Suicide inactivation of peroxidase from Chamaerops excelsa palm tree leaves

International Journal of Biological Macromolecules 49 (2011) 1078–1082

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Suicide inactivation of peroxidase from Chamaerops excelsa palm tree leaves Nazaret Hidalgo Cuadrado a , Galina G. Zhadan b , Manuel G. Roig a,∗ , Valery L. Shnyrov b a b

Departamento de Química Física, Facultad de Ciencias Químicas, Universidad de Salamanca, 37008 Salamanca, Spain Departamento de Bioquímica y Biología Molecular, Universidad de Salamanca, 37007, Spain

a r t i c l e

i n f o

Article history: Received 27 July 2011 Received in revised form 31 August 2011 Accepted 1 September 2011 Available online 8 September 2011 Keywords: Chamaerops excelsa peroxidase Hydrogen peroxide Mechanism-based inactivation Suicide inactivation Ageing

a b s t r a c t The concentration and time-dependences and the mechanism of the inactivation of Chamaerops excelsa peroxidase (CEP) by hydrogen peroxide were studied kinetically with four co-substrates (2,2 -azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), guaiacol, o-dianisidine and o-phenylenediamine). The turnover number (r) of H2 O2 required to complete the inactivation of the enzyme varied for the different substrates, the enzyme most resistant to inactivation (r = 4844) with ABTS being the most useful substrate for biotechnological applications, opening a new avenue of enquiry with this peroxidase. © 2011 Elsevier B.V. All rights reserved.

1. Introduction Peroxidases (EC 1.11.1.7; donor: hydrogen peroxide oxidorreductase) form a group of enzymes that utilize H2 O2 to oxidize a second (reducing) substrate. These enzymes share a similar catalytic cycle in which H2 O2 reacts with the resting ferric enzyme to form the intermediate Compound I, which carries two oxidizing equivalents. Compound I is subsequently reduced by reactions with two reducing substrate molecules. The first of these reduction steps generates the intermediate, Compound II, which is then further reduced back to the ferric native enzyme. Palm peroxidase reacts with a large variety of reducing substrates, using H2 O2 as an oxidizing agent [1–3]. In the absence of reducing substrates, an excess of H2 O2 leads to inactivation of the enzyme, in this case the H2 O2 acting as a suicide substrate of peroxidase and being irreversibly bound to its active site [4,5]. However, it has been suggested by several authors [6,7] that the HRP inactivation by hydrogen peroxide is due to the formation of one or several non-active enzyme products likely through the formation of Compound III (peroxyl-FeIII porphyrin). The oxidative inactivation of peroxidases is mechanismbased. The molecular mechanism underlying this hydrogen

∗ Corresponding author. Tel.: +34 923 294 487; fax: +34 923 294 579. E-mail address: [email protected] (M.G. Roig). 0141-8130/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.ijbiomac.2011.09.001

peroxide-mediated inactivation is extraordinarily complex because a multitude of reactions can occur subsequent to the reaction of the haem iron with the hydroperoxide. Despite the peculiarities among the different peroxidases, a common inactivation mechanism comprising several stages can be proposed. In the absence of substrate, or when exposed to high concentrations of hydrogen peroxide, peroxidases show the kinetic behavior of suicide inactivation, in which hydrogen peroxide is the suicide substrate that converts Compound II into a highly reactive peroxy-iron(III) porphyrin free-radical named Compound III [8]. Compound III is not part of the peroxidase cycle, but is produced under excessive exposure of protonated Compound II to oxidative species in a reaction partially mediated by free superoxide radical [9]. Despite representing different structural groups, kinetic models for the hydrogen peroxide-mediated inactivation of horseradish peroxidase (HRP) [5], ascorbate peroxidase (APX) [10], peroxidase from the Royal Palm Tree (RPTP) [11], microperoxidase-11 [12] and CEP are similar in that they are time-dependent and show saturation kinetics. From the stoichiometry of the inactivation, it has been concluded that for APX only 2.5 molecules of hydrogen peroxide are required per active site to generate the inactivation form [10], in contrast to the 265 molecules required for HRP [5]. This difference is due to the low catalytic activity of HRP, which is absent in APX [13]. For APX peroxidase, inactivation is correlated with enzyme bleaching, suggesting haem destruction [10]. Another factor in this difference is the glycosylation of the enzyme, which appears to be significant in protecting the enzyme from inactivation [5].

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2. Experimental

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calibration kit (4.45–9.6) from Bio-Rad Laboratories (Hercules, CA, USA).

2.1. Materials Analytical or extra-pure-grade polyethyleneglycol (PEG), guaiacol (2-methoxyphenol), ammonium sulfate, sodium phosphate and Tris–HCl were purchased from Sigma Chemical Co. (St. Louis MO, USA) and were used without further purification. H2 O2 was from Merck (Darmstadt, Germany). Superdex-200 columns and PhenylSepharose CL-4B columns were from GE Healthcare Bio-Sciences AB (Uppsala, Sweden). TSK-Gel DEAE-5PW was purchased from Tosoh Co. (Tokyo, Japan). Cellulose membrane tubing for dialysis (avg. flat width 3.0 in.) was purchased from Sigma Chemical Co.; slide A-lyzer dialysis cassettes (extra-strength, 3–12 mL capacity, 10,000 MWCO) were from Millipore Corp. (Billerica, MA, USA). All other reagents were of the highest purity available. The water used for preparing the solutions was double-distilled and then subjected to a de-ionisation process.

2.2. Enzyme purification CEP was purified from palm tree Chamaerops excelsa as described [14,15] but with essential modifications. Leaves (1820 g) from three-year-old palm tree were milled and homogenized in 7.28 L distilled water for 22–24 h at room temperature. Excess material was removed by vacuum filtration and centrifugation (10,000 × g, 277 K for 15 min). Pigments were extracted by phase separation over 20–22 h at 277 K after the addition to the supernatant of solid PEG to 14% (w/v) and solid ammonium sulfate to 10% (w/v). Two phases were formed after addition of ammonium sulfate: an upper polymer phase (dark brown), which contained pigments, phenols, polyphenols, oxidized phenols and PEG, and lower aqueous phase (yellow) containing peroxidase. Each phase consisted of 50% of the initial volume. These phases were separated and the phase containing peroxidase activity was centrifugated. The clear supernatant containing peroxidase activity was titrated with ammonium sulfate to a conductivity value of 232 mS cm−1 and was applied on a Phenyl-Sepharose column (1.5 cm × 35 cm) equilibrated with 100 mM phosphate buffer, pH 6.5, with 1.7 M ammonium sulfate, which has the same conductivity as the sample. The enzyme was eluted with 100 mM phosphate buffer, pH 6.5, plus 0.2 M ammonium sulfate at a flow rate of 1 mL min−1 . 15 mL fractions were collected and those showing peroxidase activity were dialyzed against 5 mM Tris buffer, pH 9.3, for 72 h, with constant stirring at 277–278 K. These fractions were membrane-concentrated (Amicon, 10 kDa cutoff) to 15 mL and applied onto a TSK-Gel DEAE-5PW column (1 cm × 30 cm) equilibrated with 5 mM Tris buffer, pH 9.3. Elution was carried out with a linear 0–300 mM NaCl gradient in the same buffer at a flow rate of 1 mL min−1 . The fractions with peroxidase activity were collected, membrane-concentrated (Amicon, 10 kDa cutoff), and applied on a Superdex-200 column equilibrated with 5 mM Tris buffer, pH 9.3. Elution was carried out in the same buffer at a flow rate of 1 mL min−1 . Finally, the peroxidase was dialyzed against distilled water and freeze-dried. The purity of the CEP was determined by SDS-PAGE as described by Fairbanks et al. [16] on a Bio-Rad Minigel device using a flat block with 12% polyacrylamide concentration; by gel filtration, which was performed using a Superdex-200 10/30 HR column in an FPLC Amersham Äkta System; and by UV–visible spectrophotometry (RZ = A403 /A280 = 2.8–3.0). Analytical isoelectrofocusing was performed on a Mini IEF cell model 111 (Bio-Rad Laboratories, Hercules, CA, USA) using Ampholine PAG-plates, pH 3.5–9.5 (GE Healthcare Bio-sciences AB, Uppsala, Sweden). The electrophoretic conditions and Silver Staining Kit Protein were as recommended by manufacturer. The standards used were from a broad-range pI

2.3. Inactivation experiments CEP was inactivated at 25 ◦ C in 10-mL incubations of universal buffer, pH 7.0, for each substrate except ABTS, whose pH was 3.0, containing a fixed amount of the enzyme (136 nM). The reason for studying the inactivation at acidic pH has been the fact that other plant peroxidases have been characterized at this pH, so that comparison can be done. The reactions were started by the addition of H2 O2 (over a range of concentrations). At specified time intervals, 5-␮L aliquots of the incubation mixtures were transferred to cuvettes containing 2 mL of an assay mixture composed of an optimum amount of substrate and H2 O2 (0.032 mM ABTS and 1.12 mM H2 O2 ; 3.0 mM o-dianisidine and 0.1 mM H2 O2 ; 2.0 mM o-phenylenediamine and 0.75 mM H2 O2 ; 18 mM guaiacol and 4.9 mM H2 O2 ). Peroxidase activity was measured by the increase in absorbance at the characteristic wavelengths for these substrates [11]; neither the substrate nor the CEP present in the assay mixture interfered with the measurement. A minimum of three incubation assays for each peroxide concentration was performed. The residual enzymatic activity (AR ) was taken as the enzymatic activity remaining (At ) as a percentage of the initial activity (A0 ). Table 1 shows all the parameters of the substrates used in the inactivation assays. The residual peroxidase activity of all the substrates was assayed after 24 h. 3. Results and discussion 3.1. Determination of the partitioning ratio (r) for the inactivation of CEP by H2 O2 Using plots of the percent residual activity for each substrate against the peroxide/enzyme ratio, the ratio required in each case for 100% inactivation can be obtained from the intercept of the fitted line. From this value, the partitioning ratio can be calculated using the following equation: AR =

At 1 [H2 O2 ] =1− , A0 1 + r [CEP]

where AR is the residual activity, At and A0 are the activities at time (t) (end of the reaction) and zero, respectively, (r) is the partitioning ratio (number of catalytic cycles given by enzymes before their inactivation), and [H2 O2 ] and [CEP] are the initial concentrations of H2 O2 and enzyme [5]. Fig. 1 shows the plots of the percent residual activity against the H2 O2 /CEP ratio for the ABTS substrate, which are representative of the results obtained for different substrates. Taking into account the consumption of two moles of H2 O2 in each catalytic cycle (1 mole for the formation of Compound I and another for inactivation or catalysis) [17–19], the (r) values calculated from these plots are given in Table 2. The inactivation of each of the enzymes by H2 O2 at a range of different concentrations was also followed over time (the results for ABTS that were representative of those obtained for each substrate are shown in Fig. 2). 3.2. Kinetics of inactivation by H2 O2 The inactivation of each of the enzymes by H2 O2 at a range of different concentrations was also followed over time (as an example, the results for ABTS are shown in Fig. 2). A plot of the natural logarithms of the percentage residual activities (ln AR ) against time afforded straight lines (Fig. 3), with slopes

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Table 1 Parameters of the different substrates (S).

ABTS Guaiacol o-Dianisidine o-Phenylenediamine

pH

Range [H2 O2 ] (mM)

 (nm)

[S] (mM)

[H2 O2 ] (mM)

3.0 7.0 7.0 7.0

40–500 30–500 30–200 30–200

414 470 420 445

0.032 18 3.0 2.0

1.12 4.9 0.1 0.75

Table 2 app app Apparent kinetic constants calculated for the inactivation of peroxidase variants by H2 O2 . (r), turnover number; kinact , inactivation rate constant; KI , dissociation constant app app app app app app of the [Compound I·H2 O2 ] complex; 1/KI , affinity constant of enzyme and H2 O2 ; kcat , rate constant of catalysis; kcat /KI , efficiency of catalysis; kinact /KI , efficiency of inactivation.

ABTS Guaiacol o-Dianisidine o-Phenylenediamine

app

app

r

kinact (s−1 )

KI

4844 3711 951 2147

3.24 3.85 1.27 1.79

845 204 39.6 87.1

(mM)

app

1/KI

app

(M−1 )

1.18 4.90 25.2 11.5

app

app

kcat (s−1 )

kcat /KI

15,694 14,287 1209 3839

18,573 70,034 30,530 44,076

(s−1 M−1 )

app

app

kinact /KI

(s−1 M−1 )

3.83 18.87 32.07 20.55

equivalent to the observed rate constants of the inactivation (kobs ) [10]. ln

[Aa ] = −kobs t [A0 ]

The inactivation of CEP by H2 O2 clearly showed saturation kinetics, as seen from the hyperbolic curves fitted to the plot of kobs against [H2 O2 ] (Fig. 4). Unlike the inactivation of HRP by H2 O2 with different substrates, the inactivation of CEP exhibits single-phase kinetics, unlike what has been found for HRP (biphasic) [4,5,20]. The observed hyperbolic nature of the plots of kobs against the H2 O2 concentration provided clear evidence that the inactivation process had saturation kinetics. From these data, the observed first-order rate constants of inactivation (kobs ) can be calculated using the equation: app

kobs = Fig. 1. Sensitivity to inactivation of CEP at different molar ratios of H2 O2 . CEP was incubated with molar excesses of peroxide in universal buffer (10 mM, pH 3.0). When the reaction was complete (18–24 h incubation) the percentage residual activities were measured with ABTS and other different substrates.

Fig. 2. Time-dependence and kinetics of the inactivation of CEP by H2 O2 . Inactivation of CEP followed over time. CEP (136 nM) was incubated with H2 O2 at concentrations of 40 (), 75 (), 250 (䊉), 350 (), 500 () mM.

kinac [H2 O2 ] app KI

+ [H2 O2 ]

,

where the kobs (the first-order inactivation rate constant) [21] values were obtained from the component in a fit of the data by a linear app regression model; kinac is a first-order inactivation rate constant, app is an inhibitor-binding constant. and KI Double-reciprocal plots of rate vs. inhibitor concentration curves are often used as tools to examine the formation of

Fig. 3. Least-squares linearized plots of percentage of remaining activity vs. time data for selected concentrations ([CEP] = 136 nM at 25 ◦ C, pH 3.0). H2 O2 concentrations: 40 (), 75 (), 250 (䊉), 350 (), 500 () mM.

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the turnover numbers (r) above, the values of the catalytic constant app (kcat ), using the relationship: app

r=

Fig. 4. Kinetic curve of inactivation. Direct and double reciprocal plot (insert) of kobs (first-order inactivation rate constant) against the H2 O2 concentration.

complexes between inhibitors and enzymes [22]. The apparent kinetic constants calculated for the different forms of this enzyme are shown in Table 2. In the absence of reducing substrates and at high H2 O2 concentrations, CEP is inactivated in a time- and H2 O2 concentrationdependent process, exhibiting suicide or mechanism-based inactivation kinetics. Similar behavior has been reported by other authors [4,5,11,20,22] in studies of H2 O2 -mediated inactivation under identical experimental conditions of temperature, pH, and the concentrations of enzyme and H2 O2 . However, differences exist in the shapes of the inactivation curves, in the magnitudes and rates of inactivation at specific H2 O2 concentrations, and in the maximal app rate of inactivation (kinac ) in comparison with the results reported in the present study. The inactivation process (Fig. 2 and Table 2) involves the participation of two pathways: one reversible and other irreversible, which may or may not function independently of each other and whose individual contribution to the overall inactivation process appears to depend on the H2 O2 concentration. For enzyme inactivation, a second molecule of hydrogen peroxide would be required, but not for inactivation itself but for the formation of Compound III. Once Compound III is formed, a fraction of that population would be transformed into a non-active species. An alternative interpretation would be that once a molecule of Compound III is formed, it has a certain probability to decay into a non-active species against decaying into an active species that would engage again in the catalytic cycle. Several authors [4,5,22] have proposed the existence of a partitioning between the two pathways for Compound I. Their model, based on studies at high H2 O2 concentrations, invokes a further partitioning between these two inactivation pathways and suggests the existence of a catalytic reaction in which H2 O2 is consumed with relatively little harm to the enzyme [22]. The model is in fact a simplification of the full kinetic approach developed previously by Arnao et al. [21], although it does offer a reasonable approximation suitable for comparative purposes, as required here. Double-reciprocal plots of the rate of inhibition against the inhibitor concentration have often been used to examine the formation of complexes between enzymes and inhibitors [22–24]. We used linear least-squares fits to the data (Fig. 3) to determine the relationship between the different enzymes and H2 O2 . For each variant we obtained the apparent values of the app rate constant of inactivation (kinac ) and the dissociation constant app (KI ) of the enzyme/inhibitor-complex and, in conjunction with

kcat

app

kinact

The results obtained are given in Table 2 and are summarized below. app The value of kinac with H2 O2 for guaiacol is greater than for the other substrates but Compound I of CEP had the highest affinity app app for H2 O2 (1/KI ), and the highest catalytic rate constant (kcat ) app app for ABTS. Furthermore, the efficiency of catalysis (kcat /KI ) was app app greater for guaiacol and the efficiency of inactivation (kinact /KI ) was also greater for the o-dianisidine substrate. The accessibility to the substrate towards the reducing binding site in the haem pocket and its affinity for the product are important for enzyme activity, as was observed with o-dianisidine. Substrate specificity may be altered by changes affecting the reducing binding site [25] and it may be possible to further adjust the specificity and level of activity of CEP by judicious changes in this region. The glycosylation of CEP appears to be significant in protecting the enzyme from inactivation with the different substrates. However, CEP is the most active of all currently known peroxidases, suggesting that the active site of the enzyme has evolved not only for improving catalytic efficiency but also for preventing inactivation by the highly reactive H2 O2 substrate. 4. Conclusions The kinetics of inactivation of CEP in the oxidation of different substrates studied (ABTS, guaiacol, o-dianisidine and o-phenylenediamine) by hydrogen peroxide shows suicide inactivation behavior similar to that of most classical peroxidases [4,5,10,13]. The model used in these experiments [10] provides satisfactory parameters for the inactivation kinetics by hydrogen peroxide, showing the high capacity of the enzyme to act on the studied substrates at different pHs. In the case of ABTS at pH 3, pH at which most enzymes are not able to function, CEP performs a turnover of molecules up to 4844 and exhibits an apparent rate constant of catalysis of 15,694 s−1 . For the rest of substrates, at pH 7, the kinetic parameters were turnover numbers 3711, 951, 2147 and the apparent rate constant of catalysis of 14,287 s−1 , 1209 s−1 and 3839 s−1 for guaiacol, o-dianisidine and o-phenylenediamine, respectively. These values may indicate that CEP exhibits an extraordinary performance against inactivation by hydrogen peroxide, opening the possibility of further studies related to the mechanisms of exchange of hydrogen peroxide in peroxidases and pointing, besides other studies [15], that CEP acts as a very robust enzyme. References [1] L. Watanabe, P.R. de Moura, L. Bleicher, A.S. Nascimiento, L.S. Zamorano, J.J. Calvete, L. Sanz, A. Pérez, S. Bursakov, M.G. Roig, V.L. Shnyrov, I. Polikarpov, J. Struct. Biol. 169 (2010) 226–242. [2] I.Y. Sakharov, B.M.K. Vesga, I.V. Sakharova, Biochemistry (Moscow) 67 (2002) 1043–1047. [3] I.Y. Sakharov, Biochemistry (Moscow) 69 (2004) 823–829. [4] M.B. Arnao, M. Acosta, J.A. del Río, F. García-Cánovas, Biochim. Biophys. Acta 1038 (1990) 85–89. [5] A.N.P. Hiner, J. Hernández-Ruiz, F. García-Cánovas, A.T. Smith, M.B. Arnao, M. Acosta, Eur. J. Biochem. 234 (1995) 506–512. [6] M. Puiu, A. R˘aducan, I. Babaligea, D. Oancea, Bioprocess Biosyst. Eng. 31 (2008) 579–586. [7] S.M. Aitken, M. Ouellet, M.D. Percival, A.M. English, Biochem. J. 375 (2003) 613–621. [8] R. Nakaijima, I. Yamazaki, J. Biol. Chem. 262 (1987) 2576–2581. [9] S.S. Adediran, A.M. Lambeir, Eur. J. Biochem. 186 (1989) 571–576. [10] A.N.P. Hiner, J.N. Rodriguez-López, M.B. Arnao, E.L. Raven, F. García-Canovas, M. Acosta, Biochem. J. 348 (2000) 321–328.

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