- Email: [email protected]
Machinery of protein folding and unfolding
231
Machinery of protein folding and unfolding Xiaodong Zhang*, Fabienne Beuron and Paul S Freemont† During the past two years, a large amount of biochemical, biophysical and low- to high-resolution structural data have provided mechanistic insights into the machinery of protein folding and unfolding. It has emerged that dual functionality in terms of folding and unfolding might exist for some systems. The majority of folding/unfolding machines adopt oligomeric ring structures in a cooperative fashion and utilise the conformational changes induced by ATP binding/hydrolysis for their specific functions. Addresses Centre for Structural Biology, Department of Biological Sciences, Imperial College of Science, Technology and Medicine, Flowers Building, South Kensington, London SW7 2AZ, UK *e-mail: [email protected] †e-mail: [email protected] Current Opinion in Structural Biology 2002, 12:231–238 0959-440X/02/$ — see front matter © 2002 Elsevier Science Ltd. All rights reserved. Abbreviations AAA+ extended family of ATPase associated with a variety of cellular activities CCT cytosolic chaperonin containing the peptide tcp1 (tail-less complex polypeptide 1) Clp caseinolytic protease EM electron microscopy Hsl heat shock locus Hsp heat shock protein NSF N-ethylmaleimide-sensitive factor PDB Protein Data Bank TPR tetratricopeptide repeat
Introduction All biological activities and functions require proteins and multiprotein complexes. Proteins must therefore be able to fold into appropriate structures and assemble into specific complexes to carry out their biological activities. Misfolded proteins or misassembled complexes can malfunction and such events can be detrimental to all living organisms, including humans [1]. Successful protein folding is a multistep process that results in a protein structure often representing the lowest free energy state. To achieve this process in a cellular environment, which predominantly favours aggregation and nonproductive folding, the assistance of protein complexes called chaperones is required. Chaperones are crucial to maintaining the native protein conformation and preventing nonspecific aggregation, whereas the chaperonin proteins directly assist the folding process. To maintain appropriate concentration levels, properly folded proteins and their complexes need to be tightly regulated. Following biological activity, large complexes need to be disassembled to either dispose of or recycle particular components for further activity. One of the most efficient ways to reduce protein levels is via proteolysis. A
number of molecular machines have evolved to perform both the unfolding (‘unfoldase’) and proteolysis activities. Among these more specialised chaperone-assisted proteases are HslUV (heat shock locus UV), FtsH (a membranebound and ATP-dependent metalloprotease), ClpAP (caseinolytic protease AP) and ClpXP. The disassembly of oligomeric protein complexes into their constituent folded or partially folded parts requires a different class of machine. ‘Disassembler’ proteins include the ATPases NSF (N-ethylmaleimide-sensitive factor) and p97, both of which recycle essential components during vesicle membrane fusion [2]. Interestingly, many of the unfoldases and disassemblers belong to the large AAA+ (ATPase associated with a variety of cellular activities) family of ATPase proteins [3], indicating a possible common molecular mechanism that is applied to different biological functions. One general feature among all of the unfolding/folding machines is their dependence on successive ATP binding and hydrolysis cycles for activity. In this review, we will focus on recent structural and biochemical advances concerning some of these proteins, with the aim of summarising any common structural or mechanistic features that are emerging.
Mechanism of protein folding — chaperonins and chaperones Molecular chaperones comprise several highly conserved families of related proteins that are crucial for the maintenance of native protein conformation [4–6]. Chaperones are subdivided into different classes, as defined by their size, cellular distribution and function [7]. The chaperonins are large oligomeric ring-like proteins divided into two subfamilies: group I comprising the GroEL–GroES system and group II the thermosome [8]. Group I chaperonins — GroEL–GroES
A comprehensive structural picture of chaperonin-assisted protein folding has been obtained from extensive studies of the Escherichia coli GroEL–GroES system. From both cryo-electron microscopy (cryo-EM) and high-resolution crystal structures, mechanistic insights have been obtained that describe the folding cycle in terms of GroEL–GroES conformation and nucleotide-binding state [9–12]. GroEL consists of two seven-subunit ring structures (back to back), with each subunit formed of three domains (equatorial, intermediate and apical). Upon the positive cooperative binding of GroES and ATP to one ring, conformational changes result in a large closed cavity into which unfolded peptides are displaced. ATP hydrolysis in the first ring and binding of ATP to the second ring results in the release of both the folded or partially unfolded substrate and GroES, with both rings working out of phase. Currently, little is known about the conformation of GroEL substrates during the folding cycle. This past year
232
Macromolecular assemblages
has seen several biochemical studies aimed at addressing these issues. Hayer-Hartl and co-workers [13••], using a rapid inhibition assay and GroEL mutants that prevent substrate re-association, showed that the GroEL–GroES cage is essential for folding without aggregation. They concluded that confinement of unfolded proteins in the folding cavity increases the flux of intermediates toward the native folded state. Horwich and colleagues [14••], also using GroEL mutants defective in substrate binding, showed that GroEL substrates bind via multivalent interactions, which could provide a mechanism for further unfolding of bound misfolded substrates. Studies measuring protein compactness upon GroEL and ATP binding [15,16] suggest a ‘substrate stretching’ model. However, deuterium exchange experiments studying bound GroEL substrates argue against a forced mechanical unfolding mechanism as part of the intermediary folding pathway [17••]. A molecular dynamics study of GroEL by Karplus and colleagues [18] has provided some details of intermediate conformational changes during the GroEL allosteric cycle. They conclude that the individual GroEL monomers encode the necessary allosteric motions for heptamer cooperativity. Fersht and colleagues [19,20] suggest that, through evolution, GroEL has acquired a wide range of activities and does not rely on a single mechanism for all substrates. Indeed, it has been shown that GroEL–GroES can mediate the iterative binding of nonproductive quaternary complexes to promote dissociation and subunit exchange between heterodimers containing a misfolded monomer [21]. However, it still remains unresolved as to how GroEL–GroES could rescue misfolded non-native intermediates without exerting direct mechanical unfolding upon these intermediates. Group II chaperonins — thermosome and CCT
The archaebacterial thermosome (composed of two subunits: α and β) and eukaryotic CCT (cytosolic chaperonin containing the peptide tcp1 [tail-less complex polypeptide 1] comprising eight different but related subunits) belong to the group II subfamily of chaperonins. Group II subunits are organised similarly to GroEL, with two large domains (equatorial and apical) linked by hinge regions to a smaller (intermediate) domain. The crystal structure of the thermosome with and without nucleotide reveals a closed conformation with a built-in ‘lid’ domain that might substitute for the co-chaperonin GroES [22]. Recent structural information, primarily from cryo-EM, on two archaebacterial thermosomes and on the chaperonin CCT has provided further mechanistic insights into chaperonin function. By fitting the atomic coordinates of the thermosome comprising only α subunits into cryo-EM maps, Schoehn et al. [23•] have analysed the domain movements that occur between the co-existing open, asymmetric and closed conformations. The apical domain is highly flexible. These three conformations, previously described for the eukaryotic chaperonin CCT [24], most probably represent various
states in the ATPase cycle, which leads to encapsulation of the folding substrate. Considerable structural information from cryo-EM has recently been obtained that describes the conformational changes during CCT-facilitated folding of actin and tubulin [25,26,27••,28]. The authors reconcile their data in the following model: partially folded tubulin substrate proteins bind to apo-CCT at the apical domain of one ring. ATP binding induces the closure of the cavity in both rings. The ADP-Pi conformational state of the CCT–β tubulin complex exhibits a closed symmetric conformation, similar to the ATP conformation, with the substrate remaining at the apical domain. It is hypothesised that release of the nucleotide and subsequent expulsion of the folded substrate might occur upon movement of the apical domains or might involve other cellular cofactors. Specific CCT subunits are characterised by their different intrinsic affinity for ATP. Hence, through sequential ATP binding and hydrolysis, CCT assists partially folded actin and tubulin substrates to reach their final native state, with their nucleotide-binding pocket in place. Chaperones — Hsp90, Hsp70 (DnaK) and prefoldin
Among the chaperone group of proteins, eukaryotic Hsp90 (heat shock protein 90) and Hsp70 (and its prokaryotic homologue DnaK) have received much attention recently. Hsp90 is involved in the folding and assembly of proteins with roles in signal transduction, cell regulation and transcriptional activation [29,30]. Hsp90 operates as part of a multichaperone complex that includes numerous co-chaperones such as Hsp70 (which provides substrates for Hsp90-mediated folding), p23 and Hop (which acts as a scaffold for Hsp90 and Hsp70). Although much is known about the biochemical role of Hsp90, little is known about its inherent ATPase activity and the conformational changes induced by nucleotide binding and hydrolysis. Recent studies have tried to resolve some of these issues. ATP binding to Hsp90 has been shown to induce dimerisation of the N-terminal domain and substrate release involves ATP hydrolysis, an activity that is enhanced by the co-chaperone p23 [31,32]. This has led to the proposal that Hsp90 acts via a molecular clamp mechanism, similar to DNA gyrase and MutL, whereby opening and closing of a substrate-binding site is mediated by transient N-terminal dimerisation [32]. Kinetic analysis now shows that ATP hydrolysis is the rate-limiting step and that the C-terminal region of Hsp90 is essential for both ATP binding and enhancing hydrolysis rates [33]. The crystal structure of two TPR (tetratricopeptide repeat) domains of the Hop adaptor protein bound to target Hsp90/Hsp70-specific peptides [34••] has provided details of the assembly of a Hsp90–Hsp70 complex. However, a number of mechanistic and structural issues still remain unresolved, including a full structural description of the Hsp90 ATPase/substratebinding cycle. Several recent studies on the prokaryotic Hsp70 chaperone DnaK have been aimed at understanding the relationship
Machinery of protein folding and unfolding Zhang, Beuron and Freemont
between substrate affinity, nucleotide binding and co-chaperone-induced ATP hydrolysis. ATP binding to the N-terminal domain induces conformational changes that open the cavity to allow substrate binding and release. Subsequent ATP hydrolysis, induced by co-chaperone DnaJ (Hsp40), results in a closed state. Nucleotide exchange is a key element of substrate release and thus is critical to Hsp70 chaperone function. Recent studies identified structural elements within the DnaK family that define the interaction with the nucleotide exchange factors Bag-1 and GrpE [35,36]. Structures of Bag-1 bound to the ATPase domain of Hsc70 (the constitutively expressed cytosolic isoform of Hsp70) now provide direct insights into Hsp70 nucleotide exchange [37,38]. Hsp70 can be functionally substituted by the ubiquitous prefoldin (GimC) chaperone. Prefoldin is a heterohexamer that can interact with nascent polypeptides in vitro. The crystal structure of prefoldin reveals a unique quaternary structure that is unlike that of any other chaperone and it therefore forms a novel class. The structure comprises a double β-barrel assembly linking coiled coils with protruding flexible ‘tentacles’ containing hydrophobic patches required for peptide binding [39]. Unlike Hsp70 and the chaperonins, prefoldin is a novel class of chaperone that does not require ATP. It uses multiple interaction sites for substrate binding to shield non-native polypeptide from aggregation before passing it to other chaperones for completion of folding.
Mechanism of protein unfolding — chaperone-assisted protein degradation In eukaryotes, protein degradation is mainly achieved by a multiprotein complex, the 26S proteasome. In prokaryotes, a number of specialised chaperone-assisted proteases are used for a similar function. These include the two-component systems ClpX/ClpP (ClpXP), ClpA/ClpP (ClpAP) and HslU/HslV (HslUV) [40]. Recently, a number of HslU and HslUV structures have become available that provide significant mechanistic understanding of their function. Three different HslUV complexes have now been reported. The complex reported by Bochtler and colleagues [41••] positions the flexible intermediate (I) domain of HslU to interact with HslV. Subsequently, Sousa et al. [42••], using X-ray crystallography and small-angle X-ray solution scattering data, presented an alternative complex, in which a HslV dodecamer in the centre of the complex interacts with one HslU hexamer on each side, with the I domains distal. Cryo-EM 2D projection data at 30 Å resolution are consistent with this latter model [43•]. A third complex structure [44•,45•] also revealed a HslV dodecamer bound to the flat surface of a HslU hexamer, with the I domains distal. However, all three structures clearly identify a sixfold symmetry match between HslU and HslV, ruling out any mechanism of protease activation by symmetry mismatch [41••,42••,44•]. These structural data unambiguously illustrate that, like many other ATPase machines, HslU and HslV assemble
233
into hexameric ring structures that stack coaxially and at least one HslU hexamer is bound to a HslV dodecamer. It is also proposed that HslU binding to HslV could induce conformational changes in HslV to activate its peptidase activity [42••]. ATP binding in HslU has been proposed to enable HslU central pore opening. Furthermore, substrate recognised by the I domain would be first unfolded through I domain relocation, which is induced by HslU ATP hydrolysis, before translocation through the central pore into the protease chamber [42••,46•]. However, considerable dispute exists over which structural configuration represents the active HslUV complex. More crucially, the nature of substrate denaturation using ATP hydrolysis and the correlation between the ATP hydrolysis cycle and HslU pore opening remain unresolved. To understand the mechanism of these crucial steps, structures of HslUV complexed with different substrates at different stages, including substrate recognition, substrate unfolding and translocation, will be required. The Clp/Hsp100 chaperones are also AAA+ ATPases and function as substrate-specificity components of the ATP-dependent Clp proteases and also in preventing the aggregation and disassembly of transient multicomponent complexes involved in transcription, replication and the cell cycle [47]. ClpA, in the presence of ATP, mediates the unfolding of proteins that are bound following the recognition of specific terminal amino acid sequences (the ssrA peptide tag added to nascent proteins when ribosomes stall during translation). In the absence of the associated protease, ClpP, substrate proteins are released from ClpA or ClpX into solution, where they can spontaneously refold or become bound by other components, such as chaperonins or other Hsp100 family members [48–50]. In the absence of high-resolution structures of either ClpX or ClpA, recent biophysical studies have been informative. EM studies have helped to locate the substrate-binding site (positioned axially on the distal surface of the ATPase) and show that the unfolding and translocation of peptide substrates could proceed in discreet steps [43•,51]. It appears that ClpXP, ClpAP and the proteasome unravel folded substrates sequentially from the point of attachment at the degradation signal [52]. In multidomain proteins, independent stable domains are degraded sequentially.
Mechanism of protein complex disassembly – p97 and NSF Protein complexes often need to be disassembled as part of a regulatory system or to recycle certain components. A good example of this latter process is the recycling of the SNARE pairs formed after vesicle membrane fusion, whereby ATP hydrolysis is required for their disassembly. SNARE complexes comprise highly stable membraneattached helical coiled-coil structures that are formed after membrane fusion events [53]. Two homologous AAA+ ATPases, p97 and NSF, have been implicated in
234
Macromolecular assemblages
Figure 1 Structural comparisons. Ribbon representations of crystal structures of the ATPases discussed in this review in complex with ADP, with ADP similarly orientated: (a) ‘disassembler’ p97 D1 (PDB code 1E32); (b) ‘unfoldase’ HslU (PDB code 1G41); (c) group I chaperonin GroEL (PDB code 1AON); (d) group II chaperonin thermosome α subunit (PDB code 1A6E); (e) chaperone Hsp90 (PDB code 1BYQ); (f) chaperone Hsp70 (PDB code 1HJO).
disassembling these SNARE complexes via ATP hydrolysis and in association with specific adaptors [54,55]. The activities and mechanism of NSF in mediating membrane fusion have been well covered in a recent review [2]. Here, we focus on recent structural studies of p97, which reveal the organisation of p97 and its interaction with its cofactor, p47. p97 is a homohexamer and, like NSF, contains two AAA+ domains, D1 and D2. Crystal structures and cryo-EM analysis of p97 now show that its N-terminal (N) domain is located at the periphery of D1 [56••,57•] and that D1 and D2 hexamers stack coaxially [56••]. NMR studies also show that UBX, a ubiquitin-like domain situated at the C terminus of p47, interacts with the N-domain of p97 [58]. Comparison with the structure of NSF D2 bound to AMPPNP [59,60] reveals a possible conformational change upon ATP hydrolysis that could induce protomer–protomer rearrangements. This would also result in a closed central pore in the ADP state. This change would cascade to the relocation of the linker
between N and D1, generating mechanical force for SNARE disassembly via p47 [56••]. Based on the observation from EM that the D1 and D2 rings have different protomer arrangements, and the possible conformational changes between different nucleotide states, a ratchet mechanism for p97 has been proposed. In this model, D1 and D2 exist in different nucleotide states, and ATP hydrolysis in one domain induces nucleotide exchange in the other. It was also speculated that D1 and D2 pack in a tail-to-tail arrangement to accommodate the cage-like feature observed in the EM reconstruction, although alternative packing arrangements are possible. The proposed ratchet mechanism, however, is independent of the exact packing arrangement between D1 and D2. It is notable that GroEL–GroES also utilises two ATPase rings in different nucleotide-bound conformations as part of its activity cycle. A different study, using cryo-EM 2D projection analysis, proposed an alternative model for SNARE disassembly [57•]. In this model, nucleotide-free p97 binds loosely to
Machinery of protein folding and unfolding Zhang, Beuron and Freemont
235
Table 1 ATPase cycle and corresponding functional stages*.
GroEL CCT†
ATP binding
ATP hydrolysis
ADP release
Oligomeric state
i) Allows GroES binding ii) Releases GroES, ADP and substrate in opposite ring Induces closing of cavity
i) Weakens GroES interaction ii) Allows ATP and substrate binding in opposite ring Allosteric substrate folding
Resulting from ATP hydrolysis in opposite ring
Heptamer
Probably requires other cellular Octamer factors Hsp90 i) Induces N-domain dimerisation and i) Opens clamp to allow substrate Unknown Dimer substrate clamping release ii) Promotes p23 binding ii) Dissociates p23 Hsp70 Induces open conformation: low affinity for Induces closed conformation: Requires nucleotide exchange Monomer substrate traps substrates factor Bag-1/GrpE HslU Enables substrate binding and translocation Polypeptide unfolding Allosteric mechanism? Hexamer p97 Substrate binding Disassembly Ratchet mechanism? Hexamer ClpX/ Oligomerisation and substrate binding Substrate unfolding and Unknown Hexamer ClpA translocation *In the GroEL–GroES chaperone system, ATP binding not only step is not yet well characterised. However, an allosteric mechanism promotes GroES binding, but also allows GroES, ADP and has been proposed for the HslU hexamer, whereby ATP hydrolysis in substrate release in the opposite ring. ATP hydrolysis, on the other one subunit would promote ADP release in the next subunit [41]. hand, weakens the interaction with GroES and allows ATP and This mechanism, combined with the open pore state associated with substrate binding to occur in the opposite ring. In Hsp70, ATP ATP binding and substrate translocation, implies that denatured binding would induce an open state, resulting in low-affinity polypeptides would ‘spiral’ through the pore. This could be a general substrate binding, whereas ATP hydrolysis would induce a closed feature of unfoldases. For p97, two models have been proposed. It is state, thereby trapping substrates. A nucleotide exchange factor is possible that both ATP binding and hydrolysis generate required to remove the ADP. For Hsp90, ATP binding locks conformational changes driving SNARE disassembly. It is unclear substrates by clamping the substrate via ATP-induced dimerisation. how ADP is released in this system, though the proposed ratchet In the chaperonin-assisted protease systems (ClpX, HslU, FtsH mechanism would allow ADP release from the opposite ring upon AAA+ domain), ATP binding not only allows substrate binding, but ATP hydrolysis. †The thermosome has been omitted from this table also results in an open pore, which allows denatured peptides to because the various nucleotide states have not been identified pass to the protease chamber. ATP hydrolysis has been implicated unambiguously and the X-ray studies have revealed limited changes as being responsible for polypeptide denaturation. The ADP release upon nucleotide binding or hydrolysis [22].
monomeric p47 and, upon ATP binding, the ordered N-domain generates mechanical force to disassemble SNARE complexes. Given that the D1 and D2 domains are both active ATPases and that they may exist in different nucleotide-bound states, defining the actual p97 nucleotide state becomes nontrivial. Interestingly, both p97 models [56••,57•] propose that the relocation of the N-domain induced by ATP binding/hydrolysis mediates target protein disassembly, as previously suggested for NSF [59,61].
ATP binding/hydrolysis is the driving force for protein folding/unfolding One common element used by many molecular machines is their ability to successively bind and hydrolyse ATP. For motor proteins (kinesin and myosin), small structural changes in the nucleotide-binding site during the ATPase cycle are amplified into larger movements that provide the necessary force and motion [62]. Current structural and functional data now show that the majority of the molecular machines used for protein folding/unfolding also couple ATP binding, ATP hydrolysis and ADP release to appropriate conformational changes and functional states. A representative ATPase structure (monomer) of each of the chaperone systems discussed is shown in Figure 1, with bound ADP oriented similarly. It is notable that, in all of the systems, the nucleotide is bound between subdomains (intradomain binding). This provides a structural explanation for how nucleotide binding and hydrolysis could induce subdomain reorientation, in order to generate mechanical forces. In
certain cases, such as p97 and HslU, the nucleotide is also bound between protomers (interdomain binding) and ATP hydrolysis might require residues from an adjacent protomer, as proposed for p97 [56••]. Furthermore, most of the folding/unfolding systems utilise single or multiple oligomeric ring structures. These oligomeric ATPases allow ATP binding and hydrolysis to be either concerted, to provide a single conformational transition (force) (such as in GroEL, NSF/p97), or allosteric, to allow multiple conformational transitions between protomers (such as in HslU, ClpX and CCT). Two-ring ATPase structures such as GroEL allow further adaptations, including allosteric transitions between the individual rings.
Conclusions An ATP hydrolysis cycle can be divided into three steps: ATP binding, ATP hydrolysis, and ADP and Pi release. Each system uses one or multiple steps to generate the necessary conformational changes and mechanical forces for their functions (summarised in Table 1). A common feature from a number of molecular machines, including unfoldases, disassemblers and certain chaperone systems, is that ATP binding allows substrate release, as well as substrate binding, whereas ATP hydrolysis induces substrate remodelling (unfolding or disassembly). However, the significant amount of structural information available and the diverse mechanistic models proposed demonstrates the complexity of the protein folding/unfolding machinery. Not only does each system adapt different stages of the
236
Macromolecular assemblages
ATPase cycle to actively perform folding/unfolding, but they also utilise cooperativity between different subunits within the machinery. Furthermore, each system might use different mechanisms for different targets, as proposed for GroEL–GroES. It is becoming increasingly apparent, however, that a combination of high- and medium-resolution structures that capture various stages of the functional cycle, together with biochemical and biophysical studies, is required for a deeper understanding of these complex machineries.
Acknowledgements We wish to thank V Pye and I Dreveny for critically reading the manuscript.
References and recommended reading Papers of particular interest, published within the annual period of review, have been highlighted as:
• of special interest •• of outstanding interest
15. Hammarstrom P, Persson M, Owenius R, Lindgren M, Carlsson U: Protein substrate binding induces conformational changes in the chaperonin GroEL. A suggested mechanism for unfoldase activity. J Biol Chem 2000, 275:22832-22838. 16. Hammarstrom P, Persson M, Carlsson U. Protein compactness measured by fluorescence resonance energy transfer. Human carbonic anhydrase II is considerably expanded by the interaction of GroEL. J Biol Chem 2001, 276:21765-21775. 17. ••
Chen J, Walter S, Horwich AL, Smith DL: Folding of malate dehydrogenase inside the GroEL-GroES cavity. Nat Struct Biol 2001, 8:721-728. Using NMR and mass spectrometry methods, the authors found that, when bound to GroEL, a core of malate dehydrogenase (MDH) secondary structure was protected from proton exchange and that, upon ATP and GroES binding, this core was only partially deprotected. Their results are consistent with hydrogen bonds between the sides of the cavity and MDH substrate being broken, as opposed to forced mechanical unfolding. 18. Ma J, Sigler PB, Xu Z, Karplus M: A dynamic model for the allosteric mechanism of GroEL. J Mol Biol 2000, 302:303-313. 19. Chatellier J, Hill F, Fersht AR: From minichaperone to GroEL 2: importance of avidity of the multisite ring structure. J Mol Biol 2000, 304:883-896.
1.
Slavotinek AM, Biesecker LG: Unfolding the role of chaperones and chaperonins in human disease. Trends Genet 2001, 17:528-535.
20. Chatellier J, Hill F, Foster NW, Goloubinoff P, Fersht AR: From minichaperone to GroEL 3: properties of an active single-ring mutant of GroEL. J Mol Biol 2000, 304:897-910.
2.
May AP, Whiteheart SW, Weis WI: Unraveling the mechanism of the vesicle transport ATPase NSF, the N-ethylmaleimide-sensitive factor. J Biol Chem 2001, 276:21991-22004.
3.
Ogura T, Wilkinson AJ: AAA+ superfamily ATPases: common structure-diverse function. Genes Cells 2001, 6:575-597.
21. Wynn RM, Song JL, Chuang DT: GroEL / GroES promote dissociation / reassociation cycles of a heterodimeric intermediate during alpha(2)beta(2) protein assembly. Iterative annealing at the quaternary structure level. J Biol Chem 2000, 275:2786-2794.
4.
Grantcharova V, Alm EJ, Baker D, Horwich AL: Mechanisms of protein folding. Curr Opin Struct Biol 2001, 11:70-82.
5.
Gottesman ME, Hendrickson WA: Protein folding and unfolding by Escherichia coli chaperones and chaperonins. Curr Opin Microbiol 2000, 3:197-202.
6.
Carrascosa JL, Llorca O, Valpuesta JM: Structural comparison of prokaryotic and eukaryotic chaperonins. Micron 2001, 32:43-50.
7.
Ben-Zvi AP, Goloubinoff P: Mechanisms of disaggregation and refolding of stable protein aggregates by molecular chaperones. J Struct Biol 2001, 135:84-93.
8.
Kusmierczyk AR, Martin J: Chaperonins: keeping a lid on folding proteins. FEBS Lett 2001, 21:343-347.
9.
Feltham JL, Gierasch LM: GroEL-substrate interactions: molding the fold, or folding the mold? Cell 2000, 100:193-196.
10. Thirumalai D, Lorimer GH: Chaperonin-mediated protein folding. Annu Rev Biophys Biomol Struct 2001, 30:245-269. 11. Melki R: Nucleotide-dependent conformational changes of the chaperonin containing tcp-1. J Struct Biol 2001, 135:170-175. 12. Steinbacher S, Ditzel L: Nucleotide binding to the thermoplasma thermosome: implications for the functional cycle of group II chaperonins. J Struct Biol 2001, 135:147-156. 13. Brinker A, Pfeifer G, Kerner MJ, Naylor DJ, Hartl FU, Hayer-Hartl M: •• Dual function of protein confinement in chaperonin-assisted protein folding. Cell 2001, 107:223-233. The folding of Rubisco and rhodenase using a modified GroEL mutant structure that blocks the rebinding of non-native proteins was studied. The results showed that folding within the GroEL cage is independent of ATP hydrolysis cycles and significantly faster than in free solution. This argues that the confinement of unfolded proteins within the folding cavity increases the flux of folding intermediates toward the native folded state. 14 ••
Farr GW, Furtak K, Rowland MB, Ranson NA, Saibil HR, Kirchhausen T, Horwich AL: Multivalent binding of nonnative substrate proteins by the chaperonin GroEL. Cell 2000, 100:561-573. This paper describes studies of mutant GroEL complexes binding to three different substrates, namely Rubisco, malate dehydrogenase (MDH) and rhodenase. For Rubisco and MDH, consecutive wild-type GroEL apical domains are preferred for binding. This could be explained by the need for a continuous hydrophobic binding surface or by the requirement of the apical domain to act cooperatively in substrate binding. A multivalent binding model is proposed for GroEL.
22. Ditzel L, Löwe J, Stock D, Stetter KO, Huber H, Huber R, Steinbacher S: Crystal structure of the thermosome, the archaeal chaperonin and homolog of CCT. Cell 1998, 93:125-138. 23. Schoehn G, Hayes M, Cliff M, Clarke AR, Saibil HR: Domain • rotations between open, closed and bullet-shaped forms of the thermosome, an archaeal chaperonin. J Mol Biol 2000, 301:323-332. The native chaperonin thermophilic factor 55 (TF55) is a heat shock protein from the archaea Sulfolobus shibatae; α, β and γ subunits have been identified. The 3D reconstruction of TF55 (ninefold symmetry) described in this report constitutes the first characterisation of three coexisting conformations in solution: open, asymmetric (‘bullet’ shaped) and closed. The observation of a ‘bullet’ asymmetric form for a group II chaperonin reveals a close parallel with the GroEL system. 24. Llorca O, Smyth MG, Carrascosa JL, Willison KR, Radermacher M, Steinbacher S, Valpuesta JM: 3D reconstruction of the ATP-bound form of CCT reveals the asymmetric folding conformation of a type II chaperonin. Nat Struct Biol 1999, 6:639-642. 25. Llorca O, McCormack EA, Hynes G, Grantham J, Cordell J, Carrascosa JL, Willison KR, Fernandez JL, Valpuesta JM: Eukaryotic type II chaperonin CCT interacts with actin through specific subunits. Nature 1999, 402:693-696. 26. Llorca O, Martin-Benito J, Ritco-Vonsovici M, Grantham J, Hynes GM, Willison KR, Carrascosa JL, Valpuesta JM: Eukaryotic chaperonin CCT stabilizes actin and tubulin folding intermediates in open quasi-native conformations. EMBO J 2000, 19:5971-5979. 27. ••
Llorca O, Martin-Benito J, Grantham J, Ritco-Vonsovici M, Willison KR, Carrascosa JL, Valpuesta JM: The ‘sequential allosteric ring’ mechanism in the eukaryotic chaperonin-assisted folding of actin and tubulin. EMBO J 2001, 20:4065-4075. CCT-bound actin is resolved as rod-shaped densities (‘stretched’ actin as opposed to ‘V’-shaped molecules) in which small and large domains are identifiable. The exact binding pattern has been mapped using monoclonal antibodies and 2D averaging. The binding of one molecule of tubulin causes substantial conformational changes in apo-CCT, which adopts a semiclosed conformation [26]. Five specific subunits using two modes of interaction are involved in the binding of tubulin to CCT. Interestingly, the location of the substrate-binding sites at the tips of the apical domains of CCT is different for actin and tubulin. 28. Llorca O, Martin-Benito J, Gomez-Puertas P, Ritco-Vonsovici M, Willison KR, Valpuesta JM: Analysis of the interaction between the eukaryotic chaperonin CCT and its substrates actin and tubulin. J Struct Biol 2001, 135:205-218. 29. Pearl LH, Prodromou C: Structure and in vivo function of Hsp90. Curr Opin Struct Biol 2000, 10:46-51.
Machinery of protein folding and unfolding Zhang, Beuron and Freemont
30. Young JC, Moarefi I, Hartl FU: Hsp90: a specialized but essential protein-folding tool. J Cell Biol 2001, 154:267-273 31. Young JC, Hartl FU: Polypeptide release by Hsp90 involves ATP hydrolysis and is enhanced by the co-chaperone p23. EMBO J 2000, 19:5930-5940. 32. Prodromou C, Panaretou B, Chohan S, Siligardi G, O’Brien R, Ladbury JE, Roe SM, Piper PW, Pearl LH: The ATPase cycle of Hsp90 drives a molecular ‘clamp’ via transient dimerization of the N-terminal domains. EMBO J 2000, 19:4383-4392. 33. Weikl T, Abelmann K, Buchner J: An unstructured C-terminal region of the Hsp90 co-chaperone p23 is important for its chaperone function. J Mol Biol 1999, 293:685-691. 34. Scheufler C, Brinker A, Bourenko G, Pegoraro S, Moroder L, •• Bartunik H, Hartl FU, Moarefi I: Structure of TPR domain-peptide complexes: critical elements in the assembly of the Hasp70-Hasp90 multichaperone machine. Cell 2000, 101:199-210. The crystal structure of two TPR domains of the adaptor protein Hop bound to Hsp90/Hsp70-specific peptides reveals the first molecular insights into the assembly of the Hsp70–Hsp90 multichaperone complex. Interestingly, the Hop TPR–peptide structures show some similarity to 14-3-3–peptide complexes, indicating a common ancestral mode of protein–peptide binding. 35. Brehmer D, Rüdiger S, Gässler CS, Klostermeier D, Packschies L, Reinstein J, Mayer MP, Bukau B: Tuning of chaperone activity of Hsp70 proteins by modulation of nucleotide exchange. Nat Struct Biol 2001, 8:427-432. 36. Han W, Christen P: Mutations in the interdomain linker region of DnaK abolish the chaperone action of the DnaK/DnaJ/GrpE system. FEBS Lett 2001, 497:55-58. 37.
Briknarova K, Takayama S, Brive L, Havert ML, Knee DA, Velasco J, Homma S, Cabezas E, Stuart J, Hoyt DW et al.: Structural analysis of BAG1 cochaperone and its interactions with Hsc70 heat shock protein. Nat Struct Biol 2001, 8:349-352.
237
44. Wang J, Song JJ, Franklin MC, Kamtekar S, Im YJ, Rho SH, Seong IS, • Lee CS, Chung CH, Eom SH: Crystal structures of the HslVU peptidase-ATPase complex reveal an ATP-dependent proteolysis mechanism. Structure 2001, 9:177-184. The authors describe an alternative arrangement of HslUV from E. coli at 3.0 Å (see [41••]), with a HslV dodecamer bound to the flat surface of a HslU hexamer, leaving the I domain of HslU distal. The size of the HslV pore changes between free and complexed HslV, and the authors argue that binding to HslU induces pore size changes in HslV, probably allowing substrate release. 45. Wang J, Song JJ, Seong IS, Franklin MC, Kamtekar S, Eom SH, • Chung CH: Nucleotide-dependent conformational changes in a protease-associated ATPase HslU. Structure 2001, 9:1107-1116. The authors compared all available HslU structures and proposed that nucleotide binding, rather than hydrolysis, induces the largest conformational changes in subdomain arrangement. Furthermore, they propose that different functional states exist during the ATP hydrolysis cycle and that these states correspond to the different stages of protein unfolding and translocation. 46. Song HK, Hartmann C, Ramachandran R, Bochtler M, Behrendt R, • Moroder L, Huber R: Mutational studies on HslU and its docking mode with HslV. Proc Natl Acad Sci USA 2000, 97:14103-14108. These authors presented systematic mutagenesis data and various peptidase activity data to dissect the roles of various regions in HslU. Deletion in the I domain in general had no effect on peptidase activity, although it abolishes the degradation of folded proteins, suggesting that the I domain is involved in substrate recognition and/or unfolding. The authors argue that the peptidase and caseinolytic activities might not require a precise complex between HslU and HslV, and different HslUV complexes might be involved in the degradation of different substrates 47.
Jenal U, Fuchs T: An essential protease involved in bacterial cell-cycle control. EMBO J 1998, 17:5658-5669.
48. Weber-Ban EU, Reid BG, Miranker AD, Horwich AL: Global unfolding of a substrate protein by the Hsp100 chaperone ClpA. Nature 1999, 401:90-93.
38. Sondermann H, Scheufler C, Schneider C, Hohfeld J, Hartl FU, Moarefi I: Structure of a Bag/Hsc70 complex: convergent functional evolution of Hsp70 nucleotide exchange factors. Science 2001, 29:1553-1557.
49. Singh S, Grimaud R, Hoskins J, Wickner S, Maurizi MR: Unfolding and internalization of proteins by the ATP-dependent proteases ClpXP and ClpAP. Proc Natl Acad Sci USA 2000, 97:8898-8903.
39. Siegert R, Leroux MR, Scheufler C, Hartl FU, Moarefi I: Structure of the molecular chaperone prefoldin: unique interaction of multiple coiled coil tentacles with unfolded proteins. Cell 2000, 102:621-632.
50. Kim YI, Burton RE, Burton BM, Sauer RT, Baker TA: Dynamics of substrate denaturation and translocation by the ClpXP degradation machine. Mol Cell 2000, 5:639-648.
40. Zwickl P, Baumeister W, Steven A: Dis-assembly lines: the proteasome and related ATPase-assisted proteases. Curr Opin Struct Biol 2000, 10:242-250.
51. Ortega J, Singh S, Ishikawa T, Maurizi MR, Steven AC: Visualization of substrate binding and translocation by the ATP-dependent protease, ClpXP. Mol Cell 2000, 6:1515-1521.
41. Bochtler M, Hartmann C, Song HK, Bourenkov GP, Bartunik HD, •• Huber R: The structures of HsIU and the ATP-dependent protease HsIU-HsIV. Nature 2000, 403:800-805. Crystal structures of HslU in different nucleotide-bound states and of the HslUV complex from E. coli were presented. The structure of the HslUV complex is the first for a complete set of components of an ATP-dependent protease. It also clarified the sixfold symmetry match between HslU and HslV, ruling out a mechanism of protease activation by symmetry mismatch. The complex structure reveals a dodecamer of HslV bound to the I domains of a HslU hexamer on both sides. The various HslU structures with different nucleotide bound allow the authors to speculate an allosteric mechanism for HslU. These structures also establish that HslU belongs to the AAA+ superfamily.
52. Lee C, Schwartz MP, Prakash S, Iwakura M, Matouschek A: ATP-dependent proteases degrade their substrates by processively unraveling them from the degradation signal. Mol Cell 2001, 7:627-637.
42. Sousa MC, Trame CB, Tsuruta H, Wilbanks SM, Reddy VS, •• McKay DB: Crystal and solution structures of an HslUV protease-chaperone complex. Cell 2000, 103:633-643. The authors report the structures of a HslUV complex from H. influenzae at 3.4 Å in the presence of ATP and of ADP-bound HslU at 2.3 Å. Their data show an alternative HslUV complex from that of E. coli [41••], in that a HslV dodecamer in the centre of the complex interacts with the flat part of HslU, while the I domain of HslU is distal. Small-angle X-ray scattering experiments and peptidase activity assays were used to support the crystal structure being an active complex. 43. Ishikawa T, Maurizi MR, Belnap D, Steven AC: Docking of • components in a bacterial complex. Nature 2000, 408:667-668. This method allowed the direct quantification of the protein densities inside ClpP in a stalled ClpAP complex. A mass of 41 kDa (1.5 RepA subunits) was calculated to be able to fit inside ClpP. Presumably, unfolding is driven by multivalent contacts between the surrounding ClpA subunits and the substrate protein. ATP binding or hydrolysis probably drives a conformational change in ClpA that pulls the protein structure apart, in an action analogous to that of GroEL. However, at the resolution of this work (30 Å), no significant conformational changes were observed in ClpA upon ATP hydrolysis.
53. Hay JC: SNARE complex structure and function. Exp Cell Res 2001, 271:10-21. 54. Sollner T, Bennett MK, Whiteheart SW, Scheller RH, Rothman JE: A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell 1993, 75:409-418. 55. Rabouille C, Kondo H, Newman R, Hui N, Freemont P, Warren G: Syntaxin 5 is a common component of the NSF- and p97-mediated reassembly pathways of Golgi cisternae from mitotic Golgi fragments in vitro. Cell 1998, 92:603-610. 56. Zhang X, Shaw A, Bates PA, Newman RH, Gowen B, Orlova E, •• Gorman MA, Kondo H, Dokurno P, Lally J et al.: Structure of the AAA ATPase p97. Mol Cell 2000, 6:1473-1484. The crystal structure of the N and D1 domains of p97 at 2.9 Å, and a 3D cryo-EM reconstruction of murine p97 at 18 Å are presented. The p97 crystal structure demonstrates that the N domain of p97 is located at the periphery of the D1 hexamer and that ADP is bound between protomers. Furthermore, this structure reveals a role for the AAA+ minimum consensus (the characteristic sequence for AAA+ superfamily proteins) in sensing Pi and transferring any conformational changes upon ATP hydrolysis to protomer–protomer rearrangement. By comparison with the NSF D2–AMPPNP structure, the authors argue that ATP hydrolysis could cause subdomains to move closer, resulting in the linker between N and D1 being pushed out from the hexamer ring. By combining the EM reconstruction with the crystal structure, they propose a ratchet mechanism for p97, in which D1 and D2 exist in different nucleotide states and ATP hydrolysis in one domain induces nucleotide exchange in the other.
238
Macromolecular assemblages
57. •
Rouiller I, Butel VM, Latterich M, Milligan RA, Wilson-Kubalek EM: A major conformational change in p97 AAA ATPase upon ATP binding. Mol Cell 2000, 6:1485-1490. The authors used cryo-EM single-particle analysis to generate 2D average projections of p97 at 20–25 Å resolution in different nucleotide-bound states. They observed large conformational changes between the nucleotide-free and ATP-bound states, and small changes between the ATP- and ADP-bound states, and argued that ATP binding, instead of hydrolysis, is the major conformational step in the p97 ATP hydrolysis cycle. They extended their study to the p97–p47 complex and observed that, upon binding to p47, no significant conformational change in p97 was observed. Furthermore, they observe that major differences exist between the ATP-bound and nucleotide-free/ADP-bound states, and that six p47s bind to the periphery of the p97 hexamer in the presence of ATP. 58. Yuan X, Shaw A, Zhang X, Kondo H, Lally J, Freemont PS, Matthews S: Solution structure and interaction surface of the
C-terminal domain from p47: a major p97-cofactor involved in SNARE disassembly. J Mol Biol 2001, 311:255-263. 59. Yu RC, Jahn R, Brunger AT: NSF N-terminal domain crystal structure: models of NSF function. Mol Cell 1999, 4:97-107. 60. Lenzen CU, Steinmann D, Whiteheart SW, Weis WI: Crystal structure of the hexamerization domain of N-ethylmaleimide-sensitive fusion protein. Cell 1998, 94:525-536. 61. May AP, Misura KM, Whiteheart SW, Weis WI: Crystal structure of the amino-terminal domain of N-ethylmaleimide-sensitive fusion protein. Nat Cell Biol 1999, 1:175-182. 62. Sablin EP, Fletterick RJ: Nucleotides switches in molecular motors: structural analysis of kinesins and myosins. Curr Opin Struct Biol 2001, 11:716-724.
Machinery of protein folding and unfolding Xiaodong Zhang*, Fabienne Beuron and Paul S Freemont† During the past two years, a large amount of biochemical, biophysical and low- to high-resolution structural data have provided mechanistic insights into the machinery of protein folding and unfolding. It has emerged that dual functionality in terms of folding and unfolding might exist for some systems. The majority of folding/unfolding machines adopt oligomeric ring structures in a cooperative fashion and utilise the conformational changes induced by ATP binding/hydrolysis for their specific functions. Addresses Centre for Structural Biology, Department of Biological Sciences, Imperial College of Science, Technology and Medicine, Flowers Building, South Kensington, London SW7 2AZ, UK *e-mail: [email protected] †e-mail: [email protected] Current Opinion in Structural Biology 2002, 12:231–238 0959-440X/02/$ — see front matter © 2002 Elsevier Science Ltd. All rights reserved. Abbreviations AAA+ extended family of ATPase associated with a variety of cellular activities CCT cytosolic chaperonin containing the peptide tcp1 (tail-less complex polypeptide 1) Clp caseinolytic protease EM electron microscopy Hsl heat shock locus Hsp heat shock protein NSF N-ethylmaleimide-sensitive factor PDB Protein Data Bank TPR tetratricopeptide repeat
Introduction All biological activities and functions require proteins and multiprotein complexes. Proteins must therefore be able to fold into appropriate structures and assemble into specific complexes to carry out their biological activities. Misfolded proteins or misassembled complexes can malfunction and such events can be detrimental to all living organisms, including humans [1]. Successful protein folding is a multistep process that results in a protein structure often representing the lowest free energy state. To achieve this process in a cellular environment, which predominantly favours aggregation and nonproductive folding, the assistance of protein complexes called chaperones is required. Chaperones are crucial to maintaining the native protein conformation and preventing nonspecific aggregation, whereas the chaperonin proteins directly assist the folding process. To maintain appropriate concentration levels, properly folded proteins and their complexes need to be tightly regulated. Following biological activity, large complexes need to be disassembled to either dispose of or recycle particular components for further activity. One of the most efficient ways to reduce protein levels is via proteolysis. A
number of molecular machines have evolved to perform both the unfolding (‘unfoldase’) and proteolysis activities. Among these more specialised chaperone-assisted proteases are HslUV (heat shock locus UV), FtsH (a membranebound and ATP-dependent metalloprotease), ClpAP (caseinolytic protease AP) and ClpXP. The disassembly of oligomeric protein complexes into their constituent folded or partially folded parts requires a different class of machine. ‘Disassembler’ proteins include the ATPases NSF (N-ethylmaleimide-sensitive factor) and p97, both of which recycle essential components during vesicle membrane fusion [2]. Interestingly, many of the unfoldases and disassemblers belong to the large AAA+ (ATPase associated with a variety of cellular activities) family of ATPase proteins [3], indicating a possible common molecular mechanism that is applied to different biological functions. One general feature among all of the unfolding/folding machines is their dependence on successive ATP binding and hydrolysis cycles for activity. In this review, we will focus on recent structural and biochemical advances concerning some of these proteins, with the aim of summarising any common structural or mechanistic features that are emerging.
Mechanism of protein folding — chaperonins and chaperones Molecular chaperones comprise several highly conserved families of related proteins that are crucial for the maintenance of native protein conformation [4–6]. Chaperones are subdivided into different classes, as defined by their size, cellular distribution and function [7]. The chaperonins are large oligomeric ring-like proteins divided into two subfamilies: group I comprising the GroEL–GroES system and group II the thermosome [8]. Group I chaperonins — GroEL–GroES
A comprehensive structural picture of chaperonin-assisted protein folding has been obtained from extensive studies of the Escherichia coli GroEL–GroES system. From both cryo-electron microscopy (cryo-EM) and high-resolution crystal structures, mechanistic insights have been obtained that describe the folding cycle in terms of GroEL–GroES conformation and nucleotide-binding state [9–12]. GroEL consists of two seven-subunit ring structures (back to back), with each subunit formed of three domains (equatorial, intermediate and apical). Upon the positive cooperative binding of GroES and ATP to one ring, conformational changes result in a large closed cavity into which unfolded peptides are displaced. ATP hydrolysis in the first ring and binding of ATP to the second ring results in the release of both the folded or partially unfolded substrate and GroES, with both rings working out of phase. Currently, little is known about the conformation of GroEL substrates during the folding cycle. This past year
232
Macromolecular assemblages
has seen several biochemical studies aimed at addressing these issues. Hayer-Hartl and co-workers [13••], using a rapid inhibition assay and GroEL mutants that prevent substrate re-association, showed that the GroEL–GroES cage is essential for folding without aggregation. They concluded that confinement of unfolded proteins in the folding cavity increases the flux of intermediates toward the native folded state. Horwich and colleagues [14••], also using GroEL mutants defective in substrate binding, showed that GroEL substrates bind via multivalent interactions, which could provide a mechanism for further unfolding of bound misfolded substrates. Studies measuring protein compactness upon GroEL and ATP binding [15,16] suggest a ‘substrate stretching’ model. However, deuterium exchange experiments studying bound GroEL substrates argue against a forced mechanical unfolding mechanism as part of the intermediary folding pathway [17••]. A molecular dynamics study of GroEL by Karplus and colleagues [18] has provided some details of intermediate conformational changes during the GroEL allosteric cycle. They conclude that the individual GroEL monomers encode the necessary allosteric motions for heptamer cooperativity. Fersht and colleagues [19,20] suggest that, through evolution, GroEL has acquired a wide range of activities and does not rely on a single mechanism for all substrates. Indeed, it has been shown that GroEL–GroES can mediate the iterative binding of nonproductive quaternary complexes to promote dissociation and subunit exchange between heterodimers containing a misfolded monomer [21]. However, it still remains unresolved as to how GroEL–GroES could rescue misfolded non-native intermediates without exerting direct mechanical unfolding upon these intermediates. Group II chaperonins — thermosome and CCT
The archaebacterial thermosome (composed of two subunits: α and β) and eukaryotic CCT (cytosolic chaperonin containing the peptide tcp1 [tail-less complex polypeptide 1] comprising eight different but related subunits) belong to the group II subfamily of chaperonins. Group II subunits are organised similarly to GroEL, with two large domains (equatorial and apical) linked by hinge regions to a smaller (intermediate) domain. The crystal structure of the thermosome with and without nucleotide reveals a closed conformation with a built-in ‘lid’ domain that might substitute for the co-chaperonin GroES [22]. Recent structural information, primarily from cryo-EM, on two archaebacterial thermosomes and on the chaperonin CCT has provided further mechanistic insights into chaperonin function. By fitting the atomic coordinates of the thermosome comprising only α subunits into cryo-EM maps, Schoehn et al. [23•] have analysed the domain movements that occur between the co-existing open, asymmetric and closed conformations. The apical domain is highly flexible. These three conformations, previously described for the eukaryotic chaperonin CCT [24], most probably represent various
states in the ATPase cycle, which leads to encapsulation of the folding substrate. Considerable structural information from cryo-EM has recently been obtained that describes the conformational changes during CCT-facilitated folding of actin and tubulin [25,26,27••,28]. The authors reconcile their data in the following model: partially folded tubulin substrate proteins bind to apo-CCT at the apical domain of one ring. ATP binding induces the closure of the cavity in both rings. The ADP-Pi conformational state of the CCT–β tubulin complex exhibits a closed symmetric conformation, similar to the ATP conformation, with the substrate remaining at the apical domain. It is hypothesised that release of the nucleotide and subsequent expulsion of the folded substrate might occur upon movement of the apical domains or might involve other cellular cofactors. Specific CCT subunits are characterised by their different intrinsic affinity for ATP. Hence, through sequential ATP binding and hydrolysis, CCT assists partially folded actin and tubulin substrates to reach their final native state, with their nucleotide-binding pocket in place. Chaperones — Hsp90, Hsp70 (DnaK) and prefoldin
Among the chaperone group of proteins, eukaryotic Hsp90 (heat shock protein 90) and Hsp70 (and its prokaryotic homologue DnaK) have received much attention recently. Hsp90 is involved in the folding and assembly of proteins with roles in signal transduction, cell regulation and transcriptional activation [29,30]. Hsp90 operates as part of a multichaperone complex that includes numerous co-chaperones such as Hsp70 (which provides substrates for Hsp90-mediated folding), p23 and Hop (which acts as a scaffold for Hsp90 and Hsp70). Although much is known about the biochemical role of Hsp90, little is known about its inherent ATPase activity and the conformational changes induced by nucleotide binding and hydrolysis. Recent studies have tried to resolve some of these issues. ATP binding to Hsp90 has been shown to induce dimerisation of the N-terminal domain and substrate release involves ATP hydrolysis, an activity that is enhanced by the co-chaperone p23 [31,32]. This has led to the proposal that Hsp90 acts via a molecular clamp mechanism, similar to DNA gyrase and MutL, whereby opening and closing of a substrate-binding site is mediated by transient N-terminal dimerisation [32]. Kinetic analysis now shows that ATP hydrolysis is the rate-limiting step and that the C-terminal region of Hsp90 is essential for both ATP binding and enhancing hydrolysis rates [33]. The crystal structure of two TPR (tetratricopeptide repeat) domains of the Hop adaptor protein bound to target Hsp90/Hsp70-specific peptides [34••] has provided details of the assembly of a Hsp90–Hsp70 complex. However, a number of mechanistic and structural issues still remain unresolved, including a full structural description of the Hsp90 ATPase/substratebinding cycle. Several recent studies on the prokaryotic Hsp70 chaperone DnaK have been aimed at understanding the relationship
Machinery of protein folding and unfolding Zhang, Beuron and Freemont
between substrate affinity, nucleotide binding and co-chaperone-induced ATP hydrolysis. ATP binding to the N-terminal domain induces conformational changes that open the cavity to allow substrate binding and release. Subsequent ATP hydrolysis, induced by co-chaperone DnaJ (Hsp40), results in a closed state. Nucleotide exchange is a key element of substrate release and thus is critical to Hsp70 chaperone function. Recent studies identified structural elements within the DnaK family that define the interaction with the nucleotide exchange factors Bag-1 and GrpE [35,36]. Structures of Bag-1 bound to the ATPase domain of Hsc70 (the constitutively expressed cytosolic isoform of Hsp70) now provide direct insights into Hsp70 nucleotide exchange [37,38]. Hsp70 can be functionally substituted by the ubiquitous prefoldin (GimC) chaperone. Prefoldin is a heterohexamer that can interact with nascent polypeptides in vitro. The crystal structure of prefoldin reveals a unique quaternary structure that is unlike that of any other chaperone and it therefore forms a novel class. The structure comprises a double β-barrel assembly linking coiled coils with protruding flexible ‘tentacles’ containing hydrophobic patches required for peptide binding [39]. Unlike Hsp70 and the chaperonins, prefoldin is a novel class of chaperone that does not require ATP. It uses multiple interaction sites for substrate binding to shield non-native polypeptide from aggregation before passing it to other chaperones for completion of folding.
Mechanism of protein unfolding — chaperone-assisted protein degradation In eukaryotes, protein degradation is mainly achieved by a multiprotein complex, the 26S proteasome. In prokaryotes, a number of specialised chaperone-assisted proteases are used for a similar function. These include the two-component systems ClpX/ClpP (ClpXP), ClpA/ClpP (ClpAP) and HslU/HslV (HslUV) [40]. Recently, a number of HslU and HslUV structures have become available that provide significant mechanistic understanding of their function. Three different HslUV complexes have now been reported. The complex reported by Bochtler and colleagues [41••] positions the flexible intermediate (I) domain of HslU to interact with HslV. Subsequently, Sousa et al. [42••], using X-ray crystallography and small-angle X-ray solution scattering data, presented an alternative complex, in which a HslV dodecamer in the centre of the complex interacts with one HslU hexamer on each side, with the I domains distal. Cryo-EM 2D projection data at 30 Å resolution are consistent with this latter model [43•]. A third complex structure [44•,45•] also revealed a HslV dodecamer bound to the flat surface of a HslU hexamer, with the I domains distal. However, all three structures clearly identify a sixfold symmetry match between HslU and HslV, ruling out any mechanism of protease activation by symmetry mismatch [41••,42••,44•]. These structural data unambiguously illustrate that, like many other ATPase machines, HslU and HslV assemble
233
into hexameric ring structures that stack coaxially and at least one HslU hexamer is bound to a HslV dodecamer. It is also proposed that HslU binding to HslV could induce conformational changes in HslV to activate its peptidase activity [42••]. ATP binding in HslU has been proposed to enable HslU central pore opening. Furthermore, substrate recognised by the I domain would be first unfolded through I domain relocation, which is induced by HslU ATP hydrolysis, before translocation through the central pore into the protease chamber [42••,46•]. However, considerable dispute exists over which structural configuration represents the active HslUV complex. More crucially, the nature of substrate denaturation using ATP hydrolysis and the correlation between the ATP hydrolysis cycle and HslU pore opening remain unresolved. To understand the mechanism of these crucial steps, structures of HslUV complexed with different substrates at different stages, including substrate recognition, substrate unfolding and translocation, will be required. The Clp/Hsp100 chaperones are also AAA+ ATPases and function as substrate-specificity components of the ATP-dependent Clp proteases and also in preventing the aggregation and disassembly of transient multicomponent complexes involved in transcription, replication and the cell cycle [47]. ClpA, in the presence of ATP, mediates the unfolding of proteins that are bound following the recognition of specific terminal amino acid sequences (the ssrA peptide tag added to nascent proteins when ribosomes stall during translation). In the absence of the associated protease, ClpP, substrate proteins are released from ClpA or ClpX into solution, where they can spontaneously refold or become bound by other components, such as chaperonins or other Hsp100 family members [48–50]. In the absence of high-resolution structures of either ClpX or ClpA, recent biophysical studies have been informative. EM studies have helped to locate the substrate-binding site (positioned axially on the distal surface of the ATPase) and show that the unfolding and translocation of peptide substrates could proceed in discreet steps [43•,51]. It appears that ClpXP, ClpAP and the proteasome unravel folded substrates sequentially from the point of attachment at the degradation signal [52]. In multidomain proteins, independent stable domains are degraded sequentially.
Mechanism of protein complex disassembly – p97 and NSF Protein complexes often need to be disassembled as part of a regulatory system or to recycle certain components. A good example of this latter process is the recycling of the SNARE pairs formed after vesicle membrane fusion, whereby ATP hydrolysis is required for their disassembly. SNARE complexes comprise highly stable membraneattached helical coiled-coil structures that are formed after membrane fusion events [53]. Two homologous AAA+ ATPases, p97 and NSF, have been implicated in
234
Macromolecular assemblages
Figure 1 Structural comparisons. Ribbon representations of crystal structures of the ATPases discussed in this review in complex with ADP, with ADP similarly orientated: (a) ‘disassembler’ p97 D1 (PDB code 1E32); (b) ‘unfoldase’ HslU (PDB code 1G41); (c) group I chaperonin GroEL (PDB code 1AON); (d) group II chaperonin thermosome α subunit (PDB code 1A6E); (e) chaperone Hsp90 (PDB code 1BYQ); (f) chaperone Hsp70 (PDB code 1HJO).
disassembling these SNARE complexes via ATP hydrolysis and in association with specific adaptors [54,55]. The activities and mechanism of NSF in mediating membrane fusion have been well covered in a recent review [2]. Here, we focus on recent structural studies of p97, which reveal the organisation of p97 and its interaction with its cofactor, p47. p97 is a homohexamer and, like NSF, contains two AAA+ domains, D1 and D2. Crystal structures and cryo-EM analysis of p97 now show that its N-terminal (N) domain is located at the periphery of D1 [56••,57•] and that D1 and D2 hexamers stack coaxially [56••]. NMR studies also show that UBX, a ubiquitin-like domain situated at the C terminus of p47, interacts with the N-domain of p97 [58]. Comparison with the structure of NSF D2 bound to AMPPNP [59,60] reveals a possible conformational change upon ATP hydrolysis that could induce protomer–protomer rearrangements. This would also result in a closed central pore in the ADP state. This change would cascade to the relocation of the linker
between N and D1, generating mechanical force for SNARE disassembly via p47 [56••]. Based on the observation from EM that the D1 and D2 rings have different protomer arrangements, and the possible conformational changes between different nucleotide states, a ratchet mechanism for p97 has been proposed. In this model, D1 and D2 exist in different nucleotide states, and ATP hydrolysis in one domain induces nucleotide exchange in the other. It was also speculated that D1 and D2 pack in a tail-to-tail arrangement to accommodate the cage-like feature observed in the EM reconstruction, although alternative packing arrangements are possible. The proposed ratchet mechanism, however, is independent of the exact packing arrangement between D1 and D2. It is notable that GroEL–GroES also utilises two ATPase rings in different nucleotide-bound conformations as part of its activity cycle. A different study, using cryo-EM 2D projection analysis, proposed an alternative model for SNARE disassembly [57•]. In this model, nucleotide-free p97 binds loosely to
Machinery of protein folding and unfolding Zhang, Beuron and Freemont
235
Table 1 ATPase cycle and corresponding functional stages*.
GroEL CCT†
ATP binding
ATP hydrolysis
ADP release
Oligomeric state
i) Allows GroES binding ii) Releases GroES, ADP and substrate in opposite ring Induces closing of cavity
i) Weakens GroES interaction ii) Allows ATP and substrate binding in opposite ring Allosteric substrate folding
Resulting from ATP hydrolysis in opposite ring
Heptamer
Probably requires other cellular Octamer factors Hsp90 i) Induces N-domain dimerisation and i) Opens clamp to allow substrate Unknown Dimer substrate clamping release ii) Promotes p23 binding ii) Dissociates p23 Hsp70 Induces open conformation: low affinity for Induces closed conformation: Requires nucleotide exchange Monomer substrate traps substrates factor Bag-1/GrpE HslU Enables substrate binding and translocation Polypeptide unfolding Allosteric mechanism? Hexamer p97 Substrate binding Disassembly Ratchet mechanism? Hexamer ClpX/ Oligomerisation and substrate binding Substrate unfolding and Unknown Hexamer ClpA translocation *In the GroEL–GroES chaperone system, ATP binding not only step is not yet well characterised. However, an allosteric mechanism promotes GroES binding, but also allows GroES, ADP and has been proposed for the HslU hexamer, whereby ATP hydrolysis in substrate release in the opposite ring. ATP hydrolysis, on the other one subunit would promote ADP release in the next subunit [41]. hand, weakens the interaction with GroES and allows ATP and This mechanism, combined with the open pore state associated with substrate binding to occur in the opposite ring. In Hsp70, ATP ATP binding and substrate translocation, implies that denatured binding would induce an open state, resulting in low-affinity polypeptides would ‘spiral’ through the pore. This could be a general substrate binding, whereas ATP hydrolysis would induce a closed feature of unfoldases. For p97, two models have been proposed. It is state, thereby trapping substrates. A nucleotide exchange factor is possible that both ATP binding and hydrolysis generate required to remove the ADP. For Hsp90, ATP binding locks conformational changes driving SNARE disassembly. It is unclear substrates by clamping the substrate via ATP-induced dimerisation. how ADP is released in this system, though the proposed ratchet In the chaperonin-assisted protease systems (ClpX, HslU, FtsH mechanism would allow ADP release from the opposite ring upon AAA+ domain), ATP binding not only allows substrate binding, but ATP hydrolysis. †The thermosome has been omitted from this table also results in an open pore, which allows denatured peptides to because the various nucleotide states have not been identified pass to the protease chamber. ATP hydrolysis has been implicated unambiguously and the X-ray studies have revealed limited changes as being responsible for polypeptide denaturation. The ADP release upon nucleotide binding or hydrolysis [22].
monomeric p47 and, upon ATP binding, the ordered N-domain generates mechanical force to disassemble SNARE complexes. Given that the D1 and D2 domains are both active ATPases and that they may exist in different nucleotide-bound states, defining the actual p97 nucleotide state becomes nontrivial. Interestingly, both p97 models [56••,57•] propose that the relocation of the N-domain induced by ATP binding/hydrolysis mediates target protein disassembly, as previously suggested for NSF [59,61].
ATP binding/hydrolysis is the driving force for protein folding/unfolding One common element used by many molecular machines is their ability to successively bind and hydrolyse ATP. For motor proteins (kinesin and myosin), small structural changes in the nucleotide-binding site during the ATPase cycle are amplified into larger movements that provide the necessary force and motion [62]. Current structural and functional data now show that the majority of the molecular machines used for protein folding/unfolding also couple ATP binding, ATP hydrolysis and ADP release to appropriate conformational changes and functional states. A representative ATPase structure (monomer) of each of the chaperone systems discussed is shown in Figure 1, with bound ADP oriented similarly. It is notable that, in all of the systems, the nucleotide is bound between subdomains (intradomain binding). This provides a structural explanation for how nucleotide binding and hydrolysis could induce subdomain reorientation, in order to generate mechanical forces. In
certain cases, such as p97 and HslU, the nucleotide is also bound between protomers (interdomain binding) and ATP hydrolysis might require residues from an adjacent protomer, as proposed for p97 [56••]. Furthermore, most of the folding/unfolding systems utilise single or multiple oligomeric ring structures. These oligomeric ATPases allow ATP binding and hydrolysis to be either concerted, to provide a single conformational transition (force) (such as in GroEL, NSF/p97), or allosteric, to allow multiple conformational transitions between protomers (such as in HslU, ClpX and CCT). Two-ring ATPase structures such as GroEL allow further adaptations, including allosteric transitions between the individual rings.
Conclusions An ATP hydrolysis cycle can be divided into three steps: ATP binding, ATP hydrolysis, and ADP and Pi release. Each system uses one or multiple steps to generate the necessary conformational changes and mechanical forces for their functions (summarised in Table 1). A common feature from a number of molecular machines, including unfoldases, disassemblers and certain chaperone systems, is that ATP binding allows substrate release, as well as substrate binding, whereas ATP hydrolysis induces substrate remodelling (unfolding or disassembly). However, the significant amount of structural information available and the diverse mechanistic models proposed demonstrates the complexity of the protein folding/unfolding machinery. Not only does each system adapt different stages of the
236
Macromolecular assemblages
ATPase cycle to actively perform folding/unfolding, but they also utilise cooperativity between different subunits within the machinery. Furthermore, each system might use different mechanisms for different targets, as proposed for GroEL–GroES. It is becoming increasingly apparent, however, that a combination of high- and medium-resolution structures that capture various stages of the functional cycle, together with biochemical and biophysical studies, is required for a deeper understanding of these complex machineries.
Acknowledgements We wish to thank V Pye and I Dreveny for critically reading the manuscript.
References and recommended reading Papers of particular interest, published within the annual period of review, have been highlighted as:
• of special interest •• of outstanding interest
15. Hammarstrom P, Persson M, Owenius R, Lindgren M, Carlsson U: Protein substrate binding induces conformational changes in the chaperonin GroEL. A suggested mechanism for unfoldase activity. J Biol Chem 2000, 275:22832-22838. 16. Hammarstrom P, Persson M, Carlsson U. Protein compactness measured by fluorescence resonance energy transfer. Human carbonic anhydrase II is considerably expanded by the interaction of GroEL. J Biol Chem 2001, 276:21765-21775. 17. ••
Chen J, Walter S, Horwich AL, Smith DL: Folding of malate dehydrogenase inside the GroEL-GroES cavity. Nat Struct Biol 2001, 8:721-728. Using NMR and mass spectrometry methods, the authors found that, when bound to GroEL, a core of malate dehydrogenase (MDH) secondary structure was protected from proton exchange and that, upon ATP and GroES binding, this core was only partially deprotected. Their results are consistent with hydrogen bonds between the sides of the cavity and MDH substrate being broken, as opposed to forced mechanical unfolding. 18. Ma J, Sigler PB, Xu Z, Karplus M: A dynamic model for the allosteric mechanism of GroEL. J Mol Biol 2000, 302:303-313. 19. Chatellier J, Hill F, Fersht AR: From minichaperone to GroEL 2: importance of avidity of the multisite ring structure. J Mol Biol 2000, 304:883-896.
1.
Slavotinek AM, Biesecker LG: Unfolding the role of chaperones and chaperonins in human disease. Trends Genet 2001, 17:528-535.
20. Chatellier J, Hill F, Foster NW, Goloubinoff P, Fersht AR: From minichaperone to GroEL 3: properties of an active single-ring mutant of GroEL. J Mol Biol 2000, 304:897-910.
2.
May AP, Whiteheart SW, Weis WI: Unraveling the mechanism of the vesicle transport ATPase NSF, the N-ethylmaleimide-sensitive factor. J Biol Chem 2001, 276:21991-22004.
3.
Ogura T, Wilkinson AJ: AAA+ superfamily ATPases: common structure-diverse function. Genes Cells 2001, 6:575-597.
21. Wynn RM, Song JL, Chuang DT: GroEL / GroES promote dissociation / reassociation cycles of a heterodimeric intermediate during alpha(2)beta(2) protein assembly. Iterative annealing at the quaternary structure level. J Biol Chem 2000, 275:2786-2794.
4.
Grantcharova V, Alm EJ, Baker D, Horwich AL: Mechanisms of protein folding. Curr Opin Struct Biol 2001, 11:70-82.
5.
Gottesman ME, Hendrickson WA: Protein folding and unfolding by Escherichia coli chaperones and chaperonins. Curr Opin Microbiol 2000, 3:197-202.
6.
Carrascosa JL, Llorca O, Valpuesta JM: Structural comparison of prokaryotic and eukaryotic chaperonins. Micron 2001, 32:43-50.
7.
Ben-Zvi AP, Goloubinoff P: Mechanisms of disaggregation and refolding of stable protein aggregates by molecular chaperones. J Struct Biol 2001, 135:84-93.
8.
Kusmierczyk AR, Martin J: Chaperonins: keeping a lid on folding proteins. FEBS Lett 2001, 21:343-347.
9.
Feltham JL, Gierasch LM: GroEL-substrate interactions: molding the fold, or folding the mold? Cell 2000, 100:193-196.
10. Thirumalai D, Lorimer GH: Chaperonin-mediated protein folding. Annu Rev Biophys Biomol Struct 2001, 30:245-269. 11. Melki R: Nucleotide-dependent conformational changes of the chaperonin containing tcp-1. J Struct Biol 2001, 135:170-175. 12. Steinbacher S, Ditzel L: Nucleotide binding to the thermoplasma thermosome: implications for the functional cycle of group II chaperonins. J Struct Biol 2001, 135:147-156. 13. Brinker A, Pfeifer G, Kerner MJ, Naylor DJ, Hartl FU, Hayer-Hartl M: •• Dual function of protein confinement in chaperonin-assisted protein folding. Cell 2001, 107:223-233. The folding of Rubisco and rhodenase using a modified GroEL mutant structure that blocks the rebinding of non-native proteins was studied. The results showed that folding within the GroEL cage is independent of ATP hydrolysis cycles and significantly faster than in free solution. This argues that the confinement of unfolded proteins within the folding cavity increases the flux of folding intermediates toward the native folded state. 14 ••
Farr GW, Furtak K, Rowland MB, Ranson NA, Saibil HR, Kirchhausen T, Horwich AL: Multivalent binding of nonnative substrate proteins by the chaperonin GroEL. Cell 2000, 100:561-573. This paper describes studies of mutant GroEL complexes binding to three different substrates, namely Rubisco, malate dehydrogenase (MDH) and rhodenase. For Rubisco and MDH, consecutive wild-type GroEL apical domains are preferred for binding. This could be explained by the need for a continuous hydrophobic binding surface or by the requirement of the apical domain to act cooperatively in substrate binding. A multivalent binding model is proposed for GroEL.
22. Ditzel L, Löwe J, Stock D, Stetter KO, Huber H, Huber R, Steinbacher S: Crystal structure of the thermosome, the archaeal chaperonin and homolog of CCT. Cell 1998, 93:125-138. 23. Schoehn G, Hayes M, Cliff M, Clarke AR, Saibil HR: Domain • rotations between open, closed and bullet-shaped forms of the thermosome, an archaeal chaperonin. J Mol Biol 2000, 301:323-332. The native chaperonin thermophilic factor 55 (TF55) is a heat shock protein from the archaea Sulfolobus shibatae; α, β and γ subunits have been identified. The 3D reconstruction of TF55 (ninefold symmetry) described in this report constitutes the first characterisation of three coexisting conformations in solution: open, asymmetric (‘bullet’ shaped) and closed. The observation of a ‘bullet’ asymmetric form for a group II chaperonin reveals a close parallel with the GroEL system. 24. Llorca O, Smyth MG, Carrascosa JL, Willison KR, Radermacher M, Steinbacher S, Valpuesta JM: 3D reconstruction of the ATP-bound form of CCT reveals the asymmetric folding conformation of a type II chaperonin. Nat Struct Biol 1999, 6:639-642. 25. Llorca O, McCormack EA, Hynes G, Grantham J, Cordell J, Carrascosa JL, Willison KR, Fernandez JL, Valpuesta JM: Eukaryotic type II chaperonin CCT interacts with actin through specific subunits. Nature 1999, 402:693-696. 26. Llorca O, Martin-Benito J, Ritco-Vonsovici M, Grantham J, Hynes GM, Willison KR, Carrascosa JL, Valpuesta JM: Eukaryotic chaperonin CCT stabilizes actin and tubulin folding intermediates in open quasi-native conformations. EMBO J 2000, 19:5971-5979. 27. ••
Llorca O, Martin-Benito J, Grantham J, Ritco-Vonsovici M, Willison KR, Carrascosa JL, Valpuesta JM: The ‘sequential allosteric ring’ mechanism in the eukaryotic chaperonin-assisted folding of actin and tubulin. EMBO J 2001, 20:4065-4075. CCT-bound actin is resolved as rod-shaped densities (‘stretched’ actin as opposed to ‘V’-shaped molecules) in which small and large domains are identifiable. The exact binding pattern has been mapped using monoclonal antibodies and 2D averaging. The binding of one molecule of tubulin causes substantial conformational changes in apo-CCT, which adopts a semiclosed conformation [26]. Five specific subunits using two modes of interaction are involved in the binding of tubulin to CCT. Interestingly, the location of the substrate-binding sites at the tips of the apical domains of CCT is different for actin and tubulin. 28. Llorca O, Martin-Benito J, Gomez-Puertas P, Ritco-Vonsovici M, Willison KR, Valpuesta JM: Analysis of the interaction between the eukaryotic chaperonin CCT and its substrates actin and tubulin. J Struct Biol 2001, 135:205-218. 29. Pearl LH, Prodromou C: Structure and in vivo function of Hsp90. Curr Opin Struct Biol 2000, 10:46-51.
Machinery of protein folding and unfolding Zhang, Beuron and Freemont
30. Young JC, Moarefi I, Hartl FU: Hsp90: a specialized but essential protein-folding tool. J Cell Biol 2001, 154:267-273 31. Young JC, Hartl FU: Polypeptide release by Hsp90 involves ATP hydrolysis and is enhanced by the co-chaperone p23. EMBO J 2000, 19:5930-5940. 32. Prodromou C, Panaretou B, Chohan S, Siligardi G, O’Brien R, Ladbury JE, Roe SM, Piper PW, Pearl LH: The ATPase cycle of Hsp90 drives a molecular ‘clamp’ via transient dimerization of the N-terminal domains. EMBO J 2000, 19:4383-4392. 33. Weikl T, Abelmann K, Buchner J: An unstructured C-terminal region of the Hsp90 co-chaperone p23 is important for its chaperone function. J Mol Biol 1999, 293:685-691. 34. Scheufler C, Brinker A, Bourenko G, Pegoraro S, Moroder L, •• Bartunik H, Hartl FU, Moarefi I: Structure of TPR domain-peptide complexes: critical elements in the assembly of the Hasp70-Hasp90 multichaperone machine. Cell 2000, 101:199-210. The crystal structure of two TPR domains of the adaptor protein Hop bound to Hsp90/Hsp70-specific peptides reveals the first molecular insights into the assembly of the Hsp70–Hsp90 multichaperone complex. Interestingly, the Hop TPR–peptide structures show some similarity to 14-3-3–peptide complexes, indicating a common ancestral mode of protein–peptide binding. 35. Brehmer D, Rüdiger S, Gässler CS, Klostermeier D, Packschies L, Reinstein J, Mayer MP, Bukau B: Tuning of chaperone activity of Hsp70 proteins by modulation of nucleotide exchange. Nat Struct Biol 2001, 8:427-432. 36. Han W, Christen P: Mutations in the interdomain linker region of DnaK abolish the chaperone action of the DnaK/DnaJ/GrpE system. FEBS Lett 2001, 497:55-58. 37.
Briknarova K, Takayama S, Brive L, Havert ML, Knee DA, Velasco J, Homma S, Cabezas E, Stuart J, Hoyt DW et al.: Structural analysis of BAG1 cochaperone and its interactions with Hsc70 heat shock protein. Nat Struct Biol 2001, 8:349-352.
237
44. Wang J, Song JJ, Franklin MC, Kamtekar S, Im YJ, Rho SH, Seong IS, • Lee CS, Chung CH, Eom SH: Crystal structures of the HslVU peptidase-ATPase complex reveal an ATP-dependent proteolysis mechanism. Structure 2001, 9:177-184. The authors describe an alternative arrangement of HslUV from E. coli at 3.0 Å (see [41••]), with a HslV dodecamer bound to the flat surface of a HslU hexamer, leaving the I domain of HslU distal. The size of the HslV pore changes between free and complexed HslV, and the authors argue that binding to HslU induces pore size changes in HslV, probably allowing substrate release. 45. Wang J, Song JJ, Seong IS, Franklin MC, Kamtekar S, Eom SH, • Chung CH: Nucleotide-dependent conformational changes in a protease-associated ATPase HslU. Structure 2001, 9:1107-1116. The authors compared all available HslU structures and proposed that nucleotide binding, rather than hydrolysis, induces the largest conformational changes in subdomain arrangement. Furthermore, they propose that different functional states exist during the ATP hydrolysis cycle and that these states correspond to the different stages of protein unfolding and translocation. 46. Song HK, Hartmann C, Ramachandran R, Bochtler M, Behrendt R, • Moroder L, Huber R: Mutational studies on HslU and its docking mode with HslV. Proc Natl Acad Sci USA 2000, 97:14103-14108. These authors presented systematic mutagenesis data and various peptidase activity data to dissect the roles of various regions in HslU. Deletion in the I domain in general had no effect on peptidase activity, although it abolishes the degradation of folded proteins, suggesting that the I domain is involved in substrate recognition and/or unfolding. The authors argue that the peptidase and caseinolytic activities might not require a precise complex between HslU and HslV, and different HslUV complexes might be involved in the degradation of different substrates 47.
Jenal U, Fuchs T: An essential protease involved in bacterial cell-cycle control. EMBO J 1998, 17:5658-5669.
48. Weber-Ban EU, Reid BG, Miranker AD, Horwich AL: Global unfolding of a substrate protein by the Hsp100 chaperone ClpA. Nature 1999, 401:90-93.
38. Sondermann H, Scheufler C, Schneider C, Hohfeld J, Hartl FU, Moarefi I: Structure of a Bag/Hsc70 complex: convergent functional evolution of Hsp70 nucleotide exchange factors. Science 2001, 29:1553-1557.
49. Singh S, Grimaud R, Hoskins J, Wickner S, Maurizi MR: Unfolding and internalization of proteins by the ATP-dependent proteases ClpXP and ClpAP. Proc Natl Acad Sci USA 2000, 97:8898-8903.
39. Siegert R, Leroux MR, Scheufler C, Hartl FU, Moarefi I: Structure of the molecular chaperone prefoldin: unique interaction of multiple coiled coil tentacles with unfolded proteins. Cell 2000, 102:621-632.
50. Kim YI, Burton RE, Burton BM, Sauer RT, Baker TA: Dynamics of substrate denaturation and translocation by the ClpXP degradation machine. Mol Cell 2000, 5:639-648.
40. Zwickl P, Baumeister W, Steven A: Dis-assembly lines: the proteasome and related ATPase-assisted proteases. Curr Opin Struct Biol 2000, 10:242-250.
51. Ortega J, Singh S, Ishikawa T, Maurizi MR, Steven AC: Visualization of substrate binding and translocation by the ATP-dependent protease, ClpXP. Mol Cell 2000, 6:1515-1521.
41. Bochtler M, Hartmann C, Song HK, Bourenkov GP, Bartunik HD, •• Huber R: The structures of HsIU and the ATP-dependent protease HsIU-HsIV. Nature 2000, 403:800-805. Crystal structures of HslU in different nucleotide-bound states and of the HslUV complex from E. coli were presented. The structure of the HslUV complex is the first for a complete set of components of an ATP-dependent protease. It also clarified the sixfold symmetry match between HslU and HslV, ruling out a mechanism of protease activation by symmetry mismatch. The complex structure reveals a dodecamer of HslV bound to the I domains of a HslU hexamer on both sides. The various HslU structures with different nucleotide bound allow the authors to speculate an allosteric mechanism for HslU. These structures also establish that HslU belongs to the AAA+ superfamily.
52. Lee C, Schwartz MP, Prakash S, Iwakura M, Matouschek A: ATP-dependent proteases degrade their substrates by processively unraveling them from the degradation signal. Mol Cell 2001, 7:627-637.
42. Sousa MC, Trame CB, Tsuruta H, Wilbanks SM, Reddy VS, •• McKay DB: Crystal and solution structures of an HslUV protease-chaperone complex. Cell 2000, 103:633-643. The authors report the structures of a HslUV complex from H. influenzae at 3.4 Å in the presence of ATP and of ADP-bound HslU at 2.3 Å. Their data show an alternative HslUV complex from that of E. coli [41••], in that a HslV dodecamer in the centre of the complex interacts with the flat part of HslU, while the I domain of HslU is distal. Small-angle X-ray scattering experiments and peptidase activity assays were used to support the crystal structure being an active complex. 43. Ishikawa T, Maurizi MR, Belnap D, Steven AC: Docking of • components in a bacterial complex. Nature 2000, 408:667-668. This method allowed the direct quantification of the protein densities inside ClpP in a stalled ClpAP complex. A mass of 41 kDa (1.5 RepA subunits) was calculated to be able to fit inside ClpP. Presumably, unfolding is driven by multivalent contacts between the surrounding ClpA subunits and the substrate protein. ATP binding or hydrolysis probably drives a conformational change in ClpA that pulls the protein structure apart, in an action analogous to that of GroEL. However, at the resolution of this work (30 Å), no significant conformational changes were observed in ClpA upon ATP hydrolysis.
53. Hay JC: SNARE complex structure and function. Exp Cell Res 2001, 271:10-21. 54. Sollner T, Bennett MK, Whiteheart SW, Scheller RH, Rothman JE: A protein assembly-disassembly pathway in vitro that may correspond to sequential steps of synaptic vesicle docking, activation, and fusion. Cell 1993, 75:409-418. 55. Rabouille C, Kondo H, Newman R, Hui N, Freemont P, Warren G: Syntaxin 5 is a common component of the NSF- and p97-mediated reassembly pathways of Golgi cisternae from mitotic Golgi fragments in vitro. Cell 1998, 92:603-610. 56. Zhang X, Shaw A, Bates PA, Newman RH, Gowen B, Orlova E, •• Gorman MA, Kondo H, Dokurno P, Lally J et al.: Structure of the AAA ATPase p97. Mol Cell 2000, 6:1473-1484. The crystal structure of the N and D1 domains of p97 at 2.9 Å, and a 3D cryo-EM reconstruction of murine p97 at 18 Å are presented. The p97 crystal structure demonstrates that the N domain of p97 is located at the periphery of the D1 hexamer and that ADP is bound between protomers. Furthermore, this structure reveals a role for the AAA+ minimum consensus (the characteristic sequence for AAA+ superfamily proteins) in sensing Pi and transferring any conformational changes upon ATP hydrolysis to protomer–protomer rearrangement. By comparison with the NSF D2–AMPPNP structure, the authors argue that ATP hydrolysis could cause subdomains to move closer, resulting in the linker between N and D1 being pushed out from the hexamer ring. By combining the EM reconstruction with the crystal structure, they propose a ratchet mechanism for p97, in which D1 and D2 exist in different nucleotide states and ATP hydrolysis in one domain induces nucleotide exchange in the other.
238
Macromolecular assemblages
57. •
Rouiller I, Butel VM, Latterich M, Milligan RA, Wilson-Kubalek EM: A major conformational change in p97 AAA ATPase upon ATP binding. Mol Cell 2000, 6:1485-1490. The authors used cryo-EM single-particle analysis to generate 2D average projections of p97 at 20–25 Å resolution in different nucleotide-bound states. They observed large conformational changes between the nucleotide-free and ATP-bound states, and small changes between the ATP- and ADP-bound states, and argued that ATP binding, instead of hydrolysis, is the major conformational step in the p97 ATP hydrolysis cycle. They extended their study to the p97–p47 complex and observed that, upon binding to p47, no significant conformational change in p97 was observed. Furthermore, they observe that major differences exist between the ATP-bound and nucleotide-free/ADP-bound states, and that six p47s bind to the periphery of the p97 hexamer in the presence of ATP. 58. Yuan X, Shaw A, Zhang X, Kondo H, Lally J, Freemont PS, Matthews S: Solution structure and interaction surface of the
C-terminal domain from p47: a major p97-cofactor involved in SNARE disassembly. J Mol Biol 2001, 311:255-263. 59. Yu RC, Jahn R, Brunger AT: NSF N-terminal domain crystal structure: models of NSF function. Mol Cell 1999, 4:97-107. 60. Lenzen CU, Steinmann D, Whiteheart SW, Weis WI: Crystal structure of the hexamerization domain of N-ethylmaleimide-sensitive fusion protein. Cell 1998, 94:525-536. 61. May AP, Misura KM, Whiteheart SW, Weis WI: Crystal structure of the amino-terminal domain of N-ethylmaleimide-sensitive fusion protein. Nat Cell Biol 1999, 1:175-182. 62. Sablin EP, Fletterick RJ: Nucleotides switches in molecular motors: structural analysis of kinesins and myosins. Curr Opin Struct Biol 2001, 11:716-724.