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Are Flavonoids Effective Antioxidants in Plants? Twenty Years of Our Investigation

Giovanni Agati, Cecilia Brunetti, Alessio Fini and 5 more

Antioxidants · 2020 · Review · CC BY

doi:10.3390/antiox9111098

Abstract

Whether flavonoids play significant antioxidant roles in plants challenged by photooxidative stress of different origin has been largely debated over the last few decades. A critical review of the pertinent literature and our experimentation as well, based on a free-of-scale approach, support an important antioxidant function served by flavonoids in plants exposed to a wide range of environmental stressors, the significance of which increases with the severity of stress. On the other side, some questions need conclusive answers when the putative antioxidant functions of plant flavonoids are examined at the level of both the whole-cell and cellular organelles. This partly depends upon a conclusive, robust, and unbiased definition of “a plant antioxidant”, which is still missing, and the need of considering the subcellular re-organization that occurs in plant cells in response to severe stress conditions. This likely makes our deterministic-based approach unsuitable to unveil the relevance of flavonoids as antioxidants in extremely complex biological systems, such as a plant cell exposed to an ever-changing stressful environment. This still poses open questions about how to measure the occurred antioxidant action of flavonoids. Our reasoning also evidences the need of contemporarily evaluating the changes in key primary and secondary components of the antioxidant defense network imposed by stress events of increasing severity to properly estimate the relevance of the antioxidant functions of flavonoids in an in planta situation. In turn, this calls for an in-depth analysis of the sub-cellular distribution of primary and secondary antioxidants to solve this still intricate matter.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 22, 2026. Paragraph numbers are ours, so an answer can point at one.

1. Introduction

It has been known for decades that UV, particularly UV-B radiation, triggers the biosynthesis of flavonoids (for recent review articles see), the vast class of phenylpropanoids comprising more than 8000 structures with a huge array of decorations. The flavonoid metabolic pathway was raised during the water-to-land transition of plants, when plants were challenged against an abrupt increase in UV-B radiation. This led to the hypothesis that flavonoids have an almost exclusive UV-B screening role in photoprotection. It was suggested that flavonoids have the peculiar capacity to absorb strongly over the UV-B portion of the solar spectrum, while offering negligible shielding to visible light radiation, thereby protecting DNA and PSII from damage. Nonetheless, the very same UV-B absorbing features belong to the vast majority of phenylpropanoids; a wide range of polyphenols display even greater capacity to shield sensitive leaf targets from high UV-B irradiance compared to flavonoids. It has been inferred, therefore, that unveiling how and how much flavonoids serve photoprotective functions in the network of UV-screening phenylpropanoids synthesized by current-day land plants needs deeper analysis.

The first time we encountered flavonoids was when examining leaves of Phillyrea latifolia adapted to deep shade (5% of full solar irradiance) or full sunlight. We observed that glandular trichomes, distributed on both the adaxial and the abaxial leaf surface in fully developed leaves, exclusively synthesized hydroxycinnamic acid (HCA) or flavonoid derivatives in the shade or in full sunlight, respectively. The almost exclusive distribution of flavonoids in glandular trichomes of leaves growing in full sunlight was confirmed later using multispectral fluorescence microimaging (Figure 1). These observations led to the hypothesis that flavonoids have functional roles beyond the absorption of UV, particularly of UV-B radiation in photoprotection. In fact, flavonoids (particularly the dihydroxy B-ring-substituted structure detected in our studies) are less effective than HCA in absorbing UV-B wavelengths. In a series of successive experiments, we also observed a steep increase in the ratio of dihydroxy to monohydroxy B-ring-substituted flavonoids in response to high light irradiance, in the presence or in the absence of UV radiation. Based on the relative light screening and antioxidant properties of mono- and dihydroxy B-ring-substituted flavonoids (reviewed in), we hypothesized that flavonoids might serve major antioxidant functions in photoprotection. We conducted studies, at very different scale levels, to support this hypothesis over the last two decades. Here, we discuss the significance of flavonoids as antioxidants in plants and their relative contribution to the integrated and modular network of antioxidant defenses plants activate when exposed to photooxidative stress of increasing severity. The matter is still under hot debate, in part because an unequivocal definition of a plant antioxidant is still missing.

2. Major Antioxidant Functions of Flavonoids in Photoprotection: Free-of-Scale Evidence

When plants moved from water to colonize land they were challenged by a wide range of “novel” environmental pressures, including, but not limited to, an abrupt increase in UV-B radiation. It has been believed for decades that the rise of flavonoid metabolism aimed at equipping plants with a more effective UV-B screening shield as compared to algae. However, this does not seem to be exactly the case (for critical reviews, see). First, a range of algal species synthesizes a variety of mycosporine-like amino acids, which are more effective than flavonoids in absorbing UV-B wavelengths. Second, flavonoids have minimum molar extinction coefficients (ε) over the 290-315, UV-B portion of the solar spectrum. Third, in plant organs most exposed to UV-B radiation, the synthesis of hydroxycinnamic acid (HCA) derivative declines in favor of flavonoid biosynthesis, even though HCAs absorb UV-B more efficiently than flavonoids. Fourth, the biosynthesis of quercetin (or dihydroxy B-ring-substituted flavonoids) is strongly favored with respect to kaempferol (or monohydroxy B-ring-substituted flavonoids) biosynthesis following exposure to UV-B radiation, though quercetin and kaempferol have similar UV-B screening efficiency. Finally, a series of relatively recent experiments have shown UV-B radiation is not a prerequisite, and blue light is more effective than UV-B light in stimulating flavonoid biosynthesis. There is increasing evidence suggesting that during the water-to-land transition, water stress and excessive light including, but not limited to UV-B radiation, drove most molecular innovations enabling early plants to successfully colonize land.

Caldwell and colleagues questioned the functional significance of flavonoids as mere UV-B absorbing pigments in UV-B exposed plants, approx. 40 years ago: they indeed noted that UV-B radiation promotes the biosynthesis of metabolites maximally absorbing over the UV-A portion of the solar spectrum. There are very few plant species synthesize flavonoids acylated with a range of HCA moieties, which may offer an effective shield against the penetration of the entire spectrum of UV wavelengths in the leaf (Figure 2).

We note that the vast majority of early studies examining the photoprotective functions of flavonoids exposed plants to unrealistic UV-B doses for very short time periods (e.g., see). Consequently, flavonoids exclusively accumulated in leaf epidermal cells, further reinforcing the idea of their exclusive roles as UV-B screening pigments. We also observe that the concentration of UV-B absorbing compounds, as estimated through absorption at 300-330 nm of tissue extracts, was in some instances assumed to represent the flavonoid concentration. This is actually wrong, as different phenylpropanoid classes effectively absorb over this portion of the solar spectrum. In other instances, identification of flavonoids was erroneously assessed based on the absorption spectra of isolated compounds. Finally, while studies examining the UV-B responses of mutants lacking the biosynthesis of a range of flavonoids conclusively showed their essential roles against UV-B damage, they did not offer a conclusive explanation about how flavonoids afford photo-protection. It is logically hard to perform experiments with the aim of unveiling the molecular events that trigger the flavonoid biosynthesis and, at the same time of assessing their functional roles in plant-environment interactions: it is simply a matter of scale (scaling-up). Results arising from growth chamber experiments, in which model plant species are exposed to unrealistic light irradiance, should not be considered as informative to describe the complexity of plants growing in their natural habitats. High UV and visible light irradiance are just two of the range of concomitant environmental stressors plants face in the field. The picture is complicated further by the observation that flavonoids occur in different plant organs, tissues and subcellular compartments in plants growing in full sun-exposed leaves (Figure 3; for a review see). Exploring the functional roles of flavonoids in photoprotection is an ecophysiological, not only a “molecular biology” challenge. This, when coupled with our constitutive inability to leave well-traced paths, has made complex the study of the primary roles played by flavonoids in photoprotection.

Helen Stafford hypothesized that a nascent and, hence, inefficient metabolism unlikely produced flavonoid at concentrations sufficient to provide an effective UV-B screening, thereby suggesting that flavonoids have alternative roles (i.e., developmental regulators) in response to UV-B stress. A few years later, Landry et al. and Sheahan first hypothesized for flavonoids an antioxidant, perhaps primary, function in UV-B treated leaves. As reported above, this hypothesis has been corroborated later by the steep increase in the ratio of dihydroxy to monohydroxy B-ring-substituted flavonoid glycosides in response to UV-B in a range of species. The capacity to scavenge reactive oxygen species (ROS) of glycosylated flavonoids is mostly conferred by the catechol group in 3′-4′ position, since the most reactive and, hence, the most reducing OH-group in 3-position is preferentially glycosylated. In other words, quercetin, but not kaempferol derivatives, are effective antioxidants. Interestingly, an increase in dihydroxy to monohydroxy B-ring-substituted flavonoids has been also reported in response to visible light, drought and salinity stress (in the absence of UV radiation,). A preferential increase in the concentration of the antioxidant quercetin glycosides was also shown in response to low temperature or nutrient deficiency in high light growing plants. These observations are consistent with the notion of flavonoid biosynthesis being activated by over-reduction of photosynthetic electron transport chain (PETC), a common feature, together with ROS production plants face when exposed to different abiotic stressors. Notably, that application of far-red light (700–780 nm), which is known to induce the oxidation of the PETC, depresses both the biosynthesis of flavonoids and the ratio of quercetin to kaempferol. This supports the view the redox and/or ROS signals are key drivers for the biosynthesis of flavonols (and for the biosynthesis of other secondary metabolites as well). This conforms to the notion that the activity of R2R3MYB proteins, regulating key genes of flavonol biosynthesis, is also under strict redox control, even because of posttranslational modifications.

It has been reasonably inferred, therefore, that the biosynthesis of flavonoids is activated by alteration in the redox potential of the cell and, in turn, flavonoids may help to limit oxidative damage primarily acting as antioxidants and not as light screeners in photoprotection. These are likely ancestral functions of flavonoids, as early land plants underwent severe desiccation and large temperature fluctuations, in addition to high light intensity, when they moved from water. In other words, “an excess of UV-B radiation” was just one (possibly not the most detrimental) of the “novel” environmental pressures plants faced when moving on land. Early land plants had to live with and adapt to an ever-changing ROS environment from the very beginning. The evolution of flavonoid (and the phenylpropanoid) metabolism as well as of TFs regulating their biosynthesis represent key components of the suite of traits whose evolution has been responsible for the increased complexity of land plants, thereby allowing their radiation toward less favorable environments. For example, in a range of modern land plants, a suite of flavonoid and hydroxycinnamate structures accumulate in plants growing under partial shading, with the monohydroxy B-ring-substituted structures dominating the phenylpropanoid pool. These compounds are mostly distributed in the external leaf tissues, such as the epidermal cells and different types of trichomes (Figure 2), and may well represent a constitutively (i.e., in plants growing in partial shading) efficient shield against UV-B radiation. The observation that monohydroxy B-ring-substituted phenylpropanoids are almost unresponsive, whereas dihydroxy B-ring-substituted flavones and flavonols increase in concentration because of UV-B, UV-A as well as visible light irradiance, offers strong support to a major antioxidant role of flavonoids in photoprotection. We note indeed that photoprotection must be “measured” only when plants experience light irradiance largely exceeding that actually used for photosynthesis, i.e., under strong photoinhibitory conditions.

3. Do Flavonoids Play Relevant Antioxidant Functions in Plants? A Matter of Definition

The actual role of flavonoids as effective antioxidants in both plants and animals has been extensively debated over the last few decades. Though flavonoids have great potential to counter the detrimental effects of the massive generation of reactive oxygen species (ROS), authoritative criticisms have been raised in many instances. Barry Halliwell, who provided a series of definitions of “antioxidants”, has always been critical of the effective ability of flavonoids to “delay, prevent or remove oxidative damage to a target molecule”, a widely accepted definition of antioxidant given in Gutteridge and Halliwell. This is because the flavonoid structures with the greatest ability to reduce (i.e., flavonoid aglycones) the levels of oxidants are usually in very low concentrations in human tissues. This, in turn, implies flavonoids cannot play systematic antioxidant functions. Even more, Halliwell, and Gutteridge and Halliwell suggested that the beneficial effects of flavonoids on human health might be also due to their pro-oxidant actions, thereby activating a range of endogenous antioxidant defenses. The Gutteridge–Halliwell definition of antioxidant was shaped over time taking into account novel evidence from studies performed on humans, but has been largely used by plant biologists as well. Being wide in its nature, this definition is rather robust, since it applies to organisms in different kingdoms, and includes not only the substance ability to quench/scavenge oxidants, but also to limit the rate at which oxidants form, i.e., the substance ability to “avoid” oxidative stress. In other words, the way through which a substance “protects” a target molecule from oxidative damage does not matter.

However, this has sparked and continues to spark debate among plant biologists. For instance, the capacity of flavonoids, both the colorless (e.g., flavones and flavonols) and the colored structures (i.e., anthocyanins) to strongly absorb over the UV- and the visible portion of the solar spectrum may effectively protect so-called sensitive targets from uncontrolled ROS-generation, and from the consequential oxidative damage. Similar reasoning also applies to relevant products of the MEP pathway, such as the volatile isoprene and the non-volatile xanthophyll, zeaxanthin. Isoprene and zeaxanthin may both increase the rigidity of thylakoid membranes in high light-grown plants concomitantly exposed to heat and water stress (drought stress, sensu stricto), and preserve lipids membrane from peroxidation: this is an antioxidant function, sensu Gutteridge and Halliwell. Even though these “indirect” functions should be regarded as antioxidant, the vast majority of plant scientists commonly refer to more restrictive definitions of antioxidants. The very same reasoning also applies to the definition of “oxidative damage”, and Halliwell posed the “debatable” question of direct or indirect biomolecular damage caused by an attack of ROS upon the constituents of living organisms.

Definitions delineate the frame, within which we have to reason to assess the effective antioxidant functions of any substance in an in planta condition. Therefore, any definition must be unbiased and robust, as much as possible. Foyer and Noctor defined antioxidant as a compound that “outcompetes others in reacting with ROS to give a relatively stable oxidized product”. The authors specifically refer to the quenching/scavenging of ROS, and introduce the concept of effective antioxidant, partly using an older definition of antioxidant given in 1995 by Halliwell and Gutteridge: “any substance that, when present at low concentrations compared with those of an oxidizable substrate, significantly delays or prevents oxidation of that substrate”. Hernandez et al. reasoned on the relevance of flavonoids as an antioxidant in plants, and defined an antioxidant as “a molecule that donates electrons or hydrogen atoms, yields an antioxidant derived radical that is efficiently quenched by other electron or hydrogen sources to prevent cellular damage; and whose properties are spatially and temporally correlated with oxidative stress events”. This is a restrictive definition compared to those given by Gutteridge and Halliwell and Foyer and Noctor. Within the framework delineated by their definition, Hernandez et al. posed serious concerns on the relevance of flavonoids as antioxidants in an in planta situation, since flavonoid radicals have not been detected in the vast majority of studies. We note, however, that by just following the authors’ definition, antioxidant-derived radicals are to be efficiently reduced to prevent oxidative/cellular damage. It is conceivable, therefore, that the phenoxyl radicals, the products of flavonoid oxidation, are at the trace level in healthy cells and unlikely detectable even with the most sensitive equipment. In actuality, Hernandez et al. detected quinones (i.e., the phenoxyl radicals ECQ and EGCGQ) of both epicatechin (EC) and epigallo catechin gallate (EGCG) (the reduced flavonoid forms) in tea plants exposed to severe drought stress, when relative water content (RWC) dropped down below 60% and maximum PSII quantum yield for photochemistry, Fv/Fm, was as low as 0.5. It is likely this study was not the most appropriate to unveil the actual antioxidant functions of flavonoids in an in planta situation, since the concentration of ECQ was 25 times higher than that of EC, and EGCGQ and EGCG did not differ much in concentration. We reason that either (perhaps both) the reduction system of flavanols was severely compromised and/or, more probably, damage to the vacuolar membrane occurred, under the severe drought stress event plants were faced with. We therefore suggest that the detection of flavonoid radicals is not the best-suited estimate of their occurred antioxidant action.

The required spatio-temporal correlation between antioxidants and oxidative stress events proposed by Hernandez et al. also poses further questions. For instance, products synthesized through the plastidial MEP pathway, such as β-carotene and isoprene, have been reported as actual quenchers of singlet oxygen, based upon the generation of their oxidation products. Since the production of isoprene and its oxidation products (methyl vinyl ketone and metacrolein) decreases under prolonged stress conditions unlikely may accomplish increased antioxidant protection when oxidative stress becomes more severe. Similarly, the concentrations of a suite of β-carotene endo-peroxides and of the volatile β-cycocitral generated by singlet oxygen-induced oxidation of β-carotene, decline as light stress become particularly severe. Similar observations have been also reported in the case of well-recognized antioxidants, such as antioxidant enzymes and ascorbate. There is vast and compelling evidence indeed that the activity of antioxidant enzymes may steeply decline during stress progression, when their action should be at the maximum. This is the case of plants suffering from high temperature or drought stress when concomitantly exposed to high solar irradiance, as also recently shown for the large declines in antioxidant enzyme activities in Platanus × acerifolia leaves during the hottest hours of the day. There is consensus indeed, that “stress-sensitive” species or genotypes are unable to maintain an efficient system for ROS removal when long exposed to stress. This is because large restrictions in the plant’s ability to use radiant energy for carbon fixation, translates into severe excess light stress antioxidant enzymes are unable to counter efficiently.

There is also evidence that the level of ascorbate (ASC) declines markedly, as also observed for the activity of ascorbate peroxidase, in the chloroplast, while steeply increasing in the vacuole because of both high light and drought stress. This stress-induced subcellular ASC redistribution decreases the usually (under non-stress conditions) high reducing the capacity of the chloroplast stroma. This poses some concerns on the effective antioxidant role of ASC (based on the spatio-temporal relationship between oxidative stress event and proper antioxidant location,) when the excess of radiant energy becomes particularly severe. There is old evidence that ASC is a relatively poor substrate for vacuolar peroxidase aimed at reducing H2O2 likely entering the vacuole at considerable rates under the most severe stressful conditions. Notably, the vacuole is the preferential site of flavonoid accumulation, although in some species colorless flavonoids have been detected in the chloroplast and in the nucleus. The functional significance of chloroplast- and nuclear-located flavonoids in preserving cells from irreversible oxidative damage has been already explored, and it will be briefly discussed in the next section. In contrast to ASC, flavonoids are excellent substrates for vacuolar class III peroxidases (POX),. More than two decades ago, Yamasaki et al. proposed a model in which phenoxyl radicals generated by the action of POX are recycled back to their reduced forms by ascorbate, and by mono-dehydroascorbate reductase as well, which act as “secondary antioxidants”. At that time, the model assumed an exclusive distribution of flavonoids in the vacuole of epidermal cells, thus raising the question about how H2O2 generated in mesophyll cells might reach that compartment. There is now compelling evidence that H2O2 is transported at both intra- and inter-cellular level through the action of the very same H2O transporters: H2O2 and H2O have indeed very similar physico-chemical properties. There is evidence that aquaporins are involved indeed in oxidative stress responses.

In addition, we note that the epidermis is not the tissue with the highest flavonoid level in plants living under natural conditions. The concentration of epidermal flavonoids, usually estimated by fluorescence microscopy of Naturstoff reagent-stained tissues, largely exceeds that in mesophyll tissues, simply because of the small volume in which epidermal flavonoids are dissolved (Figure 3). The whole-leaf content of flavonoids may reach as much as 8-10 micromoles in plants long-exposed to full sunlight, but less than 1 micro mole may accumulate in the vacuoles of epidermal layers, because of their relatively low solubility in the aqueous cellular milieu. Flavonoids located in the vacuoles of mesophyll cells are therefore close enough to ROS generating organelles (Figure 3), and hence optimally suited to buffer large alterations from ROS homeostasis. Furthermore, the view that flavonoids must be located in the very same cell organelle of ROS generation is likely to be strongly re-thought: ROS are not only involved cell-to-cell, but also in leaf-to-leaf communications, as is the case of light-induced control of stomata aperture. There is also intriguing evidence that collapsed chloroplasts are included in the vacuole following both UV-B and high light stress (so-called chlorophagy), as usually occurs in response to broad oxidative stress conditions. We likely have to re-draw the picture representing the complexity of the dynamic cellular organization during the most severe stressful conditions, to delineate the actual scenario within which biological processes occur. Autophagy is functional in ROS detoxification indeed, and there is a possibility that flavonoids might be involved in this process.

Recent findings from Gloria Muday’s lab have offered conclusive support for an effective role of flavonoids as in vivo scavengers of H2O2 (for a review, see Chapman et al.). In both Arabidopsis and tomato, flavonols, particularly quercetin mostly occurring in the cytoplasm and in the nucleus, greatly reduced the levels of H2O2 (estimated by fluorescence microscopy), thereby antagonizing ABA-induced stomatal closure. Flavonols were also effective in preserving the integrity and the growth of pollen tubes upon heat stress in tomato, by decreasing ROS levels. Agati et al., 2007, also offered compelling evidence for the effective ability of quercetin (and luteolin) derivatives to quench singlet oxygen (1O2) generated by high PAR irradiance using cross-sections of P. latifolia leaves with largely different levels of flavonoids. It was shown that dihydroxy B-ring-substituted flavonoids accumulated in the chloroplast outer envelope membrane and, hence optimally suited to quench 1O2 (Figure 4).

We observe that albeit the formation of flavonoid oxidation products is conclusive proof of their occurred in vivo antioxidant actions, the very same strong evidence comes from their capacity to reduce the level of oxidative stress. We suggest that the reduction of oxidative stress may be well suited to estimate the antioxidant role of flavonoids in an in planta situation. As already noted the occurrence of flavonoid oxidation products may be transient and largely dependent upon the severity of stress indeed, and hence, unlikely suitable for systemic measurements. Instead, newly developed imaging techniques seem to be suitable for quantifying ROS, and hence oxidative stress, not only at the cell and organ, but also at the whole-plant level. We believe that a plant antioxidant is best defined in terms of its ability to “counter” oxidative stress and damage: this ability is suitably “quantified” by measurements of “oxidative stress reduction” more than by the formation of oxidation products.

4. Conclusions: Still Open Questions

In the previous section, we offered compelling evidence of in vivo ROS scavenging by “cytoplasm-located” flavonoids (Figure 4 A–D). Nonetheless, there are very few examples of this subcellular flavonoid distribution, which also includes nuclear flavonoids. The potential antioxidant effects of nuclear-located flavonoids are of the greatest significance, as they include not only ROS quenching, but also the ability of antioxidant flavonol glycosides to chelate transition metal ions, thereby limiting the generation of ROS: this has to be regarded as an antioxidant function sensu lato. Nonetheless, studies aimed at exploring the sub-cellular distribution of flavonoids have not increased much in number over the last few years, and future investigation aimed at extending our knowledge on the sub-cellular distribution of specific flavonoid classes may help to solve the matter.

That said, the discussion about the effective antioxidant role of flavonoids in plants has been (and is still) based upon their vacuolar distribution, as observed in the vast majority of plants. In turn, this imposes to reason about the contribution of the H2O2-scavenging by vacuolar located flavonoids in modulating stress-induced changes in whole-cell redox homeostasis. The very same reasoning applies to the actual contribution of the vacuole in the maintenance of cellular redox homeostasis under stressful conditions, e.g., during excessive light. There are suggestions, indeed, that H2O2 may enter the vacuole at low rates to have major significance in altering the whole-cell redox homeostasis. We note that the rates at which H2O2 may diffuse out of the chloroplast (or the peroxisome) when the activity and the concentration of primary antioxidants decline greatly in response to severe stress are practically impossible to quantify. Additionally, we observe that the concentration range within which H2O2 activates signaling pathways to enhance further stress resistance or instead irreversibly damages cell metabolism and functioning appears very narrow. We highlight methods used to quantify H2O2 are conducted at the level of the whole-organ and, hence, are unsuitable to detect the small changes in H2O2 concentration that may occur in a very short time level in different subcellular compartments, and are responsible for the subsequent cell fate. Our deterministic approach encounters, once again, insurmountable scale-level obstacles.

In our opinion, the conclusive question regards the role of flavonoids relative to that of the modular and integrated network of antioxidant defenses plants activate in response to stress conditions of increasing severity. Plants face photo-oxidative stress of increased severity not only on long- (days/weeks) but also on a short-term (hour) basis. For instance, solar irradiance is in a large excess of that used for photosynthesis during the central hours of the day (midday depression of photosynthesis), because of stomatal limitations and supra-optimal leaf temperatures. This, in turn, leads to reductions in both Rubisco activation, thereby imposing additional biochemical limitations to photosynthesis and PSII photochemical efficiency. As already noted, excess excitation energy-induced large alterations in redox homeostasis are the very conditions that depress the activity of primary antioxidants, while promoting the biosynthesis of antioxidant flavonoids. Unveiling the extent to which primary and secondary antioxidants vary, on a short-term basis, in response to abrupt changes in excess radiant energy may help to solve the question of the actual significance of flavonoids in the modular and integrated network of antioxidant defenses. The notion that large alterations in the cell redox state trigger the biosynthesis of flavonoids leads us to hypothesize that flavonoids may complement the functions of primary antioxidants when their ROS detoxifying capacity declines in plants suffering from severe photooxidative stress. The matter needs future investigation aimed at estimating the changes in the whole spectrum of primary and secondary antioxidant defenses in plants with constitutively different levels of antioxidant flavonoids (using both natural or induced variation), challenged against the excess of radiant energy of increasing severity induced by abiotic stressors of different origin.

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  25. 25.

    Delwiche C.F., Cooper E.D. The evolutionary origin of a terrestrial flora. Curr. Biol. 2005;25:R899-R910.

    Not in our library10.1016/j.cub.2015.08.029

  26. 26.

    Mittler R. ROS are good. Trends Plant Sci. 2017;22:11-19.

    Not in our library10.1016/j.tplants.2016.08.002

  27. 27.

    Horandl E., Hadacek F. Oxygen, life forms, and the evolution of sexes in multicellular eukaryotes. Heredity. 2020;125:1-14.

    Not in our library10.1038/s41437-020-0317-9

  28. 28.

    Tena G. From algae to land plants. Nat. Plants. 2020;6:594.

    Not in our library10.1038/s41477-020-0712-5

  29. 29.

    Stafford H.A. Flavonoid evolution: An enzymic approach. Plant Physiol. 1991;96:680-685.

    Not in our library10.1104/pp.96.3.680

  30. 30.

    Pollastri S., Tattini M. Flavonols, old compounds for old roles. Ann. Bot. 2011;108:1225-1233.

    Not in our library10.1093/aob/mcr234

  31. 31.

    Agati G., Azzarello E., Pollastri S., Tattini M. Flavonoids as antioxidants in plants: Location and functional significance. Plant Sci. 2012;196:67-76.

    Not in our library10.1016/j.plantsci.2012.07.014

  32. 32.

    Schwinn K.E., Davies K.M. Flavonoids. Annual Plant Reviews 14: Plant Pigments and Their Manipulation. 2004;92-149.

    Not in our library

  33. 33.

    Liu Y., Fang S., Yang W., Shang X., Fu X. Light quality affects flavonoid production and related gene expression in Cyclocarya paliurus. J. Photochem. Photobiol. B Biol. 2018;179:66-73.

    Not in our library10.1016/j.jphotobiol.2018.01.002

  34. 34.

    Rai N., Neugart S., Yan Y., Wang F., Siipola S.M., Lindfors A.V., Winkler J.B., Albert A., Brosché M., Lehto T., et al. How do cryptochromes and UVR8 interact in natural and simulated sunlight? J. Exp. Bot. 2019;70:4975-4990.

    Not in our library10.1093/jxb/erz236

  35. 35.

    Siipola S.M., Kotilainen T., Sipari N., Morales L.O., Lindfors A.V., Robson T.M., Aphalo P.J. Epidermal UV-A absorbance and wholeleaf flavonoid composition in pea respond more to solar blue light than to solar UV radiation. Plant Cell Environ. 2015;38:941-952.

    Not in our library10.1111/pce.12403

  36. 36.

    Charron A.J., Quatrano R.S. Between a rock and a dry place: The water-stressed moss. Mol. Plant. 2009;2:478-486.

    Not in our library10.1093/mp/ssp018

  37. 37.

    Stevenosn S.R., Kamisugi Y., Trinh C.H., Schmutz J., Jenkins J.W., Grimwood J., Muchero W., Tuskan G.A., Rensing S.A., Lang D., et al. Genetic analysis of Physcomitrella patens identifies ABSCISIC ACID NON-RESPONSIVE, a regulator of ABA responses unique to basal land plants and required for desiccation tolerance. Plant Cell. 2016;28:1310-1327.

    Not in our library10.1105/tpc.16.00091

  38. 38.

    Farrant J.M., Moore J.P. Programming desiccation-tolerance: From plants to seeds to resurrection plants. Curr. Opin. Plant Biol. 2011;14:340-345.

    Not in our library10.1016/j.pbi.2011.03.018

  39. 39.

    De Vries J., Archibald J.M. Plant evolution: Landmarks on the path to terrestrial life. New Phytol. 2018;217:1428-1434.

    Not in our library10.1111/nph.14975

  40. 40.
    Evo-physio: on stress responses and the earliest land plants · 2020
    No notice
  41. 41.

    Caldwell M.M., Robberecht R., Flint S.D. Internal filters: Prospects for UV-acclimation in higher plants. Physiol. Plant. 1983;58:445-450.

    Not in our library10.1111/j.1399-3054.1983.tb04206.x

  42. 42.

    Tattini M., Matteini P., Saracini E., Traversi M.L., Giordano C., Agati G. Morphology and biochemistry of non-glandular trichomes in Cistus salvifolius L. leaves growing in extreme habitats of the Mediterranean basin. Plant Biol. 2007;9:411-419.

    Not in our library10.1055/s-2006-924662

  43. 43.

    Teramura A.H., Sullivan J.H. Effects of UV-B radiation on photosynthesis and growth of terrestrial plants. Photosynth. Res. 1994;39:463-473.

    Not in our library10.1007/bf00014599

  44. 44.

    Li J., Ou-Lee T.M., Raba R., Amundson R.G., Last R.L. Arabidopsis flavonoid mutants are hypersensitive to UV-B irradiation. Plant Cell. 1993;5:171-179.

    Not in our library10.2307/3869583

  45. 45.

    Lois R. Accumulation of UV-absorbing flavonoids induced by UV-B radiation in Arabidopsis thaliana L. I. Mechanisms of UV-resistance in Arabidopsis. Planta. 1994;194:498-503.

    Not in our library10.1007/bf00714462

  46. 46.

    Strid Å., Chow W.S., Anderson J.M. UV-B damage and protection at the molecular level in plants. Photosynth. Res. 1994;39:475-489.

    Not in our library10.1007/bf00014600

  47. 47.

    Bucholz G., Ehmann B., Wellmann E. Ultraviolet light inhibition of phytochrome-induced flavonoid biosynthesis and DNA photolyase formation in mustard cotyledons (Sinapis alba L.). Plant Physiol. 1995;108:227-234.

    Not in our library10.1104/pp.108.1.227

  48. 48.

    Markham K.R., Ryan K.G., Bloor S.J., Mitchell K.A. An increase in luteolin: Apigenin ratio in Marchantia polymorpha on UV-B enhancement. Phytochemistry. 1998;48:791-794.

    Not in our library10.1016/s0031-9422(97)00875-3

  49. 49.

    Middleton E.M., Teramura A.H. The role of flavonol glycosides and carotenoids in protecting soybean from ultraviolet-B damage. Plant Physiol. 1993;103:741-752.

    Not in our library10.1104/pp.103.3.741

  50. 50.

    Nogues S., Baker N.R. Effects of drought on photosynthesis in Mediterranean plants grown under enhanced UV-B radiation. J. Exp. Bot. 2000;51:1309-1317.

    Not in our library10.1093/jexbot/51.348.1309

  51. 51.

    Bieza K., Lois R. An Arabidopsis mutant tolerant to lethal ultraviolet-B levels shows constitutively elevated accumulation of flavonoids and other phenolics. Plant Physiol. 2001;126:1105-1115.

    Not in our library10.1104/pp.126.3.1105

  52. 52.

    Lois R., Buchanan B.-B. Severe sensitivity to ultraviolet radiation in an Arabidopsis mutant deficient in flavonoid accumulation II. Mechanisms of UV-resistance in Arabidopsis. Planta. 1994;194:504-509.

    Not in our library10.1007/bf00714463

  53. 53.

    Booij-James I.S., Dube S.K., Jansen M.A.K., Edelman M., Mattoo A.K. Ultraviolet-B radiation impacts light-mediated turnover of the Photosystem II reaction center heterodimer in Arabidopsis mutants altered in phenolic metabolism. Plant Physiol. 2000;124:1275-1283.

    Not in our library10.1104/pp.124.3.1275

  54. 54.

    Landry L.G., Chapple C.C.S., Last R.L. Arabidopsis mutants lacking phenolic sunscreen exhibit enhanced ultraviolet-B-induced injury and oxidative damage. Plant Physiol. 1995;109:1159-1166.

    Not in our library10.1104/pp.109.4.1159

  55. 55.

    Olsson L.C., Veit M., Bornman J.F. Epidermal transmittance and phenolic composition of atrazine-tolerant and atrazine-sensitive cultivars of Brassica napus grown under enhanced UV-B radiation. Physiol. Plant. 1999;107:259-266.

    Not in our library10.1034/j.1399-3054.1999.100302.x

  56. 56.

    Hofmann R.W., Swinny E.E., Bloor S.J., Markham K.R., Ryan K.G., Campbell B.D., Jordan B.R., Fountain D.W. Responses of nine Trifolium repens L. populations to ultraviolet-B radiation: Differential flavonol glycoside accumulation and biomass production. Ann. Bot. 2000;86:527-537.

    Not in our library10.1006/anbo.2000.1216

  57. 57.

    Hofmann R.W., Campbell B.D., Fountain D.W., Jordan B.R., Greer D.H., Hunt D.Y., Hunt C.L. Multivariate analysis of intraspecific responses to UV-B radiation in white clover (Trifolium repens L.). Plant Cell Environ. 2001;24:917-927.

    Not in our library10.1046/j.1365-3040.2001.00749.x

  58. 58.

    Berli F.J., Moreno D., Piccoli P., Hespanhol-Viana L., Silva M.F., Bressan-Smith R., Cavagnaro J.B., Borrini R. Abscisic acid is involved in the response of grape (Vitis vinifera L.) cv. malbec leaf tissues to ultraviolet-B radiation by enhancing ultraviolet-absorbing compounds, antioxidant enzymes and membrane sterols. Plant Cell Environ. 2010;33:1-10.

    Not in our library10.1111/j.1365-3040.2009.02044.x

  59. 59.

    Olsen K.M., Slimestad R., Lea U.S., Brede C., Løvdal T., Ruoff P., Verheul M., Lillo C. Temperature and nitrogen effects on regulators and products of the flavonoid pathway: Experimental and kinetic model studies. Plant Cell Environ. 2009;32:286-299.

    Not in our library10.1111/j.1365-3040.2008.01920.x

  60. 60.

    Lillo C., Lea U.S., Ruoff P. Nutrient depletion as a key factor for manipulating gene expression and product formation in different branches of the flavonoid pathway. Plant Cell Environ. 2008;31:587-601.

    Not in our library10.1111/j.1365-3040.2007.01748.x

  61. 61.

    Løvdal T., Olsen K.M., Slimestad R., Verheul M., Lillo C. Synergetic effects of nitrogen depletion, temperature, and light on the content of phenolic compounds and gene expression in leaves of tomato. Phytochemistry. 2010;71:605-613.

    Not in our library10.1016/j.phytochem.2009.12.014

  62. 62.

    Bathia C., Pandey A., Gaddam S.R., Hoecker U., Trivedi P.K. Low temperature-enhanced flavonol synthesis requires light-associated regulatory components in Arabidopsis thaliana. Plant Cell Physiol. 2018;59:2099-2112.

    Not in our library10.1093/pcp/pcy132

  63. 63.

    Akhtar T.A., Lees H.A., Lampi M.A., Enstone D., Brain R.A., Greenberg B.M. Photosynthetic redox imbalance influences flavonoid biosynthesis in Lemma gibba. Plant Cell Environ. 2010;33:1205-1219.

    Not in our library10.1111/j.1365-3040.2010.02140.x

  64. 64.

    Babu S., Akhtar T.A., Lampi M.A., Tripuranthakam S., Dixon G.R., Greenberg B.M. Similar stress responses are elicited by copper and ultraviolet radiation in the aquatic plant Lemma gibba: Implication of reactive oxygen species as common signals. Plant Cell Physiol. 2003;44:1320-1329.

    Not in our library10.1093/pcp/pcg160

  65. 65.

    Gerhardt K.E., Lampi M.A., Greenberg B.M. The effect of far-red light on plant growth and flavonoid accumulation in Brassica napus in the presence of ultraviolet B radiation. Photochem. Photobiol. 2008;84:1445-1454.

    Not in our library10.1111/j.1751-1097.2008.00362.x

  66. 66.

    Fanciullino A.L., Bidel L.P.R., Urban L. Carotenoid responses to environmental stimuli: Integrating redox and carbon controls into a fruit model. Plant Cell Environ. 2013;37:273-289.

    Not in our library10.1111/pce.12153

  67. 67.

    Selmar D., Kleinweichter M. Stress enhances the synthesis of secondary plant products: The impact of stress-related over-reduction on the accumulation of natural products. Plant Cell Physiol. 2013;54:817-826.

    Not in our library10.1093/pcp/pct054

  68. 68.

    Galant A., Koester R.P., Ainsworth E.A., Hicks L.M., Jez J.M. From climate change to molecular response: Redox proteomics of ozone-induced responses in soybean. New Phytol. 2012;194:220-229.

    Not in our library10.1111/j.1469-8137.2011.04037.x

  69. 69.

    Viola A.L., Camoirano A., Gonzalez D.H. Redox-dependent modulation of anthocyanin biosynthesis by the TCP transcription factor TCP15 during exposure to high light intensity conditions in Arabidopsis. Plant Physiol. 2016;170:74-85.

    Not in our library10.1104/pp.15.01016

  70. 70.

    Guehmann S., Vorbrueggen G., Kalkbrenner F., Moelling K. Reduction of a conserved Cys is essential for Myb DNA-binding. Nucleic Acids Res. 1992;20:2279-2286.

    Not in our library10.1093/nar/20.9.2279

  71. 71.

    Heine G.F., Hernandez J.M., Grotewold E. Two cysteines in plant R2R3MYB domains participate in REDOX-dependent DNA binding. J. Biol. Chem. 2004;279:37878-37885.

    Not in our library10.1074/jbc.m405166200

  72. 72.

    Taylor L.P., Grotewold E. Flavonoids as developmental regulators. Curr. Opin. Plant Biol. 2005;8:317-323.

    Not in our library10.1016/j.pbi.2005.03.005

  73. 73.

    Serpa V., Vernal J., Lamattina L., Grotewold E. Inhibition of AtMYB2 DNA-binding by nitric oxide involves cysteine S-nitrosylation. Biochem. Biophys. Res. Commun. 2007;361:1048-1053.

    Not in our library10.1016/j.bbrc.2007.07.133

  74. 74.

    Page M., Sultana N., Paszkievicz K., Florance H., Smirnoff N. The influence of ascorbate on anthocyanin accumulation during high light acclimation in Arabidopsis thaliana: Further evidence for redox control of anthocyanin synthesis. Plant Cell Environ. 2012;35:388-404.

    Not in our library10.1111/j.1365-3040.2011.02369.x

  75. 75.
    Genomes of early-diverging streptophyte algae shed light on plant terrestrialization · 2019
    No notice
  76. 76.

    Pierangelini M., Glaser K., Mikhailyuk T., Karsten U., Holzinger A. Light and dehydration but not temperature drive photosynthetic adaptations of basal streptophytes (Hormidiella, Streptosarcina and Streptofilum) living in terrestrial habitats. Microb. Ecol. 2019;77:380-393.

    Not in our library10.1007/s00248-018-1225-x

  77. 77.

    Jiao C., Sorensen I., Sun X., Sun H., Behar H., Alseekh S., Philippe G., Lopez K.P., Sun L., Reed R., et al. The Penium margaritaceum genome: Hallmarks of the origins of land plants. Cell. 2020;181:1097-1111.

    Not in our library10.1016/j.cell.2020.04.019

  78. 78.

    Reuber S., Bornman J.F., Weissenbock G. A flavonoid mutant of barley (Hordeum vulgare L.) exhibits increased sensitivity to UV-B radiation in the primary leaf. Plant Cell Environ. 1996;19:593-601.

    Not in our library10.1111/j.1365-3040.1996.tb00393.x

  79. 79.

    Burchard P., Bilger W., Weissenböck G. Contribution of hydroxycinnamates and flavonoids to epidermal shielding of UV-A and UV-B radiation in developing rye primary leaves as assessed by ultraviolet-induced chlorophyll fluorescence measurements. Plant Cell Environ. 2000;23:1373-1380.

    Not in our library10.1046/j.1365-3040.2000.00633.x

  80. 80.

    Landi M., Agati G., Fini A., Guidi L., Sebastiani F., Tattini M. Unveiling the shade nature of cyanic leaves: A view from the “blue absorbing side” of anthocyanins. Plant Cell Environ. 2020.

    Not in our library10.1111/pce/13818

  81. 81.

    Swain T., Cody V., Middleton E., Harborne J.B. Plant flavonoids in biology and medicine. Progress in Clinical and Biological Research. 1986;Volume 213:1-14.

    Not in our library

  82. 82.

    Halliwell B. Are polyphenols antioxidants or pro-oxidants? What do we learn from cell culture and in vivo studies? Arch. Biochem. Biophys. 2008;476:107-112.

    Not in our library10.1016/j.abb.2008.01.028

  83. 83.

    Williams R.J., Spencer J.P.E., Rice-Evans C. Flavonoids: Antioxidants or signalling molecules? Free Radic. Biol. Med. 2004;36:838-849.

    Not in our library10.1016/j.freeradbiomed.2004.01.001

  84. 84.

    Virgili F., Marino M. Regulation of cellular signaling from nutritional molecules: A specific roles of phytochemical beyond antioxidant activity. Free Radic. Biol. Med. 2008;45:1205-1216.

    Not in our library10.1016/j.freeradbiomed.2008.08.001

  85. 85.

    Halliwell B. The wanderings of a free radical. Free Radic. Biol. Med. 2009;46:531-542.

    Not in our library10.1016/j.freeradbiomed.2008.11.008

  86. 86.
    Flavonoids as Antioxidants and Developmental Regulators: Relative Significance in Plants and Humans · 2013
    No notice
  87. 87.

    Gutteridge J.M.C., Halliwell B. Antioxidants: Molecules, medicines, and myths. Biochem. Biophys. Res. Commun. 2010;393:561-564.

    Not in our library10.1016/j.bbrc.2010.02.071

  88. 88.

    Lesne A. Robustness: Confronting lessons from physics and biology. Biol. Rev. 2008;83:509-532.

    Not in our library10.1111/j.1469-185x.2008.00052.x

  89. 89.

    Landi M., Tattini M., Gould K.S. Multiple functional roles of anthocyanins in plant-environment interactions. Environ. Exp. Bot. 2015;119:4-17.

    Not in our library10.1016/j.envexpbot.2015.05.012

  90. 90.

    Havaux M., Dall’Osto L., Bassi R. Zeaxanthin has enhanced antioxidant capacity with respect to all other xanthophylls in Arabidopsis leaves and functions independent of binding to PSII antennae. Plant Physiol. 2007;145:1506-1520.

    Not in our library10.1104/pp.107.108480

  91. 91.

    Vickers C., Gershenzon J., Lerdau M.T., Loreto F. A unified mechanism of action for volatile isoprenoids in plant abiotic stress. Nat. Chem. Biol. 2009;5:283-291.

    Not in our library10.1038/nchembio.158

  92. 92.

    Beckett M., Loreto F., Velikova V., Brunetti C., Di Ferdinando M., Tattini M., Calfapietra C., Farrant J.M. Photosynthetic limitations and volatile and non-volatile isoprenoids in the poikilochlorophyllous resurrection plant Xerophyta humilis during dehydration and rehydration. Plant Cell Environ. 2012;35:2061-2074.

    Not in our library10.1111/j.1365-3040.2012.02536.x

  93. 93.

    Tattini M., Loreto F., Fini A., Guidi L., Brunetti C., Velikova V., Gori A., Ferrini F. Isoprenoids and phenylpropanoids are part of the antioxidant defense orchestrated daily by drought-stressed Platanus × acerifolia plants during Mediterranean summers. New Phytol. 2015;207:613-626.

    Not in our library10.1111/nph.13380

  94. 94.

    Halliwell B. Biochemistry of oxidative stress. Biochem. Soc. Tans. 2007;35:1147-1150.

    Not in our library10.1042/bst0351147

  95. 95.

    Halliwell B., Whiteman M. Measuring reactive species and oxidative damage in vivo and in cell culture: How should you do it and what do the results mean? Br. J. Pharmacol. 2004;142:231-255.

    Not in our library10.1038/sj.bjp.0705776

  96. 96.

    Gutteridge J.M.C., Halliwell B. Mini-Review: Oxidative stress, redox stress or redox success? Biochem. Biophys. Res. Commun. 2018;502:183-186.

    Not in our library10.1016/j.bbrc.2018.05.045

  97. 97.

    Halliwell B., Aeschbach R., Loliger J., Auroma O.I. The characterization of antioxidants. Food. Chem. Toxicol. 1995;33:601-617.

    Not in our library10.1016/0278-6915(95)00024-v

  98. 98.

    Hernandez I., Alegre L., Munné-Bosch S. Enhanced oxidation of flavan-3-ols and proanthocyanidin accumulation in water-stressed tea plants. Phytochemistry. 2006;67:1120-1126.

    Not in our library10.1016/j.phytochem.2006.04.002

  99. 99.

    Haffek H.P., Yakir D. Protection of isoprene against singlet oxygen in leaves. Plant Physiol. 2002;129:269-277.

    Not in our library10.1104/pp.010909

  100. 100.

    Zeinali N., Altarawneh M., Li D., Nu’rait A., Dlugogorski B.Z. New mechanistic insights: Why do plants produce isoprene? ACS Omega. 2016;1:220-225.

    Not in our library10.1021/acsomega.6b00025

  101. 101.

    Ramel F., Birtic S., Cuiné S., Triantaphylidès C., Ravanat J.-L., Havaux M. Chemical quenching of singlet oxygen by carotenoids in plants. Plant Physiol. 2012;158:1267-1278.

    Not in our library10.1104/pp.111.182394

  102. 102.

    Ramel F., Birtic S., Ginies C., Soubigou-Taconnat L., Triantaphylidès C., Havaux M. Carotenoid oxidation products are stress signals that mediate gene responses to singlet oxygen in plants. Proc. Natl. Acad. Sci. USA. 2012;109:5535-5540.

    Not in our library10.1073/pnas.1115982109

  103. 103.

    Tattini M., Velikova V., Vickers C., Brunetti C., Di Ferdinando M., Trivellini A., Fineschi S., Agati G., Ferrini F., Loreto F. Isoprene production in transgenic tobacco alters isoprenoid, non-structural carbohydrate and phenylpropanoid metabolism, and protects photosynthesis from drought stress. Plant Cell Environ. 2014;37:1950-1964.

    Not in our library10.1111/pce.12350

  104. 104.

    D’Alessandro S., Ksas B., Havaux M. Decoding β-cyclocitral-mediated retrograde signaling reveals the role of a detoxification response in plant tolerance to photooxidative stress. Plant Cell. 2018;30:2495-2511.

    Not in our library10.1105/tpc.18.00578

  105. 105.

    D’Alessandro S., Havaux M. Sensing β-carotene oxidation in photosystem II to master plant stress tolerance. New Phytol. 2019;223:1776-1783.

    Not in our library10.1111/nph.15924

  106. 106.

    Saunier A., Ormeno E., Worthman H., Temime-Roussel B., Lecareux C., Boissard C., Fernandez C. Chronic drought decreases anabolic and catabolic BVOC emissions of Quercus pubescens in a Mediterranean forest. Front. Plant Sci. 2017;8:71.

    Not in our library10.3389/fpls.2017.00071

  107. 107.

    Fini A., Brunetti C., Loreto F., Centritto M., Ferrini F., Tattini M. Isoprene responses and functions in plants challenged by environmental pressures associated to climate change. Front. Plant Sci. 2017;8:1281.

    Not in our library10.3389/fpls.2017.01281

  108. 108.

    Cappellin L., Loreto F., Biasioli F., Pastore P., McKinney K. A mechanism for biogenic production and emission of MEK from MVK decoupled from isoprene biosynthesis. Atmos. Chem. Phys. 2019;19:3125-3135.

    Not in our library10.5194/acp-19-3125-2019

  109. 109.

    Yanez-Serrano A.M., Mahlau L., Fasbender L., Byron J., Williams J., Kreuzwieser J., Wierner C. Heat stress causes enhanced use of cytosolic pyruvate for isoprene biosynthesis. J. Exp. Bot. 2019;70:5827-5838.

    Not in our library10.1093/jxb/erz353

  110. 110.

    Monson R.K., Winkler B., Rosenstiel T.N., Block K., Merl-Pham J., Strauss S.H., Ault K., Maxfield J., Moore D.J.P., Trahan N.A., et al. High productivity in hybrid-poplar plantations without isoprene emission to the atmosphere. Proc. Natl. Acad. Sci. USA. 2020;117:1596-1605.

    Not in our library10.1073/pnas.1912327117

  111. 111.

    Sharma P., Dubey R.S. Modulation of nitrate reductase activity in rice seedlings under aluminium toxicity and water stress: Role of osmolytes as enzyme protectants. J. Plant Physiol. 2005;16:854-864.

    Not in our library10.1016/j.jplph.2004.09.011

  112. 112.

    Liu J., Wang C., Wang Z., Zhang C., Lu S., Liu J. The antioxidant and free radical scavenging activities of extract and fractions from corn silk (Zea mays L.) and related flavone glycosides. Food Chem. 2011;126:261-269.

    Not in our library10.1016/j.foodchem.2010.11.014

  113. 113.

    Fini A., Brunetti C., Di Ferdinando M., Ferrini F., Tattini M. Stress-induced flavonoid biosynthesis and the antioxidant machinery of plants. Plant Signal. Behav. 2011;6:709-711.

    Not in our library10.4161/psb.6.5.15069

  114. 114.

    Fini A., Guidi L., Ferrini F., Brunetti C., Di Ferdinando M., Biricolti S., Pollastri S., Calamai L., Tattini M. Drought stress has contrasting effects on antioxidant enzymes activity and phenylpropanoid biosynthesis in Fraxinus ornus leaves: An excess light stress affair? J. Plant Physiol. 2012;169:929-939.

    Not in our library10.1016/j.jplph.2012.02.014

  115. 115.

    Liu Y.J., Zhang W., Wang Z.B., Ma L., Guo Y.P., Ren X.L., Mei L.X. Influence of shading on photosynthesis and antioxidative activities of enzymes in apple trees. Photosynthetica. 2019;57:857-865.

    Not in our library10.32615/ps.2019.081

  116. 116.

    Peltzer D., Polle A. Diurnal fluctuations of antioxidative systems in leaves of field-grown beech trees (Fagus sylvatica): Responses to light and temperature. Physiol. Plant. 2001;111:158-164.

    Not in our library10.1034/j.1399-3054.2001.1110205.x

  117. 117.

    Mubarakshina M.M., Ivanov B.N., Naydov I.A., Hillier W., Badger M.R., Krieger-Liszkay A. Production and diffusion of chloroplastic H2O2 and its implication to signaling. J. Exp. Bot. 2010;161:3577-3587.

    Not in our library10.1093/jxb/erq171

  118. 118.

    Yan N., Xu X.-F., Wang Z.-D., Huang J.-Z., Guo D.-P. Interactive effects of temperature and light intensity on photosynthesis and antioxidant enzyme activity in Zizania latifolia Turcz. plants. Photosynthetica. 2013;51:127-138.

    Not in our library10.1007/s11099-013-0009-2

  119. 119.

    Brunetti C., Guidi L., Sebastiani F., Tattini M. Isoprenoids and phenylpropanoids are key components of the antioxidant defense system of plants facing severe excess light stress. Environ. Exp. Bot. 2015;119:54-62.

    Not in our library10.1016/j.envexpbot.2015.04.007

  120. 120.

    Zechmann B., Stumpe M., Mauch F. Immunocytochemical determination of the subcellular distribution of ascorbate in plants. Planta. 2011;233:1-12.

    Not in our library10.1007/s00425-010-1275-x

  121. 121.

    Koffler B.E., Lushin-Ebengreuth N., Staibentheiner E., Muller M., Zechmann B. Compartment specific response of antioxidants to drought stress in Arabidopsis. Plant Sci. 2014;227:133-144.

    Not in our library10.1016/j.plantsci.2014.08.002

  122. 122.

    Takahama U. Oxidation of vacuolar and apoplastic phenolic substrates by peroxidase: Physiological significance of the oxidation reactions. Phytochem. Rev. 2004;3:207-219.

    Not in our library10.1023/b:phyt.0000047805.08470.e3

  123. 123.

    Sakihama Y., Mano J., Sano S., Asada K., Yamasaki H. Reduction of phenoxyl radicals mediated by mono dehydroascorbate reductase. Biochem. Biophys. Res. Commun. 2000;279:949-954.

    Not in our library10.1006/bbrc.2000.4053

  124. 124.

    Polle A. Dissecting the superoxide dismutase-ascorbate-glutathione-pathway in chloroplasts by metabolic modeling. Computer simulations as a step towards flux analysis. Plant Physiol. 2001;126:445-462.

    Not in our library10.1104/pp.126.1.445

  125. 125.

    Ferreres F., Figueiredo R., Bettencourt S., Carqueijeiro I., Oliveira J., GilIzquierdo A., Pereira D.M., Valentao P., Andrade P.B., Duarte P., et al. Identification of phenolic compounds in isolated vacuoles of the medicinal plant Catharanthus roseus and their interaction with vacuolar class III peroxidase: An H2O2 affair? J. Exp. Bot. 2011;62:2841-2854.

    Not in our library10.1093/jxb/erq458

  126. 126.

    Zipor G., Oren-Shamir M. Do vacuolar peroxidases act as plant caretakers? Plant Sci. 2013;199–200:41-47.

    Not in our library10.1016/j.plantsci.2012.09.018

  127. 127.

    Wang L.-L., Chen A.-P., Zhong N.-Q., Liu N., Wu X.-M., Wang F., Yang C.-L., Romero M.F., Xia G.-X. The Thellungiella salsuginea tonoplast aquaporin TsTIP1; 2 functions in protection against multiple abiotic stresses. Plant Cell Physiol. 2014;55:148-161.

    Not in our library10.1093/pcp/pct166

  128. 128.

    Yamasaki H., Sakihama Y., Ikehara N. Flavonoid-peroxidase reaction as a detoxification mechanism of plant cells against H2O2. Plant Physiol. 1997;115:1405-1412.

    Not in our library10.1104/pp.115.4.1405

  129. 129.

    Takahama U., Oniki T. A peroxidase/phenolics/ascorbate system can scavenge hydrogen peroxide in plant cells. Physiol. Plant. 1997;101:845-852.

    Not in our library10.1111/j.1399-3054.1997.tb01072.x

  130. 130.

    Stonebloom S., Brunkard J.O., Cheung A.C., Jiang K., Feldman L., Zambryski P. Redox states of plastids and mitochondria differentially regulate intercellular transport via plasmodesmata. Plant Physiol. 2012;158:190-199.

    Not in our library10.1104/pp.111.186130

  131. 131.

    Tian S., Wang X., Li P., Wang H., Ji H., Xie J., Qiu Q., Shen D., Dong H. Plant aquaporin AtPIP1;4 links apoplastic H2O2 induction to disease immunity pathways. Plant Physiol. 2016;171:1635-1650.

    Not in our library10.1104/pp.15.01237

  132. 132.

    Choi W.G., Miller G., Wallace I., Harper J., Mittler R., Gilroy S. Orchestrating rapid long-distance signaling in plants with Ca2+, ROS and electrical signals. Plant J. 2017;80:688-707.

    Not in our library10.1111/tpj.13492

  133. 133.

    Rodrigues O., Reshetnyak G., Grondin A., Saijo Y., Leonhardt N., Maurel C., Verdoucq L. Aquaporins facilitate hydrogen peroxide entry into guard cells to mediate ABA- and pathogen-triggered stomatal closure. Proc. Natl. Acad. Sci. USA. 2017;114:9200-9205.

    Not in our library10.1073/pnas.1704754114

  134. 134.

    Bienert G.B., Schjoerring J.K., Jahn T.P. Membrane transport of hydrogen peroxide. Biochim. Biophys. Acta. 2006;1758:994-1003.

    Not in our library10.1016/j.bbamem.2006.02.015

  135. 135.

    Bienert G.P., Chaumont F. Aquaporin-facilitated transmembrane diffusion of hydrogen peroxide. Biochim. Biophys. Acta. 2014;1840:1596-1604.

    Not in our library10.1016/j.bbagen.2013.09.017

  136. 136.

    Kim Y.X., Steudle E. Gating of aquaporins by light and reactive oxygen species in leaf parenchyma cells of the midrib of Zea mays. J. Exp. Bot. 2009;60:547-556.

    Not in our library10.1093/jxb/ern299

  137. 137.

    Wudick M.M., Li X., Valentini V., Geldner N., Chory J., Lin J., Maurel C., Luu D.-T. Subcellular redistribution of root aquaporins induced by hydrogen peroxide. Mol. Plant. 2015;8:1103-1114.

    Not in our library10.1016/j.molp.2015.02.017

  138. 138.

    Deng S.-P., Yang Y.-L., Cheng X.-X., Li W.-R., Cai J.-Y. Synthesis, spectroscopic study and radical scavenging activity of kaempferol derivatives: Enhanced water solubility and antioxidant activity. Int. J. Mol. Sci. 2019;20:975.

    Not in our library10.3390/ijms20040975

  139. 139.

    Zandalinas S.I., Mittler R. ROS-induced ROS release in plant and animal cells. Free Radic. Biol. Med. 2018;122:21-27.

    Not in our library10.1016/j.freeradbiomed.2017.11.028

  140. 140.

    Devireddy A.R., Zandalinas S.I., Gomez-Cadenas A., Blumwald E., Mittler R. Coordinating the overall stomatal response of plants: Rapid leaf-to-leaf communication during light stress. Sci. Signal. 2018;11:eaam9514.

    Not in our library10.1126/scisignal.aam9514

  141. 141.

    Fichman Y., Miller G., Mittler R. Whole-plant live imaging of reactive oxygen species. Mol. Plant. 2019;12:1203-1210.

    Not in our library10.1016/j.molp.2019.06.003

  142. 142.

    Izumi M., Ishida H., Nakamura S., Idema J. Entire photodamaged chloroplasts are transported to the central vacuole by autophagy. Plant Cell. 2017;29:377-394.

    Not in our library10.1105/tpc.16.00637

  143. 143.

    Nakamura S., Izumi M. Regulation of chlorophagy during photoinhibition and senescence: Lessons from mitophagy. Plant Cell Physiol. 2018;59:1135-1143.

    Not in our library10.1093/pcp/pcy096

  144. 144.

    Xiong Y., Contento A.L., Nguyen P.Q., Bassham D.C. Degradation of oxidized proteins by autophagy during oxidative stress in Arabidopsis. Plant Physiol. 2007;143:291-299.

    Not in our library10.1104/pp.106.092106

  145. 145.

    Yoshimoto K., Jikumaru Y., Kamiya Y., Kusano M., Consonni C., Panstruga R., Ohsumi Y., Shirasu K. Autophagy negatively regulates cell death by controlling NPR1-dependent salicylic acid signaling during senescence and the innate immune response in Arabidopsis. Plant Cell. 2009;21:2914-2927.

    Not in our library10.1105/tpc.109.068635

  146. 146.

    Pérez-Pérez M.E., Lemaire S.D., Crespo J.L. Control of autophagy in Chlamydomonas is mediated through redox dependent inactivation of the ATG4 protease. Plant Physiol. 2016;172:2219-2234.

    Not in our library10.1104/pp.16.01582

  147. 147.

    Signorelli S., Tarkowski L.P., vand den Ende W., Bassham D.C. Linking autophagy to abiotic and biotic stress responses. Trends Plant Sci. 2019;24:413-430.

    Not in our library10.1016/j.tplants.2019.02.001

  148. 148.

    Chapman J.M., Muhlemann J.K., Gayomba S.R., Muday G.K. RBOH-dependent ROS synthesis and ROS scavenging by plant specialized metabolites to modulate plant development and stress responses. Chem. Res. Toxicol. 2019;32:370-396.

    Not in our library10.1021/acs.chemrestox.9b00028

  149. 149.

    Watkins J.M., Hechler P.J., Muday G.K. Ethylene-induced flavonol accumulation in guard cells suppresses reactive oxygen species and moderates stomatal aperture. Plant Physiol. 2014;164:1707-1717.

    Not in our library10.1104/pp.113.233528

  150. 150.

    Watkins J.M., Chapman J.M., Muday G.K. Abscisic acid-induced reactive oxygen species are modulated by flavonols to control stomata aperture. Plant Physiol. 2017;175:1807-1825.

    Not in our library10.1104/pp.17.01010

  151. 151.

    Muhlemann J.K., Younts T.L.B., Muday G.K. Flavonols control pollen tube growth and integrity by regulating ROS homeostasis during high temperature stress. Proc. Natl. Acad. Sci. USA. 2018;115:E11188-E11197.

    Not in our library10.1073/pnas.1811492115

  152. 152.

    Agati G., Matteini P., Goti A., Tattini M. Chloroplast-located flavonoids can scavenge singlet oxygen. New Phytol. 2007;174:77-89.

    Not in our library10.1111/j.1469-8137.2007.01986.x

  153. 153.

    Choi W.G., Swanson S.J., Gilroy S. High-resolution imaging of Ca2+, redox status, ROS and pH using GFP biosensors. Plant J. 2012;70:118-128.

    Not in our library10.1111/j.1365-313x.2012.04917.x

  154. 154.

    Rehman A.U., Szabo M., Deak Z., Larkum A., Ralph P., Vaas P. Symbiodinium sp. cells produce light-induced intra- and extracellular singlet oxygen, which mediates photodamage of the photosynthetic apparatus and has the potential to interact with the animal host in coral symbiosis. New Phytol. 2016;212:472-484.

    Not in our library10.1111/nph.14056

  155. 155.

    Prasad A., Sedlarova M., Balukova A., Rac M., Pospisil P. Reactive oxygen species as a response to wounding: In vivo imaging in Arabidopsis thaliana. Front. Plant Sci. 2020;10:1660.

    Not in our library10.3389/fpls.2019.01660

  156. 156.

    Feucht W., Treutter D., Dithmar H., Polster J. Microspore development of three coniferous species: Affinity of nuclei for flavonoids. Tree Physiol. 2008;28:1783-1791.

    Not in our library10.1093/treephys/28.12.1783

  157. 157.

    Feucht W., Schmidt M., Treutter D. Flavanols and flavonols in the nuclei of conifer genotypes with different growth. Forests. 2014;5:2122-2135.

    Not in our library10.3390/f5092122

  158. 158.

    Choudhury F.K., Rivero R.M., Blumwald E., Mittler R. Reactive oxygen species, abiotic stress and stress combination. Plant J. 2017;90:856-867.

    Not in our library10.1111/tpj.13299

  159. 159.

    Smirnoff N., Arnaud D. Hydrogen peroxide metabolism and functions in plants. New Phytol. 2018;221:1197-1214.

    Not in our library10.1111/nph.15488

  160. 160.

    Borisova-Mubarakshina M.M., Kozuleva M.A., Rudenko N.N., Naydov I.A., Klenina I.B., Ivanov B.N. Photosynthetic electron flow to oxygen and diffusion of hydrogen peroxide through the chloroplast envelope via aquaporins. Biochim. Biophys. Acta. 2012;1817:1314-1321.

    Not in our library10.1016/j.bbabio.2012.02.036

  161. 161.

    Sewelam N., Jaspert N., Van Der Kelen K., Tognetti V.B., Schmitz J., Frerigmann H., Stahl E., Zeier J., Van Breusegem F., Maurino V.G. Spatial H2O2 signaling specificity: H2O2 from chloroplasts and peroxisomes modulates the plant transcriptome differentially. Mol. Plant. 2014;7:1191-1210.

    Not in our library10.1093/mp/ssu070

  162. 162.

    Cheeseman J.M. Hydrogen peroxide and plant stress: A challenging relationship. Plant Stress. 2007;1:4-15.

    Not in our library

  163. 163.

    Farooq M., Rehman A., Wahid A., Siddique K.H.M., Pessarakli M. Photosynthesis under heat stress. Handbook of Photosynthesis. 2016;697-701.

    Not in our library

  164. 164.

    Feller U., Crafts-Brandner S.J., Salvucci M.E. Moderately high temperatures inhibit ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) activase-mediated activation of Rubisco. Plant Physiol. 1998;116:539-546.

    Not in our library10.1104/pp.116.2.539

  165. 165.

    Salvucci M.E., Crafts-Brandner S.J. Mechanisms for deactivation of rubisco under moderate heat stress. Physiol. Plant. 2004;122:513-519.

    Not in our library10.1111/j.1399-3054.2004.00419.x

  166. 166.
    Rubisco and Rubisco Activase Play an Important Role in the Biochemical Limitations of Photosynthesis in Rice, Wheat, and Maize under High Temperature and Water Deficit · 2017
    No notice
  167. 167.

    Mullineaux P., Karpinski S. Signal transduction in response to excess light: Getting out of the chloroplast. Curr. Opin. Plant Biol. 2002;5:43-48.

    Not in our library10.1016/s1369-5266(01)00226-6

  168. 168.

    Hatier J.-H.B., Gould K.S. Foliar anthocyanins as modulators of stress signals. J. Theor. Biol. 2008;253:625-627.

    Not in our library10.1016/j.jtbi.2008.04.018

  169. 169.

    De la Haba P., De la Mata L., Molina E., Agüera E. High temperature promotes early senescence in primary leaves of sunflower (Helianthus annuus L.) plants. Can. J. Plant Sci. 2014;94:459-669.

    Not in our library10.4141/cjps2013-276

  170. 170.

    Li B., Fan R., Guo S., Wang P., Zhu X., Fan Y., Chen Y., He K., Kumar A., Shi J., et al. The Arabidopsis MYB transcription factor, MYB111 modulates salt responses by regulating flavonoid biosynthesis. Environ. Exp. Bot. 2019;166:103807.

    Not in our library10.1016/j.envexpbot.2019.103807

  171. 171.

    Gayomba S.R., Muday G.K. Flavonols regulate root hair development by modulating accumulation of reactive oxygen species in the root epidermis. Development. 2020;147:dev185189.

    Not in our library10.1242/dev.185819

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