Allosteric Mechanisms in Chaperonin Machines - American Chemical

Review pubs.acs.org/CR

Allosteric Mechanisms in Chaperonin Machines Ranit Gruber† and Amnon Horovitz*,† †

Department of Structural Biology, Weizmann Institute of Science, Rehovot 76100, Israel ABSTRACT: Chaperonins are nanomachines that facilitate protein folding by undergoing energy (ATP)-dependent movements that are coordinated in time and space owing to complex allosteric regulation. They consist of two back-to-back stacked oligomeric rings with a cavity at each end where protein substrate folding can take place. Here, we focus on the GroEL/GroES chaperonin system from Escherichia coli and, to a lesser extent, on the more poorly characterized eukaryotic chaperonin CCT/TRiC. We describe their various functional (allosteric) states and how they are affected by substrates and allosteric effectors that include ATP, ADP, nonfolded protein substrates, potassium ions, and GroES (in the case of GroEL). We also discuss the pathways of intra- and inter-ring allosteric communication by which they interconvert and the coupling between allosteric transitions and protein folding reactions.

CONTENTS

1. INTRODUCTION

1. Introduction 1.1. Overview of Allosteric Models 1.2. Potential Allosteric Mechanisms in RingShaped Machines 1.3. Chaperonins: A Primer 2. Experimental Analysis of Allostery in Chaperonins 2.1. Intra-Ring Allostery with Respect to ATP and Its Analogs 2.2. Intra-Ring Allostery with Respect to K+ Ions 2.3. Inter-Ring Allostery in ATP and GroES Binding and the Nested Model 2.4. Allosteric Effects of ADP 2.5. Allosteric Effects of Nonfolded Protein Substrates 3. Structural Basis of Allostery in Chaperonins 3.1. Structural Basis of Intra-Ring Allostery 3.2. Structural Basis of Inter-Ring Allostery 4. Role of Allostery in Chaperonin Function 4.1. Intra-Ring Allostery in Chaperonin Function 4.2. Inter-Ring Allostery in Chaperonin Function 5. Concluding Remarks Author Information Corresponding Author Notes Biographies Acknowledgments References

6588 6588

1.1. Overview of Allosteric Models

The term allostery derives from the Greek words allos and stereos that mean “other” and “solid (object)”, respectively, in accordance with the fact that it refers to the effect of a perturbation at one (regulatory) site in a molecule on the activity at another distinct and often remote (active) site. Such perturbations can be due to ligand binding or to mutations that are often then considered as being allosteric. The term was first used by Monod and Jacob1 in connection with the findings of Changeux2 that L-threonine deaminase is inhibited by its end product, L-isoleucine. They suggested that L-isoleucine inhibits L-threonine deaminase by binding to a site that does not overlap with the enzyme’s active site. These studies, therefore, indicated that feed-back inhibition in metabolic networks can be achieved by allosteric regulation of enzyme activity. They also stimulated the development of several models described below that could account for such action at a distance. These models were developed with oligomeric proteins in mind since all early examples for allostery, such as hemoglobin and Lthreonine deaminase, were known to be multisubunit. Allosteric regulation in oligomeric proteins is usually reflected in plots of initial reaction velocity or fractional saturation as a function of substrate or effector concentration. Sigmoidicity in such plots indicates the existence of positive cooperativity in ligand binding (or activity), i.e., that ligand binding to one site increases the affinity (or activity) of other sites for the same ligand (homotropic cooperativity) or other ligands (heterotropic cooperativity). In the absence of cooperativity, such plots are hyperbolic, whereas negative

6590 6590 6591 6591 6593 6593 6595 6596 6596 6596 6598 6599 6599 6600 6601 6601 6601 6601 6601 6601 6601

Special Issue: Protein Ensembles and Allostery Received: September 20, 2015 Published: January 4, 2016 © 2016 American Chemical Society

6588

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

low affinity for the substrate and a relaxed (R) state with higher affinity for the substrate (L = [T]/[R]). Substrate binding shifts the equilibrium from the T to R states in a concerted fashion, i.e., asymmetric states in which some of the subunits in a given oligomer are in the T state conformation and others in the R state conformation are not allowed. Owing to the conservation of symmetry, substrate binding causes also unbound subunits to switch from the T to the R state conformation, thereby generating unoccupied high-affinity sites that give rise to cooperativity. The extent of cooperativity in the MWC model is determined by the values of the allosteric equilibrium constant, L, and the relative affinities of the substrate for the T and R states (c = KT/KR). The main appeal of the MWC model is its elegance and that only two parameters, L and c, are needed to fit the data. Its main drawbacks are that it cannot account for negative cooperativity and that data fitting in the case of nonexclusive binding to the R state (c ≠ 0) can be difficult.6 The MWC model has been used extensively in biology.7 Its basic assumption, that the apo state of a protein is in equilibrium between low- and high-affinity conformations and that the ligand selects the most favored conformation thereby leading to a population shift, is also at the heart of the more recently popular “conformational selection” model.8 By contrast with the MWC model, ligand binding-promoted conformational changes in the KNF model take place in a sequential fashion, i.e., asymmetric states are allowed (Figure 1). In the KNF model, the ligand-bound conformation is induced upon ligand binding and does not exist in the absence of the ligand. Cooperativity in this model is due to progressive changes in the intersubunit interaction energies. The main advantage of the KNF model is that, unlike the MWC model, it can account for negative cooperativity. Its principal drawback is that it requires more parameters for data fitting. In the case of positive cooperativity, it is impossible to distinguish between the MWC and the KNF models from plots of initial rates (or fractional saturation) as a function of substrate concentration because such plots are insensitive to the presence of ligation intermediates. Recently, however, distinguishing between these models has become possible owing to advances in singlemolecule techniques and native mass spectrometry.9 Common to both the MWC and the KNF models is that cooperativity is attributed to ligand-promoted conformational changes that are concerted (MWC), sequential (KNF), or a combination of both.10 Cooperativity can, however, also arise owing to a change in the extent of conformational fluctuations about a mean structure in the absence of a change in the mean structure itself.11,12 As mentioned above, cooperativity in the KNF model arises owing to changes in the intersubunit interaction energies. Such changes are expected to be particularly large when ligand binding favors subunit dissociation or association. Liganddriven subunit assembly/disassembly, therefore, constitutes a potential mechanism for attaining unusually high cooperativity. The ligand may have a higher affinity for the monomer than the oligomer, in which case ligand binding will promote dissociation of the oligomer. Alternatively, it may have a higher affinity for the oligomeric state, in which case ligand binding will drive oligomer assembly. In the absence of V system effects, both scenarios will result in positive cooperativity in ligand binding, with respect to ligand concentration, even if all the binding sites in the oligomer have the same affinities for the ligand. Such an allosteric mechanism has been observed, for example, in members of the ClpB/Hsp104 ring-shaped

cooperativity in ligand binding (or activity), which is harder to distinguish, is reflected in slightly flattened hyperbolic curves. The extent of cooperativity and whether it is positive or negative can be determined by fitting data of initial reaction velocity or fractional saturation as a function of substrate concentration to the Hill equation

Y̅ =

K[S]n 1 + K[S]n

(1)

where Y̅ is the fraction of sites that are bound, [S] is the substrate concentration, K is the apparent binding constant, and n is the Hill coefficient. The Hill equation was derived assuming that binding occurs in an all-or-none fashion, i.e., that n is equal to the total number of binding sites, N.3 In practice, the value of n is usually noninteger and smaller than the total number of binding sites because of the existence of ligation intermediates. The Hill equation remains, however, useful for quantifying the extent of positive cooperativity (i.e., 1 < n < N) and detecting lack of cooperativity (n = 1) or apparent negative cooperativity (0 < n < 1). Given that it is usually easier to measure more accurately the velocity of an enzyme’s reaction than its fractional saturation, it is often assumed that Y̅ = V/ Vmax, i.e., that the catalytic rate constant does not depend on the extent of saturation of the binding sites (K systems). Cooperativity can, however, also involve changes in the catalytic properties as a function of the extent of saturation by substrates or effectors (V systems). Several models, in particular, the Monod−Wymam−Changeux (MWC) model4 and the Koshland−Némethy−Filmer (KNF) model,5 were put forward in the 1960s to account for cooperativity in ligand binding by oligomeric proteins. According to the MWC model (Figure 1), the oligomeric protein is in equilibrium, in the absence of substrates or effectors, between two states: a so-called tense (T) state with

Figure 1. General scheme for the binding of a substrate to a tetramer in different allosteric states.194 Squares and circles represent subunit conformations with low and high affinity for the substrate (S), respectively. States on the diagonal that are surrounded by the dashed red line are those considered by the KNF model. States in the left and right columns that are surrounded by the dashed red line correspond to the T and R states, respectively, in the MWC model. Free substrate is omitted for clarity. 6589

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

and are found in Nature. For example, it has been reported21 for the bacteriophage φ29 packaging motor, a homopentameric ring ATPase, that both ATP binding and hydrolysis are sequential. By contrast, structural studies of the large tumor antigen (LTag) of simian virus 40, an AAA+ protein that is a hexameric helicase, have shown that it can be fully occupied by ATP or ADP but not by a mixture of both, thereby suggesting a concerted hydrolysis mechanism.22 It should be noted that full occupancy by ATP depends on its concentration and does not necessarily imply that the MWC model (which refers to conformational changes that are concerted and not to concerted binding) is applicable. As mentioned above, concerted all-or-none binding should be reflected in values of Hill coefficients that are equal to the number of binding sites, but such values are rarely measured. In addition to the sequential and concerted mechanisms, studies on ClpX, a bacterial hexameric ring that unfolds proteins and translocates them into ClpP for degradation, have indicated that probabilistic conformational switching can also occur.23 This review will focus on the allosteric mechanisms of chaperonins and, in particular, on those of GroEL and CCT/TRiC that display concerted and sequential intra-ring allostery, respectively, with regard to ATP binding and/or hydrolysis. Allostery in chaperonins has been reviewed before,24−27 but here, in addition to discussion of structure and function, more attention will also be given to the thermodynamic and kinetic aspects of the subject.

chaperone family which undergo nucleotide-dependent assembly reactions and display positive cooperativity in ATPase activity, with respect to both ATP and protein subunit concentrations.13 1.2. Potential Allosteric Mechanisms in Ring-Shaped Machines

Ring-shaped nanomachines are ubiquitous and found in all forms of life. Examples include DNA binding proteins14 and chaperone rings involved in mediating protein folding and degradation,15 many of which are hexamers that belong to the AAA+ (ATPases Associated with diverse cellular Activities) superfamily of P-loop NTPases. The ring-shaped architecture of DNA and RNA binding proteins enables them to encircle double-stranded DNA or RNA and translocate along them. In the case of chaperone rings, their central cavities can provide a supportive environment for protein folding16−18 or unfolding and translocation into degradation chambers.19 It is justified to consider these ring-shaped proteins as machines because their parts move in a coordinated fashion that is most often fueled by ATP consumption and coupled to the work they carry out. A common misconception is that the free energy change upon ATP hydrolysis can be harnessed to do work, but in general, that is probably not true for biological systems where most of the released heat is rapidly dissipated. Interestingly, however, it was reported recently that the heat released during catalysis by an enzyme enhances its diffusion.20 Hence, it is possible that some machines have evolved to “channel” the energy released upon hydrolysis toward a useful purpose associated with their function. Regardless, it is likely that the action of most machines is driven mainly by conformational changes due to ATP binding and hydrolysis and that cooperativity in these reactions is responsible for their coordinated movements. The appreciation that allostery is crucial for the mechanisms of molecular machines has contributed to the renewed interest in understanding allosteric phenomena. The MWC and KNF models can be employed to describe concerted and sequential allostery, respectively, due to ATP binding by molecular machines, but many of them also display allostery in hydrolysis. Many combinations of types of allostery in ATP binding and hydrolysis can be considered (Figure 2)

1.3. Chaperonins: A Primer

Chaperonins are nanodevices that assist protein folding by undergoing conformational changes that are controlled by ATP binding and hydrolysis. They consist of two back-to-back stacked oligomeric rings with a cavity at each end where protein substrate folding can take place. The chaperonins can be classified into two groups: group I, members of which are found in eubacteria, mitochondria, and chloroplasts;17,18,28,29 and group II, whose members are found in archaea and the eukaryotic cytosol.30−33 Group I chaperonins consist of two identical (as in GroEL from Escherichia coli) or nonidentical (as in chloroplast chaperonins) homoheptameric rings. By contrast, group II chaperonins consist of two identical eight- or ninemembered hetero-oligomeric rings that are made up of two types of subunits as in the thermosome,34 the archaeal chaperonin from Thermoplasma acidophilum, or eight types of subunits as in the case of the eukaryotic cytoplasmic chaperonin CCT (chaperonin-containing t-complex polypeptide 1 (TCP1); also called TRiC). The order of the eight subunits of CCT/ TRiC in the ring was established recently by determining which arrangement is most consistent with inter-residue distance restraints obtained using chemical cross-linking and mass spectrometry.35,36 This order was also found to give the best fit to crystallographic data for CCT/TRiC,37 thereby providing independent proof for the correct order and yielding a revised structure of this chaperonin.38 The crystal structures of GroEL,39,40 the thermosome,34 and yeast CCT/TRiC38 show that subunits of group I and II chaperonins have a similar architecture (Figure 3). Each subunit consists of three domains: (i) an equatorial domain that contains an ATP binding site and is involved in inter-ring contacts; (ii) an apical domain that is positioned at the opening of the central cavity and binds unfolded or misfolded protein substrates;41 and (iii) an intermediate domain that connects the apical and equatorial domains and has a crucial role in the

Figure 2. General scheme showing various combinations of concerted and sequential models of ATP binding and hydrolysis by a dimer. For example, conformational changes associated with both ATP binding and hydrolysis can be sequential (dashed blue pathway) as observed in the case of the bacteriophage φ29 packaging motor.21 Alternatively, both ATP binding-promoted conformational changes and hydrolysis can take place in a a concerted fashion (continuous blue pathway). Many other mechanisms are possible. 6590

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

There is much disagreement regarding the mechanisms by which chaperonins assist folding. Some studies suggest that the GroEL/ES system assists folding via a “passive” mechanism, i.e., substrate encapsulation in the GroEL/ES cavity leads to prevention of aggregation but does not result in changes in the folding energy landscape that can give rise to enhancement of folding rates.58,59 In other studies, however, it was found that substrate encapsulation in the GroEL/ES cavity results in acceleration of folding relative to the rate in bulk solution owing to confinement and the negatively charged surface of the cavity walls.60−62 Simulations have suggested that the charged surface of the cavity walls increases the surface water density, thereby favoring the burial of hydrophobic residues that accompanies folding.63 A recent experimental study showed, however, that the density and dynamics of the cavity-confined water are similar to those of bulk water, but only the upper region of the cavity was probed in that work.64 It has also been proposed that chaperonins assist protein folding by an iterative annealing mechanism, i.e., that they unfold kinetically trapped folding intermediates, thereby providing them with a further opportunity to fold.65 Evidence for this “active” mechanism of action was provided by Rye and co-workers, who showed that ATP binding to GroEL66 and the C-terminal tails of GroEL67 promote protein substrate unfolding and, thus, drives folding. By contrast, it was reported by others that GroEL- and CCT/ TRiC-mediated folding do not require ATP and in-cage confinement and that, in the case of GroEL, the role of ATP and GroES is only to promote the dissociation of sticky substrates that inhibit it if not removed.68 Apo GroEL also has unfolding activity as indicated, for example, by the recent report that it accelerates the interconversion rate between the native state and a well-defined folding intermediate of a triple mutant of Fyn SH3 by about 20-fold.69 Finally, GroES, ATP, and incage confinement are also not relevant in the case of assisted folding in the presence of the apical domain of GroEL by itself.70 In principle, the various conflicting mechanisms of chaperonin action are not mutually exclusive since a universal mechnism of action may not exist. Different protein substrates can exist in multiple unfolded, misfolded, or partly folded states (hereafter referred to collectively as nonfolded states) depending on the substrate and denaturing conditions, all of which have the potential to interact with GroEL differently. In other words, it is possible that different mechanisms operate depending on the substrate and the conditions. One important potential implication of the existence of a diversity of mechanisms would be that obligate substrates of GroEL might include proteins other than those found to be encapsulated in the GroE cavity.51,52 It should also be noted, however, that the properties of chaperonins may reflect not only the wide spectrum of client proteins but also selection for carrying out moonlighting functions that are not related to protein folding.71

Figure 3. Chaperonin structures. (a, b, and c) Side views of apo GroEL (PDB ID 4HEL), the GroEL−GroES bullet-shaped complex (PDB ID 1AON),45 and the GroEL−GroES2 football-shaped complex (PDB ID 4PKO),120 respectively. Also shown are (d) a side view of the closed form of CCT/TRiC (PDB ID 4V8R)38 and (e) a top view of the open form of CCT/TRiC in which the order of the subunits is given (PDB ID 4B2T).38 The apical (A), intermediate (I), and equatorial (E) domains of GroEL and CCT/TRiC are colored in red, green, and blue, respectively.

allosteric signaling. The folding function of group I chaperonins is aided by a cochaperonin, such as GroES in E. coli, that is a ring-shaped homoheptamer.42,43 ATP-dependent44 binding of GroES to the apical domains of GroEL forms a secluded compartment with hydrophilic walls45 in which substrate proteins as large as 70 kDa can fold in isolation from bulk solution. By contrast, group II chaperonins contain a so-called “helical protrusion”, a sequence located at the tip of their apical domains that provides a built-in lid instead of the GroES-like detachable lid that they lack.46 It was shown that the ATPdependent conformational changes and folding function of an archaeal chaperonin (Thermococcus sp. strain KS-1) are impaired upon deletion of its helical protrusions.47 It is still not clear what distinguishes chaperonin substrates from other proteins. Early studies showed that GroEL can bind in vitro to a multitude of E. coli proteins48 and also to artificial polypeptides with random sequences,49 thereby suggesting that it is very promiscuous. Back-of-the-envelope calculations indicated, however, that the amount of GroEL in an E. coli cell under normal conditions is enough to facilitate the folding of no more than 5% of the proteins within a cell.50 This conundrum was resolved by showing51 that only ∼250 E. coli proteins interact in vivo with GroEL under normal conditions out of which 84 (or only 49 according to Taguchi and coworkers52) are obligate substrates that require both GroEL and GroES for folding. Many of the obligate substrates are TIM barrels, but other TIM barrel proteins fold in a GroELindependent manner. Thus, it is still not understood why some proteins require GroEL/ES for folding and others do not.53 Clients of the CCT/TRiC system have also been identified, and they include β-actin,54 α- and β-tubulin,55 and a few hundred other proteins.56,57 Here, too, an understanding of the features that render proteins to be chaperonin dependent is still lacking.

2. EXPERIMENTAL ANALYSIS OF ALLOSTERY IN CHAPERONINS 2.1. Intra-Ring Allostery with Respect to ATP and Its Analogs

The first evidence for allostery in chaperonins was obtained by showing that plots of initial rates of ATP hydrolysis by GroEL, as a function of ATP concentration from 0 to 200 μM, are sigmoidal with a Hill coefficient of about 1.9.72 The ATPase reaction velocity was found in that study to display a linear 6591

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

chaperonins is that different types of subunits within a ring can possess different affinities for ATP as reported for CCT/ TRiC.83 Such site heterogeneity cannot be distinguished from negative intra-ring cooperativity in ATP binding (since in both cases the affinity decreases with the extent of binding) that would mask positive intra-ring cooperativity that exists owing to site−site interactions. In general, the weaker positive intra-ring cooperativity in group II chaperonins is likely to be due to its relatively weak intersubunit intra-ring interactions as reflected in the flexibility of their apical domains (Figure 4).84,85 The

dependence on GroEL concentration, thereby indicating that the observed cooperativity is not due to an ATP-promoted assembly (or disassembly) reaction that yields a more active species. Shortly afterwards, it was reported73 that plots of fractional saturation of GroEL by the product ADP, as a function of ATP concentration in the same range as before72 and in the presence of GroES, are also sigmoidal. The values of the Hill coefficient were estimated to be 2.8 and 1.5 for ATP and ADP, respectively. These results showed that the cooperativity is due to K system effects (i.e., to ATP binding and not hydrolysis), but it remained unclear whether the K system effects require the presence of GroES. This issue was resolved by labeling GroEL with pyrene-maleimide and monitoring the change in fluorescence as a function of nucleotide concentration.74 Plots of the extent of nucleotidepromoted pyrene fluorescence enhancement as a function of ATP, the ATP analog AMP-PNP, and ADP concentration, in the absence of GroES, were found to be sigmoidal with Hill coefficients of about 4, 3, and 3, respectively. These results indicated, therefore, that cooperativity in ATP binding and hydrolysis by GroEL, with respect to ATP, is indeed due to K system effects. Measurements of initial rates of ATP hydrolysis and nucleotide-promoted pyrene fluorescence enhancement, as a function of ATP concentration, were also carried out for a nonlabeled and pyrene-labeled single-ring version of GroEL, respectively.75 The single-ring version of GroEL, termed SR1, was designed based on the crystal structure of GroEL39 by introducing the mutations Arg452 → Glu, Glu461 → Ala, Ser463 → Ala, and Val464 → Ala that essentially abolish interring interactions.76 The Hill coefficients for initial rates of ATP hydrolysis by SR1 and ATP-promoted fluorescence enhancement of pyrenyl-SR1, both with respect to ATP, were found to be 2.87 (±0.16) and 2.5, respectively.75 A value of 2.5 was also obtained for the Hill coefficient for ATP-promoted fluorescence enhancement of pyrenyl−GroEL.75 These results provided additional evidence that cooperativity in GroEL is not due to an ATP-promoted assembly or diassembly reaction (e.g., a shift from an inactive double ring to an active single ring). Finally, evidence for K system effects was also provided by transient kinetic studies that showed there is a sigmoidal77 or bi-sigmoidal78 dependence of the value of the rate constant of the ATP-promoted conformational change of SR1 on ATP concentration. Taken together, all these studies established, therefore, that the heptameric ring of GroEL is a K system-type allosteric unit. The extent of intra-ring allostery appears to vary between group I and group II chaperonins. As discussed above, the group I homo-oligomeric chaperonin GroEL displays positive intra-ring cooperativity in ATP binding, with respect to ATP, but in the group II hetero-oligomeric CCT/TRiC79 and the homo-oligomeric archaeal chaperonin from Methanococcus maripaludis80 it is relatively weak, and in the hetero-oligomeric T. acidophilum thermosome it appears to be absent.81 In the case of bovine CCT/TRiC, for example, the value of the Hill coefficient for its allosteric transition at low ATP concentrations, in the presence of 50 mM K+ ions, is 2.00 (±0.25),79 which is lower than the corresponding value for GroEL of 2.41 (±0.13) that was determined under the same conditions.82 It should be noted that the value of the Hill coefficient for CCT/ TRiC is lower than that for GroEL, although the opposite might have been expected given that CCT/TRiC is made up of more subunits. One possible explanation for the weaker intraring positive cooperativity in the hetero-oligomeric group II

Figure 4. Top views of the apical and equatorial domains of GroEL and CCT/TRiC. Comparison of the top views of (a) the apical domains of apo GroEL (PDB ID 1OEL)195 and (b) the open state of CCT/TRiC (PDB ID 4B2T)38 shows that there are strong inter-apical domain interactions in GroEL that can account, in part, for its concerted intra-ring allosteric transitions. These interactions include inter-apical domain salt bridges between Glu255 (red) and Lys207 (green). The inter-apical domain interactions in CCT/TRiC are weak, thereby accounting for their flexibility and the nonconcerted intra-ring allosteric transitions of this chaperone. By contrast, strong interactions exist between the equatorial domains in both (c) apo GroEL (PDB ID 1OEL)195 and (d) the open state of CCT/TRiC (PDB ID 4B2T).38

sigmoidal dependence of the value of the rate constant of the ATP-promoted conformational change of CCT/TRiC on ATP concentration indicates that positive intra-ring allostery also in this chaperonin is due to K system effects.86 Several studies have established that the allosteric switch of chaperonins requires the γ-phosphate of ATP and that the switch is unusually selective with regard to the orientation of the γ-phosphate and the geometry and size of its analogs. Plots of the extent of the change in fluorescence of pyrene-labeled GroEL as a function of the concentration of ADP−AlFx, ADP− BeFx, and ADP−GaFx (x indicates that the number of bound fluoride ions can vary) were found to be sigmoidal. In addition, the SAXS patterns of GroEL in complex with these analogs were found to be similar to that of GroEL during steady-state ATP hydrolysis. Taken together, these data indicated that these analogs are able to effect the allosteric switch of GroEL.87 ADP−AlFx can also effect the allosteric switch of group II 6592

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

2.3. Inter-Ring Allostery in ATP and GroES Binding and the Nested Model

chaperonins as indicated by (i) the crystal structure of the thermosome in complex with ADP and AlFx34 and (ii) the finding that ADP and metal fluorides induce the transition of CCT/TRiC from a high- to low-affinity state for protein substrates (i.e., from R to T as discussed below).88 By contrast, ScFx and Vi in complex with ADP were found to be unable to promote GroEL’s allosteric switch possibly owing to their larger size and lack of tetrahedral geometry. ATPγS and AMP-PNP were also found to be unable to effect GroEL’s allosteric switch, thus indicating that it is also sensitive to the larger ionic radius of S relative to O and the slightly different angle of Pβ−NH− Pγ.87 Remarkably, addition of AlFx or BeFx but not Vi to a complex of SR1−ADP−GroES with an encapsulated protein substrate, rhodanese, was found to trigger productive folding, thereby highlighting the importance of the γ-phosphate of ATP and the ability of metal fluorides to mimic it.89 The latter work also highlighted the importance of intra-ring allostery for the folding function as will be discussed later on.

Plots of initial rates of ATP hydrolysis by the Arg197 → Ala GroEL mutant, as a function of ATP concentration from 0 to 100 μM, were found to be biphasic96 and not monophasic (i.e., one sigmoid) as in the studies of wild-type GroEL reported before.72−74 The first phase, at very low ATP concentrations, corresponds to a hyperbolic increase in the rate of ATP hydrolysis, and the second phase corresponds to a decrease in activity that plateaus at higher ATP concentrations (Figure 5).

2.2. Intra-Ring Allostery with Respect to K+ Ions

The large spread in the values of the Hill coefficients that were determined in the early measurements72−75 of cooperativity in ATP binding and hydrolysis by GroEL indicated that it is sensitive to factors that varied between these studies. An advantage of the MWC model is that it can easily account for how cooperativity in the binding of one ligand (e.g., ATP), with respect to that ligand, is affected by the presence of other types of ligands termed allosteric effectors. In the case of exclusive binding of a ligand (e.g., ATP) to the R state, increasing concentrations of an allosteric effector that binds to the T or R states will increase or decrease cooperativity, respectively. In the case of nonexclusive binding of the ligand (ATP) to the R state, increasing concentrations of an allosteric effector that binds preferentially to the T state will first increase and then decrease the extent of cooperativity.90 One of the ligands that affects allostery in GroEL is the K+ ion. The ATPase activity of GroEL is NH4+ or K+ dependent91 and is positively cooperative with respect to K+ concentration.92 Owing to symmetry in coupling relations, there is also an effect of K+ ions on cooperativity in ATP binding by GroEL, with respect to ATP, which decreases with increasing K+ concentration.92 Analysis of the effects of K+ ions on ATP hydrolysis by SR1 established that K+ increases the affinity of GroEL for ATP without affecting the rate of hydrolysis.93 This finding is consistent with structural work in which K+ was found to be in contact with the α phosphate of the nucleotide.94 In addition to this K+ binding site, another site between subunits and near the ring−ring interface was identified in crystallographic work in which thallium was used to reveal potential K+ binding sites.95 Hence, it appears that K+ may affect allostery in GroEL by altering both c (= KT/KR) via binding to the nucleotide binding site and L (= [T]/[R]) via binding at the intersubunit interface. Some of the variation in the estimates of the Hill coefficient in early studies can, therefore, be attributed to differences in K+ concentrations which were 10 mM in some studies72,73 and 50 mM in others.74 Other sources of variability may have been the contamination of solutions of ATP (or its analogs) by ADP and the presence of nonfolded substrates in the GroEL preparations. The allosteric effects of nonfolded proteins and ADP will be discussed below.

Figure 5. Role of the Arg197−Glu386 salt bridge in allosteric communication in GroEL. (a) In the T state of GroEL, Arg197 in the apical domain of one subunit forms a salt bridge with Glu386 in the intermediate domain of an adjacent subunit in the same ring.195 (b) Measurements of initial rates of ATP hydrolysis by the Arg197 → Ala GroEL mutant, as a function of ATP concentration and in the presence of 10 mM K+ ions, reveal two phases corresponding to the allosteric transitions of the two rings. Only one sigmoidal phase is observed when one of the rings is GroES-bound. The results show that breaking the Arg197−Glu386 salt bridge weakens both positive intraring and negative inter-ring cooperativities. The weakened positive cooperativity is reflected in the absence of sigmoidicity and midpoints at low ATP concentrations ( L′2 (= [TR·ES]/[RR·ES])).107 Given that nonfolded proteins have a lower affinity for the R state (see below), such an allosteric effect can lead to protein substrate release from the ring distal to GroES, thereby providing a mechanism for facilitating the folding of proteins that are too large to be encapsulated in the cavity of the cis ring.108 It also follows from L2 > L′2 that stabilization of the trans ring T state, for example, upon protein substrate binding93 or by mutagenesis,109 would accelerate GroES release from the cis ring as indeed has been observed. The opposite effect, i.e. slower release of GroES, results upon stabilization of the R state by mutation.110 The nested model was also able to account for the observations that Dixon plots of the inverse of initial ATPase

mutant, both transitions are observed at low ATP concentrations because the mutation destabilizes the T state and, therefore, less ATP is required to shift the equilibrium toward the R state. Hence, it was predicted that such biphasic behavior should also be observed in the case of wild-type GroEL by extending the range of ATP concentrations at which initial rates of ATP hydrolysis are measured (to 0.8 mM), as was indeed found to be the case.97 A biphasic dependence of initial rates of ATP hydrolysis on ATP concentration has also been observed for CCT/TRiC,79 the archaeal chaperonin Mm-cpn,80 and the thermosome,81 and it, therefore, seems to be a universal feature of chaperonins. Biphasic behavior is also seen in transient kinetic measurements of the value of the observed rate constant of the allosteric transitions of CCT/TRiC86 and GroEL mutants98−100 as a function of ATP concentration. In the case of GroEL, which lacks tryptophan residues, different tryptophan substitutions were introduced so that the rate constant of the allosteric transition could be determined by monitoring the change in fluorescence as a function of time after rapid mixing with ATP. The transient kinetic data indicated that the observed biphasic behavior is not due to some form of substrate inhibition or activation.101 The assignment of each phase to the respective allosteric transition of one of the rings is supported by the monophasic behavior observed in the ATPase activity of GroEL when one of its rings is GroES-bound and, thus, not hydrolyzing ATP72 and by the monophasic transient kinetic data for SR1.77 It is also supported by experiments with a chimeric chaperonin (that contains the sequence of E. coli GroEL from position 1 to 364 and the sequence of Cpn60-1 from the bacterium Rhizobium leguminosarum from position 365 to 547), which is in equilibrium between single and double rings. Initial rates of ATP hydrolysis by this chimera, as a function of ATP concentration, are monophasic (one sigmoid) at low concentrations of the chimera but biphasic (bi-sigmoidal) when it is present at higher concentrations.102 A nested allosteric model was put forward to account for the biphasic plots of initial rates of ATP hydrolysis by GroEL as a function of ATP concentration.97 Nested allosteric models were first developed to describe certain linkage phenomena in hemoglobin103 but are particularly useful for describing allostery in large systems with hierarchical structure such as chaperonins. The hierarchical structure in such systems suggests that a corresponding hierarchy in allosteric interactions may also exist. In the nested model for GroEL (Figure 6), each ring is in equilibrium between tense (T) and relaxed (R) states with relatively low and high affinities for ATP, respectively, in accordance with the MWC representation. In the presence of increasing concentrations of ATP, the GroEL double ring

Figure 6. Scheme for the different allosteric states of GroEL according to the nested model. Rings in the T state have low affinity for ATP and high affinity for nonfolded substrates, whereas rings in the R state have high affinity for ATP and low affinity for nonfolded substrates. In the absence of ligands, GroEL is predominantly in the TT state. In the presence of ATP, the equilibrium is shifted toward the TR and RR states. 6594

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

lowers the extent of cooperativity. The similar values obtained from the transient and steady-state kinetic data, therefore, indicate that the transitions are concerted. Finally, native mass spectrometry was used recently to measure the populations of all coexisting states of GroEL, which differ in the number of bound ATP molecules.126 Given these distributions, it was possible to determine the values of the 14 ATP binding constants of GroEL and show that they are consistent with the predictions of the nested model. CCT/TRiC also displays nested allostery, but unlike GroEL, its intra-ring allosteric transitions appear to be sequential and not concerted. Plots of the value of the rate constant for the intra-ring allosteric switch of yeast CCT/TRiC as a function of ATP concentration were found to deviate from a bi-sigmoidal curve. A linear phase was observed at low ATP concentrations instead of the lag phase that is characteristic of such curves.127 These data were found to be consistent with a model in which the CCT/TRiC ring undergoes two parallel and sequential allosteric transitions. Electron microscopy analysis of bovine CCT/TRiC labeled with subunit-specific monoclonal antibodies enabled image averaging without imposing 8-fold symmetry.84 This work revealed that the apo state of CCT/ TRiC has considerable conformational heterogeneity that decreases with increasing ATP concentrations, and it suggested that the ATP-induced conformational changes spread around the ring in a sequential fashion. It was suggested that the existence of a sequential allosteric mechanism may also be indicated by a hierarchy in the magnitude of the phenotypic effects caused by mutations in the different ATP binding sites of CCT/TRiC.128 Subsequently, such a hierarchy in the effects of mutations introduced in the ATP binding site of each of the yeast CCT/TRiC’s subunits, in turn, was found to correspond to the order of the subunits in the ring.129 The hierarchy in the mutational effects was also found to be in good agreement with the rank order of affinities for ATP of the different subunits,83 thereby suggesting that sequential allostery in CTT/TRiC may be due, in part, to the differences in affinities of the subunits. It is still unclear, however, whether the sequential conformational change in CCT/TRiC has a fixed starting point(s) and direction around the ring or multiple pathways that may depend, for example, on the identity of the bound protein substrate. Sequential conformational changes may also occur in other group II chaperonins. For example, construction of a Thermococcus chaperonin complex that consists of fused wildtype and mutant subunits (defective in ATP hydrolysis or ATPinduced conformational changes) in different orders showed that greater impairment in function is caused when the mutant subunits are adjacent to each other.130 Finally, it is important to note that ATP hydrolysis appears to have different roles in group I and II chaperonins. In group I chaperonins, it appears to be needed for facilitating and timing the repeated cycling between their conformational states, whereas in the case of group II chaperonins, there is evidence that ATP hydrolysis causes conformational changes that are essential for their folding function.130,131

Figure 7. Scheme for different states of GroEL in the presence of ATP and GroES. In the absence of ATP, GroEL is mainly in the TT state. In the presence of ATP, the equilibrium is shifted toward the TR and RR states (L1 = [TT]/[TR] and L2 = [TR]/[RR]). GroES (designated by ES) binds to a ring in the R conformation of the TR and RR states with affinities K1 = [TR][ES]/[TR·ES] and K2 = [RR][ES]/[RR·ES], respectively. The symmetric football-shaped species is not included in this scheme. A measure of the effect of GroES on the allosteric transition of the distal (i.e., without bound GroES) ring of GroEL is given by ΔΔG = −RT ln(L′2/L2), where L′2 = [TR·ES]/[RR·ES].

velocity vs ADP concentration are biphasic111 and that GroEL seemed to display half-of-sites reactivity with respect to GroES binding.92 Given that GroES binding to GroEL is ATP dependent,44 negative inter-ring cooperativity in ATP binding is expected to favor formation of an asymmetric GroEL−GroES complex in which GroES is bound to only one of the GroEL rings. This bullet-shaped complex was already visualized in early studies of GroEL 112,113 and the Thermus thermophilus chaperonin114 and is considered by many to be the proteinfolding active species in the reaction cycle of GroEL.115 Football-shaped complexes in which GroES is bound to both rings of GroEL were, however, also visualized more than two decades ago,116−118 and several crystal structures of them have become available recently.119−121 Football formation requires overcoming the inter-ring negative allostery and is, therefore, promoted by the relatively high concentrations of ATP and K+ ions that favor the R state and were employed in these studies.119−121 The principle of Occam’s razor was behind the assumption in the nested model that the intra-ring allosteric transitions of GroEL are concerted, but there was little evidence to support this at the time the model was put forward. Several lines of evidence now indicate that the intra-ring allosteric transitions of GroEL are indeed concerted. Targeted molecular dynamics simulations showed that the intra-ring allosteric transitions are concerted owing to steric hindrance, i.e., one subunit cannot switch from its T- to R-state conformation if the other subunits do not switch as well.122 It was suggested that the intra-ring allosteric transitions of GroEL can, therefore, be described as t7 → r7 (where t and r stand for the respective conformations of a subunit in the T and R states of a ring) since they consist of large coupled tertiary structural changes but no quaternary transition that involves significant relative motions of the subunits.123 In accordance with these simulations, it was shown that locking a single subunit in the t conformation (by generating the Asp83 → Cys and Lys327 → Cys double mutant and introducing an intrasubunit disulfide cross-link at these positions) is sufficient to block the T → R transition.124 An additional line of evidence was provided by the finding that the values of the Hill coefficients for wild-type GroEL and several mutants derived from steady-state data (i.e., initial rates of ATP hydrolysis as a function of ATP concentration) are the same as those obtained for the respective GroEL variants from transient kinetic data (i.e., rate constants of the T → R switch as a function of ATP concentration).125 In general, the presence of intermediates (as in the case of sequential allosteric transitions)

2.4. Allosteric Effects of ADP

Does ADP bind preferentially and, thus, stabilize the T or R states of GroEL rings? The data indicating that ADP binds to GroEL in a noncooperative manner75 are consistent with either preferential binding of ADP to the T state or with ADP having similar affinities for the T and R states. It was shown, however, that low concentrations of ADP can stimulate ATP hydrolysis 6595

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

by GroEL,24 thereby suggesting that ADP binds preferentially to the R state. These seemingly conflicting results can be reconciled by assuming that ADP binds to a third, D, state, which is less stable than the T state (i.e., L′ (= [R]/[D]) > L (= [R]/[T])) but has the same affinity for ADP as the T state. In such a case, ADP binding to GroEL will be noncooperative but will still shift the equilibrium toward the R state. Despite lacking positive intra-ring cooperativity in ADP binding, GroEL does display strong inter-ring negative cooperativity in binding of ADP.132 Owing to this negative inter-ring allostery, ATP hydrolysis in one ring is inhibited by the presence of ADP in the other ring.133 The decrease in ATPase activity at high concentrations of ATP (as first observed in the case of the Arg197 → Ala mutant (Figure 5)) is, therefore, due to negative inter-ring allostery with respect to ADP that is enforced by the slow off rate of ADP. Evidence for this mechanism has also been provided by the finding that the Glu257 → Ala GroEL mutant, which displays a further sigmoidal increase (and not decrease) in ATPase activity at high concentrations of ATP, has a fast off rate for ADP.134 ATP hydrolysis by CCT/TRiC also displays a further sigmoidal increase (and not a decrease) in ATPase activity at high concentrations of ATP,79 thereby indicating that inter-ring cooperativity in ADP binding by CCT/TRiC is weak or that it has an off rate for ADP that is not rate limiting.

hydrolysis in one ring by stabilizing the T state of the opposite ring, thereby increasing its off rate for ADP. The finding that the mutation Glu257 → Ala in GroEL increases the off rate of ADP and stimulates ATP hydrolysis134 is interesting in this regard since this mutation is close to the protein substrate binding site and thus may mimic the effect of substrate binding. Evidence that ATP hydrolysis is stimulated by stabilizing the T state was recently provided by labeling GroEL with tetramethylrhodamine (TMR) at position 242 in the apical domain near the protein substrate-binding site.137 This position was chosen so that TMR molecules attached to adjacent subunits can form noncovalently “stacked” dimers in the T state but not in the R state. It was found that formation of dimers in the T state, which tethers pairs of GroEL subunits to each other in a manner that mimics intersubunit tethering by nonfolded proteins, stimulates ATP hydrolysis, whereas the labeling by itself has no effect. In the case of CCT/TRiC, a relatively high intrinsic off rate of ADP can explain why its rate of ATP hydrolysis displays a further sigmoidal increase (and not decrease) at high concentrations of ATP79 and can also account for the fact that its ATPase activity is not stimulated by nonfolded substrates.

2.5. Allosteric Effects of Nonfolded Protein Substrates

3.1. Structural Basis of Intra-Ring Allostery

Initial rates of ATP hydrolysis by GroEL were measured as a function of the concentration of an unfolded protein substrate, reduced and Ca2+-depleted α-lactalbumin, at two fixed concentrations of ATP, 100 and 500 μM.82 In the presence of 100 μM ATP, the ATPase activity was found to increase in a hyperbolic fashion with increasing concentrations of the unfolded protein substrate, thereby indicating that nonfolded protein substrates stimulate GroEL’s rate of ATP hydrolysis, as had also been reported before.135 In the presence of 500 μM ATP, the ATPase activity was found to first increase and then decrease in a hyperbolic fashion as a function of the concentration of the unfolded protein substrate. This biphasic behavior, which was reminiscent of the biphasic plots of ATPase activity of GroEL measured as a function of ATP concentration,97 indicated that nonfolded protein substrates and ATP shift the equilibrium in opposite directions (Figure 6). In other words, nonfolded protein substrates favor the T state, the acceptor state for protein substrates, whereas ATP shifts the equilibrium toward the R state, in agreement with the finding that ATP lowers the affinity of GroEL for protein substrates.136 Structural work that followed45 showed that the lining of the cavity of the T state is continuous and hydrophobic and thus has high affinity for nonfolded substrates. By contrast, the lining of the cavity in the R-like GroES-bound state is hydrophlic and thus promotes protein substrate release and folding. The extent of cooperativity in ATP hydrolysis by GroEL with respect to ATP was determined for the TT → TR transition and found to first increase and then decrease as a function of the concentration of nonfolded α-lactalbumin.82 Given that nonfolded substrates stabilize the T state, such non-monotonic behavior indicated90 that ATP does not bind exclusively to the R state of GroEL. The stimulation of the ATPase activity of GroEL by nonfolded protein substrates is not due to a V system effect since nonfolded protein substrates do not enhance the rate of ATP hydrolysis by SR1.93 Instead, it is likely that they stimulate

A fundamental question in the field of allostery concerns the mechanism(s) by which structural changes are propagated from one ligand binding site in the protein to other distant sites. Information regarding the order and structures of intermediates between the T and the R states is needed to address this question but is usually difficult to obtain since such intermediates tend to be short-lived. One experimental approach to this problem is to use phi-value analysis first developed to characterize protein folding intermediates.138 In such an analysis, information about structures of transition and intermediate states is obtained by combining equilbrium data regarding the relative stabilities of the T and R states of the wild-type protein and a mutant (i.e., their allosteric constants, L) with transient kinetic data regarding the respective rate constants of their T → R transitions. Using this approach, it was shown that the Arg197−Glu386 salt bridge between subunits in a GroEL ring (Figure 5) is already broken in the transition state of the T → R allosteric switch.125 A phi-value type of analysis also revealed that there are two parallel pathways for the T → R transition of each ring,139 one that dominates at low ATP concentrations and a second that dominates at higher ATP concentrations (although this has been disputed140). Structural features of the transition states corresponding to these pathways remain unknown. A different approach for characterizing the pathway of GroEL’s T → R transition was based on cryo-EM reconstruction of various coexisting states of the ATP hydrolysis-defective Asp398 → Ala mutant141 following rapid mixing with ATP and freezing.142 The states were then ordered so that they formed a trajectory that showed progressive structural changes. This analysis revealed three intermediate states that follow the T state. The first state, termed Rs1, is produced by 35° sideways en bloc tilting of the intermediate and apical domains at the bottom hinge of the intermediate domains. The resulting downward motion of helix M (residues 386−409 in the intermediate domain) that was also seen

3. STRUCTURAL BASIS OF ALLOSTERY IN CHAPERONINS

6596

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

before143 brings the carboxylate oxygen of the catalytic residue Asp398 to the ATP-bound Mg2+ ion coordination cage.45 Given that this motion is required for hydrolysis, it is still unclear how the T state can be ATP hydrolysis active unless fluctuations allow the Rs1 conformation to be readily visited from the T state. The downward motion of helix M is accompanied by the breaking of the Arg197−Glu386 salt bridge, in agreement with the kinetic studies that showed that this is an early event. It was reported that the broken salt bridges of Glu386 with the apical residue Arg197 (Figure 5) are replaced by new salt bridges between Glu386 and the equatorial residue Lys80, thus reflecting the downward motion of the intermediate domain during this stage. In addition to the Arg197−Glu386 salt bridge, another intersubunit salt bridge between the apical residues Lys207 and Glu255 is broken and reportedly replaced by a new Lys245−Glu255 salt bridge. It should be noted, however, that salt bridges between Lys245 and Glu255 and between Lys80 and Glu386 are not observed in the crystal structure of an R state determined recently at 2.7 Å resolution.110 The structural consequences of breaking the Arg197−Glu386 salt bridges were visualized in an earlier low-resolution (30 Å) cryo-EM reconstruction of the apo state of the Arg197 → Ala mutant which showed loosening of the rings owing to the removal of these T state-stabilizing interactions.104 The Arg197 → Ala mutation was also used in combination with the Asp83 → Ala mutation in the equatorial domain (which breaks an intrasubunit salt bridge with Asp327 in the apical domain) to stabilize an R-like state of GroEL that was crystallized in complex with ADP.110 Formation of Rs1 may be reflected in the fast kinetic phase seen in stopped-flow studies99,100 (with an apparent rate constant whose value is greater than 300 s−1 at 300 μM ATP). It is also possible, however, that a second slower phase with a rate constant whose value displays a bi-sigmoidal dependence on ATP concentration98−100 corresponds to formation of Rs1 given that the structural changes that accompany this transition are highly cooperative. A second intermediate state, Rs2, is produced by a small rigid-body elevation of the apical domains at the top hinge of the intermediate domain during which no salt bridges are broken. The third state, Rs-open, is then formed upon a further elevation of 20° and radial outward movement of the apical domains. The transition from Rs2 to Rs-open is accompanied by breaking of the apical salt bridges, thereby freeing the apical domains and enabling them to move independently from each other. These results are in accord with the crystal structure of the R state in complex with ADP in which the equatorial domains display almost perfect 7-fold symmetry whereas the apical domains display strong asymmetry.110 The outward radial motion of the apical domains position them so that they can interact with the mobile loops of GroES.45,144 Interestingly, asymmetry in the apical domains has also been observed recently in the complex of the human mitochondrial chaperonin with its cochaperonin,121 but it is not seen in the GroEL−GroES complex.45 Taniguchi and co-workers assigned the fast phase in their study to formation of an intermediate that can bind GroES, i.e., to formation of Rs-open and not Rs1.100 Hence, the correspondence between the different phases in the kinetic studies and the trajectory suggested in the cryo-EM study is not clear. Targeted122 and unbiased145 MD simulations have also shown that the T → R transition of GroEL begins with a downward movement of helix M that is associated with breaking of the Arg197−Glu386 salt bridge. The unbiased MD

simulations also showed that the downward motion of helix M involves breaking of the Asp155−Arg395 intrasubunit salt bridge that anchors helix M to helix G (residues 155−169 in the intermediate domain). Structural and kinetic analysis showed that the mutation Asp155 → Ala converts the T → R transition of GroEL from concerted to sequential.146 The targeted MD simulations indicated that, owing to steric hindrance, one subunit cannot switch from the t to r conformation without the other subunits in the ring switching as well, thereby accounting for the concertedness of the T → R transitions. Such steric hindrance was, however, not observed in the unbiased MD study upon aligning conformations from single subunit simulations with model heptamer rings.145 The analysis of the Asp155 → Ala mutant suggests that the concertedness may also be due to the coupling between intraand intersubunit salt-bridge networks.146 The early events of the T → R transition were also observed in coarse-grained modeling studies such as anisotropic network model-based simulations of the full GroEL−GroES complex147 and in Brownian dynamic simulations of the GroEL double ring.148 The latter study also showed that rupture of the intrasubunit Asp83−Lys327 salt bridge takes place almost simultaneously with breaking of the Arg197−Glu386 salt bridge and that the apical domains become flexible. In another coarsegrained modeling study based on elastic network models, various residues that have an important role in the T → R transition were identified.149 These residues include the following: (i) Asp83 and Lys327 that were also identified as important before;124,148 (ii) Arg58 in the equatorial domain that was also found to be part of an allosteric network in a correlated mutation analysis;150 and (iii) the apical domain residues Gly244 and Glu209 that were not implicated elsewhere and whose roles remain to be studied experimentally. An additional salt bridge formed during the T → R transition between the equatorial residues Lys34 and Glu483 was identified in another unbiased MD simulation of a single subunit.151 The latter study also showed that the structural changes that accompany the T → R transition are encoded in the dynamics of the single subunits. Binding of GroES to a GroEL ring in the R state promotes a further conformational change designated by R → R′. The transition that follows next, which is associated with ATP hydrolysis in the GroES-bound (cis) ring, is designated by R′ → R″. Earlier work reported that after being released from their initial salt-bridge constraints the apical domains twist counterclockwise by ∼25° during the T → R transition.143 The R → R′ transition induced by GroES binding then causes a net ∼115° clockwise twisting of the apical domains. The counterclockwise twisting in the T → R transition and clockwise twisting in the R → R′ transition have also been observed in simulations.148 In more recent structural studies of the R state, however, such counterclockwise twisting has not been observed.110,142 Hence, it appears that only twisting by ∼100° clockwise in the R → R′ transition occurs, which is accompanied by a further elevation of the apical domains. During the R → R″ transition, outside-in concerted movements of the apical helices K (residues 340− 354) and L (residues 365−373) take place. These helices, which tilt by ∼30° during the T → R transition, rotate by an additional 40° during the R → R″ transition.148 The structural changes that occur during the R → R′ (or R → R″) transition lead to a doubling of the size of the central cavity and to a change in the polarity of its surface that is due, in part, to polar and charged residues that, in the T state, are exposed to the 6597

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

Figure 8. Inter-ring communication in GroEL and CCT/TRiC. (a) In GroEL (PDB ID 4HEL), each subunit in one ring is in contact with two neighboring subunits in the opposite ring. Signaling between ATP binding sites in opposite rings can occur via a cascade of changes in electrostatic interactions. One is between the γ-phosphate of ATP (the Asp87 residues in the ATP binding sites are shown in orange) with the positive charge at the N-cap residue of helix D (shown in green), Gly88, that is induced by the helix dipole. A second one is between the helix dipole-induced negative charge at the C-cap residue of helix D, Ala109, with the positive charge of the ε-NH3+ group of Lys105 in helix D of the subunit in the opposite ring. The distance between the Nε atom of Lys105 and the backbone O atom of Ala109 is 5.6 Å. (b) The in-register stacking of subunits in CCT/TRiC (PDB ID 4B2T) is shown for subunits CCT1 (top) and CCT7 (bottom). Owing to the in-register stacking, the helices in CCT/TRiC, which correspond to helix D in the two GroEL rings, are relatively far removed from each other and, thus, unlikely to facilitate inter-ring communication. The inter-ring distance between the Cβ atoms of Ala109, the C-cap residue of helix D in GroEL, is 4 Å, whereas the inter-ring distance between the Cβ atoms of the corresponding C-cap residues in CCT/TRiC, Gln110 in CCT1 and Glu113 in CCT7, is 20.4 Å and, thus, much larger.

exterior. The R → R″ transition involves formation of an interdomain salt bridge between Asp359 and Lys80 that in the R state is salt linked to Glu386.148 The work of Thirumalai and co-workers149 revealed additional residues that are involved in the transition to the R″ state that include Val174 and Glu191. Interestingly, the mutation Glu191 → Gly interferes with GroES binding to GroEL, and its effect can be suppressed by the mutation Val174 → Phe.152 The structural basis of intra-ring allostery in group II chaperonins remains poorly understood, in part, because highresolution structures of their different allosteric states are not available or have become available only recently.34,36−38,85,153 In addition, the high-resolution CCT/TRiC structures contain bound protein substrates, and it is, therefore, not clear whether they can be assigned to particular allosteric states. Insights into the allosteric mechanism of group II chaperonins from mutational analysis are also lacking. In fact, only the Gly345 → Asp mutation in subunit 4 of CCT/TRiC has, to date, been analyzed in some detail with regard to its effects on allosteric transitions.127 Nevertheless, it seems likely that intra-ring allostery, with respect to ATP, is weaker or absent in group II chaperonins because of the weak interactions that exist between their apical domains in the apo state as reflected in their flexibility (Figure 4).84 Normal mode analysis of the thermosome showed that multiple modes are required to describe its transition from the R″ to the T state,154 in line with its greater flexibility compared to GroEL for which a single mode was shown to provide the dominant contribution.155 The principal modes of the thermosome ring were found to be characterized by weaker long-range intersubunit correlations than those found in the dominant mode of the GroEL ring, in agreement with the weaker intra-ring cooperativity in the thermosome seen by experiment.

One inter-ring contact site involves residues Glu461, Arg452, and Val464 in a subunit in one (top) ring with the same residues in a subunit in the opposite (bottom) ring. The second inter-ring contact site involves Ala109 and Lys105 in the same subunit in the top ring and these residues in an adjacent subunit in the bottom ring. The mutation Glu461→ Lys in GroEL was found to convert the stacking from staggered to inregister,156,157 i.e., each subunit in one ring is in contact with only one subunit in the opposite ring as observed in group II chaperonins.34,37 Both positive intra-ring and negative interring cooperativities, with respect to ATP, were found to be abolished in this mutant as indicated by the monophasic and nonsigmoidal dependence of its ATPase activity as a function of ATP concentration.156,158 Negative inter-ring allostery, with respect to ATP, is also disrupted in the Arg13 → Gly and Ala126 → Val double mutant of GroEL with known crystal structure,39 but the structural basis for this is not clear.159 The effects of mutations such as Glu461→ Lys156,158 and Arg197→ Ala96 on both positive intra-ring and negative inter-ring cooperativities, with respect to ATP, show that the allosteric networks involved in both types of cooperativity are coupled. In agreement with these results, elastic network model analysis showed that a single dominant normal mode can describe the dynamic changes that underlie both the positive intra-ring and the negative inter-ring cooperativities.155 The structural basis of inter-ring cooperativity in GroEL is likely to involve helix D (residues 89−109 in the equatorial domain) which extends from the ATP binding site to the Ala109−Lys105 ring−ring contact site. Hence, information regarding the nucleotide occupancy of a subunit in one ring can be transmitted to the nucleotide binding site in a contacting subunit in the opposite ring via helix D in the two subunits (Figure 8).160 It is possible that such communication is facilitated by electrostatic interactions between (i) the γphosphate of ATP with the positive charge at the N-cap residue of helix D, Gly88, that is induced by the helix dipole and (ii) the helix dipole-induced negative charge at the C-cap residue of helix D, Ala109, with the positive charge of the ε-NH3+ group

3.2. Structural Basis of Inter-Ring Allostery

In GroEL, the back-to-back stacking of the two rings is staggered such that each subunit in one ring is in contact with two neighboring subunits in the opposite ring (Figure 8).39 6598

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

onins is, therefore, expected to be very different from that of GroEL.

of Lys105 in helix D of the subunit in the opposite ring (Figure 8). Owing to the interaction between the γ-phosphate of ATP with the positive charge at the N-cap of helix D, ATP binding to both rings would draw helices D away from each other and weaken the inter-ring interaction.110 Alternatively, owing to interactions (i) and (ii) above, ATP binding to a subunit in one ring would draw the N-cap of helix D in the subunit of the other ring away from the ATP binding site, thereby reducing its affinity for ATP. Both mechanisms would lead to inter-ring negative cooperativity in ATP binding and, thus, favor asymmetric binding of ATP to one ring and ADP to the other ring. A similar mechanism could also explain inter-ring negative cooperativity in ADP binding133 if we assume that the α- and/or β-phosphates of ADP also interact with the positive charge at the N-cap of helix D. ATP hydrolysis in the GroES-bound cis ring leads to structural changes other than those at the ATP binding sites of the trans ring. The most striking change is found between the equatorial domains of the trans ring, where the main intra-ring contact, an intersubunit β-sheet interaction between residues 4−25, 37−51, and 519−523, is disrupted in the ADP-bound complex.160 This change has been attributed160 to pivoting of the trans equatorial domains about the fixed Glu461−Arg452 contact that accompanies the closer approach between the D helices upon ATP hydrolysis described above. As a result, the trans apical domains become more expanded and slightly less twisted, thereby increasing the affinity of this ring for protein substrates. The change in the intersubunit β-sheet interaction may explain why the Cys519 → Ser GroEL mutant dissociates into monomers in the presence of 5 mM ADP.161 It is also consistent with the stronger intra-ring coupling of the equatorial domains in the cis ring compared to the trans ring revealed in a study of the Markov propagation of information through the network of interactions in the GroEL−GroES complex.162 The latter study also identified two pathways of communication between the ATP binding sites in the trans ring and GroES bound to the cis ring (that may be related to the two pathways identified by phi-value analysis139) and showed that an opposite direction of information flow exists within cis and trans rings, consistent with negative inter-ring cooperativity and other previous coarse-grained simulations.163 Inter-ring allostery has also been observed with regard to protein substrate binding. Fisher and co-workers showed that binding of nonfolded glutamine synthetase to one ring of GroEL causes substantial structural changes in the opposite unoccupied ring.164 They found that both the cis and the trans apical domains rotate counterclockwise, thereby decreasing the affinity for GroES. These structural data help to explain why stabilization of the trans ring T state upon protein substrate binding accelerates GroES release from the cis ring, as has been observed.93 Regarding group II chaperonins, normal-mode analysis of the thermosome identified residues in the stem loop of the equatorial domain that are important for inter-ring communication and revealed coupling between equatorial and apical domain residues.154 Hence, protein substrates may affect inter-ring allostery also in group II chaperonins. Little else is known, however, about the structural basis of inter-ring communication in group II chaperonins. Given that the backto-back stacking of the subunits in group II chaperonins is in register, inter-ring communication via the helices that correspond to helix D in GroEL is unlikely (Figure 8). The mechanism of inter-ring communication in group II chaper-

4. ROLE OF ALLOSTERY IN CHAPERONIN FUNCTION 4.1. Intra-Ring Allostery in Chaperonin Function

Allostery provides a mechanism for switching between different functional states. In hemoglobin, for example, it facilitates the transition between oxygen binding and release states. In chaperonins, intra-ring allostery facilitates the switching between the binding (T) and release (R) states of protein substrates. The value of the Hill constant for the T → R transition of GroEL is ∼2.5 (ref 97) and, thus, relativly low for an assembly of seven subunits. Similar values have been found for hemoglobin165 and yeast glyceraldehyde-3-phosphate dehydrogenase,166 although they are tetramers. This relatively low value of the Hill constant probably reflects evolutionary optimization of both protein substrate binding, which favors the T state and thus pushed its value up, and protein substrate release from the R state, which pushed its value down. A negative linear relationship was found between the value of the rate constant of GroEL-mediated mouse DHFR refolding and the value of the Hill coefficient, thereby indicating that stabilization of the T state can indeed retard folding.167 These experimental results were found to be in agreement with simulations of folding of a lattice chain of single-bead residues in a confined environment.168 Folding times were calculated as a function of the hydrophobic−hydrophilic cycle time of the cavity walls for different fractions of the overall time in which the cavity walls are hydrophobic. This fraction is directly related to the allosteric equilibrium constant for the T → R transition and, thus, to the extent of intra-ring positive cooperativity. In the case of cycling times that are comparable to the folding times of the lattice protein, an inverse correlation was found between the folding rate and the fraction of time in which the cavity walls are hydrophobic. A more recent kinetic modeling study by Thirumalai and co-workers indicated, however, that the allosteric constant of the T → R transition has only a small effect on the efficiency of GroEL (using parameters for Rubisco) and that the main effect is due to the rate of the R → R″ transition.169 It should be noted, however, that GroES is not obligatory for GroEL-assisted folding of mouse DHFR whereas folding of Rubisco requires the full GroE system. It has been suggested that the intra-ring allosteric transitions of GroEL have evolved to be concerted in order to facilitate synchronized release of the different parts of a substrate that are GroEL bound. Such release would increase the folding efficiency of single-domain proteins, which are more common in prokaryotes than in eukaryotes,170 in cases where nonlocal interactions form at an early stage of the folding pathway. By contrast, it was suggested84,171 that the intra-ring allosteric transitions of the group II chaperonin CCT/TRiC have evolved to be sequential in order to facilitate domain-by-domain release and, thus, more efficient folding of multidomain proteins that are more common in eukaryotes. Initial support for this hypothesis came from studying the folding of single- and double-domain substrate proteins on a 2D square lattice in the presence of a chaperonin that was represented by a cavity wall with which the substrate protein can interact.171 It was found that single-domain proteins benefit more from concerted switches of the cavity walls from hydrophobic to hydrophilic, whereas double-domain proteins benefit more from sequential changes. Experimental support for this hypothesis was obtained 6599

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

conundrum was resolved by the finding obtained using fluorescence correlation spectroscopy that ∼50% of SR1 molecules are in the T state even at saturating ATP concentrations.180 The conflict with the thermodynamic models of allostery4,5 is due to the fact that the productbound states of the protein are usually neglected in these models since product release is assumed to be fast. It was shown,180 however, that the fraction of molecules in the ATPbound R state can be smaller than 1 if the rate of product (i.e., ADP) release is slow relative to the rate of hydrolysis.93,134 Consequently, it is not surprising that ADP release is the ratelimiting step in the reaction cycles of other machines such as myosin VIII.181 In the case of CCT/TRiC, its ability to function despite having a relatively fast rate of ADP release is consistent with the success in crystallizing it in complex with ATP and protein substrate (actin) together.37

by taking advantage of the Asp155 → Ala GroEL mutant that undergoes sequential T → R transitions.146 The folding yields of the CyPet and YPet domains in a CyPet-YPet chimera and that of the entire chimera were determined, in the presence of wild-type GroEL or the Asp155 → Ala mutant, by measuring the intrinsic fluorescence of the domains and the FRET between the two domains. The results showed that the domains are released in a concerted fashion in the presence of wild-type GroEL but not in the presence of the Asp155 → Ala mutant.172 The hypothesis was also tested by monitoring the folding of two GroES-independent substrates, mouse DHFR and GFP, fused to a GroES-dependent substrate, rhodanese, in the presence of wild-type GroEL or the Asp155 → Ala mutant, at different concentrations of ATP. The dependence of the folding yields of mouse DHFR and GFP on ATP concentration was found to be sigmoidal in the presence of wild-type GroEL and biphasic in the presence of the Asp155 → Ala mutant,173 thus mirroring their respective concerted and sequential transitions. Taken together, these studies showed that sequential intra-ring allosteric transitions can lead to sequential release and folding of substrate domains. In the case of CCT/TRiC, a test of this mechanism remains to be done. Substrate proteins are usually attached to multiple subunits of GroEL in the T state174 and can therefore undergo stretching when the substrate binding sites move apart during the T → R and R → R′ allosteric transitions. Such stretching can lead to unfolding of misfolded substrates, thereby giving them further opportunity to fold correctly as described by the iterative annealing model.65 Evidence for such a mechanism was reported by Rye and co-workers, who showed that ATP binding to a Rubisco-bound trans ring of the GroEL−GroES complex results in a drop in FRET that reflects substrate expansion and unfolding.66 Such ATP-promoted stretching has also been observed for the SR1-bound Val8 → Gly; Tyr283 → Asp double mutant of MBP.175 The load that is overcome during the T → R transition which drives substrate stretching was estimated using the TMR labeling strategy devised by Corsepius and Lorimer.137 They showed that each intersubunit tether (owing to TMR stacking) in the T state of GroEL stabilizes it relative to the R state by 2.6 ± 1.0 kJ/mol. Given that each ring can contain up to three tethers and that all the tethers are broken during the T → R transition (as expected for a concerted transition97), it follows that a single ring of the GroEL nanomachine can do work and overcome a load of 7.8 ± 1.7 kJ/mol upon its transition from the T to the R state. This value of 7.8 ± 1.7 kJ/mol is equivalent to the free energy of breaking only two typical solvent-exposed noncovalent interactions in proteins,176 but the maximal load that GroEL can overcome may actually be greater. It was calculated177 from ΔG = FΔX that this amount of work is sufficient to stretch a protein by ΔX = 0.2 nm if its interaction force, F, with GroEL is 70 pN as found for denatured β-lactamase.178 Such ATPinduced stretching may also occur in the case of CCT/TRiCbound substrates.179 The above-mentioned functions attributed to intra-ring allostery, e.g., switching between different functional states and forced unfolding, all require cycling between the T and the R conformational states. It follows, however, from thermodynamic models of allostery4,5 and the affinities for ATP of the T and R states93 that, in the presence of high concentrations of ATP (such as those found in the cell), the equilibrium of GroEL should be fully shifted toward the R state, thereby preventing substrate binding which is to the T state. This

4.2. Inter-Ring Allostery in Chaperonin Function

Inter-ring signaling is crucial for the function of GroEL since the discharge of GroES and an encapsulated substrate from the cis ring requires ATP binding to the trans ring.141 The complex of SR1 with GroES is, therefore, relatively long lived76 since inter-ring signaling is absent. It was also shown that (i) ATP hydrolysis in the cis ring precedes ATP binding to the trans ring133,141 and (ii) ATP hydrolysis in the cis ring is the ratelimiting step of the GroE reaction cycle in the presence of an encapsulated substrate.182 These results, which are in accord with the negative inter-ring allostery of GroEL, with respect to ATP, indicated that the time a substrate spends being encapsulated in the GroE cavity is controlled by the distribution of lifetimes of the ATP-bound cis ring. One may expect that this distribution of lifetimes of about 10 s was tuned during evolution to be compatible with the folding requirements of GroEL’s obligatory substrates. Given, however, that the GroE machine is required for the folding of many different essential proteins,51 it is unlikely that its properties (e.g., the lifetime of the ATP-bound cis ring) have evolved to be optimal for any one particular substrate.183 Accordingly, it was found that introducing the Arg13 → Gly; Ala126 → Val double mutation, which disrupts inter-ring allostery in GroEL, impairs GroELassisted folding in vivo in a substrate-selective manner.184 These results also highlighted the importance of inter-ring allostery in vivo. Given the crucial role of inter-ring signaling for the function of GroEL, it seemed reasonable to assume that mitochondrial hsp60, which purifies as a single ring, is able to mediate protein folding because it forms a transient double-ring intermediate.185 In support of this proposal, an equilbrium between single- and double-ring forms was found in the case of some single-ring GroEL variants,102 and human mitochondrial hsp60 was crystallized recently in the football-shaped form.121 It was shown, however, that mutations which block assembly of mitochondrial hsp60 into double rings do not abolish its folding activity.186 It, therefore, appears that a drop in affinity of mitochondrial hsp60 for its hsp10 cochaperonin upon ATP hydrolysis is sufficient to trigger their dissociation and the release of encapsulated substrate without a need in the case of this chaperonin system for a signal from the opposite ring.186 By contrast with the model for the reaction cycle suggested by these previous studies,133,182 it was shown that both rings of GroEL can become occupied simultaneously with ATP and GroES, thereby forming symmetric GroEL−GroES2 “football” particles.187 These symmetric particles were found to be the 6600

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

dominant form in the presence of substrate protein,188,189 and they convert into the asymmetric GroEL−GroES “bullet” complexes when the population of the nonfolded substrate protein declines during folding.187 The formation of the symmetric species was found to occur under conditions that are more physiological (i.e., 37 °C and in the presence of nonfolded substrates and 0.1−0.2 M K+ ions) than those used before by others to study the asymmetric species. It remains to be shown, however, that the symmetric particle is the dominant form also when both ADP and ATP are present at a concentration ratio that is physiological115 (estimates of the ATP/ADP concentration ratio in E. coli range from 3 to 10190,191). The studies showing that the symmetric paricles are the functional species in the GroE reaction cycle187,189 were also challenged115 on other grounds that include their employment of a substrate (reduced and Ca2+-depleted αlactalbumin) that is unable to fold. Several questions arise if we assume that the symmetric species is indeed the dominant functional form in GroEL’s reaction cycle. One question concerns the reason(s) for conservation of inter-ring negative allostery if the asymmetric species is not the main functional form. Although it was reported that inter-ring communication is dispensable in the case of a group II chaperonin192 (in agreement with the apparent absence of a communication route that corresponds to the helix D-mediated pathway in GroEL), the conservation of a feature during evolution usually indicates that it does have an important functional role. A second question concerns the implications of the symmetric species being the dominant functional form for the distribution of encapsulation times. Lorimer and co-workers proposed that asymmetry between rings in the extent of hydrolysis, which develops after formation of the fully symmetric particle, determines which GroES molecule will depart first.187 The process is, however, assumed to be stochastic, so that the first GroES molecule to bind can also be the first to depart. Consequently, a broad distribution of encapsulation times with an average of ∼1 s is expected instead of the more narrow distribution with an average encapsulation time of ∼10 s when the asymmetric species is the dominant functional form. These, on average, 10-fold shorter encapsulation times are compatible with forced unfolding and iterative annealing mechanisms of action65 but may not be sufficient for partial folding to occur especially when considering that many of the obligate substrates are relatively slow folders. More generally, a stochastic mechanism of GroES departure may seem unlikely given the deterministic nature of machine function.

structure that takes place in a domino-like fashion, whereas that of GroEL is more likely to be due to a redistribution of its conformational ensemble. The differences in the allosteric mechanisms of GroEL and CCT/TRiC stem, in part, from the respective staggered vs in-register back-to-back stacking of the rings and presence vs absence of intra-ring apical domain interactions in these chaperonins. The implications of the different allosteric mechanisms of GroEL and CCT/TRiC for their function require further study. It is unlikely, however, that the allosteric properties of chaperonins are dispensable, although some functional chaperonin forms lack allostery and specific conditions can be found where allostery may not be required.

AUTHOR INFORMATION Corresponding Author

*Phone: ++972 8 9343399. Fax: ++972 8 9343029. E-mail: [email protected]. Notes

The authors declare no competing financial interest. Biographies Ranit Gruber completed her undergraduate studies in Chemistry at the Ben-Gurion University of the Negev and her graduate studies in Structural Biology at the Weizmann Institute of Science in Rehovot. Her thesis work on “Elucidating allosteric pathways in chaperonins” was carried out under the supervision of Prof. Amnon Horovitz. She is currently a postdoctoral fellow in the laboratory of Prof. A. Horovitz. Amnon Horovitz completed his undergraduate and graduate studies in Biochemistry at the Hebrew University of Jerusalem. His thesis work on “Additivity in the effects of amino acid substitutions on protein− protein interactions” was carried out under the supervision of Profs. M. Rigbi and R. D. Levine. In 1989, he joined the laboratory of Prof. Alan Fersht in Cambridge, England, as a postdoctoral fellow, where he worked on developing the double-mutant cycle method and applying it to study protein folding and stability. In 1991, he joined the faculty of the Department of Structural Biology at the Weizmann Institute of Science in Israel where he has been since. He was chair of the Department of Structural Biology from 2000 to 2006 and has been Full Professor since 2004. He has won several awards including the Hestrin Prize of the Israel Biochemical Society (1989) and the Zimmer Award of the University of Cincinnati (2008). He is currently President of the Israel Society for Biochemistry and Molecular Biology.

ACKNOWLEDGMENTS This work was supported by grant 158/12 of the Israel Science Foundation and by the Minerva Foundation with funding from the Federal German Ministry for Education and Research. A.H. is an incumbent of the Carl and Dorothy Bennett Professorial Chair in Biochemistry.

5. CONCLUDING REMARKS Given that important advances are still being made in the study of allostery in hemoglobin193 more than 100 years after the work of Hill,3 it is remarkable how much progress has been made in understanding the allosteric mechanism of GroEL in less than three decades of research. By contrast, lack of structure−function correlations and high-resolution structural data have impeded similar progress in elucidating the allosteric mechanism of the much more complex CCT/TRiC machine. It is clear, however, that the allosteric mechanisms of GroEL and CCT/TRiC differ despite the fact that both display positive intra- and negative inter-ring cooperativities with respect to ATP and that they have similar subunit architectures. Intra-ring allostery in CCT/TRiC seems to be due to an ATP bindingtriggered propagation of a conformational change through its

REFERENCES (1) Monod, J.; Jacob, F. Teleonomic Mechanisms in Cellular Metabolism, Growth, and Differentiation. Cold Spring Harbor Symp. Quant. Biol. 1961, 26, 389−401. (2) Changeux, J. P. The Feedback Control Mechanism of Biosynthetic L-Threonine Deaminase by L-Isoleucine. Cold Spring Harbor Symp. Quant. Biol. 1961, 26, 313−318. (3) Hill, A. V. The Possible Effects of the Aggregation of the Molecules of Haemoglobin on Its Dissociation Curves. J. Physiol. 1910, 40, IV−VII. 6601

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

(4) Monod, J.; Wyman, J.; Changeux, J. P. On the Nature of Allosteric Transitions: A Plausible Model. J. Mol. Biol. 1965, 12, 88− 118. (5) Koshland, D. E., Jr.; Némethy, G.; Filmer, D. Comparison of Experimental Binding Data and Theoretical Models in Proteins Containing Subunits. Biochemistry 1966, 5, 365−385. (6) Milo, R.; Hou, J. H.; Springer, M.; Brenner, M. P.; Kirschner, M. W. The Relationship Between Evolutionary and Physiological Variation in Hemoglobin. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 16998−17003. (7) Changeux, J. P. Allostery and the Monod-Wyman-Changeux Model After 50 Years. Annu. Rev. Biophys. 2012, 41, 103−133. (8) Boehr, D. D.; Nussinov, R.; Wright, P. E. The Role of Dynamic Conformational Ensembles in Biomolecular Recognition. Nat. Chem. Biol. 2009, 5, 789−796. (9) Sharon, M.; Horovitz, A. Probing Allosteric Mechanisms Using Native Mass Spectrometry. Curr. Opin. Struct. Biol. 2015, 34, 7−16. (10) Eigen, M. Kinetics of Reaction Control and Information Transfer in Enzymes and Nucleic Acids. Nobel Symp. 1967, 5, 333− 369. (11) Cooper, A.; Dryden, D. T. Allostery Without Conformational Change. A Plausible Model. Eur. Biophys. J. 1984, 11, 103−109. (12) Townsend, P. D.; Rodgers, T. L.; Pohl, E.; Wilson, M. R.; McLeish, T. C.; Cann, M. J. Global Low-Frequency Motions in Protein Allostery: CAP as a Model System. Biophys. Rev. 2015, 7, 175− 182. (13) Hodson, S.; Marshall, J. J.; Burston, S. G. Mapping the Road to Recovery: The ClpB/Hsp104 Molecular Chaperone. J. Struct. Biol. 2012, 179, 161−171. (14) Kelman, Z.; Finkelstein, J.; O’Donnell, M. Protein Structure: Why Have Six-Fold Symmetry? Curr. Biol. 1995, 5, 1239−1242. (15) Horwich, A. L.; Weber-Ban, E. U.; Finley, D. Chaperone Rings in Protein Folding and Degradation. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 11033−11040. (16) Thirumalai, D.; Klimov, D. K.; Lorimer, G. H. Caging Helps Proteins Fold. Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 11195−11197. (17) Horwich, A. L.; Fenton, W. A. Chaperonin-Mediated Protein Folding: Using a Central Cavity to Kinetically Assist Polypeptide Chain Folding. Q. Rev. Biophys. 2009, 42, 83−116. (18) Hayer-Hartl, M.; Bracher, A.; Hartl, F. U. The GroEL-GroES Chaperonin Machine: A Nano-Cage for Protein Folding. Trends Biochem. Sci. 2016, 41, 62−76. (19) Sauer, R. T.; Baker, T. A. AAA+ Proteases: ATP-Fueled Machines of Protein Destruction. Annu. Rev. Biochem. 2011, 80, 587− 612. (20) Riedel, C.; Gabizon, R.; Wilson, C. A.; Hamadani, K.; Tsekouras, K.; Marqusee, S.; Pressé, S.; Bustamante, C. The Heat Released During Catalytic Turnover Enhances the Diffusion of an Enzyme. Nature 2015, 517, 227−230. (21) Chistol, G.; Liu, S.; Hetherington, C. L.; Moffitt, J. R.; Grimes, S.; Jardine, P. J.; Bustamante, C. High Degree of Coordination and Division of Labor Among Subunits in a Homomeric Ring ATPase. Cell 2012, 151, 1017−1028. (22) Gai, D.; Zhao, R.; Li, D.; Finkielstein, C. V.; Chen, X. S. Mechanisms of Conformational Change for a Replicative Hexameric Helicase of SV40 Large Tumor Antigen. Cell 2004, 119, 47−60. (23) Stinson, B. M.; Nager, A. R.; Glynn, S. E.; Schmitz, K. R.; Baker, T. A.; Sauer, R. T. Nucleotide Binding and Conformational Switching in the Hexamaric Ring of a AAA+ Machine. Cell 2013, 153, 628−639. (24) Horovitz, A.; Fridmann, Y.; Kafri, G.; Yifrach, O. Review: Allostery in Chaperonins. J. Struct. Biol. 2001, 135, 104−114. (25) Horovitz, A.; Willison, K. R. Allosteric Regulation of Chaperonins. Curr. Opin. Struct. Biol. 2005, 15, 646−651. (26) Saibil, H. R.; Fenton, W. A.; Clare, D. K.; Horwich, A. L. Structure and Allostery of the Chaperonin GroEL. J. Mol. Biol. 2013, 425, 1476−1487. (27) Skjærven, L.; Cuellar, J.; Martinez, A.; Valpuesta, J. M. Dynamics, Flexibility, and Allostery in Molecular Chaperonins. FEBS Lett. 2015, 589, 2522−2532.

(28) Thirumalai, D.; Lorimer, G. H. Chaperonin-Mediated Protein Folding. Annu. Rev. Biophys. Biomol. Struct. 2001, 30, 245−269. (29) Lin, Z.; Rye, H. S. GroEL-Mediated Protein Folding: Making the Impossible, Possible. Crit. Rev. Biochem. Mol. Biol. 2006, 41, 211− 239. (30) Gutsche, I.; Essen, L. O.; Baumeister, W. Group II Chaperonins: New TRiC(k)s and Turns of a Protein Folding Machine. J. Mol. Biol. 1999, 293, 295−312. (31) Horwich, A. L.; Fenton, W. A.; Chapman, E.; Farr, G. W. Two Families of Chaperonin: Physiology and Mechanism. Annu. Rev. Cell Dev. Biol. 2007, 23, 115−145. (32) Bigotti, M. G.; Clarke, A. R. Chaperonins: The Hunt for the Group II Mechanism. Arch. Biochem. Biophys. 2008, 474, 331−339. (33) Yébenes, H.; Mesa, P.; Muñoz, I. G.; Montoya, G.; Valpuesta, J. M. Chaperonins: Two Rings for Folding. Trends Biochem. Sci. 2011, 36, 424−432. (34) Ditzel, L.; Löwe, J.; Stock, D.; Stetter, K. O.; Huber, H.; Huber, R.; Steinbacher, S. Crystal Structure of the Thermosome, the Archaeal Chaperonin and Homolog of CCT. Cell 1998, 93, 125−138. (35) Kalisman, N.; Adams, C. M.; Levitt, M. Subunit Order of Eukaryotic TRiC/CCT Chaperonin by Cross-linking, Mass Spectrometry, and Combinatorial Homology Modeling. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 2884−2889. (36) Leitner, A.; Joachimiak, L. A.; Bracher, A.; Mönkemeyer, L.; Walzthoeni, T.; Chen, B.; Pechmann, S.; Holmes, S.; Cong, Y.; Ma, B.; et al. The Molecular Architecture of the Eukaryotic Chaperonin TRiC/CCT. Structure 2012, 20, 814−825. (37) Dekker, C.; Roe, S. M.; McCormack, E. A.; Beuron, F.; Pearl, L. H.; Willison, K. R. The Crystal Structure of Yeast CCT Reveals Intrinsic Asymmetry of Eukaryotic Cytosolic Chaperonins. EMBO J. 2011, 30, 3078−3090. (38) Kalisman, N.; Schröder, G. F.; Levitt, M. The Crystal Structures of the Eukaryotic Chaperonin CCT Reveal Its Functional Partitioning. Structure 2013, 21, 540−549. (39) Braig, K.; Otwinowski, Z.; Hegde, R.; Boisvert, D. C.; Joachimiak, A.; Horwich, A. L.; Sigler, P. B. The Crystal Structure of the Bacterial Chaperonin GroEL at 2.8 Å. Nature 1994, 371, 578−586. (40) Bartolucci, C.; Lamba, D.; Grazulis, S.; Manakova, E.; Heumann, H. Crystal Structure of Wild-Type Chaperonin GroEL. J. Mol. Biol. 2005, 354, 940−951. (41) Fenton, W. A.; Kashi, Y.; Furtak, K.; Horwich, A. L. Residues in Chaperonin GroEL Required for Polypeptide Binding and Release. Nature 1994, 371, 614−619. (42) Hunt, J. F.; Weaver, A. J.; Landry, S. J.; Gierasch, L.; Deisenhofer, J. The Crystal Structure of the GroES Co-Chaperonin at 2.8 Å Resolution. Nature 1996, 379, 37−45. (43) Mande, S. C.; Mehra, V.; Bloom, B. R.; Hol, W. G. Structure of the Heat Shock Protein Chaperonin-10 of Mycobacterium leprae. Science 1996, 271, 203−207. (44) Chandrasekhar, G. N.; Tilly, K.; Woolford, C.; Hendrix, R.; Georgopoulos, C. Purification and Properties of the GroES Morphogenetic Protein of Escherichia coli. J. Biol. Chem. 1986, 261, 12414−12419. (45) Xu, Z.; Horwich, A. L.; Sigler, P. B. The Crystal Structure of the Asymmetric GroEL-GroES-(ADP)7 Chaperonin Complex. Nature 1997, 388, 741−750. (46) Klumpp, M.; Baumeister, W.; Essen, L. O. Structure of the Substrate Binding Domain of the Thermosome, an Archaeal Group II Chaperonin. Cell 1997, 91, 263−270. (47) Iizuka, R.; So, S.; Inobe, T.; Yoshida, T.; Zako, T.; Kuwajima, K.; Yohda, M. Role of the Helical Protrusion in the Conformational Change and Molecular Chaperone Activity of the Archaeal Group II Chaperonin. J. Biol. Chem. 2004, 279, 18834−18839. (48) Viitanen, P. V.; Gatenby, A. A.; Lorimer, G. H. Purified Chaperonin 60 (groEL) Interacts with the Nonnative States of a Multitude of Escherichia coli Proteins. Protein Sci. 1992, 1, 363−369. (49) Aoki, K.; Motojima, F.; Taguchi, H.; Yomo, T.; Yoshida, M. GroEL Binds Artificial Proteins with Random Sequences. J. Biol. Chem. 2000, 275, 13755−13758. 6602

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

(50) Lorimer, G. H. A Quantitative Assessment of the Role of the Chaperonin Proteins in Protein Folding in Vivo. FASEB J. 1996, 10, 5−9. (51) Kerner, M. J.; Naylor, D. J.; Ishihama, Y.; Maier, T.; Chang, H. C.; Stines, A. P.; Georgopoulos, C.; Frishman, D.; Hayer-Hartl, M.; Mann, M.; et al. Proteome-Wide Analysis of Chaperonin-Dependent Protein Folding in Escherichia coli. Cell 2005, 122, 209−220. (52) Fujiwara, K.; Ishihama, Y.; Nakahigashi, K.; Soga, T.; Taguchi, H. A Systematic Survey of in Vivo Obligate Chaperonin-Dependent Substrates. EMBO J. 2010, 29, 1552−1564. (53) Azia, A.; Unger, R.; Horovitz, A. What Distinguishes GroEL Substrates from Other Escherichia coli Proteins? FEBS J. 2012, 279, 543−550. (54) Gao, Y.; Thomas, J. O.; Chow, R. L.; Lee, G. H.; Cowan, N. J. A Cytoplasmic Chaperonin that Catalyzes β-Actin Folding. Cell 1992, 69, 1043−1050. (55) Yaffe, M. B.; Farr, G. W.; Miklos, D.; Horwich, A. L.; Sternlicht, M. L.; Sternlicht, H. TCP1 Complex is a Molecular Chaperone in Tubulin Biogenesis. Nature 1992, 358, 245−248. (56) Dekker, C.; Stirling, P. C.; McCormack, E. A.; Filmore, H.; Paul, A.; Brost, R. L.; Costanzo, M.; Boone, C.; Leroux, M. R.; Willison, K. R. The Interaction Network of the Chaperonin CCT. EMBO J. 2008, 27, 1827−1839. (57) Yam, A. Y.; Xia, Y.; Lin, H. T.; Burlingame, A.; Gerstein, M.; Frydman, J. Defining the TRiC/CCT Interactome Links Chaperonin Function to Stabilization of Newly Made Proteins with Complex Topologies. Nat. Struct. Mol. Biol. 2008, 15, 1255−1262. (58) Horwich, A. L.; Apetri, A. C.; Fenton, W. A. The GroEL/GroES cis Cavity as a Passive Anti-Aggregation Device. FEBS Lett. 2009, 583, 2654−2662. (59) Tyagi, N. K.; Fenton, W. A.; Deniz, A. A.; Horwich, A. L. Double Mutant MBP Refolds at Same Rate in Free Solution as Inside the GroEL/GroES Chaperonin Chamber When Aggregation in Free Solution is Prevented. FEBS Lett. 2011, 585, 1969−1672. (60) Tang, Y. C.; Chang, H. C.; Roeben, A.; Wischnewski, D.; Wischnewski, N.; Kerner, M. J.; Hartl, F. U.; Hayer-Hartl, M. Structural Features of the GroEL-GroES Nano-Cage Required for Rapid Folding of Encapsulated Protein. Cell 2006, 125, 903−914. (61) Georgescauld, F.; Popova, K.; Gupta, A. J.; Bracher, A.; Engen, J. R.; Hayer-Hartl, M.; Hartl, F. U. GroEL/ES Chaperonin Modulates the Mechanism and Accelerates the Rate of TIM-Barrel Domain Folding. Cell 2014, 157, 922−934. (62) Gupta, A. J.; Haldar, S.; Miličić, G.; Hartl, F. U.; Hayer-Hartl, M. Active Cage Mechanism of Chaperonin-Assisted Protein Folding Demonstrated at Single-Molecule Level. J. Mol. Biol. 2014, 426, 2739− 2754. (63) England, J. L.; Lucent, D.; Pande, V. S. A Role for Confined Water in Chaperonin Function. J. Am. Chem. Soc. 2008, 130, 11838− 11839. (64) Franck, J. M.; Sokolovski, M.; Kessler, N.; Matalon, E.; GordonGrossman, M.; Han, S. I.; Goldfarb, D.; Horovitz, A. Probing Water Density and Dynamics in the Chaperonin GroEL Cavity. J. Am. Chem. Soc. 2014, 136, 9396−9403. (65) Todd, M. J.; Lorimer, G. H.; Thirumalai, D. ChaperoninFacilitated Protein Folding: Optimization of Rate and Yield by an Iterative Annealing Mechanism. Proc. Natl. Acad. Sci. U. S. A. 1996, 93, 4030−4035. (66) Lin, Z.; Madan, D.; Rye, H. S. GroEL Stimulates Protein Folding Through Forced Unfolding. Nat. Struct. Mol. Biol. 2008, 15, 303−311. (67) Weaver, J.; Rye, H. S. The C-terminal Tails of the Bacterial Chaperonin GroEL Stimulate Protein Folding by Directly Altering the Conformation of a Substrate Protein. J. Biol. Chem. 2014, 289, 23219− 23232. (68) Priya, S.; Sharma, S. K.; Sood, V.; Mattoo, R. U. H.; Finka, A.; Azem, A.; De Los Rios, P.; Goloubinoff, P. GroEL and CCT are Catalytic Unfoldases Mediating Out-of-Cage Polypeptide Refolding Without ATP. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 7199−7204.

(69) Libich, D. S.; Tugarinov, V.; Clore, G. M. Intrinsic Unfoldase/ Foldase Activity of the Chaperonin GroEL Directly Demonstrated Using Multinuclear Relaxation-Based NMR. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 8817−8823. (70) Chatellier, J.; Hill, F.; Lund, P. A.; Fersht, A. R. In Vivo Activities of GroEL Minichaperones. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 9861−9866. (71) Henderson, B.; Fares, M. A.; Lund, P. A. Chaperonin 60: A Paradoxical, Evolutionary Conserved Protein Family with Multiple Moonlighting Functions. Biol. Rev. Camb. Philos. Soc. 2013, 88, 955− 987. (72) Gray, T. E.; Fersht, A. R. Cooperativity in ATP Hydrolysis by GroEL Is Increased by GroES. FEBS Lett. 1991, 292, 254−258. (73) Bochkareva, E. S.; Lissin, N. M.; Flynn, G. C.; Rothman, J. E.; Girshovich, A. S. Positive Cooperativity in the Functioning of Molecular Chaperone GroEL. J. Biol. Chem. 1992, 267, 6796−6800. (74) Jackson, G. S.; Staniforth, R. A.; Halsall, D. J.; Atkinson, T.; Holbrook, J. J.; Clarke, A. R.; Burston, S. G. Binding and Hydrolysis of Nucleotides in the Chaperonin Catalytic Cycle: Implications for the Mechanism of Assisted Protein Folding. Biochemistry 1993, 32, 2554− 2563. (75) Inobe, T.; Makio, T.; Takasu-Ishikawa, E.; Terada, T. P.; Kuwajima, K. Nucleotide Binding to the Chaperonin GroEL: NonCooperative Binding of ATP Analogs and ADP, and Cooperative Effect of ATP. Biochim. Biophys. Acta, Protein Struct. Mol. Enzymol. 2001, 1545, 160−173. (76) Weissman, J. S.; Hohl, C. M.; Kovalenko, O.; Kashi, Y.; Chen, S.; Braig, K.; Saibil, H. R.; Fenton, W. A.; Horwich, A. L. Mechanism of GroEL Action: Productive Release of Polypeptide from a Sequestered Position Under GroES. Cell 1995, 83, 577−587. (77) Poso, D.; Clarke, A. R.; Burston, S. G. A Kinetic Analysis of the Nucleotide-Induced Allosteric Transitions in a Single-Ring Mutant of GroEL. J. Mol. Biol. 2004, 338, 969−977. (78) Amir, A.; Horovitz, A. Kinetic Analysis of ATP-dependent InterRing Communication in GroEL. J. Mol. Biol. 2004, 338, 979−988. (79) Kafri, G.; Willison, K. R.; Horovitz, A. Nested Allosteric Interactions in the Cytoplasmic Chaperonin Containing TCP-1. Protein Sci. 2001, 10, 445−449. (80) Kusmierczyk, A. R.; Martin, J. Nested Cooperativity and Salt Dependence of the ATPase Activity of the Archaeal Chaperonin Mmcpn. FEBS Lett. 2003, 547, 201−204. (81) Bigotti, M. G.; Clarke, A. R. Cooperativity in the Thermosome. J. Mol. Biol. 2005, 348, 13−26. (82) Yifrach, O.; Horovitz, A. Allosteric Control by ATP of NonFolded Protein Binding to GroEL. J. Mol. Biol. 1996, 255, 356−361. (83) Reissmann, S.; Joachimiak, L. A.; Chen, B.; Meyer, A. S.; Nguyen, A.; Frydman, J. A Gradient of ATP Affinities Generates an Asymmetric Power Stroke Driving the Chaperonin TRiC/CCT Folding Cycle. Cell Rep. 2012, 2, 866−877. (84) Rivenzon-Segal, D.; Wolf, S. G.; Shimon, L.; Willison, K. R.; Horovitz, A. Sequential ATP-Induced Allosteric Transitions of the Cytoplasmic Chaperonin Containing TCP-1 Revealed by EM Analysis. Nat. Struct. Mol. Biol. 2005, 12, 233−237. (85) Muñoz, I. G.; Yébenes, H.; Zhou, M.; Mesa, P.; Serna, M.; Park, A. Y.; Bragado-Nilsson, E.; Beloso, A.; de Cárcer, G.; Malumbres, M.; et al. Crystal Structure of the Open Conformation of the Mammalian Chaperonin CCT in Complex with Tubulin. Nat. Struct. Mol. Biol. 2011, 18, 14−19. (86) Kafri, G.; Horovitz, A. Transient Kinetic Analysis of ATPInduced Allosteric Transitions in the Eukaryotic Chaperonin Containing TCP-1. J. Mol. Biol. 2003, 326, 981−987. (87) Inobe, T.; Kikushima, K.; Makio, T.; Arai, M.; Kuwajima, K. The Allosteric Transition of GroEL Induced by Metal Fluoride-ADP Complexes. J. Mol. Biol. 2003, 329, 121−134. (88) Melki, R.; Cowan, N. J. Facilitated Folding of Actins and Tubulins Occurs via a Nucleotide-Dependent Interaction Between Cytoplasmic Chaperonin and Distinctive Folding Intermediates. Mol. Cell. Biol. 1994, 14, 2895−2904. 6603

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

by cis- and trans-Only GroEL-GroES Complexes. EMBO J. 2003, 22, 3220−3230. (109) Fridmann, Y.; Kafri, G.; Danziger, O.; Horovitz, A. Dissociation of the GroEL-GroES Asymmetric Complex Is Accelerated by Increased Cooperativity in ATP Binding to the GroEL Ring Distal to GroES. Biochemistry 2002, 41, 5938−5944. (110) Fei, X.; Yang, D.; LaRonde-LeBlanc, N.; Lorimer, G. H. Crystal Structure of a GroEL-ADP Complex in the Relaxed Allosteric State at 2.7 Å Resolution. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, E2958− E2966. (111) Bochkareva, E. S.; Girshovich, A. S. ATP Induces Non-Identity of Two Rings in Chaperonin GroEL. J. Biol. Chem. 1994, 269, 23869− 23871. (112) Saibil, H.; Dong, Z.; Wood, S.; auf der Mauer, A. Binding of Chaperonins. Nature 1991, 353, 25−26. (113) Langer, T.; Pfeifer, G.; Martin, J.; Baumeister, W.; Hartl, F. U. Chaperonin-Mediated Protein Folding: GroES Binds to One End of the GroEL Cylinder, Which Accommodates the Protein Substrate Within Its Central Cavity. EMBO J. 1992, 11, 4757−4765. (114) Ishii, N.; Taguchi, H.; Sumi, M.; Yoshida, M. Structure of Holo-Chaperonin Studied with Electron Microscopy. Oligomeric Cpn10 on Top of Two Layers of Cpn60 Rings with Two Stripes Each. FEBS Lett. 1992, 299, 169−174. (115) Haldar, S.; Gupta, A. J.; Yan, X.; Miličić, G.; Hartl, F. U.; Hayer-Hartl, M. Chaperonin-Assisted Protein Folding: Relative Population of Asymmetric and Symmetric GroEL:GroES Complexes. J. Mol. Biol. 2015, 427, 2244−2255. (116) Schmidt, M.; Rutkat, K.; Rachel, R.; Pfeifer, G.; Jaenicke, R.; Viitanen, P.; Lorimer, G.; Buchner, J. Symmetric Complexes of GroE Chaperonins as Part of the Functional Cycle. Science 1994, 265, 656− 659. (117) Azem, A.; Kessel, M.; Goloubinoff, P. Characterization of a Functional GroEL14(GroES7)2 Chaperonin Hetero-Oligomer. Science 1994, 265, 653−656. (118) Llorca, O.; Marco, S.; Carrascosa, J. L.; Valpuesta, J. M. The Formation of Symmetrical GroEL-GroES Complexes in the Presence of ATP. FEBS Lett. 1994, 345, 181−186. (119) Koike-Takeshita, A.; Arakawa, T.; Taguchi, H.; Shimamura, T. Crystal Structure of a Symmetric Football-Shaped GroEL:GroES2ATP14 Complex Determined at 3.8 Å Reveals Rearrangement Between Two GroEL Rings. J. Mol. Biol. 2014, 426, 3634−3641. (120) Fei, X.; Ye, X.; LaRonde, N. A.; Lorimer, G. H. Formation and Structures of GroEL:GroES2 Chaperonin Footballs, the ProteinFolding Functional Form. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 12775−12780. (121) Nisemblat, S.; Yaniv, O.; Parnas, A.; Frolow, F.; Azem, A. Crystal Structure of the Human Mitochondrial Chaperonin Symmetrical Football Complex. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 6044−6049. (122) Ma, J.; Sigler, P. B.; Xu, Z.; Karplus, M. A Dynamic Model for the Allosteric Mechanism of GroEL. J. Mol. Biol. 2000, 302, 303−313. (123) Ma, J.; Karplus, M. The Allosteric Mechanism of the Chaperonin GroEL: A Dynamic Analysis. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 8502−8507. (124) Shiseki, K.; Murai, N.; Motojima, F.; Hisabori, T.; Yoshida, M.; Taguchi, H. Synchronized Domain-Opening Motion of GroEL Is Essential for Communication Between the Two Rings. J. Biol. Chem. 2001, 276, 11335−11338. (125) Yifrach, O.; Horovitz, A. Mapping the Transition State of the Allosteric Pathway of GroEL by Protein Engineering. J. Am. Chem. Soc. 1998, 120, 13262−13263. (126) Dyachenko, A.; Gruber, R.; Shimon, L.; Horovitz, A.; Sharon, M. Allosteric Mechanisms Can be Distinguished Using Structural Mass Spectrometry. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 7235−7239. (127) Shimon, L.; Hynes, G. M.; McCormack, E. A.; Willison, K. R.; Horovitz, A. ATP-Induced Allostery in the Eukaryotic Chaperonin CCT Is Abolished by the Mutation G345D in CCT4 that Renders Yeast Temperature-Sensitive for Growth. J. Mol. Biol. 2008, 377, 469− 477.

(89) Chaudhry, C.; Farr, G. W.; Todd, M. J.; Rye, H. S.; Brunger, A. T.; Adams, P. D.; Horwich, A. L.; Sigler, P. B. Role of the γ-Phosphate of ATP in Triggering Protein Folding by GroEL-GroES: Function, Structure and Energetics. EMBO J. 2003, 22, 4877−4887. (90) Rubin, M. M.; Changeux, J. P. On the Nature of Allosteric Transitions: Implications of Non-Exclusive Ligand Binding. J. Mol. Biol. 1966, 21, 265−274. (91) Viitanen, P. V.; Lubben, T. H.; Reed, J.; Goloubinoff, P.; O’Keefe, D. P.; Lorimer, G. H. Chaperonin-Facilitated Refolding of Ribulose Bisphosphate Carboxylase and ATP Hydrolysis by Chaperonin 60 (groEL) are Potassium Dependent. Biochemistry 1990, 29, 5665−5671. (92) Todd, M. J.; Viitanen, P. V.; Lorimer, G. H. Hydrolysis of Adenosine 5′-Triphosphate by Escherichia coli GroEL: Effects of GroES and Potassium Ion. Biochemistry 1993, 32, 8560−8567. (93) Grason, J. P.; Gresham, J. S.; Widjaja, L.; Wehri, S. C.; Lorimer, G. H. Setting the Chaperonin Timer: The Effects of K+ and Substrate Protein on ATP Hydrolysis. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 17334−17338. (94) Wang, J.; Boisvert, D. C. Structural Basis for GroEL-Assisted Protein Folding from the Crystal Structure of (GroEL-KMgATP)14 at 2.0 Å Resolution. J. Mol. Biol. 2003, 327, 843−855. (95) Kiser, P. D.; Lorimer, G. H.; Palczewski, K. Use of Thallium to Identify Monovalent Cation Binding Sites in GroEL. Acta Crystallogr., Sect. F: Struct. Biol. Cryst. Commun. 2009, 65, 967−971. (96) Yifrach, O.; Horovitz, A. Two Lines of Allosteric Communication in the Oligomeric Chaperonin GroEL are Revealed by the Single Mutation Arg196→Ala. J. Mol. Biol. 1994, 243, 397−401. (97) Yifrach, O.; Horovitz, A. Nested Cooperativity in the ATPase Activity of the Oligomeric Chaperonin GroEL. Biochemistry 1995, 34, 5303−5308. (98) Yifrach, O.; Horovitz, A. Transient Kinetic Analysis of Adenosine 5′-Triphosphate Binding-Induced Conformational Changes in the Allosteric Chaperonin GroEL. Biochemistry 1998, 37, 7083− 7088. (99) Cliff, M. J.; Kad, N. M.; Hay, N.; Lund, P. A.; Webb, M. R.; Burston, S. G.; Clarke, A. R. A Kinetic Analysis of the NucleotideInduced Allosteric Transitions of GroEL. J. Mol. Biol. 1999, 293, 667− 684. (100) Taniguchi, M.; Yoshimi, T.; Hongo, K.; Mizobata, T.; Kawata, Y. Stopped-Flow Fluorescence Analysis of the Conformational Changes in the GroEL Apical Domain: Relationships Between Movements in the Apical Domain and the Quaternary Structure of GroEL. J. Biol. Chem. 2004, 279, 16368−16376. (101) LiCata, V. J.; Allewell, N. M. Is Substrate Inhibition a Consequence of Allostery in Aspartate Transcarbamylase? Biophys. Chem. 1997, 64, 225−234. (102) Erbse, A.; Yifrach, O.; Jones, S.; Lund, P. A. Chaperone Activity of a Chimeric GroEL Protein That Can Exist in a Single or Double Ring Form. J. Biol. Chem. 1999, 274, 20351−20357. (103) Wyman, J. Allosteric Linkage. J. Am. Chem. Soc. 1967, 89, 2202−2218. (104) White, H. E.; Chen, S.; Roseman, A. M.; Yifrach, O.; Horovitz, A.; Saibil, H. R. Structural Basis of Allosteric Changes in the GroEL Mutant Arg197→Ala. Nat. Struct. Biol. 1997, 4, 690−694. (105) Schoehn, G.; Quaite-Randall, E.; Jiménez, J. L.; Joachimiak, A.; Saibil, H. R. Three Conformations of an Archaeal Chaperonin, TF55 from Sulfolobus shibatae. J. Mol. Biol. 2000, 296, 813−819. (106) Llorca, O.; Smyth, M. G.; Carrascosa, J. L.; Willison, K. R.; Radermacher, M.; Steinbacher, S.; Valpuesta, J. M. 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. (107) Inbar, E.; Horovitz, A. GroES Promotes the T to R Transition of the GroEL Ring Distal to GroES in the GroEL-GroES Complex. Biochemistry 1997, 36, 12276−12281. (108) Farr, G. W.; Fenton, W. A.; Chaudhuri, T. K.; Clare, D. K.; Saibil, H. R.; Horwich, A. L. Folding with and Without Encapsulation 6604

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

(128) Lin, P.; Sherman, F. The Unique Hetero-Oligomeric Nature of the Subunits in the Catalytic Cooperativity of the Yeast Cct Chaperonin Complex. Proc. Natl. Acad. Sci. U. S. A. 1997, 94, 10780−10785. (129) Amit, M.; Weisberg, S. J.; Nadler-Holly, M.; McCormack, E. A.; Feldmesser, E.; Kaganovich, D.; Willison, K. R.; Horovitz, A. Equivalent Mutations in the Eight Subunits of the Chaperonin CCT Produce Dramatically Different Cellular and Gene Expression Phenotypes. J. Mol. Biol. 2010, 401, 532−543. (130) Kanzaki, T.; Iizuka, R.; Takahashi, K.; Maki, K.; Masuda, R.; Sahlan, M.; Yébenes, H.; Valpuesta, J. M.; Oka, T.; Furutani, M.; et al. Sequential Action of ATP-Dependent Subunit Conformational Change and Interaction Between Helical Protrusions in the Closure of the Built-in-Lid of Group II Chaperonins. J. Biol. Chem. 2008, 283, 34773−34784. (131) Douglas, N. R.; Reissmann, S.; Zhang, J.; Chen, B.; Jakana, J.; Kumar, R.; Chiu, W.; Frydman, J. Dual Action of ATP Hydrolysis Couples Lid Closure to Substrate Release into the Group II Chaperonin Chamber. Cell 2011, 144, 240−252. (132) Burston, S. G.; Ranson, N. A.; Clarke, A. R. The Origins and Consequences of Asymmetry in the Chaperonin Reaction Cycle. J. Mol. Biol. 1995, 249, 138−152. (133) Kad, N. M.; Ranson, N. A.; Cliff, M. J.; Clarke, A. R. Asymmetry, Commitment and Inhibition in the GroE ATPase Cycle Impose Alternating Functions on the Two GroEL Rings. J. Mol. Biol. 1998, 278, 267−278. (134) Danziger, O.; Shimon, L.; Horovitz, A. Glu257 in GroEL Is a Sensor Involved in Coupling Polypeptide Substrate Binding to Stimulation of ATP Hydrolysis. Protein Sci. 2006, 15, 1270−1276. (135) Martin, J.; Langer, T.; Boteva, R.; Schramel, A.; Horwich, A. L.; Hartl, F. U. Chaperonin-Mediated Protein Folding at the Surface of GroEL Through a ‘Molten Globule’-Like Intermediate. Nature 1991, 352, 36−42. (136) Staniforth, R. A.; Burston, S. G.; Atkinson, T.; Clarke, A. R. Affinity of Chaperonin-60 for a Protein Substrate and Its Modulation by Nucleotides and Chaperonin-10. Biochem. J. 1994, 300, 651−658. (137) Corsepius, N. C.; Lorimer, G. H. Measuring How Much Work the Chaperone GroEL Can Do. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, E2451−E2459. (138) Fersht, A. R.; Matouschek, A.; Serrano, L. The Folding of an Enzyme: I. Theory of Protein Engineering Analysis of Stability and Pathway of Protein Folding. J. Mol. Biol. 1992, 224, 771−782. (139) Horovitz, A.; Amir, A.; Danziger, O.; Kafri, G. Φ Value Analysis of Heterogeneity in Pathways of Allosteric Transitions: Evidence for Parallel Pathways of ATP-Induced Conformational Changes in a GroEL Ring. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 14095−14097. (140) Inobe, T.; Kuwajima, K. Φ Value Analysis of an Allosteric Transition of GroEL Based on a Single-Pathway Model. J. Mol. Biol. 2004, 339, 199−205. (141) Rye, H. S.; Burston, S. G.; Fenton, W. A.; Beechem, J. M.; Xu, Z.; Sigler, P. B.; Horwich, A. L. Distinct Actions of cis and trans ATP Within the Double Ring of the Chaperonin GroEL. Nature 1997, 388, 792−798. (142) Clare, D. K.; Vasishtan, D.; Stagg, S.; Quispe, J.; Farr, G. W.; Topf, M.; Horwich, A. L.; Saibil, H. R. ATP-Triggered Conformational Changes Delineate Substrate-Binding and -Folding Mechanics of the GroEL Chaperonin. Cell 2012, 149, 113−123. (143) Ranson, N. A.; Farr, G. W.; Roseman, A. M.; Gowen, B.; Fenton, W. A.; Horwich, A. L.; Saibil, H. R. ATP-Bound States of GroEL Captured by Cryo-Electron Microscopy. Cell 2001, 107, 869− 879. (144) Landry, S. J.; Zeilstra-Ryalls, J.; Fayet, O.; Georgopoulos, C.; Gierasch, L. M. Characterization of a Functionally Important Mobile Domain of GroES. Nature 1993, 364, 255−258. (145) Sliozberg, Y.; Abrams, C. F. Spontaneous Conformational Changes in the E. coli GroEL Subunit from All-Atom Molecular Dynamics Simulations. Biophys. J. 2007, 93, 1906−1916.

(146) Danziger, O.; Rivenzon-Segal, D.; Wolf, S. G.; Horovitz, A. Conversion of the Allosteric Transition of GroEL from Concerted to Sequential by the Single Mutation Asp-155→Ala. Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 13797−13802. (147) Yang, Z.; Májek, P.; Bahar, I. Allosteric Transitions of Supramolecular Systems Explored by Network Models: Application to Chaperonin GroEL. PLoS Comput. Biol. 2009, 5, e1000360. (148) Hyeon, C.; Lorimer, G. H.; Thirumalai, D. Dynamics of Allosteric Transitions in GroEL. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 18939−18944. (149) Tehver, R.; Chen, J.; Thirumalai, D. Allostery Wiring Diagrams in the Transitions that Drive the GroEL Reaction Cycle. J. Mol. Biol. 2009, 387, 390−406. (150) Kass, I.; Horovitz, A. Mapping Pathways of Allosteric Communication in GroEL by Analysis of Correlated Mutations. Proteins: Struct., Funct., Genet. 2002, 48, 611−617. (151) Skjaerven, L.; Grant, B.; Muga, A.; Teigen, K.; McCammon, J. A.; Reuter, N.; Martinez, A. Conformational Sampling and NucleotideDependent Transitions of the GroEL Subunit Probed by Unbiased Molecular Dynamics Simulations. PLoS Comput. Biol. 2011, 7, e1002004. (152) Klein, G.; Georgopoulos, C. Identification of Important Amino Acid Residues that Modulate Binding of Escherichia coli GroEL to Its Various Cochaperones. Genetics 2001, 158, 507−517. (153) Huo, Y.; Hu, Z.; Zhang, K.; Wang, L.; Zhai, Y.; Zhou, Q.; Lander, G.; Zhu, J.; He, Y.; Pang, X.; et al. Crystal Structure of Group II Chaperonin in the Open State. Structure 2010, 18, 1270−1279. (154) Jayasinghe, M.; Shrestha, P.; Wu, X.; Tehver, R.; Stan, G. Weak Intra-Ring Allosteric Communications of the Archaeal Chaperonin Thermosome Revealed by Normal Mode Analysis. Biophys. J. 2012, 103, 1285−1295. (155) Zheng, W.; Brooks, B. R.; Thirumalai, D. Allosteric Transitions in the Chaperonin GroEL are Captured by a Dominant Normal Mode That Is Most Robust to Sequence Variations. Biophys. J. 2007, 93, 2289−2299. (156) Sewell, B. T.; Best, R. B.; Chen, S.; Roseman, A. M.; Farr, G. W.; Horwich, A. L.; Saibil, H. R. A Mutant Chaperonin with Rearranged Inter-Ring Electrostatic Contacts and TemperatureSensitive Dissociation. Nat. Struct. Mol. Biol. 2004, 11, 1128−1133. (157) Cabo-Bilbao, A.; Spinelli, S.; Sot, B.; Agirre, J.; Mechaly, A. E.; Muga, A.; Guérin, D. M. Crystal Structure of the TemperatureSensitive and Allosteric-Defective GroELE461K. J. Struct. Biol. 2006, 155, 482−492. (158) Sot, B.; Galán, A.; Valpuesta, J. M.; Bertrand, S.; Muga, A. Salt Bridges at the Inter-Ring Interface Regulate the Thermostat of GroEL. J. Biol. Chem. 2002, 277, 34024−34029. (159) Aharoni, A.; Horovitz, A. Inter-Ring Communication Is Disrupted in the GroEL Mutant Arg13→Gly; Ala126→Val with Known Crystal Structure. J. Mol. Biol. 1996, 258, 732−735. (160) Ranson, N. A.; Clare, D. K.; Farr, G. W.; Houldershaw, D.; Horwich, A. L.; Saibil, H. R. Allosteric Signaling of ATP Hydrolysis in GroEL-GroES Complexes. Nat. Struct. Mol. Biol. 2006, 13, 147−152. (161) Horovitz, A.; Bochkareva, E. S.; Yifrach, O.; Girshovich, A. S. Prediction of an Inter-Residue Interaction in the Chaperonin GroEL from Multiple Sequence Alignment is Confirmed by Double-Mutant Cycle Analysis. J. Mol. Biol. 1994, 238, 133−138. (162) Chennubhotla, C.; Bahar, I. Markov Propagation of Allosteric Effects in Biomolecular Systems: Application to GroEL-GroES. Mol. Syst. Biol. 2006, 2, 36. (163) Keskin, O.; Bahar, I.; Flatow, D.; Covell, D. G.; Jernigan, R. L. Molecular Mechanisms of Chaperonin GroEL-GroES Function. Biochemistry 2002, 41, 491−501. (164) Falke, S.; Tama, F.; Brooks, C. L., 3rd.; Gogol, E. P.; Fisher, M. T. The 13 Å Structure of a Chaperonin GroEL-Protein Substrate Complex by Cryo-Electron Microscopy. J. Mol. Biol. 2005, 348, 219− 230. (165) Edelstein, S. J. Extensions of the Allosteric Model for Haemoglobin. Nature 1971, 230, 224−227. 6605

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606

Chemical Reviews

Review

Mitochondrial Chaperonin 60 Functions as a Single Toroidal Ring. J. Biol. Chem. 1992, 267, 695−698. (186) Nielsen, K. L.; Cowan, N. J. A Single Ring Is Sufficient for Productive Chaperonin-Mediated Folding in Vivo. Mol. Cell 1998, 2, 93−99. (187) Yang, D.; Ye, X.; Lorimer, G. H. Symmetric GroEL:GroES2 Complexes are the Protein-Folding Functional Form of the Chaperonin Nanomachine. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, E4298−E4305. (188) Sameshima, T.; Iizuka, R.; Ueno, T.; Funatsu, T. Denatured Proteins Facilitate the Formation of the Football-Shaped GroEL(GroES)2 Complex. Biochem. J. 2010, 427, 247−254. (189) Ye, X.; Lorimer, G. H. Substrate Protein Switches GroE Chaperonins from Asymmetric to Symmetric Cycling by Catalyzing Nucleotide Exchange. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, E4289− E4297. (190) Albe, K. R.; Butler, M. H.; Wright, B. E. Cellular Concentrations of Enzymes and Their Substrates. J. Theor. Biol. 1990, 143, 163−195. (191) Tran, Q. H.; Unden, G. Changes in the Proton Potential and the Cellular Energetics of Escherichia coli During Growth by Aerobic and Anaerobic Respiration or by Fermentation. Eur. J. Biochem. 1998, 251, 538−543. (192) Yamamoto, Y. Y.; Abe, Y.; Moriya, K.; Arita, M.; Noguchi, K.; Ishii, N.; Sekiguchi, H.; Sasaki, Y. C.; Yohda, M. Inter-Ring Communication is Dispensable in the Reaction Cycle of Group II Chaperonins. J. Mol. Biol. 2014, 426, 2667−2678. (193) Viappiani, C.; Abbruzzetti, S.; Ronda, L.; Bettati, S.; Henry, E. R.; Mozzarelli, A.; Eaton, W. A. Experimental Basis for a New Allosteric Model for Multisubunit Proteins. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 12758−12763. (194) Hammes, G. G.; Wu, C. W. Regulation of Enzyme Activity. The Activity of Enzymes Can Be Controlled by a Multiplicity of Conformational Equilibria. Science 1971, 172, 1205−1211. (195) Braig, K.; Adams, P. D.; Brünger, A. T. Conformational Variability in the Refined Structure of the Chaperonin GroEL at 2.8 Å Resolution. Nat. Struct. Biol. 1995, 2, 1083−1094.

(166) Kirschner, K.; Gallego, E.; Schuster, I.; Goodall, D. Cooperative Binding of Nicotinamide-Adenine Dinucleotide to Yeast Glyceraldehyde-3-Phosphate Dehydrogenase. I. Equilibrium and Temperature-Jump Studies at pH 8.5 and 40 °C. J. Mol. Biol. 1971, 58, 29−50. (167) Yifrach, O.; Horovitz, A. Coupling Between Protein Folding and Allostery in the GroE Chaperonin System. Proc. Natl. Acad. Sci. U. S. A. 2000, 97, 1521−1524. (168) Betancourt, M. R.; Thirumalai, D. Exploring the Kinetic Requirements for Enhancement of Protein Folding Rates in the GroEL Cavity. J. Mol. Biol. 1999, 287, 627−644. (169) Tehver, R.; Thirumalai, D. Kinetic Model for the Coupling Between Allosteric Transitions in GroEL and Substrate Protein Folding and Aggregation. J. Mol. Biol. 2008, 377, 1279−1295. (170) Netzer, W. J.; Hartl, F. U. Recombination of Protein Domains Facilitated by Co-Translational Folding in Eukaryotes. Nature 1997, 388, 343−349. (171) Jacob, E.; Horovitz, A.; Unger, R. Different Mechanistic Requirements for Prokaryotic and Eukaryotic Chaperonins: A Lattice Study. Bioinformatics 2007, 23, i240−i248. (172) Papo, N.; Kipnis, Y.; Haran, G.; Horovitz, A. Concerted Release of Substrate Domains from GroEL by ATP Is Demonstrated with FRET. J. Mol. Biol. 2008, 380, 717−725. (173) Kipnis, Y.; Papo, N.; Haran, G.; Horovitz, A. Concerted ATPInduced Allosteric Transitions in GroEL Facilitate Release of Protein Substrate Domains in an All-or-None Manner. Proc. Natl. Acad. Sci. U. S. A. 2007, 104, 3119−3124. (174) Farr, G. W.; Furtak, K.; Rowland, M. B.; Ranson, N. A.; Saibil, H. R.; Kirchhausen, T.; Horwich, A. L. Multivalent Binding of Nonnative Substrate Proteins by the Chaperonin GroEL. Cell 2000, 100, 561−573. (175) Sharma, S.; Chakraborty, K.; Müller, B. K.; Astola, N.; Tang, Y. C.; Lamb, D. C.; Hayer-Hartl, M.; Hartl, F. U. Monitoring Protein Conformation Along the Pathway of Chaperonin-Assisted Folding. Cell 2008, 133, 142−153. (176) Fersht, A. R.; Serrano, L. Principles of Protein Stability Derived from Protein Engineering Experiments. Curr. Opin. Struct. Biol. 1993, 3, 75−83. (177) Horovitz, A. Putting Handcuffs on the Chaperonin GroEL. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 10884−10885. (178) Vinckier, A.; Gervasoni, P.; Zaugg, F.; Ziegler, U.; Lindner, P.; Groscurth, P.; Plückthun, A.; Semenza, G. Atomic Force Microscopy Detects Changes in the Interaction Forces Between GroEL and Substrate Proteins. Biophys. J. 1998, 74, 3256−3263. (179) Villebeck, L.; Persson, M.; Luan, S. L.; Hammarström, P.; Lindgren, M.; Jonsson, B. H. Conformational Rearrangements of TailLess Complex Polypeptide 1 (TCP-1) Ring Complex (TRiC)-Bound Actin. Biochemistry 2007, 46, 5083−5093. (180) Frank, G. A.; Goomanovsky, M.; Davidi, A.; Ziv, G.; Horovitz, A.; Haran, G. Out-of-Equilibrium Conformational Cycling of GroEL Under Saturating ATP Concentrations. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 6270−6274. (181) Haraguchi, T.; Tominaga, M.; Matsumoto, R.; Sato, K.; Nakano, A.; Yamamoto, K.; Ito, K. Molecular Characterization and Subcellular Localization of Arabidopsis Class VIII Myosin, ATM1. J. Biol. Chem. 2014, 289, 12343−12355. (182) Rye, H. S.; Roseman, A. M.; Chen, S.; Furtak, K.; Fenton, W. A.; Saibil, H. R.; Horwich, A. L. GroEL-GroES Cycling: ATP and Nonnative Polypeptide Direct Alternation of Folding-Active Rings. Cell 1999, 97, 325−338. (183) Kawe, M.; Plückthun, A. GroEL Walks the Fine Line: The Subtle Balance of Substrate and Co-Chaperonin Binding by GroEL. A Combinatorial Investigation by Design, Selection and Screening. J. Mol. Biol. 2006, 357, 411−426. (184) Fridmann, Y.; Ulitzur, S.; Horovitz, A. In Vivo and in Vitro Function of GroEL Mutants with Impaired Allosteric Properties. J. Biol. Chem. 2000, 275, 37951−37956. (185) Viitanen, P. V.; Lorimer, G. H.; Seetharam, R.; Gupta, R. S.; Oppenheim, J.; Thomas, J. O.; Cowan, N. J. Mammalian 6606

DOI: 10.1021/acs.chemrev.5b00556 Chem. Rev. 2016, 116, 6588−6606