Involvement of surface cysteines in activity and multimer formation of thimet oligopeptidase

J.A. Sigman1, M.L. Sharky1, S.T. Walsh1, A. Pabon2, M.J. Glucksman2 and A.J. Wolfson1,3

1Department of Chemistry, Wellesley College, Wellesley, MA 02481 and 2FUHS/Chicago Medical School, Midwest Proteome Center and Department of Biochemistry and Molecular Biology, Chicago, IL 60064, USA

3 To whom correspondence should be addressed. e-mail: awolfson{at}wellesley.edu


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Thimet oligopeptidase is a metalloenzyme involved in regulating neuropeptide processing. Three cysteine residues (246, 248, 253) are known to be involved in thiol activation of the enzyme. In contrast to the wild-type enzyme, the triple mutant (C246S/C248S/C253S) displays increased activity in the absence of dithiothreitol. Dimers, purportedly formed through cysteines 246, 248 and 253, have been thought to be inactive. However, analysis of the triple mutant by native gel electrophoresis reveals the existence of dimers and multimers, implying that oligomer formation is mediated by other cysteines, probably on the surface, and that some of these forms are enzymatically active. Isolation and characterization of iodoacetate-modified monomers and dimers of the triple mutant revealed that, indeed, certain dimeric forms of the enzyme are still fully active, whereas others show reduced activity. Cysteine residues potentially involved in dimerization were identified by modeling of thimet oliogopeptidase to its homolog, neurolysin. Five mutants were constructed; all contained the triple mutation C246S/C248S/C253S and additional substitutions. Substitutions at C46 or C682 and C687 prevented multimer formation and inhibited dimer formation. The C46S mutant had enzymatic activity comparable to the parent triple mutant, whereas that of C682S/C687S was reduced. Thus, the location of intermolecular disulfide bonds, rather than their existence per se, is relevant to activity. Dimerization close to the N-terminus is detrimental to activity, whereas dimerization near the C-terminus has little effect. Altering disulfide bond formation is a potential regulatory factor in the cell owing to the varying oxidation states in subcellular compartments and the different compartmental locations and functions of the enzyme.

Keywords: dimerization/disulfide bonds/homology model/thimet oligopeptidase


    Introduction
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 Abstract
 Introduction
 Materials and methods
 Results
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 References
 
Endopeptidase EC 3.4.24.15 (thimet oligopeptidase, TOP) is a Zn(II)-dependent enzyme that preferentially cleaves peptides 5–17 amino acids in length (Barrett et al., 1995Go; Camargo et al., 1997Go; Oliveira et al., 2001Go). Increased interest in the enzyme has been generated from the findings that it cleaves several important physiological neuropeptides, including bradykinin, neurotensin and gonadotropin-releasing hormone in the extracellular milieu (Orlowski et al., 1983Go; Camargo et al., 1987Go, 1997; Dahms and Mentlein, 1992Go; Lew et al., 1995Go; Vincent et al., 1997Go). TOP has been shown to posses ß-secretase activity in amyloid precursor processing (Koike et al., 1999Go) and to be a necessary element in the pathway leading to the degradation of the ß-amyloid precursor of Alzheimer’s disease (Yamin et al., 1999Go). An intracellular (cytosolic) role of the enzyme appears to be the ability to bind and degrade selectively MHC class I oligopeptides involved in subsequent antigen presentation (Silva et al., 1999Go).

Based on the primary sequence, the enzyme belongs to the family of zinc metalloendopeptidases that includes neurolysin, neutral endopeptidase and angiotensin-converting enzyme and is evolutionarily related to thermolysin, bacterial elastase and neutral protease (Glucksman et al., 1992Go; Pierotti et al., 1990Go; Barrett et al., 1995Go) TOP is a 78 kDa enzyme and is unique among other thermolysin-like metallopeptidases in its activation by thiol reducing agents (Tisljar and Barrett, 1990Go; Shrimpton et al., 1997Go; Smith et al., 2000Go). Although some classes of proteins are activated by dimerization, TOP is unusual in that reduction of multimers, rather than their formation, promotes its activation. Despite many biochemical studies, there is a paucity of structural information about TOP and only recently has the structure of neurolysin, its closest homolog, sharing >60% sequence identity (Dauch et al., 1995Go), been solved by X-ray diffraction (Brown et al., 2001Go).

This paper describes the activity of TOP modulated by cysteines as revealed by chemical modification and mutagenesis of cysteine residues. Presented is a reproducible method for cysteine modification of both monomeric and dimeric forms of TOP without loss of activity. The experimental results support predictions made as a result of homology modeling of TOP using neurolysin as the template structure. The model and experimental data provide further insight into thiol activation of TOP.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Glutathione Sepharose, Sephacryl S-200 and PD-10 columns were obtained from Amersham Pharmacia Biotech (Piscataway, NJ). Protein gels were 12% Tris–glycine gels from Invitrogen (Carlsbad, CA) and stained with Gelcode blue stain from Pierce (Rockford, IL). The 7-methoxycoumarin-4-acetyl-Pro–Leu–Gly–Pro–Lys-dinitrophenol (MCA) fluorescent substrate was obtained from Bachem (King of Prussia, PA). Centricon YM-50 concentrators were purchased from Millipore (Bedford, MA). Tris(2-carboxyethyl)phosphine (TCEP) was obtained from Pierce. All other reagents were purchased from Sigma Chemical (St. Louis, MO).

Site-directed mutagenesis and protein expression

Double-stranded site-directed mutagenesis of TOP was performed as described previously (Cummins et al., 1999Go) with the following modifications. The plasmid vector GEXTCys, encoding the triple cysteine mutant (tCys) (C246S/C248S/C253S), was utilized as the template. Prokaryotic codon usage rules were used to obviate the use of rare codons. Protein expression and characterization were carried out as described previously (Cummins et al., 1999Go). At least two independent preparations of wild-type protein and each mutant were purified, with similar homogeneity and yields.

Recombinant thimet oligopeptidase (TOP)

Cell cultures containing the over-expressed recombinant TOP and all mutants were grown, expressed and purified utilizing the glutathione-S-transferase ‘tag’ as previously described (Shrimpton et al., 1997Go). A 40 ml suspension of cells in 100 mM Tris buffer, pH 7.4, with 0.3 mM dithiothreitol (DTT) was lysed by passing twice through a French press cell. After the initial purification with glutathione–Sepharose 4B and thrombin cleavage of the glutathione S-transferase ‘tag’, the enzyme was loaded on to a size-exclusion column packed with Sephacryl S-200 and eluted at 0.4 ml/min with 25 mM Tris buffer, pH 7.8, containing 125 mM NaCl. The purity of the enzyme was verified using SDS–PAGE under reducing conditions. The enzyme concentration was determined using the molar extinction coefficient {epsilon}280 = 73.11 mM–1 cm–1 calculated based on the amino acid content of the protein (Gill and Von Hippel, 1989Go).

Homology modeling

The three-dimensional structure of TOP was modeled to the fold of neurolysin using atomic coordinates (Protein Data Bank, 1I1I.pdb) from the recently determined X-ray crystal structure of neurolysin (Brown et al., 2001Go). The initial model was generated in SWISS PDBviewer version 3.72 (Peitsch, 1995Go; Guex and Peitsch, 1997Go; Guex et al., 1999Go) using the program Magic-Fit in the viewer software suite. Missing from the model are the first 11 N-terminal residues of TOP, KPPAACAGDVV, owing to lack of homology and disorder in the reported structure of neurolysin. This was also the case for the last 10 C-terminal residues, QVEGCEPPAC. The refined fitting procedure of the initial TOP model and the neurolysin template utilized SWISS-MODEL (version 3.5) (Guex and Peitsch, 1997Go). The quality of the model was evaluated both by using the suite of programs within the SWISS PDB viewer and by visual inspection. A 100-cycle steepest descents energy minimization of the returned model was performed using the GROMOS 43B1 force fields. The model generated was consistent with that presented by Ray et al. (Ray et al., 2002Go).

Kinetic assays

All kinetic assays were performed using a Cary Eclipse spectrofluorimeter. Cleavage of the fluorogenic MCA substrate (Wolfson et al., 1996Go) was monitored by the increase in emission at 400 nm over time using {lambda}ex = 325 nm. Less than 10% of the substrate was consumed. Assays were performed in duplicate at 37°C with 25 mM Tris buffer, pH 7.8, adjusted to a conductivity of 15 mS/cm2 with NaCl. DTT, when present, was 0.3 mM and TCEP 0.5 mM. MCA substrate concentration was calculated based on the molar extinction coefficient {epsilon}365 (17.3 mM–1 cm–1) of the 2,4-dinitrophenol. The change in fluorescence intensity over time was converted to rate of product formation using a calibration curve calculated for the peptide product, 7-methoxycoumarin-4-acetyl-Pro–Leu-OH. The pseudo-first-order rate constant, k (s–1), of enzyme activity was determined under conditions in which [E] << [S] << Km and calculated from the slope of the linear least-squares fit to the graph of initial substrate concentration (µM) versus rate of product formation (µM/s). Each plot was defined by 4–5 points and had an r2 value of at least 0.99. Kinetic parameters were determined using a hyperbolic fit to the plot of substrate concentration (µM) versus rate of product formation (µM/s) and by Eadie–Hofstee plots under conditions in which [S] is above and below Km.

Modification of cysteine residues

The modification procedure is a variation of the protocol followed by Caccia et al. (Caccia et al., 1992Go). TOP in 25 mM Tris buffer, pH 7.8, was incubated at room temperature for 30 min to allow partial dimerization to occur. If only monomers were desired, the protein was instead incubated for 1 h in buffer containing 0.3 mM DTT. A PD-10 column, for iodoacetic acid–buffer exchange, was equilibrated with a 25 mM phosphate buffer, 5 mM EDTA, pH 8.0, containing a 400-fold molar excess of iodoacetic acid relative to the protein. After eluting from the column, the protein, in iodoacetic acid–buffer, was incubated for 1 h at room temperature to allow the modification reaction to continue. Upon completion of the reaction, the protein was loaded on to the Sephacryl S-200 gel filtration column described above, equilibrated with 25 mM Tris buffer, pH 7.8, containing 125 mM NaCl, to separate the monomers and dimers.


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 Results
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 References
 
Location of cysteine residues

Of the differences between TOP and neurolysin, one of the more important is the location of Cys residues in the two enzymes. Previous work has shown that TOP is activated by low concentrations (0.1–1.0 mM) of thiol-based reductants such as DTT and 2-mercaptoethanol (Tisljar and Barrett, 1990Go), whereas there is no thiol activation of neurolysin (Serizawa et al., 1995Go). Subsequent research found that low activity was also associated with protein dimer formation (Shrimpton et al., 1997Go). Hence it seemed likely that some Cys residues near the active site might be responsible for forming protein dimers that block substrate access to the active site. Of the 14 Cys residues in TOP, seven (Cys7, Cys18, Cys46, Cys246, Cys253, Cys682 and Cys687) are not conserved in neurolysin. Shown in Figure 1 is a surface representation of the TOP model with the sulfur of Cys residues highlighted in yellow. None of the surface residues is predicted to be within the large cleft that leads to the active site; however, it is apparent that several non-conserved Cys residues could have side chains oriented towards the surface and exposed to solvent with the potential for intermolecular disulfide bonding. The N-terminal residue, Cys7, and two C-terminal residues, Cys682 and Cys687, could not be included in the model owing to disorder in the corresponding regions of the crystallized enzyme neurolysin, but it is likely that these residues are also on or near the surface. Hence any of these residues could be responsible for protein dimerization. Two of these have already been well studied, Cys246 and Cys 253. Mutation of these residues to Ser has been shown to increase the activity of the protein without the need for thiol reducing agents (Shrimpton et al., 1997Go). As has been noted by Ray et al. (Ray et al., 2002Go) and also apparent in Figure 1, these cysteine residues are not predicted to be near the active site [both are >30 Å from the active site Zn(II)] but at the opening of the large cleft that allows substrate access to the active site (Brown et al., 2001Go).



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Fig. 1. Surface representation of the TOP model. Several of the Cys sulfur atoms, shown in yellow, are exposed to the surface and serve as possible points of protein dimerization. The positions of the triple Cys implicated in loss of thiol activation, Cys245, Cys247 and Cys253, are shown just above the entrance to the active-site cleft. At the bottom of the active-site cleft is the catalytyic Zn(II), shown in green.

 
However, native PAGE indicates that even the triple mutant (C246S/C248S/C253S), tCys, forms dimers and multimers (Figure 2, lane 2). This observation, together with the model, suggests that not all dimerization processes would necessarily lead to decreased activity. Other Cys residues on the opposing face of the protein may be available for, and participate in, intermolecular disulfide bond formation without occluding the active site (Figure 1).



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Fig. 2. Native polyacrylamide gel of wild-type TOP and engineered forms. The amount of enzyme in each lane was 10 pmol. Lane 1, triple mutant (tCys); lane 2, tCys with C7S; lane 3, tCys with C46S; lane 4, tCys with C682S; lane 5, tCys with C687S; lane 6, tCys with C682S/C687S; lane 7, wt+3; lane 8, tCys+3; lane 9, wild-type TOP.

 
Chemical modification studies

Monomers and dimers of tCys were isolated for activity assays. To accomplish this we allowed the mutant partially to dimerize and then chemically modified the remaining Cys residues with iodoacetate in the mixture to prevent further dimerization. The monomers and dimers were then separated by gel filtration chromatography as described above. The dimerization is reversible with the addition of DTT. Figure 3 shows the rate of MCA substrate cleavage by wild-type TOP, tCys and the iodoacetate-modified monomers and dimers of the modified tCys. The slopes of the initial rate plots of the monomeric and dimeric forms of the triple mutant are nearly identical, indicating that, although the enzyme is forming dimers through surface cysteines, dimerization of this form does not inhibit activity.



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Fig. 3. Initial rate plots of substrate cleavage by tCys, circles; iodoacetic acid modified tCys monomer, squares; iodoacetic acid modified tCys dimer, triangles; wild-type (wt) TOP, diamonds. All reactions were performed in 25 mM Tris, pH 7.8, without DTT, except the wt buffer, which contained 0.3 mM DTT. As shown, the modification only slightly affected the rate of substrate cleavage overall and the rate of cleavage by the dimer is identical with that by the monomer. Lines represent linear least-squares fits of the data points.

 
Mutagenesis

Five site-directed mutant forms of tCys were constructed, converting one or two additional cysteines to serines. In addition to these single and double mutants, two triple mutants were created, with C46, C682 and C687 converted to serines, one with tCys (tCys+3) and one with wild-type (wt+3) as scaffold. A representative native electropherogram is shown in Figure 2. As can be seen, the degree of dimerization and multimerization varied, depending on the location of the substitution. Two mutants, those with the substitution at position 46 (lane 3) or both 682/687 (lane 6), were consistently observed to dimerize to only a limited extent and not to form higher order multimers, whereas multimers and dimers were always present for other forms with single mutations. Mutants with substitutions at either 682 or 687 formed many dimers and higher order multimers. The mutant form wt+3 dimerized to only a limited extent (lane 7), while only the monomeric form for tCys+3 was apparent (lane 8).

Enzymatic activity for all forms was determined (Table I). All of the mutant forms except that based on the wild-type were slightly more active in the absence of the thiol reagent DTT than in its presence (data not shown). Note that, despite their similar dimerization profiles, the C46S form was more active than C682S/C687S. Monomers and dimers of C46S and C682S/C687S were separated by gel filtration, as described for chemically modified TOP. Monomers in both cases had comparable activities. Dimers of C46S were as active as monomers, but dimers of C682S/C687S were considerably less active (Figure 4).


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Table I. Activity of various Cys->Ser mutants of TOP
 


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Fig. 4. Comparison of Vmax/Km for iodoacetate-modified monomers and dimers. The two enzymes, tCys with C46S and tCys with C682S/C687S, were modified with iodoacetate and separated by gel filtration as described in the text. Enzymatic activity was determined in 25 mM Tris buffer with 10 mM CaCl2 at pH 7.3 over a range of substrate concentrations spanning Km. Dark bars, iodoacetate-modified mixture; striped bars, monomers; grey bars, dimers.

 
The triply substituted forms wt+3 and tCys+3 displayed activities comparable to their parent forms; that is, wt+3 had low activity in the absence of DTT and higher activity in its presence, whereas tCys+3 was fully active in the absence of DTT and slightly inhibited when the reductant was added (Table I). The phosphine reducing agent TCEP (Getz et al., 1999Go; Han et al., 1999Go) increased the activity of wt+3 above the level attained with DTT.


    Discussion
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 Abstract
 Introduction
 Materials and methods
 Results
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 References
 
A unusual aspect of TOP is its thiol activation. The enzyme has been shown to exhibit a >8-fold increase in activity in the presence of reducing agents such as DTT, ß-mercaptoethanol and glutathione. Dimer formation via intermolecular disulfide bridges involving Cys246, -248 and -253 was proposed to block substrate access to the active site (Shrimpton et al., 1997Go). A triple mutation of these Cys residues to Ser abolished the thiol dependence of the enzyme. As shown in Figure 1, this cluster of three Cys residues is predicted to be at the protein surface in domain I. Although these residues are not near the active site, an intermolecular dimerization involving any one of these residues might block the opening to the substrate access channel. Another possibility is that suggested by Ray et al. (Ray et al., 2002Go), that dimerization either induces a conformational change that prevents substrate binding or prevents a conformational change required for activity.

Early work suggested that Cys483, the residue closest to the active site zinc, might be responsible for the thiol dependence (Pierotti et al., 1990Go). However, mutating this residue had no effect on the thiol dependence of the enzyme (Shrimpton et al., 1997Go) and in fact this residue is conserved in neurolysin, which does not exhibit activation by thiol reagents (Serizawa et al., 1995Go). The model, suggesting that the side chain of Cys483 is not available for disulfide bond formation, is consistent with these experimental observations.

The results of iodoacetate modification are of interest in the light of Ray et al.’s prediction of a negatively charged binding site in TOP (Ray et al., 2002Go). The larger, positively charged thiol reagent NEM has been shown to abolish activity in this enzyme and we have also observed this to be the case under the same modification conditions used with iodoacetate (data not shown). However, the negatively charged iodoacetate may react more slowly and only with surface residues, under the limiting conditions used for this study.

Cys248, a residue conserved in neurolysin, is also implicated in protein dimerization. This residue appears to be in a well-conserved region and an electronic environment similar to that of Cys249 in neurolysin. Involvement of Cys248 in disulfide bridges is probably dependent on the presence of other surface residues found in TOP but not in neurolysin. For instance, a disulfide bond formed between Cys46 and 248, which would block the active site of one TOP molecule, is not possible in neurolysin. This is supported by the fact that TOP readily forms multimers in solution. Even the triple Cys mutant, with decreased apparent molecular mass and radius compared with wild-type, showed a significant drop in these parameters when DTT was added (Shrimpton et al., 1997Go), indicating that some dimers and oligomers were present in the absence of reductant. Evidence is presented here that additional Cys residues in TOP are involved in protein dimerization. The gel in Figure 2 indicates the degree of dimerization and multimerization dependent upon the availability of various Cys residues. The triple Cys mutant, shown in lane 1 of Figure 2, is primarily in the dimeric form with some multimer formation. The additional change of C7, C682 or C867 with Ser onto the tCys mutant scaffold resulted in proteins with the same or increased dimer and multimer formation compared with the tCys alone (Figure 2, lanes 2, 4 and 5, respectively). However, neither the C46S nor the C682S/687S double mutant (Figure 2, lanes 3 and 6) was observed to form multimers. Based on this result, C46 seems to be the most likely position, followed by C682 and 687, for intermolecular/interdisulfide cross-links. Mutation of either C682 or C687 appears to increase the tendency to form dimers and multimers. We speculate that an intramolecular disulfide between these two residues exists in the natural form. When one is replaced by serine, the other is freer to participate in intermolecular dimers. The gel results suggest that the tCys form has two sites that can form intermolecular disulfide bonds, one near the N-terminus (C46) and one near the C-terminus (C682 or C687). When both are reactive, dimers form and consequently multimers. Mutation of either site leads to forms that can only dimerize; mutation of both (in the tCys+3 form) precludes dimerization. Hence the experimental results and self-consistent support from the model imply that intermolecular disulfide bond formation in TOP is not simply a ‘head-to-head’ process. If that were the case, there would be no higher-order forms than dimers.

We have isolated monomers and dimers of tCys, C46S and C682S/C687S through specific modification of cysteine residues using iodoacetate. As shown in Figure 3, the dimer form of the triple Cys mutant exhibits the same activity towards the quenched fluorescent substrate as monomers. Mutants with substitutions of Ser for Cys at position 46 also form dimers with activity comparable to monomers (Figure 4).

In contrast to C46, mutants with substitutions at positions 682 and 687 form dimers with lower activity. These results suggest that dimerization through cysteines 682 and 687 on the C-terminal face of TOP does not impede substrate access, whereas dimerization on the N-terminal face leads to lower activity. This result is supported by the model, which suggests that the C-terminus should lie on the side of the enzyme opposite to the active site cleft.

Furthermore, the form with triple substitution in the wild-type background (wt+3) dimerizes to only a small extent while displaying low enzymatic activity in the absence of DTT. This result might suggest that thiol activation is due to disruption of intramolecular disulfides rather than intermolecular disulfide bonds. However, loss of activity due to the formation of intramolecular disulfides between C246, C248 or C253 cannot explain the original observation by Shrimpton et al. of increased activity with stepwise mutations of those three residues. We, therefore, favor the interpretation that disulfide formation among the original set of cysteines (Shrimpton et al., 1997Go) promotes a conformational change, as proposed by Ray et al. (Ray et al., 2002Go). Those authors concluded that dimerization does not sterically block substrate access based on the assumption that only cysteines 246, 248 and 253 were involved in intermolecular disulfide bonds. Our results demonstrate that other cysteine residues are involved in dimer formation. The proposed conformational change may be triggered by disulfides through cysteines other than these three. The presence of both inter- and intramolecular disulfides bonds cannot be ruled out by the model or experimental results. A conformational change involved in catalytic activity is further supported by our data relating to the pH dependence of TOP (Sigman et al., 2003Go).

High concentrations (above 1 mM) of DTT inhibit TOP (Lew et al., 1995Go) either by disrupting key intramolecular disulfide bridges (Morales and Woessner, 1977Go) or by binding to the catalytic zinc (Barrett and Brown, 1990Go). Since we observed inhibition by DTT even for the wt+3 form, the inhibitory role of DTT probably results from its binding to zinc. This is supported by the observed effect of the reducing agent TCEP on TOP activity. The phosphorus in TCEP is sterically hindered and therefore not expected to inhibit activity by binding to zinc and it does, indeed, increase activity for the wt+3 above the levels attained with DTT.

Disulfide bond formation or dimerization could have physiological significance since there are several different roles for TOP assigned to different subcellular compartments as well as outside the cell. TOP is found predominantly in the cytosol (a relatively reductive environment) (Crack et al., 1999Go). Considering the similarities between the neural and immune systems, it is of considerable interest that the neuropeptide processing enzyme TOP acts as the primary peptidase that degrades MHC I peptides in the cytosol of immune antigen presenting cells (Silva et al., 1999Go; Saric et al., 2001Go).

Outside of the cell in the extracellular milieu, TOP has its most classically described role: in physiologically relevant neuropeptide processing on the external face of the pre-synaptic space where substrate binding and specific activity of the enzyme are exquisitely regulated (Ferro et al., 1999Go). Very close to the surface of the presynapse, where neurotransmitters are released to interact with their cognate receptors, regulation of TOP would occur by local fluxes in redox potential. Lastly, in the nucleus, a fairly oxidative compartment, TOP is relatively abundant (Crack et al., 1999Go) with no known substrates, but a targeted nuclear localization signal (Resendes et al., 1999Go; Fu et al., 2001Go). At present, there is no purported role for nuclear TOP, but changes in cell cycle cause translocation of TOP from the nucleus to the cytosol (unpublished observations).

Endogenous reducing agents such as glutathione and thioredoxin are present inside and outside the cell and can presumably regulate the multimerization of the endogenous enzyme (Smith et al., 2000Go). Alternatively, surface cysteines remote from the active site may play a role in membrane association (Crack et al., 1999Go).

The results presented here extend and elaborate on the earlier findings of Shrimpton et al. (Shrimpton et al., 1997Go) and provide experimental verification for some of the predictions of Ray et al. (Ray et al., 2002Go). The role of surface cysteines in modulating TOP activity helps to explain the apparent paradox that the three cysteine residues (246, 248, 253) required for thiol activation are not in the active site, nor would dimers involving these residues necessarily block substrate access. The explanation seems to be that dimerization is not equivalent to inactivation. Our results suggest that in wild-type TOP there are three regions for potential disulfide bond formation: at the entrance to the active-site cleft (C246/C248/C253), near the N-terminus (C46) and near the C-terminus (C682/C687). When disulfides involving C246/C248/C253 form, there is a decrease of 8–15-fold in activity, when involving C46, a decrease of ~2-fold, and there is no loss in activity when disulfides involving C682/C687 form.


    Acknowledgements
 
This work was supported by a Howard Hughes Medical Institute Undergraduate Science Education Program Grant and a National Science Foundation Research Experiences for Undergraduates Award to Wellesley College, by the NIH/National Institute of Neurological Disorders and Stroke grant no. NS 39892 (M.J.G.) and by a Camille and Henry Dreyfus Scholar/Fellow Award (A.J.W.).


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Barrett,A.J. and Brown,M.A. (1990) Biochem. J., 271, 701–706.[ISI][Medline]

Barrett,A.J., Brown,M.A., Dando,P.M., Knight,C.G., McKie,N., Rawlings,N.D. and Serizawa,A. (1995) Methods Enzymol., 248, 529–556.[ISI][Medline]

Brown,C.K., Madauss,K., Lian,W., Beck,M.R., Tolbert,W.D. and Rodgers,D.W. (2001) Proc. Natl Acad. Sci. USA, 98, 3127–3132.[Abstract/Free Full Text]

Caccia,P. et al. (1992) Eur. J. Biochem., 204, 649–655.[Abstract]

Camargo,A.C., Oliveira,E.B., Toffoletto,O., Metters,K.M. and Rossier,J. (1987) J. Neurochem., 48, 1258–1263.[ISI][Medline]

Camargo,A.C.M., Gomes,M.D., Reichl,A.P., Ferro,E.S., Jacchieri,S., Hirata,I.Y. and Julianos,L. (1997) Biochem. J., 324, 517–522.[ISI][Medline]

Crack,P.J., Wu,T.J., Cummins,P.M., Ferro,E.S., Tullai,J.W., Glucksman,M.J. and Roberts,J.L. (1999) Brain Res., 835, 113–124.[CrossRef][ISI][Medline]

Cummins,P.M., Pabon,A., Margulies,E.H. and Glucksman,M.J. (1999) J. Biol. Chem., 274, 16003–16009.[Abstract/Free Full Text]

Dahms,P. and Mentlein,R. (1992) Eur. J. Biochem., 208, 145–154.[Abstract]

Dauch,P., Vincent,J.-P. and Checler,F. (1995) J. Biol. Chem., 270, 27266–27271.[Abstract/Free Full Text]

Ferro,E.S., Tullai,J.W., Glucksman,M.J. and Roberts,J.L. (1999) DNA Cell Biol., 18, 781–789.[CrossRef][ISI][Medline]

Fu,Z., Chakraborti,T., Morse,S., Bennett,G.S. and Shaw,G. (2001) Exp. Cell Res., 269, 275–286.[CrossRef][ISI][Medline]

Getz,E.B., Xiao,M., Chakrabarty,T., Cooke,R. and Selvin,P.R. (1999) Anal. Biochem., 273, 73–80.[CrossRef][ISI][Medline]

Gill,S.C. and Von Hippel,P.H. (1989) Anal. Biochem., 182, 319–326.[ISI][Medline]

Glucksman,M.J., Orlowski,M. and Roberts,J.L. (1992) Biophys. J., 62, 119–122.[ISI][Medline]

Guex,N. and Peitsch,M.C. (1997) Electrophoresis, 18, 2714–2723.[ISI][Medline]

Guex,N., Diemand,A. and Peitsch,M.C. (1999) Trends Biochem. Sci., 24, 364–367.[CrossRef][ISI][Medline]

Han,J., Clark,C., Han,G., Chuand,T.-C. and Han,P. (1999) Anal. Biochem., 268, 404–407.[CrossRef][ISI][Medline]

Koike,H. et al. (1999) J. Biochem. (Tokyo), 126, 235–242.[Abstract]

Lew,R.A., Hey,N.J., Tetaz,T.J., Glucksman,M.J., Roberts,J.L. and Smith,A.I. (1995) Biochem. Biophys. Res. Commun., 209, 788–795.[CrossRef][ISI][Medline]

Morales,T.I. and Woessner,J.F.,Jr. (1977) J. Biol. Chem., 252, 4855–4860.[Abstract]

Oliveira,V., Campos,M., Melo,R.L., Ferro,E.S., Camargo,A.C., Juliano,M.A. and Juliano,L. (2001) Biochemistry, 40, 4417–4425.[CrossRef][ISI][Medline]

Orlowski,M., Michaud,C. and Chu,T.G. (1983) Eur. J. Biochem., 135, 81–88.[Abstract]

Peitsch,M.C. (1995) Bio/Technology, 13, 658–660.[ISI]

Pierotti,A., Dong,K.W., Glucksman,M.J., Orlowski,M. and Roberts,J.L. (1990) Biochemistry, 29, 10323–10329.[ISI][Medline]

Ray,K., Hines,C.S. and Rodgers,D.W. (2002) Protein Sci., 11, 2237–2246.[Abstract/Free Full Text]

Resendes,M.N., Dobransky,T., Ferguson,S.S. and Rylett,R.J. (1999) J. Biol. Chem., 274, 19417–19421.[Abstract/Free Full Text]

Saric,T., Beninga,J., Graef,C.I., Akopian,T.N., Rock,K.L. and Goldberg,A.L. (2001) J. Biol. Chem., 276, 36474–36481.[Abstract/Free Full Text]

Serizawa,A., Dando,P.M. and Barrett,A.J. (1995) J. Biol. Chem., 270, 2092–2098.[Abstract/Free Full Text]

Shrimpton,C.N., Glucksman,M.J., Lew,R.A., Tullai,J.W., Margulies,E.H., Roberts,J.L. and Smith,A.I. (1997) J. Biol. Chem., 272, 17395–17399.[Abstract/Free Full Text]

Sigman,J.A., Edwards,S.R., Pabon,A., Glucksman,M.J. and Wolfson,A.J. (2003) FEBS Lett., 545, 224–228.[CrossRef][ISI][Medline]

Silva,C.L., Portaro,F.C.V., Bonato,V.L.D., de Camargo,A.C.M. and Ferro,E.S. (1999) Biochem. Biophys. Res. Commun., 255, 591–595.[CrossRef][ISI][Medline]

Smith,A.I., Shrimpton,C.N., Norman,U.M., Clarke,I.J., Wolfson,A.J. and Lew,R.A. (2000) Biochem. Soc. Trans, 28, 430–434.[ISI][Medline]

Tisljar,U. and Barrett,A.J. (1990) Biochem. J., 267, 531–533.[ISI][Medline]

Vincent,B., Jiracek,J., Noble,F., Loog,M., Roques,B., Dive,V., Vincent,J.-P. and Checler,F. (1997) Eur. J. Pharmacol., 334, 49–53.[CrossRef][ISI][Medline]

Wolfson,A.J., Shrimpton,C.N., Lew,R.A. and Smith,A.I. (1996) Biochem. Biophys. Res. Commun., 229, 341–348.[CrossRef][ISI][Medline]

Yamin,R., Malgeri,E.G., Sloane,J.A., McGraw,W.T. and Abraham,C.R. (1999) J. Biol. Chem., 274, 18777–18784.[Abstract/Free Full Text]

Received March 1, 2003; revised June 17, 2003; accepted June 23, 2003.





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