. Ding, F., Dokholyan, N. V., Buldyrev, S. V., Stanley,H. E. & Shakhnovich, E. I. (2002). Moleculardynamics simulation of the SH3 domain aggregationsuggests a generic amyloidogenesis mechanism.J. Mol. Biol. 324, 851–857.

60. Knaus, K. J., Morillas, M., Sweietnicki, W., Malone,M., Surewicz, W. K. & Yee, V. C. (2001). Crystal struc-

ture of the human prion protein reveals a mechanismfor oligomerization. Nature Struct. Biol. 8, 770–774.

61. Silow, M., Tan, Y. J., Fersht, A. R. & Oliveberg, M.(1999). Formation of short-lived protein aggregatesdirectly from the coil in two-state folding.Biochemistry, 38, 13006–13012.

62. Ekiel, I., Abrahamson, M., Fulton, D. B., Lindahl, P.,Storer, A. C., Levadoux, W. et al. (1997). NMRstructural studies of human cystatin C dimers andmonomers J. Mol. Biol. 271, 266–277.

63. Hayes, M. V., Sessions, R. B., Brady, R. L. & Clarke,A. R. (1999). Engineered assembly of intertwinedoligomers of an immunoglobulin chain. J. Mol. Biol.285, 1857–1867.

64. Libonati, M., Bertoldi, M. & Sorrentino, S. (1996). Theactivity on double-stranded RNA of aggregates ofribonuclease A higher than dimers increases as afunction of the size of the aggregates. Biochem. J.318, 287–290.

65. Dieckmann, T., Mitschang, L., Hofmann, M., Kos, J.,Turk, V., Auerswald, E. A. et al. (1993). The structuresof native phosphorylated chicken cystatin and of arecombinant unphosphorylated in solution. J. Mol.Biol. 234, 1048–1059.

66. Engh, R. A., Dieckmann, T., Bode, W., Auerswald,E. A., Turk, V., Huber, R. & Oschkinat, H. (1993).Conformational variability of chicken cystatin: com-parison of structures determined by X-ray diffractionand NMR spectroscopy. J. Mol. Biol. 234, 1060–1069.

67. Parker, M. J., Lorch, M., Sessions, R. B. & Clarke, A. R.(1998). Thermodynamic properties of transient inter-mediates and transition states in the folding of twocontrasting protein structures. Biochemistry, 37,2538–2545.

68. Zurdo, J., Guijarro, J. I., Jimenez, J. L., Saibil, H. R. &Dobson, C. R. (2001). Dependence on solution con-ditions of aggregation and amyloid formation by anSH3 domain. J. Mol. Biol. 311, 325–340.

69. Conway, K. A., Harper, J. D. & Lansbury, P. T., Jr(2000). Fibrils formed in vitro from a-synuclein andtwo mutant forms linked to Parkinson’s disease aretypical amyloid. Biochemistry, 39, 2552–2563.

70. Staniforth, R. A., Dean, J. L. E., Zhong, Q., Clarke,A. R., Zerovnik, E. & Waltho, J. P. (2000). The majortransition state in protein folding need not involvethe immobilization of side chains. Proc. Natl Acad.Sci. USA, 97, 5790–5795.

71. Newcomer, M. E. (2001). Trading places. NatureStruct. Biol. 8, 282–284.

72. Martin, J. R., Craven, C. J., Jerala, R., Kroon-Zitko, L.,Zerovnik, E., Turk, V. & Waltho, J. P. (1995). Thethree-dimensional solution structure of human stefinA. J. Mol. Biol. 246, 331–343.

73. Calero, M., Pawlick, M., Soto, C., Castano, E. M.,Sigurdsson, E. M., Kumar, A. et al. (2001). Distinctproperties of wild-type and the amyloidogenichuman cystatin C variant of hereditary cerebralhemorrhage with amyloidosis, Icelandic type.J. Neurochem. 77, 628–637.

74. Zerovnik, E., Pompe-Novak, M., Skarabot, M.,Ravnikar, M., Musevi, I. & Turk, V. (2001). Humanstefin B readily forms amyloid fibrils in vitro. Biochim.Biophys. Acta, 1594, 1–5.

75. Baskakov, I. V., Legname, G., Baldwin, M. A.,Prusiner, S. B. & Cohen, F. E. (2002). Pathwaycomplexity of prion protein assembly into amyloid.J. Biol. Chem. 277, 21140–21148.

76. Ding, T. T., Lee, S.-J., Rochet, J.-C. & Lansbury, P. T.,Jr (2002). Annular a-synuclein protofibrils are


produced when spherical protofibrils are incubatedin solution or bound to brain-derived membranes.Biochemistry, 41, 10209–10217.

77. Anastasi, A., Brown, M. A., Kembhavi, A. A.,Nicklin, M. J. H., Sayers, C. A., Sunter, D. C. &Barrett, A. J. (1983). Cystatin, a protein inhibitor of

cysteine proteinases—improved purification fromegg white. Biochem. J. 211, 129–138.

78. Klunk, W. E., Jacob, R. F. & Mason, R. P. (1999).Quantifying amyloid by Congo red spectral shiftassay. Methods Enzymol. 309, 285–305.

Edited by P. T. Lansbury Jr

(Received 13 January 2003; received in revised form 12 August 2003; accepted 1 December 2003)


cystatin forms a tetramer through structural rearrangement of domain-swapped dimers prior to...

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