Productive and Nonproductive Intermediates in the Folding of Denatured Rhodanese
2000; Elsevier BV; Volume: 275; Issue: 1 Linguagem: Inglês
10.1074/jbc.275.1.63
ISSN1083-351X
AutoresMarkandeswar Panda, Boris Gorovits, Paul M. Horowitz,
Tópico(s)Biomedical Research and Pathophysiology
ResumoThe competition between protein aggregation and folding has been investigated using rhodanese (thiosulfate:cyanide sulfurtransferase, EC 2.8.1.1) as a model. During folding from a urea-denatured state, rhodanese rapidly forms associated species or intermediates, some of which are large and/or sticky. The early removal of such particles by filtration results in a decreased refolding yield. With time, a portion of the smaller aggregates can partition back first to intermediates and then to refolded protein, while a fraction of these irreversibly form unproductive higher aggregates. Dynamic light scattering measurements indicate that the average sizes of the aggregates formed during rhodanese folding increase from 225 to 325 nm over 45 min and they become increasingly heterogeneous. Glycerol addition or the application of high hydrostatic pressure improved the final refolding yields by stabilizing smaller particles. Although addition of glycerol into the refolding mixture blocks the formation of unproductive aggregates, it cannot dissociate them back to productive intermediates. The presence of 3.9 m urea keeps the aggregates small, and they can be dissociated to monomers by high hydrostatic pressure even after 1 h of incubation. These studies suggest that early associated intermediates formed during folding can be reversed to give active species. The competition between protein aggregation and folding has been investigated using rhodanese (thiosulfate:cyanide sulfurtransferase, EC 2.8.1.1) as a model. During folding from a urea-denatured state, rhodanese rapidly forms associated species or intermediates, some of which are large and/or sticky. The early removal of such particles by filtration results in a decreased refolding yield. With time, a portion of the smaller aggregates can partition back first to intermediates and then to refolded protein, while a fraction of these irreversibly form unproductive higher aggregates. Dynamic light scattering measurements indicate that the average sizes of the aggregates formed during rhodanese folding increase from 225 to 325 nm over 45 min and they become increasingly heterogeneous. Glycerol addition or the application of high hydrostatic pressure improved the final refolding yields by stabilizing smaller particles. Although addition of glycerol into the refolding mixture blocks the formation of unproductive aggregates, it cannot dissociate them back to productive intermediates. The presence of 3.9 m urea keeps the aggregates small, and they can be dissociated to monomers by high hydrostatic pressure even after 1 h of incubation. These studies suggest that early associated intermediates formed during folding can be reversed to give active species. 2-mercaptoethanol The renaturation of a denatured protein typically involves a number of intermediate states (on-pathway or off-pathway) leading to either correctly folded or misfolded structures (1Baldwin R.L. Fold. Des. 1995; 1: R1-R8Abstract Full Text Full Text PDF PubMed Scopus (235) Google Scholar, 2Hagihara Y. Goto Y. Fink A.L. Goto Y. Molecular Chaperones in the Life Cycle of Proteins. Marcel Dekker, New York1998: 1-33Google Scholar). Protein aggregation is one of the main side reactions during refolding, and it often results from interactions among partially folded intermediates (3Jaenicke R. Seckler R. Wetzel R. Advances in Protein Chemistry. Academic Press, New York1997: 1-59Google Scholar). In general, this aggregation, although ubiquitous and important for basic and applied problems, is difficult to study. The enzyme rhodanese (thiosulfate:cyanide sulfurtransferase, EC 2.8.1.1) is an interesting model for studying issues related to such problems in protein folding (4Mendoza J.A. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 16973-16976Abstract Full Text PDF PubMed Google Scholar, 5Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13587-13591Abstract Full Text PDF PubMed Google Scholar, 6Horowitz P.M. Butler M. J. Biol. Chem. 1993; 268: 2500-2504Abstract Full Text PDF PubMed Google Scholar, 7Gorovits B.M. McGee W.A. Horowitz P.M. Biochim. Biophys. Acta. 1998; 1382: 120-128Crossref PubMed Scopus (31) Google Scholar, 8Gorovits B.M. Horowitz P.M. Biochemistry. 1998; 37: 6132-6135Crossref PubMed Scopus (94) Google Scholar). This monomeric protein containing 293 amino acids (mass = 33 kDa) is folded into two independent, equal-size domains, and its crystal structure is available (9Ploegman J.H. Drent G. Kalk K.H. Hol W.G. Heinrikson R.L. Keim P. Weng L. Russell J. Nature. 1978; 273: 124-129Crossref PubMed Scopus (214) Google Scholar). The domains are tightly associated and the interdomain surface is highly hydrophobic. Rhodanese contains four cysteine residues, Cys-63, Cys-247, Cys-254, and Cys-263, which are all reduced in the active protein (9Ploegman J.H. Drent G. Kalk K.H. Hol W.G. Heinrikson R.L. Keim P. Weng L. Russell J. Nature. 1978; 273: 124-129Crossref PubMed Scopus (214) Google Scholar). During unfolding/refolding in the presence of denaturants, disulfides can form among these cysteines, leading to misfolded structures in addition to aggregation. The disulfide-containing species leading to misfolding have been characterized (10Horowitz P.M. Hua S. Biochim. Biophys. Acta. 1995; 1249: 161-167Crossref PubMed Scopus (10) Google Scholar) with SDS-polyacrylamide gel electrophoresis, and the formation of aggregates during unfolding has been reported earlier (5Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13587-13591Abstract Full Text PDF PubMed Google Scholar, 11Tandon S. Horowitz P.M. J. Biol. Chem. 1989; 264: 9859-9866Abstract Full Text PDF PubMed Google Scholar). It has been established that during unfolding, the presence of reductants such as β-ME1 and thiosulfate prevent the formation of most of the disulfides and better refolding yields are observed (5Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13587-13591Abstract Full Text PDF PubMed Google Scholar). However, refolding at low temperature (e.g. 12 °C), even in the presence of 200 mmβ-ME, 50 mm thiosulfate, and at very low protein concentration (e.g. 3.6 μg/ml), led to incomplete recovery of activity (5Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13587-13591Abstract Full Text PDF PubMed Google Scholar, 11Tandon S. Horowitz P.M. J. Biol. Chem. 1989; 264: 9859-9866Abstract Full Text PDF PubMed Google Scholar). Folding intermediates formed during chemical or thermal denaturation often contain similar levels of secondary structure as the native proteins, but they contain a decreased number of tertiary contacts (12Ptitsyn O.B. Creighton T.E. Protein Folding. W. H. Freeman and Co., New York1992: 243-300Google Scholar), partially or completely dissociated domains (13Horowitz P.M. Criscimagna N.L. J. Biol. Chem. 1990; 265: 2576-2583Abstract Full Text PDF PubMed Google Scholar), and/or incorrectly formed disulfide bonds (14Ewbank J.J. Creighton T.E. Nature. 1991; 350: 518-520Crossref PubMed Scopus (134) Google Scholar). As a result, such intermediates tend to be highly hydrophobic, and consequently they can easily form large aggregates and precipitate. It was suggested that, at intermediate concentrations of denaturant, the two domains of rhodanese dissociate (7Gorovits B.M. McGee W.A. Horowitz P.M. Biochim. Biophys. Acta. 1998; 1382: 120-128Crossref PubMed Scopus (31) Google Scholar, 13Horowitz P.M. Criscimagna N.L. J. Biol. Chem. 1990; 265: 2576-2583Abstract Full Text PDF PubMed Google Scholar) to produce a form of the protein that is able to form dimers, trimers, and higher oligomers (6Horowitz P.M. Butler M. J. Biol. Chem. 1993; 268: 2500-2504Abstract Full Text PDF PubMed Google Scholar). Formation of similar intermediates leading to incomplete refolding can be detected where aggregation is a major side-reaction. Indeed, the protein folding process can be described as a kinetic competition between correct folding and aggregation (15Kiefhaber T. Rudolph R. Kohler H.H. Buchner J. Bio/Technology. 1991; 9: 825-829Crossref PubMed Scopus (390) Google Scholar). Such competition has been observed in vitro (16Zettlmeissl G. Rudolph R. Jaenicke R. Biochemistry. 1979; 18: 5567-5571Crossref PubMed Scopus (274) Google Scholar) as well as in vivo (17King J. Haase C. Yu M.-H. Oxender D.L. Fox C.F. Protein Engineering. Alan R. Liss, Inc., New York1987: 109-121Google Scholar,18Klein J. Dhurjati P. Appl. Environ. Microbiol. 1995; 61: 1220-1225Crossref PubMed Google Scholar). Quantitative models have been proposed to explain the formation of aggregates during protein folding (15Kiefhaber T. Rudolph R. Kohler H.H. Buchner J. Bio/Technology. 1991; 9: 825-829Crossref PubMed Scopus (390) Google Scholar). Aggregation can be prevented by isolating intermediates from each other by complexing them with molecular chaperones (4Mendoza J.A. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 16973-16976Abstract Full Text PDF PubMed Google Scholar, 19****Google Scholar), utilizing detergents or mixed micelles (20Tandon S. Horowitz P.M. J. Biol. Chem. 1987; 262: 4486-4491Abstract Full Text PDF PubMed Google Scholar), or refolding at low protein concentration (5Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13587-13591Abstract Full Text PDF PubMed Google Scholar). The stabilizing effects of some co-solvents have also been partially ascribed to decreased diffusion, which prevents protein molecules from interacting with each other during the period in which they are sensitive to interaction (21Chrunyk B.A. Matthews C.R. Biochemistry. 1990; 29: 2149-2154Crossref PubMed Scopus (73) Google Scholar) along with the effects of preferential hydration (22Timasheff S.N. Shirley B.A. Methods in Molecular Biology. Humana Press, Totowa, NJ1995: 253-269Google Scholar). Several publications have demonstrated that high hydrostatic pressures can dissociate specific protein oligomers (23Paladini Jr., A.A. Weber G. Biochemistry. 1981; 20: 2587-2593Crossref PubMed Scopus (208) Google Scholar, 24Weber G. Protein Interactions. Chapman and Hall Inc., New York1992Google Scholar, 25Markley J.L.E. High-Pressure Effects in Molecular Biophysics and Enzymology. Oxford University Press, New York1996Crossref Google Scholar, 26Prehoda K.E. Mooberry E.S. Markley J.L. Biochemistry. 1998; 37: 5785-5790Crossref PubMed Scopus (62) Google Scholar). In addition, high hydrostatic pressure has recently been shown to dissociate nonspecific protein aggregates formed in vitro (8Gorovits B.M. Horowitz P.M. Biochemistry. 1998; 37: 6132-6135Crossref PubMed Scopus (94) Google Scholar), and to facilitate an increase in the folding yield by allowing intermediates to complete folding. In the present study, we have investigated the stability and relative size of aggregates formed during rhodanese folding on the efficiency of renaturation. On the basis of these studies, a minimum mechanism for rhodanese folding is proposed. The observations support the view that intermediate, dissociated species can be rescued and returned into a productive folding pathway. Recombinant bovine rhodanese (thiosulfate:cyanide sulfurtransferase, EC 2.8.1.1) was purified as described previously, and was stored at −70 °C as a crystalline suspension in 1.8m ammonium sulfate containing 1 mm sodium thiosulfate (27Miller D.M. Delgado R. Chirgwin J.M. Hardies S.C. Horowitz P.M. J. Biol. Chem. 1991; 266: 4686-4691Abstract Full Text PDF PubMed Google Scholar). For the experiments, a concentrated rhodanese stock solution (typically 10–20 mg/ml) free from ammonium sulfate was prepared by gel filtration using a G-50 Sephadex column and eluting with 0.1 m sodium phosphate, pH = 7.6. Rhodanese concentrations were determined using a value of A280 nm0.1% = 1.75 (28Sorbo B.H. Acta Chem. Scand. 1953; 7: 1129-1136Crossref Google Scholar). In the experiments for refolding and filtration of aggregates done at low protein concentrations (∼3.6 μg/ml), it was not possible to accurately determine [protein] by using the standard BCA (Pierce) or Coomassie (Pierce) assays. In these cases, a standard dilution curve was made using the fluorescence emission intensity of rhodanese from 1–20 μg/ml whose folding state was normalized by denaturing the samples in 2% SDS (final concentration). The fluorescence intensities were measured at the emission maximum, which was at 344 nm for all samples (Fig. 6, inset). Other experimental details are in the legend for Fig. 6. The standard curve was linear to 20 μg/ml with a near zero intercept and slope of 0.050 ± 0.003 cps ml/μg (see Fig. 6, inset). Quantitation by fluorescence intensity measurements of total protein after SDS denaturation avoids errors in the measurements that might result from small differences in the misfolded species, intermediates passing through the filter, and different unfolded species in the filtrate. In the present study, it was not possible to isolate the contributions of individual species to the total recovered protein. Urea was electrophoresis purity from Bio-Rad. All other reagents were of analytical grade. Solutions containing glycerol and β-ME were prepared fresh. To avoid rhodanese inactivation, glycerol stock solutions were used within 10 days. A standard buffer containing 200 mm β-ME, 50 mm sodium thiosulfate, and 50 mm Tris-HCl, pH 7.8, was used throughout this study (5Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13587-13591Abstract Full Text PDF PubMed Google Scholar). Buffers containing glycerol were made by adding appropriate amounts from a 8 m stock solution of glycerol to the other reagents and then making up the final volume of the solution followed by minor adjustment of the pH. All stock solutions were filtered through 0.2-μm (Nalgene) syringe filters. Rhodanese activity was measured by a colorimetric method (monitored at 460 nm) based on the formation of the complex between ferric ions and one of the reaction products, thiocyanate (28Sorbo B.H. Acta Chem. Scand. 1953; 7: 1129-1136Crossref Google Scholar). Aliquots of 25–50 μl of the incubating enzyme were added to 1 ml of assay mixture and incubated for 5–10 min. The assay mixture was at pH = 8.6 and consisted of 1:1:1 (volume) of 0.15m sodium thiosulfate, 0.15 m KCN, and 0.12m KH2PO4. The reaction was stopped by adding 0.5 ml of 18% formaldehyde solution. Color was developed by adding 1.5 ml of ferric nitrate solution. The presence of small amounts of β-ME, urea, or glycerol after dilution of the enzyme in the assay mixture did not interfere with the assay. The activities of the refolded rhodanese were normalized with respect to the activity of a sample of native enzyme under identical assay conditions. Unfolded rhodanese (U) was freshly prepared at 300 μg/ml by adding calculated amounts of native enzyme into 8 m urea, 50 mm Tris, 50 mm sodium thiosulfate, 1 mm β-ME, pH = 7.8, and equilibrated for at least one hour (7Gorovits B.M. McGee W.A. Horowitz P.M. Biochim. Biophys. Acta. 1998; 1382: 120-128Crossref PubMed Scopus (31) Google Scholar). Refolding was achieved by diluting the unfolded protein into the standard refolding buffer, maintaining a final protein concentration of 3.6 μg/ml. All measurements were made at 25 °C. The regain of enzyme activity was used to monitor successful refolding. The refolding time was varied for the kinetics study. At the indicated times, the protein solutions were filtered by centrifugation for 15–20 s at 6,000 × gusing a 0.22-μm Ultrafree-MC filter unit (Millipore). Identical results were achieved by filtering through Nalgene 0.2-μm (cellulose acetate) syringe filters using sterile plastic syringes (Becton-Dickinson). The filtrate was collected and analyzed for rhodanese activity, and protein concentration was measured by comparing fluorescence intensities of the solutions with that of native rhodanese under identical conditions. were monitored by dynamic light scattering measurements employing a Brookhaven laser light scattering instrument (Brookhaven Instruments, Holtsville, NY). Buffers were filtered using Nalgene 0.2-μm syringe silter, and samples were prepared as described by the manufacturer to avoid contamination by dust or presence of tiny bubbles that would contribute to scattering. The laser was set at 488 nm, and scattering was monitored at a 90o angle. One-ml samples were used in cylindrical glass tubes. All the measurements were made at 25 °C. Appropriate corrections were applied for scattering by buffer. After refolding was initiated, the distributions of particle sizes were determined at the indicated times. Data were collected and analyzed using Dynamic Light Scattering software 9KDLSW (beta version 1.2, 1995) provided by Brookhaven Instruments. Sizes were calculated using the diffusion coefficient measured by the dynamic light scattering method as described by Cleland and Wang (29Cleland J.L. Wang D.I. Biochemistry. 1990; 29: 11072-11078Crossref PubMed Scopus (146) Google Scholar). Protein aggregation under pressure was studied using a spectrofluorometer (ISS, Inc.) as described previously (8Gorovits B.M. Horowitz P.M. Biochemistry. 1998; 37: 6132-6135Crossref PubMed Scopus (94) Google Scholar). The pressure bomb employed was similar to the one described by Paladini and Weber (23Paladini Jr., A.A. Weber G. Biochemistry. 1981; 20: 2587-2593Crossref PubMed Scopus (208) Google Scholar). Native rhodanese was diluted into a solution containing 50 mm Tris-HCl, pH 7.8, 200 mm β-ME, 50 mm thiosulfate, maintaining a final urea concentration of 3.9 m and a final protein concentration of 0.3 mg/ml. After various times of incubation the samples were pressurized to between 0.001 and 2 kbar. The intensity of the scattered light was monitored at 90o using light at 400 nm. To investigate rhodanese refolding under high hydrostatic pressure, samples were pressurized up to 2 kbar at 5–10 min after dilution of the denatured protein. These samples were incubated for 70 min, depressurized, and the activities were analyzed as described above. The activity of the native protein kept at atmospheric pressure was taken as 100%. Rhodanese was covalently labeled by succinimidyl 1-pyrenebutyrate as described previously (30Gorovits B.M. Ybarra J. Seale J.W. Horowitz P.M. J. Biol. Chem. 1997; 272: 26999-27004Abstract Full Text Full Text PDF PubMed Scopus (13) Google Scholar). Briefly, protein at a final concentration of 88 μm was dissolved in triethanolamine hydrochloride (50 mm, pH 7.8). Succinimidyl 1-pyrenebutyrate was diluted from a stock solution in dimethylformamide to a final concentration of 130 μm. The mixture was incubated for 2 h at room temperature and then dialyzed against Tris-HCl (50 mm, pH 7.8) containing 0.1 m β-ME to exchange the buffer and remove unreacted labeling reagent. Pyrene-labeled rhodanese was used to monitor protein aggregation under pressure. Rhodanese covalently labeled with pyrene was diluted into the standard buffer containing 3.9 m urea. In some cases, unlabeled rhodanese was added to maintain the final protein concentration at 0.3 mg/ml. At the indicated times, samples were pressurized as described above. The polarization of the pyrene fluorescence was monitored using an ISS fluorometer (ISS, Inc.). Samples were excited at 345 nm, and the emission was detected at 400 nm. Kinetic experiments were done as discussed in Fig. 1 legend. The data were fitted to either mono- or bi-exponential first-order equations:Y = A 1 × exp (−k 1 × t) +A 2 or Y =A 1 × exp (−k 1 ×t) + A 2 × exp (−k 2 × t) +A 3, respectively. The independent variableY was the percentage of activity recovered at timet. The pseudo-first order rate constantsk 1 and k 2 and the amplitudes A 1, A 2, andA 3 were obtained from iterative non-linear least squares regression of the data using the Origin software program (MicroCal). Rhodanese at 300 μg/ml was unfolded in 8 m urea (see “Experimental Procedures”) and then diluted to give a final urea concentration of 96 mm and 4 m glycerol to allow rhodanese to refold (Fig. 1). The refolding of rhodanese is considerably slower in the presence of glycerol (7Gorovits B.M. McGee W.A. Horowitz P.M. Biochim. Biophys. Acta. 1998; 1382: 120-128Crossref PubMed Scopus (31) Google Scholar) than without it, thus permitting observation of the kinetics of the process by the methods used here. The plateaus of these kinetic traces from the single exponential fits correspond to the maximum yields (A 2 in “Experimental Procedures”). For example, under these conditions, about 35% of the activity could be recovered with a t 0.5 of ∼10 min at [rhodanese] = 3.6 μg/ml (Fig. 1, open circles). The curves in Fig. 1 show that the final yield decreased as the rhodanese concentration increased as reported earlier (7Gorovits B.M. McGee W.A. Horowitz P.M. Biochim. Biophys. Acta. 1998; 1382: 120-128Crossref PubMed Scopus (31) Google Scholar). The observed rates from single-exponential fits were k 1(obs) = 0.067 ± 0.007, 0.066 ± 0.005, 0.17 ± 0.01, 0.28 ± 0.13, and 0.27 ± 0.04 min−1 for 3.6, 10, 15, 30, and 50 μg/ml rhodanese, respectively. The calculated yields were 35, 30, 28, 18, and 14% at 3.6, 10, 15, 30, and 50 μg/ml rhodanese, respectively, when 4m glycerol was present in the standard buffer (Fig. 1). The concentration dependence of the observed rates and the maximum yields derived from the data in Fig. 1 are shown in Fig. 2 (top and bottom panels, respectively). Such non-linear dependences of both the rate and maximum regain of activity are consistent with previous results (5Mendoza J.A. Rogers E. Lorimer G.H. Horowitz P.M. J. Biol. Chem. 1991; 266: 13587-13591Abstract Full Text PDF PubMed Google Scholar). These observations highlight the complex mechanism involving competitive pathways for folding and aggregation (see “Discussion”). Samples at different protein concentrations were derived from the same unfolded rhodanese solution in 8 murea to avoid experimental errors due to any difference in unfolded states of rhodanese. In this method, the urea concentrations in the final refolding mixtures depended on the dilution ratio. This variation in urea from 0 to 1.33 m in the final refolding mixtures did not cause significant changes in the enzyme activity of control samples whether glycerol was present or not. Experiments were repeated using a protocol in which the final mixtures were supplemented with urea so that the different rhodanese concentrations were refolded at the same final urea concentration of 1.33 m (Fig. 2,bottom panel). These results show that the results do not depend on differential carryover of urea. The size distributions of the aggregates that formed during the rhodanese folding were determined by light scattering measurements performed at various times after initiating refolding, as described under “Experimental Procedures.” The results are shown in Fig. 3, where the curves represent the size distributions determined at the indicated times. For clarity, the individual curves have been displaced vertically, and, within each curve, the heights represent the relative amounts of the indicated sizes. Any information on the size and nature of individual particles within the distribution curves is beyond the scope of the present investigation because of the broadness and complexity of the distributions (31Murphy R.M. Yarmush M.L. Colton C.K. Biopolymers. 1991; 31: 1289-1295Crossref PubMed Scopus (10) Google Scholar). The average diameters of particles at the maxima of the curves and the average widths at half-height of the distributions provide a reasonable description of the multimers relevant to the aims of this study (29Cleland J.L. Wang D.I. Biochemistry. 1990; 29: 11072-11078Crossref PubMed Scopus (146) Google Scholar, 31Murphy R.M. Yarmush M.L. Colton C.K. Biopolymers. 1991; 31: 1289-1295Crossref PubMed Scopus (10) Google Scholar). Within the first minute after initiating folding, the lowest curve in Fig. 3 shows that particles with average diameters of ∼225 nm can be detected (the diameter of native rhodanese is ∼5.6 nm). The total intensity of the scattered light quickly increased within the time of manual mixing (less than 1 min), indicating that the particles observed were formed quickly. The average size of the particles increased from ∼225 nm to ∼330 nm over 44 min (highest curve, Fig. 3). The heterogeneity of the ensemble also increased during this interval as evidenced by the increase of width at half-height of the distributions from ∼94 nm at 1 min to ∼200 nm at 44 min. The fates of intermediates or particles formed during refolding were tested by a double dilution method in which refolding was initiated at a high concentration for various periods before being diluted further. Thus, refolding was started (diluting the unfolded protein into 4m glycerol-containing buffer) at 50 μg/ml rhodanese. After different times (Scheme 1), aliquots of the refolding rhodanese were further diluted to a final concentration of 3.6 μg/ml and then immediately assayed. Fig. 4 shows that, within 30 s after the refolding was started, the protein regained about 35% of the control activity, and dilution did not dissociate the aggregates that had already formed at that instant. The recoverability of activity continued to fall exponentially. The data (Fig. 4, solid circles, solid line) were fit to a biphasic rate expression (see “Experimental Procedures”), and two rate constants were evaluated for the resolved components (Fig. 4,dashed and dotted lines). The faster observed rate (k′ obs = 0.68 ± 0.07 min−1) corresponds to a half-life of 1.02 min, whereas the slower rate (k″ obs = 0.035 min−1) corresponds to a half-life of about 19.8 min. The important observation from this analysis is that the fast phase reaches a plateau corresponding to ∼12% activity in ∼5 min (Fig. 4,dotted line), after which there is a slow, further loss of activity (Fig. 4, dashed line). We have no information regarding the slow process, but it could be due to additional loss of recoverability by the formation of misfolded structures from any of the postulated intermediates. This suggests the existence of off-pathway intermediates that contribute to the loss of recoverable activity due to the time-dependent formation of aggregated species. If the species responsible for the loss were reversible aggregates formed simply because of high enzyme concentration, then they would have dissociated upon further dilution resulting in recovered activities of about 35% at these final [protein] = 3.6 μg/ml. In that case, a plot of percentage of activity recovered versus time would have been a horizontal straight line at ∼35%, instead of the observed exponential decrease in Fig. 4.FIG. 4Dilution of refolding enzyme at different stages of refolding. Unfolded rhodanese was diluted in to 4m glycerol-containing buffer at 50 μg/ml and then the refolding mixture was diluted to 3.6 μg/ml of protein at different times (Scheme 1) with the same 4 m glycerol-containing buffer. The assays were done immediately after the final dilution. Thesolid line through the data is from fit to a biphasic exponential as described under “Experimental Procedures.” The dotted line and the dashed line represent the resolved fast and slow phases that were generated using the parameters obtained from the biphasic fit of the data.View Large Image Figure ViewerDownload (PPT) Some of the off-pathway particles formed on dilution of denatured rhodanese could be filtered through a 0.22-μm filter, a process requiring between 15 and 20 s. The results are shown in Fig. 5. When the solutions were filtered early in the refolding process (closed squares, filtration at ∼3 min), the renaturation of the protein in the filtrate initially was at the expected value compared with an unfiltered sample (open squares), and the activity increased with time to give a final yield that was decreased to ∼58% of what would have been observed in the absence of filtration. Thus, a portion of the protein that would have renatured in solution was removed on the filter (compare the closed square and the open square at 45 min). However, a portion of the protein that did pass through the filter was not active initially, but it continued to fold as shown (solid squares). Thus, there were two forms of initially inactive protein that could fold in a time-dependent manner, one was retained on the filter and the second could pass the filter. An analogous result was observed for a solution filtered 9 min after initiating renaturation (closed diamonds). Here, there was a smaller total increase in recovered activity from ∼55% immediately after filtration to a maximum recovery of ∼75%, a value still below the unfiltered control. When the solution was filtered after 29 min of refolding (open diamonds), the final concentration of the active enzyme was very close to that in the unfiltered solution. It was thus apparent from the reduced final yields that filtration could remove species that were able to contribute to the yield of refolding. The time dependent increase in activity in the filtrates showed that some inactive species could pass through the filter and become active with time. The results in Fig. 6 show that the amount of total protein in the filtrate rapidly increased over the first 5 min and then more slowly, consistent with the idea that reduced recovery of activity after filtration (Fig. 5) correlates with the removal of protein rather than with inactivation of species formed during incubation. It may be noted that, in control experiments, only 2–5% of native or urea denatured rhodanese samples were lost on the 0.2-μm filters (data not shown), whereas these filters retained very large fractions of protein when refolding rhodanese
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