Anodic peroxidase isoenzymes and polyphenol oxidase activity from cucumber fruit: Tissue and substrate specificity

Phytochemistry,

Vol. 29, No. 3, pp. 705 709, 1990.

003 l-9422/90

Printed in Great Britain.

%3.00+ 0.00

Q 1990Pergamon Press plc

ANODIC PEROXIDASE ISOENZYMES AND POLYPHENOL OXIDASE ACTIVITY FROM CUCUMBER FRUIT: TISSUE AND SUBSTRATE SPECIFICITY A. RAYMOND MILLER, THOMAS J. KELLEY and CESAR V. MUJER Department of Horticulture, The Ohio State University, Ohio Agricultural Research and Development Center, Wooster, OH 44691, U.S.A. (Received

in revised form 7 August

1989)

Key Word Index-Cucumis sativus; Cucurbitaceae; fruit tissue; gel electrophoresis; polyphenol oxidase; aromatic amines; phenols; thermostability.

peroxidase isoenzymes;

Abstract-Peroxidase activity in fresh cucumber fruit was highest in the skin; followed by pericarp then carpel tissues, respectively. Polyphenol oxidase activity, by contrast, was present only in the skin. In crude extracts, both activities had pH optima near 7.0, but exhibited different temperature optima and thermostability. Using hydrogen peroxide with either benzidine, guaiacol, p-phenylenediamine, o-phenylenediamine, o-dianisidine, or 3-amino-9-ethylcarbazole as substrates, more anodic peroxidase isoenzymes were observed in the skin than the pericarp or carpel. The reactivity of the peroxidase isoenzymes toward these stains varied qualitatively and quantitatively, e.g. benzidine stained more and different isoenzymes with less relative intensity than did p-phenylenediamine. Further, no staining occurred with the above substrates in the absence of hydrogen peroxide, or with Fast Blue BB base, caffeic acid, syringaldazine or polyphenol oxidase substrates (catechol, hydroquinone, L-DOPA), regardless of hydrogen peroxide availability. These data show that cucumber fruit anodic peroxidase isoenzymes vary widely in substrate specificity. Further, the high thermostability of cucumber peroxidase and polyphenol oxidase activity suggests that these enzymes may play a role in the darkening of processed cucumber products.

INTRODUCTION

RESULTS AND DISCUSSION

Peroxidases (donor; hydrogen peroxide oxidoreductase, EC 1.11.1.7) and polyphenol oxidases (donor: oxygen oxidoreductase, EC 1.14.18.1) are ubiquitous in higher plants. Peroxidases are thought to be involved in lignification, hormone metabolism, and response to stress [ 11, whereas polyphenol oxidases mediate oxidative browning of wounded or senescing tissues [24]. In maize roots, peroxidase and polyphenol oxidase activities co-electrophorese and share certain metal inhibitor and substrate specificities, which suggests they may be the same protein [S]. Other investigators, however, have shown that while peroxidase and polyphenol oxidase have similar M,s and ammonium sulphate solubilities, they can be separated by DEAE-cellulose chromatography [3, 63, or on the basis of intracellular locations [4, 71. In cucumbers, peroxidase activity is quite high [S] and exists in several isoenzymic forms [9-111. Cucumber peroxidase is also detectable in processed cucumber products [12, 131. This thermostable peroxidase activity may account for adverse flavour, colour and odour development during storage of processed cucumbers [14-161, but polyphenol oxidase activity could also cause these changes. Polyphenol oxidase activity in cucumber fruit has not been reported. The experiments reported in this paper were conducted to determine whether cucumbers possess polyphenol oxidase activity and initially characterize cucumber peroxidase and polyphenol oxidase activity using tissue, pH, temperature, isoenzyme and substrate specificity studies.

Peroxidase activity in freshly harvested cucumber fruit was highest in the skin (166 units/mg protein; 1 unit = 1 A 470 nm/min) using guaiacol and hydrogen peroxide as substrates. Pericarp and carpel tissues had significantly less activity (80 and 52 units/mg protein, respectively). By contrast, polyphenol oxidase activity was quantifiable, using catechol as the substrate, only in the skin tissue (0.183 units/mg protein; 1 unit = 1 A 420 nm/min). Polyphenol oxidase activity was extremely low in the pericarp and carpel and could not be detected consistently. This activity was not enhanced by 0.1% sodium dodecylsulphate, regardless of tissue (data not shown) which indicated the absence of latent polyphenol oxidases observed in some other genera [ 171. A similar tissue distribution was obtained when activities were expressed on a fresh weight basis. No peroxidase activity was detected in the absence of hydrogen peroxide, and hydrogen peroxide inhibited the polyphenol oxidase reaction. To our knowledge, this is the first report of polyphenol oxidase activity in cucumbers. Because peroxidase and polyphenol oxidase activity could have similar effects on processed cucumber quality, the activities were further characterized according to pH and temperature optima, thermostability, isoenzyme profile, and substrate specificity. Peroxidase and polyphenol oxidase activities in crude extracts exhibited similar pH-activity profiles with optima near pH 7. Peroxidase also exhibited a minor shoulder at pH 4, while polyphenol oxidase had almost no activity at this pH. At pH 10, polyphenol oxidase 705

706

A. R. MII:LER

activity was still 60% of that observed at pH 7, whereas peroxidase activity had decreased to only 10% of that at pH 7. Peroxidase and polyphenol oxidase did exhibit distinctly different temperature optima and thermostability responses. Peroxidase activity was optimum near 60” with a shoulder at 40’ (Fig. 1). Polyphenol oxidase activity,

by contrast,

had

a single

temperature

optimum

et a/.

at SO”. In addition, the Q10 between 20 and 30” for peroxidase was 1.32, while for polyphenol oxidase it was 1.57. Further, thermostability experiments showed that peroxidase activity exhibited a significant (25%) thermal enhancement effect at 50” relative to the 25” control, and nearly complete inactivation at 90” (Fig. 2). Polyphenol oxidase activity showed no such thermal enhancement, but like peroxidase. required 9&100” for inactivation.

6

/

w Peroxidase 0 Polyphenoloxidase ,

I 0

I 20

I 40

I 60

Temperature Fig. I. The effect of reaction 0.1 M Na-phosphate

8C

(“1

temperature on extractable cucumber peroxidase and polyphenol buffer (pH 6.5). Each point represents the mean of duplicate

n

40

60

oxidase activity assays.

in

Peroxidase

80

100

120

Treatment temperature to 1 Fig. 2. Thermostability of extractable cucumber peroxidase and polyphenol oxidase activity in 0.1 M Naphosphate buffer (pH 6.5). Aliquots of enzyme extract were incubated for 10 min at the specified temperature, cooled on ice for 5 min, then assayed at 25”. Data are presented as per cent of25” treatment (control) and each point represents the mean of duplicate assays.

Cucumber

peroxidase

The high thermostability of these enzymes suggests that polyphenol oxidase and/or peroxidase may be involved in adverse flavour, colour and odour development in processed cucumbers [14-161, and may partially explain the presence of peroxidase in pasteurized cucumber products [12, 133. Cucumber peroxidase isoenzymes have not been characterized according to tissue or substrate specificity and preliminary experiments by us indicated that differences existed. Therefore, following anionic gel electrophoresis of extracts from skin, pericarp and carpel tissues, gels were incubated with different peroxidase substrates. We found, in the presence of hydrogen peroxide, that only guaiacol and the aromatic amines benzidine, o-dianisidine, p-phenylenediamine, o-phenylenediamine and 3amino-9-ethylcarbazole formed coloured bands (Fig. 3). Other reported peroxidase substrates (caffeic acid, Fast Blue BB base and syringaldazine) did not serve as stains for cucumber anodic isoenzymes, and no bands were observed with any potential substrate in the absence of hydrogen peroxide. Some of the substrates tested (e.g. pphenylenediamine) could be oxidized by lactase [7]. However, our data indicate that lactase is not present in cucumber fruit due to a lack of activity toward these substrates in the absence of hydrogen peroxide. Further, other heme-containing enzymes (e.g. catalase and cytochromes) could oxidize these substrates in the presence of hydrogen peroxide, thus leading to artifacts. The separation though, of these peroxidase-like activities from true peroxidase isoenzymes would require immunological and/or molecular biology approaches beyond the scope

and polyphenol

oxidase

of the present paper. Apparently, all the isoenzymes detected with the various substrates in the presence of hydrogen peroxide were heme-containing proteins, since no banding was observed when 1 mM sodium cyanide, sodium azide, or sodium ascorbate were added to the gel incubation mixture. Skin tissue had more isoenzymes than either pericarp or carpel tissue (Fig. 3), regardless of staining substrate and ‘whether the samples were applied to the gels on an equal protein or activity basis (data not shown). The number of anodic peroxidase isoenzymes in extracts from a specific tissue corresponded with the relative amounts of total peroxidase activity present. The isoenzymes could also be roughly divided, based on their relative mobilities (M,) within the gel, into three groups; slow-, moderateand fast-migrating (M,=O-O.2, 0.3-0.6 and 0.7-1.0, respectively). Hence, skin extracts contained all three isoenzyme groups and had the highest activity with the greatest number of anodic isoenzymes, compared to pericarp and carpel extracts which generally had fastand/or slow-migrating isoenzymes only and less total activity. The number and specific isoenzymes detected at pH 6.5 by a particular substrate within a tissue varied greatly. For example, benzidine stained 10 isoenzymes in skin extracts (2 slow-, 6 moderate- and 2 fast-migrating forms), while guaiacol, o-phenylenediamine, p-phenylenediamine, o-dianisidine and 3-amino-9-ethylcarbazole stained 7, 6, 5, 4 and 4 isoenzymes, respectively (Fig. 3). The number of isoenzymes visualized with the various substrates was probably not due to proteolysis because the

Starn B~llZidirl~

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3

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101

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Fig. 3. Diagramatic representation of soluble plus ionically-bound anodic peroxidase isoenzymes in extracts from cucumber skin (S), pericarp (P), and carpel (C) tissues. Following discontinuous native polyacrylamide gel electrophoresis, isoenzymes were stained with the specified substrates in the presence of hydrogen peroxide, the mobilities of the isoenzymes recorded,.and the relative intensities of the bands were rated as indicated. Each lane contained 45 pg of total protein. The direction of electrophoretic migration was from top (-) to bottom (+).

A. R. MILLER et al.

708

extracting buffer contained the proteinase inhibitor, phenylmethylsulphonylfluoride (PMSF). However, nonspecific staining due to peroxidase-like, heme-containing enzymes (see above) may have contributed to the bands visualized. Further, the two fast-migrating isoenzymes visualized with benzidine were different from the fastmigrating isoenzymes stained by the other substrates. Several other examples of discriminating reactivity were observed. For example, guaiacol was the only substrate to stain isoenzyme M, = 0.11 and p-phenylenediamine was the only substrate to stain isoenzyme M,=0.93 in skin extracts. Even with this variation, some common staining occurred. All substrates except 3-amino-9-ethylcarbazole stained isoenzyme M, = 0.04, and isoenzyme M, = 0.47 was stained by o-phenylenediamine, o-dianisidine, and 3amino-9-ethylcarbazole. Hence, it appears that certain isoenzymes will act upon a wide range of substrates (M, = 0.04 and 0.47), while others (M, = 0.11 and 0.93) require a specific substrate. The reason(s) for this range of specificity is unknown, but may be related to active site stereochemistry, physical characteristics (solubility, pK, etc.) of each substrate or suboptimal levels of the substrates and hydrogen peroxide, although the latter were thought to be in excess. Our results concur with those of Miller and George [ll], who reported 12.-15 anodic isoenzymes in cucumber seedling extracts, using benzidine as the substrate. Presently, we observed 10 benzidine-stained isoenzymes plus 6 other distinct isoenzymes stained with the other substrates. The data, however, are in contrast with those of Staub et a[. [ 181, who were unable to detect peroxidase activity following starch gel electrophoresis of cucumber fruit extracts. These investigators gave no details on buffer, pH, substrate or reaction conditions, but simply may have used a nonreactive substrate. Because catechol was used to determine polyphenol oxidase activity, we attempted to use it as an isoenzyme stain as well. When incubated at pH 6.5 in the absence of hydrogen peroxide at either 25 or 50”, catechol only very weakly colorized entire lanes (not shown). No definite bands were evident. Similar results were obtained using LDOPA as the substrate, and gels incubated with hydroquinone lacked any evidence of staining. In the presence of hydrogen peroxide, no staining was observed with any of the polyphenol oxidase substrates. The possibility remains, however, that electrophoresis caused the dissociation of a cofactor or regulatory protein necessary for polyphenol oxidase activity. In summary, cucumber anodic peroxidase isoenzymes vary widely in their substrate specificity, but react preferentially with aromatic amines. This observation suggests that investigators should choose a peroxidase stain based on its ability to react with the specific isoenzyme(s) of interest. This approach would be of further value if the in situ biochemical functions of the isoenzymes were known or if other techniques were employed to identify specific isoenzymes. Peroxidase activity, however, does not appear to account for the observed polyphenoloxidase activity. Although the activities have similar pH optima, polyphenol oxidase does not exhibit a thermal enhancement effect like peroxidase, and they differ in temperature optima, Qlo, and stainability on anionic gels.

Development Center, Wooster, OH, and immature fruits (45-55 mm diameter, ca 16 days after pollination) were randomly selected and harvested. The fruits were washed with cool tap H,O and immediately separated with a stainless steel knife into skin, pericarp and carpel tissues for subsequent enzyme assay and isoenzyme analysis. Enzyme extraction and assay. Soluble and ionically bound enzymes were extracted by grinding 5 g of tissue at 4” in 10 ml of 1 M NaCI and 1 mM PMSF with a mortar and pestle and 10 g of acid-washed sea sand. The homogenates were incubated at 4” for 1 hr to complete the extraction, then filtered through 2 layers of cheesecloth and cleared by centrifugation at 20000 g for 20 min. The resulting supernatant was assayed directly for peroxidase and polyphenol oxidase activity. Total protein was determined by the Coomassie dye-binding assay 1191. Peroxidase activity was determined spectrophotometrically [6], using 60 mM guaiacol and 0.03% (v/v) H,O, in 0.1 M Na-Pi buffer (pH 6.5). Polyphenol oxidase activity was determined spectrophotometrically [20], using 0.1 M catechol(1,2-benzenedial) in 0.1 M Na-Pi buffer (pH 6.5). For pH-activity profiles, peroxidase and polyphenol oxidase were assayed in 0.1 M (total molarity) citrate-Na-Pi-borate buffer adjusted to the desired pH. This buffer system had no observable effect on the measured activity of either enzyme compared to Na-Pi buffer. Control peroxidase and polyphenol oxidase assays lacking enzyme were run in all experiments to correct for auto-oxidation of guaiacol and catechol, respectively. Isoenzyme separation and staining. Discontinuous anionic polyacrylamide gel electrophoresis was conducted under nondissociating conditions as previously described [7]. Following electrophoresis, gels were incubated in 150 ml of 50 mM Na-Pi buffer (pH 6.5) containing one of the following substrates [4] in the presence or absence of 0.03% (v/v) H,O,: guaiacol, benzidine, o-phenylenediamine, p-phenylenediamine, syringaldazine, Fast Blue BB base, and caffeic acid at 0.05%; o-dianisidine and 3-amino-Y-ethylcarbazole at 0.02% and; catechol, hydroquinone and dihydroxyphenylalanine (L-DOPA) at 0.1%. Fast Blue BB base, caffeic acid, o-phenylenediamine, p-phenylenediamine and 3-amino-9-ethylcarbazole were first dissolved in 0.5 ml ofdimethylsulphoxide. o-Dianisidine was first dissolved in 0.5 ml of 1 M HCI. Following electrophoresis and isoenzyme visualization with each stain, the relative mobilities of the isoenzymes were recorded and stain intensity was rated as specified on the electrophoretogram. Photography of the gels for recording data was of limited use as the products of the reactions diffused readily. Tissues from 3 fruits were combined and extracted for enzyme assays and isoenzyme analysis. Assays and gels were done in duplicate and each experiment was repeated at least 3 times. The data presented are from representative experiments.

Acknowledgements~We thank Dr Robert Clements (USDA/ ARS), for advice on native PAGE, Christel Velbinger and Mark Jameson for their excellent technical assistance, and Bonnie Beck for typing the manuscript. Salaries and research support provided in part by State and Federal funds appropriated to the Ohio Agricultural Research and Development Center, The Ohio State University. Journal Article No. 17-89.

REFERENCES EXPERIMENTAL Hunt material. plants

were

I. Gaspar,

Cucumber (Cucumis sativus L. cv. ‘Heinz 3534’)

grown

at the Ohio

Agricultural

Research

and

T., Prel, C., Thorpe, T. A. and Greppin, H. (1981) Peroxidases, 197Ck1980. University of Geneva Press, Geneva.

Cucumber

peroxidase

2. Wissemann, K. W. and Montgomery, M. W. (1985) PIant Physiol. 78, 256. 3. Tremolieres, M. and Bieth, J. G. (1984) Phytochemistry 23, 501. 4. Vaughn, K. C. and Duke, S. 0. (1984) Physiol. Plant 60, 106. 5. Grison, R. and Pilet, P.-E. (1985) Phytochemistry 24, 2519. 6. Van Loon, L. C. (1971) Phytochemistry 10, 503. 18, 193. 7. Mayer, A. M. and Harel, E. (1979) Phytochemistry 8. Miller, A. R., Dalmasso, J. P. and Kretchman, D. W. (1987) J. Am. Sot. Hort. Sci. 112, 666. 9. Miller, A. R. and Kelley, T. J. (1989) HortScience 24, 650. 10. Dane, F. K. (1983) in Isozymes in Plant Genetics and Breeding, Purr B (Tanksley, S. D. and Orton, T. J., eds), p. 391. Elsevier, Amsterdam. 11. Miller, G. A. and George, W. L., Jr. (1979) HortScience 14, 24.

and polyphenol

oxidase

709

12. Buescher, R. W. and McGuire, C. (1986) J. Food Sci. 51, 1079. 13. Buescher, R. W., McGuire, C. and Skulman, B. (1987) J. Food Sci. 52, 228. 14. Anderson, E. E., Ruder, L. F., Esselen, W. B., Nebesky, E. A. and Labbee, M. (1951) Food Technol. 5, 364. 15. Labbee, M. D. and Esselen, W. B. (1954) Food Technol. 8,50. 16. Nebesky, E. A., Eselen, W. B. and Fellers, C. R. (1951) Food Technol. 5, 110. 17. Angleton, E. L. and Flurkey, W. H. (1984) Phytochemistry 23, 2723. 18. Staub, J. E., Kupper, R. S., Schuman, D., Wehner, T. C. and Mays, B. (1985) J. Am. Sot. Hort. Sci. 110,426. 19. Bradford, M. M. (1976) Anal. Biochem. 72, 248. 20. Wissemann, K. W. and Lee, C. Y. (1980) Am. J. Enol. Vitic. 31, 206.