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Assessments of Thiyl Radicals in Biosystems: Difficulties and New Applications The high reactivity of thiyl radicals (RS•), which results in half-lives on the order of microseconds, hinders their analysis in biological systems. This Feature reviews the contemporary approaches to assessment of RS• using EPR spin trapping, mass spectrometric, immunological, and HPLC protocols. Detcho A. Stoyanovsky,†,‡ Akihiro Maeda,†,‡ James L. Atkins,§ and Valerian E. Kagan†,‡ †
Department of Environmental and Occupational Health and ‡Center for Free Radical and Antioxidant Health, University of Pittsburgh, Pittsburgh, Pennsylvania 15213, United States § Walter Reed Army Institute of Research, Silver Spring, Maryland 20910, United States
A graphical representation of HPLC and immuno-spin-trapping analysis of thiyl radicals. Created by Robert Gates.
S
ulfhydryls play a central role in cellular homeostasis. To date, >25,000 proteins have been characterized, whereas the total number of proteins in human cells is believed to be ∼85,000 before posttranslational modifications (The Human Protein Reference Database; http://www.hprd.org).1 Most proteins contain either single or multiple sulfhydryl (SH) functions, thus representing a remarkable diversity of individual cysteine residues (Cys) in unique microenvironments. Moreover, the total intracellular content of protein thiols is comparable to that of the most abundant intracellular low molecular mass thiol, glutathione (GSH).2 The facile SH-to-SS and SH-to-SNO interconversions underlie enzymatic reactions, protein folding and trafficking, signaling by reactive oxygen and nitrogen species, and detoxification pathways (Scheme 1). In biological systems, the reactions that lead to regioselective modification of protein thiols in substrates remain largely unknown. The general mechanisms of thiolate/disulfide and thiolate/S-nitrosothiol interchange reactions are best described as bimolecular nucleophilic substitution and additionelimination reactions.36 Strained cyclic disulfides are reduced at higher rates, whereby a Brønsted relationship is followed in the values of pKa of both the nucleophilic thiol and of the thiol being displaced. In addition to nucleophilic reactions, one-electron oxidation of protein SH functions to immobilized thiyl radicals has been observed in the active sites of a number of enzymes.7 In proteins, immobilized RS• are formed via either long-range oneelectron transfer or short-range hydrogen atom abstraction from the thiol.8 For example, ribonucleotide reductases (RNRases) class I and II operate via cofactor-mediated generation of RS•, r 2011 American Chemical Society
which directly react with their substrates with concomitant regeneration of the SH function, thus closing the catalytic cycle. Another class of biologically significant reactions of sulfhydryls is their oxidation to freely diffusing RS•. Sulfhydryls reduce C- and O-centered radicals at diffusion controlled rates at the expense of their own oxidation to RS•. The latter can abstract H atoms from the methylene group of the penta-1,4-diene motif of phospholipids, alcohols, and the carbon backbone of sugars, to react with other sulfhydryls and to add to double bonds. Indeed, RS• generated during the metabolism of some xenobiotics have been shown to cause toxicity via modification of phospholipids and proteins.9 In biological systems, the participation of sulfhydryls in dynamic equilibria has made the assessment of the ratio between their redox forms overwhelmingly difficult. The analysis of pathways involving RS• is particularly complex. Because of their high reactivities, the half-lives of RS• are often in the microsecond scale. This Feature is focused on EPR spin trapping, mass spectrometric, immunological, and HPLC protocols that have been optimized for the identification of S-centered radicals in low molecular mass compounds and proteins. These techniques allow analysis of the catalytic activities of some SH-containing enzymes, metabolism of redox-sensitive xenobiotics, and identification of S-nitrosoproteins (PSNOs).
’ EPR/HPLC SPIN TRAPPING ANALYSIS OF GLUTATHIONE THIYL RADICAL A large body of literature indicates that the toxicity of a number of xenobiotics is caused by induction of RSH/RS•/RSSR redox cycles.10 Though glutathione is the most abundant cellular thiol with concentrations in the millimolar range, its one-electron oxidation to glutathione thiyl radical (Scheme 2, 8) is viewed as a toxicologically significant event.11 The advent of the EPR spin-trapping technique has considerably advanced the understanding of the mechanisms of oxidative stress.12 This technique is based on the high rates of interaction of nitrones or nitroso compounds (spin traps) with highly Published: May 18, 2011 6432
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Scheme 1
Scheme 2
reactive radical species. The paramagnetic nitroxides (spin adducts) formed as a result of reactions between spin traps and radical species are relatively stable compounds whose EPR spectra represent “structural fingerprints” of the parent radical species. Research the 1980s demonstrated that 8 reacts with 5,5dimethyl-1-pyrroline N-oxide (DMPO) to form nitroxide 9, which exhibits a four-line EPR spectrum with hyperfine splitting constants (in G) of aN = 15.1 and aH = 16.2 (Scheme 2, reaction 8 f 9).13 This protocol was instrumental in the identification of the reaction sequence 7 f 8 f 12 f 13 during the one-electron oxidation of phenol to phenol phenoxyl radical (7) by myeloperoxidase (MPX) and H2O2 in the presence of GSH.14,15 However, direct EPR evidence for the formation of 9 in intact cells has not been provided thus far. In cells, the assessment of this process is complicated by the low stability of 9, the lack of apparent consumption of 6, and the low, enzymatically controlled steady-state concentrations of oxidized glutathione (GSSG), O2• and H2O2. In a cell-free system, EPR detection of 9 is feasible when steady flows of 8 are generated.15 For example, only traces of 9, as assessed by EPR, could be observed in a reaction system consisting of DMPO, GSH, MPX, and H2O2 (Figure 1A, spectrum 2). Addition of phenol to this reaction system led to a marked increase of the EPR spectrum of 9, suggesting the occurrence of reaction sequence 6 f 7 f 8 f 9. Enzymecatalyzed elimination of either GSH or H2O2 from the reaction system led to a rapid disappearance of the spectrum of 9 (Figure 1B), indicating that this nitroxide decomposes with a half-life of ∼2 min. The low stability of a series of cyclic nitroxides, including 9, has been suggested to reflect an oxidative elimination of the hydrogen atom in the C2 position of the pyrrolidine ring, with concomitant formation of 11.16,17 In this process, 9 has been
postulated to be an oxidant, whereby reduction of its nitroxide functions by 9-C2-H yields the nitrone 11 and hydroxylamine 10.18 Refs. 15 and 19 present HPLC, MS, and NMR evidence showing that 9, generated via either oxidation of GSH by 7 or via photolytic homolysis of S-nitrosoglutathione (GSNO), undergoes dehydrogenation to the stable, EPR-silent nitrone 11 without any detectable accumulation of 10. Although reaction 8 f 9 is reversible,19 the large excess of nitrone (∼0.1 M DMPO versus micromolar concentrations of 9) maintains the equilibrium shifted in favor of 9; thus, the overall process is directed toward accumulation of 11. Formation of 11 can be followed by HPLC with UV detection at 258 nm, while additional identification of this nitrone can be performed by mass spectrometric analysis (Figure 1C and D). The use of 11 as an external HPLC standard has allowed analysis of the redox cycling of a series of phenols in HL60 cells, which are rich in MPX. In cells exposed to DMPO, H2O2, and phenols, quantification of 11 established that up to 50% of the total GSH can undergo one electron oxidation to 8.15 In summary, though EPR detection of RS• using spin traps is feasible in model chemical/biochemical systems, the HPLC analysis of 11 is currently the only experimental method that allows detection and quantification of 8 in intact cells. As an HPLC standard, 11 is remarkably stable; it did not decompose to any significant extent for ∼10 years when stored at 20 °C.
’ FLUORESCENCE ANALYSIS OF GLUTATHIONE THIYL RADICAL For imaging of 8 in intact cells, Borisenko et al. have employed a bifunctional molecular probe containing nitroxide and acridine moieties, 4-((9-acridinecarbonyl)amino)-2,2,6,6-tetramethylpiperidin-1-oxyl (Ac-TEMPO; Scheme 3, 14).20 6433
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Figure 1. EPR, HPLC, and MS analysis of 9 and 11. Reactions were carried out at 20 °C in 0.1 M phosphate buffer (pH 7.4). DMPO, 6, GSH, catalase, MPX, and glutathione peroxidase (GPx) were used at concentrations of 100 mM, 0.1 mM, 1 mM, 500 units mL1, 1 unit mL1, and 5 units mL1, respectively. A) EPR spectra of a reaction solution consisting of MPX, H2O2, and DMPO (trace 1); plus GSH (trace 2); plus GSH and 6 (trace 3). B) Effects of GPx and catalase on the EPR spectrum of 9 formed in a reaction system consisting of GSH, DMPO, 6, MPX, and H2O2. C) HPLC profile of 11 formed under the reaction conditions described in B; column, C18 (4.6 250 mm, Beckman Coulter, Inc., Brea, CA); mobile phase, 50 mM phosphate buffer (pH 7.4) containing 5% methanol; flow rate, 1 mL min1; UV detection at 258 nm. Inset C and panel D, UV spectrum and mass spectral analysis of the compound eluted under the peak with retention time of 6 min. Additional experimental details and 1H NMR analysis of 11 are reported in ref. 15.
Scheme 3
Because of intramolecular quenching of its excited singlet state by the stable nitroxide radical, Ac-TEMPO exhibits an EPR spectrum but not a fluorescence spectrum.20 In contrast to alkyl radicals, which react with nitroxides to form stable N-methoxyN-methyl-methanamines, RS• convert Ac-TEMPO to a secondary amine via the intermediate formation of 15. In the course of reactions 14 f 17, the EPR signal of the nitroxide function disappears and the fluorescence signal of acridine appears. AcTEMPO has been successfully used for imaging of one-electron oxidation of GSH to 8 in intact cells.20 However, Ac-TEMPO can be used as an imaging probe for 8 in cell cultures grown in the absence of ascorbic acid. In primary cell cultures, the ascorbic acid readily reduces nitroxides to hydroxylamines and therefore can lead to false readings for 8. Furthermore, for positive identification of 8, HPLC and/or mass spectrometric experiments
are required for the identification of 17 as an end reaction product.
’ ELECTRON-TRANSFER REACTIONS IN HEMOGLOBIN (Hb-FEIII) AND MYOGLOBIN (Mb-FEIII) In contrast to 8, protein thiyl radicals have limited mobility, which facilitates a highly specific, targeted one-electron transfer reaction. Pyruvate formate lyase, benzyl succinate synthase, and the RNRase superfamily form active site thiyl radicals as part of their catalytic cycles.7,21 However, EPR has not directly identified thiyl radicals in these enzymes thus far. Recent advances in EPR spin trapping methodology have allowed pinpointing of electron-transfer reactions in proteins. These techniques have been optimized in model systems based 6434
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Scheme 4
on H2O2-dependent formation of tyrosyl phenoxyl and cysteinyl thiyl radicals in Hb-FeIII and Mb-FeIII. Hb-FeIII is essential for aerobic life as an oxygen carrier, yet can also be harmful outside of red blood cells, where it acts as a peroxidase. Hb-FeIII reacts with H2O2 to form ferryl heme and globin radicals in which the unpaired electron can be localized either on the ironporphyrin ring or on a tyrosine or cysteine residue ([Hb-FeIV=O]•).22 [Hb-FeIV=O]• can participate in a number of redox reactions and has a lifetime of ∼300 ms,23 whereas its one electron oxidation leads to the formation of an oxoferryl complex ([Hb-FeIV=O]) with a lifetime ranging from minutes to hours.24 Both [Hb-FeIV=O]• and [Hb-FeIV=O] are reactive species that can trigger oxidation with potentially toxic consequences.25 Recent studies based on a combination of immuno-spin-trapping and MS have elucidated the sequence of H2O2-triggered intramolecular electron transfer in Mb-FeIII and Hb- FeIII.24,26 Reactions of protein radicals with spin traps often yield spin adducts with broad (anisotropic) EPR spectra that are difficult to interpret.27 Furthermore, these adducts undergo dehydrogenation to EPR-silent DMPOprotein nitrones following reaction 9 f 11.17,28 With the goal of improving analytical approaches in detecting the adducts, Detweiler et al. have elegantly proposed employing an immuno-spin-trapping protocol and developed a polyclonal antibody that specifically detects the nitrone moiety on proteins.28 The immuno-spin-trapping analysis of DMPO proteins is highly sensitive and, in combination with mass spectrometric methodologies, has allowed the analysis of the centers of radical generation in proteins. This method has been central to the analysis of the intramolecular transfer of unpaired electrons in Mb-FeIII and Hb-FeIII. Electronic, EPR, and mass spectrometric data, along with immuno-spin-trapping experiments, indicates that the oxidation of either Hb-FeIII or Mb-FeIII by H2O2 proceeds via sequential formation of an Fe(IV)-oxo porphyrin radical cation (Scheme 4, 19), tyrosyl radical (20), and then protein thiyl radical (21).26
Spin trapping of radical intermediates at varying DMPO concentrations has determined the sequence of these reactions with the notion that delocalization of the unpaired electron causes the reactivity of the tyrosyl radical 20 to be considerably lower than that of the thiyl radical 21. At high concentrations (g100 mM), DMPO quantitatively reacts with 20 to form Hb-Fe(IV)(=O)Tyr42/Tyr24-DMPO [or Mb- Fe(IV)(=O)-Tyr103-DMPO; 22]. At lower concentrations of DMPO ( 500 nm), only SNO functions undergo homolysis to cysteine thiyl radicals (Scheme 6, 31 f 32). Photolysis of PSNOs in the presence of DMPO leads to the formation of protein-cysteine-S-DMPO adducts that can be identified by Western blot (Figures 2 and 3)37 or by MS after proteolytic digestion. Photolysis/immuno-spin-trapping analysis has been shown to readily detect catabolic changes in the content of PSNOs in protein extracts from HepG2 cells incubated with thioredoxin type 1 (Trxn), thioredoxin reductase, and NADPH.37 The data presented in Figures 2 and 3 illustrate the immunospin-trapping analysis of PSNOs. Phosphate buffer containing DMPO and either purified S-nitroso albumin (Alb-SNO; Figure 2A, 6436
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Figure 2. Photolysis/immuno-spin-trapping analysis of Alb-SNO. Reactions were carried out at 20 °C in 0.1 M phosphate buffer (pH 7.4) containing DMPO (0.2 M). All samples were illuminated for 20 min with a 650-W lamp emitting white light, which was filtered through a 515 nm cutoff filter (luminance of 1 106 lux). Alb-SNO (A, C) and Alb-Sx (B) were used at concentrations of 0.1, 0.2, 0.5, 1.0, and 2.0 μM, respectively (lanes 15). C) Rat liver homogenate (1 mg of protein per mL) plus Alb-SNO were incubated at 20 °C for 10 min. Thereafter, aliquots from the samples were illuminated in the presence of DMPO and subjected to Western blot analysis as reported in refs. 17, 28 and 45. Briefly, proteins were resolved by SDS-PAGE (12.5%) and then transferred onto a nitrocellulose membrane. The membrane was consecutively treated with a blocking buffer (5% BSA/casein, 1:1), antiDMPO nitrone adduct polyclonal antiserum (rabbit IgG; Cayman; Ann Arbor, MI; dilution, 1:1000), and alkaline phosphatase-conjugated goat antirabbit IgG as a secondary antibody (Pierce; Rockford, IL; dilution, 1:5000). Thereafter, the membrane was exposed to Lumi-PhosTM WB Chemiluminescent substrate (Pierce) and visualized by chemiluminescence on an autoradiography film.37
lanes 15) or Alb-Sx (pre-irradiated, photolitically decomposed Alb-SNO; Figure 2B, lanes 15) was irradiated with visible light and aliquots from the reaction solution were analyzed for Alb-SDMPO by Western blot. The method afforded sensitivity for Alb-SNO in the submicromolar range. Decomposition of AlbSNO to Alb-Sx prevented the photolytic formation of Alb-SDMPO (Figure 2B, lanes 15), indicating that SNO-dependent generation of thiyl radicals was required for tagging of the protein with DMPO. Decreased content of Alb-SNO was observed in the presence of rat liver homogenate, indicating that S-nitrosoproteins are relatively unstable in biological milieu (Figure 2C, lanes 15). In contrast, incubation of rat liver homogenate with GSNO resulted in an extensive trans-S-nitrosation of hepatic proteins (HPSH; Figure 3, lane 3).45 No proteinDMPO adducts are formed from DMPO, HPSH, and GSNO when the samples are kept in the dark (Figure 3, lane 2). Labelling time of SNO functions in PSNOs with DMPO was reduced to minutes, which minimizes the possibility for redistribution of NO among cellular thiols. Furthermore, the mild conditions of the assay and the remarkably low toxicity of DMPO (ref. 17 and the references therein) are compatible with labeling of PSNOs in intact cells. After proteolysis of protein-cysteine-SDMPO adducts, MS analysis can identify the exact cysteine residues that have undergone nitrosation (tagged by DMPO). This approach may represent an important step in the analysis of PSNOs because many proteins contain multiple cysteine residues that can be subjected to S-nitrosation. Often, however, Snitrosation of only selected (critical) thiol(s) affects the activity of the corresponding protein. For example, caspase 3 contains eight cysteine residues (Human Protein Reference Database), whereby up to 4.5 SH functions per mole of protein can undergo S-nitrosation.29,46,47 Partial denitrosation of the protein by Trxn restores its proteolytic activity.29 On the other hand, all but one of the SNO functions in the p12 subunit of caspase 3 are
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Figure 3. Photolysis/immuno-spin-trapping analysis of trans-S-nitrosated HPSH.37 Extract of HPSH (molecular mass >7 kDa) was prepared via centrifugation of rat liver homogenate through a Sephadex G 25-filled spin column. HPSH (2 mg of protein per mL) was redissolved in phosphate buffer containing GSNO (0.3 mM) and EDTA (0.1 mM), and further incubated for 20 min at 20 °C. Aliquots from the samples (2 μg of protein) were subjected to Western blot analysis either prior or after irradiation in the presence of DMPO (0.2 M) as described in Figure 2. Lane 1, irradiated HPSH and DMPO; Lane 2, nonirradiated HPSH, GSNO and DMPO; and lane 3, irradiated HPSH, GSNO and DMPO.
denitrosated by GSH; denitrosation of the protein by GSH, however, did not result in its reactivation.47 Since the active site SNO function has not been observed in a mutant form of caspase 3 lacking the active site cysteine, Zach et al. proposed that in cells, NO nitrosates this cysteine to form S-nitrosocaspase 3, which is resistant to reduction by GSH.47
’ CONCLUDING REMARKS Currently, the spin trapping of S-centered radicals with DMPO is the paramount method for analysis of these radical species. DMPO has a well established mechanistic and derivative chemistry, and an array of optimized analytical methods exists for quantitation of DMPO thioethers. Furthermore, DMPO exhibits low cytotoxicity. In spin trapping experiments, cells tolerate this nitrone in concentrations up to 0.1 M, which makes it suitable for analysis of RS• in intact cells. The spin trapping of 8 has been attained with a series of acyclic nitrones and DMPO derivatives.19,48,49 Increased stability of nitroxides has been found with some DMPO derivatives (half-life up to ∼10 min). However, their breakdown products remain incompletely characterized. Hence, the protocols for assessment of DMPO thioethers reviewed herein represent a unique approach that allows reliable analysis of the biochemistry and pathology of radical-mediated protein oxidation50 and/or of S-nitrosation.37 ’ BIOGRAPHY Detcho A. Stoyanovsky received his M.S. in Organic and Analytical Chemistry at Sofia State University (Bulgaria) and 6437
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Analytical Chemistry Ph.D. in Biochemistry at the Bulgarian Academy of Sciences. His research interests are in redox biochemistry and medicinal chemistry, as well as the development of spin-trapping methods for analysis of radical metabolites. Akihiro Maeda received his Ph.D. in Molecular Biology at Tottori University (Japan). He is now a postdoctoral fellow in the group of Dr. Kagan at the University of Pittsburgh. James L. Atkins received his M.D. and Ph.D. from the University of Maryland. His research interests are renal acidification and resuscitation from trauma. He has recently retired from his position as Chief of the Blast Induced Neurotrauma Branch in the Center of Military Psychiatry and Neuroscience at Walter Reed Army Institute of Research. Valerian E. Kagan received his Ph.D. and D. Sci. in Biophysics/Biochemistry at Moscow State University/Academy of Sciences, Russia. He is the Director, Center for Free Radical and Antioxidant Health, Graduate School of Public Health at the University of Pittsburgh. His research interests are in redox biochemistry, cell biology, molecular toxicology, and lipidomics/oxidative lipidomics. Contact Stoyanovsky and Kagan at the Department of Occupational and Environmental Health, University of Pittsburgh Graduate School of Public Health, Pittsburgh, PA 15219 (stoyanovskyd@ upmc.edu;
[email protected]).
’ ACKNOWLEDGMENT This work is supported by NIH grant U19A1068021 and by a grant from the Office of Naval Research #42237. Disclaimer: The views, opinions, and/or findings contained herein are those of the authors and should not be construed as an official position, policy, or decision of the Department of the Army, the Department of the Navy, or the Department of Defense. ’ REFERENCES (1) Davison, D.; Burke, J. IBM J. Res. Dev. 2001, 45, 439. (2) Calcutt, G. Br. J. Cancer 1964, 18, 197. (3) Barnett, J. D.; Rios, A.; Williams, L. D. J. Chem. Soc. Perkin Trans. 2 1995, 1279. (4) Bachrach, S. M.; Woody, J. T.; Mulhearn, D. C. J. Org. Chem. 2002, 67, 8983. (5) Singh, R.; Whitesides, G. M. J. Am. Chem. Soc. 1990, 112, 6304–6309. (6) Bach, R. D.; Dmitrenko, O.; Thorpe, C. J. Org. Chem. 2008, 73, 12. (7) Stubbe, J.; van Der Donk, W. A. Chem. Rev. 1998, 98, 2661. (8) Frey, P. A. Annu. Rev. Biochem. 2001, 70, 121. (9) Schoneich, C.; Dillinger, U.; von Bruchhausen, F.; Asmus, K. D. Arch. Biochem. Biophys. 1992, 292, 456. (10) Munday, R. Free Radic. Biol. Med. 1989, 7, 659. (11) Schoneich, C. Chem. Res. Toxicol. 2008, 21, 1175. (12) Janzen, E. G.; Blackburn, B. J. J. Am. Chem. Soc. 1968, 90, 5909. (13) Josephy, P. D.; Rehorek, D.; Janzen, E. G. Tetrahedron Lett. 1984, 25, 1685. (14) Stoyanovsky, D. A.; Goldman, R.; Claycamp, H. G.; Kagan, V. E. Arch. Biochem. Biophys. 1995, 317, 315. (15) Stoyanovsky, D. A.; Goldman, R.; Jonnalagadda, S. S.; Day, B. W.; Claycamp, H. G.; Kagan, V. E. Arch. Biochem. Biophys. 1996, 330, 3. (16) Black, D.; Boscacci, A. B. Aust. J. Chem. 1976, 29, 2511. (17) Mason, R. P. Free Radic. Biol. Med. 2004, 36, 1214. (18) Janzen, E. G.; Haire, D. L. Adv. Free Radic. Chem. 1990, 1, 253. (19) Potapenko, D. I.; Bagryanskaya, E. G.; Tsentalovich, Y. P.; Reznikov, V. A.; Clanton, T. L.; Khramtsov, V. V. J. Phys. Chem. B. 2004, 108, 9315. (20) Borisenko, G. G.; Martin, I.; Zhao, Q.; Amoscato, A. A.; Kagan, V. E. J. Am. Chem. Soc. 2004, 126, 9221. (21) Licht, S.; Gerfen, G. J.; Stubbe, J. Science 1996, 271, 477.
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