Abstract Beard worms (Siboglinidae, Polychaeta) lack a mouth and a digestive tract and harbour chaemosynthetic bacteria in the bacteriocytes of the trophosome. Since beard worms depend on the organic compounds produced by the bacteria for nourishment, the bacteriocytes should be efficient in exchanging various substances with body fluids. For this reason, it is important to determine how the bacteriocytes are organized in the trophosome. As the first step of the present study, the appearance of bacteriocytes was examined in routinely stained paraffin sections. Second, visualization of the actual distribution of the bacteriocytes was attempted using whole‐mount in situ hybridization with a probe of the 16S rRNA nucleotide sequence of the bacterium. After routine haematoxylin & eosin staining, the bacteriocytes appeared to be aligned in cell cords accompanied with nutrient‐deposit cells that extended from both sides of the trophosome toward the dorsal side and folded up in the coelomic spaces. In whole‐mount preparations, however, bacteriocytes with intense signals of 16S rRNA were seen three‐dimensionally as many irregular leaves arranged from both sides of the ventral vessel toward the dorsal vessel. We will discuss the physiological significance of this characteristic distribution of the bacteriocytes in the present species.
The Escherichia coli DnaK chaperone system is a canonical heat shock protein 70 (Hsp70) chaperone system comprising Hsp70, Hsp40, and a nucleotide exchange factor. Although Hsp40 is known to facilitate the effective binding of Hsp70 to substrates, the role of Hsp40 in Hsp70-substrate interactions has not yet been fully elucidated. Using the E. coli heat shock transcription factor σ(32) as a substrate in the DnaK chaperone system, we here provide new insight into the Hsp70-substrate interaction. When DnaK-σ(32) complexes formed under various conditions were analyzed by gel filtration, several DnaK-σ(32) complexes with different molecular masses were detected. The results indicated that multiple DnaK molecules simultaneously bind to σ(32), even though it has been suggested that DnaK interacts with σ(32) at a molar ratio of 1:1. Two σ(32) mutants, L201D σ(32) and I54A σ(32), which have reduced affinities for DnaK and DnaJ (Hsp40), respectively, were used to further characterize DnaK-σ(32) complex formation. Pulldown assays demonstrated that the affinity of I54A σ(32) for DnaK was similar to that of wild-type σ(32) in the absence of DnaJ, whereas L201D σ(32) exhibited an extremely low affinity for DnaK. However, in the presence of ATP and DnaJ, the yield of DnaK eluted with L201D σ(32) was much higher than that eluted with I54A σ(32). These results indicate that there are multiple DnaK binding sites on σ(32) and that DnaJ strongly promotes DnaK binding to any site in the presence of ATP, regardless of the intrinsic affinity of DnaK for the site.
The heat shock response inEscherichia coli depends on a transient increase in the intracellular level of ς32 that results from both increased synthesis and transient stabilization of normally unstable ς32. Although the membrane-bound ATP-dependent protease FtsH (HflB) plays an important role in degradation of ς32, our previous results suggested that several cytosolic ATP-dependent proteases including HslVU (ClpQY) are also involved in ς32 degradation (Kanemori, M., Nishihara, K., Yanagi, H., and Yura, T. (1997) J. Bacteriol.179, 7219–7225). We now report on the ATP-dependent proteolysis of ς32 by purified HslVU protease and its unusual dependence on high temperature: ς32 was rapidly degraded at 44 °C, but with much slower rates (∼15-fold) at 35 °C. FtsH-dependent degradation of ς32 also gave similar results. In agreement with these results in vitro, the turnover of ς32 in normally growing cells at high temperature (42 °C) was much faster than at low temperature (30 °C). Taken together with other evidence, these results suggest that the ς32 level during normal growth is primarily determined by the stability (susceptibility to proteases) and synthesis rate of ς32 set by ambient temperature, whereas fine adjustment such as transient stabilization of ς32 observed upon heat shock is brought about through monitoring changes in the cellular state of protein folding.
GroEL C138W is a mutant form of Escherichia coli GroEL, which forms an arrested ternary complex composed of GroEL, the co-chaperonin GroES and the refolding protein molecule rhodanese at 25 °C. This state of arrest could be reversed with a simple increase in temperature. In this study, we found that GroEL C138W formed both stable trans- and cis-ternary complexes with a number of refolding proteins in addition to bovine rhodanese. These complexes could be reactivated by a temperature shift to obtain active refolded protein. The simultaneous binding of GroES and substrate to the cis ring suggested that an efficient transfer of substrate protein into the GroEL central cavity was assured by the binding of GroES prior to complete substrate release from the apical domain. Stopped-flow fluorescence spectroscopy of the mutant chaperonin revealed a temperature-dependent conformational change in GroEL C138W that acts as a trigger for complete protein release. The behavior of GroEL C138W was reflected closely in its in vivo characteristics, demonstrating the importance of this conformational change to the overall activity of GroEL. GroEL C138W is a mutant form of Escherichia coli GroEL, which forms an arrested ternary complex composed of GroEL, the co-chaperonin GroES and the refolding protein molecule rhodanese at 25 °C. This state of arrest could be reversed with a simple increase in temperature. In this study, we found that GroEL C138W formed both stable trans- and cis-ternary complexes with a number of refolding proteins in addition to bovine rhodanese. These complexes could be reactivated by a temperature shift to obtain active refolded protein. The simultaneous binding of GroES and substrate to the cis ring suggested that an efficient transfer of substrate protein into the GroEL central cavity was assured by the binding of GroES prior to complete substrate release from the apical domain. Stopped-flow fluorescence spectroscopy of the mutant chaperonin revealed a temperature-dependent conformational change in GroEL C138W that acts as a trigger for complete protein release. The behavior of GroEL C138W was reflected closely in its in vivo characteristics, demonstrating the importance of this conformational change to the overall activity of GroEL. triosephosphate isomerase guanidine hydrochloride isopropyl 1-thio-β-d-galactoside The chaperonin GroEL from Escherichia coli binds numerous proteins in vivo and in vitro and facilitates their folding by providing a space within its unique quaternary structure where protein molecules can safely complete their folding processes (1Hartl F.U. Hayer-Hartl M. Science. 2002; 295: 1852-1858Crossref PubMed Scopus (2821) Google Scholar, 2Fenton W.A. Horwich A.L. Prot. Sci. 1997; 6: 743-760Crossref PubMed Scopus (330) Google Scholar). The specific steps involved in binding, isolating, and then releasing the refolding protein molecules involve a series of intricately associated conformational changes that are performed by the GroEL protein. The conformational changes that are controlled by the binding and hydrolysis of ATP and the participation of an accessory chaperonin, GroES. The presently accepted cycle of chaperonin-facilitated protein folding (3Rye H.S. Burston S.G. Fenton W.A. Beechem J.M., Xu, Z. Sigler P.B. Horwich A.L. Nature. 1997; 388: 792-798Crossref PubMed Scopus (358) Google Scholar, 4Rye H.S. Roseman A.M. Chen S. Furtak K. Fenton W.A. Saibil H.R. Horwich A.L. Cell. 1999; 97: 325-338Abstract Full Text Full Text PDF PubMed Scopus (285) Google Scholar) consists of an initial binding of the refolding protein to GroEL and subsequent binding of ATP and GroES, which drops the refolding protein molecule into a cylindrical cavity formed by seven subunits of GroEL and seven subunits of GroES. The bound ATP molecules are hydrolyzed slowly by the GroEL subunit, and this has the effect of priming the cage formed by GroEL and GroES for release. Then, an additional seven molecules of ATP are bound to another heptameric ring bound back-to-back to the initial GroEL heptamer (the trans ring), and this binding triggers the release of the refolding protein molecule, ADP, and bound GroES. The numerous conformational changes of GroEL may be localized to three distinct domains found in the GroEL subunit (5Braig K. Otwinowski Z. Hegde R. Boisvert D.C. Joachimiak A. Horwich A.L. Sigler P.B. Nature. 1994; 371: 578-586Crossref PubMed Scopus (1201) Google Scholar). The apical domain is responsible for the recognition and binding of refolding protein molecules and the co-chaperonin GroES. The equatorial domain possesses the binding site for the nucleotide ATP, which acts as the modulating factor of these conformational changes. The intermediate domain transfers various signals between the two former domains to produce a concerted movement of the GroEL subunit. Mutations in each of these domains result in alterations in the facet of the chaperonin mechanism that is most reflective of the specific role of each domain (4Rye H.S. Roseman A.M. Chen S. Furtak K. Fenton W.A. Saibil H.R. Horwich A.L. Cell. 1999; 97: 325-338Abstract Full Text Full Text PDF PubMed Scopus (285) Google Scholar, 6Fenton W.A. Kashi Y. Furtak K. Horwich A.L. Nature. 1994; 371: 614-619Crossref PubMed Scopus (578) Google Scholar, 7Kawata Y. Kawagoe M. Hongo K. Miyazaki T. Higurashi T. Mizobata T. Nagai J. Biochemistry. 1999; 38: 15731-15740Crossref PubMed Scopus (44) Google Scholar, 8Weissman J.S. Hohl C.M. Kovalenko O. Kashi Y. Chen S. Braig K. Saibil H.R. Fenton W.A. Horwich A.L. Cell. 1995; 83: 577-587Abstract Full Text PDF PubMed Scopus (393) Google Scholar, 9Yifrach O. Horovitz A. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 1521-1524Crossref PubMed Scopus (55) Google Scholar, 10Murai N. Makino Y. Yoshida M. J. Biol. Chem. 1996; 271: 28229-28234Abstract Full Text Full Text PDF PubMed Scopus (48) Google Scholar). Many subtle changes may be seen when mutations and other alterations are introduced into the intermediate domain, as this has the effect of altering the cooperation of the other two domains to complete a functional chaperonin cycle (7Kawata Y. Kawagoe M. Hongo K. Miyazaki T. Higurashi T. Mizobata T. Nagai J. Biochemistry. 1999; 38: 15731-15740Crossref PubMed Scopus (44) Google Scholar, 11Martin J. Biochemistry. 2002; 41: 5050-5055Crossref PubMed Scopus (37) Google Scholar). In a previous study, our laboratory evaluated the effects of three mutant GroEL proteins whose mutations were each located in a different domain of the GroEL subunit (7Kawata Y. Kawagoe M. Hongo K. Miyazaki T. Higurashi T. Mizobata T. Nagai J. Biochemistry. 1999; 38: 15731-15740Crossref PubMed Scopus (44) Google Scholar). Two of these mutants, GroEL T89W and GroEL Y203C, produced predictable effects in their respective domains: GroEL T89W (12Mizobata T. Kawagoe M. Hongo K. Nagai J. Kawata Y. J. Biol. Chem. 2000; 275: 25600-25607Abstract Full Text Full Text PDF PubMed Scopus (9) Google Scholar), in the ATP hydrolytic abilities localized in the equatorial domain, and GroEL Y203C, in the substrate-binding characteristics of the apical domain. However, the most interesting mutant was localized in the intermediate domain. GroEL C138W was a mutant that was apparently indistinguishable from the wild-type chaperonin under normal experimental conditions. However, when we introduced into the experimental field a refolding protein molecule, bovine rhodanese, at 25 °C, we observed a complete arrest of both rhodanese refolding and ATP hydrolysis by GroEL. This arrest was seemingly unrelated to the presence or absence of GroES. Further experiments showed that at 25 °C, GroEL C138W formed a stable ternary complex in which rhodanese and GroES were simultaneously bound to the apical domain. More interestingly, this ternary complex could be induced to complete the chaperonin cycle by a simple increase in temperature to 37 °C. This result was interpreted to reflect the transfer of conformational signals between the equatorial domain and the apical domain, which were necessary for a coordinated movement of the GroEL subunit. In the present article, we show some further results obtained using the novel characteristics of this mutant chaperonin. We found that the arrested ternary complex formed by GroEL C138W was a general phenomenon, and similar arrested complexes could be formed with a number of refolding proteins in addition to bovine rhodanese, with molecular masses ranging from 33 to 85 kDa. Furthermore, bothcis-ternary complexes and trans-ternary complexes formed in this manner could be induced to resume folding upon a simple increase in the experimental temperature. Stopped-flow fluorescence analysis of the introduced tryptophan residue showed that a conformational change with an apparent rate constant of 82 s−1 in the presence of 1 mm ATP at 37 °C was strongly affected by this mutation. This conformational change became undetectable when the temperature was decreased from 37 to 25 °C, indicating that the mutation specifically rendered this conformational change temperature-dependent. We confirm in this report a previous assertion that this conformational change governs the release of substrate protein from the GroEL apical domain (13Jackson G.S. Staniforth R.A. Halsall D.J. Atkinson T. Holbrook J.J. Clarke A.R. Burston S.G. Biochemistry. 1993; 32: 2554-2563Crossref PubMed Scopus (238) Google Scholar), and from the nature of the arrested complex, propose a sequence of events which occur at the apical domain of GroEL immediately after ATP binding, that seemingly assures the efficient transfer of bound protein molecules into the GroEL central cavity. GroEL C138W was expressed in E. coli JM109/pUCESL cells and purified at 4 °C according to published protocols (14Mizobata T. Kawata Y. Biochim. Biophys. Acta. 1994; 1209: 83-88Crossref PubMed Scopus (34) Google Scholar). The substrate proteins phosphofructokinase, luciferase, pyruvate kinase, triosephosphate isomerase (TIM),1 aldolase, rhodanese, and adenylate kinase were obtained from Sigma. Gp23 protein was isolated from cells possessing a plasmid (pET23a(+)/gp23), which contained the structural gene for phage T4 gp23 protein. The gene was isolated from bacteriophage T4 using PCR (the phage samples were a generous gift from Dr. Fumio Arisaka of the Tokyo Institute of Technology) and ligated into the EcoRI/BamHI restriction sites of pET23a(+). In vivo activities of GroEL C138W were evaluated using E. coli strain KY1156, which possesses a groESL operon placed under the control of thelacUV5 promoter. E. coli KY1156 was constructed as follows. λlacUV5p-groESgroEL, a derivative of Charon 25, was first constructed by in vivo recombination (15Kanemori M. Mori H. Yura T. J. Bacteriol. 1994; 176: 4235-4242Crossref PubMed Google Scholar). The resultant phage carries a groESL operon placed under thelacUV5 promoter and lacI q genes, which were originally contained in plasmid pKV1561. This phage specimen was used to infect E. coli strain MC4100 [F− araD Δ(argF-lac)U169 rpsL relA flbB deoC ptsF rbsR]. Then, the ΔgroE68::tet mutation of KY1880 (15Kanemori M. Mori H. Yura T. J. Bacteriol. 1994; 176: 4235-4242Crossref PubMed Google Scholar) was transduced into this λ-lysogenic strain in the presence of IPTG by phage P1 to produce strain KY1156, whose genomic copy of groESL had been disrupted bytetr, and which possesses an IPTG-induciblegroESL operon within the lysogenic insert of λlacUV5p-groESgroEL. Taka-amylase A was obtained by purification and recrystallization from a commercial product, Takadiastase Sankyo (16Kawata Y. Hongo K. Mizobata T. Nagai J. Protein Eng. 1998; 11: 1293-1298Crossref PubMed Scopus (13) Google Scholar). Guanidine hydrochloride (GdnHCl) was obtained from Wako Pure Chemicals. Other reagents were obtained commercially. Each substrate protein was extensively unfolded in the presence of 4 m GdnHCl and subsequently diluted 100-fold into Refolding Buffer A (50 mm Tris-HCl, pH 7.4, containing 100 mm KCl, 10 mm MgCl2, and 2 mm 2-mercaptoethanol) containing an equimolar concentration of GroEL C138W 14-mer. Immediately afterward, 2 mm ATP was added, and each mixture was incubated for 10 min at 25 °C. Where indicated, an equimolar concentration of GroES heptamer was also added at this point. The mixture was loaded onto a Sephacryl S-300 size-exclusion column (φ 1.2 × 25.5 cm) equilibrated with Refolding Buffer A containing 2 mm ATP, either to analyze complex formation or to isolate arrested complexes for further analysis. For direct analysis of eluted fractions, 1-ml fractions were mixed with trichloroacetic acid (final concentration, 10%) to precipitate protein, and the precipitated proteins were subjected to SDS-polyacrylamide gel electrophoresis (15% gel). In order to distinguish between cis- and trans-ternary complexes, arrested chaperonin complexes that were isolated by elution from Sephacryl S-300 as described above were subjected to brief treatments with proteinase K. Samples of isolated complexes were incubated in the presence of 0.01 mg/ml proteinase K at 25 °C. At the times indicated in the figure legends, phenylmethanesulfonyl fluoride was added to a final concentration of 1 mm to stop digestion. This mixture was analyzed for remaining substrate protein using SDS-polyacrylamide gel electrophoresis and densitometry. In cases where the subunit molecular mass of the refolding protein molecule was similar to the subunit molecular mass of GroEL (causing signal overlap), the substrate proteins were visualized by covalent labeling of substrate proteins with Cy5 (Amersham Biosciences) prior to experimentation. A typical labeling reaction of substrate protein with Cy5 was performed by incubating protein samples in 0.1 msodium carbonate buffer, pH 9.5, containing a 10-fold molar concentration of labeling dye relative to the substrate protein. After labeling, the samples were desalted into 0.1 m Tris-HCl, pH 7.4, containing 0.1 m KCl and 20 mmMg(CH3COO)2 in order to remove unbound dye. In a typical labeling reaction, an average of 4 dye molecules were bound per substrate protein molecule. Quantitation of covalently labeled substrate protein was performed using a Fujifilm FLA-2000 fluorescence image analyzer. Refolding assays of Taka-amylase A and rhodanese were performed according to Kawata et al. (16Kawata Y. Hongo K. Mizobata T. Nagai J. Protein Eng. 1998; 11: 1293-1298Crossref PubMed Scopus (13) Google Scholar) and Kawata et al. (7Kawata Y. Kawagoe M. Hongo K. Miyazaki T. Higurashi T. Mizobata T. Nagai J. Biochemistry. 1999; 38: 15731-15740Crossref PubMed Scopus (44) Google Scholar), respectively. Refolding assays of TIM were performed as follows. Native TIM was denatured for 1 h in 4 m GdnHCl at a protein concentration of 1 mg/ml. To initiate refolding, this protein sample was diluted 100-fold in Refolding Buffer B (50 mm Tris-HCl, pH 7.4, 50 mm KCl, 20 mmMg(CH3COO)2, and 2 mmdithiothreitol), containing an equimolar concentration of GroEL and GroES oligomers where indicated. ATP was absent for the first 3 min, after which 2 mm ATP was added to the refolding mixture. At the indicated intervals, 2-μl aliquots of the refolding mixture were taken and added to assay mixture (Refolding Buffer B containing 40 mm glyceraldehyde 3-phosphate, 13 units/ml glycerol 3′-phosphate dehydrogenase (Roche Molecular Biochemicals) and 2 mm NADH). The reaction was followed by the decrease in absorbance at 340 nm concomitant with the production of glycerol 3-phosphate. In an additional experiment, samples of TIM complexed with GroEL, GroES, and ATP were applied to the size-exclusion column described above in the complex isolation experiments. Each collected fraction was then incubated at 37 °C for 10 min to allow resumption of refolding. These fractions were then quantitated for TIM activity, and the results were plotted relative to the fraction number. The amount of ATP that was hydrolyzed by the arrested ternary complex (GroEL-bovine rhodanese-GroES) was estimated according to the following protocol, which was modeled after the ATPase protocol of Horovitzet al. (17Horovitz A. Bochkareva E.S. Kovalenko O. Girshovich A.S. J. Mol. Biol. 1993; 231: 58-64Crossref PubMed Scopus (59) Google Scholar). Experiments were performed in ATPase buffer (50 mm Tris-HCl, pH 7.8, containing 10 mmMgCl2, 10 mm KCl, 10 mm glucose, and 1 mm dithiothreitol). Bovine rhodanese (Sigma) was incubated in rhodanese denaturing buffer (37.5 mm Tris-HCl buffer, pH 7.4, containing 6 m GdnHCl and 6 mmdithiothreitol) for 1 h to completely unfold the protein. This mixture was then diluted with the same denaturing buffer to produce a series of unfolded rhodanese samples with different rhodanese concentrations but a fixed denaturant concentration of 6 mGdn-HCl. Five μl of each unfolded rhodanese sample were then rapidly mixed with 500 μl of reaction mixture containing 60 nmGroEL C138W tetradecamer and 100 nm wild-type GroES heptamer, which had been preincubated for 10 min at 25 °C. The end result was to produce a series of samples containing differing concentrations of arrested rhodanese-GroEL C138W complex. The mixtures were further incubated for 10 min at 25 °C, whereupon the ATPase activities of each sample were assayed by addition of 70 μm ATP (1.4 μCi of [γ-32P]ATP) and incubation at 25 °C. After hydrolysis was allowed to proceed for 30 min, 60-μl aliquots from each reaction mixture were taken and mixed with 40 μl of stop solution (1 m perchloric acid and 1 mm sodium phosphate) and placed on ice for 20 min. Then, 200 μl of 20 mm ammonium molybdate was added, and this mixture was vortexed vigorously for 30 s, followed by addition of 300 μl of water-saturated isopropyl acetate and a second vortexing. The concentration of phosphomolybdate complex that partitioned into the organic phase was measured by taking 150-μl aliquots of the organic phase and counting the radioactivity by Cerenkov counting on a Wallac 1409 liquid scintillation counter. Raw values were corrected for spontaneous hydrolysis of ATP. However, the partitioning efficiency of phosphomolybdate complex into the organic phase was assumed to be 100% in the subsequent conversion of radioactivity to phosphate concentration. Stopped-flow fluorescence analysis of the changes in tryptophan-derived fluorescence of GroEL C138W were measured on an Applied Photophysics SX-17MV fluorescence spectrophotometer. The excitation wavelength was 295 nm, and a cutoff filter was used to monitor fluorescence intensities above 320 nm. All samples were prepared in stopped-flow buffer (50 mm triethanolamine, pH 7.5, containing 20 mmMgCl2 and 50 mm KCl). The experimental temperature was controlled with a water bath. Samples of GroEL C138W were mixed rapidly with ATP so that the final concentrations of GroEL and ATP were 0.5 mg/ml and 1 mm, respectively. Experiments were performed at 25 and 37 °C, and the results from five runs were averaged and were analyzed using the analysis software of the system according to either a single exponential or a sequential two-component exponential mechanism. Plasmid pACYCESLC138W was constructed by ligating a fragment of pUCESLC138W excised by EcoRI and SmaI with plasmid pACYC184 prepared by digestion with EcoRI and ScaI. This plasmid was used to transform E. coli strain KY1156 to produce strain KY1156/pACYCESLC138W. This strain was grown on Luria-Bertani plates containing the antibiotics kanamycin (50 μg/ml) and tetracycline (20 μg/ml) at either 37 or 25 °C, in both the presence and absence of 1 mm IPTG. In the absence of IPTG,E. coli KY1156 requires an alternate source of active GroEL protein to be viable at 37 °C. Preliminary experiments showed that in the presence of IPTG, the size of the colonies of E. coliKY1156/pACYCESLC138W after a 24-h incubation at 25 °C was the same as those seen after a 12-h incubation at 37 °C (data not shown). The GroEL C138W-rhodanese-GroES ternary complex observed in a previous study was postulated to be an integral part of the overall mechanism of GroEL function (7Kawata Y. Kawagoe M. Hongo K. Miyazaki T. Higurashi T. Mizobata T. Nagai J. Biochemistry. 1999; 38: 15731-15740Crossref PubMed Scopus (44) Google Scholar). However, there was the possibility that this ternary complex might have been a special intermediate, which could be observed only during the refolding reaction of rhodanese. If this were the case, the detection of such an intermediate would not hold any relevance to the overall chaperonin cycle. In this study, therefore, we performed a series of experiments in order to demonstrate the ubiquity of this complex in the refolding reactions of numerous substrate proteins, as well as the consistency in the characteristics that are observed. We used the following proteins in our evaluation: adenylate kinase from chicken (molecular mass = 21 kDa × 1), TIM from rabbit muscle (26.6 kDa × 2), bovine rhodanese (33 kDa × 1) (18Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13044-13049Abstract Full Text PDF PubMed Google Scholar), aldolase from rabbit muscle (39.2 kDa × 4), Taka-amylase A fromAspergillus oryzae (54 kDa × 1) (16Kawata Y. Hongo K. Mizobata T. Nagai J. Protein Eng. 1998; 11: 1293-1298Crossref PubMed Scopus (13) Google Scholar), Gp23 protein from T4 phage (56 kDa × 1), GroEL protein from E. coli(57 kDa × 14), pyruvate kinase from rabbit muscle (58 kDa × 4), luciferase from firefly (61 kDa × 1) (19Fridmann Y. Ulitzur S. Horovitz A. J. Biol. Chem. 2000; 275: 37951-37956Abstract Full Text Full Text PDF PubMed Scopus (12) Google Scholar), and phosphofructokinase from rabbit muscle (85.1 kDa × 4). The subunit molecular masses of the substrate proteins varied from 21 to 85 kDa, and represented a fair sampling of the various protein sizes that may be found in a typical cell. Fig. 1shows the results of a typical experiment in which a number of these proteins were renatured in the presence of GroEL C138W, ATP, and GroES at 25 °C, fractionated by size-exclusion chromatography, and analyzed by SDS-PAGE. As shown for the small protein adenylate kinase in Fig. 1 A, proteins which do not form stable ternary complexes with GroEL C138W and GroES did not co-elute with GroEL C138W under our experimental conditions. Contrary, in the case of aldolase for example, a ternary complex was detected and isolated, as indicated by the co-elution of the band corresponding to aldolase in high-molecular mass fractions containing GroEL C138W (Fig. 1 B). Aldolase was by no means the only protein to display this behavior, and as shown in Fig. 1 C, TIM, Taka-amylase A, and phosphofructokinase were also shown to form stable ternary complexes with GroEL C138W at 25 °C. Further experiments showed that of the ten proteins sampled in this study, only the smallest, adenylate kinase, was incapable of forming a stable ternary complex with GroEL C138W and GroES at 25 °C (TableI).Table IFormation of stable binary and ternary chaperonin complexes by GroEL C138W at 25 °CSubstrate proteinTotal molecular mass (kDa)Binary complexTernary complexPhosphofructokinase (rabbit)85.1 × 4++Luciferase (firefly)61 × 1++Pyruvate kinase (rabbit)58 × 4++GroEL (E. coli)57 × 14++Gp23 (T4 phage)56 × 1++Taka-amylase A (A. oryzae)54 × 1++Aldolase (rabbit)39 × 4++Rhodanese (bovine)33 × 1++TIM (rabbit)26 × 2++1-aTernary complexes were detected, but dissociated protein is also seen in the lower molecular mass fractions, suggesting a less extensive interaction with GroEL.Adenylate kinase (chicken)21 × 1+−1-a Ternary complexes were detected, but dissociated protein is also seen in the lower molecular mass fractions, suggesting a less extensive interaction with GroEL. Open table in a new tab Taking into consideration various recent studies that showed that the size of the protein molecule which could be secluded within the GroEL central cavity was restricted to proteins smaller than 60 kDa (20Houry W.A. Frishman D. Eckerskorn C. Lottspeich F. Hartl F.U. Nature. 1999; 402: 147-154Crossref PubMed Scopus (438) Google Scholar, 21Sakikawa C. Taguchi H. Makino Y. Yoshida M. J. Biol. Chem. 1999; 274: 21251-21256Abstract Full Text Full Text PDF PubMed Scopus (99) Google Scholar), it was especially interesting that phosphofructokinase, with a molecular mass of 85 kDa, was capable of forming a stable ternary complex with GroEL and GroES. Therefore, the nature of the stable ternary complex formed by GroEL C138W, GroES and each substrate protein was probed in more detail, to determine the specific orientation of GroEL, the refolding protein, and GroES. Various substrate proteins were first refolded in the presence of either GroEL C138W only or an equimolar concentration of GroEL and GroES with 2 mm ATP. The mixtures were then fractionated using size-exclusion chromatography to isolate arrested chaperonin complexes. Each purified complex was then subjected to a brief treatment with proteinase K, in order to digest any substrate protein that was accessible to the protease. Fig. 2 exemplifies the results of this experiment. As seen in the figure, we found that the substrate proteins we used could be separated largely into two groups. The first group, composed of proteins such as phosphofructokinase and pyruvate kinase, whose subunit molecular masses exceeded 55 kDa, were bound in a stable ternary complex with GroEL but displayed no differences when GroES was present, indicating that these proteins were exclusively bound in thetrans conformation in the arrested ternary complex. In contrast, proteins such as Taka-amylase A and TIM, whose subunit molecular masses fell within the boundaries dictated by the size of the central cavity of GroEL (22Xu Z. Horwich A.L. Sigler P.B. Nature. 1997; 388: 741-750Crossref PubMed Scopus (1053) Google Scholar), could be detected in significant amounts when GroES was added to the initial mixture, suggesting that a fraction of the substrate protein was shielded from digestion by the cochaperonin. Table II summarizes the relative percentages of cis ternary complex which was detected for each refolding protein. From the results in Table II, we postulate that refolding in the presence of GroEL C138W at 25 °C results in the formation of stable ternary complexes, and the specific nature of these complexes were dependent mainly on the subunit molecular mass of the refolding substrate protein.Table IIEstimation of the percentage of cis-ternary complex by brief digestion of arrested binary (− GroES) and ternary (+ GroES) chaperonin complexes with proteinase KSubstrate protein2-aThe amount of trapped protein detected at zero time (minus proteinase K) was set to 100%. (subunit molecular mass)Binary complexed (− GroES)Ternary complexed (+ GroES)TIM (26 kDa)1845Rhodanese (33 kDa)2.648.8Taka-amylase A (54 kDa)4.262.5Gp23 (56 kDa)1.33.0GroEL (57 kDa)5.214.1Luciferase (61 kDa)8.810.9Pyruvate kinase (58 kDa)17.212.02-a The amount of trapped protein detected at zero time (minus proteinase K) was set to 100%. Open table in a new tab Next, we assayed the refolding reactions of a number of proteins that formed stable ternary complexes in Figs. 1 and 2. Fig. 3 shows the results of refolding assays performed on Taka-amylase A and TIM at 25 °C, the restrictive temperature, and 37 °C, the temperature at which the behavior of GroEL C138W reverts to that of the wild-type chaperonin. For comparison, refolding assays of rhodanese in the presence of GroEL C138W are also shown in a separate panel. Taka-amylase A (16Kawata Y. Hongo K. Mizobata T. Nagai J. Protein Eng. 1998; 11: 1293-1298Crossref PubMed Scopus (13) Google Scholar) and TIM are examples of non-stringent proteins, with relatively high spontaneous refolding yields under our experimental conditions. Rhodanese is an example of a protein whose stringency during refolding is modulated by the refolding conditions (23Mendoza J.A. Demeler B. Horowitz P.M. J. Biol. Chem. 1994; 269: 2447-2451Abstract Full Text PDF PubMed Google Scholar, 24Martin J. Langer T. Boteva R. Schramel A. Horwich A.L. Hartl F.-U. Nature. 1991; 352: 36-42Crossref PubMed Scopus (728) Google Scholar). In this assay, rhodanese refolding was performed under non-permissive conditions (24Martin J. Langer T. Boteva R. Schramel A. Horwich A.L. Hartl F.-U. Nature. 1991; 352: 36-42Crossref PubMed Scopus (728) Google Scholar). As shown in the figure, at 25 °C each protein was unable to complete folding, regardless of the presence of the co-chaperonin GroES and ATP. At 37 °C, however, the refolding reaction of all three proteins in the presence of GroEL C138W was measurably improved compared with the reactions at 25 °C, demonstrating again the temperature-dependent characteristics of this chaperonin. Next, we probed the characteristics of the two types of ternary complex, cis and trans, formed by GroEL C138W, refolding protein, and GroES in detail, specifically, the fate of each type of complex after an increase in temperature to 37 °C, which removes the structural block on the mutant chaperonin. The experiment consisted of first forming the stable ternary complex at 25 °C, isolating arrested complexes by fractionation using size-exclusion chromatography, and then incubating each fraction at 37 °C to allow resumption of the refolding reaction. Each fraction was then assayed for refolded protein activity. The results are shown in Fig. 4 for the substrate protein TIM. When the fractions are incubated at 25 °C, TIM activity was detected only in the fractions corresponding to where spontaneously refolded TIM would be recovered (Fig. 4 A, 25 °C). SDS-PAGE analysis of the same fractions indicated however that TIM was still coeluted in the fractions containing GroEL/GroES, suggesting that the ternary complex was stable for the duration of the experiment. When the fractions were incubated at 37 °C, however, an additiona
Production of abnormal proteins during steady-state growth induces the heat shock response by stabilizing normally unstable sigma32 (encoded by the rpoH gene) specifically required for transcription of heat shock genes. We report here that a multicopy plasmid carrying the hslVU operon encoding a novel ATP-dependent protease inhibits the heat shock response induced by production of human prourokinase (proUK) in Escherichia coli. The overproduction of HslVU (ClpQY) protease markedly reduced the stability and accumulation of proUK and thus reduced the induction of heat shock proteins. In agreement with this finding, deletion of the chromosomal hslVU genes significantly enhanced levels of proUK and sigma32 without appreciably affecting cell growth. When the deltahslVU deletion was combined with another protease mutation (lon, clpP, or ftsH/hflB), the resulting multiple mutations caused higher stabilization of proUK and sigma32, enhanced synthesis of heat shock proteins, and temperature-sensitive growth. Furthermore, overproduction of HslVU protease reduced sigma32 levels in strains that were otherwise expected to produce enhanced levels of sigma32 due either to the absence of Lon-ClpXP proteases or to the limiting levels of FtsH protease. Thus, a set of ATP-dependent proteases appear to play synergistic roles in the negative control of the heat shock response by modulating in vivo turnover of sigma32 as well as through degradation of abnormal proteins.
We purified the nitrate reductase from the soluble fraction of Magnetospirillum magnetotacticum MS-1. The enzyme was composed of 86- and 17-kDa subunits and contained molybdenum, non-heme iron, and heme c. These properties are very similar to those of the periplasmic nitrate reductase found in Paracoccus pantotrophus. The M. magnetotacticum nap locus was clustered in seven open reading frames, napFDAGHBC. The phylogenetic analyses of NapA, NapB, and NapC suggested a close relationship between M. magnetotacticum nap genes and Escherichia coli nap genes, which is not consistent with the 16S rDNA data. This is the first finding that the α subclass of Proteobacteria possesses a napFDAGHBC-type nap gene cluster. The nap gene cluster had putative fumarate and nitrate reduction regulatory protein (Fnr) and NarL protein binding sites. Furthermore, we investigated the effect of molybdate deficiency in medium on the total iron content of the magnetosome fraction and discussed the physiological function of nitrate reductase in relation to the magnetite synthesis in M. magnetotacticum.Key words: nitrate reductase, magnetotactic bacteria, denitrification, horizontal gene transfer.
