- Email: [email protected]
Effects of phthalic anhydride modification on horseradish peroxidase stability and structure
Enzyme and Microbial Technology 36 (2005) 605–611
Effects of phthalic anhydride modification on horseradish peroxidase stability and structure Hai-Yan Songa , Jun-Hua Yaob , Jian-Zhong Liua,∗ , Shao-Jun Zhoua , Ya-Hong Xionga , Liang-Nian Jia,∗ a
Key Laboratory of Gene Engineering of Ministration of Education and Biotechnology Research Center, Zhongshan University, Guangzhou 510275, PR China b Instrumentation Analysis and Research Center, Zhongshan University, Guangzhou 510275, PR China Received 30 August 2004; accepted 16 December 2004
Abstract The thermal stability of phthalic anhydride-modified horseradish peroxidase (HRP) in organic solvents was investigated. The modification increased the tolerance of some organic solvents. In order to investigate the molecular mechanism of the stabilization, the changes between native and modified enzyme were also studied using kinetics and spectroscopic methodology. The modified HRP showed greater affinity and catalytic efficiency in some organic solvents for different substrates than native HRP. The substrate affinity and the catalytic efficiency of native and modified HRP increased with the increases of electron-donating efficiency of substituents at 4-position. The improvements of catalytic properties are related to the changes of the conformation of HRP. The modification changed the environment of both heme and tryptophan, increased ␣-helix content of HRP and decreased the tertiary structure around the aromatic acid residues in HRP. © 2004 Elsevier Inc. All rights reserved. Keywords: Horseradish peroxidase; Stability; Organic solvent; Conformation; Kinetics; Chemical modification; Phthalic anhydride; CD; Fluorescence; Electron absorption spectra
1. Introduction Horseradish peroxidase (HRP, EC 1.11.1.7) catalyzes the oxidation of aromatic compounds by hydrogen peroxide or alkyl hydroperoxide. The native enzyme consists of a single polypeptide chain with 308 amino acid resides, a heme prosthetic group and two Ca2+ ions maintaining enzyme conformation. It was widely applied in the synthesis of fine chemicals and polymer, the removal of toxic phenolics from wastewater [1,2]. To our knowledge, there is no example of the use of HRP as catalyst in industrial organic synthesis. It is mainly due to low stability and low catalytic activity in organic solvents [3]. Over the past decade, nonaqueous enzymology has emerged as a major area of biotechnology research and de∗
Corresponding authors. Tel.: +86 20 84110115; fax: +86 20 84110115. E-mail address: [email protected] (J.-Z. Liu).
0141-0229/$ – see front matter © 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.enzmictec.2004.12.018
velopment. It is investigated by numerous academic and industrial laboratories worldwide and the number of papers devoted to this topic has surged into hundreds. Although today there is little question that enzymes can function in nonaqueous media, reaction rate of enzyme in organic solvents is quite lower than in water. This limits the commercial application of nonaqueous enzymology [4,5]. Over the years, several techniques have been developed to ameliorate this loss of catalytic function, including lyophilization in the presence of lyoprotectants and excipients such as KCl, crown ethers, cyclodextrins and molecular imprinters [5], the use of site-directed mutagenesis and directed evolution [6,7], or chemical modification [7,8]. Chemical modification has now reemerged as a powerful complementary approach to site-directed mutagenesis and directed evolution [7]. Chemical modification of HRP surface has been performed to improve its stability. Acetic acid N-hydroxysuccinimide ester [9] and bifunctional N-
606
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
hydroxysuccinimide ester [10] were successfully employed to modify HRP to increase HRP’s stability in organic solvents. Our previous papers reported that modification of HRP by phthalic anhydride improved HRP’s stability and catalytic activity in aqueous buffer [11,12]. Although O’Brien et al. reported that phthalic anhydride modification enhanced HRP’s stability in DMF and THF [13], any detailed study on the mechanism of stabilization of HRP with chemical modification has not yet been reported. Thus, the aim of this work is to further investigate the stability of phthalic anhydride-modified HRP in some organic solvents. And we attempt to explain the phenomenon of stabilization from a molecular standpoint by following the reaction kinetics and by probing the changes in the enzyme structure by various spectroscopic methods after modification. These are the first reports.
HRP concentration was estimated from its Soret absorbance (molar extinction coefficient at 402 nm = 102 l mmol−1 cm−1 ) [14]. 2.4. Catalytic stability in organic solvents Organic solvent profiles of HRP samples were carried out at 30 and 60 ◦ C with exposure times of 60 and 10 min, respectively. The solvents used were methanol, dimethylformanmide (DMF), tetrahydofuran (THF) and acetonitrile (ACN). Reaction mixtures were set up with increasing percent volumes of organic solvent in 0.01 mol l−1 phosphate buffer (pH 7.0) in 10% (v/v) increments. One hundred microliters were withdrawn from each reaction mixture and assayed under the standard conditions above. 2.5. Kinetics
2. Materials and methods 2.1. Chemicals Horseradish peroxidase was purchased from Shanghai Lizhu Dong Feng Biotechnology Co. Ltd. and had a specific activity of 250 purpurogallin units/mg and RZ = 3.0. Phthalic anhydride (PA, analytical grade) was obtained from Guangzhou Chemical Reagent Factory. All other reagents were of analytic grade. 2.2. Chemical modification Chemical modification by phthalic anhydride was based on our previous method [12]. 0.15 ml of 2 mmol l−1 phthalic anhydride in DMSO in 0.01 mol l−1 phosphate buffer (pH 7.0) and 2 ml of 1 mg ml−1 HRP in 0.01 mol l−1 phosphate buffer (pH 7.0) were mixed. The reaction proceeded at 4 ◦ C for 1 h and was then dialyzed against 0.01 mol l−1 phosphate buffer (pH 7.0) at 4 ◦ C to removal excess reagent. 2.3. Peroxidase activity assay The enzyme activity was assayed by colorimetric method [10]. A reaction mixture containing 10 mmol l−1 phenol, 0.2 mmol l−1 hydrogen peroxide and 2.4 mmol l−1 4aminoantipyrin (4-AAP) in a total volume of 3.0 ml was incubated at 30 ◦ C. All regents were dissolved in 0.01 mol l−1 phosphate buffer (pH 7.0). The reaction was then started by adding 0.1 ml of diluted enzyme solution, and the initial increase in absorbance was monitored at 510 nm during 1 min. Under such conditions, the rate of formation of colored product which absorbs light at a peak wavelength of 510 nm was calculated using a molar extinction coefficient of 7100 l mol−1 cm−1 . One unit of peroxidase activity was defined as the amount of the enzyme consuming 1 mmol of hydrogen peroxide per minute under the assay conditions.
The kinetic experiments were performed using constant enzyme, 4-AAP and H2 O2 concentration as the peroxidase activity assay, and varying the concentration of substrate under the same conditions of activity assay. 2.6. Spectra Native HRP and PA-HRP was dissolved in 10 ml of 0.01 mol l−1 phosphate buffer (pH 7.0). UV–vis absorption spectra was recorded using Shimadzu UV2450 with enzyme concentration of 0.15 mol l−1 . The difference spectra were also detected using the spectrophotometer. The sample (3 ml) contained 0.15 mol l−1 enzyme and 10 mmol l−1 phenol in 0.01 mol l−1 phosphate buffer (pH 7.0) with different proportions of organic solvent. The blank consisted of the same enzyme in the same organic solvent concentration. CD experiments were carried out using a Jasco J810 spectropolarimeter. CD in the UV region (200–250 nm) was monitored with a cell of 2 mm path length with enzyme concentration of 0.5 mol l−1 . CD from near-UV region to visible region (250–700 nm) was monitored using a cell of 2 mm path length with an enzyme concentration of 4.38 mol l−1 . The CD data were expressed in terms of mean residue ellipticity, [θ], in deg cm2 dmol−1 . Fluorescence measurements were carried out using a Hitachi F4500 spectroflurimeter. The excitation and emission slit widths were set to 10 nm.
3. Results and discussion Our previous paper reported that the PA modification of HRP increased its thermostability in aqueous buffer about 10-fold [12]. Solvent tolerances of native and PA-HRP were compared in DMF, THF, ACN and methanol (Fig. 1). PAHRP showed a greater tolerance of DMF and ACN at 30 ◦ C and of THF, ACN and methanol at 60 ◦ C. Some papers also reported chemical modification improved the stability of HRP in some organic solvents. Acetylated HRP by acetic acid N-
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
hydroxysuccinimide had a greater tolerance of THF at 25 ◦ C and of DMF and methanol at 60 ◦ C [9]. The cross-linked EGHRP showed a greater tolerance of DMF and THF at 25 and 60 ◦ C [10]. O’Brien et al. reported that PA-HRP showed a greater tolerance of DMF at room temperature and of THF during a 10 min incubation at 65 ◦ C [13]. The interest results
607
are the activations of 10% acetonitrile and 20% methanol on PA-HRP both at 30 and 60 ◦ C. The activation of methanol was also found in acetylated HRP [9] and cross-linked EG-HRP [10]. In addition, the activation of acetonitrile was greater than that of methanol. Khmelnitsky et al. have reported numerous examples of enzyme activation by moderate concen-
Fig. 1. Effects of solvents on native () and PA-HRP (䊉) at 30 ◦ C for 60 min (A–C) and 60 ◦ C for 10 min (D–F).
608
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
Table 1 Kinetic parameters of native and modified horseradish peroxidase in 10% of organic solvents Solvent
HRP Km
DMF THF ACN Dioxane
PA-HRP
(mmol l−1 )
kcat
(min−1 )
1.56 × 103 2.42 × 103 2.03 × 103 7.54 × 103
16.09 3.78 17.89 20.71
kcat /Km
(l mmol−1
min−1 )
9.70 × 10 6.40 × 102 1.13 × 102 3.64 × 102
trations (10–30%) of solvents [15]. We also find that native and modified HRP were more stable in ACN than in other solvents tested at 30 ◦ C and were more stable in methanol than in other solvents tested at 60 ◦ C. HRP contains six lysine residues, Lys65, Lys84, Lys149, Lys174, Lys232 and Lys241 [16]. Lys174 is thought to interact with the heme prosthetic group. Very little of Lys84 is exposed, Lys65 and Lys149 are moderately exposed, only Lys174, Lys232 and Lys241 are well accessible [17]. In the cross-linked EG-HRP, Lys232 was completely modified and Lys174 and Lys241 were partly modified [18]. They thought the greater stability of EG-HRP likely arises from increased structural rigidity due to formation of a cross-link between Lys232 and 241. Another explanation is the modification with EGNHS neutralized positive lysine charges and formed tighter binding of a structural calcium ion. However, phthalic anhydride cannot form a cross-link. It simply covalently modified some HRP lysines. Our previous paper reported that the modification degree of amino groups from HRP with phthalic anhydride was about 52%. It indicates that three of the sixlysine -amino groups from native enzyme were modified with phthalic anhydride. PA modification also introduced a bulky benzene ring structure onto the lysine side chain. Thus, the enhanced stability of PA-HRP in some organic solvents may result from the neutralization of the positive charge on three HRP lysines and the introducing of benzene ring. Kinetic constants for phenol oxidation with native and PA-modified HRP in 10% of organic solvents are reported in Table 1. As shown, the modification increased the substrate affinity (Km ) and the catalytic efficiency (kcat /Km ) in organic solvents. In order to assay the catalytic activity of PA-HRP for different substrates, the oxidation of representative phenolic
Km (mmol l−1 )
kcat (min−1 )
kcat /Km (l mmol−1 min−1 )
9.98 1.96 20.51 8.73
1.04 × 103 1.67 × 103 2.38 × 103 3.24 × 103
1.02 × 102 8.52 × 102 1.16 × 102 3.71 × 102
Scheme 1.
substrates was chosen. The phenolics used in this work are phenol, 4-aminophenol, 4-methylphenol and 4-nitrophenol (Scheme 1 ). The kinetic parameters for these phenolics are shown in Table 2. Either in water or DMF solvents, substrate affinity and the catalytic efficiency of native and PA-modified HRP increased with the increases of electron-donating efficiency of substituents at 4-position. The similar results in THF, ACN or dioxane were obtained (data not shown). Generally, the reduction of compound II with a donor substrate is the rate-limiting step in peroxidase catalysis [19]. Electrondonating group increases electron density of oxygen atom of hydroxy group of substrate and then accelerates the reduction of compound II with the substrate, increasing enzyme activity. The effect of amino group is greater than that of methyl group. Ryu and Dordick also found that electrondonating substituents activated HRP catalysis both in water and organic solvents [20]. From Table 2, we also find that PA modification increased the affinity and the catalytic efficiency for these substrates in organic solvents. The effect might be caused by the higher surface hydrophobicity of PAHRP, which in turn may be reflected in a more favorable partition of these hydrophobic substrates to the active site.
Table 2 Kinetic parameters of native and modified horseradish peroxidase for oxidation of different substrates Solvent
Substrate
HRP Km
(mmol l−1 )
PA-HRP kcat
(min−1 )
kcat /Km
(l mmol−1
min−1 )
Km (mmol l−1 )
kcat (min−1 )
kcat /Km (l mmol−1 min−1 )
PSBa
4-AP 4-MP Phenol 4-NP
8.93 17.91 13.42 12.50
1.54 × 105 1.34 × 104 5.98 × 103 1.74 × 10
1.72 × 104 7.48 × 102 4.46 × 102 1.39
7.00 17.40 9.04 11.57
1.80 × 105 1.66 × 104 5.00 × 103 2.60 × 10
2.57 × 104 9.54 × 102 5.53 × 102 2.25
DMF
4-AP 4-MP Phenol 4-NP
6.85 14.16 16.09 7.92
6.78 × 104 8.20 × 103 1.56 × 103 3.62 × 10
9.90 × 103 5.79 × 102 9.70 × 10 4.57
4.49 6.15 9.98 9.57
6.41 × 104 6.93 × 103 1.04 × 103 6.27 × 10
1.43 × 104 1.13 × 103 1.04 × 102 6.55
a
0.01 mol l−1 phosphate buffer (pH 7.0).
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
609
Fig. 2. Absorbance spectra (A) and difference spectra (B) of native (solid line) and PA-HRP (dashed line) in 0.01 mol l−1 phosphate buffer (7.0).
The enhancement of catalytic activity resulted from the increase of surface hydrophobicity has been reported in other heme-proteins [21–23]. CD and absorption spectra in UV and UV–vis regions provide information on the structure of apo-protein and prosthetic heme of HRP [24]. Spectroscopic data used for detection of changes in HRP structure are as follows: (1) far-UV (190–260 nm) CD: changes in secondary structure of the apoprotein. (2) Near-UV (250–300 nm) CD: changes in tertiary structure of the apo-protein. (3) CD at 350–450 nm: dissociation of the prosthetic heme from the apo-protein and the structural changes of the apo-protein surrounding the heme. (4) Absorption at 350–700 nm: changes of the structure and environment of the heme. The PA modification did not bring about the shift position of Soret band (403 nm). But it resulted in shifting of the Q band (from 496 and 640 to 488 and 636 nm, respectively, Fig. 2A). The results indicate that the modification changed the environment of heme. Substrate can interact with the heme group of HRP as ligand, detected
by different spectra. Fig. 2B shows the different spectra of native or PA-modified HRP with phenol in phosphate buffer. PA-HRP showed a stronger interaction than native HRP. Similar results in organic solvents, such as DMF, THF, ACN and dioxane, were also obtained (data not shown). The substrate interactions were correlated with the increases of the substrate affinity and the catalytic efficiency both in aqueous buffer and in organic solvent after modification with PA. Torres et al. also reported that the interactions of pyrene with cytochrome c decreased with the increase of the concentration of organic solvent and were correlated with the decrease of activity when the concentration of organic solvent was increased [25]. CD spectroscopy has proved to be a useful probe of structure of protein in aqueous/organic media. Fig. 3 shows the CD of native and modified HRP in water. Native and modified HRP have identical CD spectra with negative bands at 208 and 220 nm, which agreed with the previous results [26]. The intensity of the CD bands at 208 and 222 nm increased
Fig. 3. CD spectra of native (solid line) and PA-HRP (dashed line) in 0.01 mol l−1 phosphate buffer (7.0).
610
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
Fig. 4. Fluorescence spectra of native (solid line) and PA-HRP (dashed line) in 0.01 mol l−1 phosphate buffer (7.0). Excitation wavelength was 295 nm; enzyme concentration was 7.4 mol l−1 .
(Fig. 3A) after modification indicating an increase of ␣-helix content. In near-UV region (250–350 nm), native and modified HRP have identical CD spectra with a negative band at 284 nm and a positive band at 312 nm. The intensity of the CD band at 284 nm decreased and that at 312 nm increased after modification (Fig. 3B), suggesting a loss of tertiary structure around the aromatic amino acid residues in HRP after modification. In visible region, no change of CD spectra at 403 nm was detected after modification; however, there were some changes of CD spectra in 500–700 nm region (Fig. 3B). The result is consistent with that of absorption bands above. It indicates that there is little change of the structure and environment of the heme after modification. CD spectra analysis has already been successfully used to characterize the influence of temperature and pH [27], H2 O2 [28] and organic solvent [29] on the secondary structure and activity of HRP. In these literatures [27–29], the similar trend of the intensity of CD band in UV region was correlated with enhancement of HRP’s activity. The intrinsic fluorescence of the enzyme is highly dependent on the fluorescence energy transfer from tryptophan to heme [27,30]. Thus, changes in the structure of the heme cavity affecting the distance/orientation between the heme and the tryptophan can affect the intrinsic fluorescence of the enzyme. HRP contains one tryptophan residue, Trp 117, and five tyrosine residues [16]. Both tryptophan and tyrosine residues show fluorescence. The fluorescence emission excited at 295 nm arises solely from the tryptophan residue of HRP [31]. A change in the microenvironment surrounding the tryptophan (Try 117) residue changes the emission maximum of the tryptophan fluorescence [27]. Thus, the tryptophan fluorescence was used to probe the structural change of PA-HRP compared to native HRP. Fig. 4 shows the fluorescence spectra of native and PA-modified HRP at room temperature with excitation at 295 nm (Fig. 4). The tryptophan fluorescence emission of native HRP was at 331 nm
at room temperature, which red shifted to 335 nm after PA modification (Fig. 4). The intensity of the tryptophan fluorescence emission increased after PA-modification because of a change in the relative orientation or distance between the heme and the tryptophan residue leading to a decrease in the efficiency of energy transfer [27]. This also denotes that the distance between the heme and the tryptophan residue increased [32]. Another explanation may be PA modification introduced a bulky benzene ring structure onto the lysine side chain. Fig. 4 also shows that the tryptophan fluorescence emission shifted to 335 nm from 331 nm after PA modification. The red shift indicates that the tryptophan residue became to more exposed and located in a more polar environment [27,32]. Heme absorption, CD and fluorescence are a useful conformation probe for the studying of heme-proteins. Since there is no any report on spectra of chemical modified HRP, there are not many scopes for the comparison of the present result with the reported ones. However, the changes of modified other heme-protein have been detected using spectroscopic technique. PEG-Met-hemoglobin showed up to 10 times higher activity, greater substrate affinity and the catalytic efficiency (kcat /Km ) than unmodified protein [21]. They also found that there was a red shift on the Soret band after modification. Carboxymethylated cytochrome c had greater catalytic activity and higher tryptophan fluorescence intensity than native cytochrome c [33]. Carboxymethylated cytochrome c also showed the red shift in Soret, ␣ and  bands relative to native cytochrome c. These results are almost consistent with that in this paper.
Acknowledgment We are grateful to National Natural Science Foundation of China and the Natural Science Foundation of Guangdong Province for their financial support.
References [1] Colonna S, Gaggero N, Richelmi C, Pasta P. Recent biotechnological developments in the use of peroxidases. Trends Biotechnol 1999;17:163–8. [2] Klibanov AM, Tu TM, Scoot KP. Peroxidase-catalyzed removal of phenols from coal conversion waste waters. Science 1983;221:259–61. [3] van de Velde F, van Rantwijk F, Sheldon RA. Improving the catalytic performance of peroxidases in organic synthesis. Trends Biotechnol 2001;19:73–80. [4] Klibanov AM. Why are enzymes less active in organic solvents than in water. Trends Biotechnol 1997;15:97–101. [5] Lee M-YJ, Dordick S. Enzyme activation for nonaqueous media. Curr Opin Biotechnol 2002;13:376–84. [6] Hult K, Berglund P. Engineering enzymes for improved organic synthesis. Curr Opin Biotechnol 2003;14:395–400. [7] DeSantis G, Jones JB. Chemical modification of enzymes for enhanced functionality. Curr Opin Biotechnol 1999;10:324–30.
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611 [8] Davis BG. Chemical modification of biocatalysts. Curr Opin Biotechnol 2003;14:379–86. ´ ag´ain C. Increased thermal and sol[9] Miland E, Smyth MR, O’F´ vent tolerance of acetylated horseradish peroxidase. Enzyme Microb Technol 1996;19:63–7. ´ ag´ain C. Modification of horseradish per[10] Miland E, Smyth MR, O’F´ oxidase with bifunctional N-hydroxysuccinimide esters: effects on molecular stability. Enzyme Microb Technol 1996;19:242–9. [11] Song H-Y, Liu J-Z, Xiong Y-H, Weng L-P, Ji L-N. Treatment of aqueous chlorophenol by phthalic anhydride-modified horseradish peroxidase. J Mol Catal B: Enzym 2003;22:37–44. [12] Liu J-Z, Song H-Y, Weng L-P, Ji L-N. Increased thermostability and phenol removal efficiency by chemical modified horseradish peroxidase. J Mol Catal B: Enzym 2002;18:225–32. ´ ag´ain C. Effects of phthalic anhy[13] O’Brien AM, Smith AT, O’F´ dride modification on horseradish peroxidase stability and activity. Biotechnol Bioeng 2003;81:233–40. [14] Smith AT, Sanders SA, Thorneley RNF, Burke JF, Bray RC. Characterisation of a haem active-site mutant of horseradish peroxidase, Phe41Val, with altered reactivity towards hydrogen peroxide and reducing substrates. Eur J Biochem 1992;207:507–19. [15] Khmelnitsky YL, Levashov LV, Klyachko NL, Martinek K. Engineering biocatalytic systems in organic media with low water content. Enzyme Microb Technol 1988;10:710–724. [16] Welinder KG. Amino acid sequence studies of horseradish peroxidase. Eur J Biochem 1979;93:483–502. [17] Gajhede M, Schuller DJ, Henrisksen A, Smith AT, Poulos TL. Crys´˚ resolution. Nat tal structure of horseradish peroxidase at 2.15 A Struct Biol 1997;4:1032–8. ´ ag´ain C, Nielsen PF, Welinder KG. Location of [18] O’Brien AM, O’F´ crosslinks in chemically stabilized horseradish peroxidase: implications for design of crosslinks. Biotechnol Bioeng 2001;76:277–84. [19] Colonna S, Gaggero N, Richelmi C, Pasta P. Recent biotechnological developments in the use of peroxidases. Trends Biotechnol 1999;17:163–8. [20] Ryu K, Dordick JS. How do organic solvents affect peroxidase structure and function? Biochemistry 1992;31:2588–98. [21] Torres E, Vazquez-Duhalt R. Chemical modification of hemoglobin improves biocatalytic oxidation of PHAs. Biochem Biophys Res Commun 2000;273:820–3.
611
[22] Tinoco R, Vazquez-Duhalt R. Chemical modification of cytochrome c improves their catalytic properties in oxidation of polycyclic aromatic hydrocarbons. Enzyme Microb Technol 1998;22: 8–12. [23] Garcia-Arellano H, Valderrama B, Saab-Rinc´on G, Vazquez-Duhalt R. High temperature. Bioconjugate Chem 2002;13:1336–44. [24] Strickland EH. Circular dichroism of horseradish peroxidase and its enzyme–substrate compounds. Biochim Biophys Acta 1968;151:70–5. [25] Torres E, Tinoco R, Vazquez-Duhalt R. Solvent hydrophobicity predicts biocatalytic behaviour of lignin peroxidase and cytochrome c in aqueous solution of water-miscible organic solvents. J Biotechnol 1996;49:59–67. [26] Strickland EH, Kay E, Shannon LM, Horwitz J. Peroxidase isoenzymes from horseradish roots III circular dichroism of isoenzymes and apoisoenzymes. J Biol Chem 1968;243:3560–5. [27] Chattopadhyay K, Mazumdar S. Structural and conformational stability of horseradish peroxidase: effect of temperature and pH. Biochemistry 2000;39:263–70. [28] Schmidt A, Schumacher JT, Reichelt J, Hecht H-J, Bilitewski U. Mechanistic and molecular investigations on stabilization of horseradish peroxidase C. Anal Chem 2002;74:3037–45. [29] Akita M, Tsutsumi D, Kobayashi M, Kise H. Structural change and catalytic activity of horseradish peroxidase in oxidative polymerization of phenol. Biosci Biotechnol Biochem 2001;65:1581–8. [30] Pina DG, Shnyrova AV, Gavilanes F, Rodriguez A, Leal F, Roig MG, et al. Thermally induced conformational changes in horseradish peroxidase. Eur J Biochem 2001;268:120–6. [31] Ugarova NN, Savitski AP, Berezin IV. The protoporphyrin– apoperoxidase complex as a horseradish peroxidase analog: a fluorescence study of the heme pocket. Biochim Biophys Acta 1981;662:210–9. [32] Carvalho ASL, e Melo EP, Ferreira BS, Neves-Petersen MT, Petersen SB, Aires-Barros MR. Heme and pH-dependent stability of an anionic horseradish peroxidase. Arch Biochem Biophys 2003;415:257–67. [33] Prasad S, Maiti NC, Mazumdar S, Mitra S. Reaction of hydrogen peroxide and peroxidase activity in carboxymethylated cytochrome c: spectroscopic and kinetic studies. Biochim Biophys Acta 2002;1596:63–75.
Effects of phthalic anhydride modification on horseradish peroxidase stability and structure Hai-Yan Songa , Jun-Hua Yaob , Jian-Zhong Liua,∗ , Shao-Jun Zhoua , Ya-Hong Xionga , Liang-Nian Jia,∗ a
Key Laboratory of Gene Engineering of Ministration of Education and Biotechnology Research Center, Zhongshan University, Guangzhou 510275, PR China b Instrumentation Analysis and Research Center, Zhongshan University, Guangzhou 510275, PR China Received 30 August 2004; accepted 16 December 2004
Abstract The thermal stability of phthalic anhydride-modified horseradish peroxidase (HRP) in organic solvents was investigated. The modification increased the tolerance of some organic solvents. In order to investigate the molecular mechanism of the stabilization, the changes between native and modified enzyme were also studied using kinetics and spectroscopic methodology. The modified HRP showed greater affinity and catalytic efficiency in some organic solvents for different substrates than native HRP. The substrate affinity and the catalytic efficiency of native and modified HRP increased with the increases of electron-donating efficiency of substituents at 4-position. The improvements of catalytic properties are related to the changes of the conformation of HRP. The modification changed the environment of both heme and tryptophan, increased ␣-helix content of HRP and decreased the tertiary structure around the aromatic acid residues in HRP. © 2004 Elsevier Inc. All rights reserved. Keywords: Horseradish peroxidase; Stability; Organic solvent; Conformation; Kinetics; Chemical modification; Phthalic anhydride; CD; Fluorescence; Electron absorption spectra
1. Introduction Horseradish peroxidase (HRP, EC 1.11.1.7) catalyzes the oxidation of aromatic compounds by hydrogen peroxide or alkyl hydroperoxide. The native enzyme consists of a single polypeptide chain with 308 amino acid resides, a heme prosthetic group and two Ca2+ ions maintaining enzyme conformation. It was widely applied in the synthesis of fine chemicals and polymer, the removal of toxic phenolics from wastewater [1,2]. To our knowledge, there is no example of the use of HRP as catalyst in industrial organic synthesis. It is mainly due to low stability and low catalytic activity in organic solvents [3]. Over the past decade, nonaqueous enzymology has emerged as a major area of biotechnology research and de∗
Corresponding authors. Tel.: +86 20 84110115; fax: +86 20 84110115. E-mail address: [email protected] (J.-Z. Liu).
0141-0229/$ – see front matter © 2004 Elsevier Inc. All rights reserved. doi:10.1016/j.enzmictec.2004.12.018
velopment. It is investigated by numerous academic and industrial laboratories worldwide and the number of papers devoted to this topic has surged into hundreds. Although today there is little question that enzymes can function in nonaqueous media, reaction rate of enzyme in organic solvents is quite lower than in water. This limits the commercial application of nonaqueous enzymology [4,5]. Over the years, several techniques have been developed to ameliorate this loss of catalytic function, including lyophilization in the presence of lyoprotectants and excipients such as KCl, crown ethers, cyclodextrins and molecular imprinters [5], the use of site-directed mutagenesis and directed evolution [6,7], or chemical modification [7,8]. Chemical modification has now reemerged as a powerful complementary approach to site-directed mutagenesis and directed evolution [7]. Chemical modification of HRP surface has been performed to improve its stability. Acetic acid N-hydroxysuccinimide ester [9] and bifunctional N-
606
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
hydroxysuccinimide ester [10] were successfully employed to modify HRP to increase HRP’s stability in organic solvents. Our previous papers reported that modification of HRP by phthalic anhydride improved HRP’s stability and catalytic activity in aqueous buffer [11,12]. Although O’Brien et al. reported that phthalic anhydride modification enhanced HRP’s stability in DMF and THF [13], any detailed study on the mechanism of stabilization of HRP with chemical modification has not yet been reported. Thus, the aim of this work is to further investigate the stability of phthalic anhydride-modified HRP in some organic solvents. And we attempt to explain the phenomenon of stabilization from a molecular standpoint by following the reaction kinetics and by probing the changes in the enzyme structure by various spectroscopic methods after modification. These are the first reports.
HRP concentration was estimated from its Soret absorbance (molar extinction coefficient at 402 nm = 102 l mmol−1 cm−1 ) [14]. 2.4. Catalytic stability in organic solvents Organic solvent profiles of HRP samples were carried out at 30 and 60 ◦ C with exposure times of 60 and 10 min, respectively. The solvents used were methanol, dimethylformanmide (DMF), tetrahydofuran (THF) and acetonitrile (ACN). Reaction mixtures were set up with increasing percent volumes of organic solvent in 0.01 mol l−1 phosphate buffer (pH 7.0) in 10% (v/v) increments. One hundred microliters were withdrawn from each reaction mixture and assayed under the standard conditions above. 2.5. Kinetics
2. Materials and methods 2.1. Chemicals Horseradish peroxidase was purchased from Shanghai Lizhu Dong Feng Biotechnology Co. Ltd. and had a specific activity of 250 purpurogallin units/mg and RZ = 3.0. Phthalic anhydride (PA, analytical grade) was obtained from Guangzhou Chemical Reagent Factory. All other reagents were of analytic grade. 2.2. Chemical modification Chemical modification by phthalic anhydride was based on our previous method [12]. 0.15 ml of 2 mmol l−1 phthalic anhydride in DMSO in 0.01 mol l−1 phosphate buffer (pH 7.0) and 2 ml of 1 mg ml−1 HRP in 0.01 mol l−1 phosphate buffer (pH 7.0) were mixed. The reaction proceeded at 4 ◦ C for 1 h and was then dialyzed against 0.01 mol l−1 phosphate buffer (pH 7.0) at 4 ◦ C to removal excess reagent. 2.3. Peroxidase activity assay The enzyme activity was assayed by colorimetric method [10]. A reaction mixture containing 10 mmol l−1 phenol, 0.2 mmol l−1 hydrogen peroxide and 2.4 mmol l−1 4aminoantipyrin (4-AAP) in a total volume of 3.0 ml was incubated at 30 ◦ C. All regents were dissolved in 0.01 mol l−1 phosphate buffer (pH 7.0). The reaction was then started by adding 0.1 ml of diluted enzyme solution, and the initial increase in absorbance was monitored at 510 nm during 1 min. Under such conditions, the rate of formation of colored product which absorbs light at a peak wavelength of 510 nm was calculated using a molar extinction coefficient of 7100 l mol−1 cm−1 . One unit of peroxidase activity was defined as the amount of the enzyme consuming 1 mmol of hydrogen peroxide per minute under the assay conditions.
The kinetic experiments were performed using constant enzyme, 4-AAP and H2 O2 concentration as the peroxidase activity assay, and varying the concentration of substrate under the same conditions of activity assay. 2.6. Spectra Native HRP and PA-HRP was dissolved in 10 ml of 0.01 mol l−1 phosphate buffer (pH 7.0). UV–vis absorption spectra was recorded using Shimadzu UV2450 with enzyme concentration of 0.15 mol l−1 . The difference spectra were also detected using the spectrophotometer. The sample (3 ml) contained 0.15 mol l−1 enzyme and 10 mmol l−1 phenol in 0.01 mol l−1 phosphate buffer (pH 7.0) with different proportions of organic solvent. The blank consisted of the same enzyme in the same organic solvent concentration. CD experiments were carried out using a Jasco J810 spectropolarimeter. CD in the UV region (200–250 nm) was monitored with a cell of 2 mm path length with enzyme concentration of 0.5 mol l−1 . CD from near-UV region to visible region (250–700 nm) was monitored using a cell of 2 mm path length with an enzyme concentration of 4.38 mol l−1 . The CD data were expressed in terms of mean residue ellipticity, [θ], in deg cm2 dmol−1 . Fluorescence measurements were carried out using a Hitachi F4500 spectroflurimeter. The excitation and emission slit widths were set to 10 nm.
3. Results and discussion Our previous paper reported that the PA modification of HRP increased its thermostability in aqueous buffer about 10-fold [12]. Solvent tolerances of native and PA-HRP were compared in DMF, THF, ACN and methanol (Fig. 1). PAHRP showed a greater tolerance of DMF and ACN at 30 ◦ C and of THF, ACN and methanol at 60 ◦ C. Some papers also reported chemical modification improved the stability of HRP in some organic solvents. Acetylated HRP by acetic acid N-
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
hydroxysuccinimide had a greater tolerance of THF at 25 ◦ C and of DMF and methanol at 60 ◦ C [9]. The cross-linked EGHRP showed a greater tolerance of DMF and THF at 25 and 60 ◦ C [10]. O’Brien et al. reported that PA-HRP showed a greater tolerance of DMF at room temperature and of THF during a 10 min incubation at 65 ◦ C [13]. The interest results
607
are the activations of 10% acetonitrile and 20% methanol on PA-HRP both at 30 and 60 ◦ C. The activation of methanol was also found in acetylated HRP [9] and cross-linked EG-HRP [10]. In addition, the activation of acetonitrile was greater than that of methanol. Khmelnitsky et al. have reported numerous examples of enzyme activation by moderate concen-
Fig. 1. Effects of solvents on native () and PA-HRP (䊉) at 30 ◦ C for 60 min (A–C) and 60 ◦ C for 10 min (D–F).
608
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
Table 1 Kinetic parameters of native and modified horseradish peroxidase in 10% of organic solvents Solvent
HRP Km
DMF THF ACN Dioxane
PA-HRP
(mmol l−1 )
kcat
(min−1 )
1.56 × 103 2.42 × 103 2.03 × 103 7.54 × 103
16.09 3.78 17.89 20.71
kcat /Km
(l mmol−1
min−1 )
9.70 × 10 6.40 × 102 1.13 × 102 3.64 × 102
trations (10–30%) of solvents [15]. We also find that native and modified HRP were more stable in ACN than in other solvents tested at 30 ◦ C and were more stable in methanol than in other solvents tested at 60 ◦ C. HRP contains six lysine residues, Lys65, Lys84, Lys149, Lys174, Lys232 and Lys241 [16]. Lys174 is thought to interact with the heme prosthetic group. Very little of Lys84 is exposed, Lys65 and Lys149 are moderately exposed, only Lys174, Lys232 and Lys241 are well accessible [17]. In the cross-linked EG-HRP, Lys232 was completely modified and Lys174 and Lys241 were partly modified [18]. They thought the greater stability of EG-HRP likely arises from increased structural rigidity due to formation of a cross-link between Lys232 and 241. Another explanation is the modification with EGNHS neutralized positive lysine charges and formed tighter binding of a structural calcium ion. However, phthalic anhydride cannot form a cross-link. It simply covalently modified some HRP lysines. Our previous paper reported that the modification degree of amino groups from HRP with phthalic anhydride was about 52%. It indicates that three of the sixlysine -amino groups from native enzyme were modified with phthalic anhydride. PA modification also introduced a bulky benzene ring structure onto the lysine side chain. Thus, the enhanced stability of PA-HRP in some organic solvents may result from the neutralization of the positive charge on three HRP lysines and the introducing of benzene ring. Kinetic constants for phenol oxidation with native and PA-modified HRP in 10% of organic solvents are reported in Table 1. As shown, the modification increased the substrate affinity (Km ) and the catalytic efficiency (kcat /Km ) in organic solvents. In order to assay the catalytic activity of PA-HRP for different substrates, the oxidation of representative phenolic
Km (mmol l−1 )
kcat (min−1 )
kcat /Km (l mmol−1 min−1 )
9.98 1.96 20.51 8.73
1.04 × 103 1.67 × 103 2.38 × 103 3.24 × 103
1.02 × 102 8.52 × 102 1.16 × 102 3.71 × 102
Scheme 1.
substrates was chosen. The phenolics used in this work are phenol, 4-aminophenol, 4-methylphenol and 4-nitrophenol (Scheme 1 ). The kinetic parameters for these phenolics are shown in Table 2. Either in water or DMF solvents, substrate affinity and the catalytic efficiency of native and PA-modified HRP increased with the increases of electron-donating efficiency of substituents at 4-position. The similar results in THF, ACN or dioxane were obtained (data not shown). Generally, the reduction of compound II with a donor substrate is the rate-limiting step in peroxidase catalysis [19]. Electrondonating group increases electron density of oxygen atom of hydroxy group of substrate and then accelerates the reduction of compound II with the substrate, increasing enzyme activity. The effect of amino group is greater than that of methyl group. Ryu and Dordick also found that electrondonating substituents activated HRP catalysis both in water and organic solvents [20]. From Table 2, we also find that PA modification increased the affinity and the catalytic efficiency for these substrates in organic solvents. The effect might be caused by the higher surface hydrophobicity of PAHRP, which in turn may be reflected in a more favorable partition of these hydrophobic substrates to the active site.
Table 2 Kinetic parameters of native and modified horseradish peroxidase for oxidation of different substrates Solvent
Substrate
HRP Km
(mmol l−1 )
PA-HRP kcat
(min−1 )
kcat /Km
(l mmol−1
min−1 )
Km (mmol l−1 )
kcat (min−1 )
kcat /Km (l mmol−1 min−1 )
PSBa
4-AP 4-MP Phenol 4-NP
8.93 17.91 13.42 12.50
1.54 × 105 1.34 × 104 5.98 × 103 1.74 × 10
1.72 × 104 7.48 × 102 4.46 × 102 1.39
7.00 17.40 9.04 11.57
1.80 × 105 1.66 × 104 5.00 × 103 2.60 × 10
2.57 × 104 9.54 × 102 5.53 × 102 2.25
DMF
4-AP 4-MP Phenol 4-NP
6.85 14.16 16.09 7.92
6.78 × 104 8.20 × 103 1.56 × 103 3.62 × 10
9.90 × 103 5.79 × 102 9.70 × 10 4.57
4.49 6.15 9.98 9.57
6.41 × 104 6.93 × 103 1.04 × 103 6.27 × 10
1.43 × 104 1.13 × 103 1.04 × 102 6.55
a
0.01 mol l−1 phosphate buffer (pH 7.0).
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
609
Fig. 2. Absorbance spectra (A) and difference spectra (B) of native (solid line) and PA-HRP (dashed line) in 0.01 mol l−1 phosphate buffer (7.0).
The enhancement of catalytic activity resulted from the increase of surface hydrophobicity has been reported in other heme-proteins [21–23]. CD and absorption spectra in UV and UV–vis regions provide information on the structure of apo-protein and prosthetic heme of HRP [24]. Spectroscopic data used for detection of changes in HRP structure are as follows: (1) far-UV (190–260 nm) CD: changes in secondary structure of the apoprotein. (2) Near-UV (250–300 nm) CD: changes in tertiary structure of the apo-protein. (3) CD at 350–450 nm: dissociation of the prosthetic heme from the apo-protein and the structural changes of the apo-protein surrounding the heme. (4) Absorption at 350–700 nm: changes of the structure and environment of the heme. The PA modification did not bring about the shift position of Soret band (403 nm). But it resulted in shifting of the Q band (from 496 and 640 to 488 and 636 nm, respectively, Fig. 2A). The results indicate that the modification changed the environment of heme. Substrate can interact with the heme group of HRP as ligand, detected
by different spectra. Fig. 2B shows the different spectra of native or PA-modified HRP with phenol in phosphate buffer. PA-HRP showed a stronger interaction than native HRP. Similar results in organic solvents, such as DMF, THF, ACN and dioxane, were also obtained (data not shown). The substrate interactions were correlated with the increases of the substrate affinity and the catalytic efficiency both in aqueous buffer and in organic solvent after modification with PA. Torres et al. also reported that the interactions of pyrene with cytochrome c decreased with the increase of the concentration of organic solvent and were correlated with the decrease of activity when the concentration of organic solvent was increased [25]. CD spectroscopy has proved to be a useful probe of structure of protein in aqueous/organic media. Fig. 3 shows the CD of native and modified HRP in water. Native and modified HRP have identical CD spectra with negative bands at 208 and 220 nm, which agreed with the previous results [26]. The intensity of the CD bands at 208 and 222 nm increased
Fig. 3. CD spectra of native (solid line) and PA-HRP (dashed line) in 0.01 mol l−1 phosphate buffer (7.0).
610
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611
Fig. 4. Fluorescence spectra of native (solid line) and PA-HRP (dashed line) in 0.01 mol l−1 phosphate buffer (7.0). Excitation wavelength was 295 nm; enzyme concentration was 7.4 mol l−1 .
(Fig. 3A) after modification indicating an increase of ␣-helix content. In near-UV region (250–350 nm), native and modified HRP have identical CD spectra with a negative band at 284 nm and a positive band at 312 nm. The intensity of the CD band at 284 nm decreased and that at 312 nm increased after modification (Fig. 3B), suggesting a loss of tertiary structure around the aromatic amino acid residues in HRP after modification. In visible region, no change of CD spectra at 403 nm was detected after modification; however, there were some changes of CD spectra in 500–700 nm region (Fig. 3B). The result is consistent with that of absorption bands above. It indicates that there is little change of the structure and environment of the heme after modification. CD spectra analysis has already been successfully used to characterize the influence of temperature and pH [27], H2 O2 [28] and organic solvent [29] on the secondary structure and activity of HRP. In these literatures [27–29], the similar trend of the intensity of CD band in UV region was correlated with enhancement of HRP’s activity. The intrinsic fluorescence of the enzyme is highly dependent on the fluorescence energy transfer from tryptophan to heme [27,30]. Thus, changes in the structure of the heme cavity affecting the distance/orientation between the heme and the tryptophan can affect the intrinsic fluorescence of the enzyme. HRP contains one tryptophan residue, Trp 117, and five tyrosine residues [16]. Both tryptophan and tyrosine residues show fluorescence. The fluorescence emission excited at 295 nm arises solely from the tryptophan residue of HRP [31]. A change in the microenvironment surrounding the tryptophan (Try 117) residue changes the emission maximum of the tryptophan fluorescence [27]. Thus, the tryptophan fluorescence was used to probe the structural change of PA-HRP compared to native HRP. Fig. 4 shows the fluorescence spectra of native and PA-modified HRP at room temperature with excitation at 295 nm (Fig. 4). The tryptophan fluorescence emission of native HRP was at 331 nm
at room temperature, which red shifted to 335 nm after PA modification (Fig. 4). The intensity of the tryptophan fluorescence emission increased after PA-modification because of a change in the relative orientation or distance between the heme and the tryptophan residue leading to a decrease in the efficiency of energy transfer [27]. This also denotes that the distance between the heme and the tryptophan residue increased [32]. Another explanation may be PA modification introduced a bulky benzene ring structure onto the lysine side chain. Fig. 4 also shows that the tryptophan fluorescence emission shifted to 335 nm from 331 nm after PA modification. The red shift indicates that the tryptophan residue became to more exposed and located in a more polar environment [27,32]. Heme absorption, CD and fluorescence are a useful conformation probe for the studying of heme-proteins. Since there is no any report on spectra of chemical modified HRP, there are not many scopes for the comparison of the present result with the reported ones. However, the changes of modified other heme-protein have been detected using spectroscopic technique. PEG-Met-hemoglobin showed up to 10 times higher activity, greater substrate affinity and the catalytic efficiency (kcat /Km ) than unmodified protein [21]. They also found that there was a red shift on the Soret band after modification. Carboxymethylated cytochrome c had greater catalytic activity and higher tryptophan fluorescence intensity than native cytochrome c [33]. Carboxymethylated cytochrome c also showed the red shift in Soret, ␣ and  bands relative to native cytochrome c. These results are almost consistent with that in this paper.
Acknowledgment We are grateful to National Natural Science Foundation of China and the Natural Science Foundation of Guangdong Province for their financial support.
References [1] Colonna S, Gaggero N, Richelmi C, Pasta P. Recent biotechnological developments in the use of peroxidases. Trends Biotechnol 1999;17:163–8. [2] Klibanov AM, Tu TM, Scoot KP. Peroxidase-catalyzed removal of phenols from coal conversion waste waters. Science 1983;221:259–61. [3] van de Velde F, van Rantwijk F, Sheldon RA. Improving the catalytic performance of peroxidases in organic synthesis. Trends Biotechnol 2001;19:73–80. [4] Klibanov AM. Why are enzymes less active in organic solvents than in water. Trends Biotechnol 1997;15:97–101. [5] Lee M-YJ, Dordick S. Enzyme activation for nonaqueous media. Curr Opin Biotechnol 2002;13:376–84. [6] Hult K, Berglund P. Engineering enzymes for improved organic synthesis. Curr Opin Biotechnol 2003;14:395–400. [7] DeSantis G, Jones JB. Chemical modification of enzymes for enhanced functionality. Curr Opin Biotechnol 1999;10:324–30.
H.-Y. Song et al. / Enzyme and Microbial Technology 36 (2005) 605–611 [8] Davis BG. Chemical modification of biocatalysts. Curr Opin Biotechnol 2003;14:379–86. ´ ag´ain C. Increased thermal and sol[9] Miland E, Smyth MR, O’F´ vent tolerance of acetylated horseradish peroxidase. Enzyme Microb Technol 1996;19:63–7. ´ ag´ain C. Modification of horseradish per[10] Miland E, Smyth MR, O’F´ oxidase with bifunctional N-hydroxysuccinimide esters: effects on molecular stability. Enzyme Microb Technol 1996;19:242–9. [11] Song H-Y, Liu J-Z, Xiong Y-H, Weng L-P, Ji L-N. Treatment of aqueous chlorophenol by phthalic anhydride-modified horseradish peroxidase. J Mol Catal B: Enzym 2003;22:37–44. [12] Liu J-Z, Song H-Y, Weng L-P, Ji L-N. Increased thermostability and phenol removal efficiency by chemical modified horseradish peroxidase. J Mol Catal B: Enzym 2002;18:225–32. ´ ag´ain C. Effects of phthalic anhy[13] O’Brien AM, Smith AT, O’F´ dride modification on horseradish peroxidase stability and activity. Biotechnol Bioeng 2003;81:233–40. [14] Smith AT, Sanders SA, Thorneley RNF, Burke JF, Bray RC. Characterisation of a haem active-site mutant of horseradish peroxidase, Phe41Val, with altered reactivity towards hydrogen peroxide and reducing substrates. Eur J Biochem 1992;207:507–19. [15] Khmelnitsky YL, Levashov LV, Klyachko NL, Martinek K. Engineering biocatalytic systems in organic media with low water content. Enzyme Microb Technol 1988;10:710–724. [16] Welinder KG. Amino acid sequence studies of horseradish peroxidase. Eur J Biochem 1979;93:483–502. [17] Gajhede M, Schuller DJ, Henrisksen A, Smith AT, Poulos TL. Crys´˚ resolution. Nat tal structure of horseradish peroxidase at 2.15 A Struct Biol 1997;4:1032–8. ´ ag´ain C, Nielsen PF, Welinder KG. Location of [18] O’Brien AM, O’F´ crosslinks in chemically stabilized horseradish peroxidase: implications for design of crosslinks. Biotechnol Bioeng 2001;76:277–84. [19] Colonna S, Gaggero N, Richelmi C, Pasta P. Recent biotechnological developments in the use of peroxidases. Trends Biotechnol 1999;17:163–8. [20] Ryu K, Dordick JS. How do organic solvents affect peroxidase structure and function? Biochemistry 1992;31:2588–98. [21] Torres E, Vazquez-Duhalt R. Chemical modification of hemoglobin improves biocatalytic oxidation of PHAs. Biochem Biophys Res Commun 2000;273:820–3.
611
[22] Tinoco R, Vazquez-Duhalt R. Chemical modification of cytochrome c improves their catalytic properties in oxidation of polycyclic aromatic hydrocarbons. Enzyme Microb Technol 1998;22: 8–12. [23] Garcia-Arellano H, Valderrama B, Saab-Rinc´on G, Vazquez-Duhalt R. High temperature. Bioconjugate Chem 2002;13:1336–44. [24] Strickland EH. Circular dichroism of horseradish peroxidase and its enzyme–substrate compounds. Biochim Biophys Acta 1968;151:70–5. [25] Torres E, Tinoco R, Vazquez-Duhalt R. Solvent hydrophobicity predicts biocatalytic behaviour of lignin peroxidase and cytochrome c in aqueous solution of water-miscible organic solvents. J Biotechnol 1996;49:59–67. [26] Strickland EH, Kay E, Shannon LM, Horwitz J. Peroxidase isoenzymes from horseradish roots III circular dichroism of isoenzymes and apoisoenzymes. J Biol Chem 1968;243:3560–5. [27] Chattopadhyay K, Mazumdar S. Structural and conformational stability of horseradish peroxidase: effect of temperature and pH. Biochemistry 2000;39:263–70. [28] Schmidt A, Schumacher JT, Reichelt J, Hecht H-J, Bilitewski U. Mechanistic and molecular investigations on stabilization of horseradish peroxidase C. Anal Chem 2002;74:3037–45. [29] Akita M, Tsutsumi D, Kobayashi M, Kise H. Structural change and catalytic activity of horseradish peroxidase in oxidative polymerization of phenol. Biosci Biotechnol Biochem 2001;65:1581–8. [30] Pina DG, Shnyrova AV, Gavilanes F, Rodriguez A, Leal F, Roig MG, et al. Thermally induced conformational changes in horseradish peroxidase. Eur J Biochem 2001;268:120–6. [31] Ugarova NN, Savitski AP, Berezin IV. The protoporphyrin– apoperoxidase complex as a horseradish peroxidase analog: a fluorescence study of the heme pocket. Biochim Biophys Acta 1981;662:210–9. [32] Carvalho ASL, e Melo EP, Ferreira BS, Neves-Petersen MT, Petersen SB, Aires-Barros MR. Heme and pH-dependent stability of an anionic horseradish peroxidase. Arch Biochem Biophys 2003;415:257–67. [33] Prasad S, Maiti NC, Mazumdar S, Mitra S. Reaction of hydrogen peroxide and peroxidase activity in carboxymethylated cytochrome c: spectroscopic and kinetic studies. Biochim Biophys Acta 2002;1596:63–75.