- Research article
- Open Access
Structural insight into the cooperation of chloroplast chaperonin subunits
© Zhang et al. 2016
Received: 8 January 2016
Accepted: 29 March 2016
Published: 12 April 2016
Chloroplast chaperonin, consisting of multiple subunits, mediates folding of the highly abundant protein Rubisco with the assistance of co-chaperonins. ATP hydrolysis drives the chaperonin allosteric cycle to assist substrate folding and promotes disassembly of chloroplast chaperonin. The ways in which the subunits cooperate during this cycle remain unclear.
Here, we report the first crystal structure of Chlamydomonas chloroplast chaperonin homo-oligomer (CPN60β1) at 3.8 Å, which shares structural topology with typical type I chaperonins but with looser compaction, and possesses a larger central cavity, less contact sites and an enlarged ATP binding pocket compared to GroEL. The overall structure of Cpn60 resembles the GroEL allosteric intermediate state. Moreover, two amino acid (aa) residues (G153, G154) conserved among Cpn60s are involved in ATPase activity regulated by co-chaperonins. Domain swapping analysis revealed that the monomeric state of CPN60α is controlled by its equatorial domain. Furthermore, the C-terminal segment (aa 484–547) of CPN60β influenced oligomer disassembly and allosteric rearrangement driven by ATP hydrolysis. The entire equatorial domain and at least one part of the intermediate domain from CPN60α are indispensable for functional cooperation with CPN60β1, and this functional cooperation is strictly dependent on a conserved aa residue (E461) in the CPN60α subunit.
The first crystal structure of Chlamydomonas chloroplast chaperonin homo-oligomer (CPN60β1) is reported. The equatorial domain maintained the monomeric state of CPN60α and the C-terminus of CPN60β affected oligomer disassembly driven by ATP. The cooperative roles of CPN60 subunits were also established.
Cellular protein homeostasis is regulated by molecular chaperones [1, 2]. Chaperonins are a subfamily of chaperones which have diverged into two distinct groups [1, 3, 4]. Group I chaperonins, which are found in prokaryotes and in organelles of prokaryotic origin, include GroEL/ES in E. coli, Hsp60/10 in mitochondria, and Cpn60/20 in chloroplasts [5–7]. Group II chaperonins are further divided into archaeal types (thermosome) and eukaryotic types (TRiC) [8, 9]. Both chaperonin groups share a similar molecular architecture consisting of two back-to-back stacked rings, with a central cavity in each ring that allows substrates to fold. The structure of group I chaperonin GroEL/ES is well established [10–12] and the mitochondrial Hsp60/10 crystal structure has been recently solved . Both chaperonins are cylindrical structures consisting of 14 identical subunits assembled into two heptameric rings. The crystal structures of GroEL in its various forms (GroEL apo, GroEL/ADP, GroEL/ES/ADP), GroEL mutants, and mini-chaperones have facilitated the understanding of its functional mechanism. Even the structure of mini-chaperones, which were solved in high resolution, provided details of side chains which are involved in the interaction or regulation with other proteins or molecules [10, 11, 14–16]. Each of the 14 individual subunits of the chaperonins has three domains: the apical, the equatorial, and the intermediate hinge domains. In GroEL, the apical domain (amino acid (aa) residues 189–377) recognizes substrate proteins via hydrophobic residues exposed toward the central cavity and interacts with its co-chaperonin GroES [17, 18]. The equatorial domain (aa residues 2–136 and 410–525) contains the ATP binding site and contributes to inter-ring contacts . These two domains are connected by the intermediate hinge domain (aa residues 137–188 and 378–409), which allows rigid body movement.
In the past 20 years, the asymmetric mechanism of GroEL-assisted folding has been supported, i.e. GroES binds to one GroEL ring (Bullet) and the two rings function sequentially [20, 21]. However, the observation of a symmetric GroEL-GroES2 complex, in which both GroEL rings are capped with GroES (Football), leads to the suggestions that the symmetric complexes represent a folding intermediate and that the two GroEL rings fold polypeptides simultaneously [22–26]. Recently, Haldar et al.  found that the presence of symmetric GroEL-GroES2 complexes was largely dependent on the fluoro-fluorescence pair used to label the chaperonin system and non-foldable substrates. Whether a sequential or simultaneous folding mechanism is undertaken by chaperonin in vivo is still under investigation.
Chaperonins utilize ATP to drive a conformational cycle that allows them to capture, encapsulate, fold, and release substrate proteins [28, 29]. Binding and hydrolysis of ATP induce both positive (intra-ring) and negative cooperative (inter-ring) actions of GroEL . ATP hydrolysis also promotes the partial disassembly of chloroplast chaperonin into the monomer [31, 32], a phenomenon specific to chloroplast chaperonin. The chloroplast chaperonin was first identified as a Rubisco ‘large subunit binding protein’ [33, 34]. Unlike homo-oligomeric GroEL and mitochondrial Hsp60, chloroplast chaperonin consists of multiple subunits, diverging into two distinct but related α and β types [6, 35]. Cpn60α does not assemble into tetradecameric oligomers unless incorporated into oligomers of Cpn60β subunits [6, 31]. Though the authentic chloroplast chaperonin consists of both subunit types in vivo, homo-oligomeric Cpn60β is functional in refolding model substrates with assistance from co-chaperonin in vitro [6, 36, 37]. Two highly conserved CPN60β oligomers exhibit significantly different biochemical properties [38, 39], e.g. homo-oligomers of CPN60β2, but not CPN60β1, from Chlamydomonas completely disassemble into monomers upon ATP hydrolysis. Although new information about chloroplast chaperonin is steadily being published, its structure and complicated regulatory mechanism remain unknown.
Here, we report the first crystal structure of chloroplast homo-oligomer CPN60β1 in its apo form, with a larger molecular diameter than GroEL and novel ATP binding pockets. Domain swapping between CPN60α and CPN60β revealed that the equatorial domain mediates oligomer formation and the two subunits are highly cooperative in forming functional oligomers.
Homo-oligomeric CPN60β1 bound to co-chaperonins but could not assist folding of model substrate RrRubisco
Crystal structure of the CPN60β1 oligomers
An enlarged ATP binding pocket
The equatorial domain directs the oligomer formation
C-terminal (aa residues 484–547) of CPN60β influences oligomer disassembly driven by ATP
More importantly, insertion of the E2 domain from CPN60β1 into chimeras substantially stabilized oligomers in the presence of ATP (Fig. 5a,b, chimeras S and W). We further divided the E2 domain into four segments according to sequence homologues and constructed segment swapping chimeras S1–S5 (Fig. 5c). Insertion of C-terminal fragments containing at least amino acid residues 484–547 of CPN60β1 into chimeras significantly stabilized oligomers upon ATP hydrolysis (Fig. 5c, d, chimeras S2, S3, and S4), whereas insertion of the most divergent fragment (aa residues 524–547, not visible in the crystal structure, chimera S1) or a relatively conserved fragment (aa residues 484–524, chimera S5) into the chimera had no effect on oligomer stability. The C-terminus (484–547) might stabilize CPN60β1 oligomeric states by providing large hydrophobic fragments. Replacement of amino acids in the CPN60β2 E2 domain with the corresponding amino acids of CPN60β1 yielded three further mutants (ET431RR, S451C, and RVMD468EVMK). CPN60β2 (ET431RR) and CPN60β2 (S451C) did not influence oligomer formation and disassembly upon ATP hydrolysis; however, CPN60β2 (ET431RR) changed the electrophoresis migration rate during native PAGE (Additional file 1: Figure S5A). CPN60β2 (RVMD468EVMK) did not assemble into oligomers, although a substantial portion of the protein is soluble, a phenomena similar to that observed with CPN60β2 (Q467N) (Additional file 1: Figure S5A and S5B). Structural analysis revealed that N467 and E468 of CPN60β1 form van der Waals forces between rings to stabilize the oligomers (Additional file 1: Figure S5C). Point mutants of some amino acids involved in inter-subunit interaction showed CPN60β2 (S46A) and CPN60β2 (S180G) stabilized oligomers to some extent upon ATP hydrolysis (Additional file 1: Figure S5B), and double mutants showed no cumulative effects (Additional file 1: Figure S5D). When these amino acids in CPN60β1 were replaced with the corresponding amino acids of CPN60β2, no obvious effect was observed (Additional file 1: Figure S5E). The inherent differences between the equatorial domains of CPN60β1 and CPN60β2 confer their differential response to ATP.
Individual CPN60 subunits could not complement GroEL function in E. coli . Similarly, the chaperonin chimeras (H, L, M, N), with the equatorial domain from CPN60β subunits and the apical domain from the CPN60α subunit could not complement GroEL function (Additional file 1: Figure S6A,B). Coexpression of both CPN60β subunits could partially complement GroEL function after a long incubation time (Fig. 5e) . E. coli MGM100 coexpressing CPN60β1 with chimera P, which contains the entire equatorial domain of CPN60β2, grew better than with chimera S, which has the E2 domain of CPN60β1 (Fig. 5e). Conversely, coexpression of CPN60β2 with chimera S complements GroEL function better than with chimera P (Fig. 5e), suggesting the equatorial domains from CPN60β subunits compensate for each other.
CPN60α and CPN60β subunits are highly cooperative
The multi-subunit chloroplast chaperonin Cpn60 is more complex in composition relative to its prokaryotic homologue GroEL. Cpn60, consisting of both Cpn60α and Cpn60β types, is functional both in vivo and in vitro [6, 33, 51], whereas the homo-oligomeric Cpn60β type is only functional with selective co-chaperonins in vitro [36, 52]. Recombinantly purified Chlamydomonas CPN60β1 homo-oligomers exhibit biochemical properties similar to functional hetero-oligomers, e.g. possessing high ATPase activity and high protease sensitivity, being binding model substrates and interacting with co-chaperonins . However, CPN60β1 homo-oligomers were not fully functional in refolding model substrates  (Fig. 1), which might result from a failure of allosteric movement during the functional cycle (Fig. 5). Nonetheless, due to the described functional similarity, the structure of CPN60β1 probably resembles the structure of chaperonin hetero-oligomers in vivo and is not a result of a crystallization artefact of a non-functional and, therefore, irrelevant protein. Indeed, the loosely-compacted crystal structure of CPN60β1 was consistent with the observation that Cpn60 oligomers are highly susceptible to proteolytic degradation. It is not clear why the chloroplast chaperonin might adopt a loosely compacted structure. Considering the plastid-specific co-chaperonin Cpn20 that consists of a double GroES-like domain, it is plausible that a loosely-compacted structure has been optimized to interact with Cpn20. Admittedly, the presented CPN60β1 structure may also deviate significantly from the active Cpn60 structure in vivo; however, it represents a first step toward structural and functional understanding of chloroplast chaperonin and will likely provide insights that can be exploited by future structural studies.
An enlarged nano-cage for substrate folding in CPN60
The folding nano-cage present in the crystal structure of GroEL/GroES is about 175,000 Å3 and could theoretically accommodate substrate proteins of up to 70 kDa . In reality, the functional volume of this nano-cage is smaller, probably because the 23 C-terminal amino acids not resolved in the GroEL/GroES structure protrude into the cage and occupy space. Most GroEL substrates are actually smaller than 50 kDa . In the crystal structure of CPN60, the diameter of the central cavity is about 6 Å longer than the diameter of the GroEL cage, but the largest Cpn60 substrate identified to date is the Rubisco large subunit, of only around 50 kDa in size [33, 54, 55]. Tang et al.  reported that a GroEL cage volume increased by 2–5 % remarkably decelerated the folding of large substrates, e.g. MBP (41 kDa) and RrRubisco (50 kDa), by more than 2-fold. Consistent with their results, we found that the increased size of the Cpn60 cavity corresponded to a slower rate of folding of RrRubisco relative to GroEL . Chaperonin folding of endogenous CrRubisco was not investigated in this study due to the inherent difficulty associated with the biogenesis of green type Rubisco in E. coli.
The decisive role of Cpn60α is supported by its unique equatorial domain
It is reported that the Cpn60α deletion is lethal to plants [57–59]. Here, we found that CPN60α is highly cooperative with CPN60β1 to form functional oligomers potentially by regulating the allosteric movement of CPN60 oligomers (Figs. 6 and 7). Replacement of the CPN60α apical domain by the same domain of CPN60β1 did not influence subunit cooperation and oligomer functionality, suggesting that the apical domain of CPN60α does not play an essential role (Fig. 6b). The equatorial domain of Cpn60β type, but not Cpn60α, directs the subunits to form oligomers, suggesting that the Cpn60β subunits initiate oligomer assembly  (Fig. 4). Homo-oligomeric Cpn60β type possesses high ATPase activity, substrate binding ability and interaction with co-chaperonins, and it is partially functional with selective co-chaperonin. The equatorial domain and at least one intermediate segment from CPN60α are indispensable to the functionality of oligomers (Fig. 4), suggesting that fragments from Cpn60α offer Cpn60β subunit flexibility during its allosteric movement. Unlike its GroEL homologue, Cpn60 exhibits sophisticated regulation between its subunits.
In our study, the first crystal structure of Chlamydomonas chloroplast chaperonin homo-oligomer (CPN60β1) was solved at 3.8 Å, and displayed a rigid body and structural features such as looser compaction, a larger central cavity, less contact sites, and an enlarged ATP binding pocket compared to GroEL. Interestingly, we found that the overall structure of Cpn60 resembles the GroEL allosteric intermediate state. There are two amino acids (153 and 154) conserved among Cpn60s which were involved in regulation of ATPase activity by co-chaperonins. Our domain swapping analysis revealed that the equatorial domain of CPN60α maintains its monomeric state in the absence of CPN60β (CPN60β1 or CPN60β2). This equatorial domain of CPN60α was also indispensable for functional cooperation with CPN60β1. We additionally discovered that the C-terminal fragment (aa residues 484–547) of CPN60β type influenced oligomer disassembly and allosteric rearrangement driven by ATP hydrolysis. More interestingly, two CPN60β subunits compensated for each other to rescue E. coli growth partially. Our results shed light on chloroplast chaperonin structure and cooperation between CPN60 subunits.
Plasmids cloning and protein purifications
Plasmid cloning for expression of the mitochondrial Hsp10, chaperonin chimeras, and chaperonin mutants, as well as purification of proteins are described in Additional file 1: Material and Methods.
Crystallizations and X-ray data collection and processing
Crystallizations of CPN60β1 oligomers and X-ray data collection and processing are described in Additional file 1: Material and methods.
E. coli complementation
Analysis to determine whether chaperonin chimeras complement GroEL function was performed as described previously . All chimeric or point-mutated forms of chaperonin were constructed into pOFX vector , and the resulting plasmids were transformed into E. coli MGM100 cells by electroporation (MicroPulser™, Bio-Rad). A single transformant colony was picked and grown in LB medium supplemented with 30 μg/mL kanamycin, 20 μg/mL spectinomycin and 0.02 % arabinose to an OD600 of approximately 1; 1 mL of cells was collected, carefully washed five times and resuspended in 1 mL LB. Ten-fold dilution series were made, and 7.5 μL of diluted cells were spotted onto LB agar plates supplemented with 0.2 % glucose/1 mM IPTG, 30 μg/mL kanamycin and 20 μg/mL spectinomycin. The cells were grown at 37 °C for 12–13 hours.
The ATPase activity of chaperonin was measured using a coupled enzymatic assay as described previously . ATP hydrolysis by chaperonin is linked to the oxidation of NADH to NAD+ by the coupled reaction of Pyruvate kinase and L-lactate dehydrogenase. After sequential addition of 1 mM phosphoenolpyruvate, 20 U/mL pyruvate kinase, 30 U/mL lactate dehydrogenase, and 0.5 mM NADH into the reaction buffer (50 mM HEPES/KOH pH 7.5, 10 mM KCl, 10 mM MgCl2), 1 mM ATP and/or 0.5 μM co-chaperonin was added and incubated for 3 min at 25 °C. The absorbance at 340 nm was monitored immediately after addition of 0.2 μM chaperonin for 10 min.
Analysis of chaperonin chimera and co-chaperonin interaction was performed with analytical gel filtration as described previously . Briefly, 2 μM CPN60β1 chaperonin and 10 μM co-chaperonin were incubated in buffer containing 20 mM MOPS-KOH, pH 7.5, 5 mM Mg(OAc)2, 100 mM KCl, and 2 mM ADP-AlF3 for 30 minutes at 25 °C, and the protein complexes were then separated with a Superdex 200 PC3.2/10 column (GE Healthcare) with the same buffer but using only 50 μM ADP-AlF3 at a flow rate 0.05 mL/min. Finally, 60-μL fractions were collected, resolved by 15 % SDS-PAGE and visualized with Coomassie staining.
Transmission electron microscopy
ATP (2 mM) was added to the incubation buffer (30 mM KF, 20 mM Tris-HCl pH 7.5, 10 mM MgCl2, 3 mM KAl(SO4)2). The final concentrations of chaperonin (CPN60β1) and co-chaperonin (GroES) were 0.1 μM and 0.5 μM, respectively. The reaction was incubated at 4 °C for 30 min, and the solution was subjected to ultrafiltration to remove chemical precipitates. Then, 4 μL of the supernatant was placed onto the carbon-coated grids, which had been previously hydrophilized by glow discharge; 1 min later, excess protein sample was removed by filter paper. The specimen was stained with 1 % uranyl acetate for 1 min and allowed to air dry. Images were recorded by a CCD camera system (Olympus, Tokyo, Japan) at an accelerating voltage of 120 keV on a transmission electron microscope (FEI, Eindhoven, The Netherlands).
Protein refolding and Rubisco carboxylation assay
Availability of data and materials
Coordinates and structure factor amplitudes for CPN60β1 oligomers are deposited in the Protein Data Bank under accession codes 5CDI.
Full Methods and any associated references are available in the Additional files.
We thank Dr. Andreas Bracher and Prof. Yuhang Chen for their suggestions on the crystal structures. We would like to thank Dr. Sarah Schäuffelhut for her critical reading of the manuscript. The crystal data were collected at Shanghai Synchrotron Radiation Facility (SSRF) beamline 17U. This work was supported by the State Key Laboratory of Plant Cell and Chromosome Engineering (grant no: PCCE-2012-TD-01). We are grateful to the National Natural Science Foundation of China (31170686) and the National Program on Key Basic Research Project (2011CB911102) for financial support.
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