©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Spinach Chloroplast cpn21 Co-chaperonin Possesses Two Functional Domains Fused Together in a Toroidal Structure and Exhibits Nucleotide-dependent Binding to Plastid Chaperonin 60 (*)

Franois Baneyx (1)(§), Uwe Bertsch (2)(¶), Cathy E. Kalbach (1), Saskia M. van der Vies (3), Jürgen Soll (2), Anthony A. Gatenby (1)(**)

From the (1) Molecular Biology Division, Central Research and Development, DuPont, Experimental Station, Wilmington, Delaware 19880-0328, the (2) Botanisches Institut, Olshausenstrasse 40, D-24098 Kiel, Germany, and the (3) Départment de Biochimie Médicale, Université de Genève, 1211 Genève 4, Switzerland

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS

ABSTRACT

Chloroplasts contain a 21-kDa co-chaperonin polypeptide (cpn21) formed by two GroES-like domains fused together in tandem. Expression of a double-domain spinach cpn21 in Escherichia coli groES mutant strains supports growth of bacteriophages and T5, and will also suppress a temperature-sensitive growth phenotype of a groES619 strain. Each domain of cpn21 expressed separately can function independently to support bacteriophage growth, and the N-terminal domain will additionally suppress the temperature-sensitive growth phenotype. These results indicate that chloroplast cpn21 has two functional domains, either of which can interact with GroEL in vivo to facilitate bacteriophage morphogenesis. Purified spinach cpn21 has a ring-like toroidal structure and forms a stable complex with E. coli GroEL in the presence of ADP and is functionally interchangeable with bacterial GroES in the chaperonin-facilitated refolding of denatured ribulose-1,5-bisphosphate carboxylase. Cpn21 also inhibits the ATPase activity of GroEL. Cpn21 binds with similar efficiency to both the and subunits of spinach cpn60 in the presence of adenine nucleotides, with ATP being more effective than ADP. The tandemly fused domains of cpn21 evolved early and are present in a wide range of photosynthetic eukaryotes examined, indicating a high degree of conservation of this structure in chloroplasts.


INTRODUCTION

Chloroplasts are structurally and biochemically complex organelles that satisfy the energy requirements of plant cells. Although the majority of chloroplast proteins are synthesized in the cytosol and translocated into plastids a significant number, most notably the large subunits (L)() of ribulose-1,5-bisphosphate carboxylase (Rbu-P carboxylase), are produced in the stroma. Rbu-P carboxylase L subunit polypeptides are the major products of in organello protein synthesis and assemble with imported small subunits (S) into a holoenzyme with an LS composition. During studies on the biogenesis of Rbu-P carboxylase in isolated chloroplasts, it was realized that assembly of nascent L subunit chains into LS holoenzyme does not occur rapidly and spontaneously, but involves an intermediary protein (1, 2) . This intermediary protein, later identified as chaperonin 60 (cpn60) (3) , binds to L subunits prior to their assembly into holoenzyme (1, 2, 4) . Release of L subunits from cpn60 and their subsequent oligomerization is stimulated by MgATP (5) . The observations that cpn60 also binds to imported Rbu-P carboxylase S subunits (4, 6) , and several other imported proteins (4, 7) , indicates that this molecular chaperone has a more general role in plastid development and may influence the folding of numerous chloroplast proteins.

Escherichia coli GroEL is a homologue of chloroplast cpn60 (3) and also shares in common the ability to interact with many target polypeptides to influence their folding into functional proteins (8, 9, 10, 11, 12, 13, 14, 15, 16) . From these studies it has become apparent that chaperonins regulate protein folding by stabilizing folding intermediates, thereby influencing the kinetic partitioning between aggregated (misfolded) and correctly folded proteins. The release of target proteins bound to GroEL and their subsequent progression to the native state occurs through interactions with the GroES co-chaperonin and MgATP. Even for examples where GroES is not essential for release its presence usually potentiates the discharge reaction (11, 12, 13) . The requirement for GroES in effective dissociation of GroEL-target protein complexes suggests that chloroplasts might also contain a GroES homologue to facilitate protein folding by interacting with plastid cpn60 (17) .

GroES homologues are present in mitochondria (17, 18, 19, 20) , and a functionally related protein is present in chloroplasts (21) and also encoded in a bacteriophage genome (22) . Interestingly, the chloroplast co-chaperonin is a polypeptide of about 21 kDa (cpn21), which is twice the size of bacterial and mitochondrial cpn10 and thus different from any co-chaperonin described before. Chloroplast cpn21 is comprised of two distinct cpn10-like domains fused in tandem to give a binary co-chaperonin structure (21) . Each of the two fused domains possesses several highly conserved amino acid residues that are encoded in many groES genes, suggesting that each domain could be functional. To determine the biological activity of the double-domain co-chaperonin, and each of its two separate single domains, we have constructed expression plasmids for cpn21, the N-terminal (cpn10-N) and the C-terminal (cpn10-C) domains and synthesized them in E. coli. By expressing cpn21, cpn10-N, and cpn10-C chloroplast co-chaperonins in a groES mutant strain that fails to support bacteriophage head assembly, we show that cpn21 and each separate domain are functional in vivo and facilitate bacteriophage morphogenesis. Cpn21 and cpn10-N, but not cpn10-C, also suppress the growth defect of a temperature-sensitive groES mutant strain. The functional demonstration of cpn21 activity in cells is supported by in vitro data indicating that either E. coli GroEL or chloroplast cpn60 will form a complex with spinach cpn21 co-chaperonin in the presence of adenine nucleotides. In addition, purified plastid cpn21 co-chaperonin inhibits the ATPase activity of GroEL and can effectively substitute for bacterial GroES in the chaperonin-facilitated refolding of denatured Rbu-P carboxylase. Electron micrographs of spinach cpn21 reveal a ring-like organization that is similar to E. coli GroES (23) .


EXPERIMENTAL PROCEDURES

Plasmid Construction

PCR amplification of pSOCPN70/8.1 (21) sequences was with GeneAmp (Cetus). The numbered black bars in Fig. 1 A represent oligo-nucleotide primers used for PCR and plasmid construction. Primers 1 (5`-GGTTTGGTTGTTCCCATGGCCTCAATAACCACA-3`) and 3 (5`-GAAGGAAGATGATACCATGGGTATCCTAGAAAC-3`) were used to introduce NcoI sites and methionine codons in front of the first amino acid of either the mature protein or the N-terminal domain (primer 1), or at amino acid 152 at the start of the C-terminal domain (primer 3). The initiator methionine for each translated polypeptide was designed to be within the NcoI recognition sequence (5`-CCATGG-3`). Primer 2 (5`-CAGTTTCAAGCTTACCAATTATATCACTCTTCCT-3`) introduces a stop codon at the end of the N-terminal domain followed by a HindIII site. Primer 4 (5`-TGGGCTACAAATCTAAAGCTTCAACTGGCA-3`) introduces a HindIII site in the 3`-untranslated region of the cDNA. Combinations of primers 1 and 4 were used to synthesize the cpn21 gene for plasmid pCK14, primers 1 and 2 the cpn10-N domain (pCK16) and primers 3 and 4 the cpn10-C domain (pCK19). PCR products were cut with NcoI and HindIII and ligated into the same sites in vector pKK233-2 (Pharmacia LKB Biotechnol). The complete nucleotide sequence of inserts was confirmed for all clones. Derivatives of each clone were made by transferring the NcoI/ HindIII fragments to vector pTrc99A (Pharmacia) to ensure tighter control of expression from a plasmid-encoded lacIq repressor, which was found to be essential for transformation into the minicell strain. These plasmids are pCK25 (cpn21 from pCK14), pCK26 (cpn10-N from pCK16), and pCK27 (cpn10-C from pCK19). To add histidine residues to the N terminus of cpn21, the NcoI site of pCK14 was repaired with Klenow, the gene removed with HindIII, followed by transfer to the BamHI (Klenow repaired) and HindIII sites of vector pQE30 (Qiagen) to give plasmid pCK28.


Figure 1: Plasmid construction and expression of spinach chloroplast co-chaperonins. A, the cDNA of plasmid pSOCPN70/8.1 (21) encodes the 26.8-kDa precursor of chloroplast cpn21. The transit peptide ( black), two cpn10-like domains (about 100 amino acids each, white or gray) and a linker region ( diagonal lines) are shown. Numbered black bars represent oligo-nucleotides used for PCR and plasmid construction and their hybridization sites. Plasmid pCK14 encodes full-length cpn21, pCK16 the N-terminal domain (cpn10-N), and pCK19 the C-terminal domain (cpn10-C). Expression of the pCK series is directed from a trc promoter and a lacZ ribosome-binding site in vector pKK233-2. B, expression of proteins in E. coli. Extracts from induced JM105 cells harboring pKK233-2 ( lane 1), pCK14 ( lane 2), pCK16 ( lane 3), or pCK19 ( lane 4) were resolved by SDS-PAGE (15%) and stained with Coomassie Brilliant Blue. The black arrow shows the position of cpn21 and the open arrow the position of the cpn10-N domain. Molecular weight markers ( M) are 43, 29, 18.4, 14.3, and 6.2 kDa. C, fluorograph of H-labeled cpn21 ( lane 1), cpn10-N ( lane 2), and cpn10-C ( lane 3) following synthesis in minicells, immunoprecipitation with anti- cpn21, and SDS-PAGE on 12.5% gels. The open arrows adjacent to lanes 1 and 3 mark the positions, respectively, of full-length cpn21 or cpn10. D, co-chaperonin stability in E. coli for GroES (), cpn21 (), cpn10-N (), and cpn10-C (). Proteins were labeled in minicells with [H]leucine for 5 min, chased with excess cold leucine, and sampled at various times. Radioactivity in each sample was determined as described under ``Experimental Procedures.''



Expression of Co-chaperonins

Plasmid-bearing JM105 cells were grown at 37 °C in LB medium with 0.2% glucose and 50 µg/ml ampicillin to OD = 0.5. IPTG (1 mM) induction was for 1 h, followed by passage through a French Press at 20,000 pounds/square/inch in 50 mM Tris-HCl, pH 7.7, 10 mM KCl, 10 mM MgCl, 1 mM dithiothreitol, 0.01% Tween 20 (Buffer I). To detect expression, centrifuged extracts were analyzed by SDS-PAGE and stained (Fig. 1 B). Plasmid-encoded protein synthesis was also detected in isolated minicells (24) transformed with pCK25 (cpn21), pCK26 (cpn10-N), pCK27 (cpn10-C), or pGroES following induction with IPTG and labeling with [H]leucine (DuPont NEN) (Fig. 1 C). For pulse-chase analysis minicells were labeled for 5 min, mixed with LB containing 50 µg/ml leucine, and aliquots added to acetone at various times. Samples were analyzed by SDS-PAGE (12.5%), bands excised, solubilized (Solvable, DuPont NEN), and quantitated by liquid scintillation counting. Relative synthesis rates during a 5-min labeling were determined by comparing the radioactivity incorporated into the co-chaperonin with that of plasmid-encoded -lactamase as a standard. Radioactivity was normalized to the number of leucine residues in each protein, using 31 residues for -lactamase, 18 for cpn21, 5 or cpn10-N, and 13 for cpn10-C.

Formation and Size Exclusion Chromatography of Complexes between GroEL and Co-chaperonins

Five-ml cultures of CG712(pGroES) or CG712(pCK14) were grown to early log phase in M9 minimal media, induced with 1 mM IPTG, and labeled for several generations with 100 µCi of [H]leucine (143.7 Ci/mmol, DuPont NEN). Pelleted cells were incubated on ice for 30 min in 25 µl of 50 mM Tris-HCl, pH 7.7, 5 mM EDTA, 20% sucrose, 200 µg/ml lysozyme. The cell suspension was vortexed with 100 µl of Buffer I containing 5% glycerol, subjected to three cycles of freeze-thawing, and centrifuged at 14,000 g for 15 min. The supernatant (100 µl) was passed through a Sephadex G-50 Nick column (Pharmacia) equilibrated with Buffer I containing 0.5 mM dithiothreitol and proteins in the void volume collected. Complex formation was determined by size exclusion chromatography at 23 °C using a fast protein liquid chromatography TSK column (type G3000SW, 7.5 600 mm, Pharmacia) at a flow rate of 1 ml/min. To examine the total labeled protein profile in cell extracts, 200-µl aliquots were injected onto the TSK sizing column equilibrated with Buffer I containing 0.5 mM dithiothreitol, 0.5 ml fractions collected, and the radioactivity in each fraction determined. To form a complex between GroEL and either GroES (Fig. 2 A) or cpn21 (Fig. 2 B), 200 µl of cell extracts were supplemented with 0.63 µM GroEL (14-mer) and 1 mM ADP. After a 15-min incubation at 23 °C, the sample was analyzed on the TSK sizing column as above, except that the column buffer was supplemented with 0.25 mM ADP. For each column analysis, the fractions eluting at 11-11.5 ml were combined, 100-µl aliquots precipitated with acetone, and analyzed by SDS-PAGE (15%). The radiolabeled E. coli or spinach co-chaperonins were detected by fluorography using Enhance (DuPont NEN).


Figure 2: Formation of a stable complex between GroEL and either GroES ( panel A), or spinach cpn21 ( panel B) in the presence of ADP. Strain CG712 harboring either pGroES ( panel A) or pCK14 ( panel B) were induced with IPTG and labeled for several generations with [H]leucine. Soluble cell extracts were supplemented with GroEL and incubated in either the absence () or presence () of 1 mM ADP. Samples were injected onto a TSK sizing column and the elution profile of radiolabeled proteins determined. Peak fractions containing GroEL (11-11.5 ml) were combined, acetone precipitated, and analyzed by SDS-PAGE (15%). In the fluorographs, lanes 1 (pGroES) and 3 (pCK14) contain samples prepared in the absence of ADP, and lanes 2 (pGroES) and 4 (pCK14) contain samples isolated following the addition of ADP.



Binding of GroEL and Chloroplast cpn60 to Immobilized cpn21

Strain JM105 harboring pCK28 and the lacIq repressor plasmid pREP4 (Qiagen) were grown in LB with ampicillin and kanamycin to OD = 0.5, induced with 1 mM IPTG for 2 h, harvested, and frozen. Thawed cells were broken at 20,000 pounds/square/inch in a French Press in 0.1 M Tris-HCl, pH 7.5, 0.3 M NaCl, 5 mM -mercaptoethanol (Buffer II) with 1 mM phenylmethylsulfonyl fluoride, and centrifuged at 40,000 g for 30 min. To prepare His-cpn21 resin, the supernatant was applied to 2-ml columns of Ni-NTA (Qiagen) in Buffer II, washed with Buffer II followed by 50 mM MES-NaOH, pH 6.5, 0.5 M NaCl, 10% glycerol, 5 mM -mercaptoethanol, then 25 mM imidazole, 50 mM MES-NaOH, pH 6.5, 0.3 M NaCl, 5 mM -mercaptoethanol, and finally 0.1 M Tris-HCl, pH 7.5, 10 mM KCl, 10 mM MgCl (Buffer III) containing 5 mM -mercaptoethanol. Spinach chloroplasts were isolated from 100 g of leaves (4) and lysed by incubation on ice for 15 min in 5 ml of Buffer III containing 1 mM phenylmethylsulfonyl fluoride and 10 µg/ml each of leupeptin, chymostatin, and soybean trypsin inhibitor. After centrifuging at 40,000 g for 15 min, 500-µl aliquots of the soluble protein fraction were either incubated with 10 units of apyrase for 10 min at 23 °C or supplemented with 1 mM ADP or ATP. These samples were batch incubated with 200 µl of His-cpn21 resin for 15 min at 23 °C and washed with 6 1 ml of Buffer III ± 1 mM ADP or ATP. Bound proteins were eluted with 0.3 M imidazole in Buffer III, precipitated in 5% trichloroacetic acid, washed with 80% acetone, and analyzed by SDS-PAGE (15%). To detect binding of E. coli, GroEL-purified protein was passed through a His-cpn21 column in Buffer III ± 1 mM ADP, washed, and eluted as for the chloroplast protein.

Spinach cpn21 Co-chaperonin Purification from E. coli

Spinach cpn21 was purified from JM105(pCK14) cells grown, induced, and disrupted in a French Press as described above. The crude extract was fractionated by size exclusion chromatography on a TSK G3000SW column developed at 1 ml/min in Buffer I using fast proteim liquid chromatography. Fractions corresponding to a molecular mass range of 250 to 160 kDa were pooled and exchanged into Buffer IV (20 mM MES-NaOH, pH 6.0, 1 mM dithiothreitol, 0.01% Tween 20) using Sephadex G-25 PD10 columns (Pharmacia). Proteins were loaded onto a MonoS HR 5/5 column (Pharmacia) and eluted at 1 ml/min using a 0-1 M NaCl gradient in Buffer IV. Fractions containing cpn21 (eluting at about 200 mM NaCl) were pooled and the pH raised to 6.5 with NaOH. Proteins were loaded onto the same MonoS column equilibrated in Buffer V (20 mM MES-NaOH, pH 6.5, 1 mM dithiothreitol, and 0.01% Tween 20) and eluted using a 0-1 M NaCl gradient in Buffer V. Fractions eluting at about 185 mM NaCl were supplemented with 10% glycerol and frozen at -80 °C.

Rbu-P Carboxylase Refolding Assays

Rbu-P carboxylase refolding experiments were carried out as before (10) . Reactions were initiated with 3 mM ATP, stopped with a hexokinase/glucose trap, and Rbu-P carboxylase assayed. Control reactions containing an equal amount of native Rbu-P carboxylase were prepared to establish the value associated with 100% reconstitution.


RESULTS

Synthesis and Stability of Chloroplast cpn21 and Each Separate cpn10 Domain in E. coli

A spinach cDNA clone (21) encoding the chloroplast binary co-chaperonin was used to construct a series of plasmids directing the synthesis of the complete cpn21 protein (pCK14), the cpn10-N domain (pCK16), and the cpn10-C domain (pCK19) in E. coli (Fig. 1 A). Production of the pCK14-encoded cpn21 protein and the pCK16-encoded cpn10-N domain was observed in extracts of IPTG-induced cells resolved using SDS-PAGE and stained with Coomassie Brilliant Blue (Fig. 1 B, lanes 2 and 3). Synthesis of the cpn10-C domain encoded by pCK19 was not readily detectable at steady state levels in cell extracts ( lane 4). This failure to observe the presence of cpn10-C in stained gels was due to protein instability and rapid turnover in vivo since the protein could be detected in radiolabeled E. coli minicells, although a degradation product was also apparent (Fig. 1 C, lane 3). The production in minicells of GroES, cpn21, cpn10-N, and cpn10-C allowed half-life and relative synthesis rates to be measured. Compared to a half-life of several h for GroES and about 120 min each for cpn21 and cpn10-N, that of cpn10-C was 17 min (Fig. 1 D). Relative synthesis rates of cpn21, cpn10-N, and cpn10-C were 10, 25, and 17 times, respectively, greater than plasmid-encoded -lactamase during a 5-min incubation (data not shown). Suppression of groES Mutations by cpn21 and Each cpn10 Domain

To determine if the chloroplast binary or single-domain co-chaperonin proteins were functional in E. coli and could substitute for bacterial GroES, we used two groES-defective strains, CG712 ( groES30) (15) and JZ12 ( groES619) (25) . Both the groES30 and groES619 mutations prevent bacteriophage head assembly, and bacteriophage T5 tail assembly is blocked by the groES619 mutation (26, 27, 28) . Plasmids pCK14, pCK16, pCK19, pKK233-2 (vector control), and pGroES ( E. coli groES control, 16) were independently transformed into CG712 ( groES30) and tested for their ability to support bacteriophage growth (). The plating efficiency and plaque size of on CG712 expressing full-length binary chloroplast cpn21 co-chaperonin (pCK14) were similar to that of the same strain harboring plasmid pGroES (wt groES). The cpn10-N (pCK16) or cpn10-C (pCK19) domains of the chloroplast protein expressed separately also supported growth (). Plasmid pCK16 was more effective than pCK19 as judged by its higher plating efficiency and larger plaque size, but both were considerably more efficient than the vector control (pKK233-2). The observed efficiency of plating for using pCK16 and pCK19 were comparable to the suppression levels obtained with the T4-encoded Gp31 functional analogue of GroES (22) . The reduced ability of the C-terminal domain to support growth, compared to both the binary cpn21 co-chaperonin and the cpn10-N domain, may be related to differences in stability in vivo (Fig. 1 D). This is supported by the observation that the efficiency of plating of was improved when expression levels of cpn10-C (pCK19) were increased by the addition of 0.2 mM IPTG to the top agar (data not shown).

To examine the influence of chloroplast co-chaperonins on bacteriophage T5 morphogenesis, strain JZ12 ( groES619) was transformed with the above plasmids and assayed for bacteriophage growth (). The plating efficiency of T5 on strain JZ12 harboring either pGroES or pCK14 were comparable, which suggested that the chloroplast cpn21 protein could support T5 tail assembly. Interestingly, the cpn10-N and cpn10-C domains of the spinach co-chaperonin were unable to support T5 tail assembly () in strain JZ12, although they permitted head assembly in the groES30 strain CG712. Another characteristic of the groES619 mutation is that it prevents colony formation at 43 °C. This phenotype can be suppressed by plasmid-encoded GroES (22) , and as we show here, by chloroplast cpn21 co-chaperonin encoded by pCK14 (). In contrast to the T5 growth data obtained with this strain, the cpn10-N domain was functional in suppressing the temperature-sensitive phenotype of groES619, although suppression with cpn10-C was not observed (). These differential effects of the complete binary or separated domains of cpn21 on bacteriophage assembly or cell growth in different genetic backgrounds suggest that the discharge reaction for proteins bound to GroEL may be sensitive to variations in the structure, or the cellular concentrations of the interacting co-chaperonin molecule. Similar observations have been made regarding the cellular levels of GroES and GroEL mutant proteins on bacteriophage or cell growth (25, 28) . Interactions between cpn21 and Chloroplast cpn60 or Bacterial GroEL

Binnary Complex Formation

It has been established that in the presence of ATP or ADP a stable complex is formed between GroEL and GroES (23, 29) . Co-chaperonins from mitochondria, pea chloroplasts, and bacteriophage T4 will also form a complex with E. coli GroEL in the presence of adenine nucleotides (17, 19, 20, 21, 22) , indicating considerable functional conservation in the structure of the interactive chaperonin-co-chaperonin surfaces.

To detect if similar interactions occurred between GroEL and the recombinant spinach chloroplast cpn21 co-chaperonin synthesized in E. coli, the binding experiment illustrated in Fig. 2 A was carried out. Strain CG712 ( groES30) harboring plasmids pGroES or pCK14 was induced with IPTG and labeled for several generations with [H]leucine. Extracted soluble proteins were incubated with GroEL in the presence or absence of ADP and fractionated on a TSK sizing column. On this column GroEL elutes between 10.5 and 11.5 ml. In the absence of ADP GroES (Fig. 2 A, open circles) and cpn21 (Fig. 2 B, open circles) did not form a stable complex with GroEL, and neither co-chaperonin was present in the GroEL fraction analyzed by SDS-PAGE (Fig. 2, lanes 1 and 3). In the presence of ADP, however, both GroES and cpn21 formed stable binary complexes with GroEL that were detectable by an increase in the [H]leucine present in the GroEL fraction (10.5-11.5 ml, Fig. 2, A and B, filled circles), and by the identification using SDS-PAGE of GroES (Fig. 2, lane 2) or cpn21 (Fig. 2, lane 4) in the GroEL fraction. A complex between chaperonin and co-chaperonin was not observed in the absence of GroEL (data not shown).

These data show the nucleotide-dependent formation of a complex between spinach cpn21 and E. coli GroEL, but an interaction between the cpn21 co-chaperonin and cpn60 chaperonin from chloroplasts would support the respective roles of these latter two proteins in plastid protein folding. To demonstrate this, 6 histidine residues were fused to the N terminus of cpn21 in plasmid pCK28 and synthesis of the protein induced (Fig. 3, lanes 1-3). This modification does not impair the activity of cpn21 in vivo or in vitro as determined respectively by bacteriophage plating or Rbu-P carboxylase refolding (data not shown). The histidine variant of cpn21 (His-cpn21) was immobilized to Ni-NTA resin and used as an affinity ligand to bind cpn60. The His-cpn21 resin was first tested using purified GroEL from E. coli which bound to a column of His-cpn21 only in the presence of 1 mM ADP (Fig. 3, lane 5), but not if ADP was omitted ( lane 4). In addition, the impaired ability of GroEL140 mutant protein to bind GroES (10) was also observed by the failure of purified GroEL140 to efficiently bind to the His-cpn21 resin in the presence of adenine nucleotides (data not shown). Next, chloroplast stromal extracts were incubated with His-cpn21 resin in the presence of 1 mM ATP (Fig. 3, lane 7), 1 mM ADP ( lane 8), or in the absence of either nucleotide ( lane 9). The bound proteins were released with imidazole and fractionated on an SDS-polyacrylamide gel designed to separate the and subunits of plastid cpn60 (30) . Both the and subunits of spinach cpn60 bound to His-cpn21 in the presence of ADP or ATP, with the latter causing more effective binding. Binding was not observed with stromal extracts first incubated with apyrase ( lane 9). The intensity of the and bands bound to His-cpn21 ( lanes 7 and 8) was similar to that found in crude stromal extracts ( lane 6), indicating that a preferential interaction of either cpn60 subunit with the cpn21 co-chaperonin does not occur.


Figure 3: Nucleotide-dependent binding of GroEL or chloroplast cpn60 to immobilized cpn21. Plasmid pCK28 encodes an N-terminal fusion between 6 histidine residues and cpn21. The His-cpn21 protein was synthesized in E. coli, bound to Ni-NTA columns, and washed free of contaminating proteins. Purified GroEL or soluble chloroplast proteins were incubated with the His-cpn21 resin ± adenine nucleotide, washed, eluted with imidazole, and analyzed by SDS-PAGE. Lane 1, JM105(pCK28) total preinduced proteins; lane 2, total induced JM105(pCK28) proteins; lane 3, His-cpn21 bound and eluted from Ni-NTA column (the arrow for lane 3 marks the position of His-cpn21); lanes 4 and 5 show protein eluted from column following passage of GroEL-ADP ( lane 4) and + 1 mM ADP ( lane 5) (the arrow for lane 5 marks the position of GroEL and the star the position of His-cpn21); lanes 6-9 show chloroplast extracts where 6 is the column preload, sample 7 is cpn60 bound to His-cpn21 in presence of 1 mM ATP, sample 8 is cpn60 bound in the presence of 1 mM ADP, and sample 9 is pretreated with apyrase before column loading in the absence of either ADP or ATP. The two bands marked by arrows in lanes 6-9 are the (top) and (bottom) of spinach cpn60. Lanes 1-5 were stained with Coomassie Brilliant Blue, and lanes 6-9 immunoblotted, reacted with rabbit anti-cpn60 and goat anti-rabbit IgG conjugated to alkaline phosphatase.



Chaperonin-facilitated Protein Folding

Although suppression experiments indicated that the chloroplast binary co-chaperonin could substitute for GroES to support bacteriophage growth, it is difficult to compare the relative efficiencies of the two proteins in the discharge reaction because of differences in expression levels. To compare these efficiencies, the pCK14-encoded cpn21 protein was purified from an over-expressing strain (Fig. 4) and used in several biochemical assays (Fig. 5). Both spinach co-chaperonin and E. coli GroES exhibited similar saturation profiles with respect to Rbu-P carboxylase discharge from GroEL and subsequent folding (Fig. 5 A). In addition, Rbu-P carboxylase refolding was achieved with comparable initial rates and final yield for both the chloroplast and bacterial co-chaperonins (Fig. 5 B). Interestingly, purified cpn10-N and cpn10-C were not functional in the chaperonin-facilitated Rbu-P carboxylase folding assay, even though they are functional in vivo in supporting bacteriophage assembly. Absence of activity in vitro may be related to the reduced stability of cpn10-N or cpn10-C oligomers, as judged by size exclusion chromatography (data not shown), and implies that in vivo conditions are more favorable for the single-domain oligomer assembly or stability.


Figure 4: Purification of the spinach cpn21 co-chaperonin from induced JM105 harboring pCK14 cells in a three-step process. Lane M, molecular weight markers; lane 1, total soluble proteins; lane 2, 160-250-kDa pool following size exclusion chromatography on a TSK G3000SW column; lane 3, pool eluting at 200 mM NaCl on a MonoS column developed at pH 6.0; lane 4, pool eluting at 185 mM NaCl on a MonoS column developed at pH 6.5. The position of cpn21 is indicated by an arrow.




Figure 5: Comparison of the biochemical properties of spinach cpn21 and E. coli GroES. A, cpn21 activity is saturable with respect to GroEL. The refolding of acid-denatured R. rubrum Rbu-P carboxylase (90 nM protomer) loaded onto GroEL (1.5 µM protomer) was measured in 285 µl after 1 h of incubation at room temperature and quenched with a hexokinase/glucose trap using increasing concentrations (expressed as protomers) of GroES () or cpn21 (). B, Rbu-P carboxylase refolding is achieved with comparable initial rates and final yield by cpn21 or GroES. The refolding of acid-denatured Rbu-P2 carboxylase loaded onto GroEL was performed as in panel A except that a saturating amount of GroES () or cpn21 () (0.52 µM protomer, i.e. 150 pmol) was used and that the reactions were stopped with a hexokinase/glucose trap at the indicated times. C, the hydrolysis of ATP by GroEL is inhibited by cpn21 or GroES. The rates of ATP hydrolysis by GroEL (0.1 µM protomer) were determined in 160 µl containing 9.375 µM ATP (10) and were corrected for spontaneous hydrolysis. The reaction mixture was supplemented with no additives (), 0.5 µM (protomer) of GroES (), or 0.5 µM (protomer) of cpn21 (). Control reactions for identical concentrations of GroES () or cpn21 () were performed in the absence of GroEL.



ATPase Inhibition

A partial reaction of purified GroEL in the absence of other protein components is an ATPase activity (23) that is inhibited by GroES (12, 23) and by functional homologues of GroES from mitochondria (17, 19) and bacteriophage T4 (22) . ATP hydrolysis by GroEL was measured in the absence or presence of either E. coli or spinach co-chaperonins (Fig. 5 C). Both proteins inhibited ATP hydrolysis, although full inhibition was achieved more rapidly with GroES. This may indicate that the formation of a complex between GroEL and spinach cpn21 is less efficient or requires more time. Electron Microscopy of cpn21

Fig. 6 shows electron micrographs of purified spinach cpn21 negatively stained with uranyl acetate. Two main types of structure can be observed, circular molecules with an apparent hole in the center and long strands or chains. The smaller circular objects are very similar to the surface views of the toroidal GroES protein from E. coli(23) , and we interpret these to be views of the same plane of cpn21. Individual subunits cannot be clearly resolved, but again like GroES the shape of the oligomer indicates that the cpn21 subunits are arranged with rotational symmetry around an axis through the center of the toroid. Close inspection of the chains indicates that a centrally located channel of holes continues throughout the length of the strands, giving the appearance of individual cpn21 molecules stacked on top of one another like a series of discs. The edges of the strands are indented at regular intervals possibly marking the boundaries between individual cpn21 molecules. Evolutionary Conservation of cpn21 in Photosynthetic Eukaryotes


Figure 6: Electron micrographs of cpn21. The protein was fixed with 1% gluteraldehyde and negatively stained with 1% uranyl acetate. Two magnifications are shown; in panel A the bar represents 100 nm, and in panel B the bar is equivalent to 20 nm.



The two-domain structure of chloroplast cpn21 was identified by DNA sequence analysis of a spinach cDNA clone and by isolation of a related protein from pea chloroplasts (21) . Since all other co-chaperonin proteins identified in eubacteria and mitochondria are of the cpn10 single-domain type, we surveyed a broad range of distantly related photosynthetic eukaryotes for the presence of cpn21 to examine the origin and conservation of the two-domain polypeptide. The following plants were screened: liverworts ( Marchantia polymorpha), mosses ( Dicranum flagellare), club-mosses ( Selaginella kraussiania), ferns ( Nephrolepis exaltata), gymnosperms ( Ginkgo biloba, Juniperus horizontalis, and Taxus media), monocots ( Hordeum vulgare and Zea mays), and dicots ( Arabidopsis thaliana and Petunia hybrida). Soluble extracts were separated by SDS-PAGE, transferred to nitrocellulose, and incubated with antisera against spinach cpn21. The cpn21 polypeptide could be detected in all species examined, and some of these results are shown in Fig. 7. The molecular mass of cpn21 predicted from DNA sequence analysis (21 kDa) is smaller than the mass of 24 kDa observed by SDS gel electrophoresis. In most species the cross-reacting protein co-migrated with purified spinach cpn21, but two polypeptides of slightly different sizes were observed for the moss D. flagellare (Fig. 7, lane 4). The evolutionary divergence represented by this group of species covers a time span of 4 10 or more years.


Figure 7: Immunoblot of soluble leaf extracts resolved by SDS-PAGE and incubated with anti-cpn21. Lane 1, J. horizontalis; lane 2, T. media; lane 3, G. biloba; lane 4, D. flagellare; lane 5, S. kraussiania; lane 6, Z. mays; lane 7, H. vulgare. The lower arrow indicates the position of cpn21, and the upper arrow indicates cpn60 cross-reacting with contaminating anti-cpn60 in the serum. Frozen tissue was homogenized in 0.1 M Tris-HCl, pH 7.5, 0.3 M NaCl, 5 mM -mercaptoethanol, 1 mM EDTA, 1 mM KCN, and clarified in a microfuge for 10 min. The soluble fraction was precipitated with 5% trichloroacetic acid, washed with 80% acetone, and the solubilized pellet analyzed by SDS-PAGE (12.5%). Proteins were electroblotted onto nitrocellulose and sequentially incubated with rabbit anti-cpn21 and goat anti-rabbit IgG conjugated to alkaline phosphatase.




DISCUSSION

The chaperonin family of molecular chaperones are essential components that are required for the folding of proteins within cells. Folding is facilitated by transient interactions of polypeptides in their non-native states with chaperonins, followed by a discharge step mediated by a co-chaperonin protein in the presence of MgATP (reviewed in Refs. 31, 32). The pathway results in stabilization of labile protein folding intermediates and partitioning of these toward successful isomerization to native states. The most extensively studied chaperonin is GroEL from E. coli and its co-chaperonin GroES. These two proteins have an unusual structure based on a 7-fold rotational symmetry. GroEL consists of two stacked rings with each ring comprising seven identical 60 kDa subunits and forming a double-toroid structure with a central cavity (33, 34) . GroES is also thought to comprise a ring of seven subunits, but in this case there is only a single toroid and each subunit is 10 kDa (23) . The complex molecular architecture of chaperonins clearly has important mechanistic significance since the pattern of 7-fold rotational symmetry is repeated for cpn60 homologues isolated from eukaryotes (35, 36) .

Although there is conservation in the overall organization of chaperonin subunits, differences in structure have been identified among chaperonins from certain species. For example, plant chloroplasts have the dual distinction of containing atypical chaperonins and co-chaperonins. Unlike mitochondria and most eubacteria which contain only a single type of cpn60 subunit (3, 37) , chloroplasts contain two cpn60 polypeptides (termed and ) with about 50% identity (38) . The two cpn60 polypeptides may have evolved to undertake different functions within the plastid or simply represent divergent proteins with an identical function. It is not known whether the and cpn60 subunits of chloroplasts form homo- or heterooligomeric tetradecamers. It is, therefore, intriguing that chloroplasts should possess a double-domain co-chaperonin (21) which forms an adenine nucleotide-dependent complex with both the and subunits of plastid cpn60 (Fig. 3). Based on sequence homologies between each domain in cpn21 and other published cpn10 sequences we have identified a potential oligopeptide linker located in the mature protein at Thr-103 with the sequence Thr-Asp-Asp-Val-Lys-Asp. This oligopeptide has a composition highly favorable for domain linkage (39) and is effectively positioned midway along the cpn21 polypeptide to provide an internal symmetry axis around which a pseudosymmetric structure could form following collapse and folding of the two domains. As we show here each domain of cpn21 is functional, which raises the possibility that the and subunits of chloroplast cpn60 could require different co-chaperonin interactions for maximal activity. This implies that the two co-chaperonin domains have preferred binding sites on different cpn60 molecules or different faces of the same molecule. Perhaps the fused-domain structure of the chloroplast co-chaperonin satisfies this requirement by presenting two differentially active surfaces to cpn60 in the target polypeptide discharge reaction.

Irrespective of whether the two cpn10 domains of cpn21 have different specificities in their interactions with cpn60 or , why should the chloroplast co-chaperonin maintain two functional cpn10 domains fused together rather than using separate co-chaperonin polypeptides? Domain fusion to form cpn21 was an early evolutionary event as demonstrated by its presence in all photosynthetic eukaryotes screened, and the fusion has been highly conserved suggesting that retention of the two-domain structure is advantageous in chloroplasts. Other proteins have apparently evolved by domain duplication and fusion (reviewed in Ref. 40). A domain fusion arrangement would presumably be more efficient during translocation into chloroplasts because only one transit peptide would be required to transport two functional domains, and it does present a mechanism to ensure equimolar amounts of the two active parts of the co-chaperonin.

Other advantages from domain fusion, however, may be of greater significance. For example, domains are autonomous cooperative folding units, and it has been found that domain fusion enhances the rate of folding and can improve stability by reducing the entropy of the unfolded state (41) . This occurs because the two chains are no longer independent of one another. Cpn21 is more stable than the separate cpn10-C domain synthesized in E. coli (Fig. 1 D), and domain cleavage has been shown to reduce stability in other proteins, for example the C-domain of B-crystallin (42) . Perhaps the most compelling reason for domain fusion is that it provides a mechanism for the correct association of dissimilar but related subunits (43) . As each domain collapses to its native-like structure, it is tethered to the other domain by a peptide linker, thus ensuring a high local concentration of the fused subunits with a fixed polarity. This polarity may be important in the overall assembly of the cpn21 oligomer. The simplest model for assembly of subunits into oligomers is via random collisions between subunits leading to association and shuffling to yield the native quaternary structure (44) . In the absence of domain fusion, the related but distinct cpn10-N and -C subunits may have sufficient homology to initiate toroid formation, but the constraints of 7-fold symmetry would mean any oligomer containing mixed -N and -C subunits would have suboptimal complementarity, and could not achieve the close packing expected for greatest stability. By domain fusion the problem of forming two types of co-chaperonins is simplified because each distinct domain is now linked in the same polypeptide. As the cpn21 polypeptides collide and undergo association, the close proximity and high local concentrations of the two domains should favor reshuffling to achieve the closest packing and stability.

In summary, domain fusion of cpn10 in chloroplasts suggests a strategy for production of co-chaperonin polypeptides exhibiting polarity and containing two distinct but functional surfaces that exhibit modular behavior. This implies a protein folding mechanism that requires selectivity in the binding of co-chaperonins with the and forms of cpn60 to give optimum interactions and efficiency in the polypeptide discharge reaction. Clearly, a better understanding of the manner with which chloroplast co-chaperonins interact with cpn60 will require additional structural information, the isolation and analysis of mutant plastid single- and double-domain co-chaperonins, together with the and cpn60 from plants. Such studies are in progress and may provide an explanation for the unique double-domain co-chaperonins found in chloroplasts.

  
Table: Bacteriophage and T5 growth with different groES-transformants


  
Table: Suppression of a groES619 temperature-sensitive phenotype



FOOTNOTES

*
This work was supported in part by grants from the state of Saarland (to U. B.) and the Deutsche Forschungsgemeinschaft (to J. S.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked `` advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
Present address: Dept. of Chemical Engineering, BF-10, University of Washington, Seattle, WA 98195.

Present address: Institut für Physiologische Chemie, University Krankenh Eppendorf, Martinistrasse 52, D-20246 Hamburg, Germany.

**
To whom correspondence should be addressed. Tel.: 302-695-7437; Fax: 302-695-4260.

The abbreviations used are: L, large subunit of Rbu-P carboxylase; cpn10, chaperonin 10; cpn10-N, N-terminal domain of cpn21; cpn10-C, C-terminal domain of cpn21; cpn21, chaperonin 21; cpn60, chaperonin 60; IPTG, isopropyl--Dthiogalactopyranoside; Rbu-P carboxylase, ribulose-P carboxylase; S, small subunit of Rbu-P-carboxylase; PAGE, polyacrylamide gel electrophoresis; PCR, polymerase chain reaction; MES, 4-morpholineethanesulfonic acid.


ACKNOWLEDGEMENTS

We thank C. Georgopoulos and J. Zeilstra-Ryalls for strains and bacteriophages, G. Lorimer for purified GroE proteins and Rbu-P carboxylase, J. van Breemen for electron microscopy, P. Viitanen for antisera against chaperonins, and M. Todd and P. Viitanen for discussions. REFERENCES

  • Barraclough, R., and Ellis, R. J. (1980) Biochim. Biophys. Acta 608, 19-31 [Medline]
  • Roy, H., Bloom, M., Milos, P., and Monroe, M. (1982) J. Cell Biol. 94, 20-27 [Abstract]
  • Hemmingsen, S. M., Woolford, C., van der Vies, S. M., Tilly, K., Dennis, D. T., Georgopoulos, C. P., Hendrix, R. W., and Ellis, R. J. (1988) Nature 333, 330-334 [Medline]
  • Gatenby, A. A., Lubben, T. H., Ahlquist, P., and Keegstra, K. (1988) EMBO J. 7, 1307-1314
  • Bloom, M. V., Milos, P., and Roy, H. (1983) Proc. Natl. Acad. Sci. U. S. A. 80, 1013-1017
  • Ellis, R. J., and van der Vies, S. M. (1988) Photosynth. Res. 16, 101-115
  • Lubben, T. H., Donaldson, G. K., Viitanen, P. V., and Gatenby, A. A. (1989) Plant Cell 1, 1223-1230 [Abstract]
  • Viitanen, P. V., Gatenby, A. A., and Lorimer, G. H. (1992) Protein Sci. 1, 363-369 [Abstract]
  • Goloubinoff, P., Christeller, J. T., Gatenby, A. A., and Lorimer, G. H. (1989) Nature 342, 884-889 [Medline]
  • Baneyx, F., and Gatenby, A. A. (1992) J. Biol. Chem. 267, 11637-11644 [Abstract]
  • Laminet, A. A., Ziegelhoffer, T., Georgopoulos, C., and Plückthun, A. (1990) EMBO J. 9, 2315-2319 [Abstract]
  • Martin, J., Langer, T., Boteva, R., Schramel, A., Horwich, A. L., and Hartl, F-.U. (1991) Nature 352, 36-42 [Medline]
  • Viitanen, P. V., Donaldson, G. K., Lorimer, G. H., Lubben, T. H., and Gatenby, A. A. (1991) Biochemistry 30, 9716-9723 [Medline]
  • Buchner, J., Schmidt, M., Fuchs, M., Jaenicke, R., Rudolph, R., Schmid, F. X., and Kiefhaber, T. (1991) Biochemistry 30, 1586-1591 [Medline]
  • Fayet, O., Louarn, J.-M., and Georgopoulos, C. (1986) Mol. Gen. Genet. 202, 435-445 [Medline]
  • Van Dyk, T. K., Gatenby, A. A., and LaRossa, R. A. (1989) Nature 342, 451-453 [Medline]
  • Lubben, T. H., Gatenby, A. A., Donaldson, G. K., Lorimer, G. H., and Viitanen, P. V. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 7683-7687 [Abstract]
  • Hartman, D. J., Hoogenraad, N. J., Condron, R., and Hoj, P. B. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 3394-3398 [Abstract]
  • Rospert, S., Glick, B. S., Jenö, P., Schatz, G., Todd, M. J., Lorimer, G. H., and Viitanen, P. V. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 10967-10971 [Abstract]
  • Burt, W. J. E., and Leaver, C. J. (1994) FEBS Lett. 339, 139-141 [Medline]
  • Bertsch, U., Soll, J., Seetharam, R., and Viitanen, P. V. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 8696-8700 [Abstract]
  • van der Vies, S. M., Gatenby, A. A., and Georgopoulos, C. (1994) Nature 368, 654-656 [Medline]
  • Chandrasekhar, G. N., Tilly, K., Woolford, C., Hendrix, R., and Georgopoulos, C. (1986) J. Biol. Chem. 261, 12414-12419 [Abstract]
  • Gatenby, A. A., Castleton, J. A., and Saul, M. W. (1981) Nature 291, 117-121 [Medline]
  • Zeilstra-Ryalls, J., Fayet, O., Baird, L., and Georgopoulos, C. (1993) J. Bacteriol. 175, 1134-1143 [Abstract]
  • Georgopoulos, C. P., Hendrix, R. W., Casjens, S. R., and Kaiser, A. D. (1973) J. Mol. Biol. 76, 45-60 [Medline]
  • Zweig, M., and Cummings, D. J. (1973) J. Mol. Biol. 80, 505-518 [Medline]
  • Landry, S. J., Zeilstra-Ryalls, J., Fayet, O., Georgopoulos, C., and Gierasch, L. M. (1993) Nature 364, 255-258 [Medline]
  • Lissin, N. M., Venyaminov, S. Y., and Girshovich, A. S. (1990) Nature 348, 339-342 [Medline]
  • Musgrove, J. E., Johnson, R. A., and Ellis, R. J. (1987) Eur. J. Biochem. 163, 529-534 [Abstract]
  • Hendrick, J. P., and Hartl, F.-U. (1993) Annu. Rev. Biochem. 62, 349-384 [Medline]
  • Gatenby, A. A., and Viitanen, P. V. (1994) Annu. Rev. Plant Physiol. Plant Mol. Biol. 45, 469-491
  • Braig, K., Otwinowski, Z., Hegde, R., Boisvert, D. C., Joachimiak, A., and Horwich, A. L. (1994) Nature 371, 578-586 [Medline]
  • Saibil, H. R., Zheng, D., Roseman, A. M., Hunter, A. S., Watson, G. M. F., Chen, S., auf der Mauer, A., O'Hara, B. P., Wood, S. P., Mann, N. H., Barnett, L. K., and Ellis, R. J. (1993) Curr. Biol. 3, 265-273
  • Pushkin, A. V., Tsuprun, V. L., Solovjeva, N. A., Shubin, V. V., Evstigneeva, Z. G., and Kretovich, W. L. (1982) Biochim. Biophys. Acta 704, 379-384
  • McMullin, T. W., and Hallberg, R. L. (1988) Mol. Cell. Biol. 8, 371-380 [Medline]
  • Reading, D. S., Hallberg, R. L., and Myers, A. M. (1989) Nature 337, 655-659 [Medline]
  • Martel, R., Cloney, L. P., Pelcher, L. E., and Hemmingsen, S. M. (1990) Gene ( Amst.) 94, 181-187 [Medline]
  • Argos, P. (1990) J. Mol. Biol. 211, 943-958 [Medline]
  • McLachlan, A. D. (1987) Cold Spring Harbor Symp. Quant. Biol. 52, 411-420 [Medline]
  • Liang, H., Sandberg, W. S., and Terwilliger, T. C. (1993) Proc. Natl. Acad. Sci. U. S. A. 90, 7010-7014 [Medline]
  • Mayr, E-.M., Jaenicke, R., and Glockshuber, R. (1994) J. Mol. Biol. 235, 84-88 [Medline]
  • Tang, J., James, M. N. G., Hsu, I. N., Jenkins, J. A., and Blundell, T. L. (1978) Nature 271, 618-621 [Medline]
  • Jaenicke, R. (1991) Biochemistry 30, 3147-3161 [Medline]


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