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Polyphenol Oxidases in Crops: Biochemical, Physiological and Genetic Aspects

Francesca Taranto, Antonella Pasqualone, Giacomo Mangini and 4 more

International Journal of Molecular Sciences · 2017 · Review · CC BY

doi:10.3390/ijms18020377

Abstract

Enzymatic browning is a colour reaction occurring in plants, including cereals, fruit and horticultural crops, due to oxidation during postharvest processing and storage. This has a negative impact on the colour, flavour, nutritional properties and shelf life of food products. Browning is usually caused by polyphenol oxidases (PPOs), following cell damage caused by senescence, wounding and the attack of pests and pathogens. Several studies indicated that PPOs play a role in plant immunity, and emerging evidence suggested that PPOs might also be involved in other physiological processes. Genomic investigations ultimately led to the isolation of PPO homologs in several crops, which will be possibly characterized at the functional level in the near future. Here, focusing on the botanic families of Poaceae and Solanaceae, we provide an overview on available scientific literature on PPOs, resulting in useful information on biochemical, physiological and genetic aspects.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 21, 2026. Paragraph numbers are ours, so an answer can point at one.

1. Introduction

Polyphenol oxidases (PPOs) are a group of Cu-containing enzymes that catalyse the oxidation of several phenols to o-quinones. In turn, o-quinones are highly reactive molecules that can undergo non-enzymatic secondary reactions to form brown complex polymers known as melanins and cross-linked polymers with protein functional groups.

In plants, PPOs are nearly ubiquitous and located in chloroplasts. The loss of sub-cellular compartmentalization, due to senescence, wounding, interactions with pests and pathogens and handling during postharvest processing and storage, results in contact between PPOs and vacuolar phenolic substrates. This ultimately leads to a reaction known as enzymatic browning, negatively affecting colour, flavour, nutritional properties and shelf life of food products. However, in a few cases, enzymatic browning might be beneficial, as it leads to the formation of compounds conferring characteristic flavours.

Due to the economic importance of browning, PPOs have been extensively studied in relation to their physico-chemical properties. In contrast, knowledge on role of these enzymes in plant physiology is still limited and will possibly increase by the functional characterization of PPO homologs that are now being identified in several crop species. In this review, we take advantage of recent scientific literature to provide an overview on biochemical, physiological and genetic aspects of PPOs. Focus is given to the well-studied PPOs from the botanic families of Solanaceae and Poaceae. Few highlights in other main crops are also discussed.

2.1. PPO Biochemical Properties

The conversion of phenolic substrates to o-quinones by PPOs occurs by means of two oxidation steps. The first is the hydroxylation of the ortho-position adjacent to an existing hydroxyl group (“monophenol oxidase” or “monophenolase” activity, also referred to as hydroxylase or cresolase activity). The second is the oxidation of o-dihydroxybenzenes to o-benzoquinones (“diphenol oxidase” or “diphenolase activity”, also referred to as catecholase or oxidase activity) (Figure 1). Accordingly, PPOs correspond to the following sub-classes indicated by the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NC-IUBMB): (i) monophenol monooxygenases (E.C. 1.14.18.1); and (ii) enzymes catalysing the oxidation of diphenols using oxygen as electron acceptor (subclass E.C. 1.10.3.), including catechol oxidases (E.C. 1.10.3.1) and laccases (E.C. 1.10.3.2).

Based on the specific substrate and their mechanism of action, PPOs are classified in three different types: tyrosinases, cathecol oxidases and laccases. Tyrosinases have both cresolase and catecholase activities. Catechol oxidases, also known as o-diphenol oxidases, catalyse the oxidation of o-diphenols to o-quinones. Finally, laccases are capable to oxidize a wide spectrum of aromatic compounds by a radical catalysed reaction mechanism.

PPOs have been extensively studied with respect to physico-chemical properties and PPO isoenzymes can be distinguished based on electrophoretic mobility, optimal temperature and pH and substrate specificity. PPO activity depends on the pH, which affects the binding of substrates and the catalysis. Generally, the optimal pH of PPOs ranges between 4.0 and 8.0. In several Prunoideae (almond, apricot, peach, and plum), PPOs exhibit maximal activities around pH 5.0. Similarly, apple PPOs have an optimal pH from 4.5 to 5.0. Cherry and strawberry PPOs show a narrow pH optimum at about 4.5. PPOs from pineapple, having (+)-catechin as substrate, are more active around neutral pH, whereas PPO activity in kiwi is maximal at pH 8.0. The pH optimum of grape and olive PPOs depends on the cultivar, ranging from 3.5 to 7.3 and from 4.5 to 7.0, respectively. Concerning herbaceous crops, potato and wheat PPOs show two pH optima (4.5 and 5.0 in potato; 5.3 and 6.9 in wheat), due to the presence of two ionization states of the enzyme–substrate complex and the existence of two different acid dissociation constants (pKs). In sweet potato, two different PPO isoforms exhibit different pH optima, one at pH 5.4 and the other at 6.7. Lettuce PPOs show a broad pH optimum, from 5.0 to 8.0, whereas spinach PPOs have pH optimum around 8.0. Finally, eggplant has pH optima of 5.5 and 7.5 for diphenolase and monophenolase activity, respectively.

Temperature is another important factor significantly affecting the catalytic activity of PPO, as it influences the solubility of oxygen and may lead to enzyme denaturation. The optimal PPO temperature range varies for different plant sources: 25–35 °C in lettuce, 25–45 °C in grape and 30–50 °C in olive. PPOs from strawberry and cucumber show maximum activity at 50 °C. Grape PPOs show an activity reduction of approximately 50% after 20 min at 65 °C, and a complete inactivation after 15 min at 75 °C. At 60 °C, PPOs from apple have a half-life of 30 min. PPOs from mango require more than 15 min at 80 °C for 50% loss of activity. Cocoa beans and lettuce PPOs are inactivated after 5 min at 80 and 90 °C, respectively. Thermal stability of olive PPOs is cultivar-dependent. Indeed, about 50% reduction of the initial activity was observed in cv. Donat and Gemlik after 60 min at 40 °C. The same treatment was associated with 65% reduction of PPO activity in cv. Coratina. The type of phenolic substrate can also affect PPO thermal stability. In addition, sucrose and salts in the reaction environment act as protective agents against thermal denaturation.

2.2. Enzymatic Browning in Crops and Food Processing

Browning negatively affects the commercial value of many agricultural productions, including apple, banana, cucumber, grape, mango, pear, peach, apricot, eggplant, lettuce, potato and cereals. In a few cases, it is associated with the formation of flavour compounds positively influencing the food processing quality, for instance in the production of black tea, coffee and cocoa.

The quali-quantitative composition of crop PPO phenolic substrates is variable in different crop species, resulting in different browning intensities. Catechin is the major phenolic substrate in grapes and tea, chlorogenic acid is abundant in apple, potato, sunflower, sweet potato and eggplant, and ferulic acid is mainly present in wheat. Finally, catechin, epicatechin and caffeic acid derivatives are common substrates of several fruit PPOs.

Must browning can originate from enzymatic and non-enzymatic oxidation, both resulting in the formation of o-quinones. Overall, procyanidins and monomeric catechins are associated with higher browning degree than other phenolics. Enzymatic browning depends on tyrosinases and laccases, which are produced from grape berries and moulds, respectively, and is associated with the content of hydroxycinnamates such as caffeoyltartaric acid (caftaric acid) and p-coumaroyltartaric acid (coutaric acid). In addition, peroxidases (PODs) can enhance the degradation of phenols when coexisting with PPOs. Non-enzymatic oxidation is due to the conversion of polyphenols such as (+)-catechin/(−)-epicatechin, gallic acid, and caffeic acid, which are the most readily oxidized wine constituents.

Both PPO and POD enzymatic activities play a key role in determining the phenolic profile of olive oil. PPO activity is largely present in the olive mesocarp and is relatively constant after complete fruit ripening. In contrast, peroxidase activity is mainly located in the seed. An increase in PPO activity has been observed in response to abiotic stresses. Endogenous PPOs and PODs display their maximal activity mostly during the crushing and malaxation steps of the olive oil extraction process. Considering that olive PPOs have optimal activity at 50 °C and lowest stability at 40 °C, the malaxation temperature is a crucial factor influencing the phenolic content of olive oils.

In wheat, PPOs have been reported to be responsible for browning of whole meal flour, dough and several types of end-products such as Chinese steamed bread, Middle East flat breads, noodles and the Italian Castelvetrano durum wheat bread. PPOs are closely associated with the external layers of the caryopsis constituting the bran. Therefore, the distribution of PPOs in millstreams is closely related to the effectiveness of the milling procedure in removing the bran coat, and enzymatic browning is more evident in whole-meal flour and derived end-products. At least 90% of the PPOs are removed with a flour yield over 70% (i.e., the conventional extraction rate for refined flour).

In pasta, the yellow colour is related to the carotenoid content and the level of lypoxigenase activity of semolina. The red component is instead the result of Maillard reaction during drying. Finally, brownness depends on phenolics and PPOs. Interestingly, durum wheat cultivars have significantly lower PPO activity than cultivars of hexaploid wheat. Among the pasta-making steps, dough formation is potentially the most suitable to enzymatic browning, as it facilitates the interaction between enzyme and substrates. Today, the response of consumers to the browning of pasta is changing, due to the latest introduction of new pasta types obtained with whole meal flour or other cereals believed to be healthier.

2.3. Technological Strategies for Browning Prevention

Several practices can be followed by the food industry to prevent or limit enzymatic browning, including the use of reaction inhibitors, physical treatments and the use of by-product extracts and modified atmosphere. All these methods can be used alone or in combination. Differently acting chemical inhibitors, such as reducing (ascorbic acid and sulphites), chelating (ethylenediaminetetraacetate), acid (citric and malic), and complexing compounds (cyclodextrins), inhibit melanin formation by preventing the accumulation of o-quinones, or by forming stable colourless products. Sodium sulphite, bisulphite, and metabisulphite were extensively used in the fruit and vegetable processing industry, due to their low price. However, their commercialization is banned in several Countries because of adverse effects on health. As alternatives, ascorbic acid and 4-hexylresorcinol (4-HR; 4-hexyl-1,3-benzenediol) are frequently used. In addition, cyclodextrins have a promising anti-browning effect, as shown for potato and in fruit juices. A recent study pointed out that β-cyclodextrin binds non-covalently to PPOs better than other polysaccharides and has a relevant inhibiting ability under optimum conditions.

Physical treatments such as heat, dehydration, irradiation and high pressure are another strategy to minimize browning. Blanching treatments by steam, hot water, acid/salt water solutions, and microwave are also largely used. However, to balance the degree of inactivation of PPOs and the impact on the quality of end-products, time and temperature of blanching should be carefully set up.

Microwaved extracts obtained from by-products of the agro-industry, such as thinned fruits, represent another tool for the prevention of enzymatic browning. Indeed, the microwave treatment induces the formation of Maillard reaction products (MRPs), associated with antioxidant activity. A solution of 2% extract was reported to inhibit PPOs in minimally processed peaches for eight days of storage. Finally, modified atmosphere packaging with low permeability films is another effective way for extending shelf-life of processed fruit and vegetables, as oxygen is needed to trigger browning.

3. Physiological Roles of PPOs

PPOs have been largely investigated in relation to the commercial importance of browning. However, their role in plant physiology is less clear. Several studies report a positive correlation between PPO expression and resistance/tolerance to biotic stresses. For instance, potato varieties displaying enhanced PPO activity also exhibited higher tolerance to soft rot disease caused by the bacteria Pectobacterium atrosepticum, Pectobacterium carotovorum subsp. brasiliensis and Dickeya spp.. Similarly, PPO expression could be used as a biochemical marker to predict the outcome of the interaction between different tomato genotypes and the pathogens Ralstonia solanacearum and Xanthomonas axonopodis pv. vesicatoria, causing bacterial wilt and bacterial leaf spot disease, respectively. Concerning interactions with fungi, high and rapid accumulation of localized levels of PPOs were detected in pearl millet genotypes resistant to Sclerospora graminicola, the causal agent of downy mildew; conversely, susceptible cultivars failed to accumulate PPO even after a considerable time. Likewise, rapid and localized overexpression of multiple forms of PPOs was induced in resistant wheat and chickpea genotypes challenged with Fusarium graminearum and Fusarium oxysporum f. sp. ciceri, respectively. In plant–herbivore interactions, higher PPO contents were associated with increased potato tolerance to the Colorado beetle (Leptinotarsa decemlineata).

Direct evidence showing the defensive role of PPOs came from a few functional studies. Li and Steffens found that tomato plants overexpressing a potato PPO displayed enhanced resistance to Pseudomonas syringae pv. tomato, causal agent of the bacterial speck disease. Conversely, in plants transformed with antisense PPO cDNA, a significant reduction of PPO activity resulted in increased disease susceptibility. In poplar, transgenic overexpression of PPOs enhanced resistance to herbivory by forest tent caterpillar (Malacosoma disstria). Possibly, the characterization of the PPO gene family in different plant species and the availability of powerful tools for targeted gene inactivation and editing will provide further functional evidence on the defensive role of PPOs.

Many questions need to be answered on the physiology of PPOs. While it is generally assumed that PPOs play a role in defence responses to biotic stresses, the actual mechanism by which they exert their function is debated. In fact, they may act through: (1) direct toxicity of quinones; (2) reduced bioavailability and alkylation of cellular proteins to the pathogen; (3) cross-linking of quinones with protein or other phenolics, forming physical barriers; and (4) the production of reactive oxygen species (ROS), known to play an important role in defence signalling. The involvement of PPOs in physiological circumstances not involving the disruption of cell compartmentation, resulting in the contact of PPOs with their vacuolar phenolic substrates, is seemingly counter-intuitive. However, it is known that PPO levels vary in response to drought or other forms of abiotic stresses. For example, the up-regulation of PPO genes was observed in tomato plants exposed to water stress, especially in the abscission zone of leaves and in leaf veins. However, control plants resulted less tolerant to water stress than genotypes in which PPO activity was suppressed. Possibly, PPOs might have a role in photosynthesis as an oxygen buffer or interacting with the Mehler-peroxidase reaction. However, chloroplast phenolic substrates have not been identified.

4. Genomic and Genetic Aspects

The PPO gene family has been extensively studied in Poaceae, Solanaceae and in a few other species in which PPO activity affects economically important traits. Available genomic information for land plants indicates that the PPO gene family is extremely variable in size and structure, due to lineage-specific expansion and gene loss. In fact, no putative PPO genes were identified in some species such as Arabidopsis thaliana. Different PPO homologs might be associated with very different regulatory patterns and expression profiles. However, the factors influencing the expression of individual PPO members in plants are still unclear. PPO proteins include two highly conserved copper-binding domains (CuA and CuB), each with three histidine residues, together forming a Cu2+ binding site, which has been widely used to isolate PPO cDNAs. Typically, PPO proteins also encompass a N-terminal transit peptide, determining the import of PPOs into the thylakoid lumen, a dicopper centre, and a C-terminal region. Although these three motifs are also generally conserved, rare variants have been identified. Below, we report detailed information on genomic and genetic aspects of PPOs in several crop species.

4.1. Solanaceae

Six, seven and nine single-exon PPO genes were characterized in eggplant, tomato and potato, respectively (Table S1). Mapping of these genes revealed that they are located on chromosome 8, thus suggesting an evolution based on tandem duplication events

In tomato (Solanum lycopersicum), PPO genes cluster in a 165-KB genomic region. Protein sequence conservation among family members, named PPOs A, A’, B, C, D, E and F (Table S1), ranges from 63% to 95%. Spatial and temporal differences in the expression of tomato PPO homologs were described. PPO B is highly expressed in young leaves. Inflorescence also display high expression of PPO B together with PPO E and F. PPO A and C are specifically expressed in type I and type IV trichomes, whereas PPO D transcripts are only present in type VI trichomes. Promoter functional analysis mediated by GUS (β-Glucuronidase) fusion in transgenic tomato plants indicated that PPO F is responsive to abiotic and biotic challenges, thus suggesting a role in stress defence. PPO B is localized predominantly to mitotic or apoptotic tissues and is induced by ethylene. Homology analysis of its promoter sequence revealed hits with genes involved in response to hormones, signal transduction, phenylpropanoid biosynthesis, photosynthesis, and the synthesis of fruit and seed proteins.

In potato (Solanum tuberosum L.), five PPOs were originally identified, by means of PCR amplification with primers designed on conserved PPO regions in the copper-binding domain, named PPO-P1, PPO-P2, POT32, POT33 and POT72. A very high (>92%) protein identity was detected for the interspecific potato/tomato PPO pairs POT32/PPO D, POT33/PPO B, PPO-P2/PPO E, which might be indicative of functional conservation. A recent genome-wide search allowed the identification of nine PPO-like gene models, referred to as StuPPO1–9. StuPPO1/6 and StuPPO2/3/4/5/7/8 form two genomic clusters on chromosome 8 of 47 and 144 KB, respectively, whereas StuPPO9 is located on chromosome 2. Nucleotide similarities suggest that StuPPO1–4 are allelic variants of PPO-P1/P2, POT32, POT33 and POT72, respectively. The highest PPO activity occurred in roots, stolons, tuber buds and undeveloped flowers. Moderate levels of PPO activity were observed in young leaves, mature tubers, opened flowers, while low PPO activity was present in mature leaves and stems. PPO genes did not contribute equally to the total PPO protein content in tuber tissues, with StuPPO2 playing a major role. Low PPO activity and low browning tubers were obtained when the expression of StuPPO1 to StuPPO4 were simultaneously suppressed by means of artificial microRNAs. Recently, novel putative miRNAs of PPO have been identified in relation to stress resistance and brown spot of potato.

Based on nucleotide and protein similarity, the six eggplant (Solanum melongena L.) Ppo homologs were assigned to two structural classes defined as class A (SmePPO1–3) and class B (SmePPO4–6). A linkage map based on an interspecific S. melongena × Solanum incanum population allowed the identification of a cluster of five PPO homologs (SmePPO1–5) on linkage group 8, spanning an interval of 5.1 cM. Expression studies revealed that eggplant PPO transcripts are present in tissues upon mechanical cutting or pathogen attack, and accumulate in the stem and fruits. In more detail, SmePPO1 expresses in all tissues, such as roots, young and mature leaves and pre- and post-anthesis flowers; SmePPO2 predominantly expresses in root tissues; SmePPO3 and SmePPO4 express in all tissues except for the roots; and SmePPO5 and SmePPO6 display average expression in young leaves, sprouts and fruits.

In this study, we carried out a phylogenetic analysis within PPO amino acid sequences from Solanaceae. The homologs StuPPO5 and StuPPO7 were excluded from the analysis as they do not show a complete sequence and possibly represent non-functional peptides. The phylogenetic tree shown in Figure 2 is characterized by three clades. The first clade includes tomato PPO A–D, the eggplant class A (SmePPO1–3) and potato StuPPO3/4. Interestingly, except for SmePPO3, all the clade homologs for which the expression pattern is known (SmePPO1, SmePPO2, StuPPO2, and PPO A, C, and D) are transcribed in roots. The second clade includes tomato PPO E and F and the eggplant class B (SmePPO4–6), associated with defence responses. Notably, StuPPO8 and StuPPO9, forming a phylogenetically distant clade, are the only Solanaceae PPO genes structurally characterized by introns.

4.2. Poaceae

Most cloned and characterized PPO genes in Poaceae refer to tetraploid durum wheat and exaploid common wheat, thus they occur as duplicate or triplicate sets. Most studies mapped wheat PPO genes on the group 2 of homoeologous chromosomes, however a few works localized PPO loci on chromosomes of the groups 3, 5, 6 and 7. The first chromosomal localization of PPO genes was achieved using chromosome 2A substitution lines and the N2DT2A nullisomic-tetrasomic line, displaying higher PPO activity than other cytogenetic lines. Later, PPO activity was found to be associated with a major QTL on the chromosome arm 2AL, linked to the simple sequence repeat (SSR) marker Xgwm294, accounting for more than 80% genetic variation. PPO activity was also associated with the chromosome arm 2BL, even though the phenotypic effect in this case resulted to be lower than its homoeologous region on chromosome 2AL. Finally, a major QTL was detected on chromosome arm 2DL, which explained 23% of phenotypic PPO variation. Several studies suggested that PPO genes on the homoeologous group 2 are arranged in two paralogous families, named Ppo-1 (Ppo-A1, Ppo-B1 and Ppo-D1) and Ppo-2 (Ppo-A2, Ppo-B2 and Ppo-D2). Ppo-A2, Ppo-B2 and Ppo-D2 are located 8.9, 11.4 and 10.7 cM proximal to the respective paralogs of the Ppo-1 family.

The three homoeologous genes forming the Ppo-1 family display a very similar structure, as they consist in three exons and two introns. Among these genes, an insertion of 191-bp in the second intron of the Ppo-A1 gene represents the most important nucleotide variation. In contrast, Ppo-2 genes differ in structure, as Ppo-2A and Ppo-2D display one intron, whereas Ppo-B2 displays two. Interestingly, wheat PPO genes are among the few examples of homoeologous containing introns, shown to determine alternative splicing of premRNAs, in turn influencing the level of PPO activity.

Based on the Ppo-A1 gene sequence, three functional markers (PPO18, PPO33 and MG18) were developed to identify allelic variants underlying variation of PPO activity (Table S1). The functional marker MG08 was developed for Ppo-B1 and detected four allelic variants resulting in different PPO activity. Finally, two dominant functional markers, PPO16 and PPO29, mapped on the chromosome arm 2DL, were able to discriminate genotypes with low and high PPO activity.

Both Ppo-1 and Ppo-2 genes are expressed in developing kernels and affect PPO activity. However, they show variable expression levels. Specifically, Ppo-2 genes contribute to over 70% of PPO activity, and PPO genes on chromosome 2A generate nearly 90% of the PPO transcripts.

In barley (2n = 2x = 14; genome HH), the Ppo-1 and Ppo-2 paralogs are closely linked on the long arm of chromosome 2H. The respective orthologs were mapped on the chromosome arm 2HLch of wild barley Hordeum chilense (2n = 2x = 14; genome HchHch) and hexaploid Tritordeum (2n = 6x = 42; genome AABBHchHch). Association studies showed that both genes are involved in the phenol reaction of barley spikes and grains. RT-PCR experiments revealed that Ppo-1 was significantly expressed in awns and in hulls, with the highest content at two and three weeks after flowering, while Ppo-2 was specifically expressed in caryopses two weeks after flowering.

Oryza sativa includes two subspecies, japonica and indica, that are distinguished for grain reaction to phenol treatment. Indeed, after exposure to 1%–2% aqueous phenol solution, japonica seeds do not show colour change (negative response), whereas seeds of the indica type and wild Oryza species assume a dark brown or black coloration (positive response). Particularly, discoloration was found to be controlled by a single Mendelian gene (Phr1) mapped on chromosome 4 in an interval flanked by the two markers S100 and S115. The cDNA sequence of indica Phr1 showed high similarity with PPO genes and an amino acid sequence showing 68% identity with wheat PPO.

In Figure 3, we report the results of a phylogenetic analysis carried out using nucleotide sequences of wheat and barley PPOs. As expected, we found two clearly separated clades, corresponding to homologs of the Ppo-1 and Ppo-2 families.

4.3. Other Species

PPO activity in apricot (Prunus armeniaca L.), apple (Malus domestica) and banana (Musa ssp.) has been studied in relation to enzymatic browning during post-harvest stages. In apricot, the PA-PPO gene was isolated from immature-green fruits, where it is highly expressed. In an advanced stage of fruit development, PPO activity was detected, however no PA-PPO transcripts were detected in leaves, thus indicating high stability of the PPO proteins.

A recent study demonstrated that apple PPO genes are organized in clusters as observed for tomato. Ten genes were annotated as PPOs and located on three distinct chromosomes (2, 5 and 10), including one, five and four elements, respectively. Interestingly, chromosomes 5 and 10 can be considered as homologous, due to the recent apple genome duplication. The copper-binding domains of the apple pAPO5 and pMD-PPO2 proteins shared high conserved amino acids sequences with best-hit homologs in apricot (88% and 80%, respectively), grape berry (75% and 73%, respectively) and poplar (77% in both cases). Both pAPO5 and pMD-PPO2 genes are expressed in the early stages of fruit development.

High levels of PPO activity were detected in banana flesh throughout fruit growth and ripening. Four PPO cDNA were isolated from the banana fruit: BP01, BP011, BP034 and BP035. The BP01 genomic sequence was found to contain a 85-bp intron, whereas all the others were intronless. Expression analysis revealed a different expression pattern throughout the different stages of fruit development. BP01 was present in banana flesh early in development.

A recent work reported the cloning of a PPO gene in artichoke (Cynara cardunculus var. scolymus L.). In this species, browning may occur during early post-harvest stages and negatively affects quality. Based on degenerate primers, a CsPPO sequence was identified. Cis-acting elements were revealed in the promoter region, some of which are putatively involved in the response to light and wounds. The expression of CsPPO was significantly induced 48 h after wounding the capitula, even though the browning process had started earlier.

In walnut (Juglans regia), two PPO genes were characterized, named JrPPO1 and JrPPO2, which share 80% and 71% of nucleotide and amino acid sequence identity. JrPPO1 encodes a functional enzyme expressed primarily in the leaves, hulls, and flowers, whereas JrPPO2 is more expressed in callus tissues and in roots. In walnut, the role of PPO genes has been related to pathogen resistance and the health-promoting activity of phenolic compounds as antioxidant and radical scavenging activities. Very early work suggested that polyphenol oxidase activity produced a germicidal fluid that could inhibit bacterial growth.

5. Conclusions

The investigation of genes and mechanisms associated with PPO activity represents a pivotal challenge for food technology and plant breeding, considering the negative impact of enzymatic browning on the quality of primary productions and processed foods. Importantly, in the last few years, a large body of information is being accumulated on the genetic and genomic features of PPOs and their role in plant physiology. Possibly, our understanding on the role of PPO will benefit from recent advances in plant genomics and breakthrough discoveries in functional genetics, including targeted DNA modification by genome-editing.

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    Not in our library10.1111/j.1745-4514.1980.tb00875.x

  18. 18.

    Whitaker J.R. Principles of Enzymology for the Food Sciences. 1994;271-556.

    Not in our library

  19. 19.

    Janovitz-Klapp A., Richard F., Nicolas J. Polyphenoloxidase from apple, partial purification and some properties. Phytochemistry. 1989;28:2903-2907.

    Not in our library10.1016/0031-9422(89)80250-x

  20. 20.

    Valero E., Garcia-Carmona F. pH-dependent effect of sodium chloride on latent grape polyphenol oxidase. J. Agric. Food Chem. 1998;46:2447-2451.

    Not in our library10.1021/jf971088b

  21. 21.

    Fraignier M., Marques L., Fleuriet A., Macheix J. Biochemical and immunochemical characteristics of polyphenol oxidase from different fruits of Prunus. J. Agric. Food Chem. 1995;43:2375-2380.

    Not in our library10.1021/jf00057a011

  22. 22.

    Zhou P., Smith N.L., Lee C.Y. Potential purification and some properties of Monroe apple peel polyphenol oxidase. J. Agric. Food Chem. 1993;41:532-536.

    Not in our library10.1021/jf00028a004

  23. 23.

    Wesche-Ebeling P., Montgomery M.W. Strawberry polyphenoloxidase: Extraction and partial characterization. J. Food Sci. 1990;55:1320-1324.

    Not in our library10.1111/j.1365-2621.1990.tb03925.x

  24. 24.

    Park E.Y., Luh B.S. Polyphenol oxidase of kiwifruit. J. Food Sci. 1985;50:678-684.

    Not in our library10.1111/j.1365-2621.1985.tb13771.x

  25. 25.

    Ünal M.Ü., Taş C., Şener A. Determination of biochemical properties of polyphenol oxidase from domat olives. GIDA. 2011;36:185-192.

    Not in our library

  26. 26.

    Gençer N., Sinan S., Arslan O. Kinetic properties of polyphenol oxidase obtained from various olives (Olea europea L.). Asian J. Chem. 2012;24:2159-2161.

    Not in our library

  27. 27.

    Segovia-Bravo K.A., Jarén-Galán M., García-García P., Garrido-Fernández A. Characterization of polyphenol oxidase from the Manzanilla cultivar (Olea europaea pomiformis) and prevention of browning reactions in bruised olive fruits. J. Agric. Food Chem. 2007;55:6515-6520.

    Not in our library10.1021/jf063675f

  28. 28.

    Sánchez-Ferrer A., Laveda F., Garcia-Carmona F. Partial purification of soluble potato polyphenol oxidase by partitioning in aqueous two-phase system. J. Agric. Food Chem. 1993;41:1219-1224.

    Not in our library10.1021/jf00032a010

  29. 29.

    Interesse F.S., Ruggiero P., D’Avella G., Lamparelli F. Partial purification of some properties of wheat (Triticum aestivum) o-diphenolase. J. Sci. Food Agric. 1980;31:459-466.

    Not in our library10.1002/jsfa.2740310507

  30. 30.

    Nozue M., Souri M., Arakawa D., Kojima M. Purification and characterization of two isoforms of chlorogenic acid oxidase from sweet potato cells in suspension culture. J. Plant Physiol. 1998;153:552-557.

    Not in our library10.1016/s0176-1617(98)80203-4

  31. 31.

    Heimdal H., Larsen L.M., Poll L. Characterization of polyphenol oxidase from photosynthetic and vascular lettuce tissues (Lactuca sativa). J. Agric. Food Chem. 1994;42:1428-1433.

    Not in our library10.1021/jf00043a008

  32. 32.

    Sheptovitsky Y.G., Brudvig G.W. Isolation and characterization of spinach photosystem I1 membrane-associated catalase and polyphenol oxidase. Biochemistry. 1996;35:16255-16263.

    Not in our library10.1021/bi9613842

  33. 33.

    Perez-Gilabert M., Carmona F.G. Characterization of catecholase and cresolase activities of eggplant polyphenol oxidase. J. Agric. Food Chem. 2000;48:695-700.

    Not in our library10.1021/jf990292r

  34. 34.

    Taticchi A., Esposto S., Veneziani G., Urbani S., Selvaggini R., Servili M. The influence of the malaxation temperature on the activity of polyphenoloxidase and peroxidase and on the phenolic composition of virgin olive oil. Food Chem. 2013;136:975-983.

    Not in our library10.1016/j.foodchem.2012.08.071

  35. 35.

    Serradell M.D.L.A., Rozenfeld P.A., Martinez G.A., Civello P.M., Chaves A.R., Anon M.C. Polyphenoloxidase activity from strawberry fruit (Fragaria × ananassa, Duch., cv. Selva) characterization and partial purification. J. Sci. Food Agric. 2000;80:1421-1427.

    Not in our library

  36. 36.

    Miller A.R., Kelley T.J., Mujer C.V. Anodic peroxidase isoenzymes and polyphenol oxidase activity from cucumber fruit: Tissue and substrate specificity. Phytochemistry. 1990;29:705-709.

    Not in our library10.1016/0031-9422(90)80005-2

  37. 37.

    Robinson S.P., Loveys B.R., Chacko E.K. Polyphenoloxidase enzymes in the sap and skin of mango fruit. Aust. J. Plant Physiol. 1993;20:99-107.

    Not in our library

  38. 38.

    Lee P.M., Lee K., Karim M.I.A. Biochemical studies of cocoa bean polyphenol oxidase. J. Sci. Food Agric. 1991;55:251-260.

    Not in our library10.1002/jsfa.2740550210

  39. 39.

    Lourenco E.J., Neves V.A., Silva M.A.D. Polyphenol oxidase from sweet potato: Purification and properties. J. Agric. Food Chem. 1992;40:2369-2373.

    Not in our library10.1021/jf00024a009

  40. 40.

    Reinkensmeier A., Steinbrenner K., Homann T., Bußler S., Rohn S., Rawel H.M. Monitoring the apple polyphenol oxidase-modulated adduct formation of phenolic and amino compounds. Food Chem. 2016;194:76-85.

    Not in our library10.1016/j.foodchem.2015.07.145

  41. 41.

    Gooding P.S., Bird C., Robinson S.P. Molecular cloning and characterization of banana fruit polyphenol oxidase. Planta. 2001;213:748-757.

    Not in our library10.1007/s004250100553

  42. 42.

    Rathjen A.H., Robinson S.P. Aberrant processing of polyphenol oxidase in a variegated grapevine mutant. Plant Physiol. 1992;99:1619-1625.

    Not in our library10.1104/pp.99.4.1619

  43. 43.

    Saba M.K., Moradi S. Internal browning disorder of eight pear cultivars affected by bioactive constituents and enzyme activity. Food Chem. 2016;205:257-263.

    Not in our library10.1016/j.foodchem.2016.03.022

  44. 44.

    Docimo T., Francese G., de Palma M., Mennella D., Toppino L., Lo Scalzo R., Mennella G., Tucci M. Insights in the fruit flesh browning mechanisms in Solanum melongena genetic lines with opposite post-cut behavior. J. Agric. Food Chem. 2016;64:4675-4685.

    Not in our library10.1021/acs.jafc.6b00662

  45. 45.

    Taranto F., Delvecchio L.N., Mangini G., Del Faro L., Blanco A., Pasqualone A. Molecular and physico-chemical evaluation of enzymatic browning of whole meal and dough in a collection of tetraploid wheats. J. Cereal Sci. 2012;55:405-414.

    Not in our library10.1016/j.jcs.2012.02.005

  46. 46.

    Pasqualone A., Delvecchio L.N., Mangini G., Taranto F., Blanco A. Variability of total soluble phenolic compounds and antioxidant activity in a collection of tetraploid wheat. Agric. Food Sci. 2014;23:307-316.

    Not in our library

  47. 47.

    Ullah Tea M.R., Fox P.F. Food Enzymotogy. 1991;163-187.

    Not in our library

  48. 48.

    Amorim H.W., Melo M., Fox P.F. Significance of enzymes in nonalcoholic coffee beverage. Food Enzymlogy. 1991;189-209.

    Not in our library

  49. 49.

    Jaworski A.W., Lee C.Y. Fractionation of HPLC determination of grape phenolics. J. Agric. Food Chem. 1987;35:257-259.

    Not in our library10.1021/jf00074a022

  50. 50.

    Murata M., Tsurutani M., Tomita M., Homma S., Kaneko K. Relationship between apple ripening and browning: Changes in polyphenol content and polyphenol oxidase. J. Agric. Food Chem. 1995;43:1115-1121.

    Not in our library10.1021/jf00053a001

  51. 51.

    Raymond J., Rakariyatham N., Azanza J.L. Purification and some properties of polyphenoloxidase from sunflower seeds. Phytochemistry. 1993;34:927-931.

    Not in our library10.1016/s0031-9422(00)90689-7

  52. 52.

    Stommel J.R., Whitaker B.D. Phenolic acid content and composition of eggplant fruit in a germplasm core subset. J. Am. Soc. Hortic. Sci. 2003;128:704-710.

    Not in our library

  53. 53.

    Laddomada B., Durante M., Mangini G., D’Amico L., Lenucci M.S., Simeone R., Piarulli L., Mita G., Blanco A. Genetic variation for phenolic acids concentration and composition in a tetraploid wheat (Triticum turgidum L.) collection. Genet. Resour. Crop Evol. 2016.

    Not in our library10.1007/s10722-016-0386-z

  54. 54.

    Macheix J., Fleuriet A., Billot J., Macheix J., Fleuriet A., Billot J. Phenolic compounds in fruit processing. Fruit Phenolics. 1990;239-312.

    Not in our library

  55. 55.

    Lee C.Y., Jaworski A.W. Phenolics and browning potential of white grapes grown in New York. Am. J. Enol. Vitic. 1988;39:337-340.

    Not in our library

  56. 56.

    Li H., Guo A., Wang H. Mechanisms of oxidative browning of wine. Food Chem. 2008;108:1-13.

    Not in our library10.1016/j.foodchem.2007.10.065

  57. 57.

    Ortega-García F., Blanco S., Peinado M.A., Peragón J. Polyphenol oxidase and its relationship with oleuropein concentration in fruits and leaves of olive (Olea europaea) cv. Picual trees during fruit ripening. Tree Physiol. 2008;28:45-54.

    Not in our library10.1093/treephys/28.1.45

  58. 58.

    García-Rodríguez R., Romero-Segura C., Sanz C., Sánchez-Ortiz A., Pérez A.G. Role of polyphenol oxidase and peroxidase in shaping the phenolic profile of virgin olive oil. Food Res. Int. 2011;44:629-635.

    Not in our library10.1016/j.foodres.2010.12.023

  59. 59.

    Ortega-García F., Peragón J. The response of phenylalanine ammonia lyase, polyphenol oxidase and phenols to cold stress in the olive tree (Olea europaea L. cv. Picual). J. Sci. Food Agric. 2009;89:1565-1573.

    Not in our library10.1002/jsfa.3625

  60. 60.

    Servili M., Taticchi A., Esposto S., Urbani S., Selvaggini R., Montedoro G. Influence of the decrease in oxygen during malaxation of olive paste on the composition of volatiles and phenolic compounds in virgin olive oil. J. Agric. Food Chem. 2008;56:10048-10055.

    Not in our library10.1021/jf800694h

  61. 61.

    Singh R., Sheoran I.S. Enzymic browning of whole wheat meal flour. J. Sci. Food Agric. 1972;23:121-125.

    Not in our library10.1002/jsfa.2740230115

  62. 62.

    Baik B.K., Czuchajowska Z.U., Pomeranz Y.E. Discoloration of dough for oriental noodles. Cereal Chem. 1995;72:198-204.

    Not in our library

  63. 63.

    Dexter J.E., Preston K.R., Matsuo R.R., Tipples K.H. Development of a high extraction flow for the GRL pilot mill to evaluate Canadian wheat potential for the Chinese market. Can. Inst. Food Sci. Technol. J. 1984;14:253-259.

    Not in our library10.1016/s0315-5463(84)72567-3

  64. 64.

    Faridi H., Pomeranz Y. Flat breads. Wheat: Chemistry and Technology. 1988;Volume 2:457-506.

    Not in our library

  65. 65.

    Miskelly D.M. Flour components affecting pasta and noodle color. J. Sci. Food Agric. 1984;35:463-471.

    Not in our library10.1002/jsfa.2740350417

  66. 66.

    Morris C.F., Jeffers H.C., Engle D.A. Effect of processing, formula and measurement variables on alkaline noodle color-toward an optimized laboratory system. Cereal Chem. 2000;77:77-85.

    Not in our library10.1094/cchem.2000.77.1.77

  67. 67.

    Pasqualone A., Clerici P.S.M.T. Italian durum wheat breads. Bread Consumption and Health. 2012;57-80.

    Not in our library

  68. 68.

    Hatcher D.W., Kruger J.E. Distribution of polyphenol oxidase in flour millstreams of Canadian common wheat classes milled to three extraction rates. Cereal Chem. 1993;70:51-55.

    Not in our library

  69. 69.

    Demeke T., Morris C. Molecular characterization of wheat polyphenol oxidase (PPO). Theor. Appl. Genet. 2002;104:813-818.

    Not in our library10.1007/s00122-001-0847-3

  70. 70.

    Feillet P., Autran J.C., Icard-Verniere C. Pasta brownness: An assessment. J. Cereal Sci. 2000;32:215-233.

    Not in our library10.1006/jcrs.2000.0326

  71. 71.

    Pasqualone A., Delvecchio L.N., Gambacorta G., Laddomada B., Urso V., Mazzaglia A., Ruisi P., di Miceli G. Effect of supplementation with wheat bran aqueous extracts obtained by ultrasound-assisted technologies on the sensory properties and the antioxidant activity of dry pasta. Nat. Prod. Commun. 2015;10:1739-1742.

    Not in our library

  72. 72.

    Pasqualone A., Gambacorta G., Summo C., Caponio F., di Miceli G., Flagella Z., Marrese P.P., Piro G., Perrotta C., de Bellis L., et al. Functional, textural and sensory properties of dry pasta supplemented with lyophilized tomato matrix or with durum wheat bran extracts produced by supercritical carbon dioxide or ultrasound. Food Chem. 2016;213:545-553.

    Not in our library10.1016/j.foodchem.2016.07.006

  73. 73.

    Landi M., Degl’Innocenti E., Guglielminetti L., Guidi L. Role of ascorbic acid in the inhibition of polyphenol oxidase and the prevention of browning in different browning-sensitive Lactuca sativa var. capitata (L.) and Eruca sativa (Mill.) stored as fresh-cut produce. J. Sci. Food Agric. 2013;93:1814-1819.

    Not in our library10.1002/jsfa.5969

  74. 74.

    Soares J.M., Fonseca G.G. Effect of L-ascorbic acid and sodium metabisulfite in the inhibition of the enzymatic browning of minimally processed apple. Int. J. Agric. Res. 2008;3:196-201.

    Not in our library

  75. 75.

    Richard-Forget F.C., Goupy P.M., Nicolas J.J. Cysteine as an inhibitor of enzymic browning. 2. Kinetic studies. J. Agric. Food Chem. 1992;40:2108-2113.

    Not in our library10.1021/jf00023a014

  76. 76.

    Gacchea R.N., Shete A.M., Dhole N.A., Ghole V.S. Reversible inhibition of polyphenol oxidase from apple using l-cysteine. Indian J. Chem. Technol. 2006;13:459-463.

    Not in our library

  77. 77.

    Ashie I.N.A., Simpson B.K., Smith J.P. Mechanisms for controlling enzymatic reactions in foods. Crit. Rev. Food Sci. Nutr. 1996;36:1-30.

    Not in our library10.1080/10408399609527716

  78. 78.

    Sapers G.M. Browning of foods: Control by sulfites, antioxidants, and other means. Food Technol. 1993;47:75-84.

    Not in our library

  79. 79.

    Nicolas J.J., Richard-Forget F.C., Goupy P.M., Amiot M., Aubert S.Y. Enzymatic browning reactions in apple and apple products. Crit. Rev. Food Sci. 1994;34:109-157.

    Not in our library10.1080/10408399409527653

  80. 80.

    Eskin N.A.M., Eskin N.A.M. Biochemistry of food spoilage: Enzymatic browning. Biochemistry of Foods. 1990;401-432.

    Not in our library

  81. 81.

    Martinez M.V., Whitaker J.R. The biochemistry and control of enzymatic browning. Trends Food Sci. Technol. 1995;6:195-200.

    Not in our library10.1016/s0924-2244(00)89054-8

  82. 82.

    Gultekin F., Doguc D.K. Allergic and immunologic reactions to food additives. Clin. Rev. Allerg. Immunol. 2013;45:6-29.

    Not in our library10.1007/s12016-012-8300-8

  83. 83.

    Fayad N., Marchal L., Billaud C., Nicolas J. Comparison of β-cyclodextrin effect on polyphenol oxidation catalyzed by purified polyphenol oxidase from different sources. J. Agric. Food Chem. 1997;45:2442-2446.

    Not in our library10.1021/jf9607932

  84. 84.

    Hicks K.B., Haines R.M., Tong C.B., Sapers G.M., El-Atawy Y., Irwin P.L., Seib P.A. Inhibition of enzymatic browning in fresh fruit and vegetable juices by soluble and insoluble forms of β-cyclodextrin alone or in combination with phosphates. J. Agric. Food Chem. 1996;44:2591-2594.

    Not in our library10.1021/jf960181h

  85. 85.

    Singh V., Jadhav S.B., Singhal R.S. Interaction of polyphenol oxidase of Solanum tuberosum with β-cyclodextrin: Process details and applications. Int. J. Biol. Macromol. 2015;80:469-474.

    Not in our library10.1016/j.ijbiomac.2015.07.010

  86. 86.

    Gonzalez-Fesler M., Salvatori D., Gomez P., Alzamora S.M. Convective air drying of apples as affected by blanching and calcium impregnation. J. Food Eng. 2008;87:323-332.

    Not in our library10.1016/j.jfoodeng.2007.12.007

  87. 87.

    Severini C., Baiano A., de Pilli T., Carbone B.F., Derossi A. Combined treatments of blanching and dehydration: Study on potato cubes. J. Food Eng. 2005;68:289-296.

    Not in our library10.1016/j.jfoodeng.2004.05.045

  88. 88.

    Wang R., Zhang M., Arun S., Mujumdar A.S. Effects of vacuum and microwave freeze drying on microstructure and quality of potato slices. J. Food Eng. 2010;101:131-139.

    Not in our library10.1016/j.jfoodeng.2010.05.021

  89. 89.

    Kidmose U., Martens H.J. Changes in texture, microstructure and nutritional quality of carrot slices during blanching and freezing. J. Sci. Food Agric. 1999;79:1747-1753.

    Not in our library10.1002/(sici)1097-0010(199909)79:12<1747::aid-jsfa429>3.0.co;2-b

  90. 90.

    Mukherjee S., Chattopadhyay P.K. Whirling bed blanching of potato cubes and its effects on product quality. J. Food Eng. 2007;78:52-60.

    Not in our library10.1016/j.jfoodeng.2005.09.001

  91. 91.

    Sapers G.M., Miller R.C. Heated ascorbic/citric acid solution as browning inhibitor for pre-peeled potatoes. J. Food Sci. 1995;60:762-766.

    Not in our library10.1111/j.1365-2621.1995.tb06223.x

  92. 92.

    Severini C., Baiano A., de Pilli T., Romaniello R., Derossi A. Prevention of enzymatic browning in sliced potatoes by blanching in boiling saline solutions. LWT Food Sci. Technol. 2003;36:657-665.

    Not in our library10.1016/s0023-6438(03)00085-9

  93. 93.

    Severini C., de Pilli T., Baiano A., Mastrocola D., Massini R. Study on preventing enzymatic browning of potato by microwave blanching. Sci. Aliment. 2001;21:149-160.

    Not in our library10.3166/sda.21.149-160

  94. 94.

    De Leonardis A., Lustrato G., Macciola V., Ranalli G. Application of chemical and physical agents in model systems to controlling phenoloxidase enzymes. Eur. Food Res. Technol. 2010;231:603-610.

    Not in our library10.1007/s00217-010-1312-9

  95. 95.

    Redondo D., Venturini M.E., Oria R., Arias E. Inhibitory effect of microwaved thinned nectarine extracts on polyphenol oxidase activity. Food Chem. 2016;197:603-610.

    Not in our library10.1016/j.foodchem.2015.11.009

  96. 96.

    Ngazee E., Icishahayo D., Coutinho T.A., Van der Waals J.E. Role of polyphenol oxidase, peroxidase, phenylalanine ammonia lyase, chlorogenic acid, and total soluble phenols in resistance of potatoes to soft rot. Plant Dis. 2012;96:186-192.

    Not in our library10.1094/pdis-02-11-0149

  97. 97.

    Vanitha S.C., Niranjana S.R., Umesha S. Role of phenylalanine ammonia lyase and polyphenol oxidase in host resistance to bacterial wilt of tomato. J. Phytopathol. 2009;157:552-557.

    Not in our library10.1111/j.1439-0434.2008.01526.x

  98. 98.

    Kavitha R., Umesha S. Regulation of defense-related enzymes associated with bacterial spot resistance in tomato. Phytoparasitica. 2008;36:144-159.

    Not in our library10.1007/bf02981327

  99. 99.

    Niranjan Ray S., Sarosh B.R., Shetty H.S. Induction and accumulation of polyphenol oxidase activities as implicated in development of resistance against pearl millet downy mildew disease. Funct. Plant Biol. 2006;33:563-571.

    Not in our library10.1071/fp06003

  100. 100.

    Mohammadi M., Kazemi H. Changes in peroxidase and polyphenol oxidase activities in susceptible and resistant wheat heads inoculated with Fusarium graminearum and induced resistance. Plant Sci. 2002;162:491-498.

    Not in our library10.1016/s0168-9452(01)00538-6

  101. 101.

    Raju S., Jayalakshmi S.K., Sreeramulu K. Comparative study on the induction of defense related enzymes in two different cultivars of chickpea (Cicer arietinum L.) genotypes by salicylic acid, spermin and Fusarium oxysporum f. sp. ciceri. Aust. J. Crop Sci. 2008;2:121-140.

    Not in our library

  102. 102.

    Jayalakshmi S.K., Raju S., Rani S.U., Benagi V.I., Sreeramulu K. Trichoderma harzianum L1 as a potential source for lytic enzymes and elicitor of defense responses in chickpea (Cicer arietinum L.) against wilt disease caused by Fusarium oxysporum f. sp. ciceri. Aust. J. Crop Sci. 2009;3:44-52.

    Not in our library

  103. 103.

    Castañera P., Steffens J.C., Tingey W.M. Biological performance of Colorado potato beetle larvae on potato genotypes with differing levels of polyphenol oxidase. J. Chem. Ecol. 1996;22:91-101.

    Not in our library10.1007/bf02040202

  104. 104.

    Li L., Steffens J.C. Overexpression of polyphenol oxidase in transgenic tomato plants results in enhanced bacterial disease resistance. Planta. 2002;2:239-247.

    Not in our library10.1007/s00425-002-0750-4

  105. 105.

    Thipyapong P., Melkonian J., Wolfe D.W., Steffens J.C. Suppression of polyphenol oxidases increases stress tolerance in tomato. Plant Sci. 2004;167:693-703.

    Not in our library10.1016/j.plantsci.2004.04.008

  106. 106.

    Wang J., Constabel C.P. Polyphenol oxidase overexpression in transgenic Populus enhances resistance to herbivory by forest tent caterpillar (Malacosoma disstria). Planta. 2004;220:87-96.

    Not in our library10.1007/s00425-004-1327-1

  107. 107.
    The polyphenol oxidase gene family in land plants: Lineage-specific duplication and expansion · 2012
    No notice
  108. 108.

    Araji S., Grammer T.A., Gertzen R., Anderson S.D., Mikulic-Petkovsek M., Veberic R., Phu M.L., Solar A., Leslie C.A., Dandekar A.M., et al. Novel roles for the polyphenol oxidase enzyme in secondary metabolism and the regulation of cell death in walnut. Plant Physiol. 2014;164:1191-1203.

    Not in our library10.1104/pp.113.228593

  109. 109.

    Boeck T., Winters A.L., Webb K.J., Kingston-Smith A.H. Polyphenol oxidase in leaves; is there any significance to the chloroplastic localization? J. Exp. Bot. 2015;66:3571-3579.

    Not in our library10.1093/jxb/erv141

  110. 110.

    Boeckx T., Webster R., Winters A.L., Webb K.J., Gay A., Kingston-Smith A.H. Polyphenol oxidase-mediated protection against oxidative stress is not associated with enhanced photosynthetic efficiency. Ann. Bot. 2015;116:529-540.

    Not in our library10.1093/aob/mcv081

  111. 111.

    Thipyapong P., Hunt M.D., Steffens J.C. Antisense down regulation of poly-phenol oxidase results in enhanced disease susceptibility. Planta. 2004;220:105-117.

    Not in our library10.1007/s00425-004-1330-6

  112. 112.

    Shetty SM., Chandrashekar A., Venkatesh Y.P. Eggplant polyphenol oxidase multigene family: Cloning, phylogeny, expression analyses and immunolocalization in response to wounding. Phytochemistry. 2011;72:2275-2287.

    Not in our library10.1016/j.phytochem.2011.08.028

  113. 113.

    Newman S.M., Tantasawat P., Steffens J.C. Tomato polyphenol oxidase B is spatially and temporally regulated during development and in response to ethylene. Molecules. 2011;16:493-517.

    Not in our library10.3390/molecules16010493

  114. 114.

    Chi M., Bhagwat B., Lane D., Tang G., Su Y., Sun R., Oomah B.D., Wiersma P.A., Xiang Y. Reduced polyphenol oxidase gene expression and enzymatic browning in potato (Solanum tuberosum L.) with artificial microRNAs. BMC Plant Biol. 2014;14:62.

    Not in our library10.1186/1471-2229-14-62

  115. 115.

    Newman S.M., Eannetta N.T., Yu H., Prince J.P., de Vicente M.C., Tanksley S.D., Steffens J.C. Organization of the tomato polyphenol oxidase gene family. Plant Mol. Biol. 1993;21:1035-1051.

    Not in our library10.1007/bf00023601

  116. 116.

    Thipyapong P., Steffens J.C. Tomato polyphenol oxidase: Differential response of the polyphenol oxidase F promoter to injuries and wound signals. Plant Physiol. 1997;115:409-418.

    Not in our library10.1104/pp.115.2.409

  117. 117.

    Thygesen P.W., Dry I.B., Robinson S.P. Polyphenol oxidase in potato. Plant Physiol. 1995;109:525-531.

    Not in our library10.1104/pp.109.2.525

  118. 118.

    Chi M., Liu C., Su Y., Tong Y., Liu H. Bioinformatic prediction of upstream microRNAs of PPO and novel microRNAs in potato. Can. J. Plant Sci. 2015;95:871-877.

    Not in our library10.4141/cjps-2014-308

  119. 119.

    Gramazio P., Prohens J., Plazas M., Andújar I., Castillo E., Herraiz F.J., Knapp S., Meyer S.R., Vilanova S. Location of the chlorogenic acid biosynthesis pathway and polyphenol oxidase genes in a new interspecific anchored linkage map of eggplant. BMC Plant Biol. 2014;14:350.

    Not in our library10.1186/s12870-014-0350-z

  120. 120.

    Jiménez M., Dubcovsky J. Chromosome location of genes affecting polyphenol oxidase activity in seeds of common and durum wheat. Plant Breed. 1999;118:395-398.

    Not in our library10.1046/j.1439-0523.1999.00393.x

  121. 121.

    Zhang L.P., Ge X.X., He Z.H., Wang D.S., Yan J., Xia X.C., Sutherland M.W. Mapping QTLs for polyphenol oxidase activity in a DH population from common wheat. Acta Agron. Sin. 2005;31:7-10.

    Not in our library

  122. 122.

    Watanabe N., Takeuchi A., Nakayama A. Inheritance and chromosomal location of the homoeologous genes affecting phenol colour reaction of kernels in durum wheat. Euphytica. 2004;139:87-93.

    Not in our library10.1007/s10681-004-2255-7

  123. 123.

    Mares D.J., Campbell A.W. Mapping components of flour and noodle colour in Australian wheat. Aust. J. Agric. Res. 2001;52:1297-1309.

    Not in our library10.1071/ar01048

  124. 124.

    Jukanti A.K., Bruckner P.L., Fischer A.M. Evaluation of wheat polyphenol oxidase genes. Cereal Chem. 2004;81:481-485.

    Not in our library10.1094/cchem.2004.81.4.481

  125. 125.

    Beecher B.S., Carter H.A., See D.E. Genetic mapping of new seed-expressed polyphenol oxidase genes in wheat (Triticum aestivum L.). Theor. Appl. Genet. 2012;124:1463-1473.

    Not in our library10.1007/s00122-012-1801-2

  126. 126.

    Beecher B.S., Skinner D.Z. Molecular cloning and expression analysis of multiple polyphenol oxidase genes in developing wheat (Triticum aestivum L.) kernels. J. Cereal Sci. 2011;53:371-378.

    Not in our library10.1016/j.jcs.2011.01.015

  127. 127.

    Martin J.M., Berg J.E., Hofer P., Kephart K.D., Nash D., Bruckner P.L. Allelic variation of polyphenol oxidase genes impacts on Chinese raw noodle color. J. Cereal Sci. 2011;54:387-394.

    Not in our library10.1016/j.jcs.2011.08.003

  128. 128.

    Taranto F., Mangini G., Pasqualone A., Gadaleta A., Blanco A. Mapping and allelic variations of Ppo-B1 and Ppo-B2 gene-related polyphenol oxidase activity in durum wheat. Mol. Breed. 2015;35:80.

    Not in our library10.1007/s11032-015-0272-y

  129. 129.

    Sun Y., He Z., Ma W., Xia X. Alternative splicing in the coding of Ppo-A1 directly influences the polyphenol oxidase activity in common wheat (Triticum aestivum L.). Funct. Integr. Genom. 2011;11:85-93.

    Not in our library10.1007/s10142-010-0201-4

  130. 130.

    Sun D.J., He Z.H., Xia X.C., Zhang L.P., Morris C.F., Appels R., Ma W.J., Wang H. A novel STS marker for polyphenol oxidase activity in bread wheat. Mol. Breed. 2005;16:209-218.

    Not in our library10.1007/s11032-005-6618-0

  131. 131.

    Mangini G., Taranto F., Delvecchio L.N., Pasqualone A., Blanco A. Development and validation of a new Ppo-A1 marker useful for marker-assisted selection in tetraploid wheats. Mol. Breed. 2014;34:385-392.

    Not in our library10.1007/s11032-014-0041-3

  132. 132.

    He X.Y., He Z.H., Zhang L.P., Sun D.J., Morris C.F., Fuerst E.P., Xia X.C. Allelic variation of polyphenol oxidase (PPO) genes located on chromosomes 2A and 2D and development of functional markers for the PPO genes in common wheat. Theor. Appl. Genet. 2007;115:47-58.

    Not in our library10.1007/s00122-007-0539-8

  133. 133.

    Taketa S., Matsuki K., Amano S., Saisho D., Himi E., Shitsukawa N., You T., Noda K., Takeda K. Duplicate polyphenol oxidase genes on barley chromosome 2H and their functional differentiation in the phenol reaction of spikes and grains. J. Exp. Bot. 2010;61:3983-3993.

    Not in our library10.1093/jxb/erq211

  134. 134.

    Rodríguez-Suárez C., Atienza S. Polyphenol oxidase genes in Hordeum chilense and implications in tritordeum breeding. Mol. Breed. 2014;34:1867.

    Not in our library10.1007/s11032-014-0145-9

  135. 135.

    Yu Y., Tang T., Qian Q., Wang Y., Yan M., Zeng D., Han B., Wu C.I., Shi S., Li J. Independent loss of function in a polyphenol oxidase in rice: Differentiation in grain discoloration between subspecies and the role of positive selection under domestication. Plant Cell. 2008;20:2946-2959.

    Not in our library10.1105/tpc.108.060426

  136. 136.
    A multidisciplinary approach providing new insight into fruit flesh browning physiology in apple (Malus x domestica Borkh.). · 2013
    Not checked
  137. 137.

    Shekhawat U.K.S., Ganapathi T.R. Transgenic banana plants overexpressing MusabZIP53 display severe growth retardation with enhanced sucrose and polyphenol oxidase activity. Plant Cell Tissue Organ Cult. 2014;116:387-402.

    Not in our library10.1007/s11240-013-0414-z

  138. 138.

    Chevalier T., de Rigal D., Mbéguié-A-Mbéguié D., Gauillard F., Richard-Forget F., Fils-Lycaon B.R. Molecular cloning and characterization of apricot fruit polyphenol oxidase. Plant Physiol. 1999;119:1261-1269.

    Not in our library10.1104/pp.119.4.1261

  139. 139.

    Boss P.K., Gardner R.C., Janssen B.J., Ross G.S. An apple polyphenol oxidase cDNA is up-regulated in wounded tissues. Plant Mol. Biol. 1995;27:429-433.

    Not in our library10.1007/bf00020197

  140. 140.

    Kim J.Y., Seo Y.S., Kim J.E., Sung S.K., Song K.J., An G.H., Kim W.T. Two polyphenol oxidases are differentially expressed during vegetative and reproductive development and in response to wounding in the Fuji apple. Plant Sci. 2001;161:1145-1152.

    Not in our library10.1016/s0168-9452(01)00522-2

  141. 141.

    Quarta A., Mita G., Durante M., Arlorio M., de Paolis A. Isolation of a polyphenol oxidase (PPO) cDNA from artichoke and expression analysis in wounded artichoke heads. Plant Physiol. Biochem. 2013;68:52-60.

    Not in our library10.1016/j.plaphy.2013.03.020

  142. 142.

    Martínez-García P.J., Crepeau M.W., Puiu D., Gonzalez-Ibeas D., Whalen J., Stevens K.A., Paul R., Butterfield T.S., Britton M.T., Reagan R.L., et al. The walnut (Juglans regia) genome sequence reveals diversity in genes coding for the biosynthesis of nonstructural polyphenols. Plant J. 2016;5:507-532.

    Not in our library10.1111/tpj.13207

  143. 143.

    Muir R.M., Ibáñez A.M., Uratsu S.L., Ingham E.S., Leslie C.A., McGranahan G.H., Batra N., Goyal S., Joseph J., Jemmis E.D., et al. Mechanism of gallic acid biosynthesis in bacteria (Escherichia coli) and walnut (Juglans regia). Plant Mol. Biol. 2011;75:555-565.

    Not in our library10.1007/s11103-011-9739-3

  144. 144.

    Cook M.T., Bassett H.P., Thompson F., Taubenhaus J.J. Protective enzymes. Science. 1911;33:624-629.

    Not in our library10.1126/science.33.851.624

  145. 145.

    Khodadadi F., Tohidfar M., Mohayeji M., Dandekar A.M., Leslie C.A., Kluepfel D.A., Butterfield T., Vahdati K. Induction of polyphenol oxidase in walnut and its relationship to the pathogenic response to bacterial blight. J. Am. Soc. Hortic. Sci. 2016;141:119-124.

    Not in our library

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