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Reactive oxygen species: Reactions and detection from photosynthetic tissues
Reactive Oxygen Species: Reactions and Detection from Photosynthetic Tissues Heta Mattila, Sergey Khorobrykh, Vesa Havurinne, Esa Tyystj¨arvi PII: DOI: Reference:
S1011-1344(15)00322-X doi: 10.1016/j.jphotobiol.2015.10.001 JPB 10158
To appear in: Received date: Revised date: Accepted date:
11 June 2015 30 September 2015 1 October 2015
Please cite this article as: Heta Mattila, Sergey Khorobrykh, Vesa Havurinne, Esa Tyystj¨ arvi, Reactive Oxygen Species: Reactions and Detection from Photosynthetic Tissues, (2015), doi: 10.1016/j.jphotobiol.2015.10.001
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Heta Mattila, Sergey Khorobrykh, Vesa Havurinne and Esa Tyystjärvi*
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Reactive Oxygen Species: Reactions and Detection from Photosynthetic Tissues Department of Biochemistry / Molecular Plant Biology, University of Turku, 20014 Turku, Finland
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*Corresponding author, e-mail [email protected]
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Abstract
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Reactive oxygen species (ROS) have long been recognized as compounds with dual roles. They cause cellular damage by reacting with biomolecules but they also function as agents of cellular signaling. Several different oxygen-containing compounds are classified as ROS because they react, at least with certain partners, more rapidly than ground-state molecular oxygen or because they are known to have biological effects. The present review describes the typical reactions of the most important ROS. The reactions are the basis for both the detection methods and for prediction of reactions between ROS and biomolecules. Chemical and physical methods used for detection, visualization and quantification of ROS from plants, algae and cyanobacteria will be reviewed. The main focus will be on photosynthetic tissues, and limitations of the methods will be discussed.
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Contents 1. Reactive oxygen species (ROS) and free radicals ................................................................................ 5 1.1. Properties of ROS ......................................................................................................................... 5
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1.2. General principles of detection and monitoring of ROS .............................................................. 7
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2. Singlet oxygen, 1O2 .............................................................................................................................. 8 2.1. Definition and properties of 1O2 .................................................................................................. 8 2.2. Formation and action of 1O2 in photosynthetic tissues ............................................................... 8
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2.3. Reactions of 1O2 ........................................................................................................................... 9 2.3.1. Physical deactivation of 1O2 by radiative and non-radiative mechanisms............................ 9
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2.3.2. Chemical reactions of 1O2 ................................................................................................... 10 2.4. Lifetime and diffusion distance of 1O2 ....................................................................................... 12 2.5. Detection of 1O2 ......................................................................................................................... 17
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2.5.1. Luminescence at 1270 nm .................................................................................................. 17 2.5.2. EPR detectable probes ........................................................................................................ 17
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2.5.3. Dyes ..................................................................................................................................... 18 2.5.4. Fluorescent probes ............................................................................................................. 18 2.5.5. Oxygen consumption in a reaction between 1O2 and histidine .......................................... 19
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2.5.6. Genetically encoded probes ............................................................................................... 19 2.5.7. Further gene expression methods ...................................................................................... 23
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2.5.8. Indirect methods of 1O2 detection ...................................................................................... 23 2.5.9. Summary of 1O2 detection .................................................................................................. 24 3. Hydrogen peroxide, H2O2 .................................................................................................................. 24 3.1. Definition and properties of H2O2 .............................................................................................. 24 3.2. Formation and action of H2O2 in photosynthetic tissues........................................................... 24 3.3. Reactions of H2O2 ....................................................................................................................... 25 3.4. Lifetime and diffusion distance of H2O2 ..................................................................................... 26 3.5. Detection of H2O2 ....................................................................................................................... 26 3.5.1. Principles of H2O2 detection................................................................................................ 26 3.5.2. Precipitate-forming compounds ......................................................................................... 26 3.5.3. Dyes ..................................................................................................................................... 27 3.5.4. Fluorescent probes ............................................................................................................. 27 3.5.5. Chemiluminescent probe .................................................................................................... 28 3.5.6. Microelectrodes .................................................................................................................. 28 3.5.7. Genetically encoded probes ............................................................................................... 29
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3.5.8. Unspecific probes................................................................................................................ 29 3.5.9. Indirect methods of H2O2 detection ................................................................................... 29 3.5.10. Summary of H2O2 detection .............................................................................................. 29
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4. Superoxide anion radical, O2•, and hydroperoxyl radical, HO2• ...................................................... 33
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4.1. Definitions and properties of O2• and HO2• .............................................................................. 33 4.2. Formation and action of O2• and HO2• in photosynthetic tissues ............................................ 33 4.3. Reactions of O2• and HO2• ........................................................................................................ 33
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4.4. Lifetime and diffusion distance of O2• ...................................................................................... 36 4.5. Detection methods of O2• ......................................................................................................... 37
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4.5.1. Spectroscopic methods ....................................................................................................... 37 4.5.2. EPR detectable probes ........................................................................................................ 37
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4.5.3. Dyes ..................................................................................................................................... 38 4.5.4. Fluorescent probes ............................................................................................................. 40 4.5.5. Chemiluminescent probes .................................................................................................. 40
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4.5.6. Microelectrodes .................................................................................................................. 41 4.5.7. Genetically encoded probes ............................................................................................... 41
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4.5.8. Indirect methods of O2• measurement............................................................................. 41 4.5.9. Summary of O2• detection ................................................................................................. 41 5. Hydroxyl radical, HO• ........................................................................................................................ 46
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5.1. Definition and properties of HO• ............................................................................................... 46 5.2. Formation and action of HO• in photosynthetic tissues ............................................................ 46 5.3. Reactions of HO• ........................................................................................................................ 47 5.4. Lifetime and diffusion distance of HO• ...................................................................................... 48 5.5. Detection methods of HO• ......................................................................................................... 48 5.5.1 Spectroscopic methods ........................................................................................................ 48 5.5.2. EPR detectable probes ........................................................................................................ 49 5.5.3. Fluorescent probes ............................................................................................................. 50 5.5.4. Microelectrode.................................................................................................................... 51 5.5.5. Summary of HO• detection ................................................................................................. 51 6. Peroxyl radical (ROO•), alkoxyl radical (RO•) and hydroperoxides ................................................... 54 6.1. Definition and formation ........................................................................................................... 54 6.2. Reactions of ROO•, RO• and hydroperoxides ............................................................................. 54 6.3. Lifetime and diffusion distance of ROOHs ................................................................................. 56 6.4. Detection methods of ROOHs .................................................................................................... 56
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6.4.1. Spectroscopic methods ....................................................................................................... 56 6.4.2. Chemiluminescence and thermoluminescence .................................................................. 56 6.4.3. Dyes ..................................................................................................................................... 57
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6.4.4. Fluorescent probes ............................................................................................................. 57
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6.4.5. Measurements of ROOH-derived species ........................................................................... 58 7. Ozone, O3 .......................................................................................................................................... 61 8. Non-specific methods ....................................................................................................................... 63
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8.1. Non-specific methods indicating the presence of ROS .............................................................. 63 8.2. Non-specific methods indicating reactions of ROS with biomolecules ..................................... 64
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9. Concluding remarks .......................................................................................................................... 64 10. Acknowledgements......................................................................................................................... 65
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11. References ...................................................................................................................................... 65
1.1. Properties of ROS
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1. Reactive oxygen species (ROS) and free radicals
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The term "reactive oxygen species" refers to naturally occurring, oxygen-containing chemical species whose reactivity, at least towards some substances, is higher than that of ground-state oxygen [1]. In the present review, we will limit the discussion to singlet oxygen, superoxide (superoxide anion radical and hydroperoxyl radical), hydrogen peroxide, hydroxyl radical, ozone, and lipid peroxyl radicals, lipid peroxides and alcoxides. ROS are inevitably produced in photosynthetic organisms [2,3] and cause oxidative damage by reacting with biomolecules [1,4‒7], but ROS also function as cellular signals [8‒10]. Photosynthetic organisms have extensive quenching and scavenging mechanisms that alleviate the effects of ROS [11‒14]. Free radicals are defined as chemical species that contain one or more unpaired electrons and are capable of independent existence [1]. Ground-state O2 is classified as a free radical (a diradical) because the two degenerate outermost molecular orbitals are occupied by one electron each, and in accordance to Hund's rule, the spins of these two electrons are parallel in the lowest energy state of the system. Superoxide anion radical (O2•), hydroxyl radical (HO•) and perhydroxyl radical (HO2•) are also free radicals whereas singlet oxygen (1O2), hydrogen peroxide (H2O2), ozone (O3), peroxides and alcoxides are not radicals. Some free radicals (like HO• and the hydrogen radical H•) are extremely reactive, whereas some free radicals, like ions of transition metals, exhibit weak reactivity. Free radicals are affected by external magnetic fields because the net spin magnetic momentum of the unpaired electrons interacts with a magnetic field. The energy of a free radical depends on the spin configuration and on the strength of the magnetic field. Free radicals are therefore visible in electron paramagnetic resonance (EPR, also called electron spin resonance, ESR) spectroscopy. A radical with one unpaired electron (doublet, total spin 1/2) has two energy states (designated as the spin up and the spin down states) in a magnetic field, and a free radical with two unpaired electrons
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(triplet, total spin 1), has three energy levels. The total spin of a molecule containing only paired electrons is zero, and such a molecule is described as a singlet state. Singlets interact with external magnetic fields only weakly.
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In addition to the classification as radicals and non-radicals, ROS can be roughly divided into two types: (1) Free ROS, small molecules composed of oxygen and hydrogen only, and (2) incorporated ROS in which oxygen is bound to other molecules to form reactive oxygen derivatives. Table 1 presents the most important ROS. The most important reactive nitrogen species (for a review, see [15]) are listed in Table 1 but will not be further discussed.
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Table 1. Most significant ROS and reactive nitrogen species. R is a residual of an organic molecule. Non-radicals Free ROS
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Radicals
Singlet oxygen, 1O2
Superoxide anion radical, O2• Perhydroxyl radical, HO2•
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Hydroxyl radical, HO•
Hydrogen peroxide, H2O2
Ozone, O3 Incorporated ROS and reactive nitrogen species Organic peroxides, ROOH Peroxynitrite ion, ONOO Alkyl peroxynitrite, ROONO
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Peroxyl radical, ROO• Alkoxyl radical, RO• Nitric Oxide, NO• Nitrogen dioxide,NO2•
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The biological actions of ROS are selective because the reactivity of each ROS strongly depends on the ROS and on reaction conditions. Some ROS like HO• have a very high chemical activity and are able to oxidize most organic molecules. On the other hand, H2O2 has moderate reactivity and is able to migrate far from site of production. The general reactivity of the main ROS in biological environment decreases in the order HO• > 1O2 > H2O2 > O2•, although care should be taken in using this generalization, as the rate of reaction depends on the reaction partner. Ability to interconverse is an essential property of ROS. For example, 1-electron reduction of H2O2 leads to the formation of HO•, indicating that formation of a less reactive form can promote the formation of more reactive forms. Due to the interconversions, cellular effects of ROS should be considered as complex sets of reactions. Given these reactions, action mechanisms of oxygen in the cell can be understood. Production, detoxification and detection of ROS, as well as the roles of ROS as both agents of damage and as cellular signals are widely discussed (for reviews see e.g. [16‒18]). In the following, we will give an overview of the reactions of each ROS and describe methods that have been used to detect the ROS, focusing on photosynthetic organisms. The biological roles of the ROS in photosynthetic organisms will be discussed only to the extent needed for the understanding of the detection methods.
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1.2. General principles of detection and monitoring of ROS
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In pure solvents or in the gas state, concentrations of various ROS can be directly measured by absorbance, fluorescence or EPR spectroscopy. However, the most common methods for monitoring the relative amounts of ROS require the use of chemicals that preferentially react with a particular ROS, yielding a detectable product.
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Most ROS do not equilibrate throughout a complex biological material within their lifetime but either react near to the site of origin or remain confined to the producing compartment. Therefore the cellular localization of the detector substance is often of importance for understanding the results. Unfortunately, experimentally verified information about the localization of the various detector substances is seldom available, especially in plant material. In the present review, we list the available information. The logarithm of 1-octanol-water partition coefficient (LogP) is widely used to estimate hydrophobicity. As a rule, the lower the LogP value of a compound is, the more hydrophilic it is. All experimentally determined and some estimated LogP values in this review have been obtained from online chemical structure database, Chemspider [19]; for compounds not found in Chempsider, LogP values were estimated using an online tool ALOGPS 2.1 [20]. LogP values for different ROS detection compounds are presented in Tables 3‒7. Caution should be taken in interpreting LogP values as indicators of cellular localization or intake of a molecule, because cellular environment cannot be dichotomously divided into two simple immiscible phases. Furthermore, the plant and cyanobacterial cell wall may lower the cell permeability of lipophilic molecules.
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Hydrophilic chemicals can be fed to intact plant samples in the transpiration stream but lipophilic substances may require vacuum infiltration or scratching the plant surface. Side effects of these aggressive methods should be separately evaluated. Biotechnology offers the possibility to make a mutant organism that synthesizes a specific genetically encoded ROS probe in the desired tissue or cellular compartment [21]. The success of biotechnology is based on the availability of specific cellular ROS probes and so far only probes for H2O2, based on the specific sensitivity of transcription factors (OxyR from Escherichia coli and Orp1 from Saccharomyces cerevisiae) to H2O2, have been tested for the detection of endogenous ROS (see [16,22] for reviews). Promising results have been obtained also in the development of genetically encoded 1O2 probes (see Chapter 2.5.6). Because certain ROS function as cellular signals, specific changes in gene expression can often be correlated with specific ROS. However, conclusions about the causal agent causing a change in gene expression must be drawn with great care, unless the mechanism of regulation is well understood. Enzymes present in all cells can sometimes be used to modify the sensor substances, as in the design of fluorescein diacetate dyes from which cellular esterases remove the acetate moiety, producing a fluorescent, charged dye that cannot diffuse back to the extracellular space [23]. In the case of aminophenyl and hydroxyphenyl fluorescein dyes, the strength of the ROS signal may also depend on the action of a peroxidase [24]. Detection of ROS with chemical reactions is often difficult because several ROS may react with the detector chemical. Unspecific detectors can be used in the rare cases that the system is known to produce only one type of ROS and that only one type of ROS is of interest. However, measurement of the total amount of ROS in the sample cannot be done with an unspecific detector because the reaction rates of the detector with different ROS vary. If the concentration ranges of various ROS
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species in the sample have been well characterized in advance, it may also be possible to use several unspecific detector substances and utilize knowledge about their reaction rates with different ROS.
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In addition to the specificity and localization in the biological sample, ROS detector substances may have problems of biological incompatibility, stability in the light and stability of the reaction product. Some ROS detector substances are known to produce ROS. These topics will be discussed when information is available.
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2. Singlet oxygen, 1O2 2.1. Definition and properties of 1O2
A ground-state O2 molecule ( g O2) is a triplet form (also designated as 3O2) and can therefore 3
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accept two electrons from a species containing paired electrons only if a spin conversion occurs. Thus, the reactivity of O2 at room temperature is moderate because the molecule has two electrons with parallel spins. The singlet forms of O2 containing only paired electrons are much more reactive than ground-state O2. Molecular oxygen has two singlet forms. The two electrons with antiparallel spins may reside
either on two different orbitals ( g O2) or both on one orbital ( g O2). g O2 decays rapidly to 1
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form either ground-state O2 or g , and therefore the designation 1O2 will be used for g O2, 1
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unless otherwise indicated.
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O2 is a long-lived excited state, but its decay is greatly speeded up by collisions with almost any other molecule [25]. The radiative decay of 1O2 to ground-state O2 yields luminescence with maximum at 1268 nm in the gas phase [26]; in water and deuterium oxide (D2O), the peak is at 1274 nm [27]. Although 1O2 is a singlet state, it has an EPR spectrum [28]. The spectrum has been detected only in the gas phase [29]. Rapid non-radiative decay of 1O2 occurs via (i) conversion of excitation energy of 1O2 to vibrational and rotational energy of both the quencher (e.g. H2O) and O2; (ii) a charge-transfer mechanism between the quencher (e.g. phenol or diazabicyclo[2.2.2]octane, DABCO) and 1O2; or (iii) an electronic energy transfer mechanism in which the reaction between 1O2 and a singlet ground state of the quencher (e.g. a carotenoid) produces a triplet state of both the quencher and oxygen (i.e. ground-state oxygen) [25].
2.2. Formation and action of 1O2 in photosynthetic tissues In photosynthetic tissues, 1O2 is mainly formed by photosensitization (for a review see [30]). Encounter of O2 with either a singlet or a triplet excited state of a pigment molecule (sensitizer) may lead to generation of a singlet form of oxygen. A reaction between a singlet excited state of a sensitizer and O2 yields 1O2 and a triplet state of the sensitizer, whereas a reaction between O2 and a triplet state of the sensitizer produces 1O2 and a singlet ground state of the sensitizer [25]. The reactions between O2 and singlet and triplet excited states of sensitizers are also affected by the energy gap between the singlet and triplet states of the sensitizer [31]. In practice, reactions with singlet excited states are of low significance because singlet excited states are short-lived. Chlorophylls, bacteriochlorophylls, hemes and other tetrapyrroles, especially protoporphyrin IX (an intermediate of synthesis of heme and chlorophyll), as well as iron sulfur centers are known
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photosensitizers [4]. However, chlorophyll (Chl) a and b, and bacteriochlorophylls in non-oxygenic photosynthesis, are the most important natural photosensitizers of 1O2 formation in photosynthetic organisms. According to the available data, production of 1O2 in plants is almost always sensitized by the triplet state of the primary donor (3P680*) of Photosystem II (PSII), formed by charge recombination reactions (for a review see [3]). Thus, neither the triplet state of Chl a (3Chl a) produced by intersystem crossing in the antennae [32] nor the triplet state of the primary donor of Photosystem I (PSI) (3P700*) seem to sensitize 1O2 formation [33,34]. Weak triplet formation in the antennae due to photochemical and non-photochemical quenching of excitation energy, low oxygen concentration in the antenna, and quenching of the triplet states by carotenoids may contribute to the apparently negligible sensitization by many Chl triplets in plants. Experiments with the histidine method (see 2.5.5) show that at photosynthetic photon flux density (PPFD) of 2300 µmol m-2s-1 the 1 O2 quencher histidine lowers oxygen production of the cyanobacterium Synechocystis sp. PCC 6803 by 13 % [35]. This result may actually suggest massive production of 1O2 within the membranes, as most 1O2 is expected to react at the site of production [1].
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Other sources of 1O2 than PSII in plants are related to plant defense and e.g. heat stress (for reviews, see [3,36,37]). Non-photochemical formation of 1O2 with the Russell mechanism, via recombination of two peroxy radicals, has also been shown to occur in plant tissues [38,39] but the importance of this pathway has not been studied extensively.
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Detoxification of 1O2 in plant cells occurs by carotenoids and tocopherols in the thylakoid membranes (for reviews, see [5,14]) and in the cytosol by soluble scavengers including vitamins C and B6 and glutathione (GSH) [40,41]. 1
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O2 has been speculated to be the most important ROS formed in the light reactions of photosynthesis, as the triplet state of the primary donor of PSII is ubiquitously formed by charge recombination reactions. Work on the flu mutant of Arabidopsis thaliana that produces 1O2 when grown in a light-dark rhythm has revealed that the reason for cell death is not cellular damage mediated by 1O2 but 1O2-induced signaling events [42‒44]. However, the fact that programmed cell death depends on 1O2 signaling, does not rule out the possibility that 1O2 causes cellular damage.
2.3. Reactions of 1O2
2.3.1. Physical deactivation of 1O2 by radiative and non-radiative mechanisms Only radiative deactivation (reaction 1) is available for an isolated and unperturbed 1O2 molecule, and the radiative lifetime O2(1g) without any collisions is extremely long, 72 min [25]. 1
O2 3O2+ h
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Enhancement of deactivation of 1O2 by collisions of 1O2 with other molecules is one reason for variation of the lifetime of 1O2 in different media. Deactivation of 1O2 by collisions with other molecules having sufficiently high triplet-state energy can occur by conversion of excitation energy of 1O2 to vibrational energy of the other molecule. This non-radiative deactivation process, known as electronic-vibrational energy transfer, limits the lifetime of 1O2 in many solvents. The lifetime of 1O2 in H2O is 3.1 s, about 20 times less than in D2O (68 s) [25,45,46]. Effects of solvent on the lifetime of 1O2 can often provide evidence for participation of 1O2 in chemical reactions.
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In addition to the electronic-vibrational non-radiative deactivation, 1O2 can be deactivated via an electron exchange mechanism (reaction 2). This spin-allowed electronic energy transfer to a molecule with a low triplet state energy is a very efficient mechanism of 1O2 deactivation. For example, the rate of deactivation of 1O2 with carotenoids is limited by diffusion. The rate constant for β-carotene and lutein is about 1010 M-1 s-1 [47], O2 + A 3(O2 A) 3O2 + 3A,
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(2)
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where A is an acceptor molecule, for example a carotenoid. Reaction (2), also known as physical quenching of 1O2, is the main mechanism of quenching of 1O2 by carotenoids. Bimolecular rate constants of reactions between carotenoids and 1O2 can be found in [48].
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In addition to their ability to quench 1O2, carotenoids protect the photosynthetic machinery also by quenching both the triplet and singlet excited states of Chls, thereby lowering the probability of 1O2 formation. In plant photosynthesis, the xanthophyll violaxanthin is de-epoxidated in the light to anteraxanthin and zeaxanthin which participate in so called non-photochemical quenching of excitation energy, a mechanism converting the energy of the singlet excited state of Chl a to heat. For review of the photoprotective roles of carotenoids in photosynthesis, see [49].
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2.3.2. Chemical reactions of 1O2 1 O2 is an electrophilic agent and reacts preferably with electron rich organic molecules by addition to double bonds. This chemical deactivation of 1O2 proceeds via well-known mechanisms:
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ene reaction associated with formation of a hydroperoxide (3);
(3)
cycloaddition associated with dioxetane formation (4); (4)
cycloaddition reaction with aromatic compounds and formation of endoperoxides via the Diels-Alder mechanism (5); (5)
formation of charge-transfer intermediates. 1O2 can react with compounds containing heteroatom such as N or S to form charge-transfer intermediates, reactions (6) and (7) respectively. Chargetransfer intermediates are particularly important for the interaction of 1O2 with phenols and tocopherols [50]. O2 + NR3 [1O2--- +NR3]
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(6)
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O2 + SR2 [1O2--- +SR2]
1
(7)
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The charge-transfer complex can be disintegrated either by an electron exchange mechanism (physical quenching of 1O2) or by oxidation of a heteroatom, reaction (8).
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(8)
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In addition to oxidation of a heteroatom, the reaction with 1O2 can lead to carbon-heteroatom bond cleavage and formation of a corresponding aldehyde [51]. 1O2 can efficiently oxidize amines to imines with formation of HO2•, reaction (9) [52].
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The interaction of 1O2 with the heteroatom of 2,2,6,6-tetramethly-4-piperidone (TEMP) produces 2,2,6,6-tetramethyl-4-piperidone-1-oxyl (TEMPO, TAN or TEMPONE), a stable radical (reaction 10) [53,54]. The reaction of 1O2 with TEMP (k=4 x 107 M-1 s-1 in phosphate buffer at pH 8, [53]) can be used to detect 1O2 with EPR or mass spectrometry (see 2.5.2).
(10)
O2 can react with many biologically important compounds like unsaturated fatty acids of membrane lipids [55]. The reaction of 1O2 with a cis-double bond of an unsaturated fatty acid can form both conjugated and non-conjugated diene hydroperoxides (reaction 11). The rate constants of reactions of 1O2 with unsaturated fatty acids are slightly higher than 104 M-1 s-1 [41].
(11)
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Methionine, tryptophan, tyrosine, cysteine and histidine react with 1O2 to form semistable products with a rate constant around 107 M-1 s-1 [56]. Histidine has been used for detection of 1O2 [35]. 1O2 also reacts with GSH and other thiol containing compounds with rate constants around 106 M-1 s-1. Reaction of 1O2 with GSH is associated with formation of glutathione sulphinate (GSO2H), glutathione sulphoxide (GSSOG), glutathione sulphonate (GSO3H) and glutathione disulphide (GSSG) [57]. Biologically active forms of vitamin B6 react with 1O2 with rate constants around 1 x 108 M-1s-1 [58]. Oxidation of ascorbic acid and plastohydroquinone by 1O2 can proceed as 2-electron reduction of 1O2 to H2O2 [59,60].
2.4. Lifetime and diffusion distance of 1O2
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The lifetime of 1O2 is of interest because a short lifetime would predict that 1O2 reacts at the site of its production whereas a long lifetime would allow 1O2 to act as a signal molecule. The diffusion distance can be approximated from the lifetime (see e.g. [25,61]). The lifetime of 1O2 in cellular environments has been several times measured in D2O based buffers but these measurements do not give a correct picture of what happens in vivo. Current sensor technology has, however, made it possible to measure the kinetics of the 1270 nm luminescence peak after a short laser pulse from some biological materials in H2O based buffers.
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The formation and decay of the 1O2 peak, obtained by exciting a sensitizer with a short laser pulse, remain unaltered if the values of the rate constant of the decay of the triplet state of the sensitizer (kT) and the rate constant of the decay of 1O2 (kΔ) are exchanged [62] (see Appendix in Supplementary information for the modeling of the kinetics of the 1270 nm peak). For this reason, measurements of 1O2 luminescence kinetics always need knowledge about the value of kT in the same experimental system. The long lifetime of 1O2 in D2O alleviates the problem because sensitizers with a short triplet lifetime (1/kT) can be used. If the sensitizer is an intrinsic constituent of the sample, like Chl in plant photosynthesis, the requirement of the knowledge of kT places a particular weight for the correct identification of the sensitizer. Further complications arise from the heterogenity of the biological sample. Theoretical considerations based on general cellular concentrations of histidine, tryptophan and methionine residues in proteins [63] or on the concentrations of various biomolecules like carotenoids and ascorbate that react with 1O2 [64,65] suggested that the lifetime is 0.05‒1 µs in cells [63,64] and 0.2 µs in chloroplasts [65] (Table 2). Measurements of 1270 nm luminescence kinetics from animal cells have, however, yielded longer but extremely variable lifetimes, ranging from 0.4 to 10 µs in cell suspensions or in individual cells (Table 2). Ragas et al. [66] estimate that the lifetime of 1 O2 inside Escherichia coli cells is only 7 ns. The large variation may partially depend on differences between the samples, as the concentrations of substances reacting with 1O2 may vary between cell types. Also the cellular localization of the sensitizer has been shown to affect the measured lifetime [67]. An apparent source of differences is that in some studies, whole tissues have been used whereas in other studies the current laser technology has been applied to measure 1O2 lifetime from single cells (Table 2). One possible reason for long lifetimes obtained in biological materials is the presence of membranes and lipid bodies in cells. The lifetime of 1O2 is 20‒25 µs in micelles [68] and 12.2 µs in liposomes [69]. Interaction between 1O2 with a quencher or a scavenger molecule requires physical contact of the molecules, and therefore the concentrations of quenchers and scavengers in the lipophilic environment would determine how much they shorten the lifetime of 1O2. Hackbarth et al. [70] see two different lifetime components in a mammalian cell system and suggest that one
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corresponds to the membrane and the other to the aqueous phase. However, most experimental data can be explained with only one lifetime component, which suggests that 1O2 equilibrates between different types of compartments within its decay time in the heteronegeous biological environment. Strong laser pulses have also been shown to prolong the measured lifetime of 1O2, probably via depletion of cellular 1O2 quenchers [70].
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So far all 1O2 lifetime measurements from plant and cyanobacteria samples have been done from isolated PSII preparations, either in D2O based buffers [71] or more recently also in water based buffer solutions [65,72,73]. In the case of the simplest PSII reaction centre preparations that lack the QA electron acceptor, the reaction centre triplet is obviously also 1O2 sensitizer. However, the triplet yield of the recombination of the primary radical pair is very low if the electron can proceed from pheophytin to a further acceptor [71], and therefore identification of the sensitizer triplet is more difficult when functional PSII preparations are used. The lifetimes per se mainly reflect the properties of the buffer (D2O or H2O). Correlation between kinetics of decay of the triplet form of the primary donor of PSII and 1270 nm luminescence rise at different temperatures and oxygen concentrations showed that recombination of the primary radical pair produces the sensitizer triplet in PSII particles [71]. Tomo et al. [73] show that much more 1O2 is produced by PSII preparations from a cyanobacterial mutant that contains divinyl Chl a instead of the monovinyl form occurring in the wild type.
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With regard to the potential signaling role, a triplicate lifetime, after which 5 % of 1O2 still exists, could be a practical limit for the reach of the signal. Assuming, in the absence of measurements, that the lifetime of 1O2 is 3 µs in aqueous stroma of chloroplasts and further assuming that 1O2 can pass through the envelope of a typical chloroplast (5‒7 x 2.5 µm; membrane thickness ~4 nm [74]), then approximately 20 % of the volume of the chloroplast could produce 1O2 of which a significant fraction could diffuse to the cytosol. However, 1O2 produced externally to mammalian cells failed to cause DNA strand breaks while 1O2 produced inside the cells did cause breaks [75,76], indicating that in animal cells the amount of 1O2 diffusing through the plasma membrane is small. Diffusion of 1O2 through the plasma membrane or cell wall has not been measured in plants or cyanobacteria. The diffusion distance of 1O2 may also determine the effectiveness of quenchers and scavengers. In this respect, the finding that added β-carotene protects mammalian cells against 1O2-induced cell death but does not shorten the lifetime of 1O2 [77] is highly interesting. The high viscosity of the cellular environment can partly explain the lack of an effect, as a three-fold increase in viscosity from C6H14 to CCl4 is associated with a 30 % decrease of the quenching constant of 1O2 by β-carotene [48]. The short diffusion distance of 1O2 also implies that cell-impermeable detector substances can only measure extracellularly produced 1O2.
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Sensitizer
Method of estimation
Buffer solution
Independent Lifetime measurement
0.05 to 1 µs [64]
0.4‒0.18
Effective signaling distance in viscose cellular environment, µm 0.013‒0.060
< 1 µs [63]
<0.180
<0.06
0.2 µs [65]
0.08
0.03
No
10 µs [79]
0.5
0.2
No
6 µs [79]
0.4
0.15
D2O
Yes
45 µs [80]
1.2
0.4
D2O
Yes
17 µs [67]
0.7
0.25
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System
CR
IP
Table 2. Measurements of 1O2 lifetime with 1270 nm luminescence kinetics in biological systems. An estimated effective signaling distance, at which 5 % of 1 O2 survives, is given for water (D=1.8 x 10-5 cm2 s-1 at 20 °C; [78]) and for viscose cellular environment (D=2 x 10-6 cm2s-1; [61]).
Nucleus of a single neuron HeLa cells (single cell)
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Photofrin 9-acetoxy-2,7,12,17tetrakis(beta-methoxyethyl)porphycene 5,10,15,20tetrakis(N-methyl-4-pyridyl)21H,23H-porphine Chlorin
AC
Cells
Chloroplast stroma Mammalian cells Mammalian cells
Theoretical, based on known concentrations of quenchers/ scavengers Theoretical (His, Trp, Met residues) Theoretical (ascorbate) 1270 nm H2O luminescence 1270 nm H2O luminescence
TE D
Various cellular environments
MA N
T
Effective signaling distance in water, µm
1270 nm luminescence
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Yes
30‒40 µs [67]
1‒1.1
0.3‒0.4
3.2 µs [81]
0.3
0.1
0.4 µs outside of membrane [70] 0.6 µs [82]
0.1
0.04
0.1
0.04
0.03-0.18 µs [82]
0.03‒0.08
0.01‒0.025
3.7 µs; estimated 0.007 µs inside the cells [66] 44‒59 µs (airsaturated medium, 20 °C [71] 3.2 µs [65]
0.3 (based on the extracellular lifetime)
0.005 (based on the estimated cellular lifetime)
IP
Extrapolation D2O/H2O Yes from mixtures measurements in D2O/H2O mixtures 1270 nm H2O Yes luminescence
CR
Neuron (single cell)
D2O
T
5,10,15,20tetrakis(N-methyl-4-pyridyl)21H,23H-porphine
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HeLa cells (single cell)
Pheophorbide a
Leukemia cells
Aluminum tetrasulphonated phthalocyanine
1270 nm luminescence
H2O
Rat tissues in vivo
Aluminum tetrasulphonated phthalocyanine
1270 nm luminescence
H2O
Escherichia coli cells
New Methylene Blue and Zinc tetramethyltetrapyridino[3,4b:39,49-g:30,40-l:3-,4q]porphyrazinium salt
1270 nm luminescence
H2O
PSII reaction centre preparations
PSII reaction centre triplet
1270 nm luminescence
D2O
Yes
PSII reaction centre preparations
PSII reaction centre triplet (assumed)
1270 nm luminescence
H2O
No (assignment supported by effect of
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MA N
Jurkat cells
No (fitting and literature values used) No (fitting and literature values used) No (fitting and literature values used)
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H2O, D2O
Functional PSII particles from wild type and a mutant with divinyl Chl a
PSII reaction centre triplet (assumed)
1270 nm luminescence
H2O
oxygen) Yes
MA N
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No
TE D CE P AC
3.4‒4.3 µs in H2O; 17‒18.7 µs in D2O [72] 3.94‒4.80 µs [73]
T
1270 nm luminescence
IP
Chls (assumed)
CR
PSII particles
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2.5. Detection of 1O2
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2.5.1. Luminescence at 1270 nm Several recent reviews discuss the detection of 1O2 from photosyhthetic organisms [3,83‒85]. 1O2 can be measured by recording the weak 1270 nm luminescence emitted when 1O2 returns to the ground state (reaction 1; Fig. 1; for reviews see [25,84,85]). The detection of 1270 nm luminescence from biological materials is limited by the material thickness, as absorption of water [86] and biomolecules would attenuate the signal to less than one tenth per 1 cm. Therefore, thin samples are preferred. D2O is far more transparent than H2O in the near-infrared range [87].
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Due to the low phosphorescence yield and low concentration of 1O2 in biological systems, measurements of 1270 nm luminescence in photosynthetic material have been limited to generation of 1O2 with strong laser pulses, usually in isolated samples, including PSII reaction centres of pea or PSII particles from cyanobacteria [72,73,88]. Significantly more sensitive detectors than are available now will be needed for the measurement of 1270 nm luminescence from biological material under constant illumination [65]. An additional difficulty is to distinguish 1O2 phosphorescence from the tail of Chl a fluorescence, but the separation can be done by spectral comparison or by comparing the decay times [72].
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In addition to the 1270 nm "monomol" luminescence, 1O2 also emits luminescence at 634 and 703 nm. This "dimol luminescence" is emitted after a collisional formation of a (1O2)2 dimol in the gas phase [89]. Emissions originating from transitions between several excited states of the oxygen molecule are important in atmospheric science [90] but too weak for studies from biological material.
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2.5.2. EPR detectable probes TEMP and TEMPD. The two most popular chemicals used to measure 1O2 from photosynthetic material have been 2,2,6,6-tetramethylpiperidine (TEMP) and the more water-soluble 2,2,6,6tetramethyl-4-piperidinone (TEMPD); both commercially available (see Table 3 for review of chemicals used for 1O2 detection). Reaction between 1O2 and TEMP (reaction 10) or TEMPD produces a stable oxyl radical, and the reaction has been shown to be specific to 1O2 [53,91]. In the case of TEMP, the radical is called TEMPO. TEMPD is available both as a pure compound and as a hydrochloride (TEMPD-HCl); only the hydrochloride form can be used for photosynthetic applications because the other form strongly inhibits PSII [92,93]. Due to the low solubility to water, TEMP cannot be used at temperatures below 20 °C. Both TEMP (experimentally determined LogP = 2.15 [19] and TEMPD (LogP = 0.43, as estimated with ALOGPS2.1 [20]) are thought to penetrate into cyanobacterial cells [94,95]. The amount of the TEMPO radical has traditionally been measured with EPR spectroscopy [96,97]. EPR is not suitable for automated measurements of large numbers of samples, and a mass spectrometry method was recently developed for the quantification of TEMPO [98]. TEMP was found not to inhibit the PSII activity of isolated thylakoid membranes [96]. However, storage of TEMP enriches unknown impurities. In the commercially available TEMP, and also in TEMPD, these impurities are present at such a quantity that they cause strong inhibition of PSII, as demonstrated with fluorescence, thermoluminescence and oxygen evolution measurements [92].
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Vacuum distillation removes the impurities to such extent that TEMP can be used with isolated plant thylakoids without any adverse effects [93], but even distilled TEMP causes an almost immediate inhibition of PSII in live cyanobacteria (Synechocystis sp. PCC 6803, [5]) and in live diatoms Phaeodactylum tricornutum (Fig. 2).
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With distilled TEMP, 1O2 can be detected from isolated plant thylakoid membranes (e.g. [93,96,98]). TEMP has been used also with purple bacteria [99] and TEMPD has been used to detect 1O2 from isolated thylakoids of the green alga Chlamydomonas [100], but in vivo effects of the probes on the photosystems of the organism were not reported. In intact plant material, the TEMPO radical can be reduced to an EPR silent hydroxylamine, and therefore TEMPO is usually extracted with an organic solvent if EPR is used as the detection method [101].
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2.5.3. Dyes Imidazole/RNO. Imidazole-containing compounds, including imidazole itself, are rapidly oxygenated by 1O2 [102,103], and 1O2 can also be detected by measuring bleaching of p-nitroso-dimethylaniline (RNO) by imidazole [104‒106]. The method has been used to measure the action spectrum of 1O2 production in isolated spinach thylakoids [107]. The RNO method should be used with caution in plant material because imidazole has been shown to inhibit PSII oxygen evolution and carbonic anhydrase in a pH-dependent manner [108]. Both imidazole and RNO are commercially available.
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2.5.4. Fluorescent probes SOSG. Singlet Oxygen Sensor Green®, a proprietary fluorescent probe, is highly selective for 1O2. According to the manufacturer's data, SOSG does not react with H2O2 or O2•, and our unpublished experiments have confirmed the high selectivity. Reaction with 1O2 increases the fluorescence yield of SOSG at 520‒540 nm. According to the manufacturer's data, SOSG is cell-impermeable, but a study with tobacco leaves showed that a pinhole administration of SOSG to tobacco leaves leads to preferential association of the dye with the nuclei of epidermal cells [109]. Gollmer et al. [110] showed that SOSG is also able to penetrate into human cells; therefore SOSG can be used in confocal microscopy [111]. A disadvantage of SOSG is that when illuminated with wavelengths below 600 nm, the fluorescence yield of SOSG increases similarly as during exposure to 1O2 (Fig. 3; [92,109]). Furthermore, the reaction product of SOSG and 1O2 sensitizes 1O2 production [112]. Surprisingly few researchers have taken the photosensitivity of SOSG into account (for correct use of SOSG, see [92,109,113]). SOSG caused 15 % inhibition of the photochemical yield of PSII in tobacco leaves [109]. It is unclear if SOSG can be used with live cyanobacteria, as SOSG seems to penetrate only to a small fraction of the incubated cells and fluorescence from both SOSG and Chl a is only rarely detected from the same cell [114]. Illumination of isolated pumpkin thylakoids in the presence of SOSG with >650 nm light caused a very small signal which was not sensitive to D2O, anaerobicity or azide, suggesting that SOSG cannot be used to measure 1O2 from isolated thylakoids [92]. However, SOSG has been used for detecting 1O2 from isolated plant thylakoids and from intact leaves [109,113,115]. Reasons for the contrasting results about the sensitivity of SOSG in plant material are presently unknown. DanePy. Dansyl-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrole (DanePy), another fluorescent probe, is sensitive to 1O2 with minor reactivity with O2•, H2O2 and lipid radicals [116,117]. The probe is not available commercially. DanePy has been used with bean, pea, tobacco, Arabidopsis thaliana and
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Phillyrea latifolia leaves in vivo [109,118‒121] and with the photosynthetic proteobacterium Rhodobacter sphaeroides [122]. DanePy can be used for imaging [118‒120]. DanePy is fluorescent (500‒600 nm, maximum at 545 nm) but forms a non-fluorescent adduct with 1O2, and therefore the presence of 1O2 is inferred from the decrease in fluorescence yield from untreated control to a treated sample. DanePy is harmless for the PSII yield if administered to a leaf either through a pinhole or in the transpiration stream and localizes intracellularly, mainly to chloroplasts [119,120]. DanePy is not light-sensitive [109]. An oxalate derivative of DanePy was used with live Chlamydomonas cells [100] but this dye does not penetrate to leaf cells [109].
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DMAX and DPAX. The probes 9-[2-(3-carboxy-9,10- dimethyl)anthryl]-6-hydroxy-3H-xanthen-3-one (DMAX) and 9-[2-(3-carboxy-9,10- dimethyl)anthryl]-6-hydroxy-3H-xanthen-3-one (DPAX) can be used to detect 1O2 with fluorescence [123,124]. DMAX and DPAX were reported not to react with H2O2, O2• or NO• [123,124], and DPAX has been used with intact chloroplasts [125]. DMAX and DPAX are not commercially available.
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2.5.5. Oxygen consumption in a reaction between 1O2 and histidine Histidine is known as a powerful, biocompatible antioxidant because its imidazole ring reacts rapidly with 1O2 (k=5 x 107 M-1 s-1; [126]; for a review, see [127]). Histidine also protects against HO•, but this protection may be based on the efficient scavenging of metals by histidine rather than reactions between histidine and HO• [127]. Recently, a simple method in which 1O2 production is measured by monitoring the decrease of oxygen concentration in the presence of histidine [128] has been applied to 1O2 measurements from live cyanobacteria [35,129,130]. Cyanobacteria take up histidine from the medium and therefore histidine can be used for 1O2 measurements in live cyanobacteria [35]. The method was shown to be insensitive to O2• and H2O2 [35]. No reports of the use of the histidine method in plants or algae in vivo have been published, and it is possible that in eukaryotes histidine would not enter the chloroplasts that normally export histidine. The reaction between histidine and 1 O2 occurs in neutral to alkaline solutions [126]. 2.5.6. Genetically encoded probes Specific, genetically encoded probes have not been used for the detection of 1O2. However, the IFP1.4 protein containing a biliverdin (an oxidized form of bilirubin) chromophore [131] has been shown to react with 1O2 [132] and not with H2O2, O2• or HO• [132]. The protein emits far-red fluorescence (maximum at 710 nm) when excited at 684 nm, and might therefore act as a perfect genetically encoded 1O2 sensor. The combination of IFP1.4 and a 1O2 sensitizer protein has been used for measurement of protein-protein interaction [132]. Furthermore, fluorescence of the UnaG protein that contains bilirubin as a chromophore was shown to decrease due to a reaction between bilirubin and 1O2 [133], and thus this protein may also be developed into a genetically encoded 1O2 probe. These probes have not yet been applied for detection of endogenous 1O2 production.
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Specificity
Stability
Toxicity
Quantification of TEMPO with EPR [96] or mass spectrometry [98] in vitro
Specific [53,91]
Stable but instant measurements recommended [98]
Not toxic to isolated plant thylakoids [93]; Inhibits PSII in live cyanobacteria [5] and algae (Fig. 2)
Notes
Cell permeable (Synechocystis) [95]
Not toxic to isolated plant thylakoids [93]
-
1.24 (estimated)
Cell permeable (Synechococcus) [94]
Vacuum distil before use [93,92]; Extraction of TEMPO to organic solvent avoids formation of EPR silent hydroxylamine [101] Use TEMPD-HCl [93]
Does not react Stable with H2O2 or O2• [35]
Not toxic
-
-3.32 [19]
-
Only in neutral or alkalic pH [126]
No reaction with H2O2 or O2• (manufacturer's
Little or no effect on photosynthesis
Only red light (>660 nm) can be used
-
Nuclei of epidermal cells; does not localize
The reaction product sensitizes 1O2
US
Light sensitivity
MA N
TE D
Specific, similar Stable to TEMP
CE P
TEMPD (2,2,6,6- Quantification tetramethyl-4of the reaction piperidinone) product with EPR in vitro [100] Histidine Consumption of oxygen by histidine, in vitro; in vivo from cyanobacteria [35] Singlet Oxygen Fluorescence (Ex Sensor Green, 505, Em 525), in SOSG vitro and in vivo
Localization
-
Hydrophobicity (LogP) 2.15 [19]
CR
Method
AC
Compound (IUPAC name) TEMP (2,2,6,6tetramethylpipe ridine)
IP
T
Table 3. Chemicals used to detect 1O2. The stability data indicate the stability of the signal after reaction with 1O2 in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data.
Stable
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in plants [109]
[92,109,113]
Minor reaction with H2O2, O2• and lipid radicals [116,117]
No effect on PSII efficiency [109]
Imaging
-
-
-
Decrease in absorbance at 440 nm
-
-
DPAX (9-[2-(3carboxy-9,10dimethyl)anthryl ]6-hydroxy-3Hxanthen-3-one) IFP1.4 (genetically encoded probe with biliverdin chromophore) UnaG (genetically encoded probe with bilirubin
Fluorescence (Ex 492, Em 516‒517), intact chloroplasts [124]
Does not react with NO, H2O2 or O2• [123]
Decrease in fluorescence (Ex 684, Em 710)
Does not react with NO, H2O2 or O2• [132]
Decrease in absorbance (~475) and fluorescence
Aproprotein reacts with O2• at high concentration
to chloroplasts [109] Cellular; mainly chloroplasts [109,119,120]
production [112] -
Extracellular in plants [109]; cytoplasmic in Chlamydomonas [100] -
Less sensitive than DanePy [109]
-
-
IP
Fluorescence (Ex 364, Em 545 nm) is quenched by 1O2, imaging
Not sensitive
-
-
-
Imidazole inhibits PSII [108]
-
-
-
-0.08 imidazole, [19]/ 1.45 p-nitrosodimethylaniline (estimated) Soluble to water at 10 µM [123]
-
-
-
-
-
Not yet tested for endogenous 1 O2
-
-
-
-
-
Not yet tested for endogenous 1 O2
CE P
TE D
-
AC
-
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CR
DanePy (dansyl2,2,5,5tetramethyl-2,5dihydro-1Hpyrrole) DanePy-oxalate (dansyl-2,2,5,5tetramethyl-2,5dihydro-1Hpyrrole oxalate) RNO (imidazole and p-nitrosodimethylaniline)
MA N
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data)
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CE P
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MA N
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CR
IP
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(~525)
AC
chromophore)
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2.5.7. Further gene expression methods Ability to sense 1O2 may be important both in avoiding damage and in signaling [134], and photosynthetic organisms have several genes that react specifically to 1O2 (e.g. [135]), as well as to other ROS (see e.g. [136]). The mRNA levels of 1O2 inducible genes have been used to mark 1O2 levels in the cells [137]. Also reporter systems, where proteins induced by 1O2 have been fused to marker proteins, have been developed, for example in Chlamydomonas reinhardtii and Arabidopsis thaliana [138,139], respectively.
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2.5.8. Indirect methods of 1O2 detection Damage caused by 1O2. If 1O2 is not quenched or scavenged, it can damage proteins, pigments and lipids [1,140]. The amount of 1O2 can be estimated by the extent of the damage, as reactions between cell components and 1O2 sometimes have specific reaction products. Products of lipid peroxidation, as well as oxidation products of carotenoids, plastoquinone and tocopherol, have been suggested to be used for detection of 1O2 [55,137,141,142], respectively. However, it is not usually very easy to make strong conclusions based on these data, and the ability of the photosynthetic organisms to quench 1O2 can also vary. The capacity of leaf extracts to quench 1O2 can be measured for example according to [143].
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Modifications of the amount or lifetime of 1O2. As the lifetime of 1O2 is lengthened 5‒20 fold in D2O compared to water [46,72], measurements can be done in D2O to facilitate 1O2 detection (e.g. [72]). However, the use of D2O in place of H2O has consequences in biological materials. Proteins are often more stable and more rigid in D2O than in water [144]. It has been shown that with photosynthetic organisms D2O reduces growth and oxygen evolution as well as slows down PSII electron transfer [145] and carbon fixation [146]. Chl is a natural sensitizer of 1O2 in photosynthetic systems but the amount of 1O2 can be increased by using with artificial 1O2 (photo)sensitizers [147]. However, the popular 1O2 generators Rose Bengal and methylene blue have been shown to affect PSII electron transfer [148]. Different 1O2 generators localize in different cellular compartments. For example, Neutral Red and Rose Bengal penetrate to the chloroplast, in contrast to methylene blue [148]. If the sensitizer penetrates weakly to the target tissue, then the amount of 1O2 produced may not be physically relevant. The extent of protection against cellular damage, obtained by a treatment with an 1O2 scavenger like azide, histidine or DABCO, has also been used to assess the importance of 1O2 in the damage mechanism (e.g. [149]). Due to the high reactivity of 1O2, scavengers that localize to the site of 1O2 production, often within a membrane, are preferred. Effects of natural quenchers. Carotenoids and tocopherols that function as the principal quenchers and scavengers of 1O2 can often be used to draw conclusions about the participation of 1O2. For example, participation of 1O2 in the oxidative inhibition of the repair of PSII after photoinhibition was supported by the finding that both in plants [150] and in cyanobacteria [151] the ability to synthesize -tocopherol protects the PSII repair mechanism but does not affect the rate of light-induced damage. The same tocopherol data suggested that 1O2 is not involved in the damaging reaction of photoinhibition, but on the other hand, a cyanobacterial mutant with a high carotenoid content is less sensitive to photoinhibition than the wild type [130].
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3. Hydrogen peroxide, H2O2 3.1. Definition and properties of H2O2
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2.5.9. Summary of 1O2 detection To conclude, 1O2 measurements are a tricky business. Direct measurements of 1O2 are still not possible from physiological conditions. When using chemical traps (see Table 3) it is important to check for the possible adverse effects on the organism or biological function, in addition to checking the sensitivity and reliability of the probe. The place where the sensor compound localizes can also affect the results. More 1O2 probes exist than are used in plant research and also new probes are being synthesized but the usability of the new probes in photosynthetic systems should be tested before extensive use [117,152,153]. For example, the 1O2 probe MVP (trans-1-(2’methoxyvinyl)pyrene) has long been used e.g. in medicine, but it could not detect 1O2 from leaves, maybe because its chemiluminescence (465 nm) overlaps with Chl a fluorescence [109].
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H2O2 is the result of reduction of oxygen by two electrons. In biological systems, H2O2 is mostly present in the neutral form because of its high pKa value (pKa1 is 11.8). H2O2 is a moderate oxidant (E0 of the pair H2O2+2H++2e-/2H2O is 1.776 V; [154]) that can oxidize compounds like thiols.
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3.2. Formation and action of H2O2 in photosynthetic tissues
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In biological systems, including photosynthetic tissues, H2O2 is mainly formed via dismutation of O2• (reaction 29), and because superoxide dismutases (SODs) are present in most cellular compartments, the formation of H2O2 depends on the formation of O2•. Furthermore, the direct 2electron reaction between 1O2 and plastohydroquinone was recently shown to produce a stoichiometric amount of H2O2 (Fig. 4; [60]). The production of H2O2 by the interaction of 1O2 and plastohydroquinone may have importance in chloroplast signaling because H2O2 produced within the membrane would avoid the stromal scavenging mechanisms [60]. In the soluble phase, 1O2 produces H2O2 in a reaction with ascorbate [59]. H2O2 can be enzymatically scavenged by catalases (CATs), peroxidases and peroxiredoxins. CATs have a relatively low affinity for the substrate, reflected by a high Michaelis constant Km for H2O2; usually around 50 mM in plant catalases [155,156], although a lower value of 17.6 mM was also reported [157]. Peroxidases have lower Km values (0.57 mM for ASC, 0.11 mM for H2O2; [158]). Ascorbate and GSH function as electron donors for different peroxidases. Peroxiredoxins form a third group of H2O2 scavenging enzymes. In peroxiredoxins, the oxidizable thiol group is a cysteine residue of the protein itself, and therefore peroxiredoxins require an activating reduction step between the catalytic rounds. H2O2 is known to be associated with reactions causing cellular damage. For example, the action of the herbicide paraquat, known also as methyl viologen, is based on formation of O2•, which inevitably leads to formation of H2O2, too. However, it is usually difficult to determine if the actual damaging agent is H2O2, O2• or HO• formed in the Fenton reaction (reaction 15). Cyanobacteria are highly sensitive to H2O2 [159] and this sensitivity can actually be used to destroy potentially poisonous cyanobacteria from lakes [160].
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In plants, H2O2 functions, together with O2•, as a signal molecule in stomatal opening, programmed cell death and various stress responses (for review, see [161]).
3.3. Reactions of H2O2
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H2O2 is particularly reactive against thiol groups [162], especially those of cysteine residues of proteins and other compounds [16], and the reactivity of H2O2 towards thiols explains the inhibitory effect of H2O2 on the thiol-group-containing enzymes of the Calvin-Benson cycle. H2O2 inactivates fructose-1,6-bisphosphatase, glyceraldehyde 3-phosphate dehydrogenase and xylulose-5-phosphate kinase [163] and the Krebs cycle enzyme aconitase [164] and reacts with several other plant proteins [165,166]. The Escherichia coli transcription factor OxyR recognizes H2O2 via a reaction with a cysteine residue [16,167]. The reaction of H2O2 with thiol groups is also a H2O2 elimination mechanism, accomplished by peroxiredoxins in plant cells [13]. Reduction of H2O2 by thiol groups is a two-step reaction passing through the mechanism described by reactions 12.1 and 12.2.
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H2O2 + RSH RSOH + H2O RSOH + RSH RSSR + H2O
(12.1) (12.2)
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The rate constants of reaction 12.1 range from 102 to 107 M-1 s-1. The thiolate anion (RS), formed in high pH, reacts more rapidly with H2O2 than the respective thiol (RSH) [162,168], and the reactivities of thiols depend on both pH and the pKa value of the thiol [162].
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A well-known reaction of H2O2 is its reduction via the Haber-Weiss mechanism (sum of reactions 13 and 14) and Fenton mechanism (reaction 15). Both mechanisms generate the very powerful oxidant HO•. The rate constant of reaction 13 in aqueous medium ranges from 0.13 to 0.23 M-1 s-1 [169,170] and the rate constant of reduction of H2O2 by HO2• (reaction 14) is 0.5 M-1 s-1. Thus, reactions (13) and (14) are relatively slow in biological material where concentrations of O2• and H2O2 are low. O2• + H2O2 O2 + OH + HO•
(13)
HO2• + H2O2 O2 + H2O + HO•
(14)
M+(n-1)+ H2O2 M+n + OH + HO•
(15)
The reduction of H2O2 by transition metals via the Fenton mechanism (reaction 15) is much more efficient than the Haber-Weiss reaction (reactions 13 and 14). The rate constant of Fenton reactions depend on the metal M (Fe and Cu are the biologically most important ones) or metal complex and pH, and range from 50 to 104 M-1 s-1 [171‒174]. The reduction of H2O2 via the Haber-Weiss and Fenton mechanisms (reactions 13‒15) causes subsequent reactions where interactions between O2•, HO2•, HO• and the reduced and oxidized metal ion can occur. In particular, O2• is able to reduce Fe3+ to Fe2+ and Cu2+ to Cu+, thereby promoting continuing production of HO• via reaction (15). The reduction of H2O2 by peroxidases is one of the main routes of H2O2 scavenging; organic compounds including ascorbate, glutathione and guaiacol can be used as electron donors. In the chloroplast, H2O2 scavenging is mainly catalyzed with an ascorbate-specific peroxidase (APX, (EC
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1.11.1.11)). The catalytic reduction of H2O2 by peroxidases proceeds via the peroxidase ping-pong mechanism (reactions 16‒18) [175]. PX-Fe(III)-P + H2O2 PX-Fe(IV)=O-P+ + H2O
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(16)
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PX-Fe(IV)=O-P+ + AH PX-Fe(IV)=O-P + A• +H+ PX-Fe(IV)=O-P+ AH PX-Fe(III)-P + A• + OH,
(18)
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where P is porphyrin.
(17)
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The rate constant of the reaction of H2O2 with APX is around 107 M-1 s-1 and the Km for H2O2 is 80 M [176,177]. Thus, APX is an efficient enzyme with high affinity to the substrate and a very fast rate of the reaction. CAT-depended scavenging of H2O2 also occurs through the peroxidase-like ping-pong mechanism where one H2O2 molecule is used as an electron donor (reactions 19.1 and 19.2). The decomposition of H2O2 by CAT, unlike the reaction catalyzed by peroxidase, is associated with O2 production. (19.1)
H2O2 + Fe(IV)=O-CAT+→ H2O + Fe(III)-CAT + O2
(19.2)
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H2O2 + Fe(III)-CAT H2O + Fe(IV)=O-CAT+
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The peroxidase-catalyzed reaction of H2O2 with a specific electron donor like luminol can be used for detection of H2O2. In this case, electron donor is oxidized to an optically detectable product (see 3.5.5). In addition to the above, H2O2 irreversibly inactivates two of the three types of SOD (Cu,ZnSOD, Fe-SOD), and APX in the absence of ascorbate [178].
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3.4. Lifetime and diffusion distance of H2O2 As a small, neutral molecule, H2O2 is expected to readily diffuse through membranes. Judging from experiments with the genetically encoded probe HyPer, the lifetime of H2O2 is 1‒3 min in mammalian cells [179] and around 10 s in Arabidopsis thaliana guard cells [180]. Both lifetimes are so long that 5 % of H2O2 would be present at 0.2‒1 mm from the site of generation in a viscous biological material.
3.5. Detection of H2O2 3.5.1. Principles of H2O2 detection H2O2 can be detected with several different compounds functioning as hydrogen donors for the reduction of H2O2. Some detector substances require a catalyst, usually a peroxidase or a metal ion, and if the study object does not contain endogenous peroxidases, horseradish peroxidase can be used together with the detector substance. 3.5.2. Precipitate-forming compounds DAB. The most widely used H2O2 detection chemical in plant research has been 3,3'diaminobenzidine (DAB) (see Table 4 for a list of chemicals used for H2O2 detection). DAB reacts with H2O2, forming a brown precipitate that can be made visible in plant material by removing Chl (Fig. 5; [181‒183]). DAB penetrates well to plant cells and it is not sensitive to light [184], the precipitate is stable [183], and DAB localizes partially to the chloroplast [185]. Unfortunately, DAB strongly lowers
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the quantum yield of PSII electron transfer in the concentration range required for imaging [184]. The inhibition of photosynthesis by DAB should be taken into account in the interpretation of results obtained with DAB from photosynthetic material.
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Cerium. H2O2 reacts with cerium chloride, forming electron-dense precipitates of cerium perhydroxides [186]. This has been used to localize H2O2 production e.g. in wheat roots [187] and Zinnia elegans stems [188] from electron transmission microscope images.
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3.5.3. Dyes MBTH/DMAB. A peroxidase-catalyzed oxidation by H2O2 couples 3-methyl-2-benzothiazoline hydrazone (MBTH) and 3-(dimethylamino) benzoic acid (DMAB) to a deep purple indamine compound that can be assayed spectrophotometrically at 590 nm [189]. Veljovic-Jovanovic et al. [190] showed that a plant leaf extract causes, in addition to a reaction caused by the H2O2 present in the extract, slow but significant unspecific increase in 590 nm absorption. The unspecific increase could be avoided with polyvinylpyrrolidone treatment. Furthermore, ascorbate inhibits the MBTHDMAB signal [190]. The effects of ubiquitous plant metabolites on the MBTH-DMAB signal call for special care to assure that the signal actually reflects the H2O2 content of the study material. The MBTH/DMAB method has been used to measure the increase in extracellular peroxidase activity in sunflower roots during defense signaling [191].
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4-aminoantipyrine/phenol. H2O2 can be measured with 4-aminoantipyrine and phenol as H2O2 oxidatively couples with them forming colourful quinone-imine (absorbance at 505 nm) [192]. Peroxidase is needed for the reaction. The method has been used to measure H2O2 from pea chloroplasts [193] and from root extracts of tomato [194].
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Starch/KI. Iodide ions (I-) are oxidized by H2O2 after which iodine (I2) is complexed with (exogenous) starch, and H2O2 can be histochemically localized by the blue color of the starch/I-complexes. Starch and iodide do not penetrate to cells [188,195] but the starch/KI-method has been used to detect H2O2 production in cut stem surfaces [195] and in peeled or partially digested leaf segments [196] or to measure extracellular H2O2 production by whole roots [197]. Age and wounding of the plant affect the results obtained with the starch method [188]. Iodide can be oxidized also by organic peroxides (ROOH) [198] and other cellular electron acceptors besides H2O2 [195]. DHBS. In the presence of peroxidases 3,5-dichloro-2-hydroxybenzenesulfonic acid (DHBS) is oxidized by H2O2 to its quinone form which then reacts with 4-aminoantipyrine (4-AAP) to produce a colorful substance that can be measured spectrophotochemically at 510 nm. The DHBS method has been used to measure extracellular H2O2 from a tobacco cell suspension [199]. 3.5.4. Fluorescent probes Scopoletin. Decrease of the fluorescence yield of scopoletin (7-hydroxy-6-methoxy-2H-1benzopyran-2-one) is a traditional in vivo method of H2O2 detection in biological systems [200]. In plant leaves, scopoletin mainly localizes to the cell wall but also penetrates to the cells [184]. The usability of scopoletin in plants in vivo is limited by the need to use ultraviolet (UV) wavelengths (346 nm) for excitation, as plant leaves strongly absorb UV light [201]. The 435 nm emission of scopoletin is also strongly absorbed by Chl a. Scopoletin has been used for measurement of H2O2 in root cells
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[202] and in the incubation media of callus cells [203] but also from leaves of Pseudowintera colorata [204].
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Eu3Tc. An europium-tetracycline complex (Eu3Tc) [205] reacts with H2O2 to form a highly fluorescent adduct (excitation at 360‒440 nm; emission at 616 nm). The fluorescence yield of Eu3Tc-H2O2 adduct is strongly dependent on pH [205] and therefore the complex should be used only in a buffer solutions at pH 6.6‒7.2. Eu3Tc is not photosensitive but lowers PSII yield [184]. Eu3Tc has not been used with other photosynthetic materials.
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Amplex Red and Amplex Ultrared. Amplex Red is 10-acetyl-3,7-dihydroxyphenoxazine which, after oxidation by H2O2, fluoresces at ~580 nm when excited at 570 nm; Amplex Ultrared is a derivative of the original reagent. The commercial kits also contain horseradish peroxidase as a catalyst. Both compounds have been tested for H2O2 imaging from leaves, and Amplex Red penetrates to cells and even to chloroplasts whereas Amplex Ultrared was found to stay mostly in the apoplast [184]. Both compounds have a strong negative effect on PSII yield [184], which limits their use in plant material. Furthermore, both compounds are sensitive to light [184]. Cyanide, salicylhydroxamic acid, and propyl gallate have been shown to inactivate the Amplex Red method in vitro [206]. Antal et al. [207] used Amplex Ultrared to measure H2O2 formation by isolated thylakoid membranes after treatments with short flashes in the dark.
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3.5.5. Chemiluminescent probe Luminol. 3-amino-phthal-hydrazide, known with the common name luminol, is oxidized by H2O2 in a peroxidase-catalyzed reaction to produce an excited state of 3-aminophthalate; the decay of the excited state yields blue luminescence peaking at 440 nm. Reaction with H2O2 requires a peroxidase or a metal ion as a catalyst, and a mixture of luminol and H2O2 has been used from the 1930's in crime scene investigation to detect the iron of haemoglobin present in blood stains. If the catalyst is provided, luminol can be used to measure H2O2, and a detection limit of 0.42 nM was obtained for cobalt-catalyzed measurement of H2O2 from seawater [208]. The luminescence of luminol is quenched by many cellular substances like ascorbic acid and phenols, which limits the usability of luminol for H2O2 measurements from intact biological materials [190]. Luminol chemiluminescence is also observed in the presence of peroxynitrite and O2• ([209]; see section 4.5.5). Plastoquinone was recently found to enhance the luminescence [60]. However, the Co(II) catalyzed reaction offers the very high sensitivity of 0.5 nM for plant extracts diluted to such extent that the quenchers and enhancers do not disturb the reaction [210]. The luminol/Co(II) method revealed an increase in H2O2 concentration in Arabidopsis thaliana leaf tissue after a shift to high light [211] and a hypoxiainduced increase in the H2O2 level of grapevine buds [212]. 3.5.6. Microelectrodes Microelectrodes. Electrochemical H2O2 microsensors (for O2• electrodes, see 4.5.6) have been developed (for a review see [213]). Ren et al. [214] developed a carbon fiber Pt-ultramicroelectrode coated with single-walled carbon nanotubes and hemoglobin (Hb/SWCNTs/CFUME), and were able to monitor the oxidative burst (H2O2) in Aloe leaves in vivo with a detection limit of 4 μM. Neither 0.1 mM ascorbic acid nor NaCl affected the electrode. A microelectrode with poly-ophenylenediamine and Pt microparticles (POPD–Pt-MP–Pt) has been used to measure H2O2 induced by Cd2+ treatment from leaves of oilseed rape in vivo [215]. Another Pt-electrode has been used in
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vitro with Arabidopsis thaliana cell suspensions [216]. The small size of current electrodes (diameter less than 10 µm [214]) facilitates measurements from intact plants.
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3.5.7. Genetically encoded probes With genetically encoded sensors there would be no need to get foreign molecules inside the cells to detect H2O2 (or O2•, see 4.5.7) and they also offer ways for locating the ROS in the cell, and the use of (fluorescence) microscopy (for reviews see [217,218]). The ability of H2O2 to oxidize the thiol group of OxyR [167]; reactions 12.1 and 12.2] was used in designing the H2O2 sensor HyPer, in which circularly permuted yellow fluorescent protein (cpYFP) was fused with the regulatory domain of OxyR [21]. HyPer functions as a genetically encoded H2O2 sensor. Hyper is claimed to be specific to H2O2 [21], and it has been used also in Arabidopsis thaliana to measure H2O2 produced in the peroxisome lumen and in the cytoplasm [180]. HyPer has excitation maxima at 420 and 500 nm and emits at 516 nm, and the oxidation of HyPer by H2O2 enhances the 500 nm peak and lowers the 420 nm peak. However, the sensor (like many fluorescent proteins) shows strong pH-dependence [179], and therefore the measurement should be coupled with measurement of pH. Many fluorescent proteins themselves produce ROS when illuminated, which may lead to photobleaching [219,220], but Hyper appears to show little photobleaching [221]. Another issue with genetically encoded probes is that they act as antioxidants, and their continuous expression may affect the physiology of the organism [222]. Recently published genetically encoded H2O2 probes (OxyFRET and PerFRET) respectively apply the Orp1 and Yap1 proteins of the H2O2 sensing system of yeast [223]; these probes have not yet been tested with plant material.
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3.5.8. Unspecific probes H2O2 has been measured from plant material [224,225] and from Chlorella cells [226] with fluorescein derivatives that react primarily with HO• [227]. Because these dyes can be used to detect more than one ROS, they will be described in Chapter 8. 3.5.9. Indirect methods of H2O2 detection Increase in the activities of H2O2 scavenging enzymes, including APX and CAT [180] or increase in the expression levels of genes coding for these enzymes may suggest an increase in the amount of H2O2 in the tissue. A considerable number of genes are affected by H2O2 in plants [136,228] and in cyanobacteria [229]. Many of these genes are also regulated by light [230], and plant chloroplasts produce H2O2 in the light. Purified H2O2 scavengers offer possibilities for simple measurements of H2O2 in vitro. For example, H2O2 production by a cell suspension has been measured via the reaction of a scavenger [231]. 3.5.10. Summary of H2O2 detection Some methods of H2O2 determination have been used long, and lots of knowledge about different aspects of their use has been accumulated whereas the caveats in newer, promising methods may still remain to be found. Unfortunately, most detection methods that have been studied with regard to their effects on photosynthesis seem to have adverse effects. This is particularly important for DAB, as this dye localizes to the chloroplasts in DAB-infiltrated leaves. Convincing determination of H2O2 requires the use of more than one method, and measurements of H2O2 from extracts are recommended to back up in vivo staining experiments.
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Specificity
Stability
Brown precipitate, Imaging, semiquantitative
Specific
Stable
Cerium
Electron transmission microscopy Absorbance at 590 nm
-
-
Ascorbate inhibits the signal [190]
-
Absorbance at 505 nm
-
Histochemical staining
Oxidized by ROOHs and cellular electron acceptors [195,198] -
Toxicity
Hydrophobicity (LogP) 0.24 (estimated)
Localization
Notes
Cellular; penetrates to chloroplasts [184] -
Too low sensitivity at acceptable concentration [184]
-
-
-
TE D
Strongly lowers PSII yield [184]
-
-
1.41 MBTH (estimated)/ 1.35 DMAB (estimated)
In vitro or from ground tissue
-
-
-
-
-
-
Peroxidase is needed for the reaction
-
-
-
1.04 iodide [19]
Cellimpermeable [195]
Wounding affects the results
-
-
-
-
-
Peroxidase is needed for the reaction
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DHBS/4-AAP (3,5- Absorbance at 510 dichloro-2nm hydroxybenzenes
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MBTH (3-methyl2benzothiazoline hydrazone) and DMAB (3(dimethylamino) benzoic acid) 4aminoantipyrine/ phenol Starch/KI
Light sensitivity Not sensitive [184]
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Method
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Compound (IUPAC name) DAB (3,3'diaminobenzidin e)
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Table 4. Methods of detection of H2O2. Light sensitivity data are for visible light, unless otherwise indicated. The stability data indicate the stability of the signal after reaction with H2O2 in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data.
-
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-
-
Signal affected by several metabolites and pH [205] -
Decrease Not sensitive to <50 % [184] in 4‒5 min in leaves t1/2 = 6‒8 min
Amplex Ultrared (modified from Amplex Red)
Fluorescence (Ex 567/ Em 580), Imaging
-
Luminol (3amino-phthalhydrazide)
Chemiluminescence at 430 nm
Hb/SWCNTs/CFU ME (Microelectrode
Amperometry
Interference by ascorbate, phenolics [190] and plastoquino ne [60] -
1.62 (estimated)
Cell wall, cytoplasm [184]
-
Lowers PSII yield [184]
-
Mostly intercellular [184]
-
Strongly lowers PSII yield [184]
1.93 (estimated)
Addition of peroxidases may be needed.
Strongly lowers PSII yield [184]
-
At least partly cellular, accumulates in chloroplasts [184] Limited penetration to cells [184]
-
1 % increase in signal in 30 min at PPFD 1200 µmol m2 -1 s [184] 30 % increase in signal in 30 min at PPFD 1200 µmol m2 -1 s [184] -
-
-0.41 (estimated)
In vitro or from ground tissue
Use of Co as catalyst facilitates measurements from plant tissues [210]
-
-
-
-
-
Not affected by ascorbic acid or NaCl [214]
-
US
MA N
TE D
CE P AC
Amplex Red (10acetyl-3,7dihydroxyphenox azine)
-
CR
Fluorescence (Ex 346/ Em 435) quenching, Quantification, imaging Reversible formation of fluorescent (Ex 400/ Em 615) Eu3Tc-H2O2 complex, Imaging Fluorescence (Ex 570/ Em 583), Imaging
Europiumtetracycline complex (Eu3Tc)
IP
ulfonic acid/ 4aminoantipyrine) Scopoletin
T
31
Linear decrease to <50 % in 15 min
Addition of peroxidases may be needed
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IP CR
Only little photobleachin g [221]
-
-
Can be targeted to desired tissue in vivo
Fluorescence depends strongly on pH [179].
-
-
Can be targeted to desired tissue in vivo
Moderate pH sensitivity [223]
MA N
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-
-
-
TE D
Fluorescence ratio (Ex 435 nm, Em 535 nm/480 nm)
Specific to H2O2 [21] but may be affected by other oxidants [179] Specific to H2O2 in vivo [223] but not tested with isolated protein
CE P
OxyFRET and PerFRET (genetically encoded probes based on Orp1 and Yap1, respectively).
Fluorescence ratio (Ex 420/500 nm, Em 516 nm)
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based on haemoglobin and carbon nanotubes) HyPer (genetically encoded probe based on OxyR)
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32
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4. Superoxide anion radical, O2•, and hydroperoxyl radical, HO2• 4.1. Definitions and properties of O2• and HO2•
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Reactions and biological activities of O2• have interested the scientific community for tens of years, and several reviews are available (see [232‒235]). O2• is 1-electron reduced form of O2. The midpoint redox potential (Em) of the pair O2/O2• at pH 7.0 is -160 mV vs Normal Hydrogen Electrode (NHE) in aqueous solution [236]. The Em of the pair O2/O2• becomes more positive if O2• is protonated to form HO2•, and therefore HO2• is a stronger oxidant than O2• [237]. However, O2• is a weak deprotonation agent in aqueous solutions. O2• can form O2•(H2O)n complexes (with n from 1 to 3 [233] and the free energy of hydration was estimated to be around 355 kJ mol-1 [238]. The pKa value for HO2• is 4.8 and therefore only 0.25 % of O2• is protonated at pH 7. In aprotic medium O2• forms a weaker solvation complex. The pKa value of HO2• in DMF was estimated to be 12, and therefore O2• can act as a strong deprotonation agent in an aprotic medium. Weak solvation of O2• in aprotic media leads to a shift of the redox potential of the pair O2/O2• in comparison with an aqueous solution. The redox potential of the pair O2/O2• in DMF was estimated to range between -550 and -600 mV vs NHE [233].
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In addition to its role as an oxidant in reactions that produce H2O2 (see reactions 32.1, 34 and 35 below), O2• can act as a reductant, which leads to conversion of O2• to O2 (reactions 23, 24, 32.2 and 33); see [237] for a compilation of reactions of O2• and HO2•. Enzymatically catalyzed dismutation of O2• proceeds via oxidation of one O2• ion and reduction of another one. The dual role of O2• is unique among ROS, as other ROS function as oxidants in biological environment.
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4.2. Formation and action of O2• and HO2• in photosynthetic tissues 1-electron reduction of oxygen by PSI is the main photosynthetic source of O2•, but small amounts of O2• are also produced by PSII [2,239] and by reduced forms of plastoquinone [60]. O2• is invariably scavenged by SODs (EC 1.15.1.1), enzymes catalyzing the dismutation reaction (reaction 29). Almost all aerobic organisms contain SOD(s), and the enzymes fall in the phylogenetic groups of Mn and Fe-SODs, Cu,Zn-SODs and Ni-SOD [240]. The Mn and Fe-SODs form one clade. Chloroplasts contain mainly Cu,Zn-SODs [240]. All SODs speed up O2• dismutation, but the reaction proceeds even in the absence of an enzyme. In addition to SODs, redox active metals like manganese ions catalyze the dismutation.
4.3. Reactions of O2• and HO2• According to its thermodynamic properties, O2• expresses dual reactivity; it acts as a weak oxidant and as a strong reducing agent in electron transfer reactions in aprotic media in the absence of proton donors. The redox potential of O2•/HO2 (data are available only for aprotic medium) is very low, -1.75 V in DMSO [232]. On the other hand, O2• is able to oxidize organic molecules via deprotonation-oxidation mechanism in the presence of protons or proton donors. Thus, the behavior of O2• in redox reactions very strongly depends on its protonation. Furthermore, O2• reacts preferably with neutral or radical partners and only slowly with negatively charged ones.
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In addition to electron-proton transfer reactions, O2• can take part in nucleophilic substitution or addition if the substrate has a low electron affinity. O2• participates in nucleophilic reactions with alkyl halides, sulfonates, phosphates, esters, acyl halides, acyl anhydrides to form peroxyl (ROO•), alkoxyl (RO•) radicals and epoxides as intermediates [232,233,241,242]. Both the nucleophilicity and the basicity of O2• depend on medium. O2• acts as a powerful nucleophilic agent in aprotic medium due to rapid protonation and dismutation in aqueous solutions.
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Reactions of O2• with organic and inorganic molecules fall in five basic categories: protonation, electron transfer reaction, nucleophilic substitution and addition, attraction of hydrogen, and addition to metal or metal complex.
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Protonation. Protonation of O2• is a highly important reaction because this reaction is the main source of HO2• and is involved in a complicated deprotonation-oxidation process as well. O2• can be protonated via a reaction with proton or a proton donor such as ascorbic acid, reactions (20) and (21) respectively. O2• + H+ HO2• O2• + AscH2 HO2• + AscH
(20) (21)
(22)
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O2• + -TocH HO2• + -Toc
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O2• can be also protonated by -tocopherol (-Toc) in aprotic medium, reaction (22):
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Electron transfer. O2• can reduce many organic and inorganic molecules such as quinones and cytochromes, reactions (23) and (24), respectively, via a 1-electron transfer mechanism. O2• + Q O2 + Q•
(23)
O2• + Cyt c(Fe3+) O2 + Cyt c(Fe2+)
(24)
Nucleophilic addition and substitution. Nucleophilic reactions of O2• with organic molecules are associated with formation of corresponding peroxy radicals which are more oxidizing than O2• itself. For example, O2• reacts with carbon tetrachloride (CCl4) to form the trichloromethyl peroxy radical [243], reaction (25). O2• + CCl4 Cl3COO• + Cl
(25)
Nucleophilic reaction of O2• with nitrones is widely used for detection of O2• with EPR. The reaction of O2• with 5-diethoxyphosphoryl-5-methyl-1-pyrroline N-oxide (DEPMPO) yields a stable DEPMPO/O2• adduct (reaction 26) [244].
(26)
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Attraction of hydrogen. Attraction of a hydrogen atom from organic molecules by O2• is an unlikely reaction. Most probably O2• dependent attraction of hydrogen is the result of its protonation. HO2• is a typical neutral radical and is able to attract a hydrogen from organic molecules such as linolenic acid [237], reaction (27).
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HO2• + LH H2O2 + L•
(27)
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Addition to metal of metal complex. A reaction of O2• with transition metals or metal complexes yields unstable and stable intermediates [233,237]. For example, O2• can react with manganese complexes Mn2+-L to form MnOO+L complexes [245], reaction (28). Mn2+-L + O2• MnOO+-L
(28)
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Complex reactions involving O2•. O2• is involved in many complicated reactions. Below we will describe only the most important O2• depended reactions occurring in photosynthetic organisms.
O2• + O2• H2O2 + O2
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Dismutation is one of the main reactions determining the lifetime of O2• and its possibilities to take part in other biochemical processes. Dismutation is the reaction between two molecules of O2• and can occur via noncatalytic and catalytic routes. Noncatalytic dismutation of O2• proceeds, in general, in aqueous medium and it can be written as a reaction (29). Dismutation is a 2-step reaction; the first is protonation of O2• (reaction 20), and the second step is a radical-radical reaction between O2• and HO2• or between two molecules of HO2•, reactions (30) and (31) respectively. (29) (30)
HO2• + HO2• H2O2 + O2
(31)
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O2• + HO2• + H+ H2O2 + O2
Dismutation of O2• is very slow in alkaline pH (0.3 M-1 s-1) because O2• is mainly deprotonated. Protonation of O2• leads to an increase in the reaction rate constant with a maximum (108 M-1 s-1) at pH 4.8, the pKa value of HO2•. In acidic pH less than 4.8, when O2• is mostly protonated, the reaction rate constant is reduced to 8.6 x 105 M-1 s-1 [237]. The catalytic route of O2• dismutation involves the SOD enzyme (EC 1.15.1.1). The general reaction can be written as follows: M(n+1)+-SOD + O2• → Mn+-SOD + O2
(32.1)
Mn+-SOD + O2• + 2H+ → M(n+1)+-SOD + H2O2,
(32.2)
where M is a metal ion present in SOD. The rate constant of O2• dismutation catalyzed by SOD is about 6.4 x 109 M-1 s-1 [246]. O2• and HO2• can react with H2O2 via the Haber-Weiss reaction (reactions 13 and 14). O2• can also mediate the reduction of H2O2 in the Fenton reaction (reaction 15) by reducing a transition metal ion, e.g. Fe3+ (reaction 33) [247].
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O2• + Fe3+ O2 + Fe2+
(33)
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In addition to the Haber-Weiss reaction, O2• can directly participate in decomposition of ROOHs. O2• can react with ROOH yielding the ozonide anion radical and aldehydes or ketones as final products [248]. O2• is able to react with benzoyl peroxide in toluene with formation of 1O2 [249]. In cells, decomposition of ROOH by O2• might be observed in hydrophobic regions like membranes.
O2• + AscH2 + H+ → H2O2 + AscH•,
(34)
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where AscH• is monodehydroascorbate radical.
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In cells, O2• is particularly reactive against iron-sulfur centres, and the ability of the Escherichia coli transcription factor SoxR to sense O2• is based on this reactivity [16]. Ascorbic acid and reduced GSH can participate in scavenging of O2•. The oxidation of ascorbic acid by O2• is a complicated reaction concerted with transfer of a hydrogen atom and a proton [250]. The general reaction of O2• with ascorbic acid (reaction 34) can be represented in the following way:
O2• + GSH + H+ → H2O2 + GS•
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O2• can react with reduced GSH to form H2O2 and gluthyl radical (GS•), reaction (35). Addition of O2 to GS• is considered as a source of peroxysulfenyl radical (GSOO•) [251]. (35)
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The rate constant of the reaction of O2• with reduced GSH was estimated to be 103 M−1 s−1 at maximum [162,252]. It is much less than the rate constant for the reaction of O2• with ascorbic acid, 3.3 x 105 M−1·s−1 at pH 7.8 [253].
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The ability of O2• to bind with transition metal complexes determines its toxicity to metalloenzymes. O2• can inactivate the ferredoxin-dependent nitrate reductase, interacting with molybdenum in its active center [254]. CAT passes into an inactive form due to interaction of O2• with its heme [255].
4.4. Lifetime and diffusion distance of O2• High pH and low concentration of O2• increase its lifetime by affecting the dismutation rate (reactions 29‒30; [256]). According to Fridovich [257] the lifetimes of 0.1 mM and 0.1 nM solutions of O2• in water are 0.05 s and 14 h, respectively. In seawater, 87‒1120 pM concentrations of O2• have been measured, with half-life of 10‒100 s [258,259]. In animal cells the half-life of O2• has been calculated to be below 50 ms, resulting in 40 µm diffusion distance [260,261]. To our knowledge, no measurements of the lifetime of O2• in plant cells have been published; in general, the lifetime would be controlled by SOD (reaction 32; [257]). Due to its negative charge, O2• cannot easily pass through membranes [262], and in cellular pH only a minor fraction would be in the protonated form. Plant PSII particles have been shown to produce 1‒2 µmol O2• (mg chl)-1 h-1 in high light [263], and extracellular O2• production by Chlamydomonas in UV-light was measured to be 0.60‒2.40 µmol g-1 min-1 [264].
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4.5. Detection methods of O2•
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4.5.1. Spectroscopic methods O2• can be detected directly by spectroscopy (absorption at 230–350 nm; [265]) or with EPR spectroscopy [266], but due to the strong absorption of cell components at UV-range and the low amount of O2•, these methods are rarely useful in biological applications. Changes in the EPR spectrum, induced by UV treatment of attached plant leaves, were attributed to O2• [267], but the origin of these signals is controversial [268].
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4.5.2. EPR detectable probes DMPO, EMPO, DEPMPO, BMPO, DCP, TMT and Tiron. O2• produced by photosynthetic organisms has often been measured with probes that, after reacting with O2•, can be detected with EPR spectroscopy (for reviews, see [269,270]; see Table 5 for a list of chemicals used for detection of O2•). The most widely used probes are 5,5-dimethyl-1-pyrroline N-oxide (DMPO), 5(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide (DEPMPO; reaction 26), 5-(ethoxycarbonyl)-5methyl-1-pyrroline N-oxide (EMPO) and 5-tert-butoxycarbonyl-5-methyl-1-pyrroline N-oxide (BMPO; Fig. 6), all of which are commercially available. DMPO has been used to detect extracellular O2• production by intact cells of a red alga Chattonella marina [271] and with EMPO, O2• has been detected from isolated PSII membranes of spinach [272]. DEPMPO has been used for O2• detection from isolated tobacco thylakoids [273], from reaction center complexes of spinach [274] and from isolated plasma membranes of maize [275]. BMPO has been used to detect O2• from cell wall extracts of pea [276] and from isolated PSII membranes of spinach [277]. The lipophilic cyclic hydroxylamine probe 1-hydroxy-4-isobutyramido-2,2,6,6-tetramethylpiperidinium (TMT) and the more hydrophilic probe 1-hydroxy-2,2,5,5-tetramethylpyrrolidine-3,4-dicarboxylic acid (DCP) have been used to detect O2• from pea thylakoids [278]. In comparison to the traditional EPR probes, also commercially available TMT and DCP are said to be more sensitive, and can be also used to detect the location of O2• production [278,279], although TMT might be re-reduced by photosynthetic electron transport chain [278]. The spin trap 4,5-dihydroxy-1,3-benzene-disulfonic acid disodium salt (Tiron) has been used to detect O2• from wheat roots [280] and from lichen tissue [281]. OXANOH and PTM-TC. The spin traps 2-ethyl-1-hydroxy-2,5,5-trimethyl-3-oxazolidine (OXANOH) and perchlorotriphenylmethyl radical-tricarboxylic acid (PTM-TC) have been used to detect O2• from spinach thylakoids [282,283], respectively, and from plant roots [284]. O2• induces oxidation of OXANOH to OXANO, but it is also auto-oxidized in the presence of metal-ions [282]. In contrast to most of the spin traps, the EPR signal of PTM-TC decreases in the presence of O2•. OXANOH and PTM-TC are not commercially available. Specificity. DMPO, EMPO, BMPO and DEPMPO are not specific to O2• but react also with other radicals (especially HO• and carbon centered radicals). Fortunately, reaction products of the spin trap with different ROS often have different EPR spectra [273,285‒287] (Fig. 6; see also 5.5.2). Thus, the limited specificity of these EPR probes is not only a drawback but also an advantage, as several ROS can be detected with one experiment. For example, Bogdanovic Pristov et al. [276] used DEPMPO to detect the carbon dioxide radical (CO2•), O2• and HO•. In contrast, the ability of cyclic hydroxylamines (e.g. TMT and DCP) with other than O2• limits the usability of these compounds
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because the different products usually have similar EPR spectra [279]. Interference with transition metal ions can be reduced by using chelating agents [279]. PTM-TC does not react with HO•, ROO•, H2O2, NO•, GSH or with L-ascorbate [288].
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Lifetime of spin adducts. A challenging feature of the EPR probes is that the spin adducts usually decay rapidly [289,290], especially the spin adducts of DMPO (t1/2 is about 45 s; [291]). The spin adducts of DEPMPO and EMPO are more stable, for EMPO t1/2 is about 8 min and for DEPMPO t1/2 is about 14 min [286,287,289], and of these two the EPR spectrum of the EMPO adduct is less complicated [287]. The half-lifetime of the BMPO adduct is 23 min [292]. The spin adduct of PTM-TC is stable [288]. Also Tiron has been reported to be quite stable (t1/2 is about 15 min), but the need for basic (pH 8.5) solution might restrict its use [280]. Efforts have been made to design more stable spin probes (for example [293]). Furthermore, cyclodextrins can be used to stabilize the spin adducts [290]. Cyclodextrins have also been shown not to significantly inhibit photosynthesis when used in appropriate concentrations [290]. With plant material, DMPO is more commonly used as HO• probe (see 5.5.2), however, the DMPO/O2• spin adduct decays to the HO• adduct and also to HO• itself [294]. Furthermore, UV-radiation can cause generation of the DMPO cation radical, which after reaction with water generates the hydroxyl adduct of DMPO [295]. Spin adducts of DEPMPO or EMPO do not decay to the HO• adducts [286,288] or decay only slowly (>20 min with DEPMPO) [269].
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Test of the effects of these probes on photosynthesis are rare. TMT and DCP do not affect the reduction of P700+ (although in the precence of DCMU, TMT was observed to slightly accelerate the reduction of P700+) [278]. Long incubations of plant roots with Tiron exert a negative effect on root growth [296]. Long (6 h) incubations of animal cells with DMPO, EMPO, BMPO and DEPMPO have been shown to slow down growth and cause death of the cells [297]. Higher concentrations than 90 mM of DMPO may be toxic to algae [271]. In vivo detection of O2• with EPR probes is still rare. Warwar et al. [284] were able to obtain onedimensional images of O2• production in a whole plant root using PTM-TC, and they also concluded that the probe stayed in the apoplastic space. More commonly used EPR probes, DMPO, BMPO and DEPMPO are thought to be membrane-permeable [298]. 4.5.3. Dyes NBT, NBD and XTT. Nitro blue tetrazolium (NBT) is a traditional O2• dye that has also been used with plant material. The reaction of NBT with O2• can be followed spectrophotometrically at 550‒560 nm. O2• has been detected with NBT from legume roots [299], pea leaves [300], diatoms [301] and from extracted leaf material of maize [302]. To quantify the signal, the biological material has to be made transparent by removing Chl [299] or centrifuged away [302], but the precipitated product of the NBT-O2• reaction can be histochemically localized. A method for quantification of O2• from stained leaves has been developed [303]. Both NBT and its reaction product are quite water insoluble. A more water soluble O2• probe sodium, 3´-(1-[phenylamino-carbonyl]-3,4- tetrazolium)-bis(4-methoxy-6-nitro) benzene-sulfonic acid hydrate (XTT), commonly used as a cell viability assay [304], has been introduced [305]. XTT has
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been used to detect O2• from tobacco cell suspensions and from cucumber leaf extracts [306,307], respectively.
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NBT can be reduced also by P680 of PSII and CO2- [83,308] and it has also been shown to generate O2•, although this reaction may not be significant [308]. Furthermore, detergents like Triton X-100 increase NBT reduction [309]. A related probe, 4-chloro-7-nitrobenzo-2-oxa-1,3-diazole (NBD), has been reported to be more selective towards O2• [310], but NBD has not been used, to our knowledge, with photosynthetic organisms. Neither is XTT an optimal O2• probe, as it has been shown that Escherichia coli suspensions can reduce XTT also anaerobically (with NADPH:XTT reductases; [311]) and short-chain sugars and phenolics reduce XTT, which makes the use of whole cell material difficult [311,312].
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Cytochrome c. Reduction of (exogenous) (ferri)cytochrome c (Cyt c, commercial) has often been used to detect O2•, as the reduction of Cyt c can be monitored spectrophotometrically (around 550 nm; [313]). The Cyt c method has been used in vitro to measure O2• from PSII-enriched membrane fragments from higher plants [263], from apoplasts of wheat roots [314] and from Chlamydomonas sp. ICE-L suspension [264]. In addition to O2•, Cyt c can be reduced by the components of the photosynthetic electron transport chain [315,316] and by ascorbate, GSH and other reductants [317]. In low and moderate light intensities (at PPFD 5–200 µmol m-2 s-1) Cyt c was mainly reduced by the plastoquinone pool and not by O2• [316]. For this reason, comparison of the results obtained with and without added SOD is always required [263]. To enhance the specificity of Cyt c to O2•, it is possible to use acetylated Cyt c [318], for example with Chlamydomonas in [264]. However, reduced Cyt c can also be reoxidized by many cell components, like cytochrome oxidases and peroxidases and by oxidants like peroxynitrite (ONOO-) [317], which calls for care with the use of the method.
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Epinephrine. Epinephrine (also called adrenaline) has been used to detect O2• as after the reaction with O2•, the absorbance of the reaction product, adrenochrome, can be measured (at about 490 nm; [319]). Epinephrine has been used to measure O2• from chestnut seeds [320] and spinach chloroplasts [313], extracellular O2• production from bryophytes and lichens [281] and roots [191], and mitochondrial O2• production from barley roots [321]. This method is not specific to O2•, as epinephrine has been shown to react also with H2O2 [322], and therefore specificity assays are always needed (see e.g. [191]). In addition, adrenochrome can be metabolized by a NADPHdependent cytochrome P450 reductase [323] and take part in redox cycling with mitochondrial complex I, resulting in fast increase in O2• production [324]. For the latter reason the authors [324] advise to measure only the initial part (60 s) of O2• production after addition of epinephrine. Hydroxylamine. O2• can be detected by following the formation of nitrite from hydroxylamine (NH2OH) with absorbance at 530 nm, but pH should be below 8 to avoid generation of O2• by autooxidation of hydroxylamine [325]. Comparison with measurements in the presence of added SOD may be needed to exclude unspecific reactions. The method has been used to measure O2• from homogenized leaves of maize [326]. Hydroxylamine penetrates well to leaf cells [327]. A drawback of this method is that hydroxylamine is known to reduce the manganese ions of the oxygen-evolving complex of PSII, which leads to loss of PSII electron transfer capacity [327,328].
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4.5.4. Fluorescent probes HE. The commercially available probe hydroethidine (HE; or dihydroethidine DHE) has been widely used to detect O2•, as its reaction product emits yellow to red fluorescence. HE is permeable to both mammalian and plant cells [329,330] and oxidized HE intercalates with DNA, which enhances the fluorescence. HE also has mitochondria-specific analog, Mito-HE or MitoSOX red [331]. Unfortunately, also HE reacts with other substances than O2•, for example with cytochromes, hemeproteins, manganese-porphyrin complex, hypochlorous acid NO and ONOO− [332‒334], but the reaction product of HE and O2• differs from the unspecific reaction products [335]. To enhance the specificity of fluorescence detection, one should use 490 nm light for excitation and detect 560‒570 nm fluorescence. However the spectra of the O2• specific and unspecific reaction products overlap so strongly [335] that reliable detection requires high performance liquid chromatography (HPLC) measurements [335,336]. HE can also be oxidized in UV, and in some conditions also in visible light [337], which prevents detection of light-induced O2• formation. Additional issues weakening the usability of HE are that is has been shown to increase the dismutation rate of O2•, and be toxic for Escherichia coli [332]. Nevertheless, HE has been used to detect O2• from plants, for example from tobacco cells and pea roots with a fluorescence microscope [338] and from spinach leaves with HPLC [339]. For HPLC detection, a simplified method has been developed [340], though, at least to our knowledge, it has not been used with plant material. Martin et al. [341] measured O2• production by Arabidopsis thaliana mitochondria by Mito-HE.
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4.5.5. Chemiluminescent probes Three commercially available chemiluminescent probes, lucigenin (N,N'-Dimethyl-9,9'-biacridinium dinitrate; luminescence at 470 nm), CLA (2-Methyl-6-phenyl-3,7-dihydroimidazo[1,2-a]pyrazin-3(7H)one; luminescence at 380 nm), a chemical derived from the luciferin of Cypridina, and luminol (see 3.5.5) have been used for detection of O2• from plant material. Lucigenin. Lucigenin has been used with suspension cultures of rose [342]. Lucigenin itself is capable of producing considerable amounts of O2• via redox cycling especially when used in a concentration exceeding 10 µM; unfortunately high concentrations are required for the detection of O2• [343,344]. Furthermore, lucigenin can also be reduced by NAD(P)H reductases [345]. Luminol and L-012. Luminol, better known as H2O2 detector substance (see e.g. [346]), has also been used to measure O2• from potato tuber discs [347], Vicia faba roots [348] and from tobacco leaves [349]. Luminol [350], and its analog 8-amino-5-chloro-7-phenylpyrido[3,4-d]pyridazine1,4(2H,3H)dione (L-012) [351] might also generate O2• [346]. All of these probes, lucigenin, luminol and L-012, also react with H2O2 and peroxynitrite, L-012 being the most insensitive towards H2O2 but having remarkable reactivity with peroxynitrite [209]. CLA. CLA has been used to detect O2• from tobacco cell suspension [352] and from the epidermal layer of Vicia faba leaves [353]. In addition to O2•, CLA has been reported to react with 1O2, and to some extent also with HO• and H2O2 [354]. CLA has even been used as a 1O2 probe (for example [354]). Addition of SOD suppressed the CLA signal completely in plant leaves [353], except for an initial spike.
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MCLA. Several CLA-derived O2• probes are commercially available, including 2-methyl-6-(4methoxyphenyl)-3,7-dihydroimidazo[1,2-a]pyrazin-3(7H)-one (MCLA). It has been reported that MCLA does not react with H2O2 or O3 but reacts with HO• and 1O2 [355]. Godrant et al. [356] measured extracellular production of O2• by cyanobacteria Trichodesmium erythraeum and concluded that at PPFD 50‒100 µmol m-2 s-1 there was no interference from 1O2, nor did addition of 100 nM H2O2 affect the signal, though some signal from blank was observed as well as quite remarkable amount of a SOD non-inhibitable signal. MCLA has also been used to measure the (extracellular) production of O2• by immobilized marine diatoms [357].
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4.5.6. Microelectrodes With a microelectrode (0.02 cm2) consisting of Cu,Zn-SOD immobilized on nanostructured ZnO disks Deng et al. [358] measured in vivo O2• production from bean sprouts. Direct electron transfer between SOD and the electrode was achieved, with detection limit of 0.2 µM. No interference from H2O2, O2, uric acid, ascorbic acid or 3,4-dihydroxyphenylacetic acid was detected. O2• has been detected in vitro from tomato cell suspension with a Pt electrode in which SOD was immobilized within carboxymethylcellulose-gelatin [359,360].
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4.5.7. Genetically encoded probes cpYFP. Recently a new fluorescent O2• probe, Mito-cpYFP, was presented [361]. It is a circularly permuted yellow fluorescent protein that was targeted to mitochondria of human osteosarcoma cells using the cytochrome C oxidase subunit IV targeting sequence, and it was reported to react only with O2•, not with other ROS, nor with Ca2+, ATP, ADP, NAD(P)+ or NAD(P)H [361]. The probe was also used to detect mitochondrial O2• production in Arabidopsis thaliana [362]. However, Schwarzländer et al. [363] reported that in the mitochondria of Arabidopsis thaliana the probe only responded to changes in pH and not to changes in O2• level. Furthermore, purified cpYFP appeared not to respond to O2• [364]. 4.5.8. Indirect methods of O2• measurement Gene expression. The transcriptomic response of Arabidopsis thaliana [136] and the dinoflagellate Pyrocystis lunula [365] to O2• has been published, but this knowledge has not been used for O2• detection. 4.5.9. Summary of O2• detection As almost all the methods used to measure O2• have flaws, reliable results can be best obtained by using at least two parallel methods [276,313,366]. Furthermore, use of SOD or other agents to confirm specificity to O2• is recommended, although the complex chemistry of some probes (NBT, luminol and lucigenin) may make the evaluation more difficult (see [346,367]). In particular, SOD cannot very well quench O2• produced inside the membranes (e.g. [282]), and the probes themselves may also generate O2• that is quenched by SOD. Different localization of different chemicals should be taken into account when comparing results obtained with different probes [278].
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Table 5. Chemicals used for the detection of O2•. The stability data indicate the stability of the signal after reaction with O2• in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data. Method
Specificity
Stability (t1/2)
DMPO (5,5-dimethyl-1pyrroline N-oxide)
EPR
Also reacts with HO• and other radicals [273,285‒287]
45 s [286,287,289]
DEPMPO (5(diethoxyphosphoryl)-5methyl-1-pyrroline-Noxide) EMPO (5(ethoxycarbonyl)-5methyl-1-pyrroline Noxide) BMPO (5-tertbutoxycarbonyl-5-methyl1-pyrroline N-oxide)
EPR
Also reacts with HO• and other radicals [273,285‒287]
8 min [286,287,289]
EPR
Also racts with HO• and other radicals [273,285‒287]
14 min [286,287,289]
Slower growth of animal cells at 25 mM [297]
EPR
Also reacts with HO• and other radicals [273,285‒287]
23 min [292]
TMT (1-hydroxy-4isobutyramido-2,2,6,6tetramethylpiperidinium)
EPR
Also reacts with HO• and other radicals [273,285‒287]
DCP (probe 1-hydroxy2,2,5,5tetramethylpyrrolidine-
EPR
-
4 h, reduced by photosyntheti c electron transport [278] 4h
Hydrophobici Localization ty (LogP) 0.34 (estimated)
Slower growth of animal cells at 25 mM [297]; No effect on photosynthesis [96] Slower growth of 0.47 animal cells at 25 (estimated) mM [297]
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Toxicity
CR
Compound (IUPAC name)
Notes Spin adduct decays to HO• spin adduct and to HO• [294], UV-light sensitive [295]
-
Spin adduct may decay slowly to HO• spin adduct [269,287,289]
0.41 (estimated)
-
Spin adduct may decay slowly to HO• spin adduct [287,289]
Death of animal cells after long incubation, slower growth at 25 mM [297] No effect on reduction of P700+ [278]
0.86 (estimated)
-
-
1.5 [279] (experiment al)
-
-
No effect on reduction of P700+ [278]
-1.3 [279] (experiment al)
-
-
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Epinephrine
-
-
EPR
No reaction with HO•, ROO, H2O2, NO, GSH or L-ascorbate [288] Reduced by P680, and CO2- [83,308]
Stable [288]
T
EPR
Long incubation slows root growth [296] -
Absorbance at 550 nm
Absorbance at 490 nm
Sugars and phenolics reduce [311,312]
Reacts with ascorbate, GSH, plastoquinone, other components of photosynthetic ETC, oxidases and peroxidases [315‒317] Reacts with H2O2 [322]
US -
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Absorbance at 550‒560 nm, histochemical staining Absorbance at 470 nm, histochemical staining
CR
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15 min [280]
Stable
TE D
XTT (Sodium,3*-(1[phenylamino-carbonyl]3,4- tetrazolium)-bis(4methoxy-6-nitro benzenesulfonic acid hydrate) Cytochrome c (Cyt c)
-
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NBT (Nitro blue tetrazolium)
EPR
AC
3,4-dicarboxylic acid) Tiron (trap 4,5-dihydroxy1,3-benzene-disulfonic acid disodium salt) OXANOH (2-ethyl-1hydroxy-2,5,5-trimethyl3-oxazolidine) PTM-TC (perchlorotriphenylmethy l radical-tricarboxylic acid)
-0.15 (estimated)
-
0.97 (estimated)
Cell permeable
Have been used in vivo in roots, but detection in vitro [296] Interference from metal-ions [282]
Lipophilic, soluble to water at 1 mM [284] 3.49 (estimated)
Apoplast [284]
Has been used in vivo in roots [284]
Cell permeable
-
-
-
1.95 (estimated)
Cell permeable
-
Reaction product can be oxidized by cell components or peroxynitrite [317] Reaction product metabolized
-
0.30 (estimated)
-
Comparison with and without added SOD is needed, enhanced specificity by the use of acetylated Cyt c [263]
-
-1.37 [19]
-
Reaction product may produce O2• [324]
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Absorbance at 530 nm
-
HE or DHE (Hydroethidine)
Fluorescence (Ex 490, Em 520 nm)
Reacts with cytochromes, hemeproteins, Mnporphyrin complex, hypochlorous acid, NO and ONOO[332‒334] Reacts with H2O2 [209,345]
-1.23 (estimated)
Cell permeable [327]
4.03 (estimated)
Cellpermeable, Mito-HE localizes in mitochondri a
Can be reduced by NAD(P)H reductases [345] -
-
3.37 (estimated)
-
Produces O2• [343,344]
-
-0.41 (estimated)
-
May produce O2• [346]
-
-
1.10 (estimated)
-
M0ay produce O2• [346]
-
-
2.02 (estimated)
-
-
-
-
2.19
-
-
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-
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CE P
Lucigenin (N,N′ -Dimethyl- Luminescenc 9,9′ -biacridinium e at 470 nm dinitrate)
Hydroxylamine inhibits oxygen evolving complex of PSII [327,328] Reaction product toxic to E. coli [332]
CR
Hydroxylamine
IP
by Cyt P450 reductase [323] -
T
44
Luminescenc e at 430 nm
L-012 (8-amino-5-chloro7-phenylpyrido[3,4d]pyridazine1,4(2H,3H)dione)
Luminescenc e at 460 nm
CLA (2-Methyl-6-phenyl3,7-dihydroimidazo[1,2a]pyrazin-3(7H)-one) MCLA (2-methyl-6-(4-
Luminescenc e at 380 nm
For reaction with H2O2, see 3.5.5; reacts with peroxynitrite [209] Reacts with H2O2 in the presence of peroxidase and with peroxynitrite [209,345] Reacts with 1O2, H2O2 and HO• [354]
Luminescenc
More specific than
AC
Luminol (5-amino-2,3dihydro-1,4phthalazinedione)
SOD may be needed to exclude unspecific reactions, produce O2• at the pH 8.5 or above [325] HPLC distinguishes specific and unspecific reaction products [292]; oxidized in UV and VIS light, enhances dismutation of O2• [332]
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CLA, reacts with 1O2 and HO• [355]
-
89.8% of sensitivity retained for 80 days [359]
-
-
-
-
-
-
-
Responds to pH, not to O2[364]
CR
-
TE D
-
-
US
Mitochondria targeted fluorescent (Ex 488, Em 505 nm) protein
-
MA N
Signal decreases less than 10 % in 6 days [358]
IP
T
(estimated)
Amperometry No interference by H2O2, O2, uric acid, ascorbic acid or 3,4dihydroxyphenylaceti c acid [358] Amperometry No response with 0.5 mM H2O2 [359]
CE P
CMC–G–SOD (Microelectrode with SOD on carboxymethylcellulosegelatin) Mito-cpYFP
e at 465 nm
AC
methoxyphenyl)-3,7dihydroimidazo[1,2a]pyrazin-3(7H)-one) Microelectrode with SOD on ZnO
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5. Hydroxyl radical, HO• 5.1. Definition and properties of HO•
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Hydroxyl radical (HO•), the result of homolytic cleavage of water (H2O --> HO• + H•), is the most reactive ROS. HO• is a very powerful oxidant; E0 of the pair HO•/H2O is +2.3 V [368]. The rate constants of reactions of HO• with many molecules range from 109 to 1010 M-1 s-1 [369,370]. HO• has one unpaired electron and is able to accept an electron from almost any molecule in its immediate vicinity [371].
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5.2. Formation and action of HO• in photosynthetic tissues
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Although HO• is known to be important especially in the chloroplast, HO• has received relatively little attention in plant biology. In biological systems the main HO• producing reaction is the metal-catalyzed Haber-Weiss reaction consisting of the Fenton reaction (reaction 15) and reduction of the transition metal ion by O2• (reaction 33) (Chapter 2; [1,372]). In spite of its high reactivity HO• plays a role in defense reactions in mammalian cells [373,374].
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Plants do not have specific scavengers for HO•, but the scavenging systems for H2O2 and O2• may protect against formation of HO• by limiting the Fenton reaction (reaction 15). It is assumed that plants prevent excess HO• production and subsequent damage by maintaining low levels of transition metals and sufficient levels of antioxidants in cellular compartments that might contain HO• [375]. The mechanisms maintaining appropriate levels of transition metals may also enable plants to target certain areas for controlled destruction [375]. For example, controlled destruction of cell wall pectins and xyloglucans for germination and elongative growth is believed to require HO• [376,377]. In the cell wall, HO• production has been proposed to involve soluble/cell wall bound proteins such as class III haemcontaining peroxidases (e.g. [372,378]), possibly in conjunction with cell wall associated SODs [379]. NADPH oxidases may also produce extracellular HO• [275,380]. Copper ions are readily available in cell walls and may catalyze Fenton chemistry, especially in roots [378,381]. Chloroplasts are likely the main intracellular sites of HO• production in plants due to the presence of transition metals (Fe, Cu, Mn) that are essential for the function of many transition metal binding proteins of the photosynthetic electron transfer chain [375,382]. Within chloroplasts the acceptor side of PSI is the principal site of H2O2 and O2• production [383,384], and therefore also HO• is thought to be mainly formed at PSI through the iron-catalyzed Haber-Weiss reaction (reactions 13‒15 and 33; [385]). However, with PSII being the main site for molecular oxygen evolution, also the role of PSII in HO• formation has been studied (reviewed by [2]). Formation of HO• at the acceptor side of PSII is thought to involve the reduction of metal center bound peroxides, such as non-heme iron bound peroxide. This reaction is similar to the Fenton reaction except that the reduction of the bound peroxide proceeds as an inner sphere electron transfer reaction [2,386]. Another possibility for HO• production at the acceptor side of PSII is the reduction of free peroxides in a typical Fenton reaction, with detached Mn2+ from the oxygen evolving complex of PSII serving as a possible reservoir of transition metal ions [386]. The oxygen evolving complex may also promote formation of HO• by acting as a source of H2O2 [387], especially after heat induced damage [388]. Yamashita et al. [388] proposed that HO• could be, along with 1O2, one
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of the major ROS causing damage to PSII, particularly under stress. Also components of the mitochondrial electron transfer chain can produce HO•, but in plants the mitochondrial contribution is thought to consist mainly of production of H2O2 that can leak to other cell compartments to yield HO• in Fenton chemistry [375,389].
5.3. Reactions of HO•
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HO• participates in several typical reactions:
abstraction of hydrogen atom with formation of H2O and radical of substrate (36);
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HO• + RH H2O + R•
(36)
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addition to double bonds with formation of a hydroxylated radical (37);
(37)
TE
AC CE P
HO• + SCN + OH + SCN•
D
electron transfer reactions leading to formation of a neutral radical (38.1) or a cation radical [390] (38.2); (38.1)
(38.2)
formation of aromatic-OH adduct due to a reaction of an aromatic compound with HO• is one of the methods for HO• detection with HPLC-MS. For example, HO• can react with phenylalanine to form isomers of tyrosine, reaction (39) [391]. Isomers of tyrosine are rather stable and not normally present in proteins and can serve as HO• traps in biological samples [392].
(39)
ACCEPTED MANUSCRIPT 48 Interaction of HO• with many aquated metal cations is an electron transfer reaction (40) that proceeds via two different mechanisms, both with a rate constant of ~108 M-1 s-1 [369]. HO• + Mn+ M(n+1)+ + OH
(40)
(41)
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HO• initiates lipid peroxidation resulting in hydrogen abstraction from a pentyl group of an unsaturated fatty acid and formation of a radical that interacts with 3O2 to form a peroxyl radical (ROO•) with a rate constant of ~108 M-1 s-1 [393], reaction (41).
5.4. Lifetime and diffusion distance of HO• The rates of the reactions of HO• with biomolecules are usually limited by diffusion, and therefore reactions of HO• with biomolecules take place close to the site of HO• generation. The lifetime of HO• is thought to be in the order of nanoseconds [394,395], and recent simulations yielded a lifetime of only 30 ps in aqueous environment [371]. The diffusion distance of HO• has been estimated to be only a few molecular diameters from the place of origin [395]. The lifetime of HO• has not been measured in vivo.
5.5. Detection methods of HO• 5.5.1 Spectroscopic methods DMSO. Dimethyl sulfoxide (DMSO) has been used to detect HO• by colorimetry (absorbance at 420 nm) of methanesulfinic acid (MSA) formed in the oxidation of DMSO by HO• [396]. DMSO is only weakly toxic [397,398] and MSA is stable at pH 5‒9 [396]. Using DMSO, HO• formation has been witnessed in bean cowpea nodules, cucumber seedlings [399] and in leaves of duckweed and ryegrass [398]. In DMSO
ACCEPTED MANUSCRIPT 49 trapping, MSA must be extracted from the samples before colorimetry because a reaction between diazonium salts and MSA is required for the colorimetric assay [396‒400].
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Deoxyribose degradation. Free deoxyribose sugars yield malondialdehyde (MDA) upon degradation by HO• [401], and this observation has been utilized in HO• detection via detection of TBA-reactive substances (TBARS, see 6.4.5) [402]. Use of deoxyribose sugars as HO• detectors has been criticized for both its accuracy and its utilization in biological systems where free deoxyribose is not abundant [402]. In plant sciences this method has nevertheless been used to detect HO• formation in soybean leaves [403] and in wall fractions of isolated onion bulb cells [404].
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5.5.2. EPR detectable probes DMPO, DEPMPO, EMPO, 4-POBN. EPR spectroscopy is the most widely used method for detection of HO• in photosynthetic materials [2,269,375]. EPR probes that are used for HO• detection (DMPO, DEPMPO, EMPO) form adducts with O2• as well (see 4.5.2; [269]), but the EPR spectra of O2• and HO• probe adducts are so distinct that they allow differentiation between the two ROS, especially in the case of DMPO and EMPO. DEPMPO adduct spectra are more difficult to analyze [269,405]. Furthermore, HO• adducts produce diamagnetic species with Fe(III) and O2• and decay rapidly; both reactions result in loss of the EPR signal [152]. So even though HO• adducts can be distinguished, the actual amount of HO• being formed in a system may not be obtained with simplistic analyses of the EPR signal [269]. Taking into account the fact that decay of e.g. DMPO/O2• adducts yields HO• and that different probes act as scavengers of many ROS [269], special care needs to be taken in planning experiments where one and the same spin trap probe is used for the detection of O2• and HO•, as the naturally occurring formation of HO• is tightly linked to the concentrations of free O2•, free peroxides and transition metals [372,406]. Additional steps to confirm the precursors of HO• production in a studied system can be taken by adding SOD and CAT individually or together into the spin trapping mixture, thus pinpointing the prevalent HO• production pathway [407]. Also chelates such as ethylenediamine-N,N,N’,N’-tetraacetic acid (EDTA), diethylenetriamine-N,N,N¢,N’,N’’-pentaacetic acid (DTPA) and deferoxamine mesylate (Desferal) are commonly added into the buffers used in OH• measurements in order to reduce the amount of free transition metals, but use of these chelators should also be considered carefully as they themselves can stimulate ROS production [408,409]. 4-pyridyl-1-oxide-N-tert-butylnitrone (4-POBN) is an EPR probe that is used more spesifically for HO• detection. 4-POBN also reacts with O2• but the adduct is so unstable compared to a HO• adduct that almost no O2• adducts are detected [269]. In concentrations that are commonly used in biological studies (50 mM), 4-POBN can react with H2O2 in a reaction involving peroxidases, yielding 4-POBN•/4POBN spin adduct [410]. In addition to the specificity of 4-POBN, adducts produced in typical experiments with 4-POBN and HO• are considered to be stable when compared to other HO• reactive EPR probes [269,411]. The stability of 4-POBN adducts is, however, a result from the common practice of using HO• scavenging agents such as DMSO, ethanol or formate in conjunction with HO• probes [269,407,412‒415]. These scavengers react with HO• to form carbon centered radicals, such as αhydroxyethyl in the case of ethanol [413], which in turn form stable adducts with the spin trap. In the case of 4-POBN, the HO• adduct is stable because 4-POBN is practically always used with ethanol and the
ACCEPTED MANUSCRIPT 50
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detected product is an adduct of 4-POBN and a carbon-centered hydroxyethyl radical [416‒420]. It is important to take the additional effects of ethanol (from 170 to 850 mM) into account when planning spin trap experiments with 4-POBN [269]. The half-life of the actual 4-POBN/HO• adduct has been shown to be only 1 min in Hank’s Buffer Solution, much shorter than the half-life of the DMPO/HO• adduct (2 h) in almost the same conditions [421]. The life time estimates for different spin trap adducts derived from organic chemistry (e.g. [152]) can be difficult to translate into biological context, and therefore EPR measurements of spin adducts should be done as fast as possible.
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Although there are many issues concerning in vivo detection of HO•, mainly the very short lifetime of HO• and the yet uncharacterized production pathways of other interfering ROS [375,415], EPR spectroscopy has been used in vivo in plants. In vivo measurements of HO• directly from tissues infiltrated with 4-POBN have been done by Renew et al. [422] and Deng et al. [420] on intact cucumber root and tobacco leaf cutouts, respectively. Measurements done by exposing the intact tissue to the spin trap and then measuring EPR from ground tissue are more numerous. With 4-POBN HO• has been detected from roots [407,418], coleoptiles [416,417], plant radicles and endosperm caps [423] and from potato tubers [419]. DMPO, in turn, has been used to detect HO• from roots by Demidchik et al. [424].
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In vitro HO• has been detected with DMPO from isolated PSII membranes [285,425] and from isolated rapeseed hypocotyl tissue [414]. DEPMPO [387,426‒428], EMPO [388,429,430] and 4-POBN [386,387] have also been used in isolated PSII membranes. DMPO has been utilized in many photosynthesis studies to examine HO• formation in isolated PSII antenna complexes [431] and isolated thylakoid membranes [97,432]. Both DEPMPO [275] and 4-POBN [380] have been used to trap HO• in isolated plant plasma membranes, whereas DEPMPO has been used to measure HO• from pea cell wall isolates [433,434] and from the apoplast of cucumber leaf [435]. 5.5.3. Fluorescent probes Fluorophorenitroxide proxyl fluorescamine (PF). Alternatives to EPR include 5-((2-carboxy)phenyl)-5hydroxy-1-((2,2,5,5-tetramethyl-1-oxypyrrolidin-3-yl)methyl)-3-phenyl-2-pyrrolin-4-one sodium salt (fluorophorenitroxide proxyl fluorescamine, PF) [421,436]. PF works as a spin trap for HO• in a similar manner as 4-POBN. A reaction of PF with the methyl radical produced in a reaction between HO• and DMSO eliminates the fluorescence quenching nitroxide group of PF [421]. Other radicals such as O2• do react with PF, but with the use of DMSO the probe is described by [421] as HO• specific. The excitation/emission wavelengths of PF are 380/485 nm. HO• formation has been fluorometrically detected from aliquots of PF media used for experimental incubations of barley root tip segments [436]. The use of UV wavelengths for excitation limits the use of PF in biological material. Terephthalic acid. Liu et al. [437] detected the formation of HO• by benzene-1,4-dicarboxylic acid (terepthalic acid, TPA) in supernatant of pea pod cell wall homogenate. TPA traps HO• and forms monohydroxy terephthalate (TPA-OH) that fluoresces at 350‒550 nm with excitation at 362 nm [437]. While the signal producing reaction of TPA is HO• specific and the formed adduct is relatively stable [437,438], TPA is known to react with other radicals without producing a fluorescent adduct [439]. UV absorption by biological materials may limit the use of TPA as well.
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5.5.4. Microelectrode An electronic HO• microsensor, based on a change in the conductivity of polyaniline occurring due to a reaction with HO• was recently published [440], though it has not yet been used with photosynthetic material.
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5.5.5. Summary of HO• detection Because of its very short lifetime and unspecific reactivity, HO• is a difficult research subject and its function remains easily overshadowed by other ROS [441]. Furthermore, conclusions drawn from the physiological responses to H2O2 might be misleading due to the possible formation of HO• from H2O2 by the Fenton reaction [442]. Most methods used to measure HO• from plant material are not very specific (Table 6). Many methods relying on absorption/emission of HO• specific detector molecules are difficult to use in plants because the absorbance and fluorescence spectra of photosynthetic tissues often overlap strongly with the wavelengths of the signal.
ACCEPTED MANUSCRIPT 52
Specificity
Stability (t1/2)
Absorbance at 420 nm
HO• specific [396]
Stable [396]
Deoxyribose (2Deoxy-D-ribose) DMPO (5,5-dimethyl1-pyrroline N-oxide)
Absorbance at 530 nm EPR
Not very specific [402] Reacts also with O2• and other radicals [413]
-
Toxicity
RI
Method
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Compound (IUPAC name) DMSO
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Table 6. Chemicals used for the detection of HO•. The stability data indicate the stability of the signal after reaction with HO• in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data.
AC
CE DEPMPO (5EPR (diethoxyphosphoryl)5-methyl-1-pyrrolineN-oxide)
Reacts also with O2• and other radicals [286]
Stable [286]
EMPO (5(ethoxycarbonyl)-5methyl-1-pyrroline Noxide) 4-POBN (4-pyridyl-1oxide-N-tert-
EPR
Reacts also with O2• and other radicals [287]
-
EPR
HO• specific [269]
HO• adduct:
Not very toxic Slower growth of animal cells at 25 mM [297]; Plant root cells viable after treatment [441]; No effect on photosynthesis [96] Slower growth of animal cells at 25 mM [297]; No effect on photosynthesis [427]
PT ED
2 h [413]
Not very toxic [397]
αhydroxye
Hydrophobicity (LogP) -1.35 [19]
-1.90 (estimated) 0.34 (estimated)
Notes Optimal pH 5‒9; detected molecule is methanesulfic acid [396] Can be used to detect carbon centered radicals [413]
0.47 (estimated)
Can be used to detect carbon centered radicals [286]
Slower growth of animal cells at 25 mM [297]
0.41 (estimated)
-
-0.20 (estimated)
Can be used to detect carbon centered radicals [287] Commonly used with ETOH
ACCEPTED MANUSCRIPT 53
Fluorescence (Ex 380, Em 485 nm)
Reacts also with O2• [421]
Fluorescence (Ex 362, Em 350‒550 nm)
PANI electrode based on conductance change of polyaniline
Conductance measurement
May react with other radicals, but signal is HO• specific [439] Specific to HO• [440]
PT
[413] H 7 optimal for HO• adduct [411]
RI
thyl radical adduct: stable for >8 min [413]
-
-
Can be used to detect carbon centered radicals [436]
Stable [438]
-
2.00 [19]
-
Stable [440]
-
-
-
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PT ED
MA
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Fluorophorenitroxide proxyl fluorescamine (5-((2carboxy)phenyl)-5hydroxy-1-((2,2,5,5tetramethyl-1oxypyrrolidin-3yl)methyl)-3-phenyl2-pyrrolin-4-one sodium salt) Terephthalic acid (Benzene-1,4dicarboxylic acid)
With O2• less than 1 min [413]; Without O2• 32 h [411] -
SC
butylnitrone)
ACCEPTED MANUSCRIPT 54
6. Peroxyl radical (ROO•), alkoxyl radical (RO•) and hydroperoxides 6.1. Definition and formation
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Peroxyl and alkoxyl radicals and hydroperoxides are best known as products of peroxidation of lipids. Other ROS, especially 1O2 and HO•, initiate reactions that lead to formation of these ROS.
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6.2. Reactions of ROO•, RO• and hydroperoxides
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The appearance of ROO• leads to formation of an alkoxyl radical (RO•) and lipid hydroperoxides (ROOH). The scheme of reactions leading to the appearance of these forms and, consequently, lipid peroxidation is well known, reactions (42‒44).
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HO• + RH R• + H2O
(42)
Initiation of the process and the appearance of lipid radicals;
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R• + 3O2 ROO•
(43)
addition of molecular oxygen to the carbon-centered radical (R•) yielding the ROO•;
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ROO• + RH ROOH + R•
(44)
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abstraction of a hydrogen from another lipid molecule by ROO•, like reaction (42). Reaction (44) is known as the chain propagation reaction because this reaction supplies R• for reaction (43) and thus leads to accumulation of ROOH. The spontaneous decomposition of ROOH (reactions 45 and 46) is unfavorable because the dissociation energies of ROO-H and RO-OH bonds are 376 kJ/mol and 184 kJ/mol, respectively [443]. Spontaneous decomposition of ROOH can be observed in perturbed conditions, e.g. at a high temperature. ROOH ROO• + H•
(45)
ROOH RO• + HO•
(46)
Non-enzymatic decomposition ROOH via either their reduction or oxidation by transition metal is one of the most significant ways of ROOH decomposition (reactions 47.1 and 47.2). ROOH + Mn+ RO• + OH + M(n+1)+
(47.1)
ROOH + M(n+1)+ ROO• + H+ + Mn+
(47.2)
Interaction of ROOH with radicals can also lead to decomposition of ROOH due to abstraction of a hydrogen atom and formation of ROO• (reaction 48). ROOH + R• ROO• + RH
(48)
ROOH can also react with the C-H bond of saturated and alkylaromatic hydrocarbon via reaction (49) [444].
ACCEPTED MANUSCRIPT 55 ROOH + RH RO• + R• + H2O2
(49)
Dismutation of ROOH yields both RO• and ROO•, reaction (50). ROOH + ROOH RO• + ROO• + H2O
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(50)
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Decomposition of ROOH is a key reaction in the chain reaction of lipid peroxidation because of additional formation of ROO• or RO• species that can abstract a hydrogen from a lipid molecule, reactions (44) and (51). ROO• + RH ROOH + R•
(51)
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The redox potentials (E0' at pH 7) of pairs RO•/ROOH and ROO•/ROOH are 1.6 V and from 0.77 V to 1.44 V, respectively [368]. The rate constant of the reaction of RO• with RH is 107 M-1 s-1, which is five orders higher than the rate constant of the reaction of ROO• with RH, 102 M-1 s-1 [168]. R•, ROO• and RO• can also interact with each other in several reactions [445,446]:
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The radical-radical reaction between two RO• can lead to formation of a carbonyl compound in excited state, reaction (52); RO• + RO• ROH + R=O*
(52)
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Reaction 52 is rather unlikely in biological systems because of the high reactivity of RO• toward lipids and proteins. A radical-radical reaction between two ROO• leads to the formation of an excited carbonyl compound (reaction 53.1) or 1O2 (reaction 53.2). The latter is called the Russell mechanism. The rate constant for the termination stage in the Russell reaction varies between 106 and 108 M-1 s-1 [445].
(53.1)
(53.2)
RO• and ROO• can react with R• to form dimers, reactions (54) and (55) [443,447]. RO• + R• ROR
(54)
ROO• + R• ROOR
(55)
Numerous experimental data on the kinetics of hydroperoxide chain process is represented in a recent monograph [444].
ACCEPTED MANUSCRIPT 56 Reactions (52) and (53.1) are associated with chemiluminescence (380–460 nm) due to relaxation of the excited carbonyl compound. This chemiluminescence is used to measure lipid peroxidation [448] (see 6.4.2).
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In reactions with lipid molecules, RO• can cause a homolytical scission (β-cleavage) of a C-C bond on either side of the alkoxyl group to form a wide range of aldehydes and oxo fatty acids [449]. Formaldehyde, acetaldehyde, propionaldehyde, acrolein, malondialdehyde, (E)-2-hexenal, (Z)-3-hexenal, 4-hydroxy-(E)-2-hexenal and other carbonyls are found to be products of degradation of polyunsaturated fatty acids [450], and therefore aldehydes can be used as markers of lipid peroxidation. Measurement of coloured substances produced in a reaction between aldehydes and thiobarbituric acid (TBA) is the most commonly applied method [451]. A new method for a comprehensive analysis of oxylipin carbonyls (quantitation and structural estimation) in plants with reverse-phase HPLC has recently been established [450,452].
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Reduction of ROOH by thiol groups of peroxiredoxins is an important scavenging mechanism. This reaction proceeds as a 2-step reaction like reaction with H2O2, reactions (55.1) and (55.2). The Km values and rate constants of the reaction between peroxiredoxins and ROOHs depend on both ROOH species and on the peroxiredoxin, and the Km values range from 40 nM to 1.7 mM and the rate constants from 104 M-1 s-1 to 107 M-1 s-1 [453].
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ROOH + RSH RSOH + ROH
(55.2)
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RSOH + RSH RSSR + H2O
(55.1)
Tocopherols and carotenoids can act as antioxidants preventing lipid peroxidation [454,455].
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6.3. Lifetime and diffusion distance of ROOHs The lifetimes of ROO•or RO• in the cells have been estimated to be 7 s and 1 µs, respectively [395]; these lifetimes would allow 5 % of the original ROS to be present at 0.16 mm or 60 nm from the site of formation in a viscous cellular environment, respectively. ROOHs are generally less reactive than ROO•or RO•.
6.4. Detection methods of ROOHs 6.4.1. Spectroscopic methods Lipid hydroperoxides contain fatty acids with conjugated dienes, and lipid hydroperoxides can be detected spectroscopically by measuring the absorbance of the dienes at 230‒235 nm. A high sensitivity for conjugated dienes is achieved with double-derivative spectroscopy [456]. However, this method can only be used for pure extracted material. 6.4.2. Chemiluminescence and thermoluminescence Excited carbonyl compounds, produced due to degradation of ROOHs, emit light at 380–460 nm. The chemiluminescence method (see Table 7 for a list of chemical methods for ROOH detection) can be applied for detection of ROOHs under both spontaneous and catalytic degradation of ROOH. ROOH-derived chemiluminescence has been measured from plant leaves with a sensitive camera [446]. ROOH-derived chemiluminescence resulting from sodium hypochlorite-induced degradation of ROOH was proposed as a sensitive assay for ROOH [457]. In plant material, addition of acridine in reaction medium significantly improved the linear correlation between luminescence intensity and amount of ROOHs [458]. A high-
ACCEPTED MANUSCRIPT 57 temperature thermoluminescence band peaking at 115‒130 °C was shown to correlate with the amount of lipid peroxides in chlorophyll-containing material [459]. The method has been frequently used for the detection of lipid peroxides and assessment of oxidative stress especially in photosynthesis research; see e.g. [460,461].
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6.4.3. Dyes Xylenol orange. Xylenol orange, used as dye for detection of ROOH is know as the ferrous xylenol orange method (FOX method). The FOX method is based on the oxidation of ferrous ions Fe(II) to ferric ion Fe(III) by ROOH in acidic conditions. The reaction of ferric ion with xylenol orange results in formation of a complex with an absorption maximum at 560 nm [462‒464]. Unfortunately, the stoichiometry between the amount of the xylenol orange complex and the amount of peroxide depends on peroxide species, ranging from 1.8 to 5.4 moles of xylenol orange complex per 1 mole of peroxide [465]. Therefore, a separate calibration curve is needed for each type of peroxide. The FOX method can be applied for both aqueous and organic solvents and used for detection of both hydrophilic (FOX1 method) and lipophilic (FOX2 method) peroxides. The FOX1 and FOX2 methods are not specific to ROOH because ferrous ions can be oxidized by other oxidizing agents. To accurately quantify ROOHs, the measurements are done in the presence and absence of triphenylphosphine which selectively reduces ROOHs to corresponding alcohols. To prevent peroxidation during analysis, butylated hydroxytoluenes are usually used [466,467].
TE
D
Thiocyanate. Thyocyanate (SCN) has also been used for detection of ROOH with the oxidation of ferrous iron. The reaction of ferric ion with the thiocyanate ion leads to formation of complexes absorbing at 470‒505 nm [462]. In contrast to the FOX methods, the thiocyanate-iron method can be used for detection of ROOH in material containing carotenoids [468].
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Iodide. ROOHs oxidize iodide to iodine and iodine react with iodide, yielding triiodide anion (I3-). I3- absorbs at 290 and 360 nm [198,469]. A modified iodometric method has recently been used for determination of ROOH in plants [470]. The iodometric method is sensitive to oxygen that must be removed from the reaction medium. Iodometry is not specific for ROOH because H2O2 can oxidize I- to I2 (see 3.5.3; [471]), and therefore H2O2 should be removed before measurement. The stoichiometry between amounts of peroxides and iodine is 1:1. This method is known as an International Dairy Federation method (IDF method, 74A:1991). 6.4.4. Fluorescent probes Diphenyl-1-pyrenylphosphine (DPPP). DPPP can be used as a fluorescent probe to monitor lipid peroxidation in cell membranes. DPPP reacts with organic hydroperoxides stoichiometrically to yield DPPP oxide (DPPPNO) which emits at 380 nm with excitation at 351 nm [472]. DPPP readily reacts with lipophilic peroxides like methyl linoleate hydroperoxide but not with tert-butyl hydroperoxide. The DPPP method can be used for measurement of lipid peroxidation within membranes. DPPP also rapidly reacts with H2O2, and therefore this method can only be used in vitro with H2O2 free samples. [3-(4-phenoxyphenylpyrenylphosphino) propyl]triphenylphosphonium iodide (MitoDPPP). The reactivity of MitoDPPP is similar as that of DPPP, and MitoDPPP can be oxidized by peroxides including H2O2 but not by free radicals like O2• or HO•. The oxidation of MitoDPPP with hydroperoxides yields MitoDPPPO which has 35 times higher fluorescence intensity than MitoDPPP (ex 350 nm, em 380 nm) [473]. The reaction rate of MitoDPPP
ACCEPTED MANUSCRIPT 58 with peroxides depends on the type of the peroxide. MitoDPPP is supposed to localize within lipid membranes and can be used for detection of ROOH in living cells.
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2-(4-diphenylphosphanylphenyl)-9-(1-hexylheptyl)anthra[2,1,9-def,6,5,10-d′ e′ f ′ ]diisoquinoline-1,3,8,10tetraone (Spy-HP). Spy-HP reacts rapidly with hydroperoxides such as m-chloroperbenzoic acid (MCPBA) and cumene hydroperoxide to form an oxidized derivative (Spy-HPO) that emits strong fluorescence at 535 nm when excitated at 524 nm. The formation of Spy-HPO shows strict 1:1 stoichiometry with the amount of peroxides and its affinity to lipophilic hydroperoxides is high, allowing measurements at nM concentration. Spy-HPO is stable against autoxidation and photobleaching and the reaction of Spy-HP with O2•, H2O2 and HO• is very slow compared to the reaction with lipophilic peroxides [474]. Spy-HP has been used to detect lipid peroxides in photosynthetic membranes [475].
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2-(4-diphenylphosphanyl-phenyl)-9-(3,6,9,12-tetraoxatridecyl)-anthra[2,1,9-def:6,5,10-d9e9f9]diisoquinoline1,3,8,10-tetraone (Liperfluo). Liperfluo is oxidized by peroxides to Liperfluo-Ox with (ex 524 nm, em 535 nm). Liperfluo reacts much more slowly with various other ROS including H2O2, O2•, HO• and NO• than with methyl linoleate hydroperoxide [476]. Liperfluo shows only little cytotoxicity in SH-SY5Y cells and can be used for detection of peroxides in cells. It was demonstrated that Liperfluo is more useful as a probe than Spy-LHP for live-cell fluorescence imaging [476]. The fluorescence yield of Liperfluo-Ox is linearly dependent on the concentration of methyl linoleate hydroperoxide.
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Mito-Bodipy-TOH is a fluorescent peroxyl radical probe absorbing at 537 nm and emitting at 572 nm. The probe consists of an α-tocopheryl moiety, a fluorogenic reporter and a mitochondria-targeting segment by which the probe targets to the inner mitochondrial membrane [477]. The probe reacts with lipid peroxides but its specificity has not been extensively studied. Mito-Bodipy-TOH has not been used with photosynthetic material. 6.4.5. Measurements of ROOH-derived species TBA reactive substances (TBARS). Measurements of TBARS are commonly used for lipid peroxidation assays. The method is based on the ability of ROOH-derived products like malondialdehyde to react with TBA to form TBA-adducts at an elevated temperature in acidic conditions. TBA-adducts can be easily detected by absorbance at 532 nm [478,479] or by fluorescence at 553 nm [480]. Other ROOH-derived aldehydes can also react with TBA to form complexes [481]. Decomposition of lipid peroxides to TBARS during analysis results in formation of radicals that can promote additional lipid peroxidation, which may lead to overestimation of ROOH content. Protective substances (like butylated hydroxytoluene) are usually used to inhibit the lipid peroxidation [482,483]. Traces of iron (II) can also increase TBARS by accelerating decomposition of ROOHs, and therefore chelators are added to the medium [483]. HPLC analysis of TBA-adducts allows measurement of the MDA-(TBA)2-adduct only [484]. Using gas chromatography-mass spectrometry (GC-MS) it has recently been shown that lipid peroxidation is accurately measured with the TBA method in most plants [485]. 2,4-dinitrophenylhydrazine (DNPH). This method is based on extraction of lipid-derived carbonyls and their derivatization with 2,4-dinitrophenylhydrazine and separation with HPLC and LC/MS [452]. The carbonyl species can be identified from the MS/MS spectrum. The structure of ROOH-derived carbonyls is determined from the position of the hydroperoxide group in the lipid molecule. A HPLC based comprehensive analysis of carbonyls using DNPH for derivatization can be used for detection of fatty acids involved in peroxidation [486].
ACCEPTED MANUSCRIPT 59
IP
T
Table 7. Methods of detection of ROOH species. All mentioned chemicals show a stable, non-light-sensitive signal. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data. Method
Specificity
Hydrophobicity (LogP)
Notes
Ferric ion xylenol orange complex (absorption maximum at 560 nm) [462]
1.85 (estimated), can be used for both aqueous and organic solvents [466]
Various stoichiometries between amount of xylenol orange complex and amount of peroxide [465]
Thiocyanate (IDF method)
Ferric ion thiocyanate complex (absorption at 470–505 nm) [462,468]
0.22 (estimated), can be used for both aqueous and organic solvents [462,468]
Stoichiometry between amount of complex and amount of peroxide is not strict [462]
Iodide
Formation I3Absorption at 290–360 nm [198,469] Oxidized form has high fluorescence (Ex 351, Em 380) [472] Oxidized form has high fluorescence (Ex 350, Em 380) [473]
Nonspecific. Triphenylphosphine shuld be used to evaluate unspecificity [463,466] Nonspecific. Triphenylphosphine should be used to evaluate unspecificity [462] -
1.04 [19]
Experiments can be carried out in anaerobic condition dou to sensitivity to oxygen [198,469] -
MitoDPPP ([3-(4phenoxyphenylpyre nylphosphino) propyl]triphenylpho sphonium iodide) Spy-HP (2-(4Oxidized form has high diphenylphosphanyl fluorescence (Ex 524, phenyl)-9-(1Em 535) [474] hexylheptyl)anthra[2 ,1,9-def,6,5,10-
US
MA N
TE D
CE P
Can react with H2O2 [472]
8.52 (estimated), lipophilic [472]
Can react with H2O2 [473]
Lipophilic [473]
-
Specific for lipohilic peroxides. Slowly react with hydrophilic peroxides and H2O2 [474]
Very lipohilic [474]
Useful only for organic solvents [474]
AC
DPPP (Diphenyl-1pyrenylphosphine)
CR
Compound (IUPAC name) Xylenol orange (FOX method)
ACCEPTED MANUSCRIPT
TE D
-
CE P
-
IP
-
US
CR
-
MA N
Specific for lipohilic peroxides. Slowly react with hydrophilic peroxides and H2O2 [476]
AC
d′ e′ f ′ ]diisoquinoline1,3,8,10-tetraone) Liperfluo 2-(4High fluorescence (Ex diphenylphosphanyl- 524, Em 535) [476] phenyl)-9-(3,6,9,12tetraoxatridecyl)anthra[2,1,9def:6,5,10d9e9f9]diisoquinolin e-1,3,8,10-tetraone) TBA Complex with MDA with maximum of absorption at 532 nm [478,479] Mito-Bodipy-TOH Fluorescence (Ex 537, Em 572) [477]
T
60
-0.39 (estimated)
TBA can react with other carbonyls in addition to MDA [481]
-
Targeted for mitochondrial inner membrane. Not yet tested with photosynthetic material.
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7. Ozone, O3
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Ozone is an oxygen molecule consisting of three oxygen atoms (O3). O3 is a strong oxidant (E0 of the couple O3 + 2H+ + 2e-/O2 + H2O is 2.076 V [154]). O3 may also take part in 1-electron reactions, forming the ozonide radical O3-. Thermodynamic calculations suggest a value 1.6 V for the E0 value of the couple O3/O3• [487], although a lower value of 1.0 V could also be suggested [487]. External O3 does not penetrate to living plant cells [488] and therefore detection of O3 will not be discussed. However, external O3 penetrates to the apoplastic space through stomata and causes the formation of other ROS in the apoplast and thereby initiates plant defense signaling [8].
(56)
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The principal reaction of O3 with organic compounds is the addition to double bonds and cleavage of unsaturated compounds (ozonolysis). O3 reacts with a double bond to form ozonide and via the Criegee mechanism, reaction (56) [489].
Alternatively to formation of secondary ozonides, the decomposition of primary ozonides in aqueous medium can yield carbonyls and H2O2. The ozonolysis of ethylene yields two molecules of formaldehyde and one molecule of H2O2, reaction (57). Formation of H2O2 in reactions resembling reaction (57) may contribute to the ozone-induced oxidative burst in plant apoplast [8].
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(57)
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The rate constant of the reaction of O3 with ethylene at 30 °C is 106 M-1 s-1 [490]. O3 can react with isoprene and others terpenes to give corresponding carbonyls [491]. It was also shown that ozonolysis of terpenes as -terpineol, limonene, and -pinene is associated with formation of HO•, reaction (58) [492].
(58)
The reaction of O3 with such amino acids as cysteine, tryptophan, methionine, tyrosine, histidine and other biologically important compounds like ascorbic acid and NADPH produces 1O2. The rate constants and yields of 1O2 formation for many relevant compounds are given by [493]. The yield of 1 O2 in the reaction of O3 with ascorbic acid was found to be pH dependent, indicating that 1O2 is produced in a reaction between O3 and the ascorbate anion, reaction (59) [494]. O3 is able to oxidize GSH to a sulfonic acid [495]. AsCH + O3 DHA + 1O2 + OH,
(59)
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where DHA is dehydroascorbate.
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In addition to the classic ozonide forming reactions, electron transfer to ozone can lead to the formation of an ozonide anion radical, reaction (60). The ozonide anion radical is protonated very fast (k 1010 M-1 s-1), reaction (61) and then decomposed to hydroxyl radical and 3O2 [487,496].
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IrCl63 + O3 IrCl62 + O3 O3 + H+ HO3
(60) (61)
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In the atmosphere, ozone promotes the appearance of nitrogen dioxide (NO2•) via reaction (62) with nitric oxide (NO•).
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O3 + NO• NO2• + O2
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8. Non-specific methods
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8.1. Non-specific methods indicating the presence of ROS
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Most direct ROS detection methods aim at detection and measurement of a specific type of ROS. Specificity is highly important because a method that detects several different ROS, each with different detection efficiency, has little predictive value for ROS-induced damage or ROS signaling. Several non-specific methods are, however, available in the market and they are sometimes used in the absence of specific ones. Non-specific reagents may also have advantages like low cytotoxicity or ability to localize to a specific cell compartment.
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H2DCFDA.The dyes 2′ ,7′ -dichlorodihydrofluorescein diacetate (H2DCFDA) and its derivatives 5-(and6)-chloromethyl-2′ ,7′ -dichlorodihydrofluorescein diacetate (CM-H2DCFDA) diffuse through the cell membrane and cell wall, but within the cell esterases remove the acetate groups and the chloromethyl group reacts with thiols. Therefore the dye cannot diffuse out of the cell, and a fluorescent oxidation product (DCF) can be detected by excitation at 492‒495 nm and emission at 517‒527 nm. CM-H2DCFDA has been used with Arabidopsis thaliana leaves; peeling off the epidermis is required to get the dye into leaf cells, and the dye responds to H2O2 treatment of the cells [23]. The chemical has also been used with cyanobacteria [130]. CM-H2DCFDA has been used as H2O2 detector (e.g. [497]), but because oxidation of CM-H2DCFDA depends on a Fenton-type reaction or on an unspecific enzymatic reaction with cytochrome c [498], a difference in the intensity of the fluorescence signal between two samples may indicate difference in H2O2, iron, copper or cytochrome c concentration. Also, UV and to a smaller degree visible light causes oxidation of DCFH [499], and DCF can produce O2• in light [500]. CellROX reagents. Molecular Probes Inc. produces several unspecific ROS detection reagents: CellROX® Deep Red Reagent with excitation and emission at 640 and 665 nm, respectively, CellROX® Orange Reagent (545/565 nm), CellROX® Green Reagent (485/520 nm) and OxyBurst (R) Green H2DCFDA, SE (2',7'-dichlorodihydrofluorescein diacetate, succinoimidyl ester). According to the website of the company (https://tools.lifetechnologies.com/content/sfs/manuals/mp10422.pdf; cited Feb 24, 2015), the Green Reagent binds to DNA and localizes to the nucleus in animal cells, and
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the OxyBurst Green H2DCFDA, SE penetrates to cells. These reagents have not been tested with photosynthetic material but might be useful, as the emission wavelengths of the Orange and Green reagents do not interfere with Chl a emission. The rate constants of these reactants with different ROS have not been published. The cell-impermeable OxyBurst Green H2HFF BSA reagent has been used for detection of apoplastic ROS production during the generation of systemic signals related to wounding [501].
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HPF and APF. 3'-(p-hydroxyphenyl) fluorescein (HPF) and 3'-(p-aminophenyl) fluorescein (APF) were originally developed for the detection of HO• and peroxynitrite ion (ONOO-) [227]. In both cases a dearylation reaction converts a non-fluorescent dye to a fluorescent one, with excitation at 490 nm and emission at 515 nm. The dyes are cell permeable and resistant to autooxidation [227] and react with most ROS, including 1O2 and ROO• [24,227]. Data about their reactivity against O2• varies [24,227]. The reaction between APF and pure H2O2 is slow but becomes significant in the presence of a peroxidase [24].
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TEMPO-9-AC. 4-((9-acridinecarbonyl)amino)-2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO-9-AC) is a free radical that is converted to a fluorescent form (with excitation at 366 nm and emission at 424 nm) in a reaction with another free radical like O2• [502]. Strong illumination of tobacco leaves after pinhole administration of TEMPO-9-AC did not cause detectable fluorescence [109]. In leaves, this probe remains mostly in the apoplast [109].
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Ultra-weak luminescence, emitted by triplet excited carbonyls, pigments excited by excited carbonyls and by 1O2 has been shown to be associated with oxidative stress in plants, and mechanisms of production of light have been elucidated [503]. Long exposure time is required for quantitative imaging of ultra-weak luminescence.
8.2. Non-specific methods indicating reactions of ROS with biomolecules Proteins, DNA, lipids and various small biomolecules are known to react with ROS in vitro, but conclusive evidence for a reaction occurring in vivo or in an isolated system like thylakoid membrane preparation is difficult to obtain. Oxidation of proteins by ROS typically produces carbonyl groups in the amino acid side chains, and these can be detected with an antibody against 2,4-dinitrophenyl hydrazine (DNP) [504]. DNP-antibodies “Oxyblot™” can be used for general oxidative stress analysis. Reactive carbonyl species (RCS), defined as α,β-unsaturated aldehydes and ketones, are products of further reactions of lipid peroxidation [505]. Lipid peroxidation is initiated by reactive oxygen species, and therefore an increase in RCS, detected by the TBARS test, indirectly indicates an action of ROS. Examples of RCS occurring in plant material include malondialdehyde, 4-hydroxy-(E)-2nonenal, formaldehyde and acrolein. Unfortunately, the TBARS test fails to detect some physiologically relevant RCS like 4-hydroxy-(E)-2-nonenal [22].
9. Concluding remarks ROS are hot topics in plant biology because they function both as agents of damage and mediators of cellular signals. Understanding of both functions needs knowledge about the properties and reactions of each ROS, and the present review attempts to give an account of the biologically most
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relevant features of ROS, including redox potentials, key reactions and their rate constants, and lifetime and diffusion distance in biological material.
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The second aim of this review was to summarize the most relevant detection methods that have been used, or that are potentially usable, for experimental material deriving from plants, algae or cyanobacteria. The main focus is in photosynthetic tissues. We try to provide a listing of the properties of the method, including the possibilities to locate the ROS in a cell, toxicity of the detector substance and stability of both the detector substance and the signal. Unfortunately, the list often remains incomplete, because some topics have not yet been studied. We hope that the introduction to the methods may prompt future research of the missing topics and future development of better detection methods.
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To minimize artifacts, the following principles should be always be followed when chemical probes are used for ROS measurements.
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1. Ensure the specificity of the probe.
2. Check the toxicity of the probe to the study material and especially to photosynthesis.
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3. Ensure that the wavelengths and intensities of light used in the experiments do not cause artefacts like generation of ROS. 4. Investigate the cellular localization if the probe is used in vivo.
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5. Do not draw quantitative conclusions if the probe is unspecific.
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10. Acknowledgements
This study was financially supported by Academy of Finland (grants 259075 and 271832) and by Nordic Energy Research (AquaFEED project).
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Figure legends
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Fig. 1. A plot of temporally and spectrally resolved luminescence obtained for Chl a in acetone. The peak at 1270 nm, decaying in 40 µs originates from 1O2. Reprinted from [72], Copyright (2003), with permission from Elsevier.
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Fig. 2. Effect of TEMP on PSII in intact cells of the diatom Phaeodactylum tricornutum. The decay of Chl a fluorescence yield after a single turnover flash was measured with the FL-200 fluorometer (Photon Systems Instruments, Brno, Czech Republic) from a P. tricornutum culture before (black dashed line) and after 3 min incubation with 10 mM TEMP that had been purified by vacuum distillation (solid red line).
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Fig. 3. Demonstration that 30 min illumination of tobacco leaves with low light (LL, PPFD 50 µmol m2 -1 s ) induces hardly detectable quenching of DanePy fluorescence, whereas increase in SOSG fluorescence is obvious after the same treatment. Reprinted from [109] with kind permission from Springer Science and Business Media.
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Fig. 4. Stoichiometric production of H2O2 in a reaction between 1O2 and plastoquinol. Reprinted from [60], Copyright (2015), with permission from Elsevier.
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Fig. 5. Formation of H2O2 and O2• in cucumber cotyledons in the light, visualized by staining with DAB and with NBT, respectively. Reprinted from [307], Copyright (2014), with permission from Elsevier.
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Fig. 6. EPR spectra of BMPO/•OOH and BMPO/•OH adducts, as indicated. The dotted lines indicate computer simulations of the respective spectra. Reprinted from [292], Copyright (2001), with permission from Elsevier.
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Highlights
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The review discusses the reactions of Reactive Oxygen Species (ROS) Detection methods of ROS have been critically reviewed Advantages and disadvantages of a large number of chemical detection methods are described The main focus of the review is in the detection or ROS from photosynthetic organisms and tissues
S1011-1344(15)00322-X doi: 10.1016/j.jphotobiol.2015.10.001 JPB 10158
To appear in: Received date: Revised date: Accepted date:
11 June 2015 30 September 2015 1 October 2015
Please cite this article as: Heta Mattila, Sergey Khorobrykh, Vesa Havurinne, Esa Tyystj¨ arvi, Reactive Oxygen Species: Reactions and Detection from Photosynthetic Tissues, (2015), doi: 10.1016/j.jphotobiol.2015.10.001
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Heta Mattila, Sergey Khorobrykh, Vesa Havurinne and Esa Tyystjärvi*
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Reactive Oxygen Species: Reactions and Detection from Photosynthetic Tissues Department of Biochemistry / Molecular Plant Biology, University of Turku, 20014 Turku, Finland
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*Corresponding author, e-mail [email protected]
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Abstract
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Reactive oxygen species (ROS) have long been recognized as compounds with dual roles. They cause cellular damage by reacting with biomolecules but they also function as agents of cellular signaling. Several different oxygen-containing compounds are classified as ROS because they react, at least with certain partners, more rapidly than ground-state molecular oxygen or because they are known to have biological effects. The present review describes the typical reactions of the most important ROS. The reactions are the basis for both the detection methods and for prediction of reactions between ROS and biomolecules. Chemical and physical methods used for detection, visualization and quantification of ROS from plants, algae and cyanobacteria will be reviewed. The main focus will be on photosynthetic tissues, and limitations of the methods will be discussed.
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Contents 1. Reactive oxygen species (ROS) and free radicals ................................................................................ 5 1.1. Properties of ROS ......................................................................................................................... 5
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1.2. General principles of detection and monitoring of ROS .............................................................. 7
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2. Singlet oxygen, 1O2 .............................................................................................................................. 8 2.1. Definition and properties of 1O2 .................................................................................................. 8 2.2. Formation and action of 1O2 in photosynthetic tissues ............................................................... 8
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2.3. Reactions of 1O2 ........................................................................................................................... 9 2.3.1. Physical deactivation of 1O2 by radiative and non-radiative mechanisms............................ 9
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2.3.2. Chemical reactions of 1O2 ................................................................................................... 10 2.4. Lifetime and diffusion distance of 1O2 ....................................................................................... 12 2.5. Detection of 1O2 ......................................................................................................................... 17
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2.5.1. Luminescence at 1270 nm .................................................................................................. 17 2.5.2. EPR detectable probes ........................................................................................................ 17
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2.5.3. Dyes ..................................................................................................................................... 18 2.5.4. Fluorescent probes ............................................................................................................. 18 2.5.5. Oxygen consumption in a reaction between 1O2 and histidine .......................................... 19
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2.5.6. Genetically encoded probes ............................................................................................... 19 2.5.7. Further gene expression methods ...................................................................................... 23
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2.5.8. Indirect methods of 1O2 detection ...................................................................................... 23 2.5.9. Summary of 1O2 detection .................................................................................................. 24 3. Hydrogen peroxide, H2O2 .................................................................................................................. 24 3.1. Definition and properties of H2O2 .............................................................................................. 24 3.2. Formation and action of H2O2 in photosynthetic tissues........................................................... 24 3.3. Reactions of H2O2 ....................................................................................................................... 25 3.4. Lifetime and diffusion distance of H2O2 ..................................................................................... 26 3.5. Detection of H2O2 ....................................................................................................................... 26 3.5.1. Principles of H2O2 detection................................................................................................ 26 3.5.2. Precipitate-forming compounds ......................................................................................... 26 3.5.3. Dyes ..................................................................................................................................... 27 3.5.4. Fluorescent probes ............................................................................................................. 27 3.5.5. Chemiluminescent probe .................................................................................................... 28 3.5.6. Microelectrodes .................................................................................................................. 28 3.5.7. Genetically encoded probes ............................................................................................... 29
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3.5.8. Unspecific probes................................................................................................................ 29 3.5.9. Indirect methods of H2O2 detection ................................................................................... 29 3.5.10. Summary of H2O2 detection .............................................................................................. 29
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4. Superoxide anion radical, O2•, and hydroperoxyl radical, HO2• ...................................................... 33
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4.1. Definitions and properties of O2• and HO2• .............................................................................. 33 4.2. Formation and action of O2• and HO2• in photosynthetic tissues ............................................ 33 4.3. Reactions of O2• and HO2• ........................................................................................................ 33
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4.4. Lifetime and diffusion distance of O2• ...................................................................................... 36 4.5. Detection methods of O2• ......................................................................................................... 37
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4.5.1. Spectroscopic methods ....................................................................................................... 37 4.5.2. EPR detectable probes ........................................................................................................ 37
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4.5.3. Dyes ..................................................................................................................................... 38 4.5.4. Fluorescent probes ............................................................................................................. 40 4.5.5. Chemiluminescent probes .................................................................................................. 40
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4.5.6. Microelectrodes .................................................................................................................. 41 4.5.7. Genetically encoded probes ............................................................................................... 41
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4.5.8. Indirect methods of O2• measurement............................................................................. 41 4.5.9. Summary of O2• detection ................................................................................................. 41 5. Hydroxyl radical, HO• ........................................................................................................................ 46
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5.1. Definition and properties of HO• ............................................................................................... 46 5.2. Formation and action of HO• in photosynthetic tissues ............................................................ 46 5.3. Reactions of HO• ........................................................................................................................ 47 5.4. Lifetime and diffusion distance of HO• ...................................................................................... 48 5.5. Detection methods of HO• ......................................................................................................... 48 5.5.1 Spectroscopic methods ........................................................................................................ 48 5.5.2. EPR detectable probes ........................................................................................................ 49 5.5.3. Fluorescent probes ............................................................................................................. 50 5.5.4. Microelectrode.................................................................................................................... 51 5.5.5. Summary of HO• detection ................................................................................................. 51 6. Peroxyl radical (ROO•), alkoxyl radical (RO•) and hydroperoxides ................................................... 54 6.1. Definition and formation ........................................................................................................... 54 6.2. Reactions of ROO•, RO• and hydroperoxides ............................................................................. 54 6.3. Lifetime and diffusion distance of ROOHs ................................................................................. 56 6.4. Detection methods of ROOHs .................................................................................................... 56
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6.4.1. Spectroscopic methods ....................................................................................................... 56 6.4.2. Chemiluminescence and thermoluminescence .................................................................. 56 6.4.3. Dyes ..................................................................................................................................... 57
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6.4.4. Fluorescent probes ............................................................................................................. 57
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6.4.5. Measurements of ROOH-derived species ........................................................................... 58 7. Ozone, O3 .......................................................................................................................................... 61 8. Non-specific methods ....................................................................................................................... 63
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8.1. Non-specific methods indicating the presence of ROS .............................................................. 63 8.2. Non-specific methods indicating reactions of ROS with biomolecules ..................................... 64
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9. Concluding remarks .......................................................................................................................... 64 10. Acknowledgements......................................................................................................................... 65
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11. References ...................................................................................................................................... 65
1.1. Properties of ROS
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1. Reactive oxygen species (ROS) and free radicals
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The term "reactive oxygen species" refers to naturally occurring, oxygen-containing chemical species whose reactivity, at least towards some substances, is higher than that of ground-state oxygen [1]. In the present review, we will limit the discussion to singlet oxygen, superoxide (superoxide anion radical and hydroperoxyl radical), hydrogen peroxide, hydroxyl radical, ozone, and lipid peroxyl radicals, lipid peroxides and alcoxides. ROS are inevitably produced in photosynthetic organisms [2,3] and cause oxidative damage by reacting with biomolecules [1,4‒7], but ROS also function as cellular signals [8‒10]. Photosynthetic organisms have extensive quenching and scavenging mechanisms that alleviate the effects of ROS [11‒14]. Free radicals are defined as chemical species that contain one or more unpaired electrons and are capable of independent existence [1]. Ground-state O2 is classified as a free radical (a diradical) because the two degenerate outermost molecular orbitals are occupied by one electron each, and in accordance to Hund's rule, the spins of these two electrons are parallel in the lowest energy state of the system. Superoxide anion radical (O2•), hydroxyl radical (HO•) and perhydroxyl radical (HO2•) are also free radicals whereas singlet oxygen (1O2), hydrogen peroxide (H2O2), ozone (O3), peroxides and alcoxides are not radicals. Some free radicals (like HO• and the hydrogen radical H•) are extremely reactive, whereas some free radicals, like ions of transition metals, exhibit weak reactivity. Free radicals are affected by external magnetic fields because the net spin magnetic momentum of the unpaired electrons interacts with a magnetic field. The energy of a free radical depends on the spin configuration and on the strength of the magnetic field. Free radicals are therefore visible in electron paramagnetic resonance (EPR, also called electron spin resonance, ESR) spectroscopy. A radical with one unpaired electron (doublet, total spin 1/2) has two energy states (designated as the spin up and the spin down states) in a magnetic field, and a free radical with two unpaired electrons
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(triplet, total spin 1), has three energy levels. The total spin of a molecule containing only paired electrons is zero, and such a molecule is described as a singlet state. Singlets interact with external magnetic fields only weakly.
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In addition to the classification as radicals and non-radicals, ROS can be roughly divided into two types: (1) Free ROS, small molecules composed of oxygen and hydrogen only, and (2) incorporated ROS in which oxygen is bound to other molecules to form reactive oxygen derivatives. Table 1 presents the most important ROS. The most important reactive nitrogen species (for a review, see [15]) are listed in Table 1 but will not be further discussed.
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Table 1. Most significant ROS and reactive nitrogen species. R is a residual of an organic molecule. Non-radicals Free ROS
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Radicals
Singlet oxygen, 1O2
Superoxide anion radical, O2• Perhydroxyl radical, HO2•
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Hydroxyl radical, HO•
Hydrogen peroxide, H2O2
Ozone, O3 Incorporated ROS and reactive nitrogen species Organic peroxides, ROOH Peroxynitrite ion, ONOO Alkyl peroxynitrite, ROONO
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Peroxyl radical, ROO• Alkoxyl radical, RO• Nitric Oxide, NO• Nitrogen dioxide,NO2•
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The biological actions of ROS are selective because the reactivity of each ROS strongly depends on the ROS and on reaction conditions. Some ROS like HO• have a very high chemical activity and are able to oxidize most organic molecules. On the other hand, H2O2 has moderate reactivity and is able to migrate far from site of production. The general reactivity of the main ROS in biological environment decreases in the order HO• > 1O2 > H2O2 > O2•, although care should be taken in using this generalization, as the rate of reaction depends on the reaction partner. Ability to interconverse is an essential property of ROS. For example, 1-electron reduction of H2O2 leads to the formation of HO•, indicating that formation of a less reactive form can promote the formation of more reactive forms. Due to the interconversions, cellular effects of ROS should be considered as complex sets of reactions. Given these reactions, action mechanisms of oxygen in the cell can be understood. Production, detoxification and detection of ROS, as well as the roles of ROS as both agents of damage and as cellular signals are widely discussed (for reviews see e.g. [16‒18]). In the following, we will give an overview of the reactions of each ROS and describe methods that have been used to detect the ROS, focusing on photosynthetic organisms. The biological roles of the ROS in photosynthetic organisms will be discussed only to the extent needed for the understanding of the detection methods.
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1.2. General principles of detection and monitoring of ROS
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In pure solvents or in the gas state, concentrations of various ROS can be directly measured by absorbance, fluorescence or EPR spectroscopy. However, the most common methods for monitoring the relative amounts of ROS require the use of chemicals that preferentially react with a particular ROS, yielding a detectable product.
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Most ROS do not equilibrate throughout a complex biological material within their lifetime but either react near to the site of origin or remain confined to the producing compartment. Therefore the cellular localization of the detector substance is often of importance for understanding the results. Unfortunately, experimentally verified information about the localization of the various detector substances is seldom available, especially in plant material. In the present review, we list the available information. The logarithm of 1-octanol-water partition coefficient (LogP) is widely used to estimate hydrophobicity. As a rule, the lower the LogP value of a compound is, the more hydrophilic it is. All experimentally determined and some estimated LogP values in this review have been obtained from online chemical structure database, Chemspider [19]; for compounds not found in Chempsider, LogP values were estimated using an online tool ALOGPS 2.1 [20]. LogP values for different ROS detection compounds are presented in Tables 3‒7. Caution should be taken in interpreting LogP values as indicators of cellular localization or intake of a molecule, because cellular environment cannot be dichotomously divided into two simple immiscible phases. Furthermore, the plant and cyanobacterial cell wall may lower the cell permeability of lipophilic molecules.
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Hydrophilic chemicals can be fed to intact plant samples in the transpiration stream but lipophilic substances may require vacuum infiltration or scratching the plant surface. Side effects of these aggressive methods should be separately evaluated. Biotechnology offers the possibility to make a mutant organism that synthesizes a specific genetically encoded ROS probe in the desired tissue or cellular compartment [21]. The success of biotechnology is based on the availability of specific cellular ROS probes and so far only probes for H2O2, based on the specific sensitivity of transcription factors (OxyR from Escherichia coli and Orp1 from Saccharomyces cerevisiae) to H2O2, have been tested for the detection of endogenous ROS (see [16,22] for reviews). Promising results have been obtained also in the development of genetically encoded 1O2 probes (see Chapter 2.5.6). Because certain ROS function as cellular signals, specific changes in gene expression can often be correlated with specific ROS. However, conclusions about the causal agent causing a change in gene expression must be drawn with great care, unless the mechanism of regulation is well understood. Enzymes present in all cells can sometimes be used to modify the sensor substances, as in the design of fluorescein diacetate dyes from which cellular esterases remove the acetate moiety, producing a fluorescent, charged dye that cannot diffuse back to the extracellular space [23]. In the case of aminophenyl and hydroxyphenyl fluorescein dyes, the strength of the ROS signal may also depend on the action of a peroxidase [24]. Detection of ROS with chemical reactions is often difficult because several ROS may react with the detector chemical. Unspecific detectors can be used in the rare cases that the system is known to produce only one type of ROS and that only one type of ROS is of interest. However, measurement of the total amount of ROS in the sample cannot be done with an unspecific detector because the reaction rates of the detector with different ROS vary. If the concentration ranges of various ROS
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species in the sample have been well characterized in advance, it may also be possible to use several unspecific detector substances and utilize knowledge about their reaction rates with different ROS.
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In addition to the specificity and localization in the biological sample, ROS detector substances may have problems of biological incompatibility, stability in the light and stability of the reaction product. Some ROS detector substances are known to produce ROS. These topics will be discussed when information is available.
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2. Singlet oxygen, 1O2 2.1. Definition and properties of 1O2
A ground-state O2 molecule ( g O2) is a triplet form (also designated as 3O2) and can therefore 3
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accept two electrons from a species containing paired electrons only if a spin conversion occurs. Thus, the reactivity of O2 at room temperature is moderate because the molecule has two electrons with parallel spins. The singlet forms of O2 containing only paired electrons are much more reactive than ground-state O2. Molecular oxygen has two singlet forms. The two electrons with antiparallel spins may reside
either on two different orbitals ( g O2) or both on one orbital ( g O2). g O2 decays rapidly to 1
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form either ground-state O2 or g , and therefore the designation 1O2 will be used for g O2, 1
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unless otherwise indicated.
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O2 is a long-lived excited state, but its decay is greatly speeded up by collisions with almost any other molecule [25]. The radiative decay of 1O2 to ground-state O2 yields luminescence with maximum at 1268 nm in the gas phase [26]; in water and deuterium oxide (D2O), the peak is at 1274 nm [27]. Although 1O2 is a singlet state, it has an EPR spectrum [28]. The spectrum has been detected only in the gas phase [29]. Rapid non-radiative decay of 1O2 occurs via (i) conversion of excitation energy of 1O2 to vibrational and rotational energy of both the quencher (e.g. H2O) and O2; (ii) a charge-transfer mechanism between the quencher (e.g. phenol or diazabicyclo[2.2.2]octane, DABCO) and 1O2; or (iii) an electronic energy transfer mechanism in which the reaction between 1O2 and a singlet ground state of the quencher (e.g. a carotenoid) produces a triplet state of both the quencher and oxygen (i.e. ground-state oxygen) [25].
2.2. Formation and action of 1O2 in photosynthetic tissues In photosynthetic tissues, 1O2 is mainly formed by photosensitization (for a review see [30]). Encounter of O2 with either a singlet or a triplet excited state of a pigment molecule (sensitizer) may lead to generation of a singlet form of oxygen. A reaction between a singlet excited state of a sensitizer and O2 yields 1O2 and a triplet state of the sensitizer, whereas a reaction between O2 and a triplet state of the sensitizer produces 1O2 and a singlet ground state of the sensitizer [25]. The reactions between O2 and singlet and triplet excited states of sensitizers are also affected by the energy gap between the singlet and triplet states of the sensitizer [31]. In practice, reactions with singlet excited states are of low significance because singlet excited states are short-lived. Chlorophylls, bacteriochlorophylls, hemes and other tetrapyrroles, especially protoporphyrin IX (an intermediate of synthesis of heme and chlorophyll), as well as iron sulfur centers are known
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photosensitizers [4]. However, chlorophyll (Chl) a and b, and bacteriochlorophylls in non-oxygenic photosynthesis, are the most important natural photosensitizers of 1O2 formation in photosynthetic organisms. According to the available data, production of 1O2 in plants is almost always sensitized by the triplet state of the primary donor (3P680*) of Photosystem II (PSII), formed by charge recombination reactions (for a review see [3]). Thus, neither the triplet state of Chl a (3Chl a) produced by intersystem crossing in the antennae [32] nor the triplet state of the primary donor of Photosystem I (PSI) (3P700*) seem to sensitize 1O2 formation [33,34]. Weak triplet formation in the antennae due to photochemical and non-photochemical quenching of excitation energy, low oxygen concentration in the antenna, and quenching of the triplet states by carotenoids may contribute to the apparently negligible sensitization by many Chl triplets in plants. Experiments with the histidine method (see 2.5.5) show that at photosynthetic photon flux density (PPFD) of 2300 µmol m-2s-1 the 1 O2 quencher histidine lowers oxygen production of the cyanobacterium Synechocystis sp. PCC 6803 by 13 % [35]. This result may actually suggest massive production of 1O2 within the membranes, as most 1O2 is expected to react at the site of production [1].
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Other sources of 1O2 than PSII in plants are related to plant defense and e.g. heat stress (for reviews, see [3,36,37]). Non-photochemical formation of 1O2 with the Russell mechanism, via recombination of two peroxy radicals, has also been shown to occur in plant tissues [38,39] but the importance of this pathway has not been studied extensively.
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Detoxification of 1O2 in plant cells occurs by carotenoids and tocopherols in the thylakoid membranes (for reviews, see [5,14]) and in the cytosol by soluble scavengers including vitamins C and B6 and glutathione (GSH) [40,41]. 1
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O2 has been speculated to be the most important ROS formed in the light reactions of photosynthesis, as the triplet state of the primary donor of PSII is ubiquitously formed by charge recombination reactions. Work on the flu mutant of Arabidopsis thaliana that produces 1O2 when grown in a light-dark rhythm has revealed that the reason for cell death is not cellular damage mediated by 1O2 but 1O2-induced signaling events [42‒44]. However, the fact that programmed cell death depends on 1O2 signaling, does not rule out the possibility that 1O2 causes cellular damage.
2.3. Reactions of 1O2
2.3.1. Physical deactivation of 1O2 by radiative and non-radiative mechanisms Only radiative deactivation (reaction 1) is available for an isolated and unperturbed 1O2 molecule, and the radiative lifetime O2(1g) without any collisions is extremely long, 72 min [25]. 1
O2 3O2+ h
(1)
Enhancement of deactivation of 1O2 by collisions of 1O2 with other molecules is one reason for variation of the lifetime of 1O2 in different media. Deactivation of 1O2 by collisions with other molecules having sufficiently high triplet-state energy can occur by conversion of excitation energy of 1O2 to vibrational energy of the other molecule. This non-radiative deactivation process, known as electronic-vibrational energy transfer, limits the lifetime of 1O2 in many solvents. The lifetime of 1O2 in H2O is 3.1 s, about 20 times less than in D2O (68 s) [25,45,46]. Effects of solvent on the lifetime of 1O2 can often provide evidence for participation of 1O2 in chemical reactions.
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In addition to the electronic-vibrational non-radiative deactivation, 1O2 can be deactivated via an electron exchange mechanism (reaction 2). This spin-allowed electronic energy transfer to a molecule with a low triplet state energy is a very efficient mechanism of 1O2 deactivation. For example, the rate of deactivation of 1O2 with carotenoids is limited by diffusion. The rate constant for β-carotene and lutein is about 1010 M-1 s-1 [47], O2 + A 3(O2 A) 3O2 + 3A,
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(2)
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where A is an acceptor molecule, for example a carotenoid. Reaction (2), also known as physical quenching of 1O2, is the main mechanism of quenching of 1O2 by carotenoids. Bimolecular rate constants of reactions between carotenoids and 1O2 can be found in [48].
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In addition to their ability to quench 1O2, carotenoids protect the photosynthetic machinery also by quenching both the triplet and singlet excited states of Chls, thereby lowering the probability of 1O2 formation. In plant photosynthesis, the xanthophyll violaxanthin is de-epoxidated in the light to anteraxanthin and zeaxanthin which participate in so called non-photochemical quenching of excitation energy, a mechanism converting the energy of the singlet excited state of Chl a to heat. For review of the photoprotective roles of carotenoids in photosynthesis, see [49].
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2.3.2. Chemical reactions of 1O2 1 O2 is an electrophilic agent and reacts preferably with electron rich organic molecules by addition to double bonds. This chemical deactivation of 1O2 proceeds via well-known mechanisms:
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ene reaction associated with formation of a hydroperoxide (3);
(3)
cycloaddition associated with dioxetane formation (4); (4)
cycloaddition reaction with aromatic compounds and formation of endoperoxides via the Diels-Alder mechanism (5); (5)
formation of charge-transfer intermediates. 1O2 can react with compounds containing heteroatom such as N or S to form charge-transfer intermediates, reactions (6) and (7) respectively. Chargetransfer intermediates are particularly important for the interaction of 1O2 with phenols and tocopherols [50]. O2 + NR3 [1O2--- +NR3]
1
(6)
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O2 + SR2 [1O2--- +SR2]
1
(7)
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The charge-transfer complex can be disintegrated either by an electron exchange mechanism (physical quenching of 1O2) or by oxidation of a heteroatom, reaction (8).
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(8)
(9)
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In addition to oxidation of a heteroatom, the reaction with 1O2 can lead to carbon-heteroatom bond cleavage and formation of a corresponding aldehyde [51]. 1O2 can efficiently oxidize amines to imines with formation of HO2•, reaction (9) [52].
1
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The interaction of 1O2 with the heteroatom of 2,2,6,6-tetramethly-4-piperidone (TEMP) produces 2,2,6,6-tetramethyl-4-piperidone-1-oxyl (TEMPO, TAN or TEMPONE), a stable radical (reaction 10) [53,54]. The reaction of 1O2 with TEMP (k=4 x 107 M-1 s-1 in phosphate buffer at pH 8, [53]) can be used to detect 1O2 with EPR or mass spectrometry (see 2.5.2).
(10)
O2 can react with many biologically important compounds like unsaturated fatty acids of membrane lipids [55]. The reaction of 1O2 with a cis-double bond of an unsaturated fatty acid can form both conjugated and non-conjugated diene hydroperoxides (reaction 11). The rate constants of reactions of 1O2 with unsaturated fatty acids are slightly higher than 104 M-1 s-1 [41].
(11)
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Methionine, tryptophan, tyrosine, cysteine and histidine react with 1O2 to form semistable products with a rate constant around 107 M-1 s-1 [56]. Histidine has been used for detection of 1O2 [35]. 1O2 also reacts with GSH and other thiol containing compounds with rate constants around 106 M-1 s-1. Reaction of 1O2 with GSH is associated with formation of glutathione sulphinate (GSO2H), glutathione sulphoxide (GSSOG), glutathione sulphonate (GSO3H) and glutathione disulphide (GSSG) [57]. Biologically active forms of vitamin B6 react with 1O2 with rate constants around 1 x 108 M-1s-1 [58]. Oxidation of ascorbic acid and plastohydroquinone by 1O2 can proceed as 2-electron reduction of 1O2 to H2O2 [59,60].
2.4. Lifetime and diffusion distance of 1O2
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The lifetime of 1O2 is of interest because a short lifetime would predict that 1O2 reacts at the site of its production whereas a long lifetime would allow 1O2 to act as a signal molecule. The diffusion distance can be approximated from the lifetime (see e.g. [25,61]). The lifetime of 1O2 in cellular environments has been several times measured in D2O based buffers but these measurements do not give a correct picture of what happens in vivo. Current sensor technology has, however, made it possible to measure the kinetics of the 1270 nm luminescence peak after a short laser pulse from some biological materials in H2O based buffers.
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The formation and decay of the 1O2 peak, obtained by exciting a sensitizer with a short laser pulse, remain unaltered if the values of the rate constant of the decay of the triplet state of the sensitizer (kT) and the rate constant of the decay of 1O2 (kΔ) are exchanged [62] (see Appendix in Supplementary information for the modeling of the kinetics of the 1270 nm peak). For this reason, measurements of 1O2 luminescence kinetics always need knowledge about the value of kT in the same experimental system. The long lifetime of 1O2 in D2O alleviates the problem because sensitizers with a short triplet lifetime (1/kT) can be used. If the sensitizer is an intrinsic constituent of the sample, like Chl in plant photosynthesis, the requirement of the knowledge of kT places a particular weight for the correct identification of the sensitizer. Further complications arise from the heterogenity of the biological sample. Theoretical considerations based on general cellular concentrations of histidine, tryptophan and methionine residues in proteins [63] or on the concentrations of various biomolecules like carotenoids and ascorbate that react with 1O2 [64,65] suggested that the lifetime is 0.05‒1 µs in cells [63,64] and 0.2 µs in chloroplasts [65] (Table 2). Measurements of 1270 nm luminescence kinetics from animal cells have, however, yielded longer but extremely variable lifetimes, ranging from 0.4 to 10 µs in cell suspensions or in individual cells (Table 2). Ragas et al. [66] estimate that the lifetime of 1 O2 inside Escherichia coli cells is only 7 ns. The large variation may partially depend on differences between the samples, as the concentrations of substances reacting with 1O2 may vary between cell types. Also the cellular localization of the sensitizer has been shown to affect the measured lifetime [67]. An apparent source of differences is that in some studies, whole tissues have been used whereas in other studies the current laser technology has been applied to measure 1O2 lifetime from single cells (Table 2). One possible reason for long lifetimes obtained in biological materials is the presence of membranes and lipid bodies in cells. The lifetime of 1O2 is 20‒25 µs in micelles [68] and 12.2 µs in liposomes [69]. Interaction between 1O2 with a quencher or a scavenger molecule requires physical contact of the molecules, and therefore the concentrations of quenchers and scavengers in the lipophilic environment would determine how much they shorten the lifetime of 1O2. Hackbarth et al. [70] see two different lifetime components in a mammalian cell system and suggest that one
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corresponds to the membrane and the other to the aqueous phase. However, most experimental data can be explained with only one lifetime component, which suggests that 1O2 equilibrates between different types of compartments within its decay time in the heteronegeous biological environment. Strong laser pulses have also been shown to prolong the measured lifetime of 1O2, probably via depletion of cellular 1O2 quenchers [70].
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So far all 1O2 lifetime measurements from plant and cyanobacteria samples have been done from isolated PSII preparations, either in D2O based buffers [71] or more recently also in water based buffer solutions [65,72,73]. In the case of the simplest PSII reaction centre preparations that lack the QA electron acceptor, the reaction centre triplet is obviously also 1O2 sensitizer. However, the triplet yield of the recombination of the primary radical pair is very low if the electron can proceed from pheophytin to a further acceptor [71], and therefore identification of the sensitizer triplet is more difficult when functional PSII preparations are used. The lifetimes per se mainly reflect the properties of the buffer (D2O or H2O). Correlation between kinetics of decay of the triplet form of the primary donor of PSII and 1270 nm luminescence rise at different temperatures and oxygen concentrations showed that recombination of the primary radical pair produces the sensitizer triplet in PSII particles [71]. Tomo et al. [73] show that much more 1O2 is produced by PSII preparations from a cyanobacterial mutant that contains divinyl Chl a instead of the monovinyl form occurring in the wild type.
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With regard to the potential signaling role, a triplicate lifetime, after which 5 % of 1O2 still exists, could be a practical limit for the reach of the signal. Assuming, in the absence of measurements, that the lifetime of 1O2 is 3 µs in aqueous stroma of chloroplasts and further assuming that 1O2 can pass through the envelope of a typical chloroplast (5‒7 x 2.5 µm; membrane thickness ~4 nm [74]), then approximately 20 % of the volume of the chloroplast could produce 1O2 of which a significant fraction could diffuse to the cytosol. However, 1O2 produced externally to mammalian cells failed to cause DNA strand breaks while 1O2 produced inside the cells did cause breaks [75,76], indicating that in animal cells the amount of 1O2 diffusing through the plasma membrane is small. Diffusion of 1O2 through the plasma membrane or cell wall has not been measured in plants or cyanobacteria. The diffusion distance of 1O2 may also determine the effectiveness of quenchers and scavengers. In this respect, the finding that added β-carotene protects mammalian cells against 1O2-induced cell death but does not shorten the lifetime of 1O2 [77] is highly interesting. The high viscosity of the cellular environment can partly explain the lack of an effect, as a three-fold increase in viscosity from C6H14 to CCl4 is associated with a 30 % decrease of the quenching constant of 1O2 by β-carotene [48]. The short diffusion distance of 1O2 also implies that cell-impermeable detector substances can only measure extracellularly produced 1O2.
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Sensitizer
Method of estimation
Buffer solution
Independent Lifetime measurement
0.05 to 1 µs [64]
0.4‒0.18
Effective signaling distance in viscose cellular environment, µm 0.013‒0.060
< 1 µs [63]
<0.180
<0.06
0.2 µs [65]
0.08
0.03
No
10 µs [79]
0.5
0.2
No
6 µs [79]
0.4
0.15
D2O
Yes
45 µs [80]
1.2
0.4
D2O
Yes
17 µs [67]
0.7
0.25
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System
CR
IP
Table 2. Measurements of 1O2 lifetime with 1270 nm luminescence kinetics in biological systems. An estimated effective signaling distance, at which 5 % of 1 O2 survives, is given for water (D=1.8 x 10-5 cm2 s-1 at 20 °C; [78]) and for viscose cellular environment (D=2 x 10-6 cm2s-1; [61]).
Nucleus of a single neuron HeLa cells (single cell)
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Photofrin 9-acetoxy-2,7,12,17tetrakis(beta-methoxyethyl)porphycene 5,10,15,20tetrakis(N-methyl-4-pyridyl)21H,23H-porphine Chlorin
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Cells
Chloroplast stroma Mammalian cells Mammalian cells
Theoretical, based on known concentrations of quenchers/ scavengers Theoretical (His, Trp, Met residues) Theoretical (ascorbate) 1270 nm H2O luminescence 1270 nm H2O luminescence
TE D
Various cellular environments
MA N
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Effective signaling distance in water, µm
1270 nm luminescence
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Yes
30‒40 µs [67]
1‒1.1
0.3‒0.4
3.2 µs [81]
0.3
0.1
0.4 µs outside of membrane [70] 0.6 µs [82]
0.1
0.04
0.1
0.04
0.03-0.18 µs [82]
0.03‒0.08
0.01‒0.025
3.7 µs; estimated 0.007 µs inside the cells [66] 44‒59 µs (airsaturated medium, 20 °C [71] 3.2 µs [65]
0.3 (based on the extracellular lifetime)
0.005 (based on the estimated cellular lifetime)
IP
Extrapolation D2O/H2O Yes from mixtures measurements in D2O/H2O mixtures 1270 nm H2O Yes luminescence
CR
Neuron (single cell)
D2O
T
5,10,15,20tetrakis(N-methyl-4-pyridyl)21H,23H-porphine
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HeLa cells (single cell)
Pheophorbide a
Leukemia cells
Aluminum tetrasulphonated phthalocyanine
1270 nm luminescence
H2O
Rat tissues in vivo
Aluminum tetrasulphonated phthalocyanine
1270 nm luminescence
H2O
Escherichia coli cells
New Methylene Blue and Zinc tetramethyltetrapyridino[3,4b:39,49-g:30,40-l:3-,4q]porphyrazinium salt
1270 nm luminescence
H2O
PSII reaction centre preparations
PSII reaction centre triplet
1270 nm luminescence
D2O
Yes
PSII reaction centre preparations
PSII reaction centre triplet (assumed)
1270 nm luminescence
H2O
No (assignment supported by effect of
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Jurkat cells
No (fitting and literature values used) No (fitting and literature values used) No (fitting and literature values used)
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H2O, D2O
Functional PSII particles from wild type and a mutant with divinyl Chl a
PSII reaction centre triplet (assumed)
1270 nm luminescence
H2O
oxygen) Yes
MA N
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No
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3.4‒4.3 µs in H2O; 17‒18.7 µs in D2O [72] 3.94‒4.80 µs [73]
T
1270 nm luminescence
IP
Chls (assumed)
CR
PSII particles
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2.5. Detection of 1O2
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2.5.1. Luminescence at 1270 nm Several recent reviews discuss the detection of 1O2 from photosyhthetic organisms [3,83‒85]. 1O2 can be measured by recording the weak 1270 nm luminescence emitted when 1O2 returns to the ground state (reaction 1; Fig. 1; for reviews see [25,84,85]). The detection of 1270 nm luminescence from biological materials is limited by the material thickness, as absorption of water [86] and biomolecules would attenuate the signal to less than one tenth per 1 cm. Therefore, thin samples are preferred. D2O is far more transparent than H2O in the near-infrared range [87].
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Due to the low phosphorescence yield and low concentration of 1O2 in biological systems, measurements of 1270 nm luminescence in photosynthetic material have been limited to generation of 1O2 with strong laser pulses, usually in isolated samples, including PSII reaction centres of pea or PSII particles from cyanobacteria [72,73,88]. Significantly more sensitive detectors than are available now will be needed for the measurement of 1270 nm luminescence from biological material under constant illumination [65]. An additional difficulty is to distinguish 1O2 phosphorescence from the tail of Chl a fluorescence, but the separation can be done by spectral comparison or by comparing the decay times [72].
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In addition to the 1270 nm "monomol" luminescence, 1O2 also emits luminescence at 634 and 703 nm. This "dimol luminescence" is emitted after a collisional formation of a (1O2)2 dimol in the gas phase [89]. Emissions originating from transitions between several excited states of the oxygen molecule are important in atmospheric science [90] but too weak for studies from biological material.
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2.5.2. EPR detectable probes TEMP and TEMPD. The two most popular chemicals used to measure 1O2 from photosynthetic material have been 2,2,6,6-tetramethylpiperidine (TEMP) and the more water-soluble 2,2,6,6tetramethyl-4-piperidinone (TEMPD); both commercially available (see Table 3 for review of chemicals used for 1O2 detection). Reaction between 1O2 and TEMP (reaction 10) or TEMPD produces a stable oxyl radical, and the reaction has been shown to be specific to 1O2 [53,91]. In the case of TEMP, the radical is called TEMPO. TEMPD is available both as a pure compound and as a hydrochloride (TEMPD-HCl); only the hydrochloride form can be used for photosynthetic applications because the other form strongly inhibits PSII [92,93]. Due to the low solubility to water, TEMP cannot be used at temperatures below 20 °C. Both TEMP (experimentally determined LogP = 2.15 [19] and TEMPD (LogP = 0.43, as estimated with ALOGPS2.1 [20]) are thought to penetrate into cyanobacterial cells [94,95]. The amount of the TEMPO radical has traditionally been measured with EPR spectroscopy [96,97]. EPR is not suitable for automated measurements of large numbers of samples, and a mass spectrometry method was recently developed for the quantification of TEMPO [98]. TEMP was found not to inhibit the PSII activity of isolated thylakoid membranes [96]. However, storage of TEMP enriches unknown impurities. In the commercially available TEMP, and also in TEMPD, these impurities are present at such a quantity that they cause strong inhibition of PSII, as demonstrated with fluorescence, thermoluminescence and oxygen evolution measurements [92].
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Vacuum distillation removes the impurities to such extent that TEMP can be used with isolated plant thylakoids without any adverse effects [93], but even distilled TEMP causes an almost immediate inhibition of PSII in live cyanobacteria (Synechocystis sp. PCC 6803, [5]) and in live diatoms Phaeodactylum tricornutum (Fig. 2).
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With distilled TEMP, 1O2 can be detected from isolated plant thylakoid membranes (e.g. [93,96,98]). TEMP has been used also with purple bacteria [99] and TEMPD has been used to detect 1O2 from isolated thylakoids of the green alga Chlamydomonas [100], but in vivo effects of the probes on the photosystems of the organism were not reported. In intact plant material, the TEMPO radical can be reduced to an EPR silent hydroxylamine, and therefore TEMPO is usually extracted with an organic solvent if EPR is used as the detection method [101].
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2.5.3. Dyes Imidazole/RNO. Imidazole-containing compounds, including imidazole itself, are rapidly oxygenated by 1O2 [102,103], and 1O2 can also be detected by measuring bleaching of p-nitroso-dimethylaniline (RNO) by imidazole [104‒106]. The method has been used to measure the action spectrum of 1O2 production in isolated spinach thylakoids [107]. The RNO method should be used with caution in plant material because imidazole has been shown to inhibit PSII oxygen evolution and carbonic anhydrase in a pH-dependent manner [108]. Both imidazole and RNO are commercially available.
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2.5.4. Fluorescent probes SOSG. Singlet Oxygen Sensor Green®, a proprietary fluorescent probe, is highly selective for 1O2. According to the manufacturer's data, SOSG does not react with H2O2 or O2•, and our unpublished experiments have confirmed the high selectivity. Reaction with 1O2 increases the fluorescence yield of SOSG at 520‒540 nm. According to the manufacturer's data, SOSG is cell-impermeable, but a study with tobacco leaves showed that a pinhole administration of SOSG to tobacco leaves leads to preferential association of the dye with the nuclei of epidermal cells [109]. Gollmer et al. [110] showed that SOSG is also able to penetrate into human cells; therefore SOSG can be used in confocal microscopy [111]. A disadvantage of SOSG is that when illuminated with wavelengths below 600 nm, the fluorescence yield of SOSG increases similarly as during exposure to 1O2 (Fig. 3; [92,109]). Furthermore, the reaction product of SOSG and 1O2 sensitizes 1O2 production [112]. Surprisingly few researchers have taken the photosensitivity of SOSG into account (for correct use of SOSG, see [92,109,113]). SOSG caused 15 % inhibition of the photochemical yield of PSII in tobacco leaves [109]. It is unclear if SOSG can be used with live cyanobacteria, as SOSG seems to penetrate only to a small fraction of the incubated cells and fluorescence from both SOSG and Chl a is only rarely detected from the same cell [114]. Illumination of isolated pumpkin thylakoids in the presence of SOSG with >650 nm light caused a very small signal which was not sensitive to D2O, anaerobicity or azide, suggesting that SOSG cannot be used to measure 1O2 from isolated thylakoids [92]. However, SOSG has been used for detecting 1O2 from isolated plant thylakoids and from intact leaves [109,113,115]. Reasons for the contrasting results about the sensitivity of SOSG in plant material are presently unknown. DanePy. Dansyl-2,2,5,5-tetramethyl-2,5-dihydro-1H-pyrrole (DanePy), another fluorescent probe, is sensitive to 1O2 with minor reactivity with O2•, H2O2 and lipid radicals [116,117]. The probe is not available commercially. DanePy has been used with bean, pea, tobacco, Arabidopsis thaliana and
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Phillyrea latifolia leaves in vivo [109,118‒121] and with the photosynthetic proteobacterium Rhodobacter sphaeroides [122]. DanePy can be used for imaging [118‒120]. DanePy is fluorescent (500‒600 nm, maximum at 545 nm) but forms a non-fluorescent adduct with 1O2, and therefore the presence of 1O2 is inferred from the decrease in fluorescence yield from untreated control to a treated sample. DanePy is harmless for the PSII yield if administered to a leaf either through a pinhole or in the transpiration stream and localizes intracellularly, mainly to chloroplasts [119,120]. DanePy is not light-sensitive [109]. An oxalate derivative of DanePy was used with live Chlamydomonas cells [100] but this dye does not penetrate to leaf cells [109].
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DMAX and DPAX. The probes 9-[2-(3-carboxy-9,10- dimethyl)anthryl]-6-hydroxy-3H-xanthen-3-one (DMAX) and 9-[2-(3-carboxy-9,10- dimethyl)anthryl]-6-hydroxy-3H-xanthen-3-one (DPAX) can be used to detect 1O2 with fluorescence [123,124]. DMAX and DPAX were reported not to react with H2O2, O2• or NO• [123,124], and DPAX has been used with intact chloroplasts [125]. DMAX and DPAX are not commercially available.
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2.5.5. Oxygen consumption in a reaction between 1O2 and histidine Histidine is known as a powerful, biocompatible antioxidant because its imidazole ring reacts rapidly with 1O2 (k=5 x 107 M-1 s-1; [126]; for a review, see [127]). Histidine also protects against HO•, but this protection may be based on the efficient scavenging of metals by histidine rather than reactions between histidine and HO• [127]. Recently, a simple method in which 1O2 production is measured by monitoring the decrease of oxygen concentration in the presence of histidine [128] has been applied to 1O2 measurements from live cyanobacteria [35,129,130]. Cyanobacteria take up histidine from the medium and therefore histidine can be used for 1O2 measurements in live cyanobacteria [35]. The method was shown to be insensitive to O2• and H2O2 [35]. No reports of the use of the histidine method in plants or algae in vivo have been published, and it is possible that in eukaryotes histidine would not enter the chloroplasts that normally export histidine. The reaction between histidine and 1 O2 occurs in neutral to alkaline solutions [126]. 2.5.6. Genetically encoded probes Specific, genetically encoded probes have not been used for the detection of 1O2. However, the IFP1.4 protein containing a biliverdin (an oxidized form of bilirubin) chromophore [131] has been shown to react with 1O2 [132] and not with H2O2, O2• or HO• [132]. The protein emits far-red fluorescence (maximum at 710 nm) when excited at 684 nm, and might therefore act as a perfect genetically encoded 1O2 sensor. The combination of IFP1.4 and a 1O2 sensitizer protein has been used for measurement of protein-protein interaction [132]. Furthermore, fluorescence of the UnaG protein that contains bilirubin as a chromophore was shown to decrease due to a reaction between bilirubin and 1O2 [133], and thus this protein may also be developed into a genetically encoded 1O2 probe. These probes have not yet been applied for detection of endogenous 1O2 production.
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Specificity
Stability
Toxicity
Quantification of TEMPO with EPR [96] or mass spectrometry [98] in vitro
Specific [53,91]
Stable but instant measurements recommended [98]
Not toxic to isolated plant thylakoids [93]; Inhibits PSII in live cyanobacteria [5] and algae (Fig. 2)
Notes
Cell permeable (Synechocystis) [95]
Not toxic to isolated plant thylakoids [93]
-
1.24 (estimated)
Cell permeable (Synechococcus) [94]
Vacuum distil before use [93,92]; Extraction of TEMPO to organic solvent avoids formation of EPR silent hydroxylamine [101] Use TEMPD-HCl [93]
Does not react Stable with H2O2 or O2• [35]
Not toxic
-
-3.32 [19]
-
Only in neutral or alkalic pH [126]
No reaction with H2O2 or O2• (manufacturer's
Little or no effect on photosynthesis
Only red light (>660 nm) can be used
-
Nuclei of epidermal cells; does not localize
The reaction product sensitizes 1O2
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Light sensitivity
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Specific, similar Stable to TEMP
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TEMPD (2,2,6,6- Quantification tetramethyl-4of the reaction piperidinone) product with EPR in vitro [100] Histidine Consumption of oxygen by histidine, in vitro; in vivo from cyanobacteria [35] Singlet Oxygen Fluorescence (Ex Sensor Green, 505, Em 525), in SOSG vitro and in vivo
Localization
-
Hydrophobicity (LogP) 2.15 [19]
CR
Method
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Compound (IUPAC name) TEMP (2,2,6,6tetramethylpipe ridine)
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Table 3. Chemicals used to detect 1O2. The stability data indicate the stability of the signal after reaction with 1O2 in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data.
Stable
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in plants [109]
[92,109,113]
Minor reaction with H2O2, O2• and lipid radicals [116,117]
No effect on PSII efficiency [109]
Imaging
-
-
-
Decrease in absorbance at 440 nm
-
-
DPAX (9-[2-(3carboxy-9,10dimethyl)anthryl ]6-hydroxy-3Hxanthen-3-one) IFP1.4 (genetically encoded probe with biliverdin chromophore) UnaG (genetically encoded probe with bilirubin
Fluorescence (Ex 492, Em 516‒517), intact chloroplasts [124]
Does not react with NO, H2O2 or O2• [123]
Decrease in fluorescence (Ex 684, Em 710)
Does not react with NO, H2O2 or O2• [132]
Decrease in absorbance (~475) and fluorescence
Aproprotein reacts with O2• at high concentration
to chloroplasts [109] Cellular; mainly chloroplasts [109,119,120]
production [112] -
Extracellular in plants [109]; cytoplasmic in Chlamydomonas [100] -
Less sensitive than DanePy [109]
-
-
IP
Fluorescence (Ex 364, Em 545 nm) is quenched by 1O2, imaging
Not sensitive
-
-
-
Imidazole inhibits PSII [108]
-
-
-
-0.08 imidazole, [19]/ 1.45 p-nitrosodimethylaniline (estimated) Soluble to water at 10 µM [123]
-
-
-
-
-
Not yet tested for endogenous 1 O2
-
-
-
-
-
Not yet tested for endogenous 1 O2
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-
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-
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DanePy (dansyl2,2,5,5tetramethyl-2,5dihydro-1Hpyrrole) DanePy-oxalate (dansyl-2,2,5,5tetramethyl-2,5dihydro-1Hpyrrole oxalate) RNO (imidazole and p-nitrosodimethylaniline)
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data)
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IP
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(~525)
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chromophore)
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2.5.7. Further gene expression methods Ability to sense 1O2 may be important both in avoiding damage and in signaling [134], and photosynthetic organisms have several genes that react specifically to 1O2 (e.g. [135]), as well as to other ROS (see e.g. [136]). The mRNA levels of 1O2 inducible genes have been used to mark 1O2 levels in the cells [137]. Also reporter systems, where proteins induced by 1O2 have been fused to marker proteins, have been developed, for example in Chlamydomonas reinhardtii and Arabidopsis thaliana [138,139], respectively.
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2.5.8. Indirect methods of 1O2 detection Damage caused by 1O2. If 1O2 is not quenched or scavenged, it can damage proteins, pigments and lipids [1,140]. The amount of 1O2 can be estimated by the extent of the damage, as reactions between cell components and 1O2 sometimes have specific reaction products. Products of lipid peroxidation, as well as oxidation products of carotenoids, plastoquinone and tocopherol, have been suggested to be used for detection of 1O2 [55,137,141,142], respectively. However, it is not usually very easy to make strong conclusions based on these data, and the ability of the photosynthetic organisms to quench 1O2 can also vary. The capacity of leaf extracts to quench 1O2 can be measured for example according to [143].
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Modifications of the amount or lifetime of 1O2. As the lifetime of 1O2 is lengthened 5‒20 fold in D2O compared to water [46,72], measurements can be done in D2O to facilitate 1O2 detection (e.g. [72]). However, the use of D2O in place of H2O has consequences in biological materials. Proteins are often more stable and more rigid in D2O than in water [144]. It has been shown that with photosynthetic organisms D2O reduces growth and oxygen evolution as well as slows down PSII electron transfer [145] and carbon fixation [146]. Chl is a natural sensitizer of 1O2 in photosynthetic systems but the amount of 1O2 can be increased by using with artificial 1O2 (photo)sensitizers [147]. However, the popular 1O2 generators Rose Bengal and methylene blue have been shown to affect PSII electron transfer [148]. Different 1O2 generators localize in different cellular compartments. For example, Neutral Red and Rose Bengal penetrate to the chloroplast, in contrast to methylene blue [148]. If the sensitizer penetrates weakly to the target tissue, then the amount of 1O2 produced may not be physically relevant. The extent of protection against cellular damage, obtained by a treatment with an 1O2 scavenger like azide, histidine or DABCO, has also been used to assess the importance of 1O2 in the damage mechanism (e.g. [149]). Due to the high reactivity of 1O2, scavengers that localize to the site of 1O2 production, often within a membrane, are preferred. Effects of natural quenchers. Carotenoids and tocopherols that function as the principal quenchers and scavengers of 1O2 can often be used to draw conclusions about the participation of 1O2. For example, participation of 1O2 in the oxidative inhibition of the repair of PSII after photoinhibition was supported by the finding that both in plants [150] and in cyanobacteria [151] the ability to synthesize -tocopherol protects the PSII repair mechanism but does not affect the rate of light-induced damage. The same tocopherol data suggested that 1O2 is not involved in the damaging reaction of photoinhibition, but on the other hand, a cyanobacterial mutant with a high carotenoid content is less sensitive to photoinhibition than the wild type [130].
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3. Hydrogen peroxide, H2O2 3.1. Definition and properties of H2O2
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2.5.9. Summary of 1O2 detection To conclude, 1O2 measurements are a tricky business. Direct measurements of 1O2 are still not possible from physiological conditions. When using chemical traps (see Table 3) it is important to check for the possible adverse effects on the organism or biological function, in addition to checking the sensitivity and reliability of the probe. The place where the sensor compound localizes can also affect the results. More 1O2 probes exist than are used in plant research and also new probes are being synthesized but the usability of the new probes in photosynthetic systems should be tested before extensive use [117,152,153]. For example, the 1O2 probe MVP (trans-1-(2’methoxyvinyl)pyrene) has long been used e.g. in medicine, but it could not detect 1O2 from leaves, maybe because its chemiluminescence (465 nm) overlaps with Chl a fluorescence [109].
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H2O2 is the result of reduction of oxygen by two electrons. In biological systems, H2O2 is mostly present in the neutral form because of its high pKa value (pKa1 is 11.8). H2O2 is a moderate oxidant (E0 of the pair H2O2+2H++2e-/2H2O is 1.776 V; [154]) that can oxidize compounds like thiols.
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3.2. Formation and action of H2O2 in photosynthetic tissues
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In biological systems, including photosynthetic tissues, H2O2 is mainly formed via dismutation of O2• (reaction 29), and because superoxide dismutases (SODs) are present in most cellular compartments, the formation of H2O2 depends on the formation of O2•. Furthermore, the direct 2electron reaction between 1O2 and plastohydroquinone was recently shown to produce a stoichiometric amount of H2O2 (Fig. 4; [60]). The production of H2O2 by the interaction of 1O2 and plastohydroquinone may have importance in chloroplast signaling because H2O2 produced within the membrane would avoid the stromal scavenging mechanisms [60]. In the soluble phase, 1O2 produces H2O2 in a reaction with ascorbate [59]. H2O2 can be enzymatically scavenged by catalases (CATs), peroxidases and peroxiredoxins. CATs have a relatively low affinity for the substrate, reflected by a high Michaelis constant Km for H2O2; usually around 50 mM in plant catalases [155,156], although a lower value of 17.6 mM was also reported [157]. Peroxidases have lower Km values (0.57 mM for ASC, 0.11 mM for H2O2; [158]). Ascorbate and GSH function as electron donors for different peroxidases. Peroxiredoxins form a third group of H2O2 scavenging enzymes. In peroxiredoxins, the oxidizable thiol group is a cysteine residue of the protein itself, and therefore peroxiredoxins require an activating reduction step between the catalytic rounds. H2O2 is known to be associated with reactions causing cellular damage. For example, the action of the herbicide paraquat, known also as methyl viologen, is based on formation of O2•, which inevitably leads to formation of H2O2, too. However, it is usually difficult to determine if the actual damaging agent is H2O2, O2• or HO• formed in the Fenton reaction (reaction 15). Cyanobacteria are highly sensitive to H2O2 [159] and this sensitivity can actually be used to destroy potentially poisonous cyanobacteria from lakes [160].
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In plants, H2O2 functions, together with O2•, as a signal molecule in stomatal opening, programmed cell death and various stress responses (for review, see [161]).
3.3. Reactions of H2O2
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H2O2 is particularly reactive against thiol groups [162], especially those of cysteine residues of proteins and other compounds [16], and the reactivity of H2O2 towards thiols explains the inhibitory effect of H2O2 on the thiol-group-containing enzymes of the Calvin-Benson cycle. H2O2 inactivates fructose-1,6-bisphosphatase, glyceraldehyde 3-phosphate dehydrogenase and xylulose-5-phosphate kinase [163] and the Krebs cycle enzyme aconitase [164] and reacts with several other plant proteins [165,166]. The Escherichia coli transcription factor OxyR recognizes H2O2 via a reaction with a cysteine residue [16,167]. The reaction of H2O2 with thiol groups is also a H2O2 elimination mechanism, accomplished by peroxiredoxins in plant cells [13]. Reduction of H2O2 by thiol groups is a two-step reaction passing through the mechanism described by reactions 12.1 and 12.2.
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H2O2 + RSH RSOH + H2O RSOH + RSH RSSR + H2O
(12.1) (12.2)
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The rate constants of reaction 12.1 range from 102 to 107 M-1 s-1. The thiolate anion (RS), formed in high pH, reacts more rapidly with H2O2 than the respective thiol (RSH) [162,168], and the reactivities of thiols depend on both pH and the pKa value of the thiol [162].
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A well-known reaction of H2O2 is its reduction via the Haber-Weiss mechanism (sum of reactions 13 and 14) and Fenton mechanism (reaction 15). Both mechanisms generate the very powerful oxidant HO•. The rate constant of reaction 13 in aqueous medium ranges from 0.13 to 0.23 M-1 s-1 [169,170] and the rate constant of reduction of H2O2 by HO2• (reaction 14) is 0.5 M-1 s-1. Thus, reactions (13) and (14) are relatively slow in biological material where concentrations of O2• and H2O2 are low. O2• + H2O2 O2 + OH + HO•
(13)
HO2• + H2O2 O2 + H2O + HO•
(14)
M+(n-1)+ H2O2 M+n + OH + HO•
(15)
The reduction of H2O2 by transition metals via the Fenton mechanism (reaction 15) is much more efficient than the Haber-Weiss reaction (reactions 13 and 14). The rate constant of Fenton reactions depend on the metal M (Fe and Cu are the biologically most important ones) or metal complex and pH, and range from 50 to 104 M-1 s-1 [171‒174]. The reduction of H2O2 via the Haber-Weiss and Fenton mechanisms (reactions 13‒15) causes subsequent reactions where interactions between O2•, HO2•, HO• and the reduced and oxidized metal ion can occur. In particular, O2• is able to reduce Fe3+ to Fe2+ and Cu2+ to Cu+, thereby promoting continuing production of HO• via reaction (15). The reduction of H2O2 by peroxidases is one of the main routes of H2O2 scavenging; organic compounds including ascorbate, glutathione and guaiacol can be used as electron donors. In the chloroplast, H2O2 scavenging is mainly catalyzed with an ascorbate-specific peroxidase (APX, (EC
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1.11.1.11)). The catalytic reduction of H2O2 by peroxidases proceeds via the peroxidase ping-pong mechanism (reactions 16‒18) [175]. PX-Fe(III)-P + H2O2 PX-Fe(IV)=O-P+ + H2O
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(16)
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PX-Fe(IV)=O-P+ + AH PX-Fe(IV)=O-P + A• +H+ PX-Fe(IV)=O-P+ AH PX-Fe(III)-P + A• + OH,
(18)
SC
where P is porphyrin.
(17)
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The rate constant of the reaction of H2O2 with APX is around 107 M-1 s-1 and the Km for H2O2 is 80 M [176,177]. Thus, APX is an efficient enzyme with high affinity to the substrate and a very fast rate of the reaction. CAT-depended scavenging of H2O2 also occurs through the peroxidase-like ping-pong mechanism where one H2O2 molecule is used as an electron donor (reactions 19.1 and 19.2). The decomposition of H2O2 by CAT, unlike the reaction catalyzed by peroxidase, is associated with O2 production. (19.1)
H2O2 + Fe(IV)=O-CAT+→ H2O + Fe(III)-CAT + O2
(19.2)
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H2O2 + Fe(III)-CAT H2O + Fe(IV)=O-CAT+
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The peroxidase-catalyzed reaction of H2O2 with a specific electron donor like luminol can be used for detection of H2O2. In this case, electron donor is oxidized to an optically detectable product (see 3.5.5). In addition to the above, H2O2 irreversibly inactivates two of the three types of SOD (Cu,ZnSOD, Fe-SOD), and APX in the absence of ascorbate [178].
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3.4. Lifetime and diffusion distance of H2O2 As a small, neutral molecule, H2O2 is expected to readily diffuse through membranes. Judging from experiments with the genetically encoded probe HyPer, the lifetime of H2O2 is 1‒3 min in mammalian cells [179] and around 10 s in Arabidopsis thaliana guard cells [180]. Both lifetimes are so long that 5 % of H2O2 would be present at 0.2‒1 mm from the site of generation in a viscous biological material.
3.5. Detection of H2O2 3.5.1. Principles of H2O2 detection H2O2 can be detected with several different compounds functioning as hydrogen donors for the reduction of H2O2. Some detector substances require a catalyst, usually a peroxidase or a metal ion, and if the study object does not contain endogenous peroxidases, horseradish peroxidase can be used together with the detector substance. 3.5.2. Precipitate-forming compounds DAB. The most widely used H2O2 detection chemical in plant research has been 3,3'diaminobenzidine (DAB) (see Table 4 for a list of chemicals used for H2O2 detection). DAB reacts with H2O2, forming a brown precipitate that can be made visible in plant material by removing Chl (Fig. 5; [181‒183]). DAB penetrates well to plant cells and it is not sensitive to light [184], the precipitate is stable [183], and DAB localizes partially to the chloroplast [185]. Unfortunately, DAB strongly lowers
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the quantum yield of PSII electron transfer in the concentration range required for imaging [184]. The inhibition of photosynthesis by DAB should be taken into account in the interpretation of results obtained with DAB from photosynthetic material.
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Cerium. H2O2 reacts with cerium chloride, forming electron-dense precipitates of cerium perhydroxides [186]. This has been used to localize H2O2 production e.g. in wheat roots [187] and Zinnia elegans stems [188] from electron transmission microscope images.
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3.5.3. Dyes MBTH/DMAB. A peroxidase-catalyzed oxidation by H2O2 couples 3-methyl-2-benzothiazoline hydrazone (MBTH) and 3-(dimethylamino) benzoic acid (DMAB) to a deep purple indamine compound that can be assayed spectrophotometrically at 590 nm [189]. Veljovic-Jovanovic et al. [190] showed that a plant leaf extract causes, in addition to a reaction caused by the H2O2 present in the extract, slow but significant unspecific increase in 590 nm absorption. The unspecific increase could be avoided with polyvinylpyrrolidone treatment. Furthermore, ascorbate inhibits the MBTHDMAB signal [190]. The effects of ubiquitous plant metabolites on the MBTH-DMAB signal call for special care to assure that the signal actually reflects the H2O2 content of the study material. The MBTH/DMAB method has been used to measure the increase in extracellular peroxidase activity in sunflower roots during defense signaling [191].
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4-aminoantipyrine/phenol. H2O2 can be measured with 4-aminoantipyrine and phenol as H2O2 oxidatively couples with them forming colourful quinone-imine (absorbance at 505 nm) [192]. Peroxidase is needed for the reaction. The method has been used to measure H2O2 from pea chloroplasts [193] and from root extracts of tomato [194].
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Starch/KI. Iodide ions (I-) are oxidized by H2O2 after which iodine (I2) is complexed with (exogenous) starch, and H2O2 can be histochemically localized by the blue color of the starch/I-complexes. Starch and iodide do not penetrate to cells [188,195] but the starch/KI-method has been used to detect H2O2 production in cut stem surfaces [195] and in peeled or partially digested leaf segments [196] or to measure extracellular H2O2 production by whole roots [197]. Age and wounding of the plant affect the results obtained with the starch method [188]. Iodide can be oxidized also by organic peroxides (ROOH) [198] and other cellular electron acceptors besides H2O2 [195]. DHBS. In the presence of peroxidases 3,5-dichloro-2-hydroxybenzenesulfonic acid (DHBS) is oxidized by H2O2 to its quinone form which then reacts with 4-aminoantipyrine (4-AAP) to produce a colorful substance that can be measured spectrophotochemically at 510 nm. The DHBS method has been used to measure extracellular H2O2 from a tobacco cell suspension [199]. 3.5.4. Fluorescent probes Scopoletin. Decrease of the fluorescence yield of scopoletin (7-hydroxy-6-methoxy-2H-1benzopyran-2-one) is a traditional in vivo method of H2O2 detection in biological systems [200]. In plant leaves, scopoletin mainly localizes to the cell wall but also penetrates to the cells [184]. The usability of scopoletin in plants in vivo is limited by the need to use ultraviolet (UV) wavelengths (346 nm) for excitation, as plant leaves strongly absorb UV light [201]. The 435 nm emission of scopoletin is also strongly absorbed by Chl a. Scopoletin has been used for measurement of H2O2 in root cells
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[202] and in the incubation media of callus cells [203] but also from leaves of Pseudowintera colorata [204].
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Eu3Tc. An europium-tetracycline complex (Eu3Tc) [205] reacts with H2O2 to form a highly fluorescent adduct (excitation at 360‒440 nm; emission at 616 nm). The fluorescence yield of Eu3Tc-H2O2 adduct is strongly dependent on pH [205] and therefore the complex should be used only in a buffer solutions at pH 6.6‒7.2. Eu3Tc is not photosensitive but lowers PSII yield [184]. Eu3Tc has not been used with other photosynthetic materials.
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Amplex Red and Amplex Ultrared. Amplex Red is 10-acetyl-3,7-dihydroxyphenoxazine which, after oxidation by H2O2, fluoresces at ~580 nm when excited at 570 nm; Amplex Ultrared is a derivative of the original reagent. The commercial kits also contain horseradish peroxidase as a catalyst. Both compounds have been tested for H2O2 imaging from leaves, and Amplex Red penetrates to cells and even to chloroplasts whereas Amplex Ultrared was found to stay mostly in the apoplast [184]. Both compounds have a strong negative effect on PSII yield [184], which limits their use in plant material. Furthermore, both compounds are sensitive to light [184]. Cyanide, salicylhydroxamic acid, and propyl gallate have been shown to inactivate the Amplex Red method in vitro [206]. Antal et al. [207] used Amplex Ultrared to measure H2O2 formation by isolated thylakoid membranes after treatments with short flashes in the dark.
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3.5.5. Chemiluminescent probe Luminol. 3-amino-phthal-hydrazide, known with the common name luminol, is oxidized by H2O2 in a peroxidase-catalyzed reaction to produce an excited state of 3-aminophthalate; the decay of the excited state yields blue luminescence peaking at 440 nm. Reaction with H2O2 requires a peroxidase or a metal ion as a catalyst, and a mixture of luminol and H2O2 has been used from the 1930's in crime scene investigation to detect the iron of haemoglobin present in blood stains. If the catalyst is provided, luminol can be used to measure H2O2, and a detection limit of 0.42 nM was obtained for cobalt-catalyzed measurement of H2O2 from seawater [208]. The luminescence of luminol is quenched by many cellular substances like ascorbic acid and phenols, which limits the usability of luminol for H2O2 measurements from intact biological materials [190]. Luminol chemiluminescence is also observed in the presence of peroxynitrite and O2• ([209]; see section 4.5.5). Plastoquinone was recently found to enhance the luminescence [60]. However, the Co(II) catalyzed reaction offers the very high sensitivity of 0.5 nM for plant extracts diluted to such extent that the quenchers and enhancers do not disturb the reaction [210]. The luminol/Co(II) method revealed an increase in H2O2 concentration in Arabidopsis thaliana leaf tissue after a shift to high light [211] and a hypoxiainduced increase in the H2O2 level of grapevine buds [212]. 3.5.6. Microelectrodes Microelectrodes. Electrochemical H2O2 microsensors (for O2• electrodes, see 4.5.6) have been developed (for a review see [213]). Ren et al. [214] developed a carbon fiber Pt-ultramicroelectrode coated with single-walled carbon nanotubes and hemoglobin (Hb/SWCNTs/CFUME), and were able to monitor the oxidative burst (H2O2) in Aloe leaves in vivo with a detection limit of 4 μM. Neither 0.1 mM ascorbic acid nor NaCl affected the electrode. A microelectrode with poly-ophenylenediamine and Pt microparticles (POPD–Pt-MP–Pt) has been used to measure H2O2 induced by Cd2+ treatment from leaves of oilseed rape in vivo [215]. Another Pt-electrode has been used in
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vitro with Arabidopsis thaliana cell suspensions [216]. The small size of current electrodes (diameter less than 10 µm [214]) facilitates measurements from intact plants.
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3.5.7. Genetically encoded probes With genetically encoded sensors there would be no need to get foreign molecules inside the cells to detect H2O2 (or O2•, see 4.5.7) and they also offer ways for locating the ROS in the cell, and the use of (fluorescence) microscopy (for reviews see [217,218]). The ability of H2O2 to oxidize the thiol group of OxyR [167]; reactions 12.1 and 12.2] was used in designing the H2O2 sensor HyPer, in which circularly permuted yellow fluorescent protein (cpYFP) was fused with the regulatory domain of OxyR [21]. HyPer functions as a genetically encoded H2O2 sensor. Hyper is claimed to be specific to H2O2 [21], and it has been used also in Arabidopsis thaliana to measure H2O2 produced in the peroxisome lumen and in the cytoplasm [180]. HyPer has excitation maxima at 420 and 500 nm and emits at 516 nm, and the oxidation of HyPer by H2O2 enhances the 500 nm peak and lowers the 420 nm peak. However, the sensor (like many fluorescent proteins) shows strong pH-dependence [179], and therefore the measurement should be coupled with measurement of pH. Many fluorescent proteins themselves produce ROS when illuminated, which may lead to photobleaching [219,220], but Hyper appears to show little photobleaching [221]. Another issue with genetically encoded probes is that they act as antioxidants, and their continuous expression may affect the physiology of the organism [222]. Recently published genetically encoded H2O2 probes (OxyFRET and PerFRET) respectively apply the Orp1 and Yap1 proteins of the H2O2 sensing system of yeast [223]; these probes have not yet been tested with plant material.
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3.5.8. Unspecific probes H2O2 has been measured from plant material [224,225] and from Chlorella cells [226] with fluorescein derivatives that react primarily with HO• [227]. Because these dyes can be used to detect more than one ROS, they will be described in Chapter 8. 3.5.9. Indirect methods of H2O2 detection Increase in the activities of H2O2 scavenging enzymes, including APX and CAT [180] or increase in the expression levels of genes coding for these enzymes may suggest an increase in the amount of H2O2 in the tissue. A considerable number of genes are affected by H2O2 in plants [136,228] and in cyanobacteria [229]. Many of these genes are also regulated by light [230], and plant chloroplasts produce H2O2 in the light. Purified H2O2 scavengers offer possibilities for simple measurements of H2O2 in vitro. For example, H2O2 production by a cell suspension has been measured via the reaction of a scavenger [231]. 3.5.10. Summary of H2O2 detection Some methods of H2O2 determination have been used long, and lots of knowledge about different aspects of their use has been accumulated whereas the caveats in newer, promising methods may still remain to be found. Unfortunately, most detection methods that have been studied with regard to their effects on photosynthesis seem to have adverse effects. This is particularly important for DAB, as this dye localizes to the chloroplasts in DAB-infiltrated leaves. Convincing determination of H2O2 requires the use of more than one method, and measurements of H2O2 from extracts are recommended to back up in vivo staining experiments.
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Specificity
Stability
Brown precipitate, Imaging, semiquantitative
Specific
Stable
Cerium
Electron transmission microscopy Absorbance at 590 nm
-
-
Ascorbate inhibits the signal [190]
-
Absorbance at 505 nm
-
Histochemical staining
Oxidized by ROOHs and cellular electron acceptors [195,198] -
Toxicity
Hydrophobicity (LogP) 0.24 (estimated)
Localization
Notes
Cellular; penetrates to chloroplasts [184] -
Too low sensitivity at acceptable concentration [184]
-
-
-
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Strongly lowers PSII yield [184]
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-
1.41 MBTH (estimated)/ 1.35 DMAB (estimated)
In vitro or from ground tissue
-
-
-
-
-
-
Peroxidase is needed for the reaction
-
-
-
1.04 iodide [19]
Cellimpermeable [195]
Wounding affects the results
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-
-
-
-
Peroxidase is needed for the reaction
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DHBS/4-AAP (3,5- Absorbance at 510 dichloro-2nm hydroxybenzenes
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MBTH (3-methyl2benzothiazoline hydrazone) and DMAB (3(dimethylamino) benzoic acid) 4aminoantipyrine/ phenol Starch/KI
Light sensitivity Not sensitive [184]
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Method
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Compound (IUPAC name) DAB (3,3'diaminobenzidin e)
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Table 4. Methods of detection of H2O2. Light sensitivity data are for visible light, unless otherwise indicated. The stability data indicate the stability of the signal after reaction with H2O2 in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data.
-
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-
-
Signal affected by several metabolites and pH [205] -
Decrease Not sensitive to <50 % [184] in 4‒5 min in leaves t1/2 = 6‒8 min
Amplex Ultrared (modified from Amplex Red)
Fluorescence (Ex 567/ Em 580), Imaging
-
Luminol (3amino-phthalhydrazide)
Chemiluminescence at 430 nm
Hb/SWCNTs/CFU ME (Microelectrode
Amperometry
Interference by ascorbate, phenolics [190] and plastoquino ne [60] -
1.62 (estimated)
Cell wall, cytoplasm [184]
-
Lowers PSII yield [184]
-
Mostly intercellular [184]
-
Strongly lowers PSII yield [184]
1.93 (estimated)
Addition of peroxidases may be needed.
Strongly lowers PSII yield [184]
-
At least partly cellular, accumulates in chloroplasts [184] Limited penetration to cells [184]
-
1 % increase in signal in 30 min at PPFD 1200 µmol m2 -1 s [184] 30 % increase in signal in 30 min at PPFD 1200 µmol m2 -1 s [184] -
-
-0.41 (estimated)
In vitro or from ground tissue
Use of Co as catalyst facilitates measurements from plant tissues [210]
-
-
-
-
-
Not affected by ascorbic acid or NaCl [214]
-
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TE D
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Amplex Red (10acetyl-3,7dihydroxyphenox azine)
-
CR
Fluorescence (Ex 346/ Em 435) quenching, Quantification, imaging Reversible formation of fluorescent (Ex 400/ Em 615) Eu3Tc-H2O2 complex, Imaging Fluorescence (Ex 570/ Em 583), Imaging
Europiumtetracycline complex (Eu3Tc)
IP
ulfonic acid/ 4aminoantipyrine) Scopoletin
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31
Linear decrease to <50 % in 15 min
Addition of peroxidases may be needed
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IP CR
Only little photobleachin g [221]
-
-
Can be targeted to desired tissue in vivo
Fluorescence depends strongly on pH [179].
-
-
Can be targeted to desired tissue in vivo
Moderate pH sensitivity [223]
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-
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TE D
Fluorescence ratio (Ex 435 nm, Em 535 nm/480 nm)
Specific to H2O2 [21] but may be affected by other oxidants [179] Specific to H2O2 in vivo [223] but not tested with isolated protein
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OxyFRET and PerFRET (genetically encoded probes based on Orp1 and Yap1, respectively).
Fluorescence ratio (Ex 420/500 nm, Em 516 nm)
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based on haemoglobin and carbon nanotubes) HyPer (genetically encoded probe based on OxyR)
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4. Superoxide anion radical, O2•, and hydroperoxyl radical, HO2• 4.1. Definitions and properties of O2• and HO2•
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Reactions and biological activities of O2• have interested the scientific community for tens of years, and several reviews are available (see [232‒235]). O2• is 1-electron reduced form of O2. The midpoint redox potential (Em) of the pair O2/O2• at pH 7.0 is -160 mV vs Normal Hydrogen Electrode (NHE) in aqueous solution [236]. The Em of the pair O2/O2• becomes more positive if O2• is protonated to form HO2•, and therefore HO2• is a stronger oxidant than O2• [237]. However, O2• is a weak deprotonation agent in aqueous solutions. O2• can form O2•(H2O)n complexes (with n from 1 to 3 [233] and the free energy of hydration was estimated to be around 355 kJ mol-1 [238]. The pKa value for HO2• is 4.8 and therefore only 0.25 % of O2• is protonated at pH 7. In aprotic medium O2• forms a weaker solvation complex. The pKa value of HO2• in DMF was estimated to be 12, and therefore O2• can act as a strong deprotonation agent in an aprotic medium. Weak solvation of O2• in aprotic media leads to a shift of the redox potential of the pair O2/O2• in comparison with an aqueous solution. The redox potential of the pair O2/O2• in DMF was estimated to range between -550 and -600 mV vs NHE [233].
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In addition to its role as an oxidant in reactions that produce H2O2 (see reactions 32.1, 34 and 35 below), O2• can act as a reductant, which leads to conversion of O2• to O2 (reactions 23, 24, 32.2 and 33); see [237] for a compilation of reactions of O2• and HO2•. Enzymatically catalyzed dismutation of O2• proceeds via oxidation of one O2• ion and reduction of another one. The dual role of O2• is unique among ROS, as other ROS function as oxidants in biological environment.
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4.2. Formation and action of O2• and HO2• in photosynthetic tissues 1-electron reduction of oxygen by PSI is the main photosynthetic source of O2•, but small amounts of O2• are also produced by PSII [2,239] and by reduced forms of plastoquinone [60]. O2• is invariably scavenged by SODs (EC 1.15.1.1), enzymes catalyzing the dismutation reaction (reaction 29). Almost all aerobic organisms contain SOD(s), and the enzymes fall in the phylogenetic groups of Mn and Fe-SODs, Cu,Zn-SODs and Ni-SOD [240]. The Mn and Fe-SODs form one clade. Chloroplasts contain mainly Cu,Zn-SODs [240]. All SODs speed up O2• dismutation, but the reaction proceeds even in the absence of an enzyme. In addition to SODs, redox active metals like manganese ions catalyze the dismutation.
4.3. Reactions of O2• and HO2• According to its thermodynamic properties, O2• expresses dual reactivity; it acts as a weak oxidant and as a strong reducing agent in electron transfer reactions in aprotic media in the absence of proton donors. The redox potential of O2•/HO2 (data are available only for aprotic medium) is very low, -1.75 V in DMSO [232]. On the other hand, O2• is able to oxidize organic molecules via deprotonation-oxidation mechanism in the presence of protons or proton donors. Thus, the behavior of O2• in redox reactions very strongly depends on its protonation. Furthermore, O2• reacts preferably with neutral or radical partners and only slowly with negatively charged ones.
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In addition to electron-proton transfer reactions, O2• can take part in nucleophilic substitution or addition if the substrate has a low electron affinity. O2• participates in nucleophilic reactions with alkyl halides, sulfonates, phosphates, esters, acyl halides, acyl anhydrides to form peroxyl (ROO•), alkoxyl (RO•) radicals and epoxides as intermediates [232,233,241,242]. Both the nucleophilicity and the basicity of O2• depend on medium. O2• acts as a powerful nucleophilic agent in aprotic medium due to rapid protonation and dismutation in aqueous solutions.
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Reactions of O2• with organic and inorganic molecules fall in five basic categories: protonation, electron transfer reaction, nucleophilic substitution and addition, attraction of hydrogen, and addition to metal or metal complex.
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Protonation. Protonation of O2• is a highly important reaction because this reaction is the main source of HO2• and is involved in a complicated deprotonation-oxidation process as well. O2• can be protonated via a reaction with proton or a proton donor such as ascorbic acid, reactions (20) and (21) respectively. O2• + H+ HO2• O2• + AscH2 HO2• + AscH
(20) (21)
(22)
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O2• + -TocH HO2• + -Toc
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O2• can be also protonated by -tocopherol (-Toc) in aprotic medium, reaction (22):
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Electron transfer. O2• can reduce many organic and inorganic molecules such as quinones and cytochromes, reactions (23) and (24), respectively, via a 1-electron transfer mechanism. O2• + Q O2 + Q•
(23)
O2• + Cyt c(Fe3+) O2 + Cyt c(Fe2+)
(24)
Nucleophilic addition and substitution. Nucleophilic reactions of O2• with organic molecules are associated with formation of corresponding peroxy radicals which are more oxidizing than O2• itself. For example, O2• reacts with carbon tetrachloride (CCl4) to form the trichloromethyl peroxy radical [243], reaction (25). O2• + CCl4 Cl3COO• + Cl
(25)
Nucleophilic reaction of O2• with nitrones is widely used for detection of O2• with EPR. The reaction of O2• with 5-diethoxyphosphoryl-5-methyl-1-pyrroline N-oxide (DEPMPO) yields a stable DEPMPO/O2• adduct (reaction 26) [244].
(26)
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Attraction of hydrogen. Attraction of a hydrogen atom from organic molecules by O2• is an unlikely reaction. Most probably O2• dependent attraction of hydrogen is the result of its protonation. HO2• is a typical neutral radical and is able to attract a hydrogen from organic molecules such as linolenic acid [237], reaction (27).
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HO2• + LH H2O2 + L•
(27)
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Addition to metal of metal complex. A reaction of O2• with transition metals or metal complexes yields unstable and stable intermediates [233,237]. For example, O2• can react with manganese complexes Mn2+-L to form MnOO+L complexes [245], reaction (28). Mn2+-L + O2• MnOO+-L
(28)
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Complex reactions involving O2•. O2• is involved in many complicated reactions. Below we will describe only the most important O2• depended reactions occurring in photosynthetic organisms.
O2• + O2• H2O2 + O2
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Dismutation is one of the main reactions determining the lifetime of O2• and its possibilities to take part in other biochemical processes. Dismutation is the reaction between two molecules of O2• and can occur via noncatalytic and catalytic routes. Noncatalytic dismutation of O2• proceeds, in general, in aqueous medium and it can be written as a reaction (29). Dismutation is a 2-step reaction; the first is protonation of O2• (reaction 20), and the second step is a radical-radical reaction between O2• and HO2• or between two molecules of HO2•, reactions (30) and (31) respectively. (29) (30)
HO2• + HO2• H2O2 + O2
(31)
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O2• + HO2• + H+ H2O2 + O2
Dismutation of O2• is very slow in alkaline pH (0.3 M-1 s-1) because O2• is mainly deprotonated. Protonation of O2• leads to an increase in the reaction rate constant with a maximum (108 M-1 s-1) at pH 4.8, the pKa value of HO2•. In acidic pH less than 4.8, when O2• is mostly protonated, the reaction rate constant is reduced to 8.6 x 105 M-1 s-1 [237]. The catalytic route of O2• dismutation involves the SOD enzyme (EC 1.15.1.1). The general reaction can be written as follows: M(n+1)+-SOD + O2• → Mn+-SOD + O2
(32.1)
Mn+-SOD + O2• + 2H+ → M(n+1)+-SOD + H2O2,
(32.2)
where M is a metal ion present in SOD. The rate constant of O2• dismutation catalyzed by SOD is about 6.4 x 109 M-1 s-1 [246]. O2• and HO2• can react with H2O2 via the Haber-Weiss reaction (reactions 13 and 14). O2• can also mediate the reduction of H2O2 in the Fenton reaction (reaction 15) by reducing a transition metal ion, e.g. Fe3+ (reaction 33) [247].
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O2• + Fe3+ O2 + Fe2+
(33)
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In addition to the Haber-Weiss reaction, O2• can directly participate in decomposition of ROOHs. O2• can react with ROOH yielding the ozonide anion radical and aldehydes or ketones as final products [248]. O2• is able to react with benzoyl peroxide in toluene with formation of 1O2 [249]. In cells, decomposition of ROOH by O2• might be observed in hydrophobic regions like membranes.
O2• + AscH2 + H+ → H2O2 + AscH•,
(34)
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where AscH• is monodehydroascorbate radical.
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In cells, O2• is particularly reactive against iron-sulfur centres, and the ability of the Escherichia coli transcription factor SoxR to sense O2• is based on this reactivity [16]. Ascorbic acid and reduced GSH can participate in scavenging of O2•. The oxidation of ascorbic acid by O2• is a complicated reaction concerted with transfer of a hydrogen atom and a proton [250]. The general reaction of O2• with ascorbic acid (reaction 34) can be represented in the following way:
O2• + GSH + H+ → H2O2 + GS•
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O2• can react with reduced GSH to form H2O2 and gluthyl radical (GS•), reaction (35). Addition of O2 to GS• is considered as a source of peroxysulfenyl radical (GSOO•) [251]. (35)
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The rate constant of the reaction of O2• with reduced GSH was estimated to be 103 M−1 s−1 at maximum [162,252]. It is much less than the rate constant for the reaction of O2• with ascorbic acid, 3.3 x 105 M−1·s−1 at pH 7.8 [253].
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The ability of O2• to bind with transition metal complexes determines its toxicity to metalloenzymes. O2• can inactivate the ferredoxin-dependent nitrate reductase, interacting with molybdenum in its active center [254]. CAT passes into an inactive form due to interaction of O2• with its heme [255].
4.4. Lifetime and diffusion distance of O2• High pH and low concentration of O2• increase its lifetime by affecting the dismutation rate (reactions 29‒30; [256]). According to Fridovich [257] the lifetimes of 0.1 mM and 0.1 nM solutions of O2• in water are 0.05 s and 14 h, respectively. In seawater, 87‒1120 pM concentrations of O2• have been measured, with half-life of 10‒100 s [258,259]. In animal cells the half-life of O2• has been calculated to be below 50 ms, resulting in 40 µm diffusion distance [260,261]. To our knowledge, no measurements of the lifetime of O2• in plant cells have been published; in general, the lifetime would be controlled by SOD (reaction 32; [257]). Due to its negative charge, O2• cannot easily pass through membranes [262], and in cellular pH only a minor fraction would be in the protonated form. Plant PSII particles have been shown to produce 1‒2 µmol O2• (mg chl)-1 h-1 in high light [263], and extracellular O2• production by Chlamydomonas in UV-light was measured to be 0.60‒2.40 µmol g-1 min-1 [264].
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4.5. Detection methods of O2•
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4.5.1. Spectroscopic methods O2• can be detected directly by spectroscopy (absorption at 230–350 nm; [265]) or with EPR spectroscopy [266], but due to the strong absorption of cell components at UV-range and the low amount of O2•, these methods are rarely useful in biological applications. Changes in the EPR spectrum, induced by UV treatment of attached plant leaves, were attributed to O2• [267], but the origin of these signals is controversial [268].
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4.5.2. EPR detectable probes DMPO, EMPO, DEPMPO, BMPO, DCP, TMT and Tiron. O2• produced by photosynthetic organisms has often been measured with probes that, after reacting with O2•, can be detected with EPR spectroscopy (for reviews, see [269,270]; see Table 5 for a list of chemicals used for detection of O2•). The most widely used probes are 5,5-dimethyl-1-pyrroline N-oxide (DMPO), 5(diethoxyphosphoryl)-5-methyl-1-pyrroline-N-oxide (DEPMPO; reaction 26), 5-(ethoxycarbonyl)-5methyl-1-pyrroline N-oxide (EMPO) and 5-tert-butoxycarbonyl-5-methyl-1-pyrroline N-oxide (BMPO; Fig. 6), all of which are commercially available. DMPO has been used to detect extracellular O2• production by intact cells of a red alga Chattonella marina [271] and with EMPO, O2• has been detected from isolated PSII membranes of spinach [272]. DEPMPO has been used for O2• detection from isolated tobacco thylakoids [273], from reaction center complexes of spinach [274] and from isolated plasma membranes of maize [275]. BMPO has been used to detect O2• from cell wall extracts of pea [276] and from isolated PSII membranes of spinach [277]. The lipophilic cyclic hydroxylamine probe 1-hydroxy-4-isobutyramido-2,2,6,6-tetramethylpiperidinium (TMT) and the more hydrophilic probe 1-hydroxy-2,2,5,5-tetramethylpyrrolidine-3,4-dicarboxylic acid (DCP) have been used to detect O2• from pea thylakoids [278]. In comparison to the traditional EPR probes, also commercially available TMT and DCP are said to be more sensitive, and can be also used to detect the location of O2• production [278,279], although TMT might be re-reduced by photosynthetic electron transport chain [278]. The spin trap 4,5-dihydroxy-1,3-benzene-disulfonic acid disodium salt (Tiron) has been used to detect O2• from wheat roots [280] and from lichen tissue [281]. OXANOH and PTM-TC. The spin traps 2-ethyl-1-hydroxy-2,5,5-trimethyl-3-oxazolidine (OXANOH) and perchlorotriphenylmethyl radical-tricarboxylic acid (PTM-TC) have been used to detect O2• from spinach thylakoids [282,283], respectively, and from plant roots [284]. O2• induces oxidation of OXANOH to OXANO, but it is also auto-oxidized in the presence of metal-ions [282]. In contrast to most of the spin traps, the EPR signal of PTM-TC decreases in the presence of O2•. OXANOH and PTM-TC are not commercially available. Specificity. DMPO, EMPO, BMPO and DEPMPO are not specific to O2• but react also with other radicals (especially HO• and carbon centered radicals). Fortunately, reaction products of the spin trap with different ROS often have different EPR spectra [273,285‒287] (Fig. 6; see also 5.5.2). Thus, the limited specificity of these EPR probes is not only a drawback but also an advantage, as several ROS can be detected with one experiment. For example, Bogdanovic Pristov et al. [276] used DEPMPO to detect the carbon dioxide radical (CO2•), O2• and HO•. In contrast, the ability of cyclic hydroxylamines (e.g. TMT and DCP) with other than O2• limits the usability of these compounds
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because the different products usually have similar EPR spectra [279]. Interference with transition metal ions can be reduced by using chelating agents [279]. PTM-TC does not react with HO•, ROO•, H2O2, NO•, GSH or with L-ascorbate [288].
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Lifetime of spin adducts. A challenging feature of the EPR probes is that the spin adducts usually decay rapidly [289,290], especially the spin adducts of DMPO (t1/2 is about 45 s; [291]). The spin adducts of DEPMPO and EMPO are more stable, for EMPO t1/2 is about 8 min and for DEPMPO t1/2 is about 14 min [286,287,289], and of these two the EPR spectrum of the EMPO adduct is less complicated [287]. The half-lifetime of the BMPO adduct is 23 min [292]. The spin adduct of PTM-TC is stable [288]. Also Tiron has been reported to be quite stable (t1/2 is about 15 min), but the need for basic (pH 8.5) solution might restrict its use [280]. Efforts have been made to design more stable spin probes (for example [293]). Furthermore, cyclodextrins can be used to stabilize the spin adducts [290]. Cyclodextrins have also been shown not to significantly inhibit photosynthesis when used in appropriate concentrations [290]. With plant material, DMPO is more commonly used as HO• probe (see 5.5.2), however, the DMPO/O2• spin adduct decays to the HO• adduct and also to HO• itself [294]. Furthermore, UV-radiation can cause generation of the DMPO cation radical, which after reaction with water generates the hydroxyl adduct of DMPO [295]. Spin adducts of DEPMPO or EMPO do not decay to the HO• adducts [286,288] or decay only slowly (>20 min with DEPMPO) [269].
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Test of the effects of these probes on photosynthesis are rare. TMT and DCP do not affect the reduction of P700+ (although in the precence of DCMU, TMT was observed to slightly accelerate the reduction of P700+) [278]. Long incubations of plant roots with Tiron exert a negative effect on root growth [296]. Long (6 h) incubations of animal cells with DMPO, EMPO, BMPO and DEPMPO have been shown to slow down growth and cause death of the cells [297]. Higher concentrations than 90 mM of DMPO may be toxic to algae [271]. In vivo detection of O2• with EPR probes is still rare. Warwar et al. [284] were able to obtain onedimensional images of O2• production in a whole plant root using PTM-TC, and they also concluded that the probe stayed in the apoplastic space. More commonly used EPR probes, DMPO, BMPO and DEPMPO are thought to be membrane-permeable [298]. 4.5.3. Dyes NBT, NBD and XTT. Nitro blue tetrazolium (NBT) is a traditional O2• dye that has also been used with plant material. The reaction of NBT with O2• can be followed spectrophotometrically at 550‒560 nm. O2• has been detected with NBT from legume roots [299], pea leaves [300], diatoms [301] and from extracted leaf material of maize [302]. To quantify the signal, the biological material has to be made transparent by removing Chl [299] or centrifuged away [302], but the precipitated product of the NBT-O2• reaction can be histochemically localized. A method for quantification of O2• from stained leaves has been developed [303]. Both NBT and its reaction product are quite water insoluble. A more water soluble O2• probe sodium, 3´-(1-[phenylamino-carbonyl]-3,4- tetrazolium)-bis(4-methoxy-6-nitro) benzene-sulfonic acid hydrate (XTT), commonly used as a cell viability assay [304], has been introduced [305]. XTT has
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been used to detect O2• from tobacco cell suspensions and from cucumber leaf extracts [306,307], respectively.
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NBT can be reduced also by P680 of PSII and CO2- [83,308] and it has also been shown to generate O2•, although this reaction may not be significant [308]. Furthermore, detergents like Triton X-100 increase NBT reduction [309]. A related probe, 4-chloro-7-nitrobenzo-2-oxa-1,3-diazole (NBD), has been reported to be more selective towards O2• [310], but NBD has not been used, to our knowledge, with photosynthetic organisms. Neither is XTT an optimal O2• probe, as it has been shown that Escherichia coli suspensions can reduce XTT also anaerobically (with NADPH:XTT reductases; [311]) and short-chain sugars and phenolics reduce XTT, which makes the use of whole cell material difficult [311,312].
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Cytochrome c. Reduction of (exogenous) (ferri)cytochrome c (Cyt c, commercial) has often been used to detect O2•, as the reduction of Cyt c can be monitored spectrophotometrically (around 550 nm; [313]). The Cyt c method has been used in vitro to measure O2• from PSII-enriched membrane fragments from higher plants [263], from apoplasts of wheat roots [314] and from Chlamydomonas sp. ICE-L suspension [264]. In addition to O2•, Cyt c can be reduced by the components of the photosynthetic electron transport chain [315,316] and by ascorbate, GSH and other reductants [317]. In low and moderate light intensities (at PPFD 5–200 µmol m-2 s-1) Cyt c was mainly reduced by the plastoquinone pool and not by O2• [316]. For this reason, comparison of the results obtained with and without added SOD is always required [263]. To enhance the specificity of Cyt c to O2•, it is possible to use acetylated Cyt c [318], for example with Chlamydomonas in [264]. However, reduced Cyt c can also be reoxidized by many cell components, like cytochrome oxidases and peroxidases and by oxidants like peroxynitrite (ONOO-) [317], which calls for care with the use of the method.
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Epinephrine. Epinephrine (also called adrenaline) has been used to detect O2• as after the reaction with O2•, the absorbance of the reaction product, adrenochrome, can be measured (at about 490 nm; [319]). Epinephrine has been used to measure O2• from chestnut seeds [320] and spinach chloroplasts [313], extracellular O2• production from bryophytes and lichens [281] and roots [191], and mitochondrial O2• production from barley roots [321]. This method is not specific to O2•, as epinephrine has been shown to react also with H2O2 [322], and therefore specificity assays are always needed (see e.g. [191]). In addition, adrenochrome can be metabolized by a NADPHdependent cytochrome P450 reductase [323] and take part in redox cycling with mitochondrial complex I, resulting in fast increase in O2• production [324]. For the latter reason the authors [324] advise to measure only the initial part (60 s) of O2• production after addition of epinephrine. Hydroxylamine. O2• can be detected by following the formation of nitrite from hydroxylamine (NH2OH) with absorbance at 530 nm, but pH should be below 8 to avoid generation of O2• by autooxidation of hydroxylamine [325]. Comparison with measurements in the presence of added SOD may be needed to exclude unspecific reactions. The method has been used to measure O2• from homogenized leaves of maize [326]. Hydroxylamine penetrates well to leaf cells [327]. A drawback of this method is that hydroxylamine is known to reduce the manganese ions of the oxygen-evolving complex of PSII, which leads to loss of PSII electron transfer capacity [327,328].
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4.5.4. Fluorescent probes HE. The commercially available probe hydroethidine (HE; or dihydroethidine DHE) has been widely used to detect O2•, as its reaction product emits yellow to red fluorescence. HE is permeable to both mammalian and plant cells [329,330] and oxidized HE intercalates with DNA, which enhances the fluorescence. HE also has mitochondria-specific analog, Mito-HE or MitoSOX red [331]. Unfortunately, also HE reacts with other substances than O2•, for example with cytochromes, hemeproteins, manganese-porphyrin complex, hypochlorous acid NO and ONOO− [332‒334], but the reaction product of HE and O2• differs from the unspecific reaction products [335]. To enhance the specificity of fluorescence detection, one should use 490 nm light for excitation and detect 560‒570 nm fluorescence. However the spectra of the O2• specific and unspecific reaction products overlap so strongly [335] that reliable detection requires high performance liquid chromatography (HPLC) measurements [335,336]. HE can also be oxidized in UV, and in some conditions also in visible light [337], which prevents detection of light-induced O2• formation. Additional issues weakening the usability of HE are that is has been shown to increase the dismutation rate of O2•, and be toxic for Escherichia coli [332]. Nevertheless, HE has been used to detect O2• from plants, for example from tobacco cells and pea roots with a fluorescence microscope [338] and from spinach leaves with HPLC [339]. For HPLC detection, a simplified method has been developed [340], though, at least to our knowledge, it has not been used with plant material. Martin et al. [341] measured O2• production by Arabidopsis thaliana mitochondria by Mito-HE.
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4.5.5. Chemiluminescent probes Three commercially available chemiluminescent probes, lucigenin (N,N'-Dimethyl-9,9'-biacridinium dinitrate; luminescence at 470 nm), CLA (2-Methyl-6-phenyl-3,7-dihydroimidazo[1,2-a]pyrazin-3(7H)one; luminescence at 380 nm), a chemical derived from the luciferin of Cypridina, and luminol (see 3.5.5) have been used for detection of O2• from plant material. Lucigenin. Lucigenin has been used with suspension cultures of rose [342]. Lucigenin itself is capable of producing considerable amounts of O2• via redox cycling especially when used in a concentration exceeding 10 µM; unfortunately high concentrations are required for the detection of O2• [343,344]. Furthermore, lucigenin can also be reduced by NAD(P)H reductases [345]. Luminol and L-012. Luminol, better known as H2O2 detector substance (see e.g. [346]), has also been used to measure O2• from potato tuber discs [347], Vicia faba roots [348] and from tobacco leaves [349]. Luminol [350], and its analog 8-amino-5-chloro-7-phenylpyrido[3,4-d]pyridazine1,4(2H,3H)dione (L-012) [351] might also generate O2• [346]. All of these probes, lucigenin, luminol and L-012, also react with H2O2 and peroxynitrite, L-012 being the most insensitive towards H2O2 but having remarkable reactivity with peroxynitrite [209]. CLA. CLA has been used to detect O2• from tobacco cell suspension [352] and from the epidermal layer of Vicia faba leaves [353]. In addition to O2•, CLA has been reported to react with 1O2, and to some extent also with HO• and H2O2 [354]. CLA has even been used as a 1O2 probe (for example [354]). Addition of SOD suppressed the CLA signal completely in plant leaves [353], except for an initial spike.
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MCLA. Several CLA-derived O2• probes are commercially available, including 2-methyl-6-(4methoxyphenyl)-3,7-dihydroimidazo[1,2-a]pyrazin-3(7H)-one (MCLA). It has been reported that MCLA does not react with H2O2 or O3 but reacts with HO• and 1O2 [355]. Godrant et al. [356] measured extracellular production of O2• by cyanobacteria Trichodesmium erythraeum and concluded that at PPFD 50‒100 µmol m-2 s-1 there was no interference from 1O2, nor did addition of 100 nM H2O2 affect the signal, though some signal from blank was observed as well as quite remarkable amount of a SOD non-inhibitable signal. MCLA has also been used to measure the (extracellular) production of O2• by immobilized marine diatoms [357].
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4.5.6. Microelectrodes With a microelectrode (0.02 cm2) consisting of Cu,Zn-SOD immobilized on nanostructured ZnO disks Deng et al. [358] measured in vivo O2• production from bean sprouts. Direct electron transfer between SOD and the electrode was achieved, with detection limit of 0.2 µM. No interference from H2O2, O2, uric acid, ascorbic acid or 3,4-dihydroxyphenylacetic acid was detected. O2• has been detected in vitro from tomato cell suspension with a Pt electrode in which SOD was immobilized within carboxymethylcellulose-gelatin [359,360].
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4.5.7. Genetically encoded probes cpYFP. Recently a new fluorescent O2• probe, Mito-cpYFP, was presented [361]. It is a circularly permuted yellow fluorescent protein that was targeted to mitochondria of human osteosarcoma cells using the cytochrome C oxidase subunit IV targeting sequence, and it was reported to react only with O2•, not with other ROS, nor with Ca2+, ATP, ADP, NAD(P)+ or NAD(P)H [361]. The probe was also used to detect mitochondrial O2• production in Arabidopsis thaliana [362]. However, Schwarzländer et al. [363] reported that in the mitochondria of Arabidopsis thaliana the probe only responded to changes in pH and not to changes in O2• level. Furthermore, purified cpYFP appeared not to respond to O2• [364]. 4.5.8. Indirect methods of O2• measurement Gene expression. The transcriptomic response of Arabidopsis thaliana [136] and the dinoflagellate Pyrocystis lunula [365] to O2• has been published, but this knowledge has not been used for O2• detection. 4.5.9. Summary of O2• detection As almost all the methods used to measure O2• have flaws, reliable results can be best obtained by using at least two parallel methods [276,313,366]. Furthermore, use of SOD or other agents to confirm specificity to O2• is recommended, although the complex chemistry of some probes (NBT, luminol and lucigenin) may make the evaluation more difficult (see [346,367]). In particular, SOD cannot very well quench O2• produced inside the membranes (e.g. [282]), and the probes themselves may also generate O2• that is quenched by SOD. Different localization of different chemicals should be taken into account when comparing results obtained with different probes [278].
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Table 5. Chemicals used for the detection of O2•. The stability data indicate the stability of the signal after reaction with O2• in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data. Method
Specificity
Stability (t1/2)
DMPO (5,5-dimethyl-1pyrroline N-oxide)
EPR
Also reacts with HO• and other radicals [273,285‒287]
45 s [286,287,289]
DEPMPO (5(diethoxyphosphoryl)-5methyl-1-pyrroline-Noxide) EMPO (5(ethoxycarbonyl)-5methyl-1-pyrroline Noxide) BMPO (5-tertbutoxycarbonyl-5-methyl1-pyrroline N-oxide)
EPR
Also reacts with HO• and other radicals [273,285‒287]
8 min [286,287,289]
EPR
Also racts with HO• and other radicals [273,285‒287]
14 min [286,287,289]
Slower growth of animal cells at 25 mM [297]
EPR
Also reacts with HO• and other radicals [273,285‒287]
23 min [292]
TMT (1-hydroxy-4isobutyramido-2,2,6,6tetramethylpiperidinium)
EPR
Also reacts with HO• and other radicals [273,285‒287]
DCP (probe 1-hydroxy2,2,5,5tetramethylpyrrolidine-
EPR
-
4 h, reduced by photosyntheti c electron transport [278] 4h
Hydrophobici Localization ty (LogP) 0.34 (estimated)
Slower growth of animal cells at 25 mM [297]; No effect on photosynthesis [96] Slower growth of 0.47 animal cells at 25 (estimated) mM [297]
US
MA N
TE D
AC
CE P
Toxicity
CR
Compound (IUPAC name)
Notes Spin adduct decays to HO• spin adduct and to HO• [294], UV-light sensitive [295]
-
Spin adduct may decay slowly to HO• spin adduct [269,287,289]
0.41 (estimated)
-
Spin adduct may decay slowly to HO• spin adduct [287,289]
Death of animal cells after long incubation, slower growth at 25 mM [297] No effect on reduction of P700+ [278]
0.86 (estimated)
-
-
1.5 [279] (experiment al)
-
-
No effect on reduction of P700+ [278]
-1.3 [279] (experiment al)
-
-
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Epinephrine
-
-
EPR
No reaction with HO•, ROO, H2O2, NO, GSH or L-ascorbate [288] Reduced by P680, and CO2- [83,308]
Stable [288]
T
EPR
Long incubation slows root growth [296] -
Absorbance at 550 nm
Absorbance at 490 nm
Sugars and phenolics reduce [311,312]
Reacts with ascorbate, GSH, plastoquinone, other components of photosynthetic ETC, oxidases and peroxidases [315‒317] Reacts with H2O2 [322]
US -
MA N
Absorbance at 550‒560 nm, histochemical staining Absorbance at 470 nm, histochemical staining
CR
IP
15 min [280]
Stable
TE D
XTT (Sodium,3*-(1[phenylamino-carbonyl]3,4- tetrazolium)-bis(4methoxy-6-nitro benzenesulfonic acid hydrate) Cytochrome c (Cyt c)
-
CE P
NBT (Nitro blue tetrazolium)
EPR
AC
3,4-dicarboxylic acid) Tiron (trap 4,5-dihydroxy1,3-benzene-disulfonic acid disodium salt) OXANOH (2-ethyl-1hydroxy-2,5,5-trimethyl3-oxazolidine) PTM-TC (perchlorotriphenylmethy l radical-tricarboxylic acid)
-0.15 (estimated)
-
0.97 (estimated)
Cell permeable
Have been used in vivo in roots, but detection in vitro [296] Interference from metal-ions [282]
Lipophilic, soluble to water at 1 mM [284] 3.49 (estimated)
Apoplast [284]
Has been used in vivo in roots [284]
Cell permeable
-
-
-
1.95 (estimated)
Cell permeable
-
Reaction product can be oxidized by cell components or peroxynitrite [317] Reaction product metabolized
-
0.30 (estimated)
-
Comparison with and without added SOD is needed, enhanced specificity by the use of acetylated Cyt c [263]
-
-1.37 [19]
-
Reaction product may produce O2• [324]
ACCEPTED MANUSCRIPT
Absorbance at 530 nm
-
HE or DHE (Hydroethidine)
Fluorescence (Ex 490, Em 520 nm)
Reacts with cytochromes, hemeproteins, Mnporphyrin complex, hypochlorous acid, NO and ONOO[332‒334] Reacts with H2O2 [209,345]
-1.23 (estimated)
Cell permeable [327]
4.03 (estimated)
Cellpermeable, Mito-HE localizes in mitochondri a
Can be reduced by NAD(P)H reductases [345] -
-
3.37 (estimated)
-
Produces O2• [343,344]
-
-0.41 (estimated)
-
May produce O2• [346]
-
-
1.10 (estimated)
-
M0ay produce O2• [346]
-
-
2.02 (estimated)
-
-
-
-
2.19
-
-
US MA N
-
TE D
CE P
Lucigenin (N,N′ -Dimethyl- Luminescenc 9,9′ -biacridinium e at 470 nm dinitrate)
Hydroxylamine inhibits oxygen evolving complex of PSII [327,328] Reaction product toxic to E. coli [332]
CR
Hydroxylamine
IP
by Cyt P450 reductase [323] -
T
44
Luminescenc e at 430 nm
L-012 (8-amino-5-chloro7-phenylpyrido[3,4d]pyridazine1,4(2H,3H)dione)
Luminescenc e at 460 nm
CLA (2-Methyl-6-phenyl3,7-dihydroimidazo[1,2a]pyrazin-3(7H)-one) MCLA (2-methyl-6-(4-
Luminescenc e at 380 nm
For reaction with H2O2, see 3.5.5; reacts with peroxynitrite [209] Reacts with H2O2 in the presence of peroxidase and with peroxynitrite [209,345] Reacts with 1O2, H2O2 and HO• [354]
Luminescenc
More specific than
AC
Luminol (5-amino-2,3dihydro-1,4phthalazinedione)
SOD may be needed to exclude unspecific reactions, produce O2• at the pH 8.5 or above [325] HPLC distinguishes specific and unspecific reaction products [292]; oxidized in UV and VIS light, enhances dismutation of O2• [332]
ACCEPTED MANUSCRIPT 45
CLA, reacts with 1O2 and HO• [355]
-
89.8% of sensitivity retained for 80 days [359]
-
-
-
-
-
-
-
Responds to pH, not to O2[364]
CR
-
TE D
-
-
US
Mitochondria targeted fluorescent (Ex 488, Em 505 nm) protein
-
MA N
Signal decreases less than 10 % in 6 days [358]
IP
T
(estimated)
Amperometry No interference by H2O2, O2, uric acid, ascorbic acid or 3,4dihydroxyphenylaceti c acid [358] Amperometry No response with 0.5 mM H2O2 [359]
CE P
CMC–G–SOD (Microelectrode with SOD on carboxymethylcellulosegelatin) Mito-cpYFP
e at 465 nm
AC
methoxyphenyl)-3,7dihydroimidazo[1,2a]pyrazin-3(7H)-one) Microelectrode with SOD on ZnO
ACCEPTED MANUSCRIPT 46
5. Hydroxyl radical, HO• 5.1. Definition and properties of HO•
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Hydroxyl radical (HO•), the result of homolytic cleavage of water (H2O --> HO• + H•), is the most reactive ROS. HO• is a very powerful oxidant; E0 of the pair HO•/H2O is +2.3 V [368]. The rate constants of reactions of HO• with many molecules range from 109 to 1010 M-1 s-1 [369,370]. HO• has one unpaired electron and is able to accept an electron from almost any molecule in its immediate vicinity [371].
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5.2. Formation and action of HO• in photosynthetic tissues
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Although HO• is known to be important especially in the chloroplast, HO• has received relatively little attention in plant biology. In biological systems the main HO• producing reaction is the metal-catalyzed Haber-Weiss reaction consisting of the Fenton reaction (reaction 15) and reduction of the transition metal ion by O2• (reaction 33) (Chapter 2; [1,372]). In spite of its high reactivity HO• plays a role in defense reactions in mammalian cells [373,374].
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Plants do not have specific scavengers for HO•, but the scavenging systems for H2O2 and O2• may protect against formation of HO• by limiting the Fenton reaction (reaction 15). It is assumed that plants prevent excess HO• production and subsequent damage by maintaining low levels of transition metals and sufficient levels of antioxidants in cellular compartments that might contain HO• [375]. The mechanisms maintaining appropriate levels of transition metals may also enable plants to target certain areas for controlled destruction [375]. For example, controlled destruction of cell wall pectins and xyloglucans for germination and elongative growth is believed to require HO• [376,377]. In the cell wall, HO• production has been proposed to involve soluble/cell wall bound proteins such as class III haemcontaining peroxidases (e.g. [372,378]), possibly in conjunction with cell wall associated SODs [379]. NADPH oxidases may also produce extracellular HO• [275,380]. Copper ions are readily available in cell walls and may catalyze Fenton chemistry, especially in roots [378,381]. Chloroplasts are likely the main intracellular sites of HO• production in plants due to the presence of transition metals (Fe, Cu, Mn) that are essential for the function of many transition metal binding proteins of the photosynthetic electron transfer chain [375,382]. Within chloroplasts the acceptor side of PSI is the principal site of H2O2 and O2• production [383,384], and therefore also HO• is thought to be mainly formed at PSI through the iron-catalyzed Haber-Weiss reaction (reactions 13‒15 and 33; [385]). However, with PSII being the main site for molecular oxygen evolution, also the role of PSII in HO• formation has been studied (reviewed by [2]). Formation of HO• at the acceptor side of PSII is thought to involve the reduction of metal center bound peroxides, such as non-heme iron bound peroxide. This reaction is similar to the Fenton reaction except that the reduction of the bound peroxide proceeds as an inner sphere electron transfer reaction [2,386]. Another possibility for HO• production at the acceptor side of PSII is the reduction of free peroxides in a typical Fenton reaction, with detached Mn2+ from the oxygen evolving complex of PSII serving as a possible reservoir of transition metal ions [386]. The oxygen evolving complex may also promote formation of HO• by acting as a source of H2O2 [387], especially after heat induced damage [388]. Yamashita et al. [388] proposed that HO• could be, along with 1O2, one
ACCEPTED MANUSCRIPT 47
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of the major ROS causing damage to PSII, particularly under stress. Also components of the mitochondrial electron transfer chain can produce HO•, but in plants the mitochondrial contribution is thought to consist mainly of production of H2O2 that can leak to other cell compartments to yield HO• in Fenton chemistry [375,389].
5.3. Reactions of HO•
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HO• participates in several typical reactions:
abstraction of hydrogen atom with formation of H2O and radical of substrate (36);
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HO• + RH H2O + R•
(36)
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addition to double bonds with formation of a hydroxylated radical (37);
(37)
TE
AC CE P
HO• + SCN + OH + SCN•
D
electron transfer reactions leading to formation of a neutral radical (38.1) or a cation radical [390] (38.2); (38.1)
(38.2)
formation of aromatic-OH adduct due to a reaction of an aromatic compound with HO• is one of the methods for HO• detection with HPLC-MS. For example, HO• can react with phenylalanine to form isomers of tyrosine, reaction (39) [391]. Isomers of tyrosine are rather stable and not normally present in proteins and can serve as HO• traps in biological samples [392].
(39)
ACCEPTED MANUSCRIPT 48 Interaction of HO• with many aquated metal cations is an electron transfer reaction (40) that proceeds via two different mechanisms, both with a rate constant of ~108 M-1 s-1 [369]. HO• + Mn+ M(n+1)+ + OH
(40)
(41)
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HO• initiates lipid peroxidation resulting in hydrogen abstraction from a pentyl group of an unsaturated fatty acid and formation of a radical that interacts with 3O2 to form a peroxyl radical (ROO•) with a rate constant of ~108 M-1 s-1 [393], reaction (41).
5.4. Lifetime and diffusion distance of HO• The rates of the reactions of HO• with biomolecules are usually limited by diffusion, and therefore reactions of HO• with biomolecules take place close to the site of HO• generation. The lifetime of HO• is thought to be in the order of nanoseconds [394,395], and recent simulations yielded a lifetime of only 30 ps in aqueous environment [371]. The diffusion distance of HO• has been estimated to be only a few molecular diameters from the place of origin [395]. The lifetime of HO• has not been measured in vivo.
5.5. Detection methods of HO• 5.5.1 Spectroscopic methods DMSO. Dimethyl sulfoxide (DMSO) has been used to detect HO• by colorimetry (absorbance at 420 nm) of methanesulfinic acid (MSA) formed in the oxidation of DMSO by HO• [396]. DMSO is only weakly toxic [397,398] and MSA is stable at pH 5‒9 [396]. Using DMSO, HO• formation has been witnessed in bean cowpea nodules, cucumber seedlings [399] and in leaves of duckweed and ryegrass [398]. In DMSO
ACCEPTED MANUSCRIPT 49 trapping, MSA must be extracted from the samples before colorimetry because a reaction between diazonium salts and MSA is required for the colorimetric assay [396‒400].
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Deoxyribose degradation. Free deoxyribose sugars yield malondialdehyde (MDA) upon degradation by HO• [401], and this observation has been utilized in HO• detection via detection of TBA-reactive substances (TBARS, see 6.4.5) [402]. Use of deoxyribose sugars as HO• detectors has been criticized for both its accuracy and its utilization in biological systems where free deoxyribose is not abundant [402]. In plant sciences this method has nevertheless been used to detect HO• formation in soybean leaves [403] and in wall fractions of isolated onion bulb cells [404].
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5.5.2. EPR detectable probes DMPO, DEPMPO, EMPO, 4-POBN. EPR spectroscopy is the most widely used method for detection of HO• in photosynthetic materials [2,269,375]. EPR probes that are used for HO• detection (DMPO, DEPMPO, EMPO) form adducts with O2• as well (see 4.5.2; [269]), but the EPR spectra of O2• and HO• probe adducts are so distinct that they allow differentiation between the two ROS, especially in the case of DMPO and EMPO. DEPMPO adduct spectra are more difficult to analyze [269,405]. Furthermore, HO• adducts produce diamagnetic species with Fe(III) and O2• and decay rapidly; both reactions result in loss of the EPR signal [152]. So even though HO• adducts can be distinguished, the actual amount of HO• being formed in a system may not be obtained with simplistic analyses of the EPR signal [269]. Taking into account the fact that decay of e.g. DMPO/O2• adducts yields HO• and that different probes act as scavengers of many ROS [269], special care needs to be taken in planning experiments where one and the same spin trap probe is used for the detection of O2• and HO•, as the naturally occurring formation of HO• is tightly linked to the concentrations of free O2•, free peroxides and transition metals [372,406]. Additional steps to confirm the precursors of HO• production in a studied system can be taken by adding SOD and CAT individually or together into the spin trapping mixture, thus pinpointing the prevalent HO• production pathway [407]. Also chelates such as ethylenediamine-N,N,N’,N’-tetraacetic acid (EDTA), diethylenetriamine-N,N,N¢,N’,N’’-pentaacetic acid (DTPA) and deferoxamine mesylate (Desferal) are commonly added into the buffers used in OH• measurements in order to reduce the amount of free transition metals, but use of these chelators should also be considered carefully as they themselves can stimulate ROS production [408,409]. 4-pyridyl-1-oxide-N-tert-butylnitrone (4-POBN) is an EPR probe that is used more spesifically for HO• detection. 4-POBN also reacts with O2• but the adduct is so unstable compared to a HO• adduct that almost no O2• adducts are detected [269]. In concentrations that are commonly used in biological studies (50 mM), 4-POBN can react with H2O2 in a reaction involving peroxidases, yielding 4-POBN•/4POBN spin adduct [410]. In addition to the specificity of 4-POBN, adducts produced in typical experiments with 4-POBN and HO• are considered to be stable when compared to other HO• reactive EPR probes [269,411]. The stability of 4-POBN adducts is, however, a result from the common practice of using HO• scavenging agents such as DMSO, ethanol or formate in conjunction with HO• probes [269,407,412‒415]. These scavengers react with HO• to form carbon centered radicals, such as αhydroxyethyl in the case of ethanol [413], which in turn form stable adducts with the spin trap. In the case of 4-POBN, the HO• adduct is stable because 4-POBN is practically always used with ethanol and the
ACCEPTED MANUSCRIPT 50
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detected product is an adduct of 4-POBN and a carbon-centered hydroxyethyl radical [416‒420]. It is important to take the additional effects of ethanol (from 170 to 850 mM) into account when planning spin trap experiments with 4-POBN [269]. The half-life of the actual 4-POBN/HO• adduct has been shown to be only 1 min in Hank’s Buffer Solution, much shorter than the half-life of the DMPO/HO• adduct (2 h) in almost the same conditions [421]. The life time estimates for different spin trap adducts derived from organic chemistry (e.g. [152]) can be difficult to translate into biological context, and therefore EPR measurements of spin adducts should be done as fast as possible.
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Although there are many issues concerning in vivo detection of HO•, mainly the very short lifetime of HO• and the yet uncharacterized production pathways of other interfering ROS [375,415], EPR spectroscopy has been used in vivo in plants. In vivo measurements of HO• directly from tissues infiltrated with 4-POBN have been done by Renew et al. [422] and Deng et al. [420] on intact cucumber root and tobacco leaf cutouts, respectively. Measurements done by exposing the intact tissue to the spin trap and then measuring EPR from ground tissue are more numerous. With 4-POBN HO• has been detected from roots [407,418], coleoptiles [416,417], plant radicles and endosperm caps [423] and from potato tubers [419]. DMPO, in turn, has been used to detect HO• from roots by Demidchik et al. [424].
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In vitro HO• has been detected with DMPO from isolated PSII membranes [285,425] and from isolated rapeseed hypocotyl tissue [414]. DEPMPO [387,426‒428], EMPO [388,429,430] and 4-POBN [386,387] have also been used in isolated PSII membranes. DMPO has been utilized in many photosynthesis studies to examine HO• formation in isolated PSII antenna complexes [431] and isolated thylakoid membranes [97,432]. Both DEPMPO [275] and 4-POBN [380] have been used to trap HO• in isolated plant plasma membranes, whereas DEPMPO has been used to measure HO• from pea cell wall isolates [433,434] and from the apoplast of cucumber leaf [435]. 5.5.3. Fluorescent probes Fluorophorenitroxide proxyl fluorescamine (PF). Alternatives to EPR include 5-((2-carboxy)phenyl)-5hydroxy-1-((2,2,5,5-tetramethyl-1-oxypyrrolidin-3-yl)methyl)-3-phenyl-2-pyrrolin-4-one sodium salt (fluorophorenitroxide proxyl fluorescamine, PF) [421,436]. PF works as a spin trap for HO• in a similar manner as 4-POBN. A reaction of PF with the methyl radical produced in a reaction between HO• and DMSO eliminates the fluorescence quenching nitroxide group of PF [421]. Other radicals such as O2• do react with PF, but with the use of DMSO the probe is described by [421] as HO• specific. The excitation/emission wavelengths of PF are 380/485 nm. HO• formation has been fluorometrically detected from aliquots of PF media used for experimental incubations of barley root tip segments [436]. The use of UV wavelengths for excitation limits the use of PF in biological material. Terephthalic acid. Liu et al. [437] detected the formation of HO• by benzene-1,4-dicarboxylic acid (terepthalic acid, TPA) in supernatant of pea pod cell wall homogenate. TPA traps HO• and forms monohydroxy terephthalate (TPA-OH) that fluoresces at 350‒550 nm with excitation at 362 nm [437]. While the signal producing reaction of TPA is HO• specific and the formed adduct is relatively stable [437,438], TPA is known to react with other radicals without producing a fluorescent adduct [439]. UV absorption by biological materials may limit the use of TPA as well.
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5.5.4. Microelectrode An electronic HO• microsensor, based on a change in the conductivity of polyaniline occurring due to a reaction with HO• was recently published [440], though it has not yet been used with photosynthetic material.
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5.5.5. Summary of HO• detection Because of its very short lifetime and unspecific reactivity, HO• is a difficult research subject and its function remains easily overshadowed by other ROS [441]. Furthermore, conclusions drawn from the physiological responses to H2O2 might be misleading due to the possible formation of HO• from H2O2 by the Fenton reaction [442]. Most methods used to measure HO• from plant material are not very specific (Table 6). Many methods relying on absorption/emission of HO• specific detector molecules are difficult to use in plants because the absorbance and fluorescence spectra of photosynthetic tissues often overlap strongly with the wavelengths of the signal.
ACCEPTED MANUSCRIPT 52
Specificity
Stability (t1/2)
Absorbance at 420 nm
HO• specific [396]
Stable [396]
Deoxyribose (2Deoxy-D-ribose) DMPO (5,5-dimethyl1-pyrroline N-oxide)
Absorbance at 530 nm EPR
Not very specific [402] Reacts also with O2• and other radicals [413]
-
Toxicity
RI
Method
MA
NU
SC
Compound (IUPAC name) DMSO
PT
Table 6. Chemicals used for the detection of HO•. The stability data indicate the stability of the signal after reaction with HO• in plant tissue. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data.
AC
CE DEPMPO (5EPR (diethoxyphosphoryl)5-methyl-1-pyrrolineN-oxide)
Reacts also with O2• and other radicals [286]
Stable [286]
EMPO (5(ethoxycarbonyl)-5methyl-1-pyrroline Noxide) 4-POBN (4-pyridyl-1oxide-N-tert-
EPR
Reacts also with O2• and other radicals [287]
-
EPR
HO• specific [269]
HO• adduct:
Not very toxic Slower growth of animal cells at 25 mM [297]; Plant root cells viable after treatment [441]; No effect on photosynthesis [96] Slower growth of animal cells at 25 mM [297]; No effect on photosynthesis [427]
PT ED
2 h [413]
Not very toxic [397]
αhydroxye
Hydrophobicity (LogP) -1.35 [19]
-1.90 (estimated) 0.34 (estimated)
Notes Optimal pH 5‒9; detected molecule is methanesulfic acid [396] Can be used to detect carbon centered radicals [413]
0.47 (estimated)
Can be used to detect carbon centered radicals [286]
Slower growth of animal cells at 25 mM [297]
0.41 (estimated)
-
-0.20 (estimated)
Can be used to detect carbon centered radicals [287] Commonly used with ETOH
ACCEPTED MANUSCRIPT 53
Fluorescence (Ex 380, Em 485 nm)
Reacts also with O2• [421]
Fluorescence (Ex 362, Em 350‒550 nm)
PANI electrode based on conductance change of polyaniline
Conductance measurement
May react with other radicals, but signal is HO• specific [439] Specific to HO• [440]
PT
[413] H 7 optimal for HO• adduct [411]
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thyl radical adduct: stable for >8 min [413]
-
-
Can be used to detect carbon centered radicals [436]
Stable [438]
-
2.00 [19]
-
Stable [440]
-
-
-
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Fluorophorenitroxide proxyl fluorescamine (5-((2carboxy)phenyl)-5hydroxy-1-((2,2,5,5tetramethyl-1oxypyrrolidin-3yl)methyl)-3-phenyl2-pyrrolin-4-one sodium salt) Terephthalic acid (Benzene-1,4dicarboxylic acid)
With O2• less than 1 min [413]; Without O2• 32 h [411] -
SC
butylnitrone)
ACCEPTED MANUSCRIPT 54
6. Peroxyl radical (ROO•), alkoxyl radical (RO•) and hydroperoxides 6.1. Definition and formation
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Peroxyl and alkoxyl radicals and hydroperoxides are best known as products of peroxidation of lipids. Other ROS, especially 1O2 and HO•, initiate reactions that lead to formation of these ROS.
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6.2. Reactions of ROO•, RO• and hydroperoxides
SC
The appearance of ROO• leads to formation of an alkoxyl radical (RO•) and lipid hydroperoxides (ROOH). The scheme of reactions leading to the appearance of these forms and, consequently, lipid peroxidation is well known, reactions (42‒44).
NU
HO• + RH R• + H2O
(42)
Initiation of the process and the appearance of lipid radicals;
MA
R• + 3O2 ROO•
(43)
addition of molecular oxygen to the carbon-centered radical (R•) yielding the ROO•;
D
ROO• + RH ROOH + R•
(44)
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abstraction of a hydrogen from another lipid molecule by ROO•, like reaction (42). Reaction (44) is known as the chain propagation reaction because this reaction supplies R• for reaction (43) and thus leads to accumulation of ROOH. The spontaneous decomposition of ROOH (reactions 45 and 46) is unfavorable because the dissociation energies of ROO-H and RO-OH bonds are 376 kJ/mol and 184 kJ/mol, respectively [443]. Spontaneous decomposition of ROOH can be observed in perturbed conditions, e.g. at a high temperature. ROOH ROO• + H•
(45)
ROOH RO• + HO•
(46)
Non-enzymatic decomposition ROOH via either their reduction or oxidation by transition metal is one of the most significant ways of ROOH decomposition (reactions 47.1 and 47.2). ROOH + Mn+ RO• + OH + M(n+1)+
(47.1)
ROOH + M(n+1)+ ROO• + H+ + Mn+
(47.2)
Interaction of ROOH with radicals can also lead to decomposition of ROOH due to abstraction of a hydrogen atom and formation of ROO• (reaction 48). ROOH + R• ROO• + RH
(48)
ROOH can also react with the C-H bond of saturated and alkylaromatic hydrocarbon via reaction (49) [444].
ACCEPTED MANUSCRIPT 55 ROOH + RH RO• + R• + H2O2
(49)
Dismutation of ROOH yields both RO• and ROO•, reaction (50). ROOH + ROOH RO• + ROO• + H2O
PT
(50)
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Decomposition of ROOH is a key reaction in the chain reaction of lipid peroxidation because of additional formation of ROO• or RO• species that can abstract a hydrogen from a lipid molecule, reactions (44) and (51). ROO• + RH ROOH + R•
(51)
NU
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The redox potentials (E0' at pH 7) of pairs RO•/ROOH and ROO•/ROOH are 1.6 V and from 0.77 V to 1.44 V, respectively [368]. The rate constant of the reaction of RO• with RH is 107 M-1 s-1, which is five orders higher than the rate constant of the reaction of ROO• with RH, 102 M-1 s-1 [168]. R•, ROO• and RO• can also interact with each other in several reactions [445,446]:
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The radical-radical reaction between two RO• can lead to formation of a carbonyl compound in excited state, reaction (52); RO• + RO• ROH + R=O*
(52)
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Reaction 52 is rather unlikely in biological systems because of the high reactivity of RO• toward lipids and proteins. A radical-radical reaction between two ROO• leads to the formation of an excited carbonyl compound (reaction 53.1) or 1O2 (reaction 53.2). The latter is called the Russell mechanism. The rate constant for the termination stage in the Russell reaction varies between 106 and 108 M-1 s-1 [445].
(53.1)
(53.2)
RO• and ROO• can react with R• to form dimers, reactions (54) and (55) [443,447]. RO• + R• ROR
(54)
ROO• + R• ROOR
(55)
Numerous experimental data on the kinetics of hydroperoxide chain process is represented in a recent monograph [444].
ACCEPTED MANUSCRIPT 56 Reactions (52) and (53.1) are associated with chemiluminescence (380–460 nm) due to relaxation of the excited carbonyl compound. This chemiluminescence is used to measure lipid peroxidation [448] (see 6.4.2).
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In reactions with lipid molecules, RO• can cause a homolytical scission (β-cleavage) of a C-C bond on either side of the alkoxyl group to form a wide range of aldehydes and oxo fatty acids [449]. Formaldehyde, acetaldehyde, propionaldehyde, acrolein, malondialdehyde, (E)-2-hexenal, (Z)-3-hexenal, 4-hydroxy-(E)-2-hexenal and other carbonyls are found to be products of degradation of polyunsaturated fatty acids [450], and therefore aldehydes can be used as markers of lipid peroxidation. Measurement of coloured substances produced in a reaction between aldehydes and thiobarbituric acid (TBA) is the most commonly applied method [451]. A new method for a comprehensive analysis of oxylipin carbonyls (quantitation and structural estimation) in plants with reverse-phase HPLC has recently been established [450,452].
MA
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Reduction of ROOH by thiol groups of peroxiredoxins is an important scavenging mechanism. This reaction proceeds as a 2-step reaction like reaction with H2O2, reactions (55.1) and (55.2). The Km values and rate constants of the reaction between peroxiredoxins and ROOHs depend on both ROOH species and on the peroxiredoxin, and the Km values range from 40 nM to 1.7 mM and the rate constants from 104 M-1 s-1 to 107 M-1 s-1 [453].
D
ROOH + RSH RSOH + ROH
(55.2)
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RSOH + RSH RSSR + H2O
(55.1)
Tocopherols and carotenoids can act as antioxidants preventing lipid peroxidation [454,455].
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6.3. Lifetime and diffusion distance of ROOHs The lifetimes of ROO•or RO• in the cells have been estimated to be 7 s and 1 µs, respectively [395]; these lifetimes would allow 5 % of the original ROS to be present at 0.16 mm or 60 nm from the site of formation in a viscous cellular environment, respectively. ROOHs are generally less reactive than ROO•or RO•.
6.4. Detection methods of ROOHs 6.4.1. Spectroscopic methods Lipid hydroperoxides contain fatty acids with conjugated dienes, and lipid hydroperoxides can be detected spectroscopically by measuring the absorbance of the dienes at 230‒235 nm. A high sensitivity for conjugated dienes is achieved with double-derivative spectroscopy [456]. However, this method can only be used for pure extracted material. 6.4.2. Chemiluminescence and thermoluminescence Excited carbonyl compounds, produced due to degradation of ROOHs, emit light at 380–460 nm. The chemiluminescence method (see Table 7 for a list of chemical methods for ROOH detection) can be applied for detection of ROOHs under both spontaneous and catalytic degradation of ROOH. ROOH-derived chemiluminescence has been measured from plant leaves with a sensitive camera [446]. ROOH-derived chemiluminescence resulting from sodium hypochlorite-induced degradation of ROOH was proposed as a sensitive assay for ROOH [457]. In plant material, addition of acridine in reaction medium significantly improved the linear correlation between luminescence intensity and amount of ROOHs [458]. A high-
ACCEPTED MANUSCRIPT 57 temperature thermoluminescence band peaking at 115‒130 °C was shown to correlate with the amount of lipid peroxides in chlorophyll-containing material [459]. The method has been frequently used for the detection of lipid peroxides and assessment of oxidative stress especially in photosynthesis research; see e.g. [460,461].
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6.4.3. Dyes Xylenol orange. Xylenol orange, used as dye for detection of ROOH is know as the ferrous xylenol orange method (FOX method). The FOX method is based on the oxidation of ferrous ions Fe(II) to ferric ion Fe(III) by ROOH in acidic conditions. The reaction of ferric ion with xylenol orange results in formation of a complex with an absorption maximum at 560 nm [462‒464]. Unfortunately, the stoichiometry between the amount of the xylenol orange complex and the amount of peroxide depends on peroxide species, ranging from 1.8 to 5.4 moles of xylenol orange complex per 1 mole of peroxide [465]. Therefore, a separate calibration curve is needed for each type of peroxide. The FOX method can be applied for both aqueous and organic solvents and used for detection of both hydrophilic (FOX1 method) and lipophilic (FOX2 method) peroxides. The FOX1 and FOX2 methods are not specific to ROOH because ferrous ions can be oxidized by other oxidizing agents. To accurately quantify ROOHs, the measurements are done in the presence and absence of triphenylphosphine which selectively reduces ROOHs to corresponding alcohols. To prevent peroxidation during analysis, butylated hydroxytoluenes are usually used [466,467].
TE
D
Thiocyanate. Thyocyanate (SCN) has also been used for detection of ROOH with the oxidation of ferrous iron. The reaction of ferric ion with the thiocyanate ion leads to formation of complexes absorbing at 470‒505 nm [462]. In contrast to the FOX methods, the thiocyanate-iron method can be used for detection of ROOH in material containing carotenoids [468].
AC CE P
Iodide. ROOHs oxidize iodide to iodine and iodine react with iodide, yielding triiodide anion (I3-). I3- absorbs at 290 and 360 nm [198,469]. A modified iodometric method has recently been used for determination of ROOH in plants [470]. The iodometric method is sensitive to oxygen that must be removed from the reaction medium. Iodometry is not specific for ROOH because H2O2 can oxidize I- to I2 (see 3.5.3; [471]), and therefore H2O2 should be removed before measurement. The stoichiometry between amounts of peroxides and iodine is 1:1. This method is known as an International Dairy Federation method (IDF method, 74A:1991). 6.4.4. Fluorescent probes Diphenyl-1-pyrenylphosphine (DPPP). DPPP can be used as a fluorescent probe to monitor lipid peroxidation in cell membranes. DPPP reacts with organic hydroperoxides stoichiometrically to yield DPPP oxide (DPPPNO) which emits at 380 nm with excitation at 351 nm [472]. DPPP readily reacts with lipophilic peroxides like methyl linoleate hydroperoxide but not with tert-butyl hydroperoxide. The DPPP method can be used for measurement of lipid peroxidation within membranes. DPPP also rapidly reacts with H2O2, and therefore this method can only be used in vitro with H2O2 free samples. [3-(4-phenoxyphenylpyrenylphosphino) propyl]triphenylphosphonium iodide (MitoDPPP). The reactivity of MitoDPPP is similar as that of DPPP, and MitoDPPP can be oxidized by peroxides including H2O2 but not by free radicals like O2• or HO•. The oxidation of MitoDPPP with hydroperoxides yields MitoDPPPO which has 35 times higher fluorescence intensity than MitoDPPP (ex 350 nm, em 380 nm) [473]. The reaction rate of MitoDPPP
ACCEPTED MANUSCRIPT 58 with peroxides depends on the type of the peroxide. MitoDPPP is supposed to localize within lipid membranes and can be used for detection of ROOH in living cells.
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2-(4-diphenylphosphanylphenyl)-9-(1-hexylheptyl)anthra[2,1,9-def,6,5,10-d′ e′ f ′ ]diisoquinoline-1,3,8,10tetraone (Spy-HP). Spy-HP reacts rapidly with hydroperoxides such as m-chloroperbenzoic acid (MCPBA) and cumene hydroperoxide to form an oxidized derivative (Spy-HPO) that emits strong fluorescence at 535 nm when excitated at 524 nm. The formation of Spy-HPO shows strict 1:1 stoichiometry with the amount of peroxides and its affinity to lipophilic hydroperoxides is high, allowing measurements at nM concentration. Spy-HPO is stable against autoxidation and photobleaching and the reaction of Spy-HP with O2•, H2O2 and HO• is very slow compared to the reaction with lipophilic peroxides [474]. Spy-HP has been used to detect lipid peroxides in photosynthetic membranes [475].
D
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2-(4-diphenylphosphanyl-phenyl)-9-(3,6,9,12-tetraoxatridecyl)-anthra[2,1,9-def:6,5,10-d9e9f9]diisoquinoline1,3,8,10-tetraone (Liperfluo). Liperfluo is oxidized by peroxides to Liperfluo-Ox with (ex 524 nm, em 535 nm). Liperfluo reacts much more slowly with various other ROS including H2O2, O2•, HO• and NO• than with methyl linoleate hydroperoxide [476]. Liperfluo shows only little cytotoxicity in SH-SY5Y cells and can be used for detection of peroxides in cells. It was demonstrated that Liperfluo is more useful as a probe than Spy-LHP for live-cell fluorescence imaging [476]. The fluorescence yield of Liperfluo-Ox is linearly dependent on the concentration of methyl linoleate hydroperoxide.
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Mito-Bodipy-TOH is a fluorescent peroxyl radical probe absorbing at 537 nm and emitting at 572 nm. The probe consists of an α-tocopheryl moiety, a fluorogenic reporter and a mitochondria-targeting segment by which the probe targets to the inner mitochondrial membrane [477]. The probe reacts with lipid peroxides but its specificity has not been extensively studied. Mito-Bodipy-TOH has not been used with photosynthetic material. 6.4.5. Measurements of ROOH-derived species TBA reactive substances (TBARS). Measurements of TBARS are commonly used for lipid peroxidation assays. The method is based on the ability of ROOH-derived products like malondialdehyde to react with TBA to form TBA-adducts at an elevated temperature in acidic conditions. TBA-adducts can be easily detected by absorbance at 532 nm [478,479] or by fluorescence at 553 nm [480]. Other ROOH-derived aldehydes can also react with TBA to form complexes [481]. Decomposition of lipid peroxides to TBARS during analysis results in formation of radicals that can promote additional lipid peroxidation, which may lead to overestimation of ROOH content. Protective substances (like butylated hydroxytoluene) are usually used to inhibit the lipid peroxidation [482,483]. Traces of iron (II) can also increase TBARS by accelerating decomposition of ROOHs, and therefore chelators are added to the medium [483]. HPLC analysis of TBA-adducts allows measurement of the MDA-(TBA)2-adduct only [484]. Using gas chromatography-mass spectrometry (GC-MS) it has recently been shown that lipid peroxidation is accurately measured with the TBA method in most plants [485]. 2,4-dinitrophenylhydrazine (DNPH). This method is based on extraction of lipid-derived carbonyls and their derivatization with 2,4-dinitrophenylhydrazine and separation with HPLC and LC/MS [452]. The carbonyl species can be identified from the MS/MS spectrum. The structure of ROOH-derived carbonyls is determined from the position of the hydroperoxide group in the lipid molecule. A HPLC based comprehensive analysis of carbonyls using DNPH for derivatization can be used for detection of fatty acids involved in peroxidation [486].
ACCEPTED MANUSCRIPT 59
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Table 7. Methods of detection of ROOH species. All mentioned chemicals show a stable, non-light-sensitive signal. LogP values have been obtained from Chemspider [19]; estimated LogP values were determined using a LogP prediction tool ALOGPS 2.1 [20]. - = no data. Method
Specificity
Hydrophobicity (LogP)
Notes
Ferric ion xylenol orange complex (absorption maximum at 560 nm) [462]
1.85 (estimated), can be used for both aqueous and organic solvents [466]
Various stoichiometries between amount of xylenol orange complex and amount of peroxide [465]
Thiocyanate (IDF method)
Ferric ion thiocyanate complex (absorption at 470–505 nm) [462,468]
0.22 (estimated), can be used for both aqueous and organic solvents [462,468]
Stoichiometry between amount of complex and amount of peroxide is not strict [462]
Iodide
Formation I3Absorption at 290–360 nm [198,469] Oxidized form has high fluorescence (Ex 351, Em 380) [472] Oxidized form has high fluorescence (Ex 350, Em 380) [473]
Nonspecific. Triphenylphosphine shuld be used to evaluate unspecificity [463,466] Nonspecific. Triphenylphosphine should be used to evaluate unspecificity [462] -
1.04 [19]
Experiments can be carried out in anaerobic condition dou to sensitivity to oxygen [198,469] -
MitoDPPP ([3-(4phenoxyphenylpyre nylphosphino) propyl]triphenylpho sphonium iodide) Spy-HP (2-(4Oxidized form has high diphenylphosphanyl fluorescence (Ex 524, phenyl)-9-(1Em 535) [474] hexylheptyl)anthra[2 ,1,9-def,6,5,10-
US
MA N
TE D
CE P
Can react with H2O2 [472]
8.52 (estimated), lipophilic [472]
Can react with H2O2 [473]
Lipophilic [473]
-
Specific for lipohilic peroxides. Slowly react with hydrophilic peroxides and H2O2 [474]
Very lipohilic [474]
Useful only for organic solvents [474]
AC
DPPP (Diphenyl-1pyrenylphosphine)
CR
Compound (IUPAC name) Xylenol orange (FOX method)
ACCEPTED MANUSCRIPT
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-
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-
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-
US
CR
-
MA N
Specific for lipohilic peroxides. Slowly react with hydrophilic peroxides and H2O2 [476]
AC
d′ e′ f ′ ]diisoquinoline1,3,8,10-tetraone) Liperfluo 2-(4High fluorescence (Ex diphenylphosphanyl- 524, Em 535) [476] phenyl)-9-(3,6,9,12tetraoxatridecyl)anthra[2,1,9def:6,5,10d9e9f9]diisoquinolin e-1,3,8,10-tetraone) TBA Complex with MDA with maximum of absorption at 532 nm [478,479] Mito-Bodipy-TOH Fluorescence (Ex 537, Em 572) [477]
T
60
-0.39 (estimated)
TBA can react with other carbonyls in addition to MDA [481]
-
Targeted for mitochondrial inner membrane. Not yet tested with photosynthetic material.
ACCEPTED MANUSCRIPT 61
7. Ozone, O3
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Ozone is an oxygen molecule consisting of three oxygen atoms (O3). O3 is a strong oxidant (E0 of the couple O3 + 2H+ + 2e-/O2 + H2O is 2.076 V [154]). O3 may also take part in 1-electron reactions, forming the ozonide radical O3-. Thermodynamic calculations suggest a value 1.6 V for the E0 value of the couple O3/O3• [487], although a lower value of 1.0 V could also be suggested [487]. External O3 does not penetrate to living plant cells [488] and therefore detection of O3 will not be discussed. However, external O3 penetrates to the apoplastic space through stomata and causes the formation of other ROS in the apoplast and thereby initiates plant defense signaling [8].
(56)
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The principal reaction of O3 with organic compounds is the addition to double bonds and cleavage of unsaturated compounds (ozonolysis). O3 reacts with a double bond to form ozonide and via the Criegee mechanism, reaction (56) [489].
Alternatively to formation of secondary ozonides, the decomposition of primary ozonides in aqueous medium can yield carbonyls and H2O2. The ozonolysis of ethylene yields two molecules of formaldehyde and one molecule of H2O2, reaction (57). Formation of H2O2 in reactions resembling reaction (57) may contribute to the ozone-induced oxidative burst in plant apoplast [8].
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(57)
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The rate constant of the reaction of O3 with ethylene at 30 °C is 106 M-1 s-1 [490]. O3 can react with isoprene and others terpenes to give corresponding carbonyls [491]. It was also shown that ozonolysis of terpenes as -terpineol, limonene, and -pinene is associated with formation of HO•, reaction (58) [492].
(58)
The reaction of O3 with such amino acids as cysteine, tryptophan, methionine, tyrosine, histidine and other biologically important compounds like ascorbic acid and NADPH produces 1O2. The rate constants and yields of 1O2 formation for many relevant compounds are given by [493]. The yield of 1 O2 in the reaction of O3 with ascorbic acid was found to be pH dependent, indicating that 1O2 is produced in a reaction between O3 and the ascorbate anion, reaction (59) [494]. O3 is able to oxidize GSH to a sulfonic acid [495]. AsCH + O3 DHA + 1O2 + OH,
(59)
ACCEPTED MANUSCRIPT 63
where DHA is dehydroascorbate.
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In addition to the classic ozonide forming reactions, electron transfer to ozone can lead to the formation of an ozonide anion radical, reaction (60). The ozonide anion radical is protonated very fast (k 1010 M-1 s-1), reaction (61) and then decomposed to hydroxyl radical and 3O2 [487,496].
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IrCl63 + O3 IrCl62 + O3 O3 + H+ HO3
(60) (61)
SC
In the atmosphere, ozone promotes the appearance of nitrogen dioxide (NO2•) via reaction (62) with nitric oxide (NO•).
NU
O3 + NO• NO2• + O2
MA
8. Non-specific methods
(62)
8.1. Non-specific methods indicating the presence of ROS
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Most direct ROS detection methods aim at detection and measurement of a specific type of ROS. Specificity is highly important because a method that detects several different ROS, each with different detection efficiency, has little predictive value for ROS-induced damage or ROS signaling. Several non-specific methods are, however, available in the market and they are sometimes used in the absence of specific ones. Non-specific reagents may also have advantages like low cytotoxicity or ability to localize to a specific cell compartment.
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H2DCFDA.The dyes 2′ ,7′ -dichlorodihydrofluorescein diacetate (H2DCFDA) and its derivatives 5-(and6)-chloromethyl-2′ ,7′ -dichlorodihydrofluorescein diacetate (CM-H2DCFDA) diffuse through the cell membrane and cell wall, but within the cell esterases remove the acetate groups and the chloromethyl group reacts with thiols. Therefore the dye cannot diffuse out of the cell, and a fluorescent oxidation product (DCF) can be detected by excitation at 492‒495 nm and emission at 517‒527 nm. CM-H2DCFDA has been used with Arabidopsis thaliana leaves; peeling off the epidermis is required to get the dye into leaf cells, and the dye responds to H2O2 treatment of the cells [23]. The chemical has also been used with cyanobacteria [130]. CM-H2DCFDA has been used as H2O2 detector (e.g. [497]), but because oxidation of CM-H2DCFDA depends on a Fenton-type reaction or on an unspecific enzymatic reaction with cytochrome c [498], a difference in the intensity of the fluorescence signal between two samples may indicate difference in H2O2, iron, copper or cytochrome c concentration. Also, UV and to a smaller degree visible light causes oxidation of DCFH [499], and DCF can produce O2• in light [500]. CellROX reagents. Molecular Probes Inc. produces several unspecific ROS detection reagents: CellROX® Deep Red Reagent with excitation and emission at 640 and 665 nm, respectively, CellROX® Orange Reagent (545/565 nm), CellROX® Green Reagent (485/520 nm) and OxyBurst (R) Green H2DCFDA, SE (2',7'-dichlorodihydrofluorescein diacetate, succinoimidyl ester). According to the website of the company (https://tools.lifetechnologies.com/content/sfs/manuals/mp10422.pdf; cited Feb 24, 2015), the Green Reagent binds to DNA and localizes to the nucleus in animal cells, and
ACCEPTED MANUSCRIPT 64
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the OxyBurst Green H2DCFDA, SE penetrates to cells. These reagents have not been tested with photosynthetic material but might be useful, as the emission wavelengths of the Orange and Green reagents do not interfere with Chl a emission. The rate constants of these reactants with different ROS have not been published. The cell-impermeable OxyBurst Green H2HFF BSA reagent has been used for detection of apoplastic ROS production during the generation of systemic signals related to wounding [501].
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HPF and APF. 3'-(p-hydroxyphenyl) fluorescein (HPF) and 3'-(p-aminophenyl) fluorescein (APF) were originally developed for the detection of HO• and peroxynitrite ion (ONOO-) [227]. In both cases a dearylation reaction converts a non-fluorescent dye to a fluorescent one, with excitation at 490 nm and emission at 515 nm. The dyes are cell permeable and resistant to autooxidation [227] and react with most ROS, including 1O2 and ROO• [24,227]. Data about their reactivity against O2• varies [24,227]. The reaction between APF and pure H2O2 is slow but becomes significant in the presence of a peroxidase [24].
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TEMPO-9-AC. 4-((9-acridinecarbonyl)amino)-2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO-9-AC) is a free radical that is converted to a fluorescent form (with excitation at 366 nm and emission at 424 nm) in a reaction with another free radical like O2• [502]. Strong illumination of tobacco leaves after pinhole administration of TEMPO-9-AC did not cause detectable fluorescence [109]. In leaves, this probe remains mostly in the apoplast [109].
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Ultra-weak luminescence, emitted by triplet excited carbonyls, pigments excited by excited carbonyls and by 1O2 has been shown to be associated with oxidative stress in plants, and mechanisms of production of light have been elucidated [503]. Long exposure time is required for quantitative imaging of ultra-weak luminescence.
8.2. Non-specific methods indicating reactions of ROS with biomolecules Proteins, DNA, lipids and various small biomolecules are known to react with ROS in vitro, but conclusive evidence for a reaction occurring in vivo or in an isolated system like thylakoid membrane preparation is difficult to obtain. Oxidation of proteins by ROS typically produces carbonyl groups in the amino acid side chains, and these can be detected with an antibody against 2,4-dinitrophenyl hydrazine (DNP) [504]. DNP-antibodies “Oxyblot™” can be used for general oxidative stress analysis. Reactive carbonyl species (RCS), defined as α,β-unsaturated aldehydes and ketones, are products of further reactions of lipid peroxidation [505]. Lipid peroxidation is initiated by reactive oxygen species, and therefore an increase in RCS, detected by the TBARS test, indirectly indicates an action of ROS. Examples of RCS occurring in plant material include malondialdehyde, 4-hydroxy-(E)-2nonenal, formaldehyde and acrolein. Unfortunately, the TBARS test fails to detect some physiologically relevant RCS like 4-hydroxy-(E)-2-nonenal [22].
9. Concluding remarks ROS are hot topics in plant biology because they function both as agents of damage and mediators of cellular signals. Understanding of both functions needs knowledge about the properties and reactions of each ROS, and the present review attempts to give an account of the biologically most
ACCEPTED MANUSCRIPT 65
relevant features of ROS, including redox potentials, key reactions and their rate constants, and lifetime and diffusion distance in biological material.
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The second aim of this review was to summarize the most relevant detection methods that have been used, or that are potentially usable, for experimental material deriving from plants, algae or cyanobacteria. The main focus is in photosynthetic tissues. We try to provide a listing of the properties of the method, including the possibilities to locate the ROS in a cell, toxicity of the detector substance and stability of both the detector substance and the signal. Unfortunately, the list often remains incomplete, because some topics have not yet been studied. We hope that the introduction to the methods may prompt future research of the missing topics and future development of better detection methods.
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To minimize artifacts, the following principles should be always be followed when chemical probes are used for ROS measurements.
MA
1. Ensure the specificity of the probe.
2. Check the toxicity of the probe to the study material and especially to photosynthesis.
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3. Ensure that the wavelengths and intensities of light used in the experiments do not cause artefacts like generation of ROS. 4. Investigate the cellular localization if the probe is used in vivo.
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5. Do not draw quantitative conclusions if the probe is unspecific.
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10. Acknowledgements
This study was financially supported by Academy of Finland (grants 259075 and 271832) and by Nordic Energy Research (AquaFEED project).
11. References
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Figure legends
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Fig. 1. A plot of temporally and spectrally resolved luminescence obtained for Chl a in acetone. The peak at 1270 nm, decaying in 40 µs originates from 1O2. Reprinted from [72], Copyright (2003), with permission from Elsevier.
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Fig. 2. Effect of TEMP on PSII in intact cells of the diatom Phaeodactylum tricornutum. The decay of Chl a fluorescence yield after a single turnover flash was measured with the FL-200 fluorometer (Photon Systems Instruments, Brno, Czech Republic) from a P. tricornutum culture before (black dashed line) and after 3 min incubation with 10 mM TEMP that had been purified by vacuum distillation (solid red line).
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Fig. 3. Demonstration that 30 min illumination of tobacco leaves with low light (LL, PPFD 50 µmol m2 -1 s ) induces hardly detectable quenching of DanePy fluorescence, whereas increase in SOSG fluorescence is obvious after the same treatment. Reprinted from [109] with kind permission from Springer Science and Business Media.
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Fig. 4. Stoichiometric production of H2O2 in a reaction between 1O2 and plastoquinol. Reprinted from [60], Copyright (2015), with permission from Elsevier.
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Fig. 5. Formation of H2O2 and O2• in cucumber cotyledons in the light, visualized by staining with DAB and with NBT, respectively. Reprinted from [307], Copyright (2014), with permission from Elsevier.
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Fig. 6. EPR spectra of BMPO/•OOH and BMPO/•OH adducts, as indicated. The dotted lines indicate computer simulations of the respective spectra. Reprinted from [292], Copyright (2001), with permission from Elsevier.
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Highlights
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The review discusses the reactions of Reactive Oxygen Species (ROS) Detection methods of ROS have been critically reviewed Advantages and disadvantages of a large number of chemical detection methods are described The main focus of the review is in the detection or ROS from photosynthetic organisms and tissues