JBC Origene Your Gene Company

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M705953200 on August 24, 2007

J. Biol. Chem., Vol. 282, Issue 43, 31429-31436, October 26, 2007
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
282/43/31429    most recent
M705953200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ma, L.-H.
Right arrow Articles by Wood, M. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ma, L.-H.
Right arrow Articles by Wood, M. J.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

Molecular Mechanism of Oxidative Stress Perception by the Orp1 Protein*

Li-Hua Ma, Christina L. Takanishi, and Matthew J. Wood, Recipient of American Heart Association Award 0635328N1

From the Department of Environmental Toxicology, University of California, Davis, California 95616

Received for publication, July 20, 2007 , and in revised form, August 21, 2007.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In this study we investigated the molecular mechanism by which the Orp1 (Gpx3) protein in Saccharomyces cerevisiae senses and reacts with hydrogen peroxide. Upon exposure to H2O2 Orp1Cys36 forms a disulfide-bonded complex with the C-terminal domain of the Yap1 protein (Yap1-cCRD). We used 4-nitrobenzo-2-oxa-1,3-diazole to identify a cysteine sulfenic acid (Cys-SOH) modification that forms on Cys36 of Orp1Cys36 upon exposure to H2O2. Under similar conditions, neither Cys82 of Orp1Cys82 nor Cys598 of Yap1 forms Cys-SOH. A homology-based molecular model of Orp1 suggests that the structure of the active site of Orp1 is similar to that found in mammalian selenocysteine glutathione peroxidases. Proposed active site residues Gln70 and Trp125 form a catalytic triad with Cys36 in the Orp1 molecular model. The remainder of the active site pocket is formed by Phe38, Asn126, and Phe127, which are evolutionarily conserved residues. We made Q70A and W125A mutants and tested the ability of these mutants to form Cys-SOH in response to H2O2. Both mutants were unable to form Cys-SOH and did not form a H2O2-inducible disulfide-bonded complex with Yap1-cCRD. The pKa of Cys36 was determined to be 5.1, which is 3.2 pH units lower than that of a free cysteine (8.3). In contrast, Orp1 Cys82 (the resolving cysteine) has a pKa value of 8.3. The pKa of Cys36 in the Q70A and W125A mutants is also 8.3, demonstrating the importance of these residues in modulating the nucleophilic character of Cys36. Finally, we show that S. cerevisiae strains with ORP1 Q70A and W125A mutations are less tolerant to H2O2 than those containing wild-type ORP1. The results of our study suggest that attempts to identify novel redox-regulated proteins and signal transduction pathways should focus on characterization of low pKa cysteines.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Oxidative stress has been defined as a cellular disturbance in the prooxidant-antioxidant balance in favor of prooxidants (1). Cellular antioxidant defense and detoxification pathways are formidable and allow organisms to limit the 105 oxidative DNA lesions that are estimated to form each day (2). An enormous body of evidence suggests that the production of reactive oxygen species (ROS)2 induces cellular damage and contributes to the etiology of degenerative diseases such as cancer (3–5). ROS are the main causes of oxidative stress and can include superoxide anion (Formula), hydrogen peroxide (H2O2), hydroxyl radical (HO·), and alkyl hydroperoxides (ROOH). For example, exposure to the prevalent environmental toxicant arsenic induces oxidative stress in the form of increased levels of ROS (6, 7). If allowed to accumulate unchecked, these molecules exceed the normal antioxidant buffering capacity of the cell, leading to indiscriminate damage of cellular components, including DNA, proteins, and lipids (8–11).

Recent studies have demonstrated that ROS, such as hydrogen peroxides and alkyl hydroperoxides, can regulate signal transduction pathways, transcription factors, and gene expression in a variety of prokaryotic and eukaryotic organisms (12–19). Of particular relevance to this work are the common cysteine thiol switches that can regulate the biological function of proteins (20, 21). For example, the Escherichia coli transcription factor OxyR contains two conserved cysteine residues that are essential for oxidant perception and regulation (22). Upon exposure to H2O2, Cys199 of OxyR reacts with H2O2 to form a transient Cys-SOH intermediate (23). The Cys199 Cys-SOH immediately reacts with Cys208 and forms a disulfide bond, which results in increased DNA binding affinity and OxyR target gene expression.

The budding yeast Saccharomyces cerevisiae is a valuable eukaryotic model for understanding the molecular and biochemical mechanisms involved in H2O2-regulated signal transduction pathways (24). The major H2O2 response pathway in S. cerevisiae involves the transcription factor Yap1 and the oxidant receptor protein Orp1 (12, 13, 25–27). The Yap1 oxidative stress response pathway is regulated by a novel disulfide bond-relay cascade and controls the expression of ~70 genes in response to H2O2 (28, 29). Orp1, which is homologous to glutathione peroxidase enzymes (hGpx), is responsible for H2O2 perception and subsequent catalysis of disulfide bond formation in Yap1 (12). The proposed peroxidase mechanism for Orp1 involves the initial H2O2 reaction with Cys36 (12). Based on the homology of Orp1 with selenocysteine containing hGpx enzymes, it has been proposed that Orp1 forms a sulfenic acid intermediate upon reaction with peroxides. Ultimately the Cys-SOH intermediate is resolved by Cys82 to form an intramolecular disulfide bond. This disulfide bond is then reduced via the thioredoxin pathway (12). For this reason, this class of non-selenocysteine hGpx-like enzymes has been classified as thioredoxin-dependent peroxidases or thiol peroxidases, with a similar function and mechanism to the peroxiredoxin family of proteins (30). The role of Orp1 in regulation of Yap1 involves the reaction of Cys598 in Yap1 with Cys36 resulting in a mixed disulfide bond complex. This complex has been captured in vivo through site-directed mutagenesis of the resolving Cys303 residue in Yap1 (12). The mixed disulfide is then resolved by Cys303 of Yap1 to form an intramolecular disulfide bond in Yap1. The Yap1 disulfide bond results in the masking of a nuclear export sequence in Yap1, nuclear accumulation of Yap1, and increased gene expression of Yap1 target genes (18, 28).

The molecular mechanism by which the Orp1 class of peroxiredoxins perceives hydrogen peroxide and alkyl hydroperoxide has not been characterized. In this study we demonstrate that upon reaction with H2O2, Orp1Cys36 forms a transient Cys-SOH on Cys36. Our data suggest that the reactivity of Cys36 is because of its extremely low pKa of 5.1. We also present a homology model of the Orp1 structure, which indicates that it has an active site structure similar to mammalian glutathione peroxidase. Mutation of the putative Gln70- or Trp125-active site residues to alanine abolishes H2O2-induced Cys36 Cys-SOH formation and reverts the pKa of Cys36 to 8.3.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals—7-Chloro-4-nitrobenzo-2-oxa-1,3-diazole (NBD-Cl), beta-NADPH, iodoacetamide, thioredoxin, and thioredoxin reductase were purchased from Sigma. Dithiothreitol (DTT), H2O2, and 2-mercaptoethanol were purchased from Fisher. (2-Pyridyl)-dithiobimane (PDT-Bimane) was purchased from Toronto Research Biochemicals. Other chemicals were of the highest quality commercially available.

Protein Cloning, Expression, and Purification—The ORP1 gene was cloned from S. cerevisiae genomic DNA, subcloned into the pRSET vector, and purified as described previously (13). Orp1 single and double point mutants were made using standard PCR-based mutagenesis. The peroxidase activity was measured following established methodology (13). The Yap1-cCRD protein was designed and purified as described previously.3 Briefly, Cys598 of Yap1 can then react with the Cys36-SOH intermediate to form a mixed disulfide-bonded complex between Orp1 and Yap1 and ultimately results in the inhibition of a nuclear export sequence in Yap1 (18, 28). This complex has been identified with mass spectrometry in vivo (12). Additionally, a C310A/C315A/C620A/C629T mutant form of Yap1 has been shown to form disulfide bonds in vivo in a manner similar to wild-type Yap1 (12). These data indicated to us that a truncated Yap1 protein with only Cys598 could serve as a useful tool in examining the interaction of Yap1 with H2O2-activated Orp1. After purification, all proteins were dialyzed into 25 mM Tris (pH 7.5), 100 mM NaCl, and 1 mM EDTA and stored at -80 °C.

Orp1 and Yap1-cCRD Interaction Assay—Prior to use, Orp1 and Yap1-cCRD proteins were reduced with 50 mM DTT for 2 h at 25 °C. After reduction, each protein was exchanged into 25 mM Tris buffer (pH 7.5), 100 mM NaCl, and 1 mM ETDA using a Hi-Trap desalting column (GE Healthcare) operating on an AKTA purifier fast protein liquid chromatograph. The protein concentrations of Yap1-cCRD and Orp1 proteins were determined using the BCA assay (Pierce). Yap1-cCRD and Orp1 were mixed together in equal molar ratios to a final concentration of 4 µM in a total reaction volume of 100 µl. As controls, reactions containing Yap1-cCRD or Orp1 alone were prepared. All reactions were initiated with the addition of 8 µM H2O2 and allowed to react for 1 min. Reactions were stopped with the addition of trichloroacetic acid and precipitated on ice for 20 min. Precipitated proteins were pelleted with centrifugation and resuspended in 100 mM Tris (pH 8.0), 1% SDS, and 20 mM iodoacetamide. Resuspended samples were incubated in the dark for 30 min at 25 °C, and 4x SDS-PAGE buffer was added. Reactions were incubated at 95 °C for 5 min, separated by nonreducing or reducing SDS-PAGE, and proteins visualized with Coomassie Brilliant Blue.

Detection of Cysteine Sulfenic Acid Modifications with NBD-Cl—NBD-Cl was used to detect the formation of Cys-SOH on purified proteins in response to H2O2 exposure (31). 100 mM stocks of NBD-Cl were prepared in Me2SO and stored in the dark at -20 °C. All experiments were performed aerobically. Each protein was reduced with 50 mM DTT for 2 h at 25 °C. After reduction, proteins were exchanged into 25 mM potassium phosphate (pH 6.0) and 1 mM ETDA using a Hi-Trap desalting column (GE Healthcare) operating on an AKTA purifier fast protein liquid chromatograph. Proteins were concentrated to a final concentration of 40 µM with an Amicon Ultra-4 (10-kDa cutoff). 200-µl oxidation reactions were initiated with the addition of H2O2 followed immediately with NBD-Cl, to a final concentration of 80 and 250 µM, respectively. Reactions were incubated for 1 h at 25 °C, and unreacted NBD-Cl was separated from protein with a 5-ml Hi-trap desalting column. NBD-Cl-labeled proteins were concentrated to 40 µM and analyzed with UV-visible spectroscopy on a Varian Cary 50 spectrophotometer.

pKa Determination of Orp1 Sulfhydryls with PDT-Bimane—The reaction of PDT-Bimane with cysteine forms pyridine-2-thione, which has a maximum absorption wavelength of 343 nm (32). This reaction has been used previously to determine the pKa value of cysteine residues in proteins (33). The reduced Orp1 proteins were diluted to 10 µM in sodium citrate or phosphate buffer ranging from pH 3.5 to 11.5. Reactions were started by addition of PDT-Bimane to a final concentration of 40 µM and rapidly mixed, and the absorbance at 343 nm was recorded over 120 min with a Varian Cary 50 Bio UV-visible spectrophotometer. The resulting 343 nm curves were fit to a single exponential (Y = Y0 + Ae-x/t1) function. The values for the inverse of the first-order rate constant (t1) were plotted against pH. The resulting curves were fit to the Henderson-Hasselbach equation, and the pKa value was determined.

Homology Model of Orp1—The three-dimensional model of Orp1 was calculated based on the x-ray crystal structures of human hGpx2 and hGpx5. The structures of each protein have yet to be published but are deposited in the Protein Data Bank codes 2HE3 and 2I3Y, respectively. The model was constructed using MODELLER 9 version 1 following the detailed steps described in the MODELLER advanced tutorial (34, 35). Briefly, the primary sequences of the Gpx2 and Gpx5 templates were initially aligned using the align multiple structure/sequence alignment module within MODELLER. The Orp1 sequence was then aligned with the aligned sequences of hGpx2 and hGpx5 using the multiple alignment module. Five models of Orp1 were calculated using the model module and were analyzed with the discrete optimized protein energy function. We selected the lowest energy structure for modeling purposes and used MacPyMOL for displaying the structure.

Analysis of Orp1 Mutations in Vivo—Yeast strains YPH499 (MATa ura3-52 lys2–801_amber ade2-101_ochre trp1-{Delta}63 his3-{Delta}200 leu2-{Delta}1) and YMJW22 described previously (YPH499:{Delta}orp1::KanR) were used and have been described previously (18). For Orp1 expression in S. cerevisiae, we used the ppp81 vector, which allows for stable integration of the plasmid DNA into the ADH1 promoter region (36). The pADH1-ORP1 overexpression constructs were constructed by amplifying the ORP1 coding region by PCR using oligonucleotides primers containing NotI (5') and BglII (3') sites. The PCR products were digested with NotI and BglII and ligated to ppp81 digested with the same restriction enzymes. The resulting plasmids were verified by DNA sequencing. The wild-type and mutant ORP1 expression strains were created by transforming PacI linearized pADH1-ORP1 into the YMJW22 strain. As a control we also constructed a vector-only strain by transforming PacI linearized ppp81 into the YMJW22 strain. The integration of the ORP1 gene into the ADH1 locus was confirmed by colony PCR analysis. For H2O2 sensitivity analysis, each strain was grown to mid-log phase (A600 = 0.7) in either YPD media or SC-URA minimal media. YPD plates were freshly prepared with 0, 0.5, 1.0, 1.5, 2.0, 2.5, and 3.0 mM H2O2 and were used immediately. Each strain was serially diluted, and 3 µl of each dilution was plated onto YPD plates. The plates were incubated at 30 °C for 2 days.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
The Formation of Orp1Cys36 and Yap1-cCRD Disulfide Bond Complex—The goal of this study was to elucidate the molecular mechanism by which Orp1 senses peroxides and ultimately reacts with Yap1. Therefore, we developed a gel shift assay for monitoring the Orp1-Yap1 protein interaction. A Yap1-cCRD expression construct was prepared, which included Yap1 amino acids Asn565 to Asn650 and where Cys620 and Cys629 were mutated to alanine and threonine, respectively. For this assay we used the Orp1Cys36 protein, which only contains the active site Cys36 and has Cys64 and Cys82 mutated to serine. Orp1Cys36 is incapable of forming a Cys36–Cys82 intramolecular disulfide bond but can still catalyze disulfide bond formation in Yap1 in response to H2O2 in vivo (12). Both Orp1Cys36 and Yap1-cCRD were purified to homogeneity, reduced with DTT, and exchanged into buffer containing EDTA. It was observed that the presence of EDTA minimized protein oxidation, thus allowing all experiments to be performed aerobically. These assays were also conducted anaerobically and showed similar results (data not shown). Yap1-cCRD and Orp1Cys36 were mixed in equal molar ratios, and H2O2 was added to the reactions. Reactions were stopped with trichloroacetic acid after 1 min, separated with reducing or nonreducing SDS-PAGE, and stained with Coomassie Brilliant Blue (Fig. 1). The gel shows that Orp1Cys36 alone treated with H2O2 forms a small amount of disulfide-bonded dimer. This has also been observed with Orp1 homologs from poplar trees (37). Yap1-cCRD appears unchanged by H2O2 treatment and migrates at the same apparent molecular weight. However, mixtures of Orp1Cys36 and Yap1-cCRD form a higher molecular weight complex upon H2O2 treatment. This complex migrates at the appropriate molecular weight of an Orp1Cys36 and Yap1-cCRD disulfide bond-linked complex. The band corresponding to the Orp1Cys36-Yap1-cCRD complex is not observed in the individual Orp1Cys36 or Yap1-cCRD reactions. When these reactions are run under reducing conditions the Orp1Cys36-Yap1-cCRD complex is not observed, further demonstrating that Orp1Cys36 and Yap1-cCRD are covalently linked via a disulfide bond. These experiments provide evidence that Yap1-cCRD can specifically react with Orp1 in an H2O2-inducible manner, and we hypothesized that an intermediate of the reaction involved Cys-SOH formation on Orp1.


Figure 1
View larger version (19K):
[in this window]
[in a new window]

  		FIGURE 1.
In vitro analysis of Orp1Cys36 and Yap1-cCRD disulfide-linked complex formation. Formation of the Orp1Cys36-Yap1-cCRD complex was performed aerobically at 25 °C (pH 7.5) with equimolar ratios of Orp1Cys36 and Yap1-cCRD and addition of H2O2. Each lane represents a reaction that was precipitated with trichloroacetic acid and separated with either reducing or nonreducing SDS-PAGE. Only when Orp1Cys36 and Yap1-cCRD are exposed to H2O2 is appreciable disulfide bond formation initiated. Under reducing conditions, the Orp1Cys36 and Yap1-cCRD disulfide-linked complex is not observed.

 
Identification of a Cys-SOH Intermediate on Cys36 of Orp1—To elucidate the molecular mechanism by which Orp1 senses H2O2, we wanted to determine whether the redox-active cysteines in Orp1 and Yap1-cCRD formed Cys-SOH in response to H2O2. We used the chemical NBD-Cl to probe Cys-SOH modification of specific cysteines on Orp1 and Yap1-cCRD. NBD-Cl has been used previously to monitor Cys-SOH modification on proteins (31). One limitation of this technique is that to obtain unambiguous results, it requires the use of proteins that contain only one cysteine residue. Therefore, we constructed and purified two forms of Orp1 that contained either Cys36 or Cys82 and Yap1-cCRD that contained only Cys598. We first reacted Orp1Cys36 with H2O2 and NBD-Cl. As controls we prepared samples of Orp1Cys36 in parallel that were untreated and treated with NBD-Cl alone. Unreacted NBD-Cl was removed from the reactions by gel filtration chromatography, and the recovered protein samples were concentrated prior to UV-visible spectrophotometry (Fig. 2A). For untreated Orp1Cys36, we did not observe any absorbance peaks between 300 and 500 nm. Orp1Cys36 samples that were treated with NBD-Cl showed a peak with maximal absorbance at 420 nm. This is the expected peak for the reaction of NBD-Cl with a free sulfhydryl group (R-S-NBD). Upon treatment with both H2O2 and NBD-Cl, Orp1Cys36 showed a new peak with maximal absorbance at 347 nm. This peak is indicative of formation of Cys-SOH on Cys36 (R-SO-NBD). Both the 347 and 420 nm peaks disappeared upon reduction with DTT, indicating that the NBD-Cl molecule was connected via Cys36 (data not shown). We repeated these sets of experiments with both Orp1Cys82 and Yap1-cCRD (Fig. 2, B and C). In both cases the spectra of NBD-Cl-treated samples looked the same with absorbance peak maxima at 420 nm, regardless of whether H2O2 was added or not. These data suggest that Orp1 reacts with H2O2 at Cys36 and forms a Cys-SOH intermediate, but that neither Cys82 of Orp1 nor Cys598 of Yap1 forms Cys-SOH in the presence of H2O2.


Figure 2
View larger version (12K):
[in this window]
[in a new window]

  		FIGURE 2.
NBD adducts of cysteine thiols and sulfenic acids. The reduced (R-SH) and oxidized (R-SOH) forms of the Orp1Cys36 give rise to distinctive spectra (A) with maximum absorption wavelength at 420 nm (R-S-NBD, dotted line) and 347 nm (R-S(O)-NBD, dashed line). The spectra of Orp1Cys36 itself is shown as a solid black line. Reacting to either Yap1-cCRD (B) or Orp1Cys82 (C) with H2O2 and NBD-Cl does not produce a 347 nm (R-S(O)-NBD, dotted line) absorption peak. For each of these proteins only the 420 nm peak (R-S-NBD, dotted line) is observed, indicating that they do not form Cys-SOH. a.u., absorbance units.

 


Figure 3
View larger version (41K):
[in this window]
[in a new window]

  		FIGURE 3.
Molecular model of the Orp1 protein. A, protein structure is shown as a ribbon diagram. The proposed active site residues Cys36, Gln70, and Trp125 are depicted by sticks with carbon, nitrogen, oxygen, and sulfur atoms colored green, red, blue, and orange, respectively. The side chain of the peroxidatic Cys82 is located ~10 Å below the Cys36-active site. B, Orp1-active site formed by evolutionarily conserved resides. On the hydrophobic left side of the pocket are Phe38 and Phe127 residues. On the polar right side of the pocket are Gln70, Trp125, and Asn126. The side chains of both Trp125 and Gln70 are within hydrogen bonding distance of the Cys36 sulfhydryl.

 
Molecular Modeling of Orp1—To better understand the molecular environment of Cys36 in the Orp1-active site, we constructed a homology model of the Orp1 protein. Previous studies have shown that proteins in the Orp1 family of peroxidases are thioredoxin-dependent and monomeric (12, 38). This differs from their human counterparts, which are glutathione-dependent, contain a seleno-cysteine in their active sites, and form tetramers in solution (39). We used the x-ray crystallographic structure of hGpx2 and hGpx5 as templates for constructing the Orp1 model. The overall fold of the lowest energy model of Orp1 is shown in Fig. 3A. The side chains of the Cys36-active site and Cys82 are displayed. Also shown are the side chains of the conserved Gln70 and Trp125 residues, which are located around Cys36 in the active site. These residues are also located in the active site of human seleno-cysteine glutathione peroxidases. In the Orp1 model, Cys82 is located on the surface of the protein and is partially solvent-exposed. It is ~10 Å away from the Cys36-active site and thus to form a disulfide bond with Cys36 and the helix, and loop containing Cys82 would have to be flexible. The Gln70 and Trp125 side chains are located within 3.5 Å of the Cys36 sulfhydryl and thus are strong candidates for modulating the reactivity of Cys36 toward peroxide. In bovine Gpx the catalytic triad of Cys52, Gln90, and Trp165 is thought to stabilize the selenol group (Se-OH) of the selenocysteine through hydrogen bonding with the imino group of the tryptophan residue and the amido group of the glutamine residue (40, 41). A detailed model of the Cys36-active site is shown in Fig. 3B. From this model it is apparent that Cys36 is located in a solvent-accessible active site pocket. The conserved Phe38 and Phe127 hydrophobic residues line one side of the active site, and conserved polar side chains that could form hydrogen bonds with Cys36 line the other side of the active site.


Figure 4
View larger version (20K):
[in this window]
[in a new window]

  		FIGURE 4.
Mutagenesis of the Orp1-active site. A, Orp1 peroxidase activity was measured using a NADPH-coupled assay. Wild-type Orp1 (solid line) activity was compared with the activity of the Orp1Q70A (dotted line) and Orp1W125A (dashed line) point mutants. B, UV-visible absorption spectra of Orp1Cys36 (solid line), Orp1Cys36/Q70A (dotted line), and Orp1Cys36/W125A (dashed line) after exposure to H2O2 and NBD-Cl. Neither Orp1Cys36/Q70A nor Orp1Cys36/W125A was able to form R-S(O)-NBD. C, reducing and nonreducing SDS-PAGE analysis of Orp1Cys36, Orp1Cys36/Q70A, and Orp1Cys36/W125A reacted with Yap1-cCRD with and without H2O2 treatment. Reactions were precipitated with trichloroacetic acid, and gels were stained with Coomassie Brilliant Blue. A.U., absorbance units.

 
Mutational Analysis of Orp1—To further examine the roles of Gln70 and Trp125 in the formation of Cys-SOH on Cys36 upon H2O2 exposure, we made four versions of the Orp1 protein. To measure the hydrogen peroxidase activity of Orp1, we mutated Gln70 and Trp125 to alanine. To measure the ability of Orp1 to form a H2O2-inducible disulfide-bonded complex with Yap1-cCRD and Cys-SOH on Cys36, we prepared Q70A and W125A mutants of Orp1 that only contained the Cys36. We monitored the hydrogen peroxidase enzymatic activity of Orp1, Orp1Q70A, and Orp1W125A using a previously established NADPH-linked assay (13). This assay showed that Orp1Q70A is no longer active as a peroxidase and that the Orp1W125A mutant retained ~10% of wild-type peroxidase activity (Fig. 4A). Next we examined the ability of Orp1Cys36/Q70A and Orp1Cys36/W125A to form Cys-SOH in response to H2O2. NBD-Cl modification experiments were conducted as described previously with similar controls. Wild-type Orp1 formed Cys-SOH as judged by the absorbance peak at 347 nm (Fig. 4B). Both Orp1Cys36/Q70A and Orp1Cys36/W125A reacted with NBD-Cl in the absence of H2O2 and displayed an absorbance peak at 420 nm. Upon addition of H2O2 and NBD-Cl, neither Orp1Cys36/Q70A nor Orp1Cys36/W125A displayed an absorbance peak at 347 nm. These data suggest that Orp1Cys36/Q70A and Orp1Cys36/W125A are incapable of forming Cys-SOH in response to H2O2. Finally, we assayed the ability of Yap1-cCRD to react with Orp1Cys36/Q70A and Orp1Cys36/W125A to form a disulfide-bonded complex. We individually mixed Orp1Cys36, Orp1Cys36/Q70A, or Orp1Cys36/W125A with Yap1-cCRD in equimolar ratios, added H2O2, and separated the reactions on reducing and nonreducing SDS-PAGE (Fig. 4C). Both Orp1Cys36/Q70A and Orp1Cys36/W125A were unable to form significant amounts of Orp1-Yap1-cCRD disulfide-bonded complex upon H2O2 addition. Taken together these results indicate that Gln70 and Trp125 influence the reactivity of Cys36. Without these residues, Orp1 cannot react with H2O2 to form a Cys-SOH intermediate and is not able to form mixed disulfide bonds with Yap1.

Analysis of the pKa of Orp1 Sulfhydryls—We hypothesized that the sulfhydryl pKa of Cys36 is lowered by its molecular environment. To quantify the pKa of Orp1Cys36 and Orp1Cys82, we used a thiol reactivity assay described previously by Brennan and co-workers (33) for measuring the pKa value of the active site cysteine in the OhrR transcription factor. Briefly, the chemical PDT-Bimane is reacted with a protein that contains a single cysteine residue over a range of pH values (32). The reaction is monitored at 343 nm as a function of time and fit to a first-order exponential function. We performed this analysis on the Orp1Cys36 protein over a range of pH values from 3.5 to 8.5 (Fig. 5A). These data show that at pH values of 3.5 and 4.0, the reaction of Orp1Cys36 with PDT-Bimane is slow and largely unchanged. This is what would be expected for a cysteine sulfhydryl that is protonated. Between the pH values of 4.5 and 6.0, the t1 for the reactions decreased ~90-fold. At pH values 7.0 and above, the reaction is extremely fast and complete within 2 min. The t1 values were normalized and plotted as function of pH (Fig. 5B). These points were fit to the Henderson-Hasselbach equation, and a pKa value of 5.1 was determined. This value is 3.2 pH units lower than the pKa of free cysteine and indicates that Cys36 would be in the thiolate form at physiological pH. We performed the similar PDT-Bimane reaction pKa analysis for Orp1Cys82, Orp1Cys36/Q70A, and Orp1Cys36/W125A (data not shown). The t1 values for the resulting first-order reaction curves were normalized and plotted with those for Orp1Cys36 (Fig. 5B). These data show that the pKa for Cys82 in Orp1 is 8.3, which is very close to the pKa of an isolated cysteine molecule. Remarkably, mutation of either Gln70 or Trp125 to alanine results in the reversion of the pKa of Cys36 to 8.3. These data strongly suggests that Gln70 and Trp125 stabilize Cys36 in its thiolate form.


Figure 5
View larger version (21K):
[in this window]
[in a new window]

  		FIGURE 5.
pKa determination of sulfhydryls with PDT-Bimane. A, reaction of Orp1Cys36 with PDT-Bimane was monitored at 343 nm at pH values ranging from 3.5 to 8.5. The increase at 343 nm results from the release of pyridyl-2-thione from PDT-Bimane. Each curve was fit to a single exponential function (solid black line), and the first-order rate constants were determined. B, inverse of the rate constants (t1) were normalized for the reactions of PDT-Bimane with Orp1Cys36, Orp1Cys36/Q70A, Orp1Cys36/W125A, and Orp1Cys82 and plotted as a function of pH. The results are fit to the Henderson-Hasselbalch equation. From these curve fits sulfhydryl pKa values of 5.1, 8.3, 8.3, and 8.3 were determined for the Orp1Cys36, Orp1Cys36/Q70A, Orp1Cys36/W125A and Orp1Cys82, respectively.

 
Analysis of Orp1 Mutations in Vivo—We sought to characterize the phenotypes of the Orp1Q70A and Orp1W125A S. cerevisiae mutants. To this end, we transformed an S. cerevisiae strain in which the ORP1 gene had already been deleted, with integrating plasmids containing the wild-type ORP1, ORP1Q70A, ORP1W125A, or ppp81 as a vector control. Correct integration of the ORP1 genes into the ADH1 promoter was verified with colony PCR analysis (data not shown). The resulting strains were assayed for their resistance to increasing concentrations of H2O2 peroxide. The parental wild-type YPH499 strain was used as a positive control. This strain showed colony forming ability up to 3.0 mM H2O2 (Fig. 6). The {Delta}orp1 strain showed increased sensitivity to H2O2. This strain was able to form colonies up to 2.0 mM H2O2. As expected, the ppp81-transformed strain showed similar H2O2 resistance levels as the {Delta}orp1 strain. Expression of Orp1 resulted in complementation of the {Delta}orp1 strain, and this strain had H2O2 resistance levels that were similar to that of the parental YPH499 strain. Expression of either Orp1Q70A or Orp1W125A resulted in strains that displayed H2O2 resistance phenotypes similar to the {Delta}orp1 strain.


Figure 6
View larger version (62K):
[in this window]
[in a new window]

  		FIGURE 6.
In vivo analysis of Orp1Q70A and Orp1W125A mutants. H2O2 resistance phenotypes of the parental S. cerevisiae strains YPH499, {Delta}orp1, ppp81 ADH1 integrating vector, ADH1 integrated Orp1, Orp1Q70A, and Orp1W125A are shown. Both YPH499 and ADH1 integrated Orp1 were able to form colonies out to 3 mM H2O2. ADH1 integrated Orp1Q70A and Orp1W125A displayed phenotypes similar to {Delta}orp1, indicating that they were nonfunctional in vivo.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The discovery that disulfide bond formation in the Yap1 transcription factor is regulated by a peroxidase-like enzyme has led to the realization that peroxidases and peroxiredoxin enzymes are not only involved in ROS scavenging but also function as peroxide sensors in vivo. The Orp1 protein is a member of a thiol peroxidase family of enzymes that is homologous to human glutathione peroxidases, yet it is functionally similar to the peroxiredoxin protein family (12). These classes of proteins have evolved to specifically react with and reduce peroxides, such as H2O2 and the less polar alkyl hydroperoxides. Additionally, as part of their catalytic mechanism, these enzymes form oxidized disulfide bonds either between cysteine pairs within the protein or with glutathione. It is thus not surprising that these classes of enzymes have evolved to use similar biochemical mechanisms to form mixed disulfide bonds with other proteins, thereby activating disulfide bond relay cascades and modulating the function of transcription factors such as Yap1 and Pap1 (12, 13, 17, 42).

The goal of this study was to investigate the molecular mechanism by which the Orp1 class of peroxiredoxin-like enzymes senses H2O2. We initially established a disulfide bond formation assay between Orp1 and a portion of the Yap1 protein that has been shown to react with Orp1 in vivo. A disulfide-bonded complex was observed between Orp1 and Yap1-cCRD immediately following treatment with H2O2. We showed that a mutant form of Orp1, in which only the active site Cys36 is present, forms a Cys-SOH intermediate upon H2O2 exposure using an NBD-Cl chemical modification assay. Cys-SOH intermediates have been shown to have extremely short half-lives and have proven difficult to identify in proteins. For this reason, the remaining nucleophilic cysteines need to be removed from the protein being studied. In the case of Orp1, Cys36-SOH can be attacked by Cys82 of Orp1 to form an intramolecular disulfide bond or by Cys598 of Yap1 to form a mixed disulfide-bonded complex. We also examined the ability of Cys82 of Orp1 or Cys598 of Yap1 to form Cys-SOH in response to H2O2. Our data showed that neither cysteine could form Cys-SOH as judged by NDB-Cl reactivity. These data provided support for the idea that Cys-SOH could be used as a reliable biochemical marker for Orp1 activity. It also suggests that H2O2-induced Cys-SOH formation may be a general characteristic of redoxactive cysteine residues.

In this study we provide a comprehensive analysis of the correlation between cysteine sulfhydryl pKa, peroxidase activity, H2O2-induced Cys-SOH formation, and Orp1-active site structure. It had been shown previously that the selenocysteine of the human glutathione peroxidase enzymes reacts with H2O2 to form a selenol intermediate prior to reaction with glutathione (40). In addition, the seleno-cysteine-active site is formed by a catalytic triad involving Gln83 and Trp157. Mutation of either of these amino acids was shown to decrease the enzymatic activity of hGpx and reduced the ability of iodoacetic acid to inhibit hGpx activity (41). Our homology model of Orp1 showed that the homologous Gln70 and Trp125 were part of an active site pocket formed around Cys36 (Fig. 3, A and B). One side of the pocket was entirely hydrophobic, lined with the conserved phenylalanine residues Phe38 and Phe127. These residues have van der Waals interactions with Cys36 and appear to effectively shield Cys36 from the bulk solvent. On the other side of the pocket are the side chains of Gln70, Trp125, and Asn126, which are all conserved residues containing side chains that can make hydrogen bonds with the sulfhydryl-active site. Our results strongly suggest that the role of Gln70 and Trp125 in Orp1 reactivity is to stabilize the thiolate form of Cys36. In the absence of H2O2, mutation of either residue abolishes the low pKa of Cys36. Although the Cys36-active site pocket is accessible to solvent, this pocket does not appear to be able to accommodate Cys36-SOH. Therefore, upon oxidation the Cys-SOH would protrude from the active site pocket into the solvent, making it more vulnerable to nucleophilic attack by another cysteine.

The global identification of redox-active cysteine residues in proteins has been the focus of many recent investigations (43–45). Gladyshev and co-workers (44) used a novel bioinformatics approach to identify redox-active cysteines in the proteomes of multiple organisms. Their approach identified many previously characterized oxidoreductase proteins, such as the glutathione peroxidase family, which contain a nucleophilic active site cysteine. Proteins that contain cysteines with abnormally low pKa values have historically been characterized by their increased reactivity with haloacetic acids. Use of haloacetic acid modification in conjunction with monitoring enzymatic activity is a well established approach for measuring cysteine pKa (46, 47). One example is Cys25 in the sulfhydryl protease papain that has apKa of ~4 (47). Our results and prior studies on nucleophilic active site cysteine suggest that one common biochemical feature of these active site cysteines is a lowered sulfhydryl pKa. In the case of Orp1, we determined the pKa of Cys36 to be ~3.2 pH units lower than that of free cysteine. It is this feature alone that contributes to the reactivity of Orp1 toward peroxides and enables it to form Cys-SOH and, eventually, disulfide bonds.

There appear to be multiple ways that enzymes and transcription factors have evolved lower pKa cysteines, thereby increasing their reactivity toward peroxides. In the case of the OxyR transcription factor, the reactivity of Cys199 appears to be modulated via an ion pair with Arg266. The x-ray crystal structure of OxyR shows that the side chain of Arg266 is buried in the hydrophobic core behind the Cys199-active site (48). Although the pKa of Cys199 has not been determined, it has been shown that Cys199 forms Cys-SOH in response to H2O2 (23). The active site composition of OxyR suggests that the positively charged and buried Arg266 side chain stabilizes Cys199 in the thiolate form. Two additional examples of redox-active cysteines that use ion pairing to lower their pKa can be seen in ArsC, an arsenate reductase found in E. coli, and in AhpC, a peroxiredoxin found in E. coli (49). In the case of ArsC, the pKa of Cys12 in its active site is 6.4 and is stabilized by the positive charge on His8. Mutation of His8 to glycine resulted in reversion of the Cys12 pKa back to that of free cysteine. Finally, an example of a low pKa cysteine that is neither part of a catalytic triad nor an ion pair can be found in the OhrR transcription factor from Bacillus subtilis (50). When reduced, OhrR can cooperatively bind its promoter and repress transcription (33). Upon reaction with organic hydroperoxides, OhrR is oxidized at Cys15 and forms a Cys-SOH. Using similar pKa determination techniques as those used in the present study, Brennan and co-workers (33) showed that OhrR has a pKa value of 5.2. The x-ray crystal structure of reduced OhrR shows that Cys15 sits at the end of an {alpha}-helix and is adjacent to two tyrosine residues. Brennan and co-workers (33) suggest that the positive macrodipole of the {alpha}-helix could explain the lower pKa value of Cys15, but biochemical studies to confirm this hypothesis have not been performed. What remains a major challenge in the field of protein redox regulation is the identification of proteins with low pKa cysteines. Identification of these cysteines will undoubtedly lead to the discovery of novel proteins that are regulated by thiol oxidation and reduction as well as other enzymes that utilize nucleophilic sulfhydryls as part of their catalytic mechanism.


   FOOTNOTES
 
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

1 To whom correspondence should be addressed: Dept. of Environmental Toxicology, University of California, Davis, One Shields Ave., Davis, CA 95616. Tel.: 530-754-2271; Fax: 530-752-3394; E-mail: mjwood{at}ucdavis.edu.

2 The abbreviations used are: ROS, reactive oxygen species; DTT, dithiothreitol; NBD-Cl, 7-chloro-4-nitrobenzo-2-oxa-1,3-diazole; PDT-Bimane, (2-pyridyl)-dithiobimane; Me2SO, dimethyl sulfoxide. Back

3 C. L. Takanishi, L. Ma, and M. J. Wood, submitted for publication. Back


   ACKNOWLEDGMENTS
 
We thank Heather Bolstad and Gisela Storz for critical reading of this manuscript.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Sies, H. (1985) in Oxidative Stress (Sies, H., ed) pp. 1-8, Academic Press, London
  2. Fraga, C. G., Shigenaga, M. K., Park, J. W., Degan, P., and Ames, B. N. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 4533-4537[Abstract/Free Full Text]
  3. Cerutti, P., and Trump, B. (1991) Cancer Cells 3, 1-7[Medline] [Order article via Infotrieve]
  4. Klaunig, J. E., Xu, Y., Isenberg, J. S., Bachowski, S., Kolaja, K. L., Jiang, J., Stevenson, D. E., and Walborg, E. F. (1998) Environ. Health Perspect. 106, 289-295[CrossRef][Medline] [Order article via Infotrieve]
  5. Trush, M. A., and Kensler, T. W. (1991) Free Radic. Biol. Med. 10, 201-209[CrossRef][Medline] [Order article via Infotrieve]
  6. Hei, T. K., Liu, S. X., and Waldren, C. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 8103-8107[Abstract/Free Full Text]
  7. Liu, S. X., Athar, M., Lippai, I., Waldren, C., and Hei, T. K. (2001) Proc. Natl. Acad. Sci. U. S. A. 98, 1643-1648[Abstract/Free Full Text]
  8. Kasai, H. (1997) Mutat. Res. 387, 147-163[CrossRef][Medline] [Order article via Infotrieve]
  9. Oliver, C. N., Starke-Reed, P. E., Stadtman, E. R., Liu, G. J., Carney, J. M., and Floyd, R. A. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 5144-5147[Abstract/Free Full Text]
  10. Reznick, A. Z., and Packer, L. (1994) Methods Enzymol. 233, 357-363[Medline] [Order article via Infotrieve]
  11. Vile, G. F., and Tyrrell, R. M. (1995) Free Radic. Biol. Med. 18, 721-730[CrossRef][Medline] [Order article via Infotrieve]
  12. Delaunay, A., Pflieger, D., Barrault, M. B., Vinh, J., and Toledano, M. B. (2002) Cell 111, 471-481[CrossRef][Medline] [Order article via Infotrieve]
  13. Mason, J. T., Kim, S. K., Knaff, D. B., and Wood, M. J. (2006) Biochemistry 45, 13409-13417[CrossRef][Medline] [Order article via Infotrieve]
  14. Ushio-Fukai, M., and Alexander, R. W. (2004) Mol. Cell. Biochem. 264, 85-97[CrossRef][Medline] [Order article via Infotrieve]
  15. Van Der Wijk, T., Blanchetot, C., Overvoorde, J., and Den Hertog, J. (2003) J. Biol. Chem. 278, 13968-13974[Abstract/Free Full Text]
  16. Veal, E. A., Findlay, V. J., Day, A. M., Bozonet, S. M., Evans, J. M., Quinn, J., and Morgan, B. A. (2004) Mol. Cell 15, 129-139[CrossRef][Medline] [Order article via Infotrieve]
  17. Vivancos, A. P., Castillo, E. A., Biteau, B., Nicot, C., Ayte, J., Toledano, M. B., and Hidalgo, E. (2005) Proc. Natl. Acad. Sci. U. S. A. 102, 8875-8880[Abstract/Free Full Text]
  18. Wood, M. J., Storz, G., and Tjandra, N. (2004) Nature 430, 917-921[CrossRef][Medline] [Order article via Infotrieve]
  19. Toledano, M. B., Delaunay, A., Monceau, L., and Tacnet, F. (2004) Trends Biochem. Sci. 29, 351-357[CrossRef][Medline] [Order article via Infotrieve]
  20. McEligot, A. J., Yang, S., and Meyskens, F. L., Jr. (2005) Annu. Rev. Nutr. 25, 261-295[CrossRef][Medline] [Order article via Infotrieve]
  21. Paget, M. S., and Buttner, M. J. (2003) Annu. Rev. Genet. 37, 91-121[CrossRef][Medline] [Order article via Infotrieve]
  22. Zheng, M., Aslund, F., and Storz, G. (1998) Science 279, 1718-1721[Abstract/Free Full Text]
  23. Lee, C., Lee, S. M., Mukhopadhyay, P., Kim, S. J., Lee, S. C., Ahn, W. S., Yu, M. H., Storz, G., and Ryu, S. E. (2004) Nat. Struct. Mol. Biol. 11, 1179-1185[CrossRef][Medline] [Order article via Infotrieve]
  24. Jamieson, D. J. (1998) Yeast 14, 1511-1527[CrossRef][Medline] [Order article via Infotrieve]
  25. Moye-Rowley, W. S., Harshman, K. D., and Parker, C. S. (1989) Genes Dev. 3, 283-292[Abstract/Free Full Text]
  26. Wu, A. L., and Moye-Rowley, W. S. (1994) Mol. Cell. Biol. 14, 5832-5839[Abstract/Free Full Text]
  27. Wemmie, J. A., Wu, A. L., Harshman, K. D., Parker, C. S., and Moye-Rowley, W. S. (1994) J. Biol. Chem. 269, 14690-14697[Abstract/Free Full Text]
  28. Delaunay, A., Isnard, A. D., and Toledano, M. B. (2000) EMBO J. 19, 5157-5166[CrossRef][Medline] [Order article via Infotrieve]
  29. Gasch, A. P., Spellman, P. T., Kao, C. M., Carmel-Harel, O., Eisen, M. B., Storz, G., Botstein, D., and Brown, P. O. (2000) Mol. Biol. Cell 11, 4241-4257[Abstract/Free Full Text]
  30. Maiorino, M., Ursini, F., Bosello, V., Toppo, S., Tosatto, S. C., Mauri, P., Becker, K., Roveri, A., Bulato, C., Benazzi, L., De Palma, A., and Flohe, L. (2007) J. Mol. Biol. 365, 1033-1046[CrossRef][Medline] [Order article via Infotrieve]
  31. Ellis, H. R., and Poole, L. B. (1997) Biochemistry 36, 15013-15018[CrossRef][Medline] [Order article via Infotrieve]
  32. Mansoor, S. E., and Farrens, D. L. (2004) Biochemistry 43, 9426-9438[CrossRef][Medline] [Order article via Infotrieve]
  33. Hong, M., Fuangthong, M., Helmann, J. D., and Brennan, R. G. (2005) Mol. Cell 20, 131-141[CrossRef][Medline] [Order article via Infotrieve]
  34. Eswar, N., John, B., Mirkovic, N., Fiser, A., Ilyin, V. A., Pieper, U., Stuart, A. C., Marti-Renom, M. A., Madhusudhan, M. S., Yerkovich, B., and Sali, A. (2003) Nucleic Acids Res. 31, 3375-3380[Abstract/Free Full Text]
  35. Fiser, A., Do, R. K., and Sali, A. (2000) Protein Sci. 9, 1753-1773[Abstract]
  36. Easlon, E., Tsang, F., Dilova, I., Wang, C., Lu, S. P., Skinner, C., and Lin, S. J. (2007) J. Biol. Chem. 272, 6161-6171
  37. Navrot, N., Collin, V., Gualberto, J., Gelhaye, E., Hirasawa, M., Rey, P., Knaff, D. B., Issakidis, E., Jacquot, J. P., and Rouhier, N. (2006) Plant Physiol. 142, 1364-1379[Abstract/Free Full Text]
  38. Avery, A. M., Willetts, S. A., and Avery, S. V. (2004) J. Biol. Chem. 279, 46652-46658[Abstract/Free Full Text]
  39. Flohe, L. (1989) in Glutathione (Dolphin, D., Avramovic, O., and Poulson, R., eds) pp. 643-731, John Wiley & Sons, Inc., New York
  40. Epp, O., Ladenstein, R., and Wendel, A. (1983) Eur. J. Biochem. 133, 51-69[Medline] [Order article via Infotrieve]
  41. Maiorino, M., Aumann, K. D., Brigelius-Flohe, R., Doria, D., van den Heuvel, J., McCarthy, J., Roveri, A., Ursini, F., and Flohe, L. (1995) Biol. Chem. Hoppe-Seyler 376, 651-660[Medline] [Order article via Infotrieve]
  42. Okazaki, S., Naganuma, A., and Kuge, S. (2005) Antioxid. Redox. Signal. 7, 327-334[CrossRef][Medline] [Order article via Infotrieve]
  43. Eaton, P. (2006) Free Radic. Biol. Med. 40, 1889-1899[CrossRef][Medline] [Order article via Infotrieve]
  44. Fomenko, D. E., Xing, W., Adair, B. M., Thomas, D. J., and Gladyshev, V. N. (2007) Science 315, 387-389[Abstract/Free Full Text]
  45. Hampton, M. B., Baty, J. W., and Winterbourn, C. C. (2006) in Redox Proteomics (Dalle-Donne, I., Scaloni, A., and Butterfield, D. A., eds) pp. 253-265, John Wiley & Sons, Inc., Hoboken, NJ
  46. Oesterhelt, D., Bauer, H., Kresze, G. B., Steber, L., and Lynen, F. (1977) Eur. J. Biochem. 79, 173-180[Medline] [Order article via Infotrieve]
  47. Sluyterman, L. A. (1967) Biochim. Biophys. Acta 139, 439-449[Medline] [Order article via Infotrieve]
  48. Choi, H., Kim, S., Mukhopadhyay, P., Cho, S., Woo, J., Storz, G., and Ryu, S. (2001) Cell 105, 103-113[CrossRef][Medline] [Order article via Infotrieve]
  49. Gladysheva, T., Liu, J., and Rosen, B. P. (1996) J. Biol. Chem. 271, 33256-33260[Abstract/Free Full Text]
  50. Fuangthong, M., and Helmann, J. D. (2002) Proc. Natl. Acad. Sci. U. S. A. 99, 6690-6695[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?



This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
282/43/31429    most recent
M705953200v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Ma, L.-H.
Right arrow Articles by Wood, M. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Ma, L.-H.
Right arrow Articles by Wood, M. J.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg