P22 tailspike trimer assembly is governed by interchain redox associations

Biochimica et Biophysica Acta 1700 (2004) 105 – 116 www.bba-direct.com

P22 tailspike trimer assembly is governed by interchain redox associations B.L. Danek 1, A.S. Robinson * Department of Chemical Engineering, University of Delaware, Newark, DE 19716, USA Received 16 April 2003; received in revised form 7 April 2004; accepted 8 April 2004 Available online 10 May 2004

Abstract Though disulfide bonds are absent from P22 tailspike protein in its native state, a disulfide-bonded trimeric intermediate has been identified in the tailspike folding and assembly pathway in vitro. The formation of disulfide bonds is critical to efficient assembly of native trimers as mutations at C-terminal cysteines reduce or inhibit trimer formation. We investigated the effect of different redox folding environments on tailspike formation to discover if simple changes in reducing potential would facilitate trimer formation. Expression of tailspike in trxB cell lines with more oxidizing cytoplasms led to lower trimer yields; however, observed assembly rates were unchanged. In vitro, the presence of any redox buffer decreased the overall yield compared to non-redox buffered controls; however, the greatest yields of the native trimer were obtained in reducing rather than oxidizing environments at pH 7. Slightly faster trimer formation rates were observed in the redox samples at pH 7, perhaps by accelerating the reduction of the disulfide-bonded protrimer to the native trimer. These rates and the effects of the redox system were found to depend greatly on the pH of the refolding reaction. Oxidized glutathione (GSSG) trapped a tailspike intermediate, likely as a mixed disulfide. This trapped intermediate was able to form native trimer upon addition of dithiothreitol (DTT), indicating that the trapped intermediate is on the assembly pathway, rather than the aggregation pathway. Thus, the presence of redox agents interfered with the ability of the tailspike monomers to associate, demonstrating that disulfide associations play an important role during the assembly of this cytoplasmic protein. D 2004 Elsevier B.V. All rights reserved. Keywords: Protein folding; Redox environment; Tailspike; Folding intermediate; Protein assembly

1. Introduction Since Anfinsen’s [1] work describing the self-sufficiency of the amino acid sequence in determining protein structure, many investigators have sought to understand the underlying guidelines that ultimately lead to native structures and

Abbreviations: DTT, dithiothreitol; DTTox, oxidized DTT; IAM, iodoacetamide; SDS, sodium dodecyl sulfate; EDTA, ethylene diaamine tetraactetic acid disodium salt; Tris, Tris(hydroxymethyl) aminomethane chloride; PAGE, polyacrylamide gel electrophoresis; GSH, reduced glutathione; GSSG, oxidized glutathione; AMP, ampicillin; LB, Luria broth; IPTG, isopropyl-D-thiogalactopyranoside; BPTI, bovine pancreatic trypsin inhibitor; RNase A, ribonuclease A; IGF-I, insulin-like growth factor-I; RCP, riboflavin carrier protein; rhm-CSFbeta, recombinant human macrophage colony stimulating factor beta * Corresponding author. Tel.: +1-302-831-0557; fax: +1-302-8316262. E-mail address: [email protected] (A.S. Robinson). 1 Current address: 608 Jimmy Fund Building, Dana Farber Cancer Institute, 44 Binney St., Boston, MA 02115, USA. 1570-9639/$ - see front matter D 2004 Elsevier B.V. All rights reserved. doi:10.1016/j.bbapap.2004.04.001

conformations. There has been significant progress in understanding the intricate and detailed interactions of small and relatively simple proteins as they proceed through their folding pathways [2– 5], and more limited strides in describing more complex and oligomeric proteins [6 – 9]. Disulfide-containing proteins have been of particular interest in the field of protein folding as the presence of disulfide bonds in folding intermediates has been used as a probe to define and quantify folding pathways and intermediates. The characterization of disulfide bonded folding intermediates has revealed an unexpected role during folding for nonnative interactions, or interactions not present in the native conformation. Non-native disulfide pairing has been implicated in a variety of different protein-folding pathways, including bovine pancreatic trypsin inhibitor (BPTI) [10 – 12], ribonuclease A (RNase) [13], insulin-like growth factor-I (IGF-I) [14], recombinant human macrophage colony stimulating factor h (rhm-CSFh) [15] and riboflavin carrier protein (RCP) [16]. One surprising development was the discovery of a non-

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native disulfide bond in a folding intermediate of P22 tailspike, a protein that contains no native disulfide bonds [17]. This indicated that the traditional route of exploring protein-folding pathways from a native structure backward could overlook crucial interactions not immediately apparent from a protein’s crystal structure or unfolding pathway. More recent studies show that transient intramolecular disulfide bonds facilitate the initial stages of cytoplasmic folding of the multimeric VP1 capsid protein in simian virus 40 infected kidney cells [18]. Thus non-native disulfide bond formation has been identified in folding pathways within both prokaryotic and eukaryotic cytoplasms. Here we study further the importance of redox environments during the folding and assembly of the P22 tailspike protein. P22 tailspike is a trimeric protein composed of three identical monomers with 666 amino acids each [19]. The native trimer is resistant to denaturation by SDS, proteolysis, and has a melting temperature greater than 80 jC [20,21]. This extreme stability is believed to result from interdigitation of h-strands between the subunits rather than intersubunit covalent linkages (Fig. 1A) [22]. Because of the large degree of hydrogen bonding and hydrophobic interaction between the subunits, folding and assembly of the tailspike trimer are coupled processes. Tailspike assembly occurs over a much longer time frame than many classically studied folding models, with a half

time of 5 min in vivo [20] and 30 min in vitro [17,23,24]. Due to the slow assembly, tailspike folding intermediates can be characterized by nondenaturing gel electrophoresis or size exclusion chromatography [17,25,26]. Using native gel electrophoresis, a trimeric folding intermediate, protrimer (Fig 1B), was found to contain interchain disulfide bonds [17], even though all the cysteine residues are in the reduced state in the native trimer [22,27]. The reduction of these interchain disulfide bonds leads to the formation of the native conformation [17]. Additionally, the sulfhydryl-modifying reagent, iodoacetamide (IAM), blocks trimer assembly, both in vivo [28] and in vitro [17], suggesting that cysteine residues are active thiol agents during folding in the cell. Investigations into the nature of these unexpected cysteine interactions first identified that critical residues are located in the C-terminal tail, as a truncation at 489 was not reactive with radiolabeled IAM [28]. Mutational studies of the three cysteines in the C-terminus, 496, 613 and 635, support the hypothesis that cysteines play significant roles during the folding and assembly of the tailspike trimer [29]. In particular, trimer formation was noticeably inhibited in mutants at these residues, and no appreciable trimer was formed when both cysteines at 613 and 635 are mutated to serine [29,30]. The presence of disulfide bonds in folding intermediates has been described more recently in both the dimer [31] and monomer [29].

Fig. 1. Structure and folding/aggregation pathway of P22 tailspike. (A) Structure of P22 tailspike (108 – 666) [22]. Subunits are shown in black, dark gray, and light gray. (B) Schematic diagram of the folding pathway for P22 tailspike [17,25,38,48]. IM, monomeric folding intermediate; IM*, monomeric aggregation intermediate; ID, dimeric folding intermediate; ID*, dimeric aggregation intermediate.

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Significant advances in understanding the key cysteine residues have been made, but the importance of the local and bulk redox environment on the efficiency of folding intermediate formation has not been comprehensively investigated. One might expect that a protein without disulfide bonds would be insensitive to redox environment. Therefore, the observations of a dependence of trimer yield and formation rate on redox environment would support the importance of thiol activity. Our goals are ultimately to understand the timing and connectivity of the bond formation in the folding intermediates. In this study we characterize the effect of altering redox conditions on the competition between tailspike trimer folding and aggregation. We additionally investigate the effect of altering the pH of folding in the presence and absence of redox reagents. We consider the specific redox characteristics of the folding intermediates as they proceed through the folding and assembly pathway.

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min to pellet cells. The cell pellet was resuspended to 8 ODml, approximately one-fourth the original culture volume, in lysis solution (50 mM Tris, pH 8.0, 2 mM EDTA, 20 mM MgSO4, 20 Ag/ml DNAse, 100 Ag/ml lysozyme, 0.1% Triton X-100) and subjected to two freeze-thaw cycles to complete lysis. The lysed cells were then centrifuged at 20,000  g for 10 min to separate the pellet (insoluble protein aggregate) from the supernatant (soluble protein). The supernatant was decanted and one-third volume 3  sodium dodecyl sulfate (SDS) sample buffer (163 mM Tris, pH 6.8, 0.25 mg/ml bromphenol blue, 5 mg/ml SDS, 50% glycerol) or one-third volume 3  nondenaturing sample buffer (15 mM Tris base, 0.12 M glycine, 0.25 mg/ml bromphenol blue, 30% glycerol) was added. The pellet was resuspended in 1  sample buffer to the original volume of the lysis solution. Samples were separated on 7.5% acrylamide gels containing 1% SDS and Coomassie-stained. 2.4. Large batch protein expression and purification

2. Materials and methods 2.1. Materials SDS, glycine, Tris, and EDTA were obtained from BioRad Laboratories (Hercules, CA). Electrophoresis grade urea was obtained from Fisher Scientific (Pittsburgh, PA). DTT, DTTox, GSH and GSSG for redox refolding buffers were obtained from Sigma (St. Louis, MO). 2.2. Cell strains AD494 [D(ara -leu)7697 araD139 lacX74 DgalE galK rpsL phoR D(phoA)PvuII DmalF3 thi trxB::kan] and parental strain DHB4 Escherichia coli cells [32] were a gift of J. Beckwith (Harvard). AD494 trxB cells were maintained on media containing 30 Ag/ml of kanamycin (KAN). Lysogens of these strains were constructed using the EDE3 Lysogenization Kit (Novagen) in order to support expression from the pET plasmids. The Origamik strain was obtained from Novagen, and cells were maintained on KAN (30 Ag/ml) and tetracycline (12.5 Ag/ml). 2.3. In vivo protein expression Bacterial cells were made competent for transformation by electroporation [33] or heat shock [34]. Cells were then transformed with plasmid DNA via electroporation (BioRad Gene Pulser) or heat shock, and plasmid selection maintained by ampicillin (AMP) addition (100 Ag/ml). Transformed cells were grown at the desired temperature in Luria broth [34] containing AMP (LB-AMP) to an OD600 of 0.5 – 0.7, and protein expression was induced by the addition of isopropyl-h-D-thiogalactopyranoside (IPTG) to 1 mM. Protein was expressed for 3 –4 h. Aliquots of cell culture were removed and centrifuged at 20,000  g for 2

Tailspike protein was produced from overexpression in wild type DHB4 E. coli cells similar to in vivo expression experiments described above. One to six liters of bacterial cultures were harvested for soluble tailspike trimers. Tailspike protein was then purified essentially as described [35] to >95% purity. 2.5. In vitro refolding studies Purified tailspike protein was denatured at 1 mg/ml in 50 mM Tris, 8 M urea, pH 3.0 for 1 h. Refolding was initiated by rapid dilution of the denatured protein to the appropriate concentration into a refolding buffer (50 mM Tris, 1 mM EDTA, pH 7.6). For redox refolding studies, freshly opened redox agents were used to make 100 mM stocks of each DTT, DTTox, GSH and GSSG in 50 mM Tris and 1 mM EDTA at the indicated pH. The redox stocks were diluted with 50 mM Tris and 1 mM EDTA at the appropriate pH to investigate various reagent concentrations. For the inert environment refolding reactions, Tris refolding buffer and 8 M urea at pH 3.0 were bubbled overnight with nitrogen to remove any dissolved oxygen. Three 30Al aliquots of the urea solution and a larger pool of refolding buffer were placed in small glass scintillation vials (5 ml), which were then placed together into a large glass vessel, sealed and purged with nitrogen. Then, 3 Al of a 10 mg/ml wild type tailspike protein stock was added to each of the urea aliquots to a final concentration of 1 mg/ml and the vessel was once again purged with nitrogen. These solutions were allowed to incubate at room temperature for 60 min to denature to monomers. Refolding was initiated by adding 270 Al of refolding buffer to the denatured chains to a final concentration of 100 Ag/ml, and the vessel was resealed and purged with nitrogen at least 15 times. Refolding was allowed to proceed under nitrogen for 5 days at room temperature before adding 1/2 sample volume of native

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sample buffer. Triplicate parallel control refolding reactions were performed under atmospheric conditions. The reactions were quenched by the addition of 1/2 sample volume of 3  nondenaturing sample buffer (15 mM Tris, 0.12 M glycine, 0.25 mg/ml bromphenol blue, 30% glycerol) and placing on wet ice. Samples were then separated on 7% nondenaturing Tris/glycine acrylamide gels and silverstained as described previously [36]. The addition of glycerol sample buffer and placement on wet ice has been shown to be sufficient in halting further chain association and preventing alterations of tailspike folding intermediate populations during the time scale of electrophoretic separation [25]. 2.6. Protein concentration quantitation and kinetic analysis Protein concentrations were determined as described previously [29]. Briefly, the trimer band densities of silver-stained nondenaturing gels were compared to a linear calibration curve defined by three to five standards of known trimer concentration. The use of nondenaturing gels to isolate temporal folding intermediates has been used for many years for the tailspike system [17,25,29,31,37,38]. One- and two-dimensional gel analyses of samples held on ice and from those run on nondenaturing gels indicate that no detectable assembly changes occur during gel separation at 4 jC [25]. Therefore, we used this approach to determine the observed rate of trimer formation with confidence that the time delay did not alter the observed trimer concentrations or folding intermediates. Silver staining of nondenaturing gels was more consistent and had a broader linear range for trimer quantification than silver or Coomassie staining of SDS gels (data not shown). Refolding yields were calculated from five to seven independent data sets. Trimer-concentration-versus-time data were fit to a first-order relationship through nonlinear least squares analysis for three to four independent reactions. This has been shown previously to be the best method to fit trimer formation with time data [39]. This analysis is not strictly accurate for describing folding kinetics under aggregating conditions as it discounts the competition with the aggregation pathway. However, it allows for a convenient comparison between the folding behaviors at various conditions. Error was determined as the standard deviation from the average of three or more independently calculated formation rates.

3. Results 3.1. Altering redox conditions affects tailspike yields in vivo In order to determine whether transient disulfide bonding would be altered by simple changes in cellular redox state, expression of tailspike protein was examined in wild-type E. coli strain DHB4(DE3), and a mutant strain lacking thio-

redoxin reductase, AD494 trxB(DE3), which leads to a more oxidizing cytoplasm. Tailspike protein expression was induced in mid-logarithmic growth by the addition of IPTG to 1 mM. Cell samples were collected 3 h after induction and lysed as described in Materials and methods. The soluble and insoluble fractions were separated by centrifugation, and the samples were separated on SDS-PAGE (Fig. 2A). Quantitation of trimer yields from five independent transformants showed that approximately 50% less trimer was produced per cell in the AD494 trxB cells than in the parental DHB4 cells (Fig. 2B). Thus, wild-type tailspike chains folded productively in both cell lines at 30 jC, but the yields were significantly lower in trxB cells, which have a more oxidizing cellular environment. When the tailspike protein levels were corrected for total protein per lane, the relative trimer levels in AD494 trxB cells increased, but still remained lower (f 60%) than that of DHB4 (data not shown). Tailspike protein was also expressed in Origamik cells (Novagen), which have a trxB gor mutation and an even

Fig. 2. Cellular redox changes lower tailspike trimer yields. (A) Coomassiestained SDS-PAGE of wild-type tailspike expression. DHB4 cells and AD494 trxB cells were transformed with either pET11a plasmid (control) or wild-type tailspike plasmid pET-tsp (TSP) and grown at 30 jC. Protein expression was induced by addition of 1 mM IPTG, cell samples were taken at 3 h following induction, and cells were lysed and fractionated into soluble (S) and pellet (P) fractions. These samples were separated by SDSPAGE, and Coomassie-stained. Pure tailspike trimer (Std) and molecular weight markers (MW) are indicated. (B) Quantitation of wild-type tailspike expression. NIH Image was used to determine protein band densities for quantitative comparisons of protein yields. Error bars represent the standard deviation from the average of five independent transformants.

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more oxidizing cellular environment than the AD494 trxB cells. In repeated experiments, very little tailspike protein was expressed, although the majority was as the soluble native trimer (data not shown). The growth rates for trxB cells and Origamik cells were slower than for the parental DHB4 cells, and transformation efficiencies were also lower, perhaps due to a rapid decline in viability in the stationary phase of growth. Lower trimer yields likely result from the lower cell viability and not a direct effect of the altered cellular cytoplasm. It is not possible to verify the redox potential in these cells experimentally; there are several disulfidebonded proteins for which the Origami cells do not improve yields (G. Georgiou, personal communication). Pulse-chase labeling of tailspike chains in wild-type and AD494 trxB cells shows that apparent trimer formation rates are the same (Bhatia and Robinson, unpublished data). The lack of any improved relative trimer assembly in these cell lines indicates that bulk changes of cytoplasmic redox conditions do not facilitate interchain interactions. 3.2. Redox reagents interfere with tailspike refolding and reduce yields Our in vivo data showed that a simple change in cytoplasmic reducing potential did not improve tailspike trimer expression. By moving to an in vitro system, we wanted to isolate tailspike folding from any cellular variables, such as lower viability and protein expression levels. Previous studies revealed that in vitro, low protein concentrations mimicked low temperature expression, while high protein concentrations mirrored physiological expression temperatures [29]. Previously, tailspike refolding yields were shown to be sensitive to dithiothreitol (DTT) addition when refolded at a total protein concentration of 30 Ag/ml [17]. However, other researchers found contradictory data using a glutathione system and a total protein concentration of 50 Ag/ml [37]. We chose to investigate refolding at a high protein concentration (100 Ag/ml) to capture interactions that more closely imitate cellular behavior. Here we tested the effects of redox buffer composition on the rates and yields of tailspike folding and assembly. We investigated the effect of a glutathione buffer system, a monothiol, at various ratios of reduced to oxidized form to alter the bulk reducing potential of the buffer, while maintaining a constant concentration of buffer components. By altering the bulk redox environment, the formation of various folding intermediates could likely be manipulated and perhaps optimized. Denatured tailspike was diluted into refolding buffer (50 mM Tris, 1 mM EDTA) at pH 7.0 with 10 mM glutathione at five ratios of reduced and oxidized species (10 mM GSH, 3:1 GSH/GSSG, 1:1 GSH/GSSG, 1:3 GSH/GSSG and 10 mM GSSG). Two hours after initiating refolding, the reactions were halted by the addition of sample buffer and

Fig. 3. In vitro refolding yields are decreased in redox buffers, but maximum yields result from the most reducing environments. (A) Tailspike refolding reactions at 100 Ag/ml after 2 h at room temperature. Tailspike trimer was denatured at 1 mg/ml in 8 M urea at pH 3.0 and diluted 10-fold into 50 mM Tris and 1 mM EDTA at a refolding pH of 7.0. Redox reagents were used at various ratios as indicated, in a total 10 mM glutathione buffer. Sample lacking redox buffer is indicated (C). Lane 1 contains a 10 Ag/ml tailspike trimer standard (Std). Redox buffers were freshly prepared from 100 mM frozen stocks of the reagents. Reactions were halted by the addition of 1/2 volume of native sample buffer (see Materials and methods) and placement on wet ice. Samples were separated on native gels and visualized by silver staining. (B) Yields from refolding reactions held at room temperature for 5 – 7 h. Samples were prepared as in (A). Control reactions were performed in 50 mM Tris, 1 mM EDTA at the indicated pH. Redox buffers were composed of 5 mM GSH (1:0), 2.5 mM GSH and 2.5 mM GSSG (1:1) and 5 mM GSSG (0:1) at the indicated pH. Trimer concentrations were quantified by comparison to a standard curve fit from quantitation of three to five trimer standard concentrations. Yields are the average of two to four independent reactions, and the error is one standard deviation.

separated on 7% nondenaturing gels (Fig. 3A). Tailspike trimer yields decreased with increasing oxidizing agent concentration, and at the most oxidizing conditions, 10 mM GSSG, no apparent trimer was observed. Therefore, the presence of GSSG interferes with tailspike assembly and prevents the formation of native trimer. The maximal rate of disulfide exchange reactions occurs when the pH of the refolding buffer is equal to the pKa of thiol group [40]. The pH of the refolding buffer was varied to change the fraction of the protein thiols

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existing as thiolate ions. Thus, we could examine the effect that these ions played in the trimerization reactions. As the pH was altered, ion states could potentially facilitate disulfide bond formation and isomerization in folding intermediates. Tailspike protein was refolded at 100 Ag/ml and room temperature, as above, in glutathione buffers at a total concentration of 5 mM and three ratios of reduced to oxidized species (1:0, 1:1 and 0:1). After 5– 7 h, 1/2 sample volume of 3  native sample buffer was added to halt each reaction, and the samples were separated on 7% nondenaturing gels. Trimer concentrations of three to five independent refolding reactions were determined by measuring gel band densities and comparison to a calibration curve of three to five trimer concentration standards. Fig. 3B shows the tailspike trimer yields at each buffer composition considered. At pH 7 (Fig. 3B), the presence of any oxidized glutathione (GSSG) considerably decreased observed trimer yields. In fact, trimer formation was significantly inhibited in the presence of GSH, as seen in Fig. 3A. At higher pHs, tailspike trimer yields became even more sensitive to the addition of oxidizing agents. At pH 8 and 8.5 (Fig. 3B), the addition of any oxidized component severely affected trimer yield and the presence of GSSG at any level completely inhibited observable trimer levels to the sensitivity of the staining. In the reducing redox agent buffers, GSH slightly decreased the trimer yield over the salt buffer at these pH values. In order to examine whether or not molecular oxygen was playing a role in the in vitro refolding reactions, we initiated refolding reactions in a nitrogen inert environment. Refolding reactions were performed, and trimer yields calculated as described in Materials and methods. The trimer yields of the reactions in the inert environment and those for a parallel experiment incubated at atmospheric conditions were within experimental error (34 F 4 and 36 F 8 Ag/ml, respectively). Thus, the absence of molecular oxygen did not affect the yield of the refolding reactions and is not playing a significant role in the in vitro refolding of tailspike. Therefore, tailspike chains do not require external agents to refold into the native conformation. 3.3. Reducing agents accelerate tailspike in vitro refolding kinetics at pH 7 An investigation of tailspike folding yield and kinetics in vivo revealed that altering the redox environment of the cytoplasm reduced the overall yield but did not affect the kinetics of trimer assembly [41]. By examining an isolated redox buffer system in vitro, we hoped to determine how redox reagents affected the rate of trimer formation, using both glutathione and dithiothreitol buffer systems. Fig. 4 shows a representative plot of tailspike trimer formation as a function of refolding time in four different refolding

Fig. 4. In vitro refolding kinetics are accelerated in the presence of reducing agents. Trimer formed as a function of time for tailspike refolding reactions in different redox buffers at pH 7.0: non-redox control (filled circles); 50 mM DTT (open circles); 50 mM DTTox (open diamonds) or 5 mM GSH (open squares). Tailspike trimer was denatured in 8 M urea at pH 3.0 and 1 mg/ml. Refolding was initiated by a 10-fold dilution into the indicated refolding buffer. Samples were taken at the indicated times, added to cold native sample buffer and quenched on wet ice and separated on nondenaturing gels. Trimer concentrations were determined by correlating silver-stained band densities to trimer standards. A representative experiment is shown. Inset highlights early time points ( < 60 min).

buffers at pH 7. Refolding reactions were performed as described in Materials and methods, with samples taken at designated times and refolding halted by the addition of sample buffer and placement on wet ice. Samples were separated through native gel electrophoresis and silverstained. Trimer concentrations at each time point were determined by measuring gel band densities and comparing to a trimer concentration standard curve. Trimer concentrations with time data were fit to a firstorder relationship to compare redox effects. The apparent trimer formation rates determined for each of the different redox buffer systems are presented in Table 1. This firstorder approximation best captures trimer formation rate data [37], although it does not provide a mechanistic description of the tailspike folding pathway. However, no other mathematical model, such as the sum of two first-order processes, improved the fit of the experimental data (data not shown). Therefore, the simplest approximation was employed here as a means to compare the effects of the different redox buffer conditions. Even though the overall yield was lower in all the redox buffers (Fig. 3B and data not shown), tailspike trimer formed faster in the presence of reducing agents than in the salt buffer or in more oxidizing conditions (Fig. 4 and Table 1). The addition of oxidizing agents seemed to have little effect on the rate of trimer formation,

B.L. Danek, A.S. Robinson / Biochimica et Biophysica Acta 1700 (2004) 105–116 Table 1 First-order rate constants for wild-type trimer formation, k ( 102 min k ( 102 min Control 50 mM DTT 25/25 mM DTT/DTTox 50 mM DTTox 5 mM GSH 5 mM GSSG

1 a

)

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3.4. Monothiol redox reagent traps productive tailspike monomer

1

)

10.2 F 1 18.2 F 3 19.1F2 12.6 F 2 25.0 F 2 NDb

a Tailspike was denatured at 1 mg/ml in 8M urea at pH 3.0. Refolding was initiated by rapid 10-fold dilution into the indicated redox buffer at 22 – 24 jC. Timed samples were separated on nondenaturing gels at 4 jC and silver-stained. Concentration versus time data was fit to a first-order relationship as described in Materials and methods. The error represents the standard deviation of fits of two to four independent experiments. b ND indicates that the kinetic constant was not determined due to absence of quantifiable trimer levels.

with these reactions proceeding at similar rates to the salt buffer reactions (Fig. 4 and Table 1). The appearance of trimer with time has a steeper slope in the early time points in the reactions containing reducing agents than the control reaction and that with oxidizing agent (Fig. 4, inset). The sharper slopes are reflected in the larger formation rates. However, the overall yields of trimer in the reactions containing redox reagents are lower than the control reaction (Figs. 3 and 4, >200 min).

As it became apparent that simple changes of the in vitro redox environment did not improve tailspike trimer refolding yields, we were interested in understanding the effect these redox reagents had on trimer formation and the folding intermediates. To this end, we separated endpoint refolding samples on nondenaturing gels, but overexposed the gels in silver staining to visualize any folding intermediates and aggregation species present. Refolding was performed at pH 8.5, as the largest effects of redox agents were observed under these conditions. Representative gels of refolding mixtures in glutathione and dithiothreitol buffers are shown in Fig. 5. Fig. 5A shows the folding intermediates and aggregation species populated at the end of a 6-h refolding reaction in various ratios of reduced to oxidized glutathione. At the most reducing conditions of 5 mM GSH (Fig. 5A, lane 1), there were no apparent folding intermediates still present after 6 h of folding. As the concentration of GSSG was increased, however, there was an increasing amount of monomeric folding intermediates (Fig. 5A, lanes 2 – 4). Upon inspection of the monomeric bands, two distinct populations were identified. As the GSSG content increased, the concentration of the monomer with the slower

Fig. 5. Oxidized glutathione traps tailspike chains in a mixed disulfide, preventing further assembly or aggregation. (A) Refolding samples from 100 Ag/ml reactions in various glutathione refolding buffers. Lane 1: 5 mM GSH; lane 2: 3.3 mM GSH and 1.7 mM GSSG; lane 3: 1.7 mM GSH and 3.3 mM GSSG; lane 4: 5 mM GSSG. Each reaction was allowed to refold for 6 h at 23 jC before separation on a nondenaturing gel. Bands were visualized through silver staining. The arrows show the two monomeric populations stalled in the presence of GSSG. (B) Refolding samples from 100 Ag/ml reactions in various dithiothreitol refolding buffers. Lane 1 has 50 mM DTT; lane 2: 33 mM DTT and 17 mM DTTox; lane 3: 17 mM DTT and 33 mM DTTox; lane 4: 50 mM DTTox. Each reaction was allowed to refold for 6 h at 23 jC before separation on a nondenaturing gel. Bands were visualized through silver staining. The arrow shows the faint amount of monomer still present in 50 mM DTTox.

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mobility increased, until, in the refolding buffer with 5 mM GSSG, only this larger monomer was apparent (Fig. 5A, lane 4). Glutathione has been shown to form relatively stable mixed disulfides with protein thiols [42,43]. It seemed likely that the slower mobility band in Fig. 5A was a tailspike polypeptide chain which had formed a complex with the oxidized glutathione. The association with glutathione prevented further productive assembly to trimer, but also did not permit the monomers to aggregate. While we are unable to completely rule out the possibility that intramolecular disulfides among the tailspike monomers are responsible for the mobility change, the discrete protein bands suggest a more homogeneous population that would result from a covalent modification with glutathione. Tailspike refolded in the presence of the dithiol redox agents did not have significant amounts of monomeric intermediates present after 6 h of folding at room temperature (Fig. 5B), although a small amount of monomer was apparent in the 50 mM DTTox sample (Fig. 5B, lane 4, arrow). The decrease in trimer formation in the oxidized dithiol buffers was coupled with an increase in aggregation that was not seen in the monothiol refolding buffer samples (Fig. 5A versus Fig. 5B, lanes 2– 4). Oxidized dithiothreitol most likely inhibited tailspike assembly at the monomer level by hindering early assembly reactions to form the disulfide-bonded monomer and dimer intermediates. Because these assembly reactions could not occur, few dimer and protrimer intermediates were observed. The build-up of monomer chains allowed for an increased partitioning to the aggregation pathway (Fig. 1B). 3.5. Redox interactions between chains are critical for trimer formation Refolding tailspike chains in a glutathione buffer system showed that oxidized glutathione trapped a monomer species that was not susceptible to aggregation. In order to test for the presence of a mixed disulfide with glutathione buffer components, tailspike refolding was initiated in three different buffers before spiking the reactions with DTT to rapidly change the redox environment. Tailspike refolding reactions in Tris refolding buffer containing either 5 mM GSH or 5 mM GSSG were incubated for 60 min at room temperature. Next, aliquots of each reaction were added to one of three buffers: Tris refolding buffer, 100 mM DTT, or 100 mM DTTox. These mixtures were incubated at room temperature for an additional 5 h to allow the reactions to go to completion. The refolding mixtures were separated on nondenaturing gels and silver-stained (Fig. 6A). Trimer yields were calculated as described in Materials and methods. Tailspike monomers that were trapped as a mixed disulfide with glutathione formed trimer upon addition of 100 mM DTT (Fig. 6A). Interestingly, almost 100% of all tailspike chains from trapped monomer formed trimer upon

Fig. 6. Monomeric intermediates trapped in a mixed disulfide dissociate to form trimer upon DTT addition. Tailspike trimer was denatured in 8 M urea at pH 3.0 before a rapid 10-fold dilution into Tris refolding buffer alone (Tris buffer), buffer with 5 mM GSH (GSH buffer), or buffer with 5 mM GSSG (GSSG buffer) at pH 8.5. After 60 min of refolding, 18-Al aliquots of each reaction were added to 2 Al of Tris refolding buffer ( + buffer, solid), 100 mM DTT ( + DTT, hatched) or 100 mM DTTox ( + DTTox, unshaded), buffered to pH 8.5. The newly diluted samples were allowed to continue refolding at room temperature for another 5 h before 10 Al of 3  native sample buffer were added to halt further folding. The samples were separated on nondenaturing gels and silver-stained. Trimer levels were quantified as described in Materials and methods. (A) Silver-stained native gel of redox spike refolding reactions after 6 total hours of folding. (B) Trimer yields from redox spike refolding reactions, determined by averaging three independent reaction yields. The error represents one standard deviation.

addition of 10 mM DTT, as the trimer yields were within experimental error of those in the Tris refolding buffer (Fig. 6B, Tris hatched versus GSSG hatched). This increase in trimer formation was not due to dilution of the refolding mixture, as samples added to buffer without any redox agents did not show any change in trimer yield (Fig. 6B, solid bars). Additionally, no trimer formation was seen in

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the GSSG refolding samples after addition of 100 mM DTTox (Fig. 6B, GSSG open bar). Therefore, it was the specific reducing action of DTT that allowed for productive trimer formation. This suggests that the GSSG trapped species is an on-pathway intermediate prevented from forming trimer by steric restraints from the protein thiolglutathione coupling.

4. Discussion The C-terminal cysteines have been shown to be critical for efficient assembly of tailspike monomers, with mutations at these residues resulting in stalled intermediates and decreased trimer yields [29]. Additionally, the protrimer [17], dimer [31] and monomer [29] folding intermediates have all been found to contain disulfide bonds. Other proteins with disulfide bonded folding intermediates have been found to fold optimally under certain redox buffer conditions [44 – 46], allowing for manipulation of external environment to favor certain intermediate conformations or pathways. Thus, we wanted to examine the effect of altering the refolding redox environment to determine whether this would facilitate tailspike trimer formation in wild-type tailspike. Typically, DTT is added to non-disulfide bonded proteins to reduce the opportunity for cysteine oxidation, with the view that this oxidation would enhance off-pathway reactions. Therefore, one might expect that reducing agents would only serve to increase the yield if no disulfides were formed. In this way, we hoped to provide insight into the role of extrinsic redox conditions for this novel assembly interaction. The expression of tailspike in cell lines with a more oxidizing cytoplasm resulted in reduced yields of native trimer (Fig. 2), with similar observed trimer formation rates for all cell lines. Thus, a simple change in redox environment did not alter tailspike folding efficiency in vivo, although other disulfide-bonded proteins have been shown to exhibit improved folding under these conditions [47]. Furthermore, in all in vitro buffer systems containing redox reagents (either favoring oxidizing or reducing conditions), the trimer yield was lower than in the control buffer that contained no redox agents (Fig. 3). Thus, altering the redox environment of in vitro refolding only interfered with or inhibited tailspike trimer formation. Here we found that the presence of oxidizing reagents had the greatest detrimental effect on trimer formation. This is in contrast to what was previously reported by Robinson and King [17], where increasing oxidizing conditions were shown to enhance trimer yields. Robinson and King performed refolding reactions at 30 Ag/ml, where productive folding predominates, while our current work was at 100 Ag/ ml, where there is significant competition with off-pathway aggregation and observed refolding rates and yields better reflect cellular results [29]. An additional difference is that Robinson and King [17] calculated yields after a 1-h reaction time, where kinetic effects contributed to the observations.

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Using the redox buffer system of Robinson and King (total DTT concentration of 10 mM) at 100 Ag/ml protein showed increases in the initial rate of trimer formation, but no significant changes in yield at each ratio investigated following a 5 –7-h reaction time in comparison to a nonredox control (data not shown). Even though the overall yield was lower in all the redox buffers (Fig. 3), tailspike trimer formed faster in the presence of reducing agents than in the salt buffer or in more oxidizing conditions at pH 7 (Fig. 4 and Table 1). This apparent contradiction may be resolved in part by considering how the refolding data were fit. Although the first-order approximation best captures trimer formation rate data [37], it does not reflect mechanistically the complexities of the entire tailspike folding pathway. The conversion of the protrimer to the native trimer has been postulated to be the rate-determining step for tailspike formation in vivo [38]. In vitro, the reduction of interchain disulfide bonds in the protrimer produces the native trimer [17]. Therefore, in the increased reducing environment of 50 mM DTT or 5 mM GSH, the faster kinetics likely resulted from more favorable conditions for the protrimer to trimer transition. Benton et al. [37] reported that protrimer concentrations were decreased in the presence of 5 mM GSH at pH 8.3. The faster formation rates observed here may reflect an increase in the protrimer to trimer conversion, but a decrease in the overall protrimer levels from reducing conditions results in the lower final trimer yields of the refolding reaction. The early folding and assembly reactions to form protrimer are not described by the first-order fit, as one would expect that only the rate-limiting step contributes greatly to the observed rates. Altering the pH of the refolding buffer had a dramatic effect on the resulting trimer yields. As the pH increased, the effect of oxidized glutathione became more pronounced, with no trimer production apparent at an 0:1 GSH/GSSG ratio at either pH 8 or pH 8.5. When trimer formation rates of the glutathione buffer systems were investigated at the elevated pHs, however, it was found that trimer formed faster in the more oxidizing reactions (data not shown). This is opposite of the results for the refolding reactions at pH 7, where the more reducing reactions formed trimer faster than the salt buffered reactions, but is consistent with the earlier Robinson and King results [17]. This indicates that the choice of buffer and refolding conditions can dramatically alter the observed trimer formation rate. Since trimer formation is not a true first-order process, final yields of trimer in all cases were lower in the presence of redox buffer than in non-redox buffer control reactions. Regardless of the conditions chosen, the interactions between cysteine residues and redox reagents are affecting the ability of the tailspike chains to associate into trimers. Preventing disulfide formation in fact leads to an enhanced aggregation propensity (Kim and Robinson, unpublished data). Thus, cysteine interactions and disulfide bonds in the

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folding intermediates are complex and sensitive to external manipulation. The differences between the tailspike trimer formation observed in the presence of dithiol redox agents and monothiol redox agents (Fig. 5) are likely due to the different nature of the thiol agents. Fig. 7 depicts a proposed scheme for tailspike association and the effects of redox agents in the refolding buffer. Because oxidized DTT inhibits trimer assembly, the pool of tailspike polypeptide chains in the refolding mixture is maintained at high concentrations. Because correct disulfide formation, and therefore assembly, is inhibited, these monomer chains are driven toward aggregation, as aggregating species increase in the refolding mixtures with high levels of DTTox. The assembly intermediates that are competent for aggregate formation have not been identified; however, it has been suggested that the presence of disulfide bonds in the productive folding intermediates, which are lacking the aggregation species [25], may be a distinguishing characteristic of the folding pathway [29]. Therefore, if DTTox prevents the correct disulfide bond formation, likely by allowing for indiscriminate disulfide pairing, these protein species would be available to participate in aggregation reactions. In contrast, glutathione appears to remove productive monomeric folding intermediates from the folding pathway, but prevent off-pathway associations. By blocking reactive sulfhydryl groups through a mixed disulfide, oxidized glutathione prevents on-pathway association but may not allow for conformational changes necessary for higher order aggregation. The tailspike chains trapped in this mixed

disulfide with glutathione retain their productive folding capability as nearly all of them form trimer upon dissociation of the disulfide bonds (Fig. 6). Benton et al. [37] showed mathematically that the protrimer is an obligate intermediate on the folding pathway by comparing models that included or excluded an onpathway trimeric intermediate. Additionally, they found that protrimer was diminished in the presence of reducing agent. Robinson and King [17] showed that the addition of reducing agent enabled the conversion of protrimer to native trimer. The question of whether or not tailspike assembly must transition through an obligate disulfidebonded intermediate has not been unambiguously addressed. However, the observation that the addition of redox agents greatly influences the yield and the rate of trimer association would be unexpected if a disulfidebonded intermediate was not a critical step. Taken together, the observations that protrimer formation is limiting in vivo [38], that a disulfide-bonded protrimer species is formed in vitro [17] and that oxidizing conditions lower trimer yields in vivo and in vitro (these results) suggest that sulfur chemistry is important in the efficient on-pathway assembly of the non-disulfide bonded tailspike trimer. The disulfide bond may help stabilize the position of the tailspike chains by preventing the shifting of hydrogen bonded-sheets necessary for oligomerization. The possible oxidizing and reducing agents in vitro are unknown, but the effects seen here with the trxB or Origamik cells suggest that the conventional cellular redox buffers are not involved in tailspike folding and assembly, at least not in any simple manner. The recent discovery of transient, non-native disulfide bonds in the mammalian viral protein, Vp1 [18], is of particular interest. The simian virus 40 complex is known to assemble in the cytoplasm, like tailspike. The fact that two viral proteins are found to transition through non-native disulfide-bonded intermediates during assembly in the highly reducing cytoplasm of their host cells could indicate the need for viral assembly processes that are independent of global redox environments. This independence may be manifested through the protein monomers acting as autonomous redox agents, facilitating disulfide shuffling reactions.

Acknowledgements

Fig. 7. Model for redox reactions and action of redox agents on the tailspike folding and assembly pathway. Addition of oxidized redox agents blocks productive association of tailspike trimer. The dithiol, DTTox, prohibits assembly and increases the partitioning to the aggregation pathway. The monothiol, GSSG, removes tailspike polypeptide chains from both the productive and aggregation pathway by trapping reactive sulfhydryl groups in a mixed disulfide.

We would like to thank H. F. Gilbert, C. Thorpe, and C. Robinson for advice and comments on the manuscript. C. Thorpe also graciously assisted with inert refolding experiments. We also thank J. King and C. Haase-Pettingell for assistance with the radioactive protein preparation; J. Kim for help in performing some refolding reaction, S. Bhatia for initial studies on DHB4 and AD494 expression; and S. Betts and R. Seckler for communication of unpublished results. This research was supported by funding from NIH R01 GM 60543.

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References [23] [1] C.B. Anfinsen, Principles that govern the folding of protein chains, Science 181 (96) (1973) 223 – 230. [2] J.R. Huth, et al., Domain-dependent protein folding is indicated by the intracellular kinetics of disulfide bond formation of human chorionic gonadotropin beta subunit, J. Biol. Chem. 267 (1992) 21396 – 21403. [3] Z.-Y. Guo, L. Shen, Y.-M. Feng, The different folding behavior of insulin and insulin-like growth factor 1 is mainly controlled by their B-Chain/Domain, Biochemistry 41 (2002) 1556 – 1567. [4] T.E. Creighton, Protein folding pathways determined using disulphide bonds, BioEssays 14 (3) (1992) 195 – 199. [5] K. Saito, E. Welker, H.A. Scheraga, Folding of a disulfide-bonded protein species with free thiol(s): competition between conformational folding and disulfide reshuffling in an intermediate of bovine pancreatic ribonuclease A, Biochemistry 40 (2001) 15002 – 15008. [6] F. Chiti, et al., Kinetic partitioning of protein folding and aggregation, Nat. Struct. Biol. 9 (2) (2002) 137 – 143. [7] I. Braakman, E. van Anken, Folding of viral envelope glycoproteins in the endoplasmic reticulum, Traffic 1 (2000) 533 – 539. [8] J.J. Pitt, E. Da Silva, J.J. Gorman, Determination of the disulfide bond arrangement of Newcastle disease virus hemagglutinin neuraminidase, J. Biol. Chem. 274 (9) (2000) 6469 – 6478. [9] P.S. Reddy, R.B. Corley, Assembly, sorting and exit of oligomeric proteins from the endoplasmic reticulum, BioEssays 20 (1998) 546 – 554. [10] D.P. Goldenberg, Kinetic analysis of the folding and unfolding of a mutant form of bovine pancreatic trypsin inhibitor lacking the cysteine-14 and cysteine-38 Thiols, Biochemistry 27 (1988) 2481 – 2489. [11] T.E. Creighton, Conformation restrictions on the pathway of folding and unfolding of the pancreatic trypsin inhibitor, J. Mol. Biol. 113 (1977) 275 – 293. [12] T.E. Creighton, N.J. Darby, J. Kemmink, The roles of partly folded intermediates in protein folding, FASEB J. 10 (1996) 110 – 118. [13] M.-C. Song, H.A. Scheraga, Formation of native structure by intermolecular thiol-disulfide exchange reactions without oxidants in the folding of bovine pancreatic ribonuclease A, FEBS 471 (2000) 177 – 181. [14] S.J. Milner, et al., Probing the disulfide folding pathway of insulinlike growth factor-I, Biotech. Bioeng. 62 (1999) 693 – 703. [15] Y.H. Zhang, et al., Conformational analysis of intermediates involved in the in vitro folding pathways of recombinant human macrophage colony stimulating factor beta by sulfhydryl group trapping and hydrogen/deuterium pulsed labeling, Biochemistry 41 (2002) 15495 – 15504. [16] P. Pattanaik, et al., Refolding of native and recombinant chicken riboflavin carrier (or binding) protein: evidence for the formation of non-native intermediates during the generation of active protein, Eur. J. Biochem. 258 (1998) 411 – 418. [17] A.S. Robinson, J.A. King, Disulphide-bonded intermediate on the folding and assembly pathway of a non-disulphide bonded protein, Nat. Struct. Biol. 4 (6) (1997) 350 – 355. [18] P.P. Li, et al., Formation of transitory intrachain and interchain disulfide bonds accompanies the folding and oligomerization of simian virus 40 Vp1 in the cytoplasm, Proc. Natl. Acad. Sci. 99 (3) (2002) 1353 – 1358. [19] R.T. Sauer, et al., Phage P22 tail protein: gene and amino acid sequence, Biochemistry 21 (1982) 5811 – 5815. [20] D. Goldenberg, P. Berget, J.A. King, Maturation of the tailspike endorhamnosidase of Salmonella phage P22, J. Biol. Chem. 257 (1982) 7864 – 7871. [21] J.M. Sturtevant, et al., Thermostability of temperature-sensitive folding mutants of the P22 Tailspike Protein, J. Biol. Chem. 264 (18) (1989) 10693 – 10698. [22] S. Steinbacher, et al., Crystal structure of P22 tailspike protein:

[24] [25]

[26]

[27]

[28]

[29]

[30]

[31]

[32] [33] [34]

[35]

[36]

[37]

[38]

[39]

[40]

[41]

[42]

[43]

[44]

[45]

115

interdigitated subunits in a thermostable trimer, Science 265 (1994) 383 – 386. A. Fuchs, C. Seiderer, R. Seckler, In Vitro folding pathway of page P22 Tailspike Protein, Biochemistry 30 (1991) 6598 – 6604. M. Danner, R. Seckler, Mechanism of phage P22 tailspike protein folding mutations, Protein Sci. 2 (1993) 1869 – 1881. M.A. Speed, D.I.C. Wang, J.A. King, Multimeric intermediates in the pathway to the aggregated inclusion body state for P22 tailspike polypeptide chains, Protein Sci. 4 (1995) 900 – 908. B.G. Lefebvre, A.S. Robinson, Tailspike intermediates generated from pressure-treated aggregates form native trimer faster than freshly denatured protein, Biotech. Bioeng. 82 (2003) 595 – 604. D. Sargent, et al., Secondary structure and thermostability of the phage P22 tailspike: analysis of Raman spectroscopy of the wild type protein and a temperature-sensitive folding mutant, J. Mol. Biol. 199 (1988) 491 – 502. S. Sather, J.A. King, Intracellular trapping of a cytoplasmic folding intermediate of the phage P22 tailspike using iodoacetamide, J. Biol. Chem. 269 (41) (1994) 25268 – 25276. B.L. Danek, A.S. Robinson, Non-native interactions between cysteines direct productive assembly of P22 tailspike protein, Biophys. J. 85 (2003) 3237 – 3247. C. Haase-Pettingell, et al., Role for cysteine residues in the in vivo folding and assembly of the phage P22 tailspike, Protein Sci. 10 (2001) 397 – 410. S. Betts, J.A. King, Cold rescue of the thermolabile tailspike intermediate at the junction between productive folding and off-pathway aggregation, Protein Sci. 7 (1998) 1516 – 1523. A.I. Derman, et al., Mutations that allow disulfide formation in the cytoplasm of Escherichia coli, Science 262 (1993) 1744 – 1747. J.C. Weaver, Electroporation: a general phenomenon for manipulating cells and tissues, J. Cell. Biochem. 51 (4) (1993) 426 – 435. J. Sambrook, E.F. Fritsch, T. Maniatis, Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, 1989. J.A. King, M.-H. Yu. Mutational analysis of protein folding pathway using the P22 tailspike endorhamnosidase, in Methods in Enzymology. In: C.W.H. Hirs, S. Timashiff, 1986, Academic Press, New York, pp. 250 – 266. S.K. Sather, J. King, Intracellular trapping of a cytoplasmic folding intermediate of the phage P22 tailspike using iodoacetamide, J. Biol. Chem. 269 (41) (1994) 25268 – 25276. C.B. Benton, J.A. King, P.L. Clark, Characterization of the protrimer intermediate in the folding pathway of the interdigitated B-helix tailspike protein, Biochemistry 41 (16) (2002) 5093 – 5103. D. Goldenberg, J.A. King, Trimeric intermediate in the in vivo folding and subunit assembly of the tail spike endorhamnosidase of bacteriophage P22, Proc. Natl. Acad. Sci. U. S. A. 79 (1982) 3403 – 3407. D. Goldenberg, D.H. Smith, J.A. King, Genetic analysis of the folding pathway for the tail spike phage P22, Proc. Natl. Acad. Sci. U. S. A. 80 (23) (1983) 7060 – 7064. H.F. Gilbert, Molecular and cellular aspects of thiol-disulfide exchange, in: A. Meister (Ed.), Advances in Enzymology, Wiley, New York, 1990, pp. 169 – 172. S.K. Bhatia, The Role of Cysteines in the Folding Pathway of P22 Tailspike Protein, in Chemical Engineering, University of Delaware, Newark, MS Thesis, 1999. K.-H.W. Lau, J.A. Thomas, Specific mixed disulfide formation with purified bovine cardiac glycogen synthase I and Glutathione, J. Biol. Chem. 258 (1983) 2321 – 2326. D.W. Walters, H.F. Gilbert, Thiol/Disulfide redox equilibrium and kinetic behavior of chicken liver fatty acid synthase, J. Biol. Chem. 261 (1986) 13135 – 13143. Y. Kawata, et al., Chaperonin GroE-facilitated refolding of disulfidebonded and reduced Taka-amylase A from Aspergillus oryzae, Protein Eng. 11 (12) (1998) 1293 – 1298. K. Maskos, M. Huber-Wunderlich, R. Glockshuber, DsbA and DsbC-

116

B.L. Danek, A.S. Robinson / Biochimica et Biophysica Acta 1700 (2004) 105–116

catalyzed oxidative folding of proteins with complex disulfide bridge patterns in vitro and in vivo, J. Mol. Biol. 325 (3) (2003) 495 – 513. [46] T.E. Creighton, Role of the environment in the refolding of reduced pancreatic trypsin inhibitor, J. Mol. Biol. 144 (1980) 521 – 550. [47] P.H. Bessette, et al., Efficient folding of proteins with multiple disul-

fide bonds in the Escherichia coli cytoplasm, Proc. Natl. Acad. Sci. 96 (1999) 13703 – 13708. [48] D.P. Goldenberg, D.H. Smith, J. King, Genetic analysis of the folding pathway For the tail spike phage P22, Proc. Natl. Acad. Sci. U. S. A. 80 (23) (1983) 7060 – 7064.