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Group II chaperonins are ATP-dependent ring-shaped complexes that bind non-native polypeptides and facilitate protein folding in archaea and eukaryotes. A built-in lid encapsulates substrate proteins within the central chaperonin chamber. Here we describe the fate of the substrate during the nucleotide cycle of group II chaperonins. The chaperonin substrate-binding sites are exposed and the lid is open in both the ATP-free and ATP-bound pre-hydrolysis states. ATP hydrolysis has a dual function in the folding cycle, triggering both lid closure and substrate release into the central chamber. Notably, substrate release can occur in the absence of a lid and lid closure can occur without substrate release. However, productive folding requires both events, so that the polypeptide is released into the confined space of the closed chamber where it folds. Our results show that ATP hydrolysis coordinates the structural and functional determinants that trigger productive folding.
Achieving correct protein folding is critical for cellular health and viability. Failure to fold and maintain protein homeostasis is associated with a growing number of diseases (Hartl and Hayer-Hartl, 2009; Powers et al., 2009). Accordingly, cell viability is dependent on a class of proteins called molecular chaperones, which bind non-native proteins and facilitate their folding (Bigotti and Clarke, 2008; Frydman, 2001; Hartl and Hayer-Hartl, 2009; Spiess et al., 2004). Among these, the group II chaperonins found in eukaryotic cells and archaea, have a unique ring-shaped structure that determines its functional characteristics (Bigotti and Clarke, 2008; Gómez-Puertas et al., 2004; Spiess et al., 2004). For instance, the eukaryotic chaperonin TRiC/CCT assists the folding of ~10% of newly translated proteins, including essential cytoskeletal proteins, cell cycle regulators and tumor suppressors (Thulasiraman et al., 1999; Yam et al., 2008). Intriguingly, many of its substrates, such as actin, cannot be folded by other chaperone systems (Spiess et al., 2004), suggesting TRiC possesses unique mechanistic features absent from other chaperones.
Group II chaperonins are large complexes consisting of two stacked rings of eight (or less frequently nine) subunits each (Bigotti and Clarke, 2008; Gómez-Puertas et al., 2004; Spiess et al., 2004). Individual subunits are generally different, ranging from one to four in archaea, to eight different subunits for TRiC/CCT. The general subunit architecture is conserved across group II chaperonins. Each subunit consists of an equatorial, ATP binding domain, an intermediate hinge domain, and an apical domain, which contains the substrate binding sites; a flexible protrusion extends from the apical domain and acts as a built-in lid. ATP binding and hydrolysis drives group II chaperonins through a conformational cycle that is not well understood. In the absence of nucleotide, the lid-containing segments are open, and the complex binds substrate. The open-state structures of, TRiC/CCT, and an archaeal chaperonin from Methanococcus maripaludis are remarkably similar (Booth et al., 2008; Pereira et al., 2010; Zhang et al., 2010). Incubation with hydrolyzable ATP induces a compact conformation, where the lid segments of each subunit form a beta-stranded iris that closes over the central cavity of the complex. The structure of this closed state is also virtually the same in eukaryotic and archaeal chaperonins (Booth et al., 2008; Cong et al., 2010; Ditzel et al., 1998; Pereira et al., 2010; Zhang et al., 2010). The presence of an intact lid is dispensable for substrate binding and ATP hydrolysis in both eukaryotic and archaeal chaperonins. However, the lid confers allosteric coupling of subunits within the complex and is essential for substrate folding (Kanzaki et al., 2008; Meyer et al., 2003; Reissmann et al., 2007). While the fully open and fully closed states are emerging in some structural detail, little is known about the trajectory of the chaperonin through the conformation cycle or how substrate folding is achieved (Bigotti and Clarke, 2008).
A number of studies using archaeal and eukaryotic chaperonins have suggested that ATP binding suffices to close the built-in lid and trigger substrate folding (Iizuka et al., 2003; Llorca et al., 2001; Villebeck et al., 2007; Stuart et al., 2010). Subsequent ATP hydrolysis would serve to reopen the lid and release the folded protein. In contrast, other studies reported that ATP binding alone is unable to close the lid or promote substrate folding (Bigotti et al., 2006; Meyer et al., 2003; Reissmann et al., 2007). Instead, these studies identified the transition state of ATP-hydrolysis as the critical step in the ATPase cycle that promotes the closed conformation (Meyer et al., 2003; Reissmann et al., 2007).
A fundamental question for group II chaperonins concerns the fate of the substrate during the ATPase cycle. The current model proposes that group II chaperonins do not release the substrate during folding (Gómez-Puertas et al., 2004; Stuart et al., 2010). Instead, ATP binding would cause the apical domains with their bound substrate to move, and this movement mechanically forces substrate folding. In this view, substrate liberation occurs after nucleotide hydrolysis, perhaps after nucleotide release and the subsequent return of the chaperonin to the open state.
Some experimental results are not reconciled easily with the “mechanical force” model. The substrate binding sites of group II chaperonins have been mapped to the vicinity of helix 11 (Spiess et al., 2006), which is unavailable to the central cavity in the ATP-induced closed state. The mechanical model of group II chaperonin action suggests that the cavity is not necessarily a folding chamber per se, rather, it is used as a mechanical scaffold for active remodeling. This led to the suggestion that the lids primarily to assist in the conformational cycle of the chaperonin (Kanzaki et al., 2008). However, ATP incubation of a group II chaperonin lacking a lid (Cpn-Δlid) produces an identical conformation to that of wild type but is unable to promote substrate folding (Reissmann et al., 2007; Zhang et al., 2010). Thus, the movement of the apical domains does not require the presence of the lid; however, their movement alone is insufficient to promote folding.
Here we use the group II chaperonin from mesophilic archaea Methanococcus maripaludis, herein Cpn, to define the fate of the polypeptide substrate during the conformational cycle of group II chaperonins. The allosteric regulation and structure of this Cpn are similar to those of TRiC/CCT (Pereira et al., 2010; Reissmann et al., 2007; Zhang et al., 2010). We find that ATP hydrolysis has a dual role in group II chaperonin function, promoting both lid closure and release of the substrate into the cavity. Importantly, both events must occur for successful substrate folding. We suggest an alternate model for group II chaperonin function, whereby folding relies on the release of the substrate into a unique chemical environment within the closed chamber.
We initially examined whether the folding reaction is completed within the closed central chamber of group II chaperonins. In principle, folding of a polypeptide with a strict chaperonin requirement, i.e. a stringent substrate, could require several cycles of Cpn binding and release (Fig. 1A). Alternatively, the substrate could fold in a single ATPase cycle event, without requiring multiple rounds of binding and release. To test these possibilities, we employed rhodanese, a stringent Cpn substrate (Martin et al., 1991). 35S-rhodanese binds to nucleotide-free Cpn in an unstructured, proteinase K (herein PK)-sensitive state (Fig. 1A, left arrow; Fig. 1B, lane 2 bottom panel and Fig. 1C for Native gel analysis). Addition of ATP induces lid closure and encapsulates the substrate within the closed chamber (Meyer et al., 2003; Reissmann et al., 2007)(Fig. 1A). Upon closure, the Cpn lid segments and the encapsulated 35S-rhodanese are protected from proteolytic digestion (Fig. 1B, lane 3). Importantly, ATP addition causes the time-dependent folding of rhodanese (Fig. 1D, red symbols). Comparing the kinetics of rhodanese folding (t1/2~12 min) with the estimated kinetics of a single round of ATP hydrolysis (Bigotti et al., 2006; Reissmann et al., 2007) indicates that completion of rhodanese folding involves several cycles of ATP binding and release. Importantly, addition of protease at any time following ATP addition interrupted the folding reaction (Fig. 1D, PK, shown for t=0 and t=13 min). Since PK can only degrade the substrate if the lid is open, this result suggests that the Cpn-substrate complex undergoes repeated cycles of ATP-driven opening and closing during the folding reaction.
We next examined whether such iterative cycling is required to achieve folding by exploiting the observation that addition of AlFx together with ATP locks group II chaperonins in a symmetrically closed state that fully encapsulates the substrate (Meyer et al., 2003)(Fig. 1A; right arrow). The ATP-AlFx-induced state of Cpn-rhodanese was locked closed, leading to full proteolytic protection of both Cpn and substrate (Fig. 1B, lane 4) and a characteristic electrophoretic migration shift on native gels (Fig. 1C). Under these conditions, ATPase cycling is interrupted (Fig. S1A), and the substrate undergoes a single round of binding and encapsulation, allowing us to evaluate whether iterative cycling is required for group II chaperonin folding (Fig. 1E). Strikingly, the rate and yield of rhodanese folding under these non-cycling conditions were identical to those observed for the actively cycling chaperonin (Fig. 1E). Furthermore, addition of PK to the ATP-AlFx reaction did not interrupt folding, confirming that there was no reopening of the Cpn and no release of the non-native substrate under these conditions. We conclude that the closed chamber of group II chaperonins is the folding active compartment. Furthermore, a single round of encapsulation in this chamber can achieve maximum rhodanese folding, with similar kinetics and yield as observed under cycling conditions. Thus, although iterative cycling does occur, it is not strictly required for Cpn-dependent folding.
To examine whether ATP binding suffices to promote the folding active state of group II chaperonins, we specifically impaired the ATPase active site by targeting Asp386, which is essential to coordinate the water molecule that participates as a nucleophile during the hydrolysis of the phosphate-anhydride bond (Cpn-D386A; Fig. 2A). Cpn-D386A cannot hydrolyze ATP but retains efficient ATP binding [data not shown and (Reissmann et al., 2007)]. Importantly, unlike Cpn-WT, Cpn-D386A is unable to fold the stringent Cpn substrates rhodanese (Fig. 2B) and malate dehydrogenase (data not shown). This demonstrates that ATP binding is insufficient to induce the fully folding-active state observed upon ATP hydrolysis.
We next assessed the proposal that ATP binding leads to partial (Clare et al., 2008) or full (Iizuka et al., 2003; Llorca et al., 2001) lid closure. To this end, the structure of ATP-bound Cpn-D386A was derived to 15 Å resolution by single particle Cryo-EM (Fig. 2C, blue). Comparison of these structures with the ATP-free and ATP-bound states of Cpn-WT, derived to 10Å and 6Å respectively, revealed the conformational changes induced by ATP binding, distinguishing them from those induced by ATP-hydrolysis (Fig. 2D; Fig. S2). ATP incubation with Cpn-WT induces lid closure, yielding a symmetrically closed structure similar to that previously obtained for Cpn-WT with ATP-AlFx (Fig. 2C cyan and Fig. S5)(Pereira et al., 2010; Zhang et al., 2010). In contrast, ATP binding to Cpn-D386A yielded an open structure that resembled the nucleotide free state (Fig. 2D for overlay and Fig. S2A, B). Further addition of AlFx did not result in closure (not shown). Despite leaving the lid open, ATP binding induced a ~20Å constriction in the chaperonin opening (Fig. 2D and Fig. S2B, 110 Å span versus 130 Å in the ATP-free state). Closer analysis of the conformational changes in a single subunit indicated that ATP binding induces an en masse rigid body tilt of the entire intermediate and apical domains towards the ATP-binding equatorial domain (Fig. S2C). We conclude that ATP-binding is insufficient to close the lid but triggers domain movements that lead, upon hydrolysis, to the closed state. These results are consistent with fluorescence experiments on the thermosome from Thermoplasma acidophilum, indicating a rapid re-arrangement attributed to ATP binding, followed by a slower re-arrangement attributed to ATP hydrolysis and lid closure (Bigotti and Clarke, 2005; Reissmann et al., 2007).
The effects of ATP binding on the conformation of both the substrate and the lid were further examined using biochemical assays (Fig. 2E–F). As described above, addition of ATP·AlFx to Cpn-WT stabilizes the closed state, locking the encapsulated substrate inside the chamber and leading to proteolytic protection of both the lids and the substrate (Fig. 1B, 1C; Fig. 2E and Fig. 2F, lane 3; top panel for Cpn, bottom panel for 35S-Rho for 35S-Rho-Cpn-WT complex). Both ATP and ATP-AlFx induce a structurally similar closed state in Cpn-WT (e.g. Fig. 2C, right panel) but the ATP-AlFx state displays a characteristic faster electrophoretic migration on native gels [Fig. 2E; a similar effect is observed for TRiC/CCT; (Meyer et al 2003)]. In contrast to Cpn-WT, incubation of Cpn-D386A with either ATP or ATP·AlFx failed to produce the signature mobility shift (Fig. 2E). Furthermore, both the lid and the substrate remained in a largely unstructured, protease sensitive state upon ATP binding (Fig. 2F, lanes 5, 6, 7), consistent with the result that ATP binding leaves Cpn in an open state (Fig. 2C, D). Importantly, the lid also remains open under conditions where only one ring binds nucleotide [0.2 mM (Reissmann et al., 2007)] or if Cpn-WT is incubated with the non-hydrolyzable ATP analogues AMPPNP or ATPγS (at either 0.2 mM or 1 mM; data not shown), further supporting the conclusion that ATP binding to either one ring or both does not suffice to close the lid.
Lid closure and substrate encapsulation are essential for folding substrates such as actin for TRiC (Meyer et al., 2003) and rhodanese (Reissmann et al., 2007) and MDH for Cpn (Fig. S3A, B). We next examined whether lid closure modulates the interaction of the substrate with the chamber. The “mechanical force” model proposes that the chaperonin does not release the substrate proteins into the closed cavity; in this scenario the chaperonin-substrate interaction persists in the closed state leading to the mechanical remodeling of the substrate conformation (Fig. 3A(i), left) (Llorca et al., 2001). Alternatively, ATP hydrolysis could promote substrate release into the closed chamber (Fig. 3A(ii), left). Since monitoring the substrate-chaperonin interaction inside the closed chamber is complicated by the presence of the lid, we exploited the previously characterized Cpn-Δlid variant that lacks the entire lid-forming segments (Pereira et al., 2010; Reissmann et al., 2007; Zhang et al., 2010). Importantly, Cpn-Δlid achieves the same ATP-induced “closed” conformation as Cpn-WT (Zhang et al., 2010) and its ATPase activity and substrate binding ability are unaffected (Reissmann et al., 2007). These features of Cpn-Δlid allowed us to distinguish between the above models (Fig. 3A, right panels). Thus, the model that proposes that the polypeptide remains associated with the chaperonin throughout the ATPase cycle predicts that the substrate will remain bound to Cpn-Δlid upon addition of ATP or ATP·AlFx (Fig. 3A(i), ‘Δlid’ right). In contrast, if ATP weakens the chaperonin-substrate interaction, the absence of the lid will allow the polypeptide to diffuse away from the chaperonin (Fig. 3A(ii), ‘Δlid’ right). Of note, Cpn-Δlid cannot promote folding of substrates such as rhodanese and MDH (Fig. S3A, B) and (Reissmann et al., 2007)), thus substrate release from Cpn-Δlid cannot be ascribed to completion of folding.
Purified 35S-rhodanese·Cpn complexes were incubated in the presence or absence of ATP for 10 min and analyzed using native gels followed by autoradiography (Fig. 3B). Cpn-WT co-migrates with the substrate under both conditions (Fig. 3B, WT), as expected given that 35S-rhodanese is encapsulated in the closed complex (See Fig. 1B and C). The small ATP-induced reduction in bound substrate is presumably due to loss through ATPase cycling and/or folding (see below, Fig. 3C). Strikingly, incubation of Cpn-Δlid with ATP led to a dramatic reduction in the amount of Cpn-bound rhodanese (Fig. 3B, Δlid). This ATP-dependent loss of rhodanese required ATP hydrolysis, as it was not observed when the Cpn-Δlid also carried the D386A mutation (Fig. 3B, Δlid/D386A). Similar results were obtained for other Cpn-bound polypeptides, including MDH (data not shown) and actin (see below Fig. 5).
The ATP-induced reduction in Cpn-substrate affinity was further evinced through the use of a “Trap”, a modified GroEL that scavenges non-native polypeptides (Fig. 3C) (Frydman and Hartl, 1996). Trap will not bind to folded rhodanese but will bind to non-native polypeptides once they are released from the Cpn (Frydman and Hartl, 1996) (Fig. 3C, see Trap lane). For all Cpn variants tested, little or no 35S-rhodanese was captured by the Trap in the absence of ATP, suggesting that rhodanese binds stably to all nucleotide free Cpn variants, and cannot be displaced by the Trap (Fig. 3C, -ATP). Addition of ATP to Cpn-WT allowed a fraction of rhodanese to bind to the more rapidly migrating Trap (Fig. 3C, WT+ATP). Comparing the WT incubations in the presence and absence of Trap (i.e. Fig. 3B and 3C) suggests that during normal ATP cycling a fraction of the substrate is released in a non-native form that rebinds to the chaperonin for another round of folding. This non-native polypeptide is captured by the Trap, which thus prevents Cpn rebinding and interrupts the cycle. Importantly, addition of ATP to Cpn-Δlid-35S-rhodanese caused a near complete transfer of the bound polypeptide to the Trap (Fig. 3C, Δlid), indicating that ATP induces substrate release from the chaperonin. Furthermore, no increase in substrate transfer to the Trap was observed upon ATP addition to Cpn-Δlid D386A (Fig. 3C, Δlid/D386A) indicating that substrate dissociation from Cpn requires ATP hydrolysis.
The experiments above show that ATP hydrolysis has a function that is completely lid-independent, namely, to release the substrate from the chaperonin binding sites. We next employed ATP·AlFx, which mimics the trigonal-bipyramidal transition state of ATP hydrolysis (Meyer et al., 2003) (Fig. 3D). As with Cpn-WT (Fig. S1A), the addition of AlFx to Cpn-Δlid immediately arrests its ATPase activity, suggesting that inhibition of ATP hydrolysis and trapping of the closed state occurs after a single cycle (Fig. S3C). Whereas incubation of Cpn-WT-35S-rhodanese with ATP·AlFx closes the chamber and encapsulates the substrate (Fig. 3E; Fig. 3F), the substrate remains protease sensitive following incubation of Cpn-Δlid-35S-rhodanese with ATP·AlFx (Fig. 3E). Native gel analysis showed that Cpn-Δlid with ATP·AlFx undergoes the same signature shift as Cpn-WT, consistent with structural analyses showing both Cpns adopt the same closed conformation upon incubation with ATP·AlFx (Pereira et al., 2010; Zhang et al., 2010). ATP·AlFx induced a complete release of a broad panel of polypeptides (Fig. 3F for 35S-rhodanese; Fig. S3D-G for other substrates), indicating that ATP hydrolysis blocks general access to the substrate binding sites. The same conclusion was reached using size exclusion chromatography of purified Cpn-35S-rhodanese complexes incubated in the presence or absence of ATP·AlFx and analyzed on a Bio-Sil SEC-400-5 column (Fig. S3H). This experiment also indicated that ATP·AlFx induces full substrate release from Cpn-Δlid.
The effect of nucleotide hydrolysis on Cpn-substrate interactions was further examined using rhodanese carrying the environmentally sensitive fluorescent moiety Nile Red (Kim et al., 2005) (herein NR-Rho; Fig. 3G–I). In free solution, NR-Rho exhibits a low fluorescence emission spectrum characteristic of an aqueous, polar environment, with a maximum at ~650nm (Fig. 3G, gray trace). However, binding to Cpn caused a fluorescence intensity increase, as well as a blue shift of the maximal intensity to ~630nm (Fig. 3G, red trace for Cpn-Δlid; similar results obtained for Cpn-WT, data not shown). This change in fluorescence upon Cpn-binding is diagnostic for rhodanese occupying a more hydrophobic environment (Kim et al., 2005). We used the maximal fluorescence at 630 nm to monitor the effect of nucleotides on the substrate-chaperonin interaction. The Cpn-Δlid-NR-Rho fluorescence signal remained stable in the absence of nucleotide (Fig. 3H, 3I; red traces). Addition of ATP produced a rapid decay in fluorescence intensity (Fig. 3H, ‘+ATP’, blue trace). This supports our previous conclusion that ATP cycling by Cpn leads to substrate release. Addition of ATP·AlFx yielded similar results (Fig. 3I, ‘+ATP·AlFx’, cyan trace), supporting the idea that the ATP hydrolysis transition state induces substrate release. We conclude that ATP hydrolysis has a dual function within the chaperonin cycle; it promotes lid closure (Fig. 2) and also triggers substrate release from the chaperonin binding sites (Fig. 3). Strikingly, the latter function is not dependent on the presence of a lid.
A simple model explaining our results is that the ATP-induced Cpn conformation no longer exposes the substrate binding sites. We tested this model using an order of addition experiment (Fig. 4). In the control condition (Fig. 4, Ctrl), substrate was added to the open, apo-Cpn, which exposes the substrate binding sites. The second condition added the substrate first, prior to incubation with ATP·AlFx (Fig. 4, S→A); this condition measured the extent of ATP·AlFx-induced substrate release. In the third condition, we incubated with ATP·AlFx first, and then added substrate to the chaperonin (Fig. 4, A→S); this measured the ability of an ATP·AlFx-preincubated closed complex to bind substrate (Fig. 4A). If the binding sites are still available in the closed state, we might expect some substrate binding for closed Cpn-Δlid in the A→S condition, which still retains a large opening allowing access to the central cavity (Pereira et al., 2010; Zhang et al., 2010). Since the pore size may restrict polypeptide entry to the cavity, and may sterically interfere with substrate binding we used both rhodanese (Fig. 4Bi) and a small 12 mer peptide substrate (herein PepB; Fig. S3G and Fig. 4Bii). The small peptide substrate should be able to freely diffuse inside the closed chamber in the Cpn-Δlid.
In the absence of nucleotide, both substrates bound to Cpn-WT and Cpn-Δlid (Ctrl; Fig. 4B, 4C and Fig. S3G). As expected, addition of ATP·AlFx to the Cpn-substrate complex (Fig. 4, S→A) promoted substrate encapsulation for Cpn-WT (WT S→A; Fig. 4B, 4C) and substrate release for Cpn-Δlid (Δlid S→A; Fig. 4B, 4C). In the case of A→S, closing the Cpn-WT chamber with ATP·AlFx precluded substrate binding, thus the closed lid blocks access to the central cavity (WT A→S; Fig. 4B; scheme in Fig. 4C). For Cpn-Δlid, substrate should bind the chaperonin in the A→S condition provided that the binding sites are still available in the closed conformation. This was not the case; instead the ATP-AlFx preincubated Cpn-Δlid was unable to bind either 35S-rhodanese or the small PepB (Fig. 4B; Cpn-Δlid compare S→A and A→S). Thus, the ATP·AlFx state of Cpn-Δlid no longer exposes the substrate binding sites. Given that the ATP·AlFx conformations of Cpn-WT and Cpn-Δlid are virtually identical (Pereira et al., 2010; Zhang et al., 2010), these experiments show that the substrate binding sites are no longer available upon ATP-hydrolysis.
What is the possible mechanism for substrate release in group II chaperonins? A structural analogy with the distantly related bacterial group I chaperonins, e.g. GroEL, is not possible, given that they use a detachable lid, GroES, which upon ATP binding, both caps the cavity and displaces the substrate. In contrast, we show that substrate release in group II chaperonins is lid independent and requires ATP-hydrolysis.
Closer examination of Cpn structures in the open and closed states led to a hypothesis for how ATP hydrolysis induces substrate eviction (Fig. 5A) (Pereira et al., 2010; Zhang et al., 2010). In the open state, the substrate binding region around helix 11 is well exposed (Fig. 5A, pink in left panel) (Spiess et al., 2006) leaving ample space to accommodate the bound substrate. In contrast, the closed state brings the apical domains from adjacent subunits into close proximity (Fig. 5A). Closure causes helix 11 to form a tightly packed interface with a loop spanning residues 327-331 in its neighboring subunit (Fig. 5A, cyan). Such lateral intra-ring contacts might displace the substrate from its binding site, causing the 327-331 region to acts as a ‘release loop for the substrate’ (herein rls loop). To disrupt this lateral interface, we made Ala substitutions in four loop residues making key contacts with helix 11 yielding the Cpn-rls variants (Fig. 5A, T327A, N328A, K330A, and D331A). To better understand the role of the rls loop within the chaperonin cycle, we used Cryo-EM to obtain a detailed structural characterization of the conformation of both Cpn-rls and Cpn-rls-Δlid in the presence or absence of ATP or ATP·AlFx (Fig. S4A for Fourier Shell Correlation analysis of resolutions; S4B for Cpn-rls-Δlid and S5 for Cpn-rls). The rls chaperonins achieve essentially the same closed state as the wild type counterparts (Fig. 5B, 5C; Fig. S4B for Cpn-rls-Δlid; Fig. S5 for Cpn-rls). Consistent with their ability to reach a closed state, the Cpn-rls mutants were competent for ATP binding and hydrolysis (data not shown).
We initially focused on Cpn-rls-Δlid, since the absence of a lid simplifies analysis of substrate release (Fig. 5D-F). Cpn-WT and Cpn-Δlid served as controls. In the absence of ATP, all chaperonins bound rhodanese and actin efficiently, as shown by native gel analysis (Fig. 5D). Strikingly, Cpn-rls-Δlid was incapable of releasing either substrate in the presence of ATP, unlike Cpn-Δlid (Fig. 5D, compare lane 6 to lane 4). This suggests that the lateral contacts between helix 11 and the rls loop 327-331 are indeed important for releasing the substrate upon ATP-hydrolysis.
We next examined the effect of the transition state mimic ATP·AlFx (Fig. 5E). Native gel analysis showed that Cpn-rls-Δlid adopts the same fast migrating conformation observed for Cpn-Δlid and Cpn-WT (Fig. 5E, Coomassie blue panel) consistent with the Cryo-EM analysis. Surprisingly, unlike ATP, incubation with ATP·AlFx caused Cpn-rls-Δlid to efficiently release all the substrates tested (Fig. 5E for rhodanese and actin). This observation was striking given the apparent similarity between the ATP and ATP·AlFx structures of Cpn-rls variants (Fig. 5E and Fig. S4B). It thus appears that, in the rls mutant the conformation promoting substrate release cannot be stably populated by ATP alone whereas ATP·AlFx can stabilize this state and evict the substrate.
Fluorescence spectroscopy provided independent support for the above conclusions. As for Cpn-Δlid, NR-Rho bound to Cpn-rls-Δlid had an emission spectrum characteristic of a hydrophobic environment (data not shown). In contrast to Cpn-Δlid (Fig. 5Fi, blue trace), ATP incubation did not cause any appreciable change in the fluorescence of NR-Rho bound to Cpn-rls-Δlid (Fig. 5Fii, blue trace), indicating that ATP alone cannot release the bound substrate. However, when AlFx was added to an ongoing incubation of NR-Rho·Cpn-rls-Δlid with ATP, the fluorescence rapidly dropped, indicating substrate release from the chaperonin (Fig. 5Fii, cyan trace). A similar reduction in fluorescence intensity was observed if ATP and AlFx were added together, but was absent if only AlFx was added (not shown). We conclude that weakening the lateral contacts between helix 11 and its neighboring subunit prevents substrate release, even though Cpn-rls-Δlid can hydrolyze ATP and achieve the closed state. However, stabilizing the post-hydrolysis state by addition of AlFx populates the conformation that evicts the substrate.
We next examined the effect of the rls mutations in the Cpn with the intact lid (herein Cpn-rls, Fig. 6). Detailed structural analyses of the ATP and ATP·AlFx induced states in both Cpn-WT and Cpn-rls revealed interesting differences between these chaperonins (Fig. 6B–F; Fig. S5). Single particle Cryo-EM reconstructions were obtained to 4–6Å for both chaperonins in the presence of either ATP or ATP·AlFx (Cpn-WT-ATP 6Å, Cpn-WT-ATP·AlFx 4.3Å, Cpn-rls-ATP 5Å, Cpn-rls- ATP·AlFx 6Å, Fig. 6 and Fig. S5). Models of these structures were then built by flexible fitting into the density map with Rosetta (Fig. 6 and Fig. S5A; see Fig. S5B for goodness of fit between model and density map) (DiMaio et al., 2009). Cpn-rls achieved a closed state with either ATP or ATP·AlFx, similar to those obtained with Cpn-WT. Notably, superimposition of the structures of Cpn-WT and Cpn-rls in the different nucleotide states revealed variations in their structure, particularly in the region corresponding to the apical domains (Fig. 6C–F; i. top view of superimposed EM density maps). These differences were also evident when comparing the apical domain regions in the respective chaperonin models (Fig. 6C–F; ii. detail of apical domain and lid for a subunit within the complex). The ATP (magenta) and ATP·AlFx (blue) states of Cpn-WT were essentially identical (Fig. 6C). Thus, ATP·AlFx generates the same closed state observed under ATP-cycling conditions (e.g. Fig. 1E). Importantly, we observed a shift in the apical domain regions between the closed Cpn-rls states induced by ATP (yellow) and ATP·AlFx (cyan) (Fig. 6D). Cpn-rls-ATP also exhibited noticeable differences with both closed WT structures (e.g. Fig. 6F). The apical domain protrusions in Cpn-rls-ATP are shifted clockwise, and the apical domains including the lid, are tilted up compared to the ATP·AlFx state, exhibiting significant variations in helix 11 (ii. red arrow) and the rls loop (ii. blue arrow). In contrast, the ATP·AlFx states of Cpn-WT and Cpn-rls were nearly identical (Fig. 6E). These structural analyses demonstrate that even though Cpn-rls can close with ATP, impairment of the helix 11/loop 327-331 contacts results in aberrant intra-ring interactions between the apical domains. This is consistent with the inability of Cpn-rls-Δlid to release the substrate in the presence of ATP (See Fig 5Fii.). Furthermore, ATP·AlFx induces a closed conformation in Cpn-rls that is indistinguishable from the closed state of Cpn-WT with either ATP or ATP-AlFx. This is consistent with, and explains, the finding that ATP·AlFx leads to substrate release in Cpn-rls-Δlid (See Fig 5Fii.)
The identification of a mechanism that evicts the bound polypeptide upon closure allowed us to test the relevance of substrate release for the folding cycle. First, the ability of Cpn-rls to encapsulate a bound substrate was examined by native gel analysis (Fig. 6G, i) and PK digestion (Fig. 6G, ii) as shown above for Cpn-WT. Incubation of the Cpn-rls with rhodanese yielded a binary complex that behaved exactly as that of Cpn-WT (Fig. 6G, i). Protease digestion analysis indicated that, in the absence of nucleotide, the substrate binds in an unstructured conformation (Fig. 6G, ii, lane 2). Importantly, incubation with either ATP or ATP·AlFx led to proteolytic protection of both the chaperonin lid segments (Fig. 6G, ii, lanes 3, 4, top panel) and the bound 35S-rhodanese (Fig. 6G, ii, lanes 3, 4, bottom panel). Thus, both ATP and ATP·AlFx induce stable lid closure and fully encapsulate the substrate within the central chamber of Cpn-rls.
Rhodanese-chaperonin complexes were prepared for Cpn-rls and Cpn-WT, which served as a control (Fig. 6H). As expected, addition of ATP or ATP-AlFx to the Cpn-WT complex induced rhodanese folding (Fig. 6H, black traces). Strikingly, addition of ATP to the Cpn-rls complex failed to promote rhodanese folding, even though the substrate was encapsulated within the closed chamber (Fig. 6H, green trace). We hypothesized that failure to fold stems from the failure to release the bound substrate into the central chamber. Therefore, we tested the effect of ATP·AlFx, which should promote substrate release (See Fig. 5). Addition of ATP-AlFx to the Cpn-rls reaction caused efficient rhodanese folding (Fig. 6H). These experiments indicate that lid closure and substrate encapsulation are by themselves, unable to promote substrate folding. Importantly, they demonstrate that substrate release into the central closed chamber is essential for productive folding by group II chaperonins.
Our study defines how the ATPase cycle of group II chaperonins modulates the interaction with substrates (Fig. 7). We find that ATP hydrolysis triggers substrate release from the chaperonin through a hitherto unanticipated mechanism involving lateral intra-ring contacts between adjacent apical domains. Given the high degree of structural and mechanistic similarity among all group II chaperonins our findings have broad implications to understand cellular folding in eukaryotes and archaea.
To resolve the role of ATP binding in group II chaperonin action we specifically impaired hydrolysis by targeting D386 (Ditzel et al., 1998). We find that ATP binding alone does not support substrate folding or lid closure, similar to previous findings for TRiC/CCT (Meyer et al., 2003). ATP binding does induce a conformational change that constricts the Cpn chamber entrance from 130Å to 110Å (Fig. S2; Fig. 7A, B). The movement results from an en bloc counterclockwise rotation of the intermediate and apical domains with respect to the equatorial, ATP binding domain (Fig. S2). Notably, a similar concerted movement of intermediate and apical domains has previously been observed during lid closure for TRiC/CCT (Booth et al., 2008). Our results indicate that ATP hydrolysis is generally required for lid closure and folding in group II chaperonins, underscoring the general conservation of architecture and mechanism between archaeal and eukaryotic chaperonins.
ATP hydrolysis has a dual role within the group II chaperonin cycle: it both triggers lid closure and releases the substrate from the apical domains into the cavity (Fig. 7). Importantly, both events are required for productive folding. Lid closure in the absence of substrate release is also insufficient to achieve folding (Fig. 6H). This contrasts with the previously proposed mechanical force model, which suggests that folding occurs through movement of the apical domains without releasing the substrate. The observation that we can generate a chaperonin state that can close the lid without releasing the substrate, raises the possibility that lid closure and substrate encapsulation precede release (Fig. 7A, B, shown in brackets). Such a mechanism would ensure that substrates are confined inside the chamber prior to their release, thereby avoiding the premature escape of non-native aggregation-prone species into the cytosol.
The released substrate folds while encapsulated in the central cavity (Fig. 1E). No folding was observed when the substrate was released into the bulk solution (Cpn-Δlid, Fig. S3A, B), indicating that the chemical and physical characteristics of the closed central chamber create a folding active compartment. The nature of this compartment will depend on the side chains exposed in the closed state as well as the effect of crowding on the solvent properties of the chamber (Tang et al., 2006). The hetero-oligomeric nature of most group II chaperonins may lead to a diversification of the chamber properties (Cong et al., 2010), which may contribute to the folding of specific substrates. While a single encapsulation step suffices for optimal folding in vitro, in the cellular context cycling on and off the chaperonin likely fulfills an important homeostatic function. Thus, each cycle may expose the substrate polypeptide to additional folding cofactors as well as quality control components thereby preventing folding-incompetent proteins from clogging the chaperonin. How the balance between processivity and clearance is achieved in vivo is an important question for future research.
ATP hydrolysis releases the bound substrate from its chaperonin binding sites through a hitherto unanticipated mechanism; namely, a conformational change that brings together vicinal apical domains. This creates a lateral interface between helix 11 of one domain and loop 327-331 in the adjacent subunit (Fig. 7B, 7C, pink and green respectively). The crystal structure suggests that formation of this lateral H-bonded network is incompatible with substrate binding. We hypothesize that these lateral inter-subunit contacts displace the substrate from its binding site (pink in Fig. 7). The precise mechanism of release will require further investigation. One possibility is that the inter-subunit interaction sterically interferes with substrate rebinding during thermal breathing of the chaperonin-substrate interaction. Alternatively, the helix 11-rls loop interaction could create an entropic zipper that displaces the substrate. Yet another model is that the rls interaction helps stabilize a conformation that cannot bind substrate. The presence of ATP-AlFx may compensate energetically for the loss of the H-bonded network between substrate binding region and rls loop, and by itself induce the subtle conformational change required to release the substrate.
The unique nature of substrate release in group II chaperonins may have important implications for hetero-oligomeric chaperonins, particularly in light of recent findings that different subunits recognize distinct motifs in the substrate (Spiess et al., 2006). Since the mechanism for substrate release depends on the nature of a specific inter-subunit interface, rather than a general GroES binding interface as observed in GroEL, the local kinetics of substrate release could vary for a specific apical domain (e.g. shading in Fig. 7B). The order of release of different regions of a substrate polypeptide from their respective subunits may be influenced by the strength of this interaction vis-à-vis the timing of conformational change and formation of the lateral interface. The ensuing sequential mechanism of substrate release from the chaperonin could provide exquisite control of the folding pathway of the substrate, which in turn contributes to the unique ability of these chaperonins to fold specific proteins. One could envision that subunit-specific substrate remodeling and/or ordered release directs substrates of group II chaperonins along specific folding trajectories. Exploring these exciting possibilities may have profound implications for our understanding and ability to control cellular folding pathways.
All Cpn variants were produced by site-directed mutagenesis; purification and functional analyses were performed as described (Reissmann et al., 2007). MDH refolding was performed as in (Hayer-Hartl, 2000). Fluorescent proteins were generated as in (Kim et al., 2005) and fluorescence measured on a Fluorolog-3 Fluorimeter (Horiba-Jobin Yvon).
Samples were embedded in vitreous ice on 400-mesh R1.2/1.3 Quantifoil grids (Quantifoil Micro Tools GmbH, Jena Germany) and imaged on a JEM3200FSC electron cryo-microscope and JEM2010F elecron cryo-microscope (JEOL Inc, Tokyo, Japan) with field emission guns. Details about the image acquisition parameters are in Table S1 of the Supplementary Materials. The image processing steps followed those described in (Baker et al., 2010). The figures were prepared using MacPyMOL (http://www.pymol.org) and UCSF Chimera (Pettersen et al., 2004).
We thank Drs. W.E. Moerner for providing Nile Red maleimide. We also thank members of the Frydman lab and Raul Andino for critical reading of the manuscript; Frank DiMaio and David Baker for advice on modeling and Erik Miller for useful discussions and Jeremy England and Erik Miller for providing PepB. This work was supported by NIH grants GM74074 (to JF); 5PN2EY016525 (to JF and WC); P41RR002250 (to WC). NRD was a recipient of pre-doctoral ARCS fellowships and was supported by NIH training grant (T32-007276). JZ was a recipient of a NIH training grant on Nanobiology (R90 DK071054 and T90 DA022885)
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