Reactive species formed on proteins exposed to singlet oxygen

Abstract

Singlet oxygen (¹O₂) is believed to be generated in biological systems by a range of endogenous processes (e.g. enzymatic and chemical reactions) and exogenous stimuli (e.g. UV or visible light in the presence of a sensitiser). Kinetic data is consistent with proteins being a major target for ¹O₂, with damage occurring preferentially at Trp, His, Tyr, Met, and Cys side-chains. Reaction with each of these residues gives rise to further reactive species. In the case of Trp and Tyr, initial poorly characterised endoperoxides are believed to undergo ring-opening reactions to give hydroperoxides, which can be reduced to the corresponding alcohols; other products arising from radical intermediates can also be generated, particularly in the presence of UV light and metal ions. With His side-chains, poorly characterised peroxides are also formed. Reaction with Met and Cys has been proposed to occur via zwitterionic peroxy intermediates. Peroxides are also generated on isolated proteins, and protein within intact cells, via¹O₂-mediated reactions. The peroxides formed on Trp, Tyr, and His peptides, as well as on proteins, have been shown to induce damage to other targets, with molecular oxidation of thiol residues an important reaction. This can result in the inactivation of cellular enzymes and the oxidation of other biological targets. Protein cross-linking and aggregation can also be induced by reactive species formed on photo-oxidised proteins, though the nature of the species that participate in such reactions is poorly understood. These secondary reactions, and particularly those involving hydroperoxides, may play a key role in the induction of secondary damage (bystander effects) in systems subject to photo-oxidation.

---

Michael J. Davies

Michael Davies was awarded his BSc (1981) and DPhil (1984) degrees from the University of York, UK. After post-doctoral studies at Brunel University in London, he was appointed as a lecturer at the University of York in 1987. He moved to Australia in late 1995 to take up a position as a Group Leader at the Heart Research Institute, Sydney, on an Australian Research Council QE2 Fellowship. He was promoted to a Senior/Professorial Fellowship in 2000, and the Deputy Directorship of the Institute in 2001. In 2003, he was awarded the Silver Medal of the International EPR Society for his work in biology/medicine. His primary research interests lie in mechanisms of protein oxidation, the biological consequences of such reactions, and the development of methods to quantify protein damage in vivo.

---

Formation of singlet oxygen in biological systems

Singlet oxygen (molecular oxygen in its ¹Δ_g state; ¹O₂) has been postulated to be an important intermediate in a range of biological systems. Evidence has been presented for the formation of ¹O₂ by a range of peroxidase enzymes (e.g. myeloperoxidase,^1,2 lactoperoxidase,³ horseradish peroxidase,⁴ and chloroperoxidase⁵) and during lipoxygenase-catalysed reactions.⁶ A number of stimulated cell types (e.g. eosinophils⁷ and macrophages⁸) have been proposed to generate this oxidant, possibly as a result of the presence of the above enzymes, though other cells, such as neutrophils, do not appear to do so.^1,91O₂ is also formed in high yield in reactions involving ozone¹⁰ and during the reaction of H₂O₂ with HOCl,¹¹ sodium molybdate,¹² or peroxynitrite.¹³ A number of endoperoxide compounds have been synthesised which undergo thermal decomposition at room or elevated temperatures, thereby allowing the specific generation of defined yields of ¹O₂.¹⁴¹O₂ has also been shown to be formed during the bimolecular termination reactions of some peroxyl radicals;¹⁵ this species may therefore be an important intermediate in lipid (and other) peroxidation reactions.

Despite the wide range of non-photochemical reactions outlined above that generate ¹O₂, the most common source of ¹O₂ in biological systems is probably exposure to UV light, or visible light in the presence of an appropriate sensitiser. Cells contain a wide range of chromophores that can act as sensitising agents for ¹O₂ formation, and the quantum yields for the formation of this species by a wide range of agents have been collated.¹⁶ Whilst in some cases it is well established what the major ¹O₂-generating pathway is (e.g. within cells that have been pre-loaded with an exogenous sensitiser before light exposure), in many cases, there is considerable doubt as to what the principal sensitising agent is, with a large number of endogenous materials able to act in this capacity.

Direct generation of ¹O₂via the absorption of light by proteins that do not contain bound or associated co-factors is rather limited. Most proteins only absorb UV light relatively weakly in the region of the spectrum (λ > 290 nm) that is incident at the Earth's surface; shorter solar wavelengths are efficiently removed by molecules in the atmosphere. The major chromophores that absorb at wavelengths longer than 280 nm are tryptophan (Trp), tyrosine (Tyr), and cystine. Histidine (His), cysteine (Cys), and phenylalanine (Phe) only absorb UV significantly at shorter wavelengths (Phe ≤ ca. 270 nm, His ≤ ca. 240 nm, Cys ≤ ca. 250 nm). None of these side-chains has a high quantum yield for ¹O₂ generation.¹⁶ All other major amino acid side-chains and backbone peptide bonds do not absorb significantly at wavelengths above ca. 230 nm. A number of oxidation products of protein side-chains (particularly of Trp) are, however, reasonably efficient ¹O₂ sensitisers.¹⁶ Direct ¹O₂ formation by proteins is therefore inefficient and probably only contributes in a minor manner to the overall flux of ¹O₂ that might be generated within a biological system, despite the widespread abundance of proteins. The majority of ¹O₂ to which proteins may be exposed is therefore likely to arise via the photochemistry of other chromophores. As energy transfer to yield ¹O₂ usually competes with electron transfer, most photosensitisers generate both ¹O₂ and radicals.¹⁷ However, as energy transfer to O₂ is often more rapid (k ≈ 1–3 × 10⁹ dm³ mol⁻¹ s⁻¹) than electron transfer (e.g. to give O₂⁻˙, typically k ≤ 1 × 10⁷ dm³ mol⁻¹ s⁻¹), the former process often predominates, though there is obviously an enormous variation depending on the sensitiser, the excitation wavelength, and the reaction conditions.¹⁷

Proteins as a target for ¹O₂

Rate constants have been determined for the reaction of ¹O₂ with a wide range of biological targets, including DNA, RNA, proteins, lipids, and sterols.¹⁸ However, due to the abundance of proteins within most biological systems (they comprise ca. 70% of the dry mass of most cells¹⁹) and the high rate constants for reaction of ¹O₂ with some protein side-chains,^18,20 it would be expected that these will be the major targets for ¹O₂. This conclusion should, however, be treated with caution, as this will obviously be affected by the potential localisation and accumulation of sensitisers within particular cellular compartments (e.g. within membranes) and the limited diffusion radius of ¹O₂.²¹¹O₂ can interact with protein side-chains by both physical quenching and chemical reaction. The former results in energy transfer and de-excitation of the singlet state, but no chemical change in the energy acceptor. For proteins, only Trp appears to give rise to significant physical quenching,²² with k ≈ 2–7 × 10⁷ dm³ mol⁻¹ s⁻¹; chemical reaction occurs with k ≈ 3 × 10⁷ dm³ mol⁻¹ s⁻¹.²² The relative contributions of these processes with Trp are affected by a number of factors, including the environment of the residue and the presence and position of substituents on the indole ring.^23,24

The rate constants for chemical reaction of ¹O₂ with the side-chains of other free amino acids vary dramatically (Table 1). Of the common amino acids, only Trp, His, Tyr, Met and Cys, undergo rapid chemical reaction at physiological pH values. All other side-chains react with k < 0.7 × 10⁷ dm³ mol⁻¹ s⁻¹ at neutral pH.¹⁸ At higher pH values, Arg and Lys react rapidly via their unprotonated forms.²⁶

Table 1 Selected rate constants for reaction of ¹O₂ with protein side-chains at pH ca. 7^18,20,25

Side-chain	Rate constant/10^7 dm^3 mol^−1 s^−1
a Chemical reaction. b Physical quenching. c pH dependent with ca. 10 × 10^7 at pH > 8, and 0.5 × 10^7 dm^3 mol^−1 s^−1 at low pH.
Trp	3^a
	2–7^b
His	3.2–9^c
Tyr	0.8
Cys	0.9
Met	1.6

---

Formation and characterisation of peroxides on amino acid side-chains

In the case of Trp, Tyr, and His, there is good evidence for the formation of multiple endo- and hydroperoxides as a result of the initial addition of ¹O₂ to each of these residues. In the case of Trp and Tyr, the nature of these peroxides is reasonably well established. The intermediates formed with His, Met, and Cys are less well defined.

In the case of free Trp, reaction with ¹O₂ yields 3α-hydroperoxypyrroloindole (1) as an isolable product. The precursor species to this compound are not well defined. It has been proposed that the initial species may be a dioxetane across the C2–C3 double bond, though there is an absence of direct evidence for this, and there are other potential intermediates, including a hydroperoxide at C3 (Scheme 1).^27–32 The chemistry of peptide-bound Trp appears to be similar to that of free Trp, which is somewhat surprising given that the nitrogen lone pair involved in the ring-closing reaction will be much less available in peptides, due to the presence of the amide bond. It would therefore be expected that the formation of species analogous to 1 would occur at a much slower rate (cf. data for Tyr below). Under such conditions other reaction pathways may become important, and the precursor peroxides may be isolable or detectable; this has yet to be investigated in detail.

Scheme 1

Detailed studies have been carried out with free Tyr and peptide-bound Tyr. In the case of the free amino acid, evidence has been presented for the generation of a number of peroxidic species. These are outlined in Scheme 2. The proposed initial intermediate—the endoperoxide—has not been detected. Ring opening of this species is proposed to give a hydroperoxide at C1 (3) and direct evidence has been obtained for the presence of this compound.³³ Evidence has also been presented for the formation of related species with other phenols.^34,35 With free Tyr, rapid ring closure involving the unprotonated amine function then ensues, resulting in the generation of the cyclised product 3a-hydroperoxy-6-oxo-2,3,3a,6,7,7a-hexahydro-1H-indol-2-carboxylic acid (4) which has been extensively characterised.^33,36–39

Scheme 2

With Tyr residues in peptides, peroxide formation is also observed (Fig. 1), with the initial species believed to be an endoperoxide analogous to that outlined in Scheme 2. Subsequent ring opening of this species has been postulated to give rise to a hydroperoxide group at C1; this species has been fully characterised by NMR.³³ The yield of this material, when generated using Rose Bengal and light of λ > 345 nm, is enhanced when the reaction is carried out in D₂O rather than H₂O, and decreased when the reaction is carried out in the presence of the ¹O₂ scavenger azide; these data are consistent with the peroxides being generated via¹O₂-mediated reactions. The ring-closure reaction with the free amino group that occurs with the free amino acid is markedly slowed in peptides where the Tyr residue is not present at the N-terminus, due to the delocalisation of the nitrogen lone pair into the amide bond, making this atom a much less efficient nucleophile in the ring-closing Michael reaction. With compounds such as 4-hydroxyphenylpropionic acid (HPPA), where such reactions cannot occur, the C1 peroxide is also the major species detected. The major peroxidic product detected with peptide-bound Tyr residues at room temperature is therefore the C1 hydroperoxide (6) formed via the reactions in Scheme 3.³³

Fig. 1 (a) Formation of peroxides on 4-hydroxyphenylpropionic acid (□), N-acetyl Tyr (○), and Gly–Tyr–Gly (△) (all 2.5 mM) in D₂O, pH 7, as a result of illumination with light (from a 250 W tungsten-filament bulb passing through a 345 nm cut-off filter) in the presence of O₂ and the photosensitiser Rose Bengal (10 µM) at 4 °C with increasing illumination time. (b) Comparison of Gly–Tyr–Gly peroxide yields in D₂O, H₂O, and H₂O with added azide (5 mM) after 60 min illumination in the presence of Rose Bengal (10 µM) and O₂ at 4 °C. Peroxide concentrations were measured using the FOX assay with H₂O₂ standards.⁴⁰ Data are the means (±SD) of triplicate determinations from a single experiment representative of several, as the absolute peroxide yields varied between experiments due to slight differences in oxygenation rates.

Scheme 3

Oxidation of free His by ¹O₂ is believed to occur via the initial formation of one or more endoperoxides, such as 8, and the consumption of a single molecule of O₂ per mole of His (Scheme 4).⁴¹ Endoperoxides analogous to 8 have been detected directly by NMR with highly substituted derivatives in organic solvents at low temperatures.⁴² As the temperature was raised, these species were observed to decompose to yield further intermediates, which could not be characterised, and, subsequently, a range of stable products. Recent studies from the author's laboratory have shown that a number of peroxides are present in reaction mixtures where the illumination is carried out at 4 °C (Fig. 2).⁴³ These peroxides have lifetimes of minutes to hours at room temperature, which is inconsistent with them being endoperoxides. It is possible that they are hydroperoxides formed by ring opening of an initial endoperoxide, by analogy with the behaviour observed for Trp and Tyr. A number of these species are sufficiently stable to allow separation from the parent compound by HPLC, and the characterisation of these species is currently being attempted. NMR analyses of these reaction systems are consistent with this hypothesis.⁴⁴

Fig. 2 Formation of peroxides (in µM) on imidazole, histidine, and imidazole and histidine derivatives in H₂O, pH 7, as a result of illumination with light (from a 250 W tungsten-filament bulb passing through a 345 nm cut-off filter) in the presence of O₂ and the photosensitiser Rose Bengal (10 µM) at 4 °C with increasing illumination time. Peroxide concentrations were measured using the FOX assay with H₂O₂ standards.⁴⁰ Data are the means (±SD) of triplicate determinations from a single experiment representative of several, as the absolute peroxide yields varied between experiments due to slight differences in oxygenation rates.

Scheme 4

The formation of reactive species during the oxidation of free Met has been examined and evidence has been presented for the formation of a zwitterionic peroxy species (Scheme 5).⁴⁵ This species undergoes subsequent reaction with a second molecule of Met to give two moles of the sulfoxide⁴⁵ (reviewed in ref. 17 and 46). The stoichiometry of O₂ to Met consumption is complex and varies with pH, with this being 1 ∶ 1 above pH 7 and 1 ∶ 2 below pH 5, consistent with other reactions also playing a role. A number of other intermediates, including a nitrogen–sulfur cyclic intermediate, have been reported (reviewed in ref. 17 and 46); subsequent hydrolysis of this species gives the sulfoxide.⁴⁵ This cyclic intermediate is only likely to be of significance with free Met, due to the involvement of the free amino group. The situation with Met residues in peptides and proteins remain to be fully elucidated, though it has been shown that the sulfoxide is a major product.^47,48

Scheme 5

As with Met, the products arising from reaction of free Cys are well established—the disulfide is formed, though not in a quantitative manner—but the mechanism that gives rise to this product is less well established.^17,25,46 In addition to the disulfide, oxyacids such as cysteic acid (RSO₃H) are formed under some conditions.^17,46 The quantitative yield of the products arising from Cys residues in proteins is not well established and it would be expected that the ratio of disulfide to oxyacids will vary with the protein structure, due to steric or electronic barriers to dimer formation. This remains to be explored in detail. Qualitative studies have shown that oxidation of free cystine occurs via a zwitterionic peroxy intermediate, which reacts further with another molecule of cystine to give two molecules of RSS(=O)R.^46,49 The effect of incorporation of cystine residues into a protein on such reactions, where steric and electronic effects are likely to inhibit subsequent processes, remains to be determined.

Peroxide formation on isolated proteins

Peroxidic species have been shown to be formed on exposure of a wide range of proteins to ¹O₂, irrespective of their composition.^39,50 Examination of a range of proteins with different amino acid compositions has shown that peroxides can be generated on Trp, Tyr, or His residues within proteins. Thus, peroxides have been detected on melittin, which contains only Trp of the major reactive residues (i.e. no Tyr, His, Met, or Cys), on trypsin inhibitor, which contains only Tyr of the major reactive side-chains, as well as other proteins that lack one or more of these reactive residues.⁵¹ These peroxides are formed with a wide range of different sensitisers, though the overall yield varies markedly.^43,52,53 The peroxide concentration can be enhanced by use of D₂O as the solvent, consistent with the mediation of ¹O₂ (Fig. 3).⁵⁰ Protein peroxide formation has also been observed with other, non-photolytic, sources of ¹O₂ (e.g. with a molybdate–H₂O₂ system; Fig. 4).⁵⁰ The exact location of these peroxides within protein structures, and the nature of these species, have not been established as yet.

Fig. 3 Formation of peroxides on bovine serum albumin (75 µM) as a result of illumination with light (from a 250 W tungsten-filament bulb passing through a 345 nm cut-off filter) in the presence of O₂ and the photosensitiser Rose Bengal (10 µM) at 4 °C in H₂O (○) and D₂O (□). Peroxide concentrations were measured using the FOX assay with H₂O₂ standards.⁴⁰ Data are the means (±SD) of triplicate determinations from a single experiment representative of several, as the absolute peroxide yields varied between experiments due to slight differences in oxygenation rates.

Fig. 4 Formation of peroxides on bovine serum albumin (75 µM) treated with a MoO₄²⁻–H₂O₂ system. Reactions were performed at 22 °C with 12.5 mM MoO₄²⁻ in D₂O (open bars), H₂O (stippled bars), or H₂O in the presence of azide (5 mM) (black bars) at pH 7.4. The reactions were initiated by the addition of H₂O₂, with further aliquots of H₂O₂ added after 1 and 10 min; 1.62 mM H₂O₂ was added in total. The reactions were allowed to proceed for 30 min, then unreacted H₂O₂ was removed by gel filtration using a PD-10 column, and protein peroxide concentrations were measured using the FOX assay with H₂O₂ standards.⁴⁰ Data are expressed as means (n = 6, from 2 separate experiments) ± SD.

Peroxide formation on proteins within cells

Peroxides can be generated, in high yield, on bulk cellular proteins extracted from cells and subsequently exposed to visible light in the presence of a sensitiser (e.g. Rose Bengal⁵⁴). Recent studies have shown that peroxides are also generated on proteins within intact viable cells on exposure to visible light in the presence of a range of sensitisers.^43,54 The yield of these peroxides depends markedly on the sensitiser employed, even when identical initial sensitiser concentrations and illumination times are employed.⁴³ This is believed to be due not only to variation in the quantum efficiency of ¹O₂ generation, but also differences in sub-cellular localisation and concentration. The enhanced yield of these materials observed when the cells are cultured in media made up with D₂O, and decreased concentration in the presence of azide, is consistent with a role for ¹O₂ in the formation of these peroxides, though these data do not exclude a role for other photochemical processes in the formation of these species.⁵⁴ In the case of THP-1 cells (a human monocyte-derived cell line), a quantification has been attempted with regard to protein peroxide formation compared to total O₂ consumed. The data obtained are consistent with a reasonably efficient conversion of O₂ into protein-bound peroxides, with ca. 15% being detected as protein peroxides.⁵⁴ This figure is likely to be an underestimate of the total flux of O₂ going via this reaction pathway, as no correction was made for potential decay or further O₂-consuming reactions of the peroxides once formed.⁵⁴ The absolute levels of protein-bound peroxide formation in these cells have been calculated as representing an incidence of peroxide groups per protein molecule of ca. 0.5.⁵⁴ This figure is of a similar order of magnitude to the level of carbonyl groups detected on proteins in aged human cells or diseased brain tissue.^55–58 Whether peroxide formation on proteins occurs in intact biological systems (e.g. skin exposed to UV light) has not yet been definitively established. The data presented above suggest that this is likely.

Decomposition of side-chain peroxides

Compound 1, formed on oxidation of Trp by ¹O₂, spontaneously decomposes to the corresponding alcohol, 3α-hydroxypyrroloindole (2), at room or elevated temperatures.³⁹ The rate of decomposition is enhanced by the presence of reducing agents such as dimethylsulfide. In the absence of reducing agents, decomposition appears to be complex and may involve both heterolytic and homolytic cleavage of the peroxide bond.^27–32 Decomposition of 1 and 2 may also yield N-formylkynurenine³² and, hence, kynurenine via hydrolysis (Scheme 1). Metal ions and UV light have been shown to catalyse the decomposition of Trp-derived peroxides to radicals.^31,51 Two of the major overall products of oxidation, N-formylkynurenine and kynurenine, are formed with both the free amino acid and Trp residues in proteins,⁵⁹ suggesting that similar pathways occur in both cases. N-Formylkynurenine and kynurenine are better photosensitisers than Trp and, hence, can give rise to further ¹O₂ formation during continued light exposure.^16,60–62 Thus, the formation of N-formylkynurenine and kynurenine on proteins can result in an escalating cycle of damage. Extended UV exposure may also exacerbate damage via the sensitised decomposition of the initial peroxides. The poor sensitising activity, and high physical quenching activity (compared to chemical reaction), of Trp residues is believed to play a key role in the initial protection of the human lens against UV damage,⁶³ but after high levels of exposure, the gradual accumulation of more potent sensitisers, such as N-formylkynurenine and kynurenine and species formed by covalent addition of free Trp oxidation products to lens proteins,^64–66 may result in an enhancement of damage. The role of Trp and related materials as protective or potentiating agents in the human lens has been reviewed elsewhere.^63,67

The hydroperoxide species formed on Tyr residues are unstable and decompose above 4 °C.³³ Thermal decomposition at ambient or lower temperatures, and in the absence of added reagents, occurs in a relatively clean manner to give 5 in the case of the free amino acid (Scheme 2), and 7 with peptide-bound Tyr (Scheme 3), though decomposition at higher temperatures or in the presence of redox-active metal ions and UV light is less clean and a number of other products are formed.^33,39 Radicals have been detected during some of these peroxide decomposition processes;^33,39 the products of these radical reactions remain to be established, though previous studies have reported that 3,4-dihydroxyphenylalanine and di-tyrosine are not products.⁵⁹ It has been reported that 5 formed via the reactions shown in Scheme 2 can be oxidised further in basic conditions, giving rise to the decarboxylated keto compound.^36,38 The ultimate fate of the dienone alcohol (7) formed from the peptide-bound Tyr peroxide (Scheme 3) has yet to be established, though it has been shown that this species can undergo Michael reactions with nucleophiles such as thiols and amines.⁵⁰ These reactions may contribute to protein cross-linking reactions.

With histidine and related imidazole compounds, the initial endoperoxides detected by NMR at low temperatures undergo rapid decomposition to give a number of further poorly characterised peroxide intermediates and, ultimately, in the case of histidine derivatives, products including derivatives of aspartic acid and asparagine, and urea.^41,42,68 These reactions, which involve ring opening, consume further molecules of O₂.⁶⁹ Some of this O₂ consumption may arise via the decomposition of peroxides to give carbon-centred radicals that subsequently react to form peroxyl radicals.³⁹ The nature of these reactions are poorly understood and remain to be elucidated, though it is clear that multiple oxygenations and ring-opening reactions occur.

As might be expected on the basis of the above data, protein-bound peroxides also decay at room temperature or above to give non-peroxide products (Fig. 5).⁵⁰ These decay kinetics are likely to reflect composite curves for each of the individual species present and are, therefore, dependent on the protein under study. A wide range of factors are likely to affect the stability of these protein-bound materials, including the accessibility of these species to external reductants (see also below), the presence of metal ions and other reductants, continued exposure to UV, and the location of these materials on the protein structure (e.g. within hydrophobic pockets or on the surface of the protein). Little is known as yet about the factors that control the exact lifetime of such species.

Fig. 5 Stability of bovine serum albumin-derived peroxides at different temperatures. Peroxides were generated as outlined in Fig. 3. After the cessation of illumination, the samples were incubated in the absence of light for up to 24 h at 4 (□), 22 (○), and 37 °C (△), with the residual peroxide levels assayed at the indicated time points, as outlined in Fig. 3. Data are expressed as means of percentage initial peroxide concentration (n = 6, from 2 separate experiments) ± SD.

Data has been presented on the lifetime of protein peroxides formed within viable cells after generation using the sensitiser Rose Bengal and visible light. The species detected had half-lives of ca. 4 h at 37 °C when the cells were incubated in the absence of light, but in complete growth medium.⁵⁴ The kinetics of decay of these species was not affected by the presence of glucose or pre-loading of the cells with ascorbate.⁵⁴ It is likely that the lifetime of intracellular protein peroxides will depend significantly on the extent of depletion of cellular reducing equivalents (which may remove these species either directly or as a result of their utilisation as enzymatic co-factors), the species on which the peroxides are formed, their intracellular location (e.g. whether these are present on membrane proteins in a hydrophobic environment or on the surface of cytosolic proteins), and the activity of protective enzymes. The influence of some of these factors on the stability of protein peroxides is discussed further below.

Transfer of damage to other biological targets

Reaction with antioxidants

Peroxides generated on peptides and proteins by ¹O₂ undergo relatively ready reaction with a range of low molecular mass thiols, including the physiologically relevant tripeptide GSH.⁷⁰ Reaction does not appear to be stoichiometric, with large excesses of thiol (up to 10-fold) required to ensure rapid and complete removal of the peroxides.⁷⁰ The products of these reactions have not been characterised in detail, though it is likely that these will include the disulfide and oxyacids (RSOH, RSO₂H, and RSO₃H).⁷¹ Equal excesses (i.e. 10-fold) of a range of different thiols have been shown to give rise to different kinetics and extents of peroxide removal, suggesting that there are multiple factors that influence these reactions.⁷⁰ Other sulfur-containing species, such as the thioethers methionine and 3,3′-thiodipropionic acid, react much less rapidly.⁷⁰ In contrast, the organoseleno compound ebselen is very efficient, and ascorbic acid and some derivatives (though not ascorbic acid-2-phosphate) are moderately effective.⁷⁰ Phenolic compounds (e.g. Trolox C, BHT, and probucol) have, as expected, little effect.⁷⁰

Reaction with other proteins

As might be expected from the above discussion, protein thiols also react with peptide and protein peroxides. It has been shown that Trp-, Tyr- and His-derived peroxides present on small peptides can deplete thiol groups present on the enzyme glyceraldehyde-3-phosphate dehydrogenase (GAPDH), with this process paralleling the loss of the peroxides.^70,72 The rate of oxidation of thiol groups on proteins is structure dependent, with both the nature and size of the peroxide and the accessibility/location of the thiol group on the target protein playing a major role.^70,72,73 Thiols present on the same protein are oxidised at varying rates by different peroxides, and the same peroxide can react at different rates with a range of protein thiols. Smaller peroxides appear to induce thiol oxidation at a more rapid rate than larger peroxides (e.g. peptide-bound versus protein-bound species⁷³), though evidence has also been obtained for electronic interactions and more subtle structural effects also playing a role (cf. the more efficient inactivation of caspase enzymes by Trp-derived peroxides compared to Tyr- and His-derived species⁷³). As with the low molecular mass thiols, the stoichiometry of thiol oxidation to peroxides removed is variable, with values ranging between 2 and 1.⁷⁰ The products generated from such thiol oxidation have not been determined, but the variation in stoichiometry is consistent with the formation of both disulfides and oxyacids, with these being generated via molecular reactions rather than radical-mediated processes. The yield of disulfide versus other products is likely to be dramatically affected by the protein structure, with both electronic and steric effects determining the accessibility of the thiol to dimerisation reactions. These thiol oxidation reactions can result in enzyme inactivation, with loss of activity of a number of enzymes detected on incubation with peptide and protein peroxides (e.g. glyceraldehyde-3-phosphate dehydrogenase, glutathione reductase, caspases, and Ca²⁺-ATPases^72–74).

There is considerable evidence for the facile formation of high molecular weight aggregates (dimers and higher species) via cross-linking reactions.⁷⁵ Some of these aggregates may arise from radical–radical termination reactions of Tyr-derived phenoxyl radicals to give dityrosine,^76–78 though it has been reported that this is not a major process with most photo-oxidised proteins.⁵⁹ Intermolecular disulfide cross-linking has been discussed above, and appears to be a common process.^17,79 UVA light has been reported to generate disulfide cross-linked proteins in the lens and in isolated proteins; this may occur via¹O₂-mediated⁸⁰ or radical reactions,^81,82 or both processes concurrently.^82,83

Carbonyl groups formed as a result of side-chain oxidation⁵² can react with amine (Lys) and guanidine (Arg) functions on other proteins via Schiff base formation. Though such reactions are reversible, subsequent Amadori rearrangments results in irreversible cross-linking. It has also been proposed that His oxidation products can react with Lys, Cys, or other His residues to give cross-links.^{47,59,84–88} These reactions may account for the consumption of Lys and Arg residues in photo-oxidised proteins (reviewed in ref. 17), and may be responsible for the ‘dark’ reaction (post-cessation of illumination) inactivation observed with some enzymes (e.g. ref. 89–91). His residues may be of particular importance in aggregate formation, as proteins that lack His residues have been reported not to form cross-links.^59,85

In contrast to aggregation, there are few reports on protein fragmentation induced by photo-oxidation intermediates.^17,24 Photo-oxidation of lysozyme in the presence of lumiflavin has been reported to give peptide fragments, with this being proposed to occur either via oxidation of Trp residues and/or the formation of radicals.⁹² Radicals generated from peroxides formed on Trp, Tyr, and His residues³⁹ have been shown to carry out intermolecular hydrogen atom abstraction reactions from α-carbon sites on model peptides to yield backbone radicals;⁵⁰ these are likely progenitors of backbone fragmentation in the presence of O₂.^71,93,94 The significance of such reactions with proteins has yet to be determined.

Reaction with prosthetic groups

Evidence has been presented for the modification of prosthetic groups within proteins structures. Exposure of catalase to ¹O₂ has been shown to generate multiple conformers with more acidic isoelectric points as a result of the modification of the heme group.^95,96 This damage is distinct from that induced by UV light per se,⁹⁵ and has been proposed as a marker of ¹O₂ formation in vivo.⁹⁷ Similar reactions are likely with other prosthetic groups with which ¹O₂ reacts rapidly (for rate constants, see ref. 18).

Reaction with other targets

There is relatively little information as yet on the potential role of ¹O₂-mediated peptide or protein peroxides in the induction of damage to biological targets such as lipids and DNA, though preliminary evidence for this type of activity has started to appear.^98,99 This type of activity, if it involves peroxides, is likely to occur via the formation of radicals from the peroxide, catalysed by continued exposure to short wavelength UV light (which has been shown to decompose Tyr-derived peroxides³³) or metal ions.^33,39,72 Analogous experiments with peptide- and protein-derived peroxides generated by high energy radiation (γ radiation or X-rays, reviewed in ref. 100), have shown that these species can induce DNA base oxidation, strand breaks, DNA–protein cross-links,^101–104 and lipid oxidation,¹⁰⁵ so such damage might also be expected to occur with ¹O₂-mediated peroxides.

Consequences of ¹O₂-mediated damage to proteins

The formation of modified proteins as a result of ¹O₂-mediated photo-oxidation can result in wide-ranging biophysical and biochemical changes in both the properties and function of proteins. These include an increased or decreased susceptibility of the oxidised protein to proteolytic enzymes, alterations in mechanical properties (e.g. of silk and collagen), an increased extent of, or susceptibility to, unfolding, changes in conformation, an increase in hydrophobicity, altered light scattering properties and optical rotation, and changes in binding of co-factors and metal ions (e.g. ref. 17, 47, 59, 106, and 107). The role of reactive species formed on the oxidised proteins in these reactions is not fully understood, though it is clear that many of the products of such oxidation are poorly repaired, and the major fate of these species within cells is catabolism (reviewed in ref. 93 and 108–110). Though many oxidised proteins are known to be removed at an enhanced rate compared to normal proteins, populations exist which are poorly removed, resulting in intracellular accumulation of modified materials, particularly in the lysosomal compartments of cells.^111,112 The build-up of such materials may have major effects on cellular function,^93,108–110 and thereby contribute in a significant manner to the initiation and development of damage induced by sources of ¹O₂, for example, that seen in skin and the lens of the eye on prolonged exposure to UV light. Furthermore, the evidence that is starting to appear on the downstream effects of peptide and protein peroxide formation, such as the inhibition of key cellular enzymes and the potential induction of DNA damage, suggests that these species may be key intermediates in cellular dysfunction and disease progression.

Acknowledgements

Due to space limitations, a large number of papers which are pertinent to this topic could not be cited; the author apologies for these omissions. The author gratefully acknowledges the important contributions of his co-workers cited in the reference list. Work carried out in the author's laboratory is supported by the Australian Research Council and National Health and Medical Research Council.

References

1. J. R. Kanofsky, J. Wright, G. E. Miles-Richardson and A. I. Tauber, Biochemical requirements for singlet oxygen production by purified human myeloperoxidase, J. Clin. Invest., 1984, 74, 1489–1495.
2. J. R. Kanofsky, J. Wright and A. I. Tauber, Effect of ascorbic acid on the production of singlet oxygen by purified human myeloperoxidase, FEBS Lett., 1985, 187, 299–301.
3. J. R. Kanofsky, Singlet oxygen production by lactoperoxidase, J. Biol. Chem., 1983, 258, 5991–5993.
4. J. R. Kanofsky, Singlet oxygen production from the peroxidase-catalyzed oxidation of indole-3-acetic acid, J. Biol. Chem., 1988, 263, 14 171–14 175.
5. J. R. Kanofsky, Singlet oxygen production by chloroperoxidase-hydrogen peroxide-halide systems, J. Biol. Chem., 1984, 259, 5596–5600.
6. J. R. Kanofsky and B. Axelrod, Singlet oxygen production by soybean lipoxygenase isozymes, J. Biol. Chem., 1986, 261, 1099–1104.
7. J. R. Kanofsky, H. Hoogland, R. Wever and S. J. Weiss, Singlet oxygen production by human eosinophils, J. Biol. Chem., 1988, 20, 9692–9696.
8. M. J. Steinbeck, A. U. Khan and M. J. Karnovsky, Extracellular production of singlet oxygen by stimulated macrophages quantified using 9,10-diphenylanthracene and perylene in a polystyrene film, J. Biol. Chem., 1993, 268, 15 649–15 854.
9. J. R. Kanofsky, Bromine derivatives of amino acids as intermediates in the peroxidase-catalyzed formation of singlet oxygen, Arch. Biochem. Biophys., 1989, 274, 229–234.
10. J. R. Kanofsky and P. Sima, Singlet oxygen production from the reactions of ozone with biological molecules, J. Biol. Chem., 1991, 266, 9039–9042.
11. C. S. Foote and S. Wexler, Olefin oxidations with excited singlet molecular oxygen, J. Am. Chem. Soc., 1964, 86, 3879–3880.
12. J. M. Aubry, B. Cazin and F. Duprat, Chemical sources of singlet oxygen. 3. Peroxidation of water-soluble singlet oxygen carriers with the hydrogen peroxide-molybdate system, J. Org. Chem., 1989, 726–728.
13. P. Di Mascio, E. J. Bechara, M. H. Medeiros, K. Briviba and H. Sies, Singlet molecular oxygen production in the reaction of peroxynitrite with hydrogen peroxide, FEBS Lett., 1994, 355, 287–289.
14. M. Nakano, Y. Kambayashi, H. Tatsuzawa, T. Komiyama and K. Fujimori, Useful ¹O₂ (¹Δ_g) generator, 3-(4′-methyl-1′-naphthyl)-propionic acid, 1′,4′-endoperoxide (NEPO), for dioxygenation of squalence (a skin surface lipid) in an organic solvent and bacterial killing in aqueous medium, FEBS Lett., 1998, 432, 9–12.
15. M. Nakano, K. Takayama, Y. Shimizu, Y. Tsuji, H. Inaba and T. Migita, Spectroscopic evidence for the generation of singlet oxygen in self-reaction of sec-peroxy radicals, J. Am. Chem. Soc., 1976, 98, 1974–1975.
16. R. W. Redmond and J. N. Gamlin, A compilation of singlet oxygen yields from biologically relevant molecules, Photochem. Photobiol., 1999, 70, 391–475.
17. R. C. Straight and J. D. Spikes, in Singlet O₂, ed. A. A. Frimer, CRC Press, Boca Raton, 1985, vol. 4, p. 91.
18. F. Wilkinson, W. P. Helman and A. B. Ross, Rate constants for the decay and reactions of the lowest electronically excited state of molecular oxygen in solution. An expanded and revised compilation, J. Phys. Chem. Ref. Data, 1995, 24, 663–1021.
19. Geigy Scientific Tables: Physical Chemistry, Composition of Blood, Hematology, Somatometric Data, ed. C. Lentner, Ciba-Geigy Ltd, Basle, 1984, vol. 3.
20. B. Monroe, in Singlet O₂, ed. A. A. Frimer, CRC Press, Boca Raton, 1985, vol. 1, p. 177.
21. J. R. Kanofsky, Quenching of singlet oxygen by human red cell ghosts, Photochem. Photobiol., 1991, 53, 93–99.
22. I. B. C. Matheson, R. D. Etheridge, N. R. Kratowich and J. Lee, The quenching of singlet oxygen by amino acids and proteins, Photochem. Photobiol., 1975, 21, 165–171.
23. M. C. Palumbo, N. A. Garcia and G. A. Arguello, The interaction of singlet molecular oxygen O₂ (¹Δ_g) with indolic derivatives. Distinction between physical and reactive quenching., J. Photochem. Photobiol., B, 1990, 7, 33–42.
24. A. Michaeli and J. Feitelson, Reactivity of singlet oxygen toward amino acids and peptides, Photochem. Photobiol., 1994, 59, 284–289.
25. M. Rougee, R. V. Bensasson, E. J. Land and R. Pariente, Deactivation of singlet molecular oxygen by thiols and related compounds, possible protectors against skin photosensitivity, Photochem. Photobiol., 1988, 47, 485–489.
26. G. Papeschi, M. Monici and S. Pinzauti, pH effect on dye sensitized photooxidation of aminoacids and albumins, Med. Biol. Environ., 1982, 10, 245–250.
27. M. Nakagawa, K. Yoshikawa and T. Hino, The photosensitized oxygenation of Nb-methyltryptamine, J. Am. Chem. Soc., 1975, 97, 6496–6501.
28. M. Nakagawa, H. Okajima and T. Hino, Photosensitized oxygenation of Nb-methoxycarbonyltryptophan methyl ester and Nb-methoxycarbonyl tryptamine. Isolation and novel transformation of a 3a-hydroperoxypyrroloindole, J. Am. Chem. Soc., 1976, 98, 635–637.
29. M. Nakagawa, H. Watanabe, S. Kodato, H. Okajima, T. Hino, J. L. Flippen and B. Witkop, A valid model for the mechanism of oxidation of tryptophan to formylkynurenine - 25 years later, Proc. Natl. Acad. Sci. U. S. A., 1977, 74, 4730–4733.
30. M. Nakagawa, H. Okajima and T. Hino, Photosensitized oxygenation of Nb-methoxycarbonyltryptamines. A new pathway to kynurenine derivatives, J. Am. Chem. Soc., 1977, 99, 4424–4429.
31. I. Saito, T. Matsuura, M. Nakagawa and T. Hino, Peroxidic intermediates in photosensitized oxygenation of tryptophan derivatives, Acc. Chem. Res., 1977, 10, 346–352.
32. R. Langlois, H. Ali, N. Brasseur, J. R. Wagner and J. E. van Lier, Biological activities of phythalocyanines – IV. Type II sensitized photooxidation of L-tryptophan and cholesterol by sulfonated metallo phthalocyanines, Photochem. Photobiol., 1986, 44, 117–123.
33. A. Wright, W. A. Bubb, C. L. Hawkins and M. J. Davies, Singlet oxygen-mediated protein oxidation: evidence for the formation of reactive side-chain peroxides on tyrosine residues, Photochem. Photobiol., 2002, 76, 35–46.
34. I. Saito, S. Kato and T. Matsuura, Photoinduced reactions. XL. Addition of singlet oxygen to monocyclic aromatic rings, Tetrahedron Lett., 1970, 3, 239–242.
35. F. Jensen and C. S. Foote, Chemistry of singlet oxygen – 48. Isolation and structure of the primary product of photooxygenation of 3,5-di-t-butyl catechol, Photochem. Photobiol., 1987, 46, 325–330.
36. E. Katsuya, K. Seya and H. Hikino, Photo-oxidation of L-tyrosine, an efficient, 1,4-chirality transfer reaction, J. Chem. Soc., Chem. Commun., 1988, 934–935.
37. F. M. Jin, J. Leitich and C. von Sonntag, The photolysis (λ = 254 nm) of tyrosine in aqueous solutions in the absence and presence of oxygen – the reaction of tyrosine with singlet oxygen, J. Photochem. Photobiol., A, 1995, 92, 147–153.
38. S. Criado, A. T. Soltermann, J. M. Marioli and N. A. Garcia, Sensitized photooxidation of di- and tripeptides of tyrosine, Photochem. Photobiol., 1998, 68, 453–458.
39. A. Wright, C. L. Hawkins and M. J. Davies, Singlet oxygen-mediated protein oxidation: evidence for the formation of reactive peroxides, Redox Rep., 2000, 5, 159–161.
40. C. Gay, J. Collins and J. M. Gebicki, Hydroperoxide assay with the ferric-xylenol orange complex, Anal. Biochem., 1999, 273, 149–155.
41. M. Tomita, M. Irie and T. Ukita, Sensitized photooxidation of histidine and its derivatives. Products and mechanism of the reaction, Biochemistry, 1969, 8, 5149–5160.
42. P. Kang and C. S. Foote, Synthesis of a C-13, N-15 labeled imidazole and characterization of the 2,5-endoperoxide and its decomposition, Tetrahedron Lett., 2000, 41, 9623–9626.
43. V. Policarpio, C. L. Hawkins and M. J. Davies, unpublished data.
44. V. Policarpio, W. A. Bubb, C. L. Hawkins and M. J. Davies, unpublished data.
45. P. K. Sysak, C. S. Foote and T.-Y. Ching, Chemistry of singlet oxygen – XXV. Photooxygenation of methionine, Photochem. Photobiol., 1977, 26, 19–27.
46. W. Ando and T. Takata, in Singlet O₂, ed. A. A. Frimer, CRC Press, Boca Raton, 1985, vol. 3, p. 1.
47. J. Dillon, R. Chiesa, R. H. Wang and M. McDermott, Molecular changes during the photooxidation of alpha-crystallin in the presence of uroporphyrin, Photochem. Photobiol., 1993, 57, 526–530.
48. E. L. Finley, M. Busman, J. Dillon, R. K. Crouch and K. L. Schey, Identification of photooxidation sites in bovine alpha-crystallin, Photochem. Photobiol., 1997, 66, 635–641.
49. C. S. Foote and J. W. Peters, Chemistry of singlet oxygen. XIV. A reactive intermediate in sulfide photooxidation, J. Am. Chem. Soc., 1971, 93, 3975–3796.
50. A. Wright, C. L. Hawkins and M. J. Davies, unpublished data.
51. C. L. Hawkins and M. J. Davies, unpublished data.
52. J. A. Silvester, G. S. Timmins and M. J. Davies, Protein hydroperoxides and carbonyl groups generated by porphyrin-induced photo-oxidation of bovine serum albumin, Arch. Biochem. Biophys., 1998, 350, 249–258.
53. J. A. Silvester, G. S. Timmins and M. J. Davies, Photodynamically-generated bovine serum albumin radicals: evidence for damage transfer and oxidation at cysteine and tryptophan residues, Free Radical Biol. Med., 1998, 24, 754–766.
54. A. Wright, C. L. Hawkins and M. J. Davies, Photo-oxidation of cells generates long-lived intracellular protein peroxides, Free Radical Biol. Med., 2003, 34, 637–647.
55. C. N. Oliver, B. W. Ahn, E. J. Moerman, S. Goldstein and E. R. Stadtman, Age-related changes in oxidized proteins, J. Biol. Chem., 1987, 262, 5488–5491.
56. L. Lyras, P. J. Evans, P. J. Shaw, P. G. Ince and B. Halliwell, Oxidative damage and motor neurone disease. Difficulties in the measurement of protein carbonyls in human brain tissue, Free Radical Res., 1996, 24, 397–406.
57. M. J. Davies, S. Fu, H. Wang and R. T. Dean, Stable markers of oxidant damage to proteins and their application in the study of human disease, Free Radical Biol. Med., 1999, 27, 1151–1163.
58. S. Linton, M. J. Davies and R. T. Dean, Protein oxidation in ageing, Exp. Gerentol., 2001, 36, 1503–1518.
59. D. Balasubramanian, X. Du and J. S. J. Zigler, The reaction of singlet oxygen with proteins, with special reference to crystallins, Photochem. Photobiol., 1990, 52, 761–768.
60. P. Walrant and R. Santus, N-formyl-kynurenine, a tryptophan photooxidation product, as a photodynamic sensitizer, Photochem. Photobiol., 1974, 19, 411–417.
61. M.-P. Pileni, R. Santus and E. J. Land, On the photosensitizing properties of N-formylkynurenine and related compounds, Photochem. Photobiol., 1978, 28, 525–529.
62. M.-P. Pileni, M. Giraud and R. Santus, Kynurenic acid. II. Photosensitizing properties, Photochem. Photobiol., 1979, 30, 257–261.
63. M. J. Davies and R. J. W. Truscott, Photo-oxidation of proteins and its role in cataractogenesis, J. Photochem. Photobiol., B, 2001, 63, 114–125.
64. J. A. Aquilina and R. J. Truscott, Cysteine is the initial site of modification of α-crystallin by kynurenine, Biochem. Biophys. Res. Commun., 2000, 276, 216–223.
65. J. A. Aquilina, J. A. Carver and R. J. Truscott, Polypeptide modification and cross-linking by oxidized 3- hydroxykynurenine, Biochemistry, 2000, 39, 16 176–16 184.
66. J. A. Aquilina and R. J. W. Truscott, Identifying sites of attachment of UV filters to proteins in older human lenses, Biochim. Biophys. Acta, 2002, 1596, 6–15.
67. J. E. Roberts, in Sun Protection in Man, ed. P. U. Giacomoni, Elsevier, Amsterdam, 2001, p. 155.
68. S. Kai and M. Suzuki, Dye-sensitized photooxidation of 2,4-disubstituted imidazoles – the formation of isomeric imidazolinones, Heterocycles, 1996, 43, 1185–1188.
69. S. H. Chang, G. M. Teshima, T. Milby, B. Gillece-Castro and E. Canova-Davis, Metal-catalyzed photooxidation of histidine in human growth hormone, Anal. Biochem., 1997, 244, 221–227.
70. P. E. Morgan, R. T. Dean and M. J. Davies, unpublished data.
71. W. M. Garrison, Reaction mechanisms in the radiolysis of peptides, polypeptides, and proteins, Chem. Rev., 1987, 87, 381–398.
72. P. E. Morgan, R. T. Dean and M. J. Davies, Inhibition of glyceraldehyde-3-phosphate dehydrogenase by peptide and protein peroxides generated by singlet oxygen attack, Eur. J. Biochem., 2002, 269, 1916–1925.
73. M. B. Hampton, P. E. Morgan and M. J. Davies, Inactivation of cellular caspases by peptide-derived tryptophan and tyrosine peroxides, FEBS Lett., 2002, 527, 289–292.
74. C. Schoneich, V. Sharov and M. J. Davies, unpublished data.
75. J. D. Goosey, J. S. Zigler Jr. and J. H. Kinoshita, Cross-linking of lens crystallins in a photodynamic system: a process mediated by singlet oxygen, Science, 1980, 208, 1278–1280.
76. P. Guptasarma and D. Balasubramanian, Dityrosine formation in the proteins of the eye lens, Curr. Eye Res., 1992, 11, 1121–1125.
77. J. D. Spikes, H. R. Shen, P. Kopeckova and J. Kopecek, Photodynamic crosslinking of proteins. III. Kinetics of the FMN- and rose bengal-sensitized photooxidation and intermolecular crosslinking of model tyrosine-containing N-(2-hydroxypropyl)methacrylamide copolymers, Photochem. Photobiol., 1999, 70, 130–137.
78. H. Shen, J. D. Spikes, C. J. Smith and J. Kopecek, Photodynamic cross-linking of proteins V. Nature of the tyrosine-tyrosine bonds formed in the FMN-sensitized intermolecular cross-linking of N-acetyl-L-tyrosine, J. Photochem. Photobiol., A, 2000, 133, 115–122.
79. R. J. Truscott and R. C. Augusteyn, Oxidative changes in human lens proteins during senile nuclear cataract formation, Biochim. Biophys. Acta, 1977, 492, 43–52.
80. M. Linetsky and B. J. Ortwerth, Quantitation of the singlet oxygen produced by UVA irradiation of human lens proteins, Photochem. Photobiol., 1997, 65, 522–529.
81. M. Linetsky, H. L. James and B. J. Ortwerth, The generation of superoxide anion by the UVA irradiation of human lens proteins, Exp. Eye Res., 1996, 63, 67–74.
82. J. Dillon, B. J. Ortwerth, C. F. Chignell and K. J. Reszka, Electron paramagnetic resonance and spin trapping investigations of the photoreactivity of human lens proteins, Photochem. Photobiol., 1999, 69, 259–264.
83. F. J. Giblin, V. R. Leverenz, V. A. Padgaonkar, N. J. Unakar, L. Dang, L. R. Lin, M. F. Lou, V. N. Reddy, D. Borchman and J. P. Dillon, UVA light in vivo reaches the nucleus of the guinea pig lens and produces deleterious, oxidative effects, Exp. Eye Res., 2002, 75, 445–458.
84. H. Verweij and J. van Steveninck, Model studies on photodynamic crosslinking, Photochem. Photobiol., 1982, 35, 265–267.
85. P. Guptasarma, D. Balasubramanian, S. Matsugo and I. Saito, Hydroxyl radical mediated damage to proteins, with special reference to the crystallins, Biochemistry, 1992, 31, 4296–4303.
86. H. R. Shen, J. D. Spikes, P. Kopecekova and J. Kopecek, Photodynamic crosslinking of proteins. I. Model studies using histidine- and lysine-containing N-(2-hydroxypropyl)methacrylamide copolymers, J. Photochem. Photobiol., B, 1996, 34, 203–210.
87. H. R. Shen, J. D. Spikes, P. Kopeckova and J. Kopecek, Photodynamic crosslinking of proteins. II. Photocrosslinking of a model protein-ribonuclease A, J. Photochem. Photobiol., B, 1996, 35, 213–219.
88. H.-R. Shen, J. D. Spikes, C. J. Smith and J. Kopecek, Photodynamic cross-linking of proteins IV. Nature of the His-His bond(s) formed in the rose bengal-photosensitized cross-linking of N-benzoyl-L-histidine, J. Photochem. Photobiol., A, 2000, 130, 1–6.
89. T. M. Dubbelman, C. Haasnoot and J. van Steveninck, Temperature dependence of photodynamic red cell membrane damage, Biochim. Biophys. Acta, 1980, 601, 220–227.
90. A. Spector, G. M. Wang and R. R. Wang, The prevention of cataract caused by oxidative stress in cultured rat lenses. II. Early effects of photochemical stress and recovery, Exp. Eye Res., 1993, 57, 659–667.
91. A. Spector, G. M. Wang, R. R. Wang, W. C. Li and N. J. Kleiman, A brief photochemically induced oxidative insult causes irreversible lens damage and cataract. II. Mechanism of action, Exp. Eye Res., 1995, 60, 483–493.
92. T. Gomyo and M. Fujimaki, Studies on changes in protein by dye sensitized photooxidation. Part 3. On the photodecomposition products of lysozyme, Agric. Biol. Chem., 1970, 34, 302–309.
93. M. J. Davies and R. T. Dean, Radical-Mediated Protein Oxidation: From Chemistry to Medicine, Oxford University Press, Oxford, 1997, pp. 1–443.
94. C. L. Hawkins and M. J. Davies, Generation and propagation of radical reactions on proteins, Biochim. Biophys. Acta, 2001, 1504, 196–219.
95. F. Lledias and W. Hansberg, Oxidation of human catalase by singlet oxygen in myeloid leukemia cells, Photochem. Photobiol., 1999, 70, 887–892.
96. F. Lledias, P. Rangel and W. Hansberg, Oxidation of catalase by singlet oxygen, J. Biol. Chem., 1998, 273, 10 630–10 637.
97. F. Lledias and W. Hansberg, Catalase modification as a marker for singlet oxygen, Methods Enzymol., 2000, 319, 110–119.
98. G. D. Ouedraogo and R. W. Redmond, Photosensitized membrane oxidation leads to remote DNA damage by secondary reactive oxygen species, Free Radical Biol. Med., 2002, 33(suppl. 2), S421.
99. L. Booth and R. W. Redmond, Can lipid peroxidation of plasma membranes induce DNA strand breaks, Free Radical Biol. Med., 2002, 33(suppl. 2), S390.
100. J. M. Gebicki, Protein hydroperoxides as new reactive oxygen species, Redox Rep., 1997, 3, 99–110.
101. S. Gebicki and J. M. Gebicki, Crosslinking of DNA and proteins induced by protein hydroperoxides, Biochem. J., 1999, 338, 629–636.
102. C. Luxford, B. Morin, R. T. Dean and M. J. Davies, Histone H1- and other protein- and amino acid-hydroperoxides can give rise to free radicals which oxidize DNA, Biochem. J., 1999, 344, 125–134.
103. C. Luxford, R. T. Dean and M. J. Davies, Radicals derived from histone hydroperoxides damage nucleobases in RNA and DNA, Chem. Res. Toxicol., 2000, 13, 665–672.
104. C. Luxford, R. T. Dean and M. J. Davies, Induction of DNA damage by oxidised amino acids and proteins, Biogerentology, 2002, 3, 95–102.
105. M. J. Davies, unpublished data.
106. C. Prinsze, T. M. Dubbelman and J. Van Steveninck, Protein damage, induced by small amounts of photodynamically generated singlet oxygen or hydroxyl radicals, Biochim. Biophys. Acta, 1990, 1038, 152–157.
107. E. Silva, C. De Landea, A. M. Edwards and E. Lissi, Lysozyme photo-oxidation by singlet oxygen: properties of the partially inactivated enzyme, J. Photochem. Photobiol., B, 2000, 55, 196–200.
108. K. J. Davies, S. W. Lin and R. E. Pacifici, Protein damage and degradation by oxygen radicals. IV. Degradation of denatured protein, J. Biol. Chem., 1987, 262, 9914–9920.
109. R. T. Dean, S. Fu, R. Stocker and M. J. Davies, Biochemistry and pathology of radical-mediated protein oxidation, Biochem. J., 1997, 324, 1–18.
110. E. R. Stadtman and R. L. Levine, Protein oxidation, Ann. N. Y. Acad. Sci., 2000, 899, 191–208.
111. A. J. Grant, W. Jessup and R. T. Dean, Inefficient degradation of oxidized regions of protein molecules, Free Radical Res. Commun., 1993, 18, 259–267.
112. A. J. Grant, W. Jessup and R. T. Dean, Accelerated endocytosis and incomplete catabolism of radical-damaged protein, Biochim. Biophys. Acta, 1992, 1134, 203–209.

---