Molecular Plant Advance Access published online on November 14, 2008
Molecular Plant, doi:10.1093/mp/ssn072
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Methods for Analysis of Protein Glutathionylation and their Application to Photosynthetic Organisms
Institut de Biotechnologie des Plantes, UMR 8618, CNRS/Université Paris-Sud 11, Bâtiment 630, Orsay 91405, Cedex, France
1 To whom correspondence should be addressed. E-mail stephane.lemaire{at}u-psud.fr, fax + 33 1 69 15 34 24, tel. +33 1 69 15 33 38
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Abstract |
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 TOP Abstract INTRODUCTIONMETHODS FOR IDENTIFICATION AND...3-GLUTATHIONYLATION IN...FUNDING |
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Protein S-glutathionylation, the reversible formation of a mixed-disulfide between glutathione and protein thiols, is involved in protection of protein cysteines from irreversible oxidation, but also in protein redox regulation. Recent studies have implicated S-glutathionylation as a cellular response to oxidative/nitrosative stress, likely playing an important role in signaling. Considering the potential importance of glutathionylation, a number of methods have been developed for identifying proteins undergoing glutathionylation. These methods, ranging from analysis of purified proteins in vitro to large-scale proteomic analyses in vivo, allowed identification of nearly 200 targets in mammals. By contrast, the number of known glutathionylated proteins is more limited in photosynthetic organisms, although they are severely exposed to oxidative stress. The aim of this review is to detail the methods available for identification and analysis of glutathionylated proteins in vivo and in vitro. The advantages and drawbacks of each technique will be discussed as well as their application to photosynthetic organisms. Furthermore, an overview of known glutathionylated proteins in photosynthetic organisms is provided and the physiological importance of this post-translational modification is discussed.
Key Words: environmental signals • oxidative and photo-oxidative stress • proteomics • signal transduction • Glutathionylation • Methods • Glutaredoxin • Glutathione
Received for publication September 3, 2008. Accepted for publication October 1, 2008.
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INTRODUCTION |
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TOPAbstractINTRODUCTIONMETHODS FOR IDENTIFICATION AND...3-GLUTATHIONYLATION IN...FUNDING |
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Glutathione is a small tripeptide (
-L-glutamyl-L-cysteinyl-glycine) existing in the reduced form (GSH) or the oxidized form (GSSG) in which two glutathione molecules are linked via a disulfide bond. It is highly abundant (1–10 mM) in most cells, with the reduced form being predominant. For example, the concentration of glutathione in plant chloroplasts is estimated at between 1 and 4.5 mM and GSSG represents 
10% of this pool (Noctor and Foyer, 1998). Glutathione is generally considered to constitute a redox buffer that maintains the intracellular environment reduced. It protects many cell types against reactive oxygen species (ROS), xenobiotics and heavy metals damage. In plants, glutathione is also considered crucial in many other processes, including flowering, cell differentiation, programmed cell death, pathogen resistance, symbiosis, or cell cycle. The multiple functions of glutathione in photosynthetic organisms have been extensively reviewed recently (Foyer and Noctor, 2005; Meyer and Hell, 2005; Ogawa, 2005; Noctor, 2006; Pauly et al., 2006; Rouhier et al., 2008). Recent studies revealed that glutathione is also involved in a post-translational modification termed glutathionylation. S-thiolation consists of the formation of a reversible mixed disulfide between a small-molecular-weight thiol and a protein cysteine thiol (Gilbert, 1984; Ziegler, 1985). Since glutathione is, by far, the most abundant low-molecular-weight thiol in cells, glutathionylation represents the major form of S-thiolation. This modification apparently occurs mainly under oxidative stress conditions but may also be important under normal conditions, especially for regeneration of several thiol peroxidases. The exact mechanism leading to protein glutathionylation in vivo is still unclear. The reverse reaction, deglutathionylation, is likely catalyzed by the disulfide oxidoreductases glutaredoxins (GRXs) (Rouhier et al., 2008). Cysteine thiols can be oxidized to sulfinic or sulfonic acid in the presence of ROS. These oxidation states are generally irreversible, except in the case of 2-cys peroxiredoxins, in which the sulfinic acid form can be regenerated by sulfiredoxin and sestrin (Jacob et al., 2006; Jönsson and Lowther, 2007). Glutathionylation can protect cysteine thiols against irreversible oxidation to sulfinic or sulfonic acid but can also alter, either positively or negatively, the activity of many proteins. Therefore, glutathionylation could be an important redox signaling mechanism allowing cells to sense and signal harmful stress conditions and trigger appropriate responses. Glutathionylation has been shown to be involved in the regulation of several signal transduction pathways. In mammals, several proteins of the NF-kappaB pathway appear to be regulated by glutathionylation (Pineda-Molina et al., 2001; Reynaert et al., 2006b; Shelton et al., 2007; Kil et al., 2008). Glutathionylation was also reported to affect the activity of many protein kinases and phosphatases (Barrett et al., 1999a; Ward et al., 2000; Humphries et al., 2002; Rao and Clayton, 2002; Ward et al., 2002; Chu et al., 2003; Cross and Templeton, 2004; Humphries et al., 2005; Leonberg and Chai, 2007). Several transcription factors are also likely glutathionylated, such as the c-Jun subunit of the AP-1 complex (Klatt et al., 1999b, 1999c), the p50 subunit of NF-kappaB (Pineda-Molina et al., 2001), or Nuclear Factor 1 (Bandyopadhyay et al., 1998). The mechanisms, targets, and functional importance of glutathionylation have been described in several recent reviews and will not be extensively described in this article (Hurd et al., 2005; Ghezzi, 2005a, 2005b; Michelet et al., 2006; Dalle-Donne et al., 2007; Gallogly and Mieyal, 2007; Ghezzi and Di Simplicio, 2007; Townsend, 2007; Dalle-Donne et al., 2008; Rouhier et al., 2008).
Considering the potential importance of glutathionylation in stress responses and adaptation, a number of methods have been developed for identifying and analyzing proteins undergoing glutathionylation. These methods, which range from the analysis of purified proteins in vitro to large-scale proteomic studies, allowed identification of nearly 200 targets in mammals. The number of known glutathionylated proteins is much more limited in photosynthetic organisms, although they are severely exposed to oxidative stress conditions. Nevertheless, there is a growing interest for glutathionylation in these organisms and the number of known targets started to increase recently with the development of proteomic-based studies. In this review, we will detail the methods available for identification and analysis of glutathionylated proteins in vivo and in vitro. The advantages and limitations of each technique will be discussed, as well as their application to photosynthetic organisms. An overview of known glutathionylated proteins in photosynthetic organisms will also be provided and the possible physiological importance of this post-translational modification will be discussed.
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METHODS FOR IDENTIFICATION AND ANALYSIS OF GLUTATHIONYLATED PROTEINS |
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TOPAbstractINTRODUCTIONMETHODS FOR IDENTIFICATION AND...3-GLUTATHIONYLATION IN...FUNDING |
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Two major types of strategies have been developed to detect glutathionylated proteins. The most widely used techniques for proteomic analyses are based on the use of labeled glutathione, by either 35S radiolabeling or biotinylation. These techniques can be used either in vivo or in vitro and allow detection of glutathione adducts on S-thiolated proteins. The second type of approach is based on the detection of glutathionylated proteins without labeling of the glutathione pool using, for example, anti-glutathione antibodies or thiol alkylation after deglutathionylation by GRXs.
Methods Based on Labeling of Glutathione
35S Radiolabeling
Radiolabeled glutathione is a convenient tool for the analysis of protein glutathionylation. It allows a very sensitive and quantitative detection of glutathionylated proteins. The mixed disulfide between glutathione and protein cysteines can be readily reduced by chemical reductants like dithiothreitol (DTT). Therefore, a control treatment of radioactive samples with DTT allows confirmation that the labeling is indeed linked to glutathionylation. In vitro, the glutathionylation of several proteins has been analyzed using 3H-GSH (Klatt et al., 1999a, 1999c, 2000; Pineda-Molina et al., 2001; Cao et al., 2005) or 35S-GSH (Lind et al., 1998; Ward et al., 2000; Manevich et al., 2004; Hidalgo et al., 2006). The glutathionylation of mitochondrial complex II was demonstrated by incubation of isolated mitochondria with 35S-GSH followed by treatment with the thiol-specific oxidant diamide (Hurd et al., 2008).
Radiolabeling of the glutathione pool by 35S-cysteine has been the most widely used method for proteomic analysis of glutathionylated proteins in vivo (Figure 1). The first step consists of inhibition of protein synthesis with cycloheximide followed by incubation in the presence of 35S-cysteine. Glutathione is synthesized from glutamate, cysteine, and glycine in two ATP-dependent reactions catalyzed by -glutamylcysteine synthetase (-ECS) and glutathione synthetase (Noctor and Foyer, 1998; Meyer and Hell, 2005). Intracellular cysteine concentrations are maintained in the low micromolar range (up to 50 µM) to avoid generation of hydroxyl radicals by a Fenton-type reaction in the presence of hydrogen peroxide and transition metals (Meyer and Hell, 2005). In E. coli, high levels of cysteine have been shown to promote oxidative DNA damage through Fenton reactions (Park and Imlay, 2003). By contrast, the reactivity of the thiol group of glutathione with transition metals being much lower, cells can maintain millimolar concentrations of glutathione. Therefore, under conditions of protein synthesis inhibition, 35S-cysteine will be mainly incorporated into glutathione molecules. In photosynthetic organisms, in addition to cycloheximide, cells are also incubated in the presence of chloramphenicol in order to block chloroplast protein synthesis (Michelet et al., 2008). After the initial labeling step, the cells are placed under conditions leading to protein S-thiolation, namely oxidative stress conditions, generally triggered by addition of external oxidants such as H2O2, diamide, or NO donors such as GSNO or PABA/NO. Alternatively, S-thiolation can be achieved by increasing intracellular ROS production such as induction of the respiratory burst in monocytes, which leads to S-thiolation of several proteins (Rokutan et al., 1991; Ravichandran et al., 1994; Seres et al., 1996). After this second step, proteins can be extracted and separated on non-reducing mono- or bidimensional gels. S-thiolated proteins can be visualized after gel drying by autoradiography or phosphor imaging technologies. DTT treatments of radioactive samples should lead to a loss of the radioactive signal, thereby confirming that the labeling is indeed linked to S-thiolation.
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This method was originally developed to study S-thiolation in human monocytes (Rokutan et al., 1991) and allowed the identification of glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as a prominent S-thiolated protein (Ravichandran et al., 1994). A similar approach allowed the identification of glutathionylated GAPDH in human endothelial cells (Schuppe-Koistinen et al., 1994) and in yeast (Grant et al., 1999). This method also led to the identification of several other individual proteins undergoing glutathionylation in mammals including carbonic anhydrase III (Chai et al., 1994), actin (Rokutan et al., 1994), ubiquitin-conjugating enzyme (Jahngen-Hodge et al., 1997), protein kinase C-
(Ward et al., 2000), pax-8 (Cao et al., 2005), cyclophilin A (Ghezzi et al., 2006), and thioredoxin (Casagrande et al., 2002). With the development of proteomic approaches, this method was further adapted for large-scale identification of S-thiolated proteins using 2D gels and peptide mass fingerprinting. The first study was performed on human T lymphocytes where 38 S-thiolated proteins could be identified after treatment with diamide or H2O2 (Fratelli et al., 2002). Subsequently, similar studies allowed identification of a number of S-thiolated proteins in several cell types and organisms (Table 1). We recently employed this strategy to identify S-thiolated proteins in the unicellular green alga Chlamydomonas reinhardtii after diamide treatment (Michelet et al., 2008). Oxidative treatment with diamide led to the identification of 25 targets, mainly chloroplastic and involved in diverse metabolic processes (detailed below).
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The 35S-cysteine labeling method has, however, several major drawbacks. The major problem resides in the necessary pretreatment with protein synthesis inhibitors that will unavoidably perturb cell physiology. Moreover, this method does not allow discriminating between the different possible types of S-thiolation, although glutathionylation is considered to be predominant. Indeed, some of the radiolabeled proteins might not be glutathionylated but cysteinylated or S-thiolated by other low-molecular-weight compounds synthesized from cysteine, such as the 398-Da molecule recently identified in B. subtilis (Lee et al., 2007). The contribution of glutathionylation to the total labeling can be estimated in the presence of buthionine sulfoximine (BSO), a specific inhibitor of
-ECS. In human T cells, more than 80% of the radiolabeling is lost in the presence of BSO, thereby confirming that glutathionylation is the major type of S-thiolation (Fratelli et al., 2002). Nevertheless, the situation might be different in other cell types or other organisms. In B. subtilis, the six S-thiolated proteins identified by 35S-cysteine labeling were all found to be cysteinylated rather than glutathionylated (Hochgräfe et al., 2007). Another limitation is linked to the necessity to perform 2D gels to visualize the S-thiolated proteins. The loading limit of 2D gels will favor the identification of abundant proteins. Though this problem might be partly overcome by fractionation of the extract before 2D electrophoresis, low abundance proteins will probably not be identified with this method. Similarly, proteins with a high or low pI or a very high or very low molecular weight will not be detected. The sensitivity of this method is also limited by the low specific activity of the 35S-labeled glutathione pool. This limitation seems particularly problematic in photosynthetic cells where cysteine might be less efficiently imported from the medium than in human cells. This problem has impaired the identification of 35S-thiolated proteins in Arabidopsis cell cultures (Dixon et al., 2005b). In Chlamydomonas, the cells had to be incubated for several hours in the presence of 35S-cysteine in order to detect S-thiolated proteins (Michelet et al., 2008) while less than 30 min are required in non-photosynthetic cells. Another drawback is that the use of this method is restricted to cell cultures, thereby avoiding studies on whole plants under physiological conditions, and strongly limiting genetic analyses. Finally, this method can only detect proteins undergoing glutathionylation during the treatment while some proteins might be already glutathionylated under basal conditions. Despite its numerous limitations and drawbacks, the 35S-cysteine labeling method has allowed identification of most known S-thiolated proteins. The method can be useful for identification of most abundant S-thiolated proteins in cell cultures while other methods will have to be employed for the analysis of low abundance proteins.
Biotinylated Glutathione
Biotinylated glutathione can be easily synthesized in vitro either in the reduced (BioGSH) or oxidized form (BioGSSG). An analog of reduced glutathione, glutathione ethyl ester (GEE), can also be used to generate a membrane-permeable form of biotinylated glutathione (BioGEE). The synthesis of all these compounds is based on the use of a water-soluble biotinylation reagent, named sulfosuccinimidyl-6-(biotinamido)hexanoate (Sulfo-NHS-Biotin) (Figure 2A). The biotinylation reagent is used to couple biotin to the primary amino groups of glutathione under mild alkaline conditions using an amine-free buffer. After the reaction is completed, any remaining biotinylation reagent is quenched by the addition of an amine-containing buffer to a 10-fold molar excess of the starting sulfo-NHS-biotin concentration. Depending on the glutathione form, the biotinylation reagent is reacted with the reduced form (GSH or GEE) at a 1:1 molar ratio, while a 2:1 molar ratio is used with the oxidized form. However, when the biotinylated reagent is coupled to reduced glutathione, it can react with the sulfur atom of glutathione, decreasing the total amount of free-thiol biotinylated glutathione (Figure 2B). In the case of oxidized glutathione, the presence of two primary amino groups at the opposite ends of GSSG leads to the incorporation of two biotin moieties into the GSSG molecule (Figure 2C). The resulting biotinylated glutathione molecules can be used as an effective marker for oxidant-induced S-glutathionylation. The presence of the biotin moiety on glutathione allows a sensitive and specific detection of glutathionylated proteins by non-reducing Western blot probed with commercially available streptavidin–horseradish peroxidase or anti-biotin antibodies.
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Biotinylated glutathione has been used to analyze the glutathionylation of several purified proteins in vitro, including p53 (Velu et al., 2007), caspase-3 (Huang et al., 2008), the cysteinyl-rich oligopeptidase EP24.15 (Demasi et al., 2008), GAPDH (Holtgrefe et al., 2008), or carbonic anhydrase III and malate dehydrogenase (Niture et al., 2005). Biotin-labeled glutathione can also be used in vivo for proteomic identification of glutathionylated proteins. When BioGSSG is used, it mimics a defined component of oxidative stress, namely a shift in the glutathione redox couple to the oxidized disulfide state. By contrast, BioGSH and BioGEE do not induce oxidative stress and they are used in combination with oxidants, such as diamide or H2O2. Biotin-labeled proteins can be affinity purified on avidin-conjugated agarose beads. After extensive washing with detergent buffer, proteins bound to avidin via a mixed-disulfide bond with biotinylated glutathione can be eluted by incubation with reducing agents, such as reduced DTT or β-mercaptoethanol, and identified by mass spectrometry. Initially, BioGEE was used to identify annexin A2 and peroxiredoxin II as glutathionylated proteins in Hela cells after an oxidant stress caused by exogenous H2O2 or generated in response to TNF-
(Sullivan et al., 2000). In Arabidopsis cell cultures treated with H2O2, BioGEE allowed visualization of 22 glutathionylated proteins, two of which, cytosolic triose phosphate isomerase (TPI) and chloroplastic fructose-1,6-bisphosphate aldolase, were identified by Edman sequencing (Ito et al., 2003). Nine other proteins glutathionylated in vivo were found in Arabidopsis dark-grown cell cultures after tert-butyl hydroperoxide (tBOOH) treatment in the presence of BioGSSG (Dixon et al., 2005b). In vitro treatments of total extracts from these Arabidopsis cells with BioGSSG followed by streptavidin agarose affinity purification and 2D gels led to the identification of 72 proteins. However, only 22 proteins could be eluted by DTT from the column under denaturing washes, suggesting that some of the targets might not be glutathionylated directly but rather associated with glutathionylated proteins. This underlines that caution should be taken when defining the conditions for washing the column before DTT elution. BioGSSG also recently allowed identification of 11 glutathionylated proteins by LC–MS/MS in rat heart during post-ischemic reperfusion (Brennan et al., 2006). S-thiolation has also been investigated with biotinylated cysteine (Eaton et al., 2002a, 2002b; Sadidi et al., 2005). Moreover, different forms of biotinylated glutathione have been employed in focused studies to demonstrate the glutathionylation of several proteins in vivo, including GAPDH (Eaton et al., 2002c), IKK-β (Reynaert et al., 2006b), mitochondrial complex I (Hurd et al., 2008), CFTR chloride channel (Wang et al., 2005), Ras (Adachi et al., 2004a), and annexin A2 (Caplan et al., 2004). Biotin-based strategies for proteomic analysis of glutathionylated proteins have several advantages compared to 35S-cysteine labeling methods. First, protein synthesis does not have to be inhibited during the oxidative stress treatment. Second, this method only detects glutathionylated proteins rather than all S-thiolation targets. Third, the affinity purification is very specific and overcomes 2D-gel limitations. The targets can be analyzed on 2D gels only loaded with glutathionylated proteins rather than total extracts, allowing detection of significantly less abundant proteins. Alternatively, eluted proteins can be analyzed by highly sensitive and high-throughput proteomic methods such as nanoLC–MS/MS. Finally, the presence of the biotin tag on the proteins of interest allows their detection by multiple methods such as immunoblotting with or without prior immunoprecipitation using biotin antibodies or HRP–avidin (Reynaert et al., 2006a), batch or column-based affinity purifications, or cellular localization by fluorescence microscopy (Brennan et al., 2006).
Despite all these advantages, much fewer glutathionylated proteins have been identified with this method than by 35S-cysteine labeling. The major drawback of the methods based on biotinylated glutathione is the presence of the bulky biotin tag on the glutathione molecule that might perturb the function of proteins interacting with glutathione and especially those controlling glutathionylation. For example, we have observed that some proteins glutathionylated with biotinylated glutathione could not be deglutathionylated by GRXs (Zaffagnini and Lemaire, unpublished results). A common drawback of both labeling methods is that they do not give access to proteins glutathionylated under basal conditions.
Anti-Glutathione Antibodies
Glutathionylated proteins can also be detected with commercially available anti-glutathione antibodies. Methods based on such antibodies are promising, since they could overcome most problems encountered with 35S and biotin labeling methods. Indeed, with anti-glutathione antibodies, glutathionylated proteins could be analyzed under more physiological conditions, since no pretreatment is required. This could allow detection of glutathionylated proteins by Western blots with 1D or 2D gels, by immunoprecipitation, or even by immunocytolocalization. Almost all published studies have been performed with a mouse monoclonal antibody (Virogen, Watertown, USA), which has proved useful to analyze, in vivo, the glutathionylation of actin (Dalle-Donne et al., 2003; Wang et al., 2003; Fiaschi et al., 2006), tubulin (Landino et al., 2004; Fernandes et al., 2005), S-crystallin (Craghill et al., 2004), HSP70 (Hoppe et al., 2004; West et al., 2006), and Type 1 calcium release channels (Aracena et al., 2005; Aracena-Parks et al., 2006). These proteins can probably be detected in total extracts because they are very abundant. Indeed, this antibody exhibits a low sensitivity that greatly limits the number of glutathionylated proteins detected. This sensitivity issue can be partly overcome by working on purified proteins or fractions enriched with the protein of interest. The anti-glutathione antibody has been used to study several purified proteins such as IKappaB (Kil et al., 2008), aconitase (Han et al., 2005), carbonic anhydrase III (Rizzello et al., 2007), tyrosine hydroxylase (Sadidi et al., 2005), or branched chain aminotransferase (Conway et al., 2008). Immunoprecipitation with a specific antibody followed by Western blot with the anti-glutathione antibody was also used for interferon regulatory protein 3 (Prinarakis et al., 2008), mitochondrial complex II (Chen et al., 2007), IKKβ (Reynaert et al., 2006b), p53 (Velu et al., 2007), or protein tyrosine phosphatase 1B (Rinna et al., 2006). However, this preliminary enrichment is only possible when the target protein is known and it is therefore not applicable for proteomic identification of unknown glutathionylated proteins. In total extracts, the anti-glutathione antibody only detects few abundant proteins (Brennan et al., 2006). Every proteomic study based on the use of this antibody led to the identification of only four or five abundant proteins such as HSP70 or actin (Craghill et al., 2004; West et al., 2006; Newman et al., 2007) (Table 1). The number of glutathionylated proteins detected appears higher with strong inducers of glutathionylation such as PABA/NO (Findlay et al., 2006; Townsend et al., 2006). A major drawback of the anti-glutathione antibody concerns its specificity. Indeed, glutathione is a very flexible molecule that can potentially exhibit hundreds of conformations, either in solution or bound to proteins (Lampela et al., 2003). Hence, the affinity of the antibody for glutathionylated proteins is likely to vary greatly, depending on the conformation of the glutathione adduct and the environment of the thiolated cysteine. We have observed, for example, that several purified plant proteins for which glutathionylation was confirmed by MALDI–TOF mass spectrometry were not detected by the antibody on Western blots (Zaffagnini and Lemaire, unpublished results). Conversely, it has been reported that some proteins containing the Glu–Cys–Gly sequence in their primary structure could react with the antibody (Demasi et al., 2008). Other commercially available antibodies appear to have the same problems of sensitivity and specificity (Murakami and Mawatari, 2003; Mawatari and Murakami, 2004; Castellano et al., 2008). Overall, the anti-glutathione antibodies currently available can prove useful for the analysis of individual proteins but do not appear to be appropriate for large-scale detection of glutathionylated proteins by proteomic approaches.
Reduction of Glutathionylated Proteins by GRX
GRXs are small disulfide oxidoreductases belonging to the thioredoxin (TRX) superfamily (Rouhier et al., 2008). GRX was the first enzyme identified as a specific glutathionyl-mixed disulfide oxidoreductase (Gravina and Mieyal, 1993). This catalysis of deglutathionylation is performed much more efficiently by GRXs than by other disulfide oxidoreductases such as TRXs or protein disulfide isomerases (Jung and Thomas, 1996; Zaffagnini et al., 2008). GRXs have also been suggested to play a role in the catalysis of glutathionylation (Starke et al., 2003; Beer et al., 2004). They constitute a multigenic family classified in different subgroups (Lillig et al., 2008). Plant genomes contain around 30 different GRX genes (Lemaire, 2004; Rouhier et al., 2004, 2006). Classical GRXs possess a CPYC active site with two vicinal cysteines forming a glutathione reducible disulfide. These GRXs can catalyze protein disulfide oxidoreduction and deglutathionylation. The latter reaction can occur through a dithiol mechanism involving the formation of a disulfide bond at the active site of GRX but also through a monothiol mechanism that only requires the most N-terminal active site cysteine (Rouhier et al., 2008). On the other hand, both cysteines are required for disulfide reduction. Therefore, a monocysteinic mutant of GRX retaining only the most N-terminal active site cysteine (CPYS) can catalyze protein deglutathionylation but cannot reduce protein disulfides anymore (Nordstrand et al., 1999). A second ubiquitous class of GRX corresponds to proteins harboring a CGFS active site sequence. Despite the presence of only one active site cysteine, some CGFS-type GRXs contain a disulfide involving a partially conserved C-terminal cysteine. These GRXs have been shown to catalyze deglutathionylation, presumably through a dithiol mechanism (Tamarit et al., 2003; Zaffagnini et al., 2008). Moreover, CGFS-type GRXs are apparently not reduced by GSH but by thioredoxin reductases (Fernandes et al., 2005; Zaffagnini et al., 2008).
Based on the properties of GRXs, a method has been developed to identify glutathionylated proteins by GRX reduction (Figure 3). This method, developed by Lind et al. (2002), is based on the use of a monocysteinic mutant of E. coli GRX3 that specifically deglutathionylates proteins in vitro in the presence of GSH. Protein extracts are initially heavily alkylated with N-ethylmaleimide (NEM) to block all free thiols. After reduction by the monocysteinic GRX, the newly accessible thiols are derivatized by NEM-biotin to tag proteins that were initially glutathionylated in the extract. These proteins are then purified by avidin affinity chromatography and identified by proteomic analysis as described above for methods using biotinylated glutathione. This approach allows detection of glutathionylated proteins in more physiological conditions than the labeling methods, since no pretreatment is required. Furthermore, one of its major advantages is to allow detection of proteins already glutathionylated under basal conditions. This approach initially allowed identification of 22 proteins undergoing glutathionylation after diamide treatment of human ECV304 endothelial cells but also 21 proteins glutathionylated in untreated cells (Lind et al., 2002). Considering the number of target proteins identified, the GRX reduction method appears quite sensitive. Combined to LC–MS/MS and elution by biotin competition or acetic acid, this method allowed identification of the glutathionylation sites of -actin, hsp60, and elongation factor 1--1 (Hamnell-Pamment et al., 2005). More recently, this method was used to study the glutathionylation of -ketoglutarate dehydrogenase (Applegate et al., 2008). A similar approach based on the use of streptavidin-FITC has been employed for in-situ visualization of glutathionylated proteins (Reynaert et al., 2006a). Additional glutathionylated proteins could be identified with the same approach by replacing the mutant GRX by other enzymes potentially catalyzing deglutathionylation, such as sulfiredoxin (Findlay et al., 2006). One limitation of this method is linked to the possible specificity of E. coli GRX3 for some glutathionylated proteins. However, this method could also be useful to explore the specificities of GRXs. Indeed, it could be interesting to determine if different proteins are identified after reduction of a protein extract by different GRX mutants.
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Other Methods
Several other methods have been proposed to detect and identify glutathionylated proteins. The GST overlay approach is based on the ability of a GST from Schistosoma japonicum to specifically bind to the glutathione moiety of glutathionylated proteins (Cheng et al., 2005). In this method, proteins from total extracts are separated by SDS–PAGE and blotted onto nitrocellulose membranes that are incubated with biotin-labeled GST. Glutathionylated proteins are then visualized by chemiluminescence after HRP–avidin incubation. This method is, as several of the previous ones, intrinsically limited by the necessity to separate the proteins on a gel and to transfer them on a membrane before glutathionylated proteins can be detected. Moreover, the specificity and sensitivity of this approach are not known. The GST overlay was used to study p53 glutathionylation after immunoprecipitation (Velu et al., 2007) and the glutathionylation of proteins in serum (Nonaka et al., 2007) but it has not yet been used for proteomic analysis.
Several studies have used immobilized glutathione or its analogs to affinity purify proteins containing reactive cysteines susceptible to undergoing glutathionylation. While these methods do not allow in vivo analyses, they may be complementary to techniques based on radiolabeled or biotinylated glutathione for the analysis of glutathionylation targets in vitro. GSH and GSSG affinity matrices allowed identification of seven candidate glutathionylated proteins by Western blot (Niture et al., 2005) and were also used to study the glutathionylation of SERCA (Adachi et al., 2004b) and p53 (Velu et al., 2007). Several proteins undergoing glutathionylation were also shown to bind to GSNO sepharose columns (Klatt et al., 2000). On a larger scale, many techniques based on spectrophotometric assays, HPLC, or mass spectrometry are available for quantification of the total content of glutathionylated proteins in protein extracts (reviewed in Dalle-Donne et al., 2008).
Conclusion
To date, although 35S-cysteine labeling has several major drawbacks, most known glutathionylated proteins were identified with this method. All the methods for proteomic analysis of glutathionylated proteins described above are complementary, since they exhibit different advantages and drawbacks (Table 2). In the future, a combination of all these methods as well as newly developed approaches will be required to get further insights into the diversity of proteins undergoing glutathionylation in vivo in different organisms, different cells, and different physiological conditions.
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3-GLUTATHIONYLATION IN PHOTOSYNTHETIC ORGANISMS |
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TOPAbstractINTRODUCTIONMETHODS FOR IDENTIFICATION AND...3-GLUTATHIONYLATION IN...FUNDING |
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Compared to mammals, much less is known on glutathionylation in photosynthetic organisms where redox proteomic approaches have been mainly focused on the identification of proteins interacting with TRXs (Schurmann and Buchanan, 2008). Overall proteomics allowed identification of around 300 putative TRX targets in different organisms and sub-cellular compartments (Buchanan and Balmer, 2005; Michelet et al., 2006). However, many glutathionylated proteins identified in mammals are found among these possible targets of TRXs. Though some proteins might be regulated by TRX and glutathionylation, this striking overlap suggests that some of the methods used to detect TRX targets and especially TRX affinity columns are also detecting glutathionylated proteins. Therefore, a number of glutathionylated proteins might be currently considered as putative TRX targets and their identification appears important to get more insights into the complex network of redox-regulated processes in plant cells.
To our knowledge, the first plant proteins reported to undergo glutathionylation were GSTs from Arabidopsis thaliana (Dixon et al., 2002). These GSTs differ from other isoforms by containing a cysteine in place of a serine in their active site and are separated in two classes. The first class corresponds to proteins with glutathione-dependent dehydroascorbate reductase (DHAR) and thioltransferase activities like GRXs. The two cytosolic DHAR–GSTs were found to undergo glutathionylation in vitro in the presence of GSSG. The mixed disulfide form was proposed to constitute a key intermediate in the catalytic mechanism for DHA reduction. The second class corresponds to Lambda GSTs (GSTL), which only possess thioltransferase activity. Two of these enzymes are glutathionylated in vitro by GSSG treatment.
A soybean protein tyrosine phosphatase (PTP) was reported to undergo oxidative inactivation during GSSG treatment in vitro (Dixon et al., 2005a). In mammals, it is well established that PTP-1B can be protected from oxidative inactivation by glutathionylation (Barrett et al., 1999a, 1999b) or by formation of a sulfenyl amide (Salmeen et al., 2003; van Montfort et al., 2003). Other phosphatases can also be protected by formation of an intramolecular disulfide (Caselli et al., 1998; Lee et al., 2002; Savitsky and Finkel, 2002). Compared to its mammalian counterparts, the plant enzyme is surprisingly insensitive to inactivation by H2O2 but hypersensitive to GSSG (Dixon et al., 2005a). The inactivation of soybean PTP by GSSG was proposed to be mediated by glutathionylation of a cysteine residue but also by a glutathionylation-triggered formation of a disulfide bond. This inactivation likely protects the enzyme under highly oxidizing conditions.
Glutathionylation of the poplar type II peroxiredoxin, PRXIIB, on its active site cysteine was found to induce dissociation of its non-covalent homodimers into monomers (Noguera-Mazon et al., 2006). This suggested the existence of a redox-dependent dimer–monomer switch in the PRX family.
Human TRX was identified among in vivo glutathionylated targets by 35S-cysteine proteomic analysis and biochemical studies revealed that the modification likely decreases its activity (Casagrande et al., 2002; Fratelli et al., 2002). While non-photosynthetic organisms usually contain one to three TRXs, multiple isoforms are present in plants (Lemaire et al., 2003; Buchanan and Balmer, 2005; Gelhaye et al., 2005; Meyer et al., 2005), some of which undergo glutathionylation in vitro. In the case of poplar mitochondrial TRXh2, glutathionylation of an additional cysteine, distinct from the two active site cysteines, appears to increase the redox potential of the enzyme and is therefore likely to affect its activity (Gelhaye et al., 2004). In Arabidopsis thaliana, nine different TRX isoforms classified into four subtypes (f, m, x and y) are present in chloroplasts (Lemaire et al., 2007). In vitro analyses on Chlamydomonas and Arabidopsis chloroplastic TRXs revealed that only f-type TRXs could undergo glutathionylation (Michelet et al., 2005). These TRXs play a major role in the light regulation of carbon metabolisms and especially the Calvin–Benson cycle. They are oxidized in the dark and reduced in the light by Ferredoxin–Thioredoxin Reductase (FTR). Once reduced, TRXs f are in turn able to reduce specific regulatory disulfides on their target enzymes, including several enzymes of the Calvin–Benson cycle that are inactive in the dark and activated under illumination by f-type TRXs. Glutathionylation of TRX f specifically affects an extra cysteine, conserved in all f-type TRXs, and sharing a similar position, close to the active site disulfide. The glutathionylation of this residue lowers the ability of TRX f to activate its target enzymes in the light. This effect is probably due to a perturbation of the interaction of TRX f with FTR resulting in a decrease of TRX f reduction in the light. Consequently, glutathionylation of TRX f is likely to decrease the activation of all TRX f targets in the light and could therefore constitute a new mechanism of regulation of photosynthetic metabolism under oxidizing conditions (Michelet et al., 2005; Lemaire et al., 2007).
Cytosolic GAPDH, a glycolytic enzyme, was one of the first targets of glutathionylation identified and is one of the best studied glutathionylated enzymes (Schuppe-Koistinen et al., 1994). GAPDH contains a highly reactive active site cysteine whose thiol group is readily oxidized to sulfenic acid in the presence of H2O2. Overoxidation to the sulfonic and sulfinic acid forms leads to irreversible inactivation of the enzyme. However, the enzyme can be protected from this inactivation by glutathionylation triggered by reaction of the cysteine sulfenic acid with reduced glutathione. In plants, cytosolic GAPDH isoforms were also shown to undergo glutathionylation in vitro (Holtgrefe et al., 2008). Plants also contain, in addition to glycolytic isoforms, two GAPDH in chloroplast, A4 and A2B2, which belong to the Calvin–Benson cycle. The NADPH-dependent activity of the major isoform, A2B2, is specifically light regulated by TRX f while the A4 isoform is not. Conversely, A4-GAPDH undergoes glutathionylation in vitro in the presence of H2O2 and GSH while A2B2-GAPDH does not (Zaffagnini et al., 2007). As in the cytosolic isoforms, glutathionylation of A4-GAPDH occurs on the active site cysteine and protects the enzyme from irreversible oxidation. These results suggest that under conditions leading to glutathionylation in chloroplasts, the activity of both GAPDH isoforms is affected, either directly in the case of A4-GAPDH or indirectly through glutathionylation of TRX f in the case of A2B2-GAPDH.
Besides the in vitro studies on purified proteins described above, three proteomic studies aimed at identifying glutathionylated proteins have been performed in photosynthetic organisms and considerably increased the number of known targets. The list of all glutathionylated proteins identified in vivo and in vitro is presented in Table 3. BioGEE allowed identification of two enzymes glutathionylated in vivo in Arabidopsis cell cultures: cytosolic TPI and the Calvin–Benson cycle enzyme fructose-1,6-bisphophate aldolase (Ito et al., 2003). In vitro, recombinant TPI appears to be inactivated by glutathionylation. A second study, on dark-grown Arabidopsis cell cultures treated with BioGSSG and tBOOH, identified nine glutathionylated proteins in vivo: tubulin and β, cytosolic GAPDH, hsc70-1, two isoforms of sucrose synthase, transducin, actin, and acetyl-CoA carboxylase (Dixon et al., 2005b). Most of these targets were previously identified in mammals, suggesting that common redox regulation mechanisms are shared between plants and animals. BioGSSG was also used to identify proteins susceptible to glutathionylation in vitro in cellular extracts from dark-grown Arabidopsis cell cultures (Dixon et al., 2005b). A total of 71 proteins glutathionylated or associated with glutathionylated proteins were identified on 2D gels by peptide mass fingerprinting after DTT elution from the streptavidin affinity column. Interestingly, two targets, methionine synthase and alcohol dehydrogenase, required, to undergo glutathionylation in vitro, a component from Arabidopsis extracts sensitive to cysteine alkylation (Dixon et al., 2005b). Surprisingly, very few chloroplastic proteins were found among glutathionylated proteins in Arabidopsis extracts but this may reflect the use of dark-grown cells. Indeed, chloroplast being the major site of ROS production in the light (Foyer et al., 1997), glutathionylation is likely to be an important regulatory mechanism in this compartment. More recently, 25 proteins glutathionylated in vivo were identified by 35S-cysteine labeling in Chlamydomonas cell cultures grown in the light (Michelet et al., 2008). Among these targets, 18 are located in the chloroplast, confirming the assumption that glutathionylation may be important in this compartment. Three of these targets were produced as recombinant proteins and purified for further analyses in vitro. The glutathionylation of chloroplastic HSP70B occurs specifically on one of its three cysteines and is likely to affect the activity of this molecular chaperone, since the targeted residue is located in the ATPase domain of the protein. In the case of chloroplastic 2-cys PRX, glutathionylation of the two conserved cysteines was found to induce a dimer/monomer transition as previously observed for poplar PRXIIB (Noguera-Mazon et al., 2006). The third studied enzyme, isocitrate lyase, is a key enzyme of acetate assimilation that is reversibly inactivated by glutathionylation of two cysteines (Michelet et al., 2008).
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Overall, in photosynthetic organisms, 36 proteins were found to undergo glutathionylation in vivo while 89 are putative targets identified in vitro (Table 3). All these targets participate in diverse biological processes such as photosynthesis, oxidative stress responses, protein folding, amino acid, ATP metabolism, etc. The presence of many proteins related to stress responses, such as peroxiredoxins or chaperones, was also observed in non-photosynthetic organisms and is not surprising considering that glutathionylation is triggered by pro-oxidizing conditions. Glutathionylation may also be important for the regulation of photosynthesis, especially the Calvin–Benson cycle. Four enzymes of this pathway are well characterized TRX targets that are activated in the light by reduction of a regulatory disulfide (Lemaire et al., 2007). Moreover, proteomic studies, based on monocysteinic affinity columns or derivatization of cysteines exposed after TRX reduction, have identified all 11 Calvin–Benson cycle enzymes as putative TRX targets. In addition to A4-GAPDH and FBP aldolase, two new Calvin–Benson cycle enzymes, phosphoglycerate kinase and ribose-5-phosphate isomerase, were found to be S-thiolated in vivo in Chlamydomonas (Michelet et al., 2008). Considering the similarities between cytosolic and chloroplastic TPI isoforms, the TPI participating in the Calvin–Benson cycle is also likely to be regulated by glutathionylation (Ito et al., 2003). Finally, the glutathionylation of TRX f likely affects the regulation of all TRX-regulated Calvin–Benson cycle enzymes. All these data suggest the existence of a complex cross-talk between TRX and glutathionylation, which deserves further examination (Michelet et al., 2006). We have proposed that glutathionylation could decrease the turnover of the Calvin–Benson cycle under oxidative stress conditions, thereby allowing a redistribution of reducing power in the chloroplast to cope with ROS (Michelet et al., 2005; Lemaire et al., 2007).
In order to understand the role and importance of protein glutathionylation in photosynthetic organisms, the enzymes controlling this post-translational modification will have to be characterized. GRXs likely play a major role in the control of deglutathionylation reactions. The analysis of GRX function is a difficult task because of the presence of 30 different GRX isoforms in plants (Lemaire, 2004; Rouhier et al., 2004, 2006). Nevertheless, during the last 5 years, knowledge on the function and biochemical properties of GRXs in photosynthetic organisms has rapidly increased (reviewed in Rouhier et al., 2008). Plant GRXs were recently shown to be involved in iron–sulfur cluster assembly/biogenesis (Feng et al., 2006; Picciocchi et al., 2007; Rouhier et al., 2007; Bandyopadhyay et al., 2008), petal development (Xing et al., 2005), and plant/pathogen interactions (Ndamukong et al., 2007). Plant GRXs probably also play a role in oxidative stress responses by reducing peroxides (Lee et al., 2002), methionine sulfoxide reductases (Vieira Dos Santos et al., 2007), or peroxiredoxins (Rouhier et al., 2001, 2002; Finkemeier et al., 2005; Gama et al., 2007). We have recently characterized the biochemical properties of two Chlamydomonas GRXs, including their ability to catalyze the deglutathionylation of A4-GAPDH (Zaffagnini et al., 2008). Cytosolic GRX1 is a classical CPYC-type GRX that is reduced by GSH and exhibits DHAR and disulfide reductase activities but can also catalyze deglutathionylation very efficiently. On the other hand, chloroplastic GRX3, which is a CGFS-type GRX, cannot use GSH as an electron donor but is efficiently reduced by FTR in the light. Moreover, GRX3 is probably only able to catalyze deglutathionylation, since it does not exhibit any activity in the DHA or disulfide reduction assays. In contrast, chloroplastic and cytosolic TRXs have almost no deglutathionylation activity. These data suggest that in photosynthetic organisms, GRXs are key enzymes for regulation of deglutathionylation processes. Moreover, these results also indicate that different GRX isoforms may exhibit different biochemical properties. This suggests that the multiple isoforms of GRXs present in plants may constitute, together with TRXs, a complex network of redox regulators that deserve further attention. In the future, the ability of GRXs to catalyze glutathionylation should be analyzed, as well as the potential role of other enzymes such as GSTs or sulfiredoxins. Genetic analyses will also have to be performed to further define the importance of these enzymes in vivo. Our knowledge on glutathionylation in photosynthetic organisms has only started to increase recently. The number of known glutathionylated proteins will probably continue to expand rapidly with the appearance of new proteomic studies. The use of highly sensitive methods under normal conditions could allow identification of enzymes whose catalytic cycle includes formation of a glutathionylated intermediate. The effect of glutathionylation on the new targets will have to be examined carefully, both in vitro and in vivo. In parallel, the physiological conditions triggering glutathionylation in vivo will have to be explored. The combination of these different approaches will be required to get further insights into the role and the importance of glutathionylation in photosynthetic organisms.
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TOPAbstractINTRODUCTIONMETHODS FOR IDENTIFICATION AND...3-GLUTATHIONYLATION IN...FUNDING |
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This work was supported by Agence Nationale de la Recherche Grant JC–45751 and Centre National de la Recherche Scientifique.
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The authors would like to thank Myroslawa Miginiac-Maslow and Laure Michelet for critical reading of the manuscript and helpful suggestions. No conflict of interest declared.
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Adachi T, Pimentel DR, Heibeck T, Hou X, Lee YJ, Jiang B, Ido Y, Cohen RA. S-glutathiolation of Ras mediates redox-sensitive signaling by angiotensin II in vascular smooth muscle cells. J. Biol. Chem. (2004a) 279:29857–29862.
Adachi T, Weisbrod RM, Pimentel DR, Ying J, Sharov VS, Schoneich C, Cohen RA. S-glutathiolation by peroxynitrite activates SERCA during arterial relaxation by nitric oxide. Nat. Med (2004b) 10:1200–1207.
Applegate MA, Humphries KM, Szweda LI. Reversible inhibition of alpha-ketoglutarate dehydrogenase by hydrogen peroxide: glutathionylation and protection of lipoic acid. Biochemistry (2008) 47:473–478.
Aracena P, Tang W, Hamilton SL, Hidalgo C. Effects of S-glutathionylation and S-nitrosylation on calmodulin binding to triads and FKBP12 binding to type 1 calcium release channels. Antioxid. Redox. Signal (2005) 7:870–881.
Aracena-Parks P, Goonasekera SA, Gilman CP, Dirksen RT, Hidalgo C, Hamilton SL. Identification of cysteines involved in S-nitrosylation, S-glutathionylation, and oxidation to disulfides in ryanodine receptor type 1. J. Biol. Chem. (2006) 281:40354–40368.
Bandyopadhyay S, et al. Chloroplast monothiol glutaredoxins as scaffold proteins for the assembly and delivery of [2Fe–2S] clusters. EMBO J (2008) 27:1122–1133.[CrossRef][Web of Science][Medline]
Bandyopadhyay S, Starke DW, Mieyal JJ, Gronostajski RM. Thioltransferase (glutaredoxin) reactivates the DNA-binding activity of oxidation-inactivated nuclear factor I. J. Biol. Chem. (1998) 273:392–397.
Barrett WC, DeGnore JP, Keng YF, Zhang ZY, Yim MB, Chock PB. Roles of superoxide radical anion in signal transduction mediated by reversible regulation of protein–tyrosine phosphatase 1B. J. Biol. Chem. (1999a) 274:34543–34546.
Barrett WC, DeGnore JP, Konig S, Fales HM, Keng YF, Zhang ZY, Yim MB, Chock PB. Regulation of PTP1B via glutathionylation of the active site cysteine 215. Biochemistry (1999b) 38:6699–6705.
Beer SM, Taylor ER, Brown SE, Dahm CC, Costa NJ, Runswick MJ, Murphy MP. Glutaredoxin 2 catalyzes the reversible oxidation and glutathionylation of mitochondrial membrane thiol proteins: implications for mitochondrial redox regulation and antioxidant DEFENSE. J. Biol. Chem (2004) 279:47939–47951.
Brennan JP, Miller JI, Fuller W, Wait R, Begum S, Dunn MJ, Eaton P. The utility of N,N-biotinyl glutathione disulfide in the study of protein S-glutathiolation. Mol. Cell Proteomics (2006) 5:215–225.
Buchanan BB, Balmer Y. Redox regulation: a broadening horizon. Annu. Rev. Plant Biol. (2005) 56:187–220.[CrossRef][Medline]
Cao X, Kambe F, Lu X, Kobayashi N, Ohmori S, Seo H. Glutathionylation of two cysteine residues in paired domain regulates DNA binding activity of Pax-8. J. Biol. Chem. (2005) 280:25901–25906.
Caplan JF, Filipenko NR, Fitzpatrick SL, Waisman DM. Regulation of annexin A2 by reversible glutathionylation. J. Biol. Chem. (2004) 279:7740–7750.
Casagrande S, Bonetto V, Fratelli M, Gianazza E, Eberini I, Massignan T, Salmona M, Chang G, Holmgren A, Ghezzi P. Glutathionylation of human thioredoxin: a possible crosstalk between the glutathione and thioredoxin systems. Proc. Natl Acad. Sci. U S A (2002) 99:9745–9749.
Caselli A, Marzocchini R, Camici G, Manao G, Moneti G, Pieraccini G, Ramponi G. The inactivation mechanism of low molecular weight phosphotyrosine–protein phosphatase by H2O2. J. Biol. Chem. (1998) 273:32554–32560.
Castellano I, Ruocco MR, Cecere F, Di Maro A, Chambery A, Michniewicz A, Parlato G, Masullo M, De Vendittis E. Glutathionylation of the iron superoxide dismutase from the psychrophilic eubacterium Pseudoalteromonas haloplanktis. Biochim. Biophys. Acta (2008) 1784:816–826.
Chai YC, Ashraf SS, Rokutan K, Johnston RB Jr, Thomas JA. S-thiolation of individual human neutrophil proteins including actin by stimulation of the respiratory burst: evidence against a role for glutathione disulfide. Arch. Biochem. Biophys (1994) 310:273–281.[CrossRef][Web of Science][Medline]
Chen YR, Chen CL, Pfeiffer DR, Zweier JL. Mitochondrial complex II in the post-ischemic heart: oxidative injury and the role of protein S-glutathionylation. J. Biol. Chem. (2007) 282:32640–32654.
Cheng G, Ikeda Y, Iuchi Y, Fujii J. Detection of S-glutathionylated proteins by glutathione S-transferase overlay. Arch. Biochem. Biophys (2005) 435:42–49.[CrossRef][Web of Science][Medline]
Chu F, Ward NE, O'Brian CA. PKC isozyme S-cysteinylation by cystine stimulates the pro-apoptotic isozyme PKC delta and inactivates the oncogenic isozyme PKC epsilon. Carcinogenesis (2003) 24:317–325.
Conway ME, Coles SJ, Islam MM, Hutson SM. Regulatory control of human cytosolic branched-chain aminotransferase by oxidation and S-glutathionylation and its interactions with redox sensitive neuronal proteins. Biochemistry (2008) 47:5465–5479.[CrossRef][Web of Science][Medline]
Craghill J, Cronshaw AD, Harding JJ. The identification of a reaction site of glutathione mixed-disulphide formation on S-crystallin in human lens. Biochem. J. (2004) 379:595–600.
Cross JV, Templeton DJ. Oxidative stress inhibits MEKK1 by site-specific glutathionylation in the ATP-binding domain. Biochem. J. (2004) 381:675–683.
Dalle-Donne I, Milzani A, Gagliano N, Colombo R, Giustarini D, Rossi R. Molecular mechanisms and potential clinical significance of S-glutathionylation. Antioxid. Redox. Signal (2008) 10:445–473.
Dalle-Donne I, Rossi R, Giustarini D, Colombo R, Milzani A. Actin S-glutathionylation: evidence against a thiol–disulphide exchange mechanism. Free Radic. Biol. Med (2003) 35:1185–1193.[CrossRef][Web of Science][Medline]
Dalle-Donne I, Rossi R, Giustarini D, Colombo R, Milzani A. S-glutathionylation in protein redox regulation. Free Radic. Biol. Med (2007) 43:883–898.[CrossRef][Web of Science][Medline]
Demasi M, Piassa Filho GM, Castro LM, Ferreira JC, Rioli V, Ferro ES. Oligomerization of the cysteinyl-rich oligopeptidase EP24.15 is triggered by S-glutathionylation. Free Radic. Biol. Med (2008) 44:1180–1190.[CrossRef][Web of Science][Medline]
Dixon DP, Davis BG, Edwards R. Functional divergence in the glutathione transferase superfamily in plants: identification of two classes with putative functions in redox homeostasis in Arabidopsis thaliana. J. Biol. Chem. (2002) 277:30859–30869.
Dixon DP, Fordham-Skelton AP, Edwards R. Redox regulation of a soybean tyrosine-specific protein phosphatase. Biochemistry (2005a) 44:7696–7703.[CrossRef][Medline]
Dixon DP, Skipsey M, Grundy NM, Edwards R. Stress-induced protein S-glutathionylation in Arabidopsis. Plant Physiol (2005b) 138:2233–2244.
Eaton P, Byers HL, Leeds N, Ward MA, Shattock MJ. Detection, quantitation, purification, and identification of cardiac proteins S-thiolated during ischemia and reperfusion. J. Biol. Chem. (2002a) 277:9806–9811.
Eaton P, Fuller W, Shattock MJ. S-thiolation of HSP27 regulates its multimeric aggregate size independently of phosphorylation. J. Biol. Chem. (2002b) 277:21189–21196.
Eaton P, Wright N, Hearse DJ, Shattock MJ. Glyceraldehyde phosphate dehydrogenase oxidation during cardiac ischemia and reperfusion. J. Mol. Cell Cardiol (2002c) 34:1549–1560.[CrossRef][Web of Science][Medline]
Feng Y, Zhong N, Rouhier N, Hase T, Kusunoki M, Jacquot JP, Jin C, Xia B. Structural insight into poplar glutaredoxin C1 with a bridging iron–sulfur cluster at the active site. Biochemistry (2006) 45:7998–8008.
Fernandes AP, Fladvad M, Berndt C, Andresen C, Lillig CH, Neubauer P, Sunnerhagen M, Holmgren A, Vlamis-Gardikas A. A novel monothiol glutaredoxin (Grx4) from Escherichia coli can serve as a substrate for thioredoxin reductase. J. Biol. Chem. (2005) 280:24544–24552.
Fiaschi T, Cozzi G, Raugei G, Formigli L, Ramponi G, Chiarugi P. Redox regulation of beta-actin during integrin-mediated cell adhesion. J. Biol. Chem. (2006) 281:22983–22991.
Findlay VJ, Townsend DM, Morris TE, Fraser JP, He L, Tew KD. A novel role for human sulfiredoxin in the reversal of glutathionylation. Cancer Res. (2006) 66:6800–6806.
Finkemeier I, Goodman M, Lamkemeyer P, Kandlbinder A, Sweetlove LJ, Dietz KJ. The mitochondrial type II peroxiredoxin F is essential for redox homeostasis and root growth of Arabidopsis thaliana under stress. J. Biol. Chem. (2005) 280:12168–12180.
Foyer CH, Noctor G. Redox homeostasis and antioxidant signaling: a metabolic interface between stress perception and physiological responses. Plant Cell. (2005) 17:1866–1875.
Foyer CH, Lelandais M, Kunert K-J. Photo-oxidative stress in plants. Physiol. Plant (1997) 92:696–717.
Fratelli M, Demol H, Puype M, Casagrande S, Villa P, Eberini I, Vandekerckhove J, Gianazza E, Ghezzi P. Identification of proteins undergoing glutathionylation in oxidatively stressed hepatocytes and hepatoma cells. Proteomics (2003) 3:1154–1161.[CrossRef][Web of Science][Medline]
Fratelli M, et al. Identification by redox proteomics of glutathionylated proteins in oxidatively stressed human T lymphocytes. Proc. Natl Acad. Sci. U S A (2002) 99:3505–3510.
Gallogly MM, Mieyal JJ. Mechanisms of reversible protein glutathionylation in redox signaling and oxidative stress. Curr. Opin. Pharmacol (2007) 7:381–391.[CrossRef]
Gama F, Keech O, Eymery F, Finkemeier I, Gelhaye E, Gardestrom P, Dietz KJ, Rey P, Jacquot J-P, Rouhier N. The mitochondrial type II peroxiredoxin from poplar. Physiologia Plantarum (2007) 129:196–206.[CrossRef]
Gelhaye E, et al. A specific form of thioredoxin h occurs in plant mitochondria and regulates the alternative oxidase. Proc. Natl Acad. Sci. U S A (2004) 101:14545–14550.
Gelhaye E, Rouhier N, Navrot N, Jacquot JP. The plant thioredoxin system. Cell Mol. Life Sci. (2005) 62:24–35.[CrossRef][Web of Science][Medline]
Ghezzi P. Regulation of protein function by glutathionylation. Free Radic. Res. (2005a) 39:573–580.[Web of Science][Medline]
Ghezzi P. Oxidoreduction of protein thiols in redox regulation. Biochem. Soc. Trans. (2005b) 33:1378–1381.[CrossRef][Web of Science][Medline]
Ghezzi P, Di Simplicio P. Glutathionylation pathways in drug response. Curr. Opin. Pharmacol (2007) 7:398–403.[CrossRef]
Ghezzi P, et al. Redox regulation of cyclophilin A by glutathionylation. Proteomics (2006) 6:817–825.[CrossRef][Web of Science][Medline]
Gilbert HF. Redox control of enzyme activities by thiol/disulfide exchange. Methods Enzymol (1984) 107:330–351.[Web of Science][Medline]
Grant CM, Quinn KA, Dawes IW. Differential protein S-thiolation of glyceraldehyde-3-phosphate dehydrogenase isoenzymes influences sensitivity to oxidative stress. Mol. Cell Biol. (1999) 19:2650–2656.
Gravina SA, Mieyal JJ. Thioltransferase is a specific glutathionyl mixed disulfide oxidoreductase. Biochemistry (1993) 32:3368–3376.[CrossRef][Medline]
Hamnell-Pamment Y, Lind C, Palmberg C, Bergman T, Cotgreave IA. Determination of site-specificity of S-glutathionylated cellular proteins. Biochem. Biophys. Res. Commun (2005) 332:362–369.[Web of Science][Medline]
Han D, Canali R, Garcia J, Aguilera R, Gallaher TK, Cadenas E. Sites and mechanisms of aconitase inactivation by peroxynitrite: modulation by citrate and glutathione. Biochemistry (2005) 44:11986–11996.
Hidalgo C, Sanchez G, Barrientos G, Aracena-Parks P. A transverse tubule NADPH oxidase activity stimulates calcium release from isolated triads via ryanodine receptor type 1 S -glutathionylation. J. Biol. Chem. (2006) 281:26473–26482.
Hochgräfe F, Mostertz J, Pother DC, Becher D, Helmann JD, Hecker M. S-cysteinylation is a general mechanism for thiol protection of Bacillus subtilis proteins after oxidative stress. J. Biol. Chem. (2007) 282:25981–25985.
Holtgrefe S, Gohlke J, Starmann J, Druce S, Klocke S, Altmann B, Wojtera J, Lindermayr C, Scheibe R. Regulation of plant cytosolic glyceraldehyde 3-phosphate dehydrogenase isoforms by thiol modifications. Physiol. Plant (2008) 133:211–228.
Hoppe G, Chai YC, Crabb JW, Sears J. Protein s-glutathionylation in retinal pigment epithelium converts heat shock protein 70 to an active chaperone. Exp. Eye Res. (2004) 78:1085–1092.[CrossRef]
Huang Z, Pinto JT, Deng H, Richie JP Jr. Inhibition of caspase-3 activity and activation by protein glutathionylation. Biochem. Pharmacol (2008) 75:2234–2244.
Humphries KM, Deal MS, Taylor SS. Enhanced dephosphorylation of cAMP-dependent protein kinase by oxidation and thiol modification. J. Biol. Chem. (2005) 280:2750–2758.
Humphries KM, Juliano C, Taylor SS. Regulation of cAMP-dependent protein kinase activity by glutathionylation. J. Biol. Chem. (2002) 277:43505–43511.
Hurd TR, Costa NJ, Dahm CC, Beer SM, Brown SE, Filipovska A, Murphy MP. Glutathionylation of mitochondrial proteins. Antioxid. Redox. Signal (2005) 7:999–1010.
Hurd TR, Raquejo R, Filipovska A, Brown S, Prime TA, Robinson AJ, Fearnley IM, Murphy MP. Complex I within oxidatively-stressed bovine heart mitochondria is glutathionylated on Cys-531 and Cys-704 of the 75-kDa subunit: potential role of Cys residues in decreasing oxidative damage. J. Biol. Chem (2008).
Ito H, Iwabuchi M, Ogawa K. The sugar-metabolic enzymes aldolase and triose-phosphate isomerase are targets of glutathionylation in Arabidopsis thaliana: detection using biotinylated glutathione. Plant Cell Physiol (2003) 44:655–660.
Jacob C, Knight I, Winyard PG. Aspects of the biological redox chemistry of cysteine: from simple redox responses to sophisticated signalling pathways. Biol. Chem. (2006) 387:1385–1397.[CrossRef][Web of Science][Medline]
Jahngen-Hodge J, Obin MS, Gong X, Shang F, Nowell TR Jr, Gong J, Abasi H, Blumberg J, Taylor A. Regulation of ubiquitin-conjugating enzymes by glutathione following oxidative stress. J. Biol. Chem. (1997) 272:28218–28226.
Jönsson TJ, Lowther WT. The peroxiredoxin repair proteins. Subcell. Biochem (2007) 44:115–141.
Jung CH, Thomas JA. S-glutathiolated hepatocyte proteins and insulin disulfides as substrates for reduction by glutaredoxin, thioredoxin, protein disulfide isomerase, and glutathione. Arch. Biochem. Biophys (1996) 335:61–72.[CrossRef][Web of Science][Medline]
Kil IS, Kim SY, Park JW. Glutathionylation regulates IkappaB. Biochem. Biophys. Res. Commun (2008) 373:169–173.[CrossRef][Web of Science][Medline]
Klatt P, Molina EP, Lamas S. Nitric oxide inhibits c-Jun DNA binding by specifically targeted S-glutathionylation. J. Biol. Chem. (1999a) 274:15857–15864.
Klatt P, Molina EP, Lamas S. Nitric oxide inhibits c-Jun DNA binding by specifically targeted S-glutathionylation. J. Biol. Chem. (1999b) 274:15857–15864.
Klatt P, Molina EP, De Lacoba MG, Padilla CA, Martinez-Galesteo E, Barcena JA, Lamas S. Redox regulation of c-Jun DNA binding by reversible S-glutathiolation. Faseb. J. (1999c) 13:1481–1490.
Klatt P, Pineda Molina E, Perez-Sala D, Lamas S. Novel application of S-nitrosoglutathione–Sepharose to identify proteins that are potential targets for S-nitrosoglutathione-induced mixed-disulphide formation. Biochem. J. (2000) 349:567–578.
Lampela O, Juffer AH, Rauk A. Conformational analysis of glutathione in aqueous solution with molecular dynamics. J. Phys. Chem. A (2003) 107:9208–9220.
Landino LM, Moynihan KL, Todd JV, Kennett KL. Modulation of the redox state of tubulin by the glutathione/glutaredoxin reductase system. Biochem. Biophys. Res. Commun (2004) 314:555–560.[CrossRef][Web of Science][Medline]
Lee JW, Soonsanga S, Helmann JD. A complex thiolate switch regulates the Bacillus subtilis organic peroxide sensor OhrR. Proc. Natl Acad. Sci. U S A (2007) 104:8743–8748.
Lee SR, Yang KS, Kwon J, Lee C, Jeong W, Rhee SG. Reversible inactivation of the tumor suppressor PTEN by H2O2. J. Biol. Chem. (2002) 277:20336–20342.
Lemaire SD. The glutaredoxin family in oxygenic photosynthetic organisms. Photosynth. Res. (2004) 79:305–318.[CrossRef][Web of Science][Medline]
Lemaire SD, Collin V, Keryer E, Issakidis-Bourguet E, Lavergne D, Miginiac-Maslow M. Chlamydomonas reinhardtii: a model organism for the study of the thioredoxin family. Plant Physiol. Biochem (2003) 41:513–521.
Lemaire SD, Michelet L, Zaffagnini M, Massot V, Issakidis-Bourguet E. Thioredoxins in chloroplasts. Curr. Genet. (2007) 51:343–365.[CrossRef][Web of Science][Medline]
Leonberg AK, Chai YC. The functional role of cysteine residues for c-Abl kinase activity. Mol. Cell Biochem (2007) 304:207–212.[CrossRef][Web of Science][Medline]
Lillig CH, Berndt C, Holmgren A. Glutaredoxin systems. Biochim. Biophys. Acta (2008) 1780:1304–1317.
Lind C, Gerdes R, Hamnell Y, Schuppe-Koistinen I, von Lowenhielm HB, Holmgren A, Cotgreave IA. Identification of S-glutathionylated cellular proteins during oxidative stress and constitutive metabolism by affinity purification and proteomic analysis. Arch. Biochem. Biophys (2002) 406:229–240.[CrossRef][Web of Science][Medline]
Lind C, Gerdes R, Schuppe-Koistinen I, Cotgreave IA. Studies on the mechanism of oxidative modification of human glyceraldehyde-3-phosphate dehydrogenase by glutathione: catalysis by glutaredoxin. Biochem. Biophys. Res. Commun (1998) 247:481–486.[CrossRef][Web of Science][Medline]
Manevich Y, Feinstein SI, Fisher AB. Activation of the antioxidant enzyme 1-CYS peroxiredoxin requires glutathionylation mediated by heterodimerization with pi GST. Proc. Natl Acad. Sci. U S A (2004) 101:3780–3785.
Mawatari S, Murakami K. Different types of glutathionylation of hemoglobin can exist in intact erythrocytes. Arch. Biochem. Biophys (2004) 421:108–114.[CrossRef][Web of Science][Medline]
Meyer AJ, Hell R. Glutathione homeostasis and redox-regulation by sulfhydryl groups. Photosynth. Res. (2005) 86:435–457.[CrossRef][Web of Science][Medline]
Meyer Y, Reichheld JP, Vignols F. Thioredoxins in Arabidopsis and other plants. Photosynth. Res. (2005) 86:419–433.[CrossRef][Web of Science][Medline]
Michelet L, et al. Glutathionylation of chloroplast thioredoxin f is a redox signaling mechanism in plants. Proc. Natl Acad. Sci. U S A (2005) 102:16478–16483.
Michelet L, Zaffagnini M, Massot V, Keryer E, Vanacker H, Miginiac-Maslow M, Issakidis-Bourguet E, Lemaire SD. Thioredoxins, glutaredoxins, and glutathionylation: new crosstalks to explore. Photosynth. Res. (2006) 89:225–245.[CrossRef][Web of Science][Medline]
Michelet L, Zaffagnini M, Vanacker H, Le Marechal P, Marchand C, Schroda M, Lemaire SD, Decottignies P. In vivo targets of S-thiolation in Chlamydomonas reinhardtii. J. Biol. Chem. (2008) 283:21571–21578.
Murakami K, Mawatari S. Oxidation of hemoglobin to methemoglobin in intact erythrocyte by a hydroperoxide induces formation of glutathionyl hemoglobin and binding of alpha-hemoglobin to membrane. Arch. Biochem. Biophys (2003) 417:244–250.[CrossRef][Web of Science][Medline]
Ndamukong I, Abdallat AA, Thurow C, Fode B, Zander M, Weigel R, Gatz C. SA-inducible Arabidopsis glutaredoxin interacts with TGA factors and suppresses JA-responsive PDF1.2 transcription. Plant J (2007) 50:128–139.[CrossRef][Medline]
Newman SF, Sultana R, Perluigi M, Coccia R, Cai J, Pierce WM, Klein JB, Turner DM, Butterfield DA. An increase in S-glutathionylated proteins in the Alzheimer's disease inferior parietal lobule, a proteomics approach. J. Neurosci. Res. (2007) 85:1506–1514.[CrossRef][Web of Science][Medline]
Niture SK, Velu CS, Bailey NI, Srivenugopal KS. S-thiolation mimicry: quantitative and kinetic analysis of redox status of protein cysteines by glutathione-affinity chromatography. Arch. Biochem. Biophys (2005) 444:174–184.[CrossRef][Web of Science][Medline]
Noctor G. Metabolic signalling in defence and stress: the central roles of soluble redox couples. Plant Cell Environ (2006) 29:409–425.[CrossRef][Medline]
Noctor G, Foyer CH. Ascorbate and glutathione: keeping active oxygen under control. Annu. Rev. Plant Physiol. Plant Mol. Biol. (1998) 49:249–279.[CrossRef]
Noguera-Mazon V, Lemoine J, Walker O, Rouhier N, Salvador A, Jacquot JP, Lancelin JM, Krimm I. Glutathionylation induces the dissociation of 1-Cys D-peroxiredoxin non-covalent homodimer. J. Biol. Chem. (2006) 281:31736–31742.
Nonaka K, et al. Serum levels of S-glutathionylated proteins as a risk-marker for arteriosclerosis obliterans. Circ. J. (2007) 71:100–105.
Nordstrand K, Slund F, Holmgren A, Otting G, Berndt KD. NMR structure of Escherichia coli glutaredoxin 3-glutathione mixed disulfide complex: implications for the enzymatic mechanism. J. Mol. Biol. (1999) 286:541–552.[CrossRef][Web of Science][Medline]
Ogawa K. Glutathione-associated regulation of plant growth and stress responses. Antioxid. Redox. Signal (2005) 7:973–981.
Park S, Imlay JA. High levels of intracellular cysteine promote oxidative DNA damage by driving the fenton reaction. J. Bacteriol (2003) 185:1942–1950.
Pauly N, Pucciariello C, Mandon K, Innocenti G, Jamet A, Baudouin E, Herouart D, Frendo P, Puppo A. Reactive oxygen and nitrogen species and glutathione: key players in the legume–Rhizobium symbiosis. J. Exp. Bot (2006) 57:1769–1776.
Picciocchi A, Saguez C, Boussac A, Cassier-Chauvat C, Chauvat F. CGFS-type monothiol glutaredoxins from the cyanobacterium Synechocystis PCC6803 and other evolutionary distant model organisms possess a glutathione-ligated [2Fe–2S] cluster. Biochemistry (2007) 46:15018–15026.[CrossRef][Web of Science][Medline]
Pineda-Molina E, Klatt P, Vazquez J, Marina A, Garcia de Lacoba M, Perez-Sala D, Lamas S. Glutathionylation of the p50 subunit of NF-
B: a mechanism for redox-induced inhibition of DNA binding. Biochemistry (2001) 40:14134–14142.[CrossRef][Web of Science][Medline]
Prinarakis E, Chantzoura E, Thanos D, Spyrou G. S-glutathionylation of IRF3 regulates IRF3-CBP interaction and activation of the IFN beta pathway. EMBO J (2008) 27:865–875.[CrossRef][Web of Science][Medline]
Rao RK, Clayton LW. Regulation of protein phosphatase 2A by hydrogen peroxide and glutathionylation. Biochem. Biophys. Res. Commun (2002) 293:610–616.[CrossRef][Web of Science][Medline]
Ravichandran V, Seres T, Moriguchi T, Thomas JA, Johnston RB Jr. S-thiolation of glyceraldehyde-3-phosphate dehydrogenase induced by the phagocytosis-associated respiratory burst in blood monocytes. J. Biol. Chem. (1994) 269:25010–25015.
Reynaert NL, Ckless K, Guala AS, Wouters EF, van der Vliet A, Janssen-Heininger YM. In situ detection of S-glutathionylated proteins following glutaredoxin-1 catalyzed cysteine derivatization. Biochim. Biophys. Acta (2006a) 1760:380–387.
Reynaert NL, et al. Dynamic redox control of NF-kappaB through glutaredoxin-regulated S-glutathionylation of inhibitory kappaB kinase beta. Proc. Natl Acad. Sci. U S A (2006b) 103:13086–13091.
Rinna A, Torres M, Forman HJ. Stimulation of the alveolar macrophage respiratory burst by ADP causes selective glutathionylation of protein tyrosine phosphatase 1B. Free Radic. Biol. Med (2006) 41:86–91.[CrossRef][Web of Science][Medline]
Rizzello A, Ciardiello MA, Acierno R, Carratore V, Verri T, di Prisco G, Storelli C, Maffia M. Biochemical characterization of a S-glutathionylated carbonic anhydrase isolated from gills of the Antarctic icefish Chionodraco hamatus. Protein J (2007) 26:335–348.[CrossRef][Web of Science][Medline]
Rokutan K, Johnston RB Jr, Kawai K. Oxidative stress induces S-thiolation of specific proteins in cultured gastric mucosal cells. Am. J. Physiol (1994) 266:G247–G254.
Rokutan K, Thomas JA, Johnston RB Jr. Phagocytosis and stimulation of the respiratory burst by phorbol diester initiate S-thiolation of specific proteins in macrophages. J. Immunol (1991) 147:260–264.[Abstract]
Rouhier N, Couturier J, Jacquot JP. Genome-wide analysis of plant glutaredoxin systems. J. Exp. Bot (2006) 57:1685–1696.
Rouhier N, et al. Functional, structural, and spectroscopic characterization of a glutathione-ligated [2Fe–2S] cluster in poplar glutaredoxin C1. Proc. Natl Acad. Sci. U S A (2007) 104:7379–7384.
Rouhier N, Gelhaye E, Jacquot JP. Glutaredoxin-dependent peroxiredoxin from poplar: protein–protein interaction and catalytic mechanism. J. Biol. Chem. (2002) 277:13609–13614.
Rouhier N, Gelhaye E, Jacquot JP. Plant glutaredoxins: still mysterious reducing systems. Cell Mol. Life Sci. (2004) 61:1266–1277.[CrossRef][Web of Science][Medline]
Rouhier N, Gelhaye E, Sautiere PE, Brun A, Laurent P, Tagu D, Gérard J, de Faÿ E, Meyer Y, Jacquot JP. Isolation and characterization of a new peroxiredoxin from poplar sieve tubes that uses either glutaredoxin or thioredoxin as a proton donor. Plant Physiol (2001) 127:1299–1309.
Rouhier N, Lemaire SD, Jacquot JP. The role of glutathione in photosynthetic organisms: emerging functions for glutaredoxins and glutathionylation. Annu. Rev. Plant Biol. (2008) 59:143–166.[CrossRef][Medline]
Sadidi M, Geddes TJ, Kuhn DM. S-thiolation of tyrosine hydroxylase by reactive nitrogen species in the presence of cysteine or glutathione. Antioxid. Redox. Signal (2005) 7:863–869.
Salmeen A, Andersen JN, Myers MP, Meng TC, Hinks JA, Tonks NK, Barford D. Redox regulation of protein tyrosine phosphatase 1B involves a sulphenyl-amide intermediate. Nature (2003) 423:769–773.[CrossRef][Web of Science][Medline]
Savitsky PA, Finkel T. Redox regulation of Cdc25C. J. Biol. Chem. (2002) 277:20535–20540.
Schuppe-Koistinen I, Moldeus P, Bergman T, Cotgreave IA. S-thiolation of human endothelial cell glyceraldehyde-3-phosphate dehydrogenase after hydrogen peroxide treatment. Eur. J. Biochem (1994) 221:1033–1037.[Web of Science][Medline]
Schurmann P, Buchanan BB. The ferredoxin/thioredoxin system of oxygenic photosynthesis. Antioxid. Redox. Signal (2008) 10:1235–1274.
Seres T, Ravichandran V, Moriguchi T, Rokutan K, Thomas JA, Johnston RB Jr. Protein S-thiolation and dethiolation during the respiratory burst in human monocytes: a reversible post-translational modification with potential for buffering the effects of oxidant stress. J. Immunol (1996) 156:1973–1980.[Abstract]
Shelton MD, Kern TS, Mieyal JJ. Glutaredoxin regulates nuclear factor kappa-B and intercellular adhesion molecule in Muller cells: model of diabetic retinopathy. J. Biol. Chem. (2007) 282:12467–12474.
Shenton D, Grant CM. Protein S-thiolation targets glycolysis and protein synthesis in response to oxidative stress in the yeast Saccharomyces cerevisiae. Biochem. J. (2003) 374:513–519.
Starke DW, Chock PB, Mieyal JJ. Glutathione-thiyl radical scavenging and transferase properties of human glutaredoxin (thioltransferase): potential role in redox signal transduction. J. Biol. Chem. (2003) 278:14607–14613.
Sullivan DM, Wehr NB, Fergusson MM, Levine RL, Finkel T. Identification of oxidant-sensitive proteins: TNF- induces protein glutathiolation. Biochemistry (2000) 39:11121–11128.[CrossRef][Web of Science][Medline]
Tamarit J, Belli G, Cabiscol E, Herrero E, Ros J. Biochemical characterization of yeast mitochondrial Grx5 monothiol glutaredoxin. J. Biol. Chem. (2003) 278:25745–25751.
Townsend DM. S-glutathionylation: indicator of cell stress and regulator of the unfolded protein response. Mol. Interv (2007) 7:313–324.
Townsend DM, Findlay VJ, Fazilev F, Ogle M, Fraser J, Saavedra JE, Ji X, Keefer LK, Tew KD. A glutathione S-transferase pi-activated prodrug causes kinase activation concurrent with S-glutathionylation of proteins. Mol. Pharmacol (2006) 69:501–508.
van Montfort RL, Congreve M, Tisi D, Carr R, Jhoti H. Oxidation state of the active-site cysteine in protein tyrosine phosphatase 1B. Nature (2003) 423:773–777.[CrossRef][Web of Science][Medline]
Velu CS, Niture SK, Doneanu CE, Pattabiraman N, Srivenugopal KS. Human p53 is inhibited by glutathionylation of cysteines present in the proximal DNA-binding domain during oxidative stress. Biochemistry (2007) 46:7765–7780.[CrossRef][Web of Science][Medline]
Vieira Dos Santos C, Laugier E, Tarrago L, Massot V, Issakidis-Bourguet E, Rouhier N, Rey P. Specificity of thioredoxins and glutaredoxins as electron donors to two distinct classes of Arabidopsis plastidial methionine sulfoxide reductases B. FEBS Lett. (2007) 581:4371–4376.[CrossRef][Web of Science][Medline]
Wang J, Tekle E, Oubrahim H, Mieyal JJ, Stadtman ER, Chock PB. Stable and controllable RNA interference: investigating the physiological function of glutathionylated actin. Proc. Natl Acad. Sci. U S A (2003) 100:5103–5106.
Wang W, Oliva C, Li G, Holmgren A, Lillig CH, Kirk KL. Reversible silencing of CFTR chloride channels by glutathionylation. J. Gen. Physiol (2005) 125:127–141.
Ward NE, Chu F, O'Brian CA. Regulation of protein kinase C isozyme activity by S-glutathiolation. Methods Enzymol (2002) 353:89–100.[Web of Science][Medline]
Ward NE, Stewart JR, Ioannides CG, O'Brian CA. Oxidant-induced S-glutathiolation inactivates protein kinase C-alpha (PKC-alpha): a potential mechanism of PKC isozyme regulation. Biochemistry (2000) 39:10319–10329.[CrossRef][Web of Science][Medline]
West MB, Hill BG, Xuan YT, Bhatnagar A. Protein glutathiolation by nitric oxide: an intracellular mechanism regulating redox protein modification. Faseb. J. (2006) 20:1715–1717.
Xing S, Rosso MG, Zachgo S. ROXY1, a member of the plant glutaredoxin family, is required for petal development in Arabidopsis thaliana. Development (2005) 132:1555–1565.
Zaffagnini M, Michelet L, Marchand C, Sparla F, Decottignies P, Le Marechal P, Miginiac-Maslow M, Noctor G, Trost P, Lemaire SD. The thioredoxin-independent isoform of chloroplastic glyceraldehyde-3-phosphate dehydrogenase is selectively regulated by glutathionylation. FEBS J (2007) 274:212–226.[CrossRef][Medline]
Zaffagnini M, Michelet L, Massot V, Trost P, Lemaire SD. Biochemical characterization of glutaredoxins from Chlamydomonas reinhardtii reveals the unique properties of a chloroplastic CGFS-type glutaredoxin. J. Biol. Chem. (2008) 283:8868–8876.
Ziegler DM. Role of reversible oxidation-reduction of enzyme thiols-disulfides in metabolic regulation. Annu. Rev. Biochem (1985) 54:305–329.[Web of Science][Medline]
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