Dipartimento Scienze Molecolari Agroalimentari, Università di Milano, Via Celoria 2, 20133 Milan, Italy
1 To whom correspondence should be addressed. e-mail: silvia.pagani{at}unimi.it
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Keywords: phosphatase activity/RhdA active-site elongation/rhodanese modules/substrate selectivity/sulfurtransferase activities
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Therefore, RhdA is the member of known rhodaneses that should suit the fundamental requirements for possible phosphatase activity. The major difference characterizing the RhdA active-site with respect to the three closely related Cdc25 phosphatases (Cdc25A, B, C) is that in Cdc25 phosphatases the activesite loop [HisCys(X)5Arg] is one residue longer than in RhdA [HisCys(X)4Arg]. The five X residues in Cdc25s form a loop whose backbone amides hydrogen bond to the phosphate of the substrate (Rudolph, 2002). The absence in RhdA of phosphatase activity supports the relevance of the length of the active-site loop for catalysis. According to this hypothesis, rational mutagenesis was applied to investigate the effect of RhdA active-site loop elongation in substrate recognition. On the structural basis detailed in our previous study (Bordo et al., 2001
), the position between amino acids His233 and His234, which are structurally equivalent in Cdc25A to Ser433 and Glu435, was considered the most appropriate for a single-residue insertion in A.vinelandii RhdA and Ala and Ser residues were chosen for the insertion. Preliminary analysis of the engineered RhdAs (RhdA-Ala and RhdA-Ser mutants) failed to detect phosphatase activity, based on p-nitrophenyl phosphate (pNPP) hydrolysis (Bordo et al., 2001
). Since pNPP is recognized as a poor substrate for Cdc25 enzymes and the search for good artificial substrates for in vitro analysis of phosphatase activity of Cdc25 enzymes is a subject of investigation (Gottlin et al., 1996
; Chen et al., 2000
; Kolmodin and Aqvist, 2000
; McCain et al., 2002
), the lack of reactivity towards pNPP was not considered sufficient to dismiss the hypothesis that elongation of the RhdA active-site loop could generate enzymes with phosphatase activity. Keeping this in mind, we have extended the previous study and analyzed the reactivity of either sulfur- or phosphate-containing compounds with the modified scaffold of RhdA.
The main goal of the present work was to provide evidence that both RhdA-Ala and RhdA-Ser mutants, but not wild-type RhdA, were able to catalyze the hydrolysis of the phosphatase artificial substrate 3-O-methylfluorescein phosphate (OMFP). The absence of any sulfurtransferase activities in both mutants indicated that specificity of recognition (S- or P-containing substrate) is essentially determined by the tailored arrangement of the catalytic loop. The results of this investigation should be taken as experimental evidence of the hypothesis (Bordo and Bork, 2002) that the structural difference in the active-site loop length (rhodaneses versus Cdc25 phosphatases) may reflect a specific mutational event (a single-residue insertion or deletion), which changed the selectivity from sulfur- to phosphate-containing substrates (or vice versa).
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
The recombinant plasmids for expression of RhdA [pQER1 (Pagani et al., 2000)] or of RhdAs with single-residue insertion [pQM22 or pQM5 (Bordo et al., 2001
)] were transformed into Escherichia coli BL21[rep4] and protein overexpression was rapidly induced by addition of 1 mM isopropyl thio-ß-D-galactoside to a mid-logarithmic culture (OD600 = 0.6). Cell-free extracts were prepared from 500 ml of culture. After 4 h of induction, cells were harvested by centrifugation and resuspended in 5 ml of 50 mM TrisHCl buffer (pH 8.0) containing 0.3 M NaCl. Cell disruption was carried out by incubation with 0.3 mg/ml lysozyme and sonication. RhdA and the RhdAs with single-residue insertion (RhdA-Ala and RhdA-Ser, respectively) were purified by chromatography on an Ni-NTA agarose column (gel volume, 8 ml). The His-tagged proteins were eluted by addition of 200 mM imidazole.
Activity assays
The discontinuous method that determines the product thiocyanate, based on the absorbance of the ferricthiocyanate complex at 460 nm, was used to determine either thiosulfate: cyanide sulfurtransferase (rhodanese, TST) or 3-mercaptopyruvate:cyanide sulfurtransferase (MST) activities (Westley, 1981). The assays lasted 12 min and 1 U of enzyme is defined as the amount of enzyme that produces 1 µmol of thiocyanate per minute at 37°C.
Sulfurtransferase activity in the presence of dithiothreitol (DTT) as acceptor substrate and thiosulfate as sulfur donor was determined spectrophotometrically by the continuous method described by Pecci et al. (Pecci et al., 1976). The rate of spontaneous autoxidation of DTT was always subtracted. One unit (U) of enzyme is defined as the amount of enzyme that oxidizes 1 µmol of dithiothreitol per minute at 37°C.
Phosphatase activity was assayed by a continuous fluorimetric method which measures the initial rates of hydrolysis of the artificial substrate OMFP (Gottlin et al., 1996). The fluorimetric measurements were carried out with an LS50 luminescence spectrometer (Perkin-Elmer) equipped with a PTP-1 Peltier temperature programmer (Perkin-Elmer) set at 20°C; the excitation and emission wavelengths were 471 and 530 nm, respectively, slit width 5 nm and data pitch 1 s. 3-O-Methylfluorescein (OMF) formation was quantitated by fluorescence and fluorescence units were converted to product concentration by using an OMF calibration curve generated by measuring the fluorescence of OMF solutions at various concentrations in the assay buffer. One unit (U) is defined as the amount of enzyme that produces 1 µmol of OMF per minute at 20°C. The assay mixture was 0.5 ml of 50 mM TrisHCl (pH 8), 0.25 mM OMFP and different enzyme concentrations; when stated, 1 mM dithiothreitol (DTT) was included in the assay mixture. In a set of experiments, the examined proteins were pre-incubated in the assay buffer containing 2 mM DTT for 1 h at 25°C before addition of OMFP; the final DTT concentration in the assay was 1 mM. The rate of spontaneous OMFP hydrolysis was always subtracted and the data presented are the averages of at least three independent determinations.
The protein concentration was determined by dye-binding colorimetric assay (Bradford, 1976).
![]() |
Results and discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Sulfurtransferase activities of wild-type and mutated forms of RhdA
The residues surrounding the catalytic Cys230 in RhdA generate a strong positive electrostatic field (Bordo et al., 2000), which is in keeping with the RhdA in vitro enzymic activity:
S2O32 + RhdA SO32 + RhdA-S
RhdA-S + CN RhdA + SCN
Even in the absence of thiosulfate, RhdA is isolated as a stable Cys230 persulfurated form (Bordo et al., 2000; Pagani et al., 2000
), a distinctive feature not shared by the mutants with elongated catalytic loop RhdA-Ala and RhdA-Ser (Bordo et al., 2001
). As shown in Table I, single-residue insertion in the catalytic loop of RhdA strongly impaired the ability to transfer sulfur in the presence of thiosulfate as sulfur donor to either cyanide (TST activity) or to the dithiol DTT (DTT-ST activity), the typical rhodanese activities. The MST of wild-type RhdA was low, compared with TST activity. Even single-residue insertion in the catalytic loop of RhdA affected MST activity; the greatest effect of the elongation was found in the sulfur transfer reactions in the presence of thiosulfate as sulfur donor. This result can be taken as evidence that productive binding of thiosulfate requires a specifically arranged catalytic loop, not maintained in the modified scaffold of RhdA-Ala and RhdA-Ser mutants. Our previous study (Pagani et al., 2000
) showed that RhdA did not follow the consensus rules of substrate donor recognition of eukaryotic sulfurtransferases (Luo and Horowitz, 1994
; Nagahara et al., 1995
; Nagahara and Nishino, 1996
). The ability to catalyze sulfur transfer from thiosulfate to cyanide of both Thr232Lys and Thr232Ala RhdAs (Pagani et al., 2000
), which were expressed in the persulfurated form, was, indeed, not affected by the change of the residue at position +2 with respect to the catalytic cysteine. The above evidence and the data presented here (Table I) might suggest that in RhdA interactions with persulfurated active-sites, cysteine should be the basis for productive binding with sulfur acceptor in the sulfur transfer reactions.
|
The rhodanese and Cdc25 phosphatase families display five or six sequential peptide NH groups, respectively, radially arranged around the thiol group of the catalytic cysteine residue, which is located at the bottom of the active-site pocket. Crystallographic analyses (Bordo et al., 2001) showed that hypophosphite, but not phosphate, is stably bound to the catalytic pocket of the desulfurated RhdA, thus suggesting that the precise size restrictions in the RhdA catalytic pocket may result in a non-productive accommodation of phosphatase substrate(s). Considering that the selection of proper substrate(s) for Cdc25-like phosphatase could be the reason for the lack of reactivity towards pNPP of the mutated RhdAs with an elongated catalytic loop (Bordo et al., 2001
), we attempted to re-evaluate the effect of single-residue insertion at the position between His233 and His234 of RhdA in generating phosphatase activity. The phosphatase activity of wild-type RhdA and of the mutated RhdAs, tailored to mimick the Cdc25 enzymes active-site loop (RhdA-Ala and RhdA-Ser) was tested by using the artificial substrate OMFP. OMFP has been proved to be an appropriate substrate for Cdc25 phosphatases (Gottlin et al., 1996
; Chen et al., 2000
), showing values of kcat/Km significantly higher than pNPP (McCain et al., 2002
). As is evident in Figure 1A, the hydrolysis product OMF was detectable only in the presence of RhdA-Ala and RhdA-Ser and its production was a linear function of the concentration in the assay of the RhdA mutants with an elongated active-site loop. The calculated values of kcat are listed in Table II, representing the results presented in Figure 1A and Table II (column A) obtained by measuring phosphatase activity in the absence of the reductant DTT, usually present in the assay for Cdc25 enzymes (Dunphy and Kumagai, 1991
). Negligible phosphatase activity was found for either wild-type RhdA or sulfur-free RhdA obtained by cyanolysis, indicating that the thiolate of the catalytic cysteine is not the only structural requirement for phosphatase activity in RhdA. The designed loop insertion mutations in RhdA were, however, effective in generating productive interaction with OMFP, although the activity kcat values for these RhdA mutants were low compared with those of Cdc25 enzymes (Gottlin et al., 1996
; Chen et al., 2000
; McCain et al., 2002
).
|
|
Effect of reductants on RhdA mutants
The reactive nature of the active-site cysteine and the fact that the sulfenic form of the cysteine is enzymatically inactive support the need for reducing agents in the phosphatase assays of Cdc25s (Claiborne et al., 1999). From the molecular point of view, the published 3D structure of Cdc25s (Fauman et al., 1998
; Reynolds et al., 1999
) demonstrated that the active-site cysteine can form an intramolecular disulfide bond with another conserved cysteine in the molecule. In a recent report (Savitsky and Finkel, 2002
), the effects of oxidative stress on the Cdc25s were examined and it was suggested that in Cdc25C, formation of a disulfide bond between Cys330 and the active-site Cys377 could rescue the protein and prevent the formation of definitely inactivated sulfinic species. In the case of the engineered RhdAs, the presence of only one cysteine residue in the molecule does not allow the formation of any intramolecular disulfide bond. In an attempt to give a rationale for the DTT effect on phosphatase activity of the engineered RhdAs (see Table II), we analyzed whether different experimental conditions of reductant addition could modulate the phosphatase activity of the mutated RhdAs (Figure 2). In the case of RhdA-Ala, comparison of the activity values in the presence of DTT (1 mM final concentration) obtained by using in the assay the enzyme pre-incubated with DTT (kcat = 0.055 min1) or not (kcat = 0.046 min1) indicated that under both experimental conditions the phosphatase activity doubled with respect to that measured in the absence of the reductant (kcat = 0.026 min1). The effectiveness of DTT in enhancing the phosphatase activity of RhdA-Ser was higher and was dependent on the experimental conditions of reductant addition, the kcat values being 0.003 min1 (in the absence of DTT), 0.013 and 0.026 min1 (with DTT only in the assay and after pre-incubation, respectively). The free thiol present in RhdA-Ser and RhdA-Ala could be the target of oxidative events. The significant increase in phosphatase activity of both RhdA mutants via reductive pressure of DTT could be ascribed to the fact that DTT could prevent the formation of a sulfenic form (i.e. Cys-SOH) of the enzyme, cysteine sulfenic acid being readily reversible by thiol reduction (Savitsky and Finkel, 2002
). Mass spectrometric measurements (data not shown) were not consistent with cysteine sulfenic acid formation on the molecules of the RhdA mutants with an elongated active-site loop. The formation of an intermolecular disulfide bond, on the other hand, has to be considered since an additional protein band showing a Mr consistent with an RhdA mutant dimeric form was observed by SDSPAGE analysis (Figure 3, lanes 12). In both RhdA-Ala and RhdA-Ser, this form was evaluated to represent
50% of the enzyme population. According to the formation of an intermolecular disulfide bond, a single protein band was detected in both mutated RhdAs by SDSPAGE analysis in the presence of ß-mercaptoethanol (Figure 3, lanes 56). Notably, the electrophoretic patterns of RhdA-Ala and RhdA-Ser after DTT treatment in the same conditions used for phosphatase activity detection showed one only protein band (Figure 3, lanes 34), indicating that DTT treatment did prevent the oxidative process involving the catalytic cysteine of the modified RhdAs. Although quantification of the observed molecular forms (i.e. monomeric versus dimeric forms) is rather speculative by SDSPAGE analysis, in the case of RhdA-Ala the effectiveness of DTT in preserving the form with the cysteine available for catalysis should explain the doubling of phosphatase activity in the presence of the reductant.
|
|
Our previous studies on Azotobacter vinelandii sulfurtransferase (Bordo et al., 2000, 2001; Pagani et al., 2000
) revealed unique structural features of RhdA which make it a suitable model for the identification of functional diversity determinants among the rhodanese superfamily enzymes. Therefore, from a comparison with the structurally homologous Cdc25 phosphatases (Fauman et al., 1998
; Hofmann et al., 1998
; Reynolds et al., 1999
; Bordo et al., 2001
), we have engineered the RhdA scaffold to produce an artificial enzyme mimicking the pocket of Cdc25 active-sites. In the mutants RhdA-Ala and RhdA-Ser with single-residue introduction via rational design (Bordo et al., 2001
), the elongation of the native RhdA catalytic loop definitely resulted in the impairment of productive interaction with substrates involved in sulfur transfer reactions. The modified catalytic loop, on the other hand, made RhdA-Ala and RhdA-Ser able to interact productively with the artificial phosphatase substrate OMFP. Taken together, these results clearly indicate that sulfurtransferase and phosphatase activities are not capable of existing concurrently. Since the identification of artificial substrates suitable for kinetic analyses of Cdc25 enzymes seems to be difficult (Gottlin et al., 1996
; Chen et al., 2000
; Kolmodin and Aqvist, 2000
; McCain et al., 2002
), the choice of the artificial substrate OMFP in the present study was dictated by the fact that it has been proven to be more efficient than pNPP for Cdc25s. Although there is still much to learn about whether specific structural interactions are responsible for the catalytic and molecular behaviors of RhdA-Ala and RhdA-Ser, their ability to catalyze OMFP hydrolysis highlighted the functional plasticity of the RhdA scaffold. Considering that the recognition of distinctive structural motives in the ubiquitous rhodanese domains should facilitate the identification of possible biological substrates (Bordo and Bork, 2002
), here, for the first time, we have identified in a model rhodanese scaffold a structural determinant for selective recognition of substrates (i.e. sulfur- or phosphate-containing compounds).
![]() |
Acknowledgements |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() |
---|
Bordo,D. and Bork,P. (2002) EMBO Rep., 3, 741746.
Bordo,D., Deriu,D., Colnaghi,R., Carpen,A., Pagani,S. and Bolognesi,M. (2000) J. Mol. Biol., 298, 691704.[CrossRef][ISI][Medline]
Bordo,D., Forlani,F., Spallarossa,A., Colnaghi,R., Carpen,A., Bolognesi,M. and Pagani,S. (2001) Biol. Chem., 382, 12451252.[ISI][Medline]
Bradford,M.M. (1976) Anal. Biochem., 72, 248254.[CrossRef][ISI][Medline]
Burow,M., Kessler,D. and Papenbrock,J. (2002) Biol. Chem., 383, 13631372.[ISI][Medline]
Chen,W., Wilborn,M. and Rudolph,J. (2000) Biochemistry, 39, 1078110789.[CrossRef][ISI][Medline]
Claiborne,A., Yeh,J.I., Luba,J., Crane,E.J., Charrier,V. and Parsonage,D. (1999) Biochemistry, 38, 1540715416.[CrossRef][ISI][Medline]
Colnaghi,R., Pagani,S., Kennedy,C. and Drummond,M. (1996) Eur. J. Biochem., 236, 240248.[Abstract]
Colnaghi,R., Cassinelli,G., Drummond,M., Forlani,F. and Pagani,S. (2001) FEBS Lett., 500, 153156.[CrossRef][ISI][Medline]
Dunphy,W.G. and Kumagai,A. (1991) Cell, 67, 189196.[ISI][Medline]
Fauman,E.B., Cogswell,J.P., Lovejoy,B., Rocque,W.J., Holmes,W., Montana,V.G., Piwnica-Worms,H., Rink, M. and Saper,M.A. (1998) Cell, 93, 617625.[ISI][Medline]
Gottlin,E.B., Xu,X., Epstein,D.M., Burke,S.P., Eckstein,J.W., Ballou,D.P. and Dixon,J.E. (1996) J. Biol. Chem., 271, 2744527449.
Hofmann,K., Bucher,P. and Kajava,A.V. (1998). J. Mol. Biol., 282, 195208.[CrossRef][ISI][Medline]
Jackson,M.D. and Denu,J.M. (2001) Chem. Rev., 101, 23132340.[CrossRef][ISI][Medline]
Kolmodin,K. and Aqvist,J. (2000) FEBS Lett., 465, 811.[CrossRef][ISI][Medline]
Luo,G. and Horowitz,P.M. (1994) J. Biol. Chem., 269, 82208225.
McCain,D.F., Catrina,I.E., Hengge,A.C. and Zhang,Z.Y. (2002) J. Biol. Chem., 277, 1119011200.
Mueller,E.G., Palenchar,P.M. and Buck,C.J. (2001) J. Biol. Chem., 276, 3358833595.
Nagahara,N. and Nishino,T. (1996) J. Biol. Chem., 271, 2739527401.
Nagahara,N., Okazaki,T. and Nishino,T. (1995) J. Biol. Chem., 270, 1623016235.
Nilsson,I. and Hoffmann,I. (2000) Prog. Cell Cycle Res., 4, 107114.[Medline]
Pagani,S., Forlani,F., Carpen,A., Bordo,D. and Colnaghi,R. (2000) FEBS Lett., 472, 307311.[CrossRef][ISI][Medline]
Palenchar,P.M., Buck,C.J., Cheng,H., Larson,T.J. and Mueller,E.G. (2000) J. Biol. Chem., 275, 82838286.
Papenbrock,J. and Schmidt,A. (2000) Eur. J. Biochem., 267, 55715579.
Pecci,L., Pensa,B., Costa,M., Cignini,P.L. and Cannella,C. (1976) Biochim. Biophys. Acta, 445, 104111.[ISI][Medline]
Ploegman,J.H., Drent,G., Kalk,K.H., Hol,W.G.J., Heinrikson,R.L., Keim,P., Weng,L. and Russel,J. (1978) Nature, 273, 124129.[ISI][Medline]
Reynolds,R.A., Yem,A.W., Wolfe,C.L., Deibel,M.R.,Jr, Chidester,C.G. and Watenpaugh,K.D. (1999) J. Mol. Biol., 293, 559568.[CrossRef][ISI][Medline]
Rudolph,J. (2002) Biochemistry, 41, 1461314623.[CrossRef][ISI][Medline]
Savitsky,P.A. and Finkel,T. (2002) J. Biol. Chem., 277, 2053520540.
Schultz,J., Milpetz,F., Bork,P. and Ponting,C.P. (1998) Proc. Natl Acad. Sci. USA, 95, 58575864.
Spallarossa,A., Donahue,J.L., Larson,T.J., Bolognesi,M. and Bordo,D. (2001) Structure, 9, 120.[ISI][Medline]
Westley,J. (1981) Methods Enzymol., 77, 285291.[Medline]
Xu,X. and Burke,S.P. (1996) J. Biol. Chem., 271, 51185124.
Received February 17, 2003; revised May 30, 2003; accepted June 6, 2003.