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Deactivation of chloroperoxidase by monosaccharides (d -glucose, d -galactose, and d -xylose)
Carbohydrate Research 370 (2013) 72–75
Contents lists available at SciVerse ScienceDirect
Carbohydrate Research journal homepage: www.elsevier.com/locate/carres
Deactivation of chloroperoxidase by monosaccharides (D-glucose, D-galactose, and D-xylose) Rixiao Jin a, Chaonan Li a, Lifei Zhi a, Yucheng Jiang a,b,⇑, Mancheng Hu a,b, Shuni Li a,b, Quanguo Zhai a,b a b
School of Chemistry and Materials Science, Shaanxi Normal University, Chang’an South Road, 199, Xi’an 710062, China Key Laboratory of Macromolecular Science of Shaanxi Province, Shaanxi Normal University, China
a r t i c l e
i n f o
Article history: Received 17 March 2012 Received in revised form 1 July 2012 Accepted 3 July 2012 Available online 14 July 2012 Keywords: Deactivation Chloroperoxidase Monosaccharide
a b s t r a c t In this work, it was found that some monosaccharides normally used to stabilize enzymes at high temperatures, however, actually caused a deactivation of chloroperoxidase (CPO). The red native CPO was converted to a stable pale species that lost enzymatic activity. This deactivation was irreversible and was sensitive to temperature. It was different from the general peroxide-mediated deactivation of CPO. Data from measurement of chlorination activity as well as UV–vis, fluorescent, and CD spectral analysis indicated that monosaccharide-induced deactivation can be attributed to precipitation of protein in the presence of monosaccharide and the interaction of the aldehyde group of sugar with amino groups, especially the terminal amino group, on proteins to form Schiff bases which then rearrange to the stable amino ketone. It is further noted that the deactivation efficiency depends on the stereostructure of monosaccharides. D-Glucose was the most efficient inactivating agents due to its aptitude for both of the interactions mentioned in the above paragraphs. The deactivation was specific to aldose. No deprivation of the heme iron was involved in this deactivation. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Chloroperoxidase (CPO) from Caldariomyces fumago is probably the most versatile known heme enzymes. It catalyzes chlorination of activated C–H bonds and reactions reminiscent of peroxidase, catalase, and cytochrome P450. CPO shares cysteine thiolate heme ligation with cytochrome P-450 monooxygenase family, while possessing a polar active site milieu similar to those of the heme peroxidases and catalase.1,2 The inactivation of CPO reported in literature mainly included the following ways: The first was a general inactivation of typical peroxidases (e.g., horseradish peroxidase) caused by excess oxidants (i.e., hydrogen peroxide) or by double substrates involving H2O2.3–7 Clark et al. described a catalytic cycle of such systems, in which hydrogen peroxide bound to the penta-coordinate ferric resting state Fe(III) to generate Compound I, a oxoferryl porphyrin p cation radical containing Fe(IV). Compound I reacts with hydrogen peroxide either in a catalase-like two-electron reduction, which releases molecular oxygen, or in two single-electron transfers in which Compound II and Compound III were involved as intermediates. During two single-electron reductions of Compound ⇑ Corresponding author. Tel.: +86 029 81530763. E-mail addresses: [email protected] (R. Jin), [email protected] (C. Li), [email protected] (L. Zhi), [email protected] (Y. Jiang), [email protected] (M. Hu), [email protected] (S. Li), [email protected] (Q. Zhai). 0008-6215/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.carres.2012.07.001
I, free radicals and superoxide anion radical such as HO2, HO, were formed. These radicals oxidize the heme and/or protein residues, leading to the inactivation of the enzyme.8 Ayala et al. pointed out that under peroxidasic conditions the most important inactivating event was the destruction of the heme group. Regarding protein oxidation, no major structural disruption such as protein cross-linking or unfolding was detected. Tryptophan (most sensitive to oxidation) oxidation was a simultaneous event, rather than the predominant cause of deactivation.9 Additionally, a catalaselike function was described,10 in which CPO was significantly resistant to inactivation by hydrogen peroxide or by organic hydroperoxides because of its intrinsic catalase activity. Nevertheless, exposure to high concentrations of hydrogen peroxide (>30 mM) could irreversibly inactivate CPO with a half-life of about 1 min. Furthermore, Hager et al. reported a suicidal formation of N-alkylporphyrins during the oxidation of terminal alkenes and alkynes. The inactivated CPO underwent spontaneous loss of the heme alkyl moiety with partial restoration of enzymatic activity.11 Our group reported an arginine-induced inactivation that might be caused mainly by the binding of guanidinium group in arginine to the acid–base catalytic group Glu183 in CPO.12 Recently, we surprisingly discovered that CPO could be inactivated by some monosaccharides (D-glucose, D-galactose, and D-xylose) in the absence of H2O2. It is well known that polyhydroxy compounds, such as monosaccharides and glycerin etc., can improve the thermal stability of enzyme because they can prevent
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enzyme from conformation change at high temperatures. Therefore, this deactivation is very interesting and could provide new insight into the nature of interaction between sugars and CPO. In this paper, UV–vis, fluorescent, and circular dichroism spectral analysis combined with the measurement of chlorination activity was employed to probe D-glucose-mediated deactivation of CPO and the mechanism involved in the deactivation process. 2. Materials and methods 2.1. Materials Chloroperoxidase was isolated from the growth medium of Caldariomyces fumago according to the method established by Morris and Hager13 with minor modifications using acetone rather than ethanol in the solvent fractionation step. The enzyme had an activity of 7800 U/mL based on the standard monochlorodimedon (MCD) chlorination assay. The Rz of CPO used in all experiments was 1.35 (Rz = purity standard = A398/A280 = 1.44 for pure enzyme). D-Glucose, D-galactose, and D-xylose were purchased from Shanghai Chemical and Medicine Co. Ltd. MCD was obtained from Sigma. Citric acid, Sodium citrate, and hydrogen peroxide (30% in aqueous solution) were obtained from Xi’an Chemical Co. Ltd. All chemicals are of analytical grade unless otherwise indicated. Enzyme assay and other optical measurements were performed on a Shimadzu UV-1700 spectrophotometer, 970 CRT fluorescence spectrometry (Shanghai analyze instrument factory) and circular dichroism (CD) spectrometer (Applied Photophysics Ltd), respectively. 2.2. Methods 2.2.1. UV–vis spectroscopy 0.2 mM CPO in buffer at pH 5.5 was incubated with different concentrations of D-glucose as long as 86 h at 4 °C, which is the storage temperature of enzyme. An appropriate dilution was needed before recording its UV–vis spectra (7.2 lM). The spectrum of native CPO with same concentration was recorded for comparison. The concentration of CPO was determined by the absorbance at 398 nm (Soret band) using an extinction coefficient of 91,200 M 1 cm 1. 2.2.2. Fluorescence measurements 0.2 mM CPO in buffer at pH 5.5 was incubated with different concentrations of D-glucose for 86 h at 4 °C. The sample was diluted appropriately (2.0 lM), and then excited at 287 nm. Fluorescence spectra were recorded from 300 to 400 nm. The spectrum of native CPO with same concentration was also recorded for comparison. 2.2.3. Circular dichroism (CD) measurements 7.2 lM CPO sample was prepared as above for CD spectra recording from 190 to 700 nm in a 200 lL quartz cuvette at 4 °C. The operation parameters were as following: path-length: 1.0 mm, time-per-point: 0.25, number of scans: 3, spectral width: 1.0 nm. 3. Results and discussion 3.1. Deactivation of CPO by D-glucose As shown in Figure 1, a red native CPO (0.2 mM) was converted to a pale green species (G-CPO) after 50 h incubation with D-glucose at 4 °C accompanied with spontaneous loss of partial
Figure 1. Red native CPO (right) was converted to a pale green species (left) after 50 h incubation with D-glucose at 4 °C, pH 5.5.
enzymatic activity as judged by the MCD assay. This deactivation was sensitive to temperature, but not sensitive to pH in the range of pH 4.5–5.5 which was the suitable pH for storage of enzyme. G-CPO was stable, and no color change was found after 10 days of standing, indicating the deactivation was an irreversible process. No deprivation of the heme iron was involved in this inactivation as confirmed by the graphite furnace atomic absorption spectrometry measurement. It was also examined that if other monosaccharide (e.g., D-galactose and D-xylose) could cause this deactivation. Similar phenomenon was observed. 3.1.1. UV spectra of G-CPO The iron(II1) porphyrin in CPO is a high-spin complex that displays a Soret band at kmax = 398 nm.14,15 In Figure 2a, the absorption of kmax had no obvious change in the presence of trace amount of D-glucose, but began to decrease obviously when Cglucose/CCPO increased to 200. This decrease in absorption continued with incubation time, meanwhile the absorption shifted from 398 nm to 409 nm gradually. However, the absorption was found to increase again with the peak position shifted back to 398 nm gradually on standing. But the final spectrum was not identical to that of native chloroperoxidase (Fig. 2b and c). The pH value did not change during this process. Moreover, the higher the Cglucose/CCPO was, the faster the decrease in absorbance. Based on the above spectral properties of G-CPO, it was postulated that there were two possible interactions involved in this process. Firstly, it was noted that the monosaccharides tested here (D-glucose, D-galactose, and D-xylose) happened to be all aldoses. So one possible interaction was the interaction of these monosaccharides with amino groups, especially the terminal amino group, on CPO to form Schiff bases which then rearrange to the stable amino ketone, as illustrated in Figure 3. This interaction might account for the observed loss of CPO’s activity due to the destruction of its conformation during this process. To further testify this speculation, fructose was incubated with CPO under the same conditions as used for glucose. As expected, almost no decrease in Soret band and enzymatic activity was observed (Fig. 4), demonstrating that the key functional groups that caused the deactivation of CPO were the –CHO group in sugars. Another reason causing absorption decrease was the precipitation of protein. CPO was heavily glycosylated and carbohydrate accounted for about 19% of the total molecular weight.1 The crystal structure showed 14 glycosylation sites with a total of 34 sugar molecules in different domains. The interactions of hydroxyl group
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a
A A
0.4 Absorbance
G L G
0.2 L
A B C D E F G H I J K L
0h 1h 3h 15h 18h 23h 25h 40h 45h 47h 61h 86h
b
c
G 0.0 300
400
500 600 700 Wavelength/nm
800
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Figure 2. UV–vis absorption spectra of CPO and G-CPO in 0.05 M citrate buffer (pH 5.5) on incubation time with the amount of D-glucose were 200 (Cglucose/CCPO) at 4 °C. b and c specified the dependence of absorption and wavelength on incubation time, respectively.
of glucose would also obstruct the interaction of hydroxyl group of glucose with carbohydrate on CPO surface and prohibited CPO from precipitation, so the decrease of intensity of Soret band was much smaller in this case. Secondly, phenol was chosen to compare the effect of phenolic hydroxyl with that of aldehyde group on this enzymatic deactivation. It was found that the peak intensity at 398 nm decreased when phenol concentration increased to 0.02 M after 50 h, however, only 32% activity drop was observed, which indicated that phenolic hydroxyl was not as effective as aldehyde group on the deactivation of CPO.
Figure 3. The interaction of aldehyde group of aldose (glucose) with amino groups among CPO surface.
0.3 A B C D E F G H I J K L
Absorbance
0.2
0.1
0h 0.5h 1h 2h 3h 4h 5h 7h 16h 17h 18h 19h
3.1.2. Fluorescence spectra of G-CPO Most iron porphyrin ring in resting protein was considered long-term non-fluorescent, other than myoglobin.16 But Trp, Tyr and Phe residues have intrinsic fluorescence. The fluorescence emission of CPO samples showed a maximum around 333 nm caused mainly by the five Trp residues in CPO.17 The Spectrofluorometric titration of CPO with glucose (Fig. 5) showed a decrease in fluorescence emission with the increase of glucose concentration, and a very minor concomitant red shift was observed which indicated the polarity of the environment around fluorophore increased. Most fluorophores in CPO situated deeply inside the protein molecule, and are surrounded by a hydrophobic environment. This little red shift of emission peak indicated a minor uncoiling of a-helix occurred in the presence of glucose. This conclusion was consistent with that concluded from CD
0.0 300
400
500 600 700 Wavelength/nm
800
900
Figure 4. UV–vis absorption spectra of CPO in 0.05 M citrate buffer (pH 5.5) on incubation time with the amount of D-fructose were 200 (Cfructose/CCPO) at 4 °C.
of D-glucose with the carbohydrate among CPO surface by hydrogen bond might account for the occurrence of precipitation of protein and decrease of absorption at 398 nm. Two experiments were designed to further study the role of aldehyde group in this deactivation. Firstly, the acetylation of glucose was carried out for masking the hydroxyl groups so as to provide steric hindrance for the access of aldehyde group to hold down the deactivation. Much smaller changes of intensity of the Soret band were observed in the presence of glucose acetate and partial activity of CPO remained. This result confirmed that aldehyde group really played a key role. On the other hand, acetylation
Figure 5. Fluorescence of 2.0 lM CPO and G-CPO in 0.05 M citrate buffer (pH 5.5). The Cglucose/CCPO was 50:1, 100:1, 200:1, 500:1, respectively.
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4. Conclusion
Figure 6. Circular dichroism of CPO and G-CPO in 0.05 M citrate buffer (pH 5.5) at 20 °C or 4 °C, respectively. The Cglucose/CCPO was 50:1 and 200:1.
spectral analysis. The main cause for the decrease in fluorescence intensity should be reasonably attributed to both the precipitation of the protein and the interaction of –CHO with amino groups on the surface of CPO. 3.1.3. Circular dichroism (CD) of G-CPO CD spectra were employed to investigate the changes of secondary structure of CPO in this inactivation (Fig. 6). The far-UV CD spectrum of CPO showed a large negative band around 208 nm, which showed CPO was a typical a-helix protein. This was in agreement with the reported data.18 The negative absorption became less strong when glucose was added in, indicating the secondary structure content of G-CPO decreased a little. This was consistent with the result from fluorescence studies. Both the changes of tertiary structure (250–350 nm) and heme active site were very trivial as judged by CD spectroscopy. In addition, Figure 6 showed this deactivation is sensitive to temperature. 3.2. The effect of sugar conformation on the deactivation of CPO It was showed that D-glucose is more effective to inactivate CPO than other sugars tested. This was possibly related to the conformation of these monosaccharides with its deactivation effect on CPO. The dominant conformation of D-glucose was 1e2e3e4e (e stands for an equatorial hydroxyl group), whereas that of D-galactose was 1e2e3e4a (a stands for axial hydroxyl group). This result suggested that 1e2e3e4e conformation is more suitable for the interaction of both the aldehyde and the hydroxyl groups from the sugars with amino groups and the hydroxyl groups of carbohydrates on CPO surface, respectively. Meanwhile, D-glucose is a pyranose whereas D-xylose is a furanose. It seemed that the deactivation was more favorable to the monosaccharide with CHOH (exo).
Some monosaccharides (D-glucose, D-galactose, and D-xylose), which are well known additives used to stabilize enzymes at high temperatures, were found, however, actually causing a deactivation of chloroperoxidase (CPO) in the absence of H2O2. Data from measurement of chlorination activity as well as UV–vis, fluorescent, and CD spectral analysis suggested that this monosaccharide-induced deactivation was probably due to two aspects: one was the interaction of aldehyde group of sugar with amino groups, especially the terminal amino group, on proteins to form Schiff bases which then rearrange to the stable amino ketone. Another was the occurrence of precipitation of protein in the presence of monosaccharide. This deactivation was irreversible and D-glucose was the most efficient due to its aptitude to interact with amino groups and to bind to the carbohydrate on CPO surface that resulted in the precipitation of protein. No deprivation of the heme iron was involved in this deactivation. Acknowledgments This work was funded by the National Natural Science Foundation of China (21176150) and the Fundamental Research Funds for the Central Universities (GK201001006). References 1. Sundaramoorthy, M.; Terner, J.; Poulos, T. L. Structure 1995, 3, 1367–1377. 2. Sundaramoorthy, M.; Terner, J.; Poulos, T. L. Chem. Biol. 1998, 5, 461–473. 3. Rodriguez-Lopez, J. N.; Hernández-Ruiz, J.; Garcia-Cánovas, F.; Thorneley, R. N. F.; Acosta, M.; Arnao, M. B. J. Biol. Chem. 1997, 272, 5469–5476. 4. Huang, Q.; Pinto, R. A.; Griebenow, K.; Schweitzer-Stenner, R.; Weber, W. J. J. Am. Chem. Soc. 2005, 127, 1431–1437. 5. Baynton, K. J.; Bewtra, J. K.; Biswas, N.; Taylor, K. E. Biochim. Biophys. Acta 1994, 1206, 272–278. 6. Nazari, K.; Mahmoudi, A.; Shahrooz, M.; Khodafarin, R.; Moosavi-Movahedi, A. A. J. Enzyme Inhib. Med. Chem. 2005, 20, 285–292. 7. Ortiz de Montellano, P. R.; David, S. K.; Ator, M. A.; Tew, D. Biochemistry 1988, 27, 5470–5476. 8. Park, J. B.; Clark, D. S. Biotechnol. Bioeng. 2006, 93, 1190–1195. 9. Ayala, M.; Batista, C. V.; Vazquez-Duhalt, R. J. Biol. Inorg. Chem. 2011, 16, 63–68. 10. Valderrama, B.; Ayala, M.; Vazquez-Duhalt, R. Chem. Biol. 2002, 9, 555–565. 11. Dexter, A. F.; Hager, L. P. J. Am. Chem. Soc. 1995, 117, 817–818. 12. Bai, C.; Bo, H.; Jiang, Y.; Hu, M.; Li, S.; Zhai, Q. Process Biochem. 2010, 45, 312– 316. 13. Morris, D. R.; Hager, L. P. J. Biol. Chem. 1966, 241, 1763–1768. 14. Egawa, T.; Proshlyskov, D. A.; Miki, H.; Makino, R.; Ogura, T.; Kitagawa, T.; Ishimura, Y. J. Biol. Inorg. Chem. 2001, 6, 46–54. 15. Wagenknecht, H. A.; Woggon, W. D. Chem. Biol. 1997, 4, 367–372. 16. Feng, Y.; Yang, H.; Gu, X.; Jiang, H.; Lu, T. Spectrosc. Spect. Anal. 2003, 23, 532– 534. 17. Nuell, M. J.; Fang, G. H.; Axley, M. J.; Kenigsberg, P.; Hager, L. P. J. Bacteriol. 1988, 170, 1007–1011. 18. Hassani, L.; Ranjbar, B.; Khajeh, K.; Naderi-Manesh, H.; Naderi-Manesh, M.; Sadeghi, M. Enzyme Microb. Technol. 2006, 38, 118–125.
Contents lists available at SciVerse ScienceDirect
Carbohydrate Research journal homepage: www.elsevier.com/locate/carres
Deactivation of chloroperoxidase by monosaccharides (D-glucose, D-galactose, and D-xylose) Rixiao Jin a, Chaonan Li a, Lifei Zhi a, Yucheng Jiang a,b,⇑, Mancheng Hu a,b, Shuni Li a,b, Quanguo Zhai a,b a b
School of Chemistry and Materials Science, Shaanxi Normal University, Chang’an South Road, 199, Xi’an 710062, China Key Laboratory of Macromolecular Science of Shaanxi Province, Shaanxi Normal University, China
a r t i c l e
i n f o
Article history: Received 17 March 2012 Received in revised form 1 July 2012 Accepted 3 July 2012 Available online 14 July 2012 Keywords: Deactivation Chloroperoxidase Monosaccharide
a b s t r a c t In this work, it was found that some monosaccharides normally used to stabilize enzymes at high temperatures, however, actually caused a deactivation of chloroperoxidase (CPO). The red native CPO was converted to a stable pale species that lost enzymatic activity. This deactivation was irreversible and was sensitive to temperature. It was different from the general peroxide-mediated deactivation of CPO. Data from measurement of chlorination activity as well as UV–vis, fluorescent, and CD spectral analysis indicated that monosaccharide-induced deactivation can be attributed to precipitation of protein in the presence of monosaccharide and the interaction of the aldehyde group of sugar with amino groups, especially the terminal amino group, on proteins to form Schiff bases which then rearrange to the stable amino ketone. It is further noted that the deactivation efficiency depends on the stereostructure of monosaccharides. D-Glucose was the most efficient inactivating agents due to its aptitude for both of the interactions mentioned in the above paragraphs. The deactivation was specific to aldose. No deprivation of the heme iron was involved in this deactivation. Ó 2012 Elsevier Ltd. All rights reserved.
1. Introduction Chloroperoxidase (CPO) from Caldariomyces fumago is probably the most versatile known heme enzymes. It catalyzes chlorination of activated C–H bonds and reactions reminiscent of peroxidase, catalase, and cytochrome P450. CPO shares cysteine thiolate heme ligation with cytochrome P-450 monooxygenase family, while possessing a polar active site milieu similar to those of the heme peroxidases and catalase.1,2 The inactivation of CPO reported in literature mainly included the following ways: The first was a general inactivation of typical peroxidases (e.g., horseradish peroxidase) caused by excess oxidants (i.e., hydrogen peroxide) or by double substrates involving H2O2.3–7 Clark et al. described a catalytic cycle of such systems, in which hydrogen peroxide bound to the penta-coordinate ferric resting state Fe(III) to generate Compound I, a oxoferryl porphyrin p cation radical containing Fe(IV). Compound I reacts with hydrogen peroxide either in a catalase-like two-electron reduction, which releases molecular oxygen, or in two single-electron transfers in which Compound II and Compound III were involved as intermediates. During two single-electron reductions of Compound ⇑ Corresponding author. Tel.: +86 029 81530763. E-mail addresses: [email protected] (R. Jin), [email protected] (C. Li), [email protected] (L. Zhi), [email protected] (Y. Jiang), [email protected] (M. Hu), [email protected] (S. Li), [email protected] (Q. Zhai). 0008-6215/$ - see front matter Ó 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.carres.2012.07.001
I, free radicals and superoxide anion radical such as HO2, HO, were formed. These radicals oxidize the heme and/or protein residues, leading to the inactivation of the enzyme.8 Ayala et al. pointed out that under peroxidasic conditions the most important inactivating event was the destruction of the heme group. Regarding protein oxidation, no major structural disruption such as protein cross-linking or unfolding was detected. Tryptophan (most sensitive to oxidation) oxidation was a simultaneous event, rather than the predominant cause of deactivation.9 Additionally, a catalaselike function was described,10 in which CPO was significantly resistant to inactivation by hydrogen peroxide or by organic hydroperoxides because of its intrinsic catalase activity. Nevertheless, exposure to high concentrations of hydrogen peroxide (>30 mM) could irreversibly inactivate CPO with a half-life of about 1 min. Furthermore, Hager et al. reported a suicidal formation of N-alkylporphyrins during the oxidation of terminal alkenes and alkynes. The inactivated CPO underwent spontaneous loss of the heme alkyl moiety with partial restoration of enzymatic activity.11 Our group reported an arginine-induced inactivation that might be caused mainly by the binding of guanidinium group in arginine to the acid–base catalytic group Glu183 in CPO.12 Recently, we surprisingly discovered that CPO could be inactivated by some monosaccharides (D-glucose, D-galactose, and D-xylose) in the absence of H2O2. It is well known that polyhydroxy compounds, such as monosaccharides and glycerin etc., can improve the thermal stability of enzyme because they can prevent
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enzyme from conformation change at high temperatures. Therefore, this deactivation is very interesting and could provide new insight into the nature of interaction between sugars and CPO. In this paper, UV–vis, fluorescent, and circular dichroism spectral analysis combined with the measurement of chlorination activity was employed to probe D-glucose-mediated deactivation of CPO and the mechanism involved in the deactivation process. 2. Materials and methods 2.1. Materials Chloroperoxidase was isolated from the growth medium of Caldariomyces fumago according to the method established by Morris and Hager13 with minor modifications using acetone rather than ethanol in the solvent fractionation step. The enzyme had an activity of 7800 U/mL based on the standard monochlorodimedon (MCD) chlorination assay. The Rz of CPO used in all experiments was 1.35 (Rz = purity standard = A398/A280 = 1.44 for pure enzyme). D-Glucose, D-galactose, and D-xylose were purchased from Shanghai Chemical and Medicine Co. Ltd. MCD was obtained from Sigma. Citric acid, Sodium citrate, and hydrogen peroxide (30% in aqueous solution) were obtained from Xi’an Chemical Co. Ltd. All chemicals are of analytical grade unless otherwise indicated. Enzyme assay and other optical measurements were performed on a Shimadzu UV-1700 spectrophotometer, 970 CRT fluorescence spectrometry (Shanghai analyze instrument factory) and circular dichroism (CD) spectrometer (Applied Photophysics Ltd), respectively. 2.2. Methods 2.2.1. UV–vis spectroscopy 0.2 mM CPO in buffer at pH 5.5 was incubated with different concentrations of D-glucose as long as 86 h at 4 °C, which is the storage temperature of enzyme. An appropriate dilution was needed before recording its UV–vis spectra (7.2 lM). The spectrum of native CPO with same concentration was recorded for comparison. The concentration of CPO was determined by the absorbance at 398 nm (Soret band) using an extinction coefficient of 91,200 M 1 cm 1. 2.2.2. Fluorescence measurements 0.2 mM CPO in buffer at pH 5.5 was incubated with different concentrations of D-glucose for 86 h at 4 °C. The sample was diluted appropriately (2.0 lM), and then excited at 287 nm. Fluorescence spectra were recorded from 300 to 400 nm. The spectrum of native CPO with same concentration was also recorded for comparison. 2.2.3. Circular dichroism (CD) measurements 7.2 lM CPO sample was prepared as above for CD spectra recording from 190 to 700 nm in a 200 lL quartz cuvette at 4 °C. The operation parameters were as following: path-length: 1.0 mm, time-per-point: 0.25, number of scans: 3, spectral width: 1.0 nm. 3. Results and discussion 3.1. Deactivation of CPO by D-glucose As shown in Figure 1, a red native CPO (0.2 mM) was converted to a pale green species (G-CPO) after 50 h incubation with D-glucose at 4 °C accompanied with spontaneous loss of partial
Figure 1. Red native CPO (right) was converted to a pale green species (left) after 50 h incubation with D-glucose at 4 °C, pH 5.5.
enzymatic activity as judged by the MCD assay. This deactivation was sensitive to temperature, but not sensitive to pH in the range of pH 4.5–5.5 which was the suitable pH for storage of enzyme. G-CPO was stable, and no color change was found after 10 days of standing, indicating the deactivation was an irreversible process. No deprivation of the heme iron was involved in this inactivation as confirmed by the graphite furnace atomic absorption spectrometry measurement. It was also examined that if other monosaccharide (e.g., D-galactose and D-xylose) could cause this deactivation. Similar phenomenon was observed. 3.1.1. UV spectra of G-CPO The iron(II1) porphyrin in CPO is a high-spin complex that displays a Soret band at kmax = 398 nm.14,15 In Figure 2a, the absorption of kmax had no obvious change in the presence of trace amount of D-glucose, but began to decrease obviously when Cglucose/CCPO increased to 200. This decrease in absorption continued with incubation time, meanwhile the absorption shifted from 398 nm to 409 nm gradually. However, the absorption was found to increase again with the peak position shifted back to 398 nm gradually on standing. But the final spectrum was not identical to that of native chloroperoxidase (Fig. 2b and c). The pH value did not change during this process. Moreover, the higher the Cglucose/CCPO was, the faster the decrease in absorbance. Based on the above spectral properties of G-CPO, it was postulated that there were two possible interactions involved in this process. Firstly, it was noted that the monosaccharides tested here (D-glucose, D-galactose, and D-xylose) happened to be all aldoses. So one possible interaction was the interaction of these monosaccharides with amino groups, especially the terminal amino group, on CPO to form Schiff bases which then rearrange to the stable amino ketone, as illustrated in Figure 3. This interaction might account for the observed loss of CPO’s activity due to the destruction of its conformation during this process. To further testify this speculation, fructose was incubated with CPO under the same conditions as used for glucose. As expected, almost no decrease in Soret band and enzymatic activity was observed (Fig. 4), demonstrating that the key functional groups that caused the deactivation of CPO were the –CHO group in sugars. Another reason causing absorption decrease was the precipitation of protein. CPO was heavily glycosylated and carbohydrate accounted for about 19% of the total molecular weight.1 The crystal structure showed 14 glycosylation sites with a total of 34 sugar molecules in different domains. The interactions of hydroxyl group
74
R. Jin et al. / Carbohydrate Research 370 (2013) 72–75
a
A A
0.4 Absorbance
G L G
0.2 L
A B C D E F G H I J K L
0h 1h 3h 15h 18h 23h 25h 40h 45h 47h 61h 86h
b
c
G 0.0 300
400
500 600 700 Wavelength/nm
800
900
Figure 2. UV–vis absorption spectra of CPO and G-CPO in 0.05 M citrate buffer (pH 5.5) on incubation time with the amount of D-glucose were 200 (Cglucose/CCPO) at 4 °C. b and c specified the dependence of absorption and wavelength on incubation time, respectively.
of glucose would also obstruct the interaction of hydroxyl group of glucose with carbohydrate on CPO surface and prohibited CPO from precipitation, so the decrease of intensity of Soret band was much smaller in this case. Secondly, phenol was chosen to compare the effect of phenolic hydroxyl with that of aldehyde group on this enzymatic deactivation. It was found that the peak intensity at 398 nm decreased when phenol concentration increased to 0.02 M after 50 h, however, only 32% activity drop was observed, which indicated that phenolic hydroxyl was not as effective as aldehyde group on the deactivation of CPO.
Figure 3. The interaction of aldehyde group of aldose (glucose) with amino groups among CPO surface.
0.3 A B C D E F G H I J K L
Absorbance
0.2
0.1
0h 0.5h 1h 2h 3h 4h 5h 7h 16h 17h 18h 19h
3.1.2. Fluorescence spectra of G-CPO Most iron porphyrin ring in resting protein was considered long-term non-fluorescent, other than myoglobin.16 But Trp, Tyr and Phe residues have intrinsic fluorescence. The fluorescence emission of CPO samples showed a maximum around 333 nm caused mainly by the five Trp residues in CPO.17 The Spectrofluorometric titration of CPO with glucose (Fig. 5) showed a decrease in fluorescence emission with the increase of glucose concentration, and a very minor concomitant red shift was observed which indicated the polarity of the environment around fluorophore increased. Most fluorophores in CPO situated deeply inside the protein molecule, and are surrounded by a hydrophobic environment. This little red shift of emission peak indicated a minor uncoiling of a-helix occurred in the presence of glucose. This conclusion was consistent with that concluded from CD
0.0 300
400
500 600 700 Wavelength/nm
800
900
Figure 4. UV–vis absorption spectra of CPO in 0.05 M citrate buffer (pH 5.5) on incubation time with the amount of D-fructose were 200 (Cfructose/CCPO) at 4 °C.
of D-glucose with the carbohydrate among CPO surface by hydrogen bond might account for the occurrence of precipitation of protein and decrease of absorption at 398 nm. Two experiments were designed to further study the role of aldehyde group in this deactivation. Firstly, the acetylation of glucose was carried out for masking the hydroxyl groups so as to provide steric hindrance for the access of aldehyde group to hold down the deactivation. Much smaller changes of intensity of the Soret band were observed in the presence of glucose acetate and partial activity of CPO remained. This result confirmed that aldehyde group really played a key role. On the other hand, acetylation
Figure 5. Fluorescence of 2.0 lM CPO and G-CPO in 0.05 M citrate buffer (pH 5.5). The Cglucose/CCPO was 50:1, 100:1, 200:1, 500:1, respectively.
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4. Conclusion
Figure 6. Circular dichroism of CPO and G-CPO in 0.05 M citrate buffer (pH 5.5) at 20 °C or 4 °C, respectively. The Cglucose/CCPO was 50:1 and 200:1.
spectral analysis. The main cause for the decrease in fluorescence intensity should be reasonably attributed to both the precipitation of the protein and the interaction of –CHO with amino groups on the surface of CPO. 3.1.3. Circular dichroism (CD) of G-CPO CD spectra were employed to investigate the changes of secondary structure of CPO in this inactivation (Fig. 6). The far-UV CD spectrum of CPO showed a large negative band around 208 nm, which showed CPO was a typical a-helix protein. This was in agreement with the reported data.18 The negative absorption became less strong when glucose was added in, indicating the secondary structure content of G-CPO decreased a little. This was consistent with the result from fluorescence studies. Both the changes of tertiary structure (250–350 nm) and heme active site were very trivial as judged by CD spectroscopy. In addition, Figure 6 showed this deactivation is sensitive to temperature. 3.2. The effect of sugar conformation on the deactivation of CPO It was showed that D-glucose is more effective to inactivate CPO than other sugars tested. This was possibly related to the conformation of these monosaccharides with its deactivation effect on CPO. The dominant conformation of D-glucose was 1e2e3e4e (e stands for an equatorial hydroxyl group), whereas that of D-galactose was 1e2e3e4a (a stands for axial hydroxyl group). This result suggested that 1e2e3e4e conformation is more suitable for the interaction of both the aldehyde and the hydroxyl groups from the sugars with amino groups and the hydroxyl groups of carbohydrates on CPO surface, respectively. Meanwhile, D-glucose is a pyranose whereas D-xylose is a furanose. It seemed that the deactivation was more favorable to the monosaccharide with CHOH (exo).
Some monosaccharides (D-glucose, D-galactose, and D-xylose), which are well known additives used to stabilize enzymes at high temperatures, were found, however, actually causing a deactivation of chloroperoxidase (CPO) in the absence of H2O2. Data from measurement of chlorination activity as well as UV–vis, fluorescent, and CD spectral analysis suggested that this monosaccharide-induced deactivation was probably due to two aspects: one was the interaction of aldehyde group of sugar with amino groups, especially the terminal amino group, on proteins to form Schiff bases which then rearrange to the stable amino ketone. Another was the occurrence of precipitation of protein in the presence of monosaccharide. This deactivation was irreversible and D-glucose was the most efficient due to its aptitude to interact with amino groups and to bind to the carbohydrate on CPO surface that resulted in the precipitation of protein. No deprivation of the heme iron was involved in this deactivation. Acknowledgments This work was funded by the National Natural Science Foundation of China (21176150) and the Fundamental Research Funds for the Central Universities (GK201001006). References 1. Sundaramoorthy, M.; Terner, J.; Poulos, T. L. Structure 1995, 3, 1367–1377. 2. Sundaramoorthy, M.; Terner, J.; Poulos, T. L. Chem. Biol. 1998, 5, 461–473. 3. Rodriguez-Lopez, J. N.; Hernández-Ruiz, J.; Garcia-Cánovas, F.; Thorneley, R. N. F.; Acosta, M.; Arnao, M. B. J. Biol. Chem. 1997, 272, 5469–5476. 4. Huang, Q.; Pinto, R. A.; Griebenow, K.; Schweitzer-Stenner, R.; Weber, W. J. J. Am. Chem. Soc. 2005, 127, 1431–1437. 5. Baynton, K. J.; Bewtra, J. K.; Biswas, N.; Taylor, K. E. Biochim. Biophys. Acta 1994, 1206, 272–278. 6. Nazari, K.; Mahmoudi, A.; Shahrooz, M.; Khodafarin, R.; Moosavi-Movahedi, A. A. J. Enzyme Inhib. Med. Chem. 2005, 20, 285–292. 7. Ortiz de Montellano, P. R.; David, S. K.; Ator, M. A.; Tew, D. Biochemistry 1988, 27, 5470–5476. 8. Park, J. B.; Clark, D. S. Biotechnol. Bioeng. 2006, 93, 1190–1195. 9. Ayala, M.; Batista, C. V.; Vazquez-Duhalt, R. J. Biol. Inorg. Chem. 2011, 16, 63–68. 10. Valderrama, B.; Ayala, M.; Vazquez-Duhalt, R. Chem. Biol. 2002, 9, 555–565. 11. Dexter, A. F.; Hager, L. P. J. Am. Chem. Soc. 1995, 117, 817–818. 12. Bai, C.; Bo, H.; Jiang, Y.; Hu, M.; Li, S.; Zhai, Q. Process Biochem. 2010, 45, 312– 316. 13. Morris, D. R.; Hager, L. P. J. Biol. Chem. 1966, 241, 1763–1768. 14. Egawa, T.; Proshlyskov, D. A.; Miki, H.; Makino, R.; Ogura, T.; Kitagawa, T.; Ishimura, Y. J. Biol. Inorg. Chem. 2001, 6, 46–54. 15. Wagenknecht, H. A.; Woggon, W. D. Chem. Biol. 1997, 4, 367–372. 16. Feng, Y.; Yang, H.; Gu, X.; Jiang, H.; Lu, T. Spectrosc. Spect. Anal. 2003, 23, 532– 534. 17. Nuell, M. J.; Fang, G. H.; Axley, M. J.; Kenigsberg, P.; Hager, L. P. J. Bacteriol. 1988, 170, 1007–1011. 18. Hassani, L.; Ranjbar, B.; Khajeh, K.; Naderi-Manesh, H.; Naderi-Manesh, M.; Sadeghi, M. Enzyme Microb. Technol. 2006, 38, 118–125.