Proteolytic processing of polyphenol oxidase from plants and fungi

Abstract

Polyphenol oxidase (PPO), a metalloenzyme containing a type-3 copper center, is produced by many species of plants, fungi, and bacteria. There is great variability in the subunit molecular mass reported for PPO, even from a single species. In some cases, experimental evidence (usually protein sequencing by Edman degradation) indicates that the variability in molecular mass for PPO from a given species is the result of proteolytic processing at the N and/or C-termini of the protein. In order to identify specific sequence regions where proteolysis occurs in PPO from most species, the experimentally established N and C-termini of these proteolyzed enzymes were compared to the protein sequences of other PPOs for which the N and C-termini have not been established by protein sequencing methods. In all cases the N-terminal proteolysis sites were located prior to a conserved arginine residue, and the C-terminal proteolysis sites were located following a conserved tyrosine motif. Based on the sites of proteolysis, molecular masses were calculated for the enzymes, and the calculated values were used to rationalize the varying molecular masses reported in the literature. To determine the structural implications of N and C-terminal proteolysis, the proteolysis sites were related to the two available PPO structures: Ipomoea batatas catechol oxidase and Streptomyces castaneoglobisporus tyrosinase. A structural “core” region that appears to be essential for structural stability and enzymatic activity was identified.

Introduction

Polyphenol oxidases (PPOs) are ubiquitous metalloenzymes occurring in species ranging from mammals to bacteria [1], [2]. These enzymes utilize molecular oxygen to catalyze the oxidation of o-diphenols to o-quinones (catecholase/diphenolase activity; EC 1.10.3.1) via a type-3 copper center [3], [4], [5]. The quinone products self-polymerize or react with various nucleophiles to yield dark colored melanin polymers. Some PPOs can also catalyze the hydroxylation of various monophenols to o-diphenols (cresolase/monophenolase activity; EC 1.14.18.1). In enzyme nomenclature the designation tyrosinase (TYR) is most commonly used for an enzyme that exhibits both catecholase and cresolase activities, while catechol oxidase (CO) generally is used to designate an enzyme that exhibits only catecholase activity. However, the designation polyphenol oxidase, or simply phenoloxidase, is very commonly used in the literature for both TYRs and COs from plants and fungi. In many cases this may be because it is unclear, or has not been determined, whether a given enzyme has cresolase activity [6], [7], [8]. The designations we use herein (PPO, TYR, or CO) for enzymes from specific plants, fungi, and bacteria are those most commonly used in the literature for a given species.

The functions of plant and fungal PPOs are not well understood, but there is evidence that the melanins produced by fungal PPOs provide a means of defense and resistance to stressors, contribute to the formation and stability of spores, and play a role in virulence [1], [2]. For PPOs of higher plants, a role in defense against insects and microorganisms has been suggested, as PPO may be upregulated in wounded tissues and melanins form a protective scab after attack upon the plant [1], [6], [9]. In addition, oxygen-scavenging in the chloroplast has been proposed [10], and PPOs have been implicated in the biosynthesis of betalains [1], [6], [11], [12], [13]. Regardless of the natural functions of PPOs in these organisms, melanin production by the enzyme is of great concern because post-harvest browning of produce results in considerable economic loss due to alteration of color, flavor, and nutritional value [6], [7], [14], [15], [16].

Many PPOs are believed to function in vivo as monomers, but a few fungal PPOs may associate to function as small oligomers (e.g., tetramers; reviewed in [2]). Regardless of the in vivo quaternary structure of a given PPO, molecular masses determined by SDS–PAGE are the “subunit molecular masses” of that PPO because SDS–PAGE causes dissociation of any oligomeric forms and unfolding of the individual subunits. One might expect subunit molecular masses for PPOs from various species to be similar, even if the oligomeric state of the functional PPO differs from species to species. However, subunit molecular masses reported in the literature for plant and fungal PPOs vary considerably, even for a single species, as described below. For a few species lower molecular mass subunits have been shown to result from proteolysis of a larger precursor form [17], [18], [19], [20], [21]. Through a review of experimental results reported in the literature and a comparative analysis of PPO protein sequences and structures, our aims are to explain the varying subunit molecular masses in terms of proteolytic cleavage sites located in specific sequence regions for PPOs from most species, and to relate this information to the three-dimensional structure of the enzyme.

Tyrosinase from the button mushroom Agaricus bisporus has been the subject of study for at least five decades by research groups around the globe. (For a review, see [22].) The subunit molecular mass of a latent form is generally reported to be ∼67 kDa [20], and that of an active form ranges from 43 to 45 kDa [20], [23], [24], [25].

The molecular masses reported for plant PPOs are also variable; a few examples follow. Vicia faba PPO (broad bean) is reported to exist as a 60–65 kDa latent form as well as a 43–45 kDa active form [18], [19]. Similarly, Vitis vinifera PPO (grape berry) exists as 60 kDa latent and 40 kDa active forms [17]. Forms of 53, 45, and 37 kDa have been reported for Metroxylon sagu PPO (sago palm) [26].

For V. faba and V. vinifera PPOs the ∼40 kDa active form has been shown to result from proteolytic cleavage of a C-terminal fragment from the ∼60 kDa latent form [18], [19], [27]; partial sequencing of the fragments by Edman degradation has established the cleavage site in each case. Similarly, the 75 kDa precursor of TYR from the ascomycete Neurospora crassa is C-terminally processed to a 46 kDa active form [21]. Protein sequence determination by electrospray mass spectrometry analysis of a 43 kDa active form of Larrea tridentata PPO (creosote bush) indicated that this PPO had undergone proteolysis of a C-terminal fragment [28]. Recent determination of the C-termini of an active 43.2 kDa TYR from the filamentous fungus Trichoderma reesei [29], [30] and an active 42 kDa TYR from the edible mushroom Pholiota nameko [31] indicated that these forms of the enzymes result from C-terminal proteolysis of the larger precursor TYRs encoded by the gene sequences.

C-terminal processing occurs for CO from the sweet potato Ipomoea batatas as well [32]. A crystal structure has been determined for an active 39 kDa form of I. batatas CO which is missing a C-terminal fragment, apparently as a result of in vivo proteolysis [33]. This form of I. batatas CO is a single-domain, globular protein containing a type-3 copper center. The I. batatas CO structure has been compared [4], [8], [34] with the structure of the homologous metalloprotein hemocyanin from Octopus dofleini [35]. O. dofleini hemocyanin has a type-3 copper center-containing domain very similar in structure to that of I. batatas CO, but it possesses an additional C-terminal domain relative to I. batatas CO. It has been proposed that the C-terminal portion of the larger, precursor I. batatas CO forms a discrete C-terminal domain (similar to that of O. dofleini hemocyanin) which covers the active site of the enzyme prior to its removal by proteolysis [8], [36], [37], [38].

Plant PPOs contain a bipartite N-terminal plastidic transit peptide directing the protein to the plastid lumen [16], [39]. This N-terminal transit peptide is cleaved off in two steps to generate the mature protein. N-terminal protein sequencing by Edman degradation of the mature protein has established the transit peptide cleavage site for I. batatas CO [32], [40], PPOE of Lycopersicon esculentum (cherry tomato) [41], POT32 of Solanum tuberosum tubers (potato) [42], Spinacia oleracea thylakoid PPO (spinach) [43], Triticum aestivum PPO (bread wheat) [44], V. faba PPO [18], [19], and V. vinifera PPO [27] (see Table 1). The transit peptide cleavage site for many other plants has been predicted based on homology. Although fungal TYRs do not have transit peptides, two reports indicate that TYRs from some fungi do undergo N-terminal proteolysis of a few residues. N-terminal protein sequencing of the active enzymes from Pycnoporus sanguineus [45] and T. reesei [29], [30] revealed that 4 and 18 residues had been cleaved from their N-termini, respectively. (In the case of T. reesei TYR, these 18 residues constitute a signal peptide. Unlike all fungal TYRs studied to date, T. reesei TYR is excreted [29].) On the other hand, N-terminal protein sequencing of N. crassa TYR indicates that this protein begins with the residue following the initial methionine encoded by the gene sequence [46].

We gathered from the literature all the available data on the N and/or C-terminal residues of PPOs as determined by N or C-terminal protein sequencing methods (or mass spectrometry or X-ray crystallography in a few cases). Primary literature references in which the N and C-terminal sequences were reported are given in Table 1. We compared the experimentally established N and/or C-termini of these PPOs with the protein sequences of other PPOs for which the N and C-termini have not been established by protein sequencing, in order to identify specific sequence regions where proteolysis is likely to occur for most PPOs. We then predicted the most likely sites of proteolysis within those sequence regions. Based on the known and predicted sites of proteolysis, we calculated molecular masses for the enzymes and used the calculated values to explain the varying subunit molecular masses reported in the literature. Finally, we related the sites of proteolysis and molecular masses to the two available PPO structures: I. batatas CO and Streptomyces castaneoglobisporus TYR.