Oxidant-specific Folding of Yap1p Regulates Both Transcriptional Activation and Nuclear Localization
2005; Elsevier BV; Volume: 280; Issue: 49 Linguagem: Inglês
10.1074/jbc.m504716200
ISSN1083-351X
AutoresKailash Gulshan, Sherry A. Rovinsky, Sean T. Coleman, W. Scott Moye‐Rowley,
Tópico(s)Endoplasmic Reticulum Stress and Disease
ResumoThe yeast transcriptional regulator Yap1p is a key determinant in oxidative stress resistance. This protein is found in the cytoplasm under non-stressed conditions but rapidly accumulates in the nucleus following oxidant exposure. There it activates transcription of genes encoding antioxidants that return the redox balance of the cell to an acceptable range. Yap1p localization to the nucleus requires the oxidant-specific formation of disulfide bonds in the N-terminal cysteine-rich domain (N-CRD) and/or the C-terminal cysteine-rich domain (C-CRD). H2O2 exposure triggers the formation of two interdomain disulfide bonds between the N-and C-CRDs. This dually disulfide-bonded structure has been argued to mask the nuclear export signal in the C-CRD that would otherwise prevent Yap1p nuclear accumulation. The C-CRD is required for wild-type H2O2 tolerance but dispensable for resistance to diamide. The Saccharomyces cerevisiae TRX2 gene, encoding a thioredoxin protein, cannot be induced by H2O2 in the presence of various mutant forms of Yap1p lacking the normally functioning C-CRD. In this work, we demonstrate that the proper folding of Yap1p in the presence of H2O2 is required for recruitment of the mediator component Rox3p to the TRX2 promoter in addition to the nuclear accumulation of Yap1p during stress by this oxidant. These data demonstrate that the dually disulfide-bonded Yap1p N- and C-CRDs form a bifunctional protein domain controlling both nuclear localization and transcriptional activation. The yeast transcriptional regulator Yap1p is a key determinant in oxidative stress resistance. This protein is found in the cytoplasm under non-stressed conditions but rapidly accumulates in the nucleus following oxidant exposure. There it activates transcription of genes encoding antioxidants that return the redox balance of the cell to an acceptable range. Yap1p localization to the nucleus requires the oxidant-specific formation of disulfide bonds in the N-terminal cysteine-rich domain (N-CRD) and/or the C-terminal cysteine-rich domain (C-CRD). H2O2 exposure triggers the formation of two interdomain disulfide bonds between the N-and C-CRDs. This dually disulfide-bonded structure has been argued to mask the nuclear export signal in the C-CRD that would otherwise prevent Yap1p nuclear accumulation. The C-CRD is required for wild-type H2O2 tolerance but dispensable for resistance to diamide. The Saccharomyces cerevisiae TRX2 gene, encoding a thioredoxin protein, cannot be induced by H2O2 in the presence of various mutant forms of Yap1p lacking the normally functioning C-CRD. In this work, we demonstrate that the proper folding of Yap1p in the presence of H2O2 is required for recruitment of the mediator component Rox3p to the TRX2 promoter in addition to the nuclear accumulation of Yap1p during stress by this oxidant. These data demonstrate that the dually disulfide-bonded Yap1p N- and C-CRDs form a bifunctional protein domain controlling both nuclear localization and transcriptional activation. Cells must detoxify reactive oxygen species that are formed during aerobic metabolism to maintain viability (1Cadenas E. Annu. Rev. Biochem. 1989; 58: 79-110Crossref PubMed Scopus (856) Google Scholar, 2Imlay J.A. Annu. Rev. Microbiol. 2003; 57: 395-418Crossref PubMed Scopus (1592) Google Scholar). Sensing the resulting changes in the oxidative environment is an essential ability enabling proper regulatory adjustment of the multiple systems that control the intracellular redox potential. Utilization of the uniquely oxidant-sensitive cysteine residue in proteins has emerged as a pervasive theme in proteins that have been designed to monitor the redox milieu (3Paget M.S. Buttner M.J. Annu. Rev. Genet. 2003; 37: 91-121Crossref PubMed Scopus (251) Google Scholar). The Saccharomyces cerevisiae transcriptional regulatory protein Yap1p has provided important insight into the mechanism of oxidative stress sensing in eukaryotic cells via changes in the redox status of its cysteine residues (see Refs. 3Paget M.S. Buttner M.J. Annu. Rev. Genet. 2003; 37: 91-121Crossref PubMed Scopus (251) Google Scholar, 4Moye-Rowley W.S. Prog. Nucleic Acid Res. Mol. Biol. 2003; 73: 251-279Crossref PubMed Scopus (82) Google Scholar, 5Linke K. Jakob U. Antioxid. Redox Signal. 2003; 5: 425-434Crossref PubMed Scopus (99) Google Scholar, 6Rodrigues-Pousada C.A. Nevitt T. Menezes R. Azevedo D. Pereira J. Amaral C. FEBS Lett. 2004; 567: 80-85Crossref PubMed Scopus (95) Google Scholar for recent reviews). Yap1p is a positive regulator of gene expression and recognizes a binding site called a Yap1p-response element (YRE) 4The abbreviations used are: YREYap1p-response elementN-CRDN-terminal cysteine-rich domainC-CRDC-terminal cysteine-rich domainUASupstream activating sequenceChIPchromatin immunoprecipitationTAPtandem affinity purificationPIPES1,4-piperazinediethanesulfonic acidmTATAmutant TATA.4The abbreviations used are: YREYap1p-response elementN-CRDN-terminal cysteine-rich domainC-CRDC-terminal cysteine-rich domainUASupstream activating sequenceChIPchromatin immunoprecipitationTAPtandem affinity purificationPIPES1,4-piperazinediethanesulfonic acidmTATAmutant TATA. that is located in the promoter of target genes (7Kuge S. Jones N. EMBO J. 1994; 13: 655-664Crossref PubMed Scopus (383) Google Scholar, 8Wu A.-L. Moye-Rowley W.S. Mol. Cell. Biol. 1994; 14: 5832-5839Crossref PubMed Google Scholar, 9Fernandes L. Rorigues-Pousada C. Struhl K. Mol. Cell. Biol. 1997; 17: 6982-6993Crossref PubMed Scopus (258) Google Scholar). In the absence of oxidative challenge, Yap1p resides in a cytoplasmic location (10Kuge S. Jones N. Nomoto A. EMBO J. 1997; 16: 1710-1720Crossref PubMed Scopus (344) Google Scholar). However, upon stress by addition of oxidants like diamide and H2O2, nuclear export of Yap1p is inhibited (11Yan C. Lee L.H. Davis L.I. EMBO J. 1998; 17: 7416-7429Crossref PubMed Scopus (203) Google Scholar, 12Kuge S. Toda T. Iizuka N. Nomoto A. Genes Cells. 1998; 3: 521-532Crossref PubMed Scopus (135) Google Scholar), and the factor rapidly accumulates in the nucleus, where it induces the expression of genes that often encode antioxidant proteins (10Kuge S. Jones N. Nomoto A. EMBO J. 1997; 16: 1710-1720Crossref PubMed Scopus (344) Google Scholar, 13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar). This oxidant-regulated nuclear localization has been the focus of much research, and important details have emerged illuminating the molecular mechanism. Yap1p contains two clusters of cysteine residues called cysteine-rich domains located at the N terminus (N-CRD) and the C terminus (C-CRD). These N- and C-CRDs provide all the cysteine residues found in the Yap1p sequence, and control of their redox status is crucial for control of Yap1p activity by oxidative stress. Previous work established that deletion of the C-CRD leads to Yap1p being constitutively located in the nucleus with transcription of an artificial Yap1p-responsive reporter plasmid and diamide tolerance being constitutively elevated (10Kuge S. Jones N. Nomoto A. EMBO J. 1997; 16: 1710-1720Crossref PubMed Scopus (344) Google Scholar, 14Wemmie J.A. Steggerda S.M. Moye-Rowley W.S. J. Biol. Chem. 1997; 272: 7908-7914Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar). Interestingly, although elimination of the C-CRD leads to diamide hyper-resistance, H2O2 tolerance is lowered relative to wild-type Yap1p (14Wemmie J.A. Steggerda S.M. Moye-Rowley W.S. J. Biol. Chem. 1997; 272: 7908-7914Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar). Yap1p-response element N-terminal cysteine-rich domain C-terminal cysteine-rich domain upstream activating sequence chromatin immunoprecipitation tandem affinity purification 1,4-piperazinediethanesulfonic acid mutant TATA. Yap1p-response element N-terminal cysteine-rich domain C-terminal cysteine-rich domain upstream activating sequence chromatin immunoprecipitation tandem affinity purification 1,4-piperazinediethanesulfonic acid mutant TATA. Important characterization of this oxidant-selective behavior of Yap1p was provided by the biochemical analysis of Yap1p after diamide and H2O2 exposure. Diamide challenge produces disulfide bonds that appear to form in either the N- or C-CRD (15Kuge S. Arita M. Murayama A. Maeta K. Izawa S. Inoue Y. Nomoto A. Mol. Cell. Biol. 2001; 21: 6139-6150Crossref PubMed Scopus (202) Google Scholar), whereas H2O2 stress induces the formation of a disulfide bond between the N- and C-CRDs (16Delaunay A. Isnard A.D. Toledano M.B. EMBO J. 2000; 19: 5157-5166Crossref PubMed Scopus (414) Google Scholar). Mutational analysis of cysteine residues in the N- and C-CRDs indicated that Cys303 and Cys598 are required for both normal H2O2 resistance and disulfide bond formation, but these same residues are dispensable for diamide tolerance (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar, 16Delaunay A. Isnard A.D. Toledano M.B. EMBO J. 2000; 19: 5157-5166Crossref PubMed Scopus (414) Google Scholar). More recent experiments have demonstrated that two disulfide bonds are formed between residues in the N- and C-CRDs (17Wood M.J. Storz G. Tjandra N. Nature. 2004; 430: 917-921Crossref PubMed Scopus (134) Google Scholar) in response to H2O2 but not diamide. The oxidant-specific defects of Yap1p mutants have been linked to specific defects in the program of Yap1p-dependent gene activation (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar, 16Delaunay A. Isnard A.D. Toledano M.B. EMBO J. 2000; 19: 5157-5166Crossref PubMed Scopus (414) Google Scholar). The thioredoxin-encoding TRX2 gene is a Yap1p regulatory target that is required for normal H2O2 resistance (7Kuge S. Jones N. EMBO J. 1994; 13: 655-664Crossref PubMed Scopus (383) Google Scholar). Yap1p mutants that cannot form the interdomain disulfide bond between the N- and C-CRDs are unable to elevate TRX2 expression in the presence of H2O2 (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar, 16Delaunay A. Isnard A.D. Toledano M.B. EMBO J. 2000; 19: 5157-5166Crossref PubMed Scopus (414) Google Scholar). Experiments using fragments of the TRX2 promoter placed upstream of a CYC1-lacZ fusion gene indicated that TRX2 promoter sequences from positions -141 to -61 prevent overexpression of β-galactosidase activity in response to the presence of a constitutively nuclear Yap1p mutant (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar). We have determined that the element blocking high-level CYC1-lacZ activation in this TRX2-CYC1-lacZ reporter corresponds to the TRX2 TATA box. Chromatin immunoprecipitation studies have indicated that the mediator component Rox3p is preferentially recruited to the TRX2 promoter during H2O2 stress. Yap1p mutants unable to form the N- and C-CRD interdomain disulfide bonds fail either to recruit Rox3p to TRX2 or to activate expression of this gene in the face of H2O2 challenge. This problem is seen even in a C-CRD mutant that is constitutively located in the nucleus. Together, these data argue that formation of properly disulfide-bonded Yap1p during H2O2 stress is required to generate a new bifunctional protein domain that allows Yap1p to accumulate in the nucleus as well as to recruit Rox3p to TRX2. Yeast Strains and Media—The yeast strains used in this study were as follows: SEY6210 (MATα leu2-3,112 ura3-52 his3-Δ200 trp1-Δ901 lys2-801 suc2-Δ9 Mel-), SM13 (MATα leu2-3,112 ura3-52 his3-Δ200 trp1-Δ901 lys2-801 suc2-Δ9 Mel- yap1-Δ2::hisG), and YSC21 (MATα leu2-3,112 ura3-52 his3-Δ200 trp1-Δ901 lys2-801 suc2-Δ9 Mel- trx2Δ::LEU2). YSC21 was generated by PCR amplification of the trx2Δ::LEU2 allele from EMY62 (provided by Dr. Eric Muller, University of Washington) (18Muller E.G. J. Biol. Chem. 1991; 266: 9194-9202Abstract Full Text PDF PubMed Google Scholar) and transformation of yeast strain SEY6210 with the resulting product. Leu+ transformants were recovered and purified. Genomic DNA was recovered from Leu+ cells, and the genome was analyzed for proper disruption of TRX2 by Southern blotting. The S. cerevisiae cells were grown in rich medium (YPD, 2% yeast extract/1% peptone/2% dextrose), minimal medium, or complete minimal medium supplemented with amino acids, adenine, and uracil as described (19Sherman F. Fink G. Hicks J. Methods in Yeast Genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1979Google Scholar). Transformation was performed using the LiOAc technique (20Ito H. Fukuda Y. Murata K. Kimura A. J. Bacteriol. 1983; 153: 163-168Crossref PubMed Google Scholar). Assays for β-galactosidase activity were carried out on permeabilized cells as described (21Guarente L. Methods Enzymol. 1983; 101: 181-191Crossref PubMed Scopus (871) Google Scholar). H2O2 resistance assays were carried out by spot test (22Wu A.-L. Wemmie J.A. Edgington N.P. Goebl M. Guevara J.L. Moye-Rowley W.S. J. Biol. Chem. 1993; 268: 18850-18858Abstract Full Text PDF PubMed Google Scholar). Plasmids—Low-copy-number URA3-containing plasmids carrying the wild-type or C629A YAP1 gene have been described (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar, 23Wemmie J.A. Wu A.-L. Harshman K.D. Parker C.S. Moye-Rowley W.S. J. Biol. Chem. 1994; 269: 14690-14697Abstract Full Text PDF PubMed Google Scholar). LEU2 variants of wild-type and C629A YAP1 were generated by moving a SacI/HindIII fragment from pSM58wt or pC629A into SacI/HindIII-cut pRS315 (pSMS51 and pRS315-C629A respectively). The TRX2- (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar) and GSH1-lacZ (8Wu A.-L. Moye-Rowley W.S. Mol. Cell. Biol. 1994; 14: 5832-5839Crossref PubMed Google Scholar) reporter plasmids have been described previously. A low-copy-number plasmid (pSC-T1) bearing the wild-type TRX2 coding region containing a BamHI site introduced after the translation start site was generated by PCR. Wild-type or mutant YRE forms of the TRX2 promoter were generated by PCR and fused to the TRX2 open reading frame (pSC-T1) or lacZ reporter plasmids (pSEYC102) (24Emr S.D. Vassarotti A. Garret J. Geller B.C. Takeda M. Douglas M.G. J. Cell Biol. 1986; 102: 523-533Crossref PubMed Scopus (139) Google Scholar). The primer used to alter the -218 YRE (GTT TAT ACT Cga gGT AAA GGA TGC TCC) introduced an XhoI site to facilitate identification. (The nucleotides changed are indicated by lowercase letters, and the restriction enzyme site introduced is underlined) The primer used to introduce changes into the -181 YRE (CTG AAC GCG tcT Aga AAG AAA AGA GCC) incorporated an XbaI site to ease identification. The mutant promoters were moved into the context of pSC-T1 or pSEYC102 as EcoRI/BamHI fragments. The 3′-deletion series was created by PCR amplification of a portion of the TRX2 promoter and were then placed upstream of the CYC1 promoter lacking its upstream activating sequence (UAS) and fused to lacZ. The forward primer for all plasmids in this series anneals at 237 bp upstream of the translation start site. Its sequence is GCG GAT CCA TGT GTA ATT GTT TAT ACT C. For the -120 deletion mutant, the reverse primer was CCG AGA TCT ATG GCT TTC TTA TAT ACT GAT. The -104 deletion mutant was created with the reverse primer CCG AGA TCT CTT TTC ATC CCC CGA ATG GCT T. For the -90 deletion mutant, the reverse primer was CCG AGA TCT TTA TTC TCT TGT CAG CTT TTC. These fragments were cloned into pCR2.1-TOPO (Invitrogen) and digested with EcoRI and BglII, and the TRX2 segments were inserted into EcoRI/BamHI-cut p314-CLZ as described (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar). The -155 and -61 deletion mutants have been described (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar). A 240-bp fragment of the TRX2 promoter containing a mutant TATA box (pSAR28) was constructed by site-directed mutagenesis and cloned into the pCR2.1-TOPO cloning vector. The primer used to mutate the site was GAA AGA GGG ATA TCA GtC GAC AAG AAA GCC ATC GG to create pSAR28, which was then cleaved with EcoRI and BamHI; and the 240-bp fragment was moved into the pSEYC102 plasmid to create a lacZ fusion (pSAR29). A second 240-bp fragment was generated that would replace the TRX2 TATA box with the TRP5 TATA box. This fragment was ligated into pSEYC102 to create a lacZ fusion (pSAR31). The primer used to produce this mutant was GAA AGA GGG ATA TCA GTA TAA GAA AAA GCC ATT CGG. All PCR products were sequenced to ensure that no errors had occurred during amplification. DNase I Footprinting—PCR products of the TRX2 promoter containing either wild-type or mutant YREs were amplified using primers -296-TRX2-For (GAT GGA TCC AAG ATC AGC ATA ACT TG) and TRX2-BglII-Rev (CCG AGA TCT TAT TGA TGT GTT ATT TAA AG) and cloned into pCR2.1-TOPO. These plasmids were digested with BamHI, treated with calf intestinal alkaline phosphatase, phosphorylated with T4 polynucleotide kinase and [γ-32P]ATP, and finally cleaved with EcoRI. The appropriate labeled fragments were isolated from an agarose gel using a Qiagen mini-elute column. DNase I protection was performed using bacterially expressed Yap1p as reported previously (25Moye-Rowley W.S. Harshman K.D. Parker C.S. Genes Dev. 1989; 3: 283-292Crossref PubMed Scopus (240) Google Scholar). Electrophoretic Mobility Shift Assay—For the TATA box-binding protein Tbp1p, a 237-bp segment of the wild-type TRX2 promoter was amplified using primers -236-TRX2-For (TGCGGATTCATGTGTAATTGTTTATAC) and TRX2-BglII-Rev. The resulting PCR fragment was cloned into pCR2.1-TOPO. A 5′-end-labeled fragment was prepared as described above for the DNase I probes. The labeled fragment was isolated and incubated with bacterially produced Tbp1p (provided by Dr. Anthony Weil, Vanderbilt University). In some cases, unlabeled competitor DNAs were included in the binding reaction to verify binding specificity. Electrophoretic mobility shift assays were performed basically as described (26Kang J.J. Auble D.T. Ranish J.A. Hahn S. Mol. Cell. Biol. 1995; 15: 1234-1243Crossref PubMed Google Scholar). Binding was carried out in 4% glycerol, 20 mm Tris-HCl (pH 8.0), 60 mm KCl, 5 mm MgCl2, 100 μg/ml bovine serum albumin, 1 mm dithiothreitol, and 20 μg/ml poly(dI-dC) for 45 min at 30 °C. DNA and protein·DNA complexes were resolved by electrophoresis on a 4% native acrylamide gel at 4 °C using 1× buffer containing 45 mm Tris (pH 8.0), 45 mm boric acid, and 1 mm EDTA. The gel was dried and visualized by autoradiography. Primer Extension Mapping of the Transcription Start Site—RNA was isolated from yeast strains grown in YPD medium. Primer CAC TGT CGT ATT CAG AAG CG was first end-labeled with 32P and then mixed with 50 μg of total RNA. This mixture was ethanol-precipitated, and nucleic acids were brought up in 20 μl of annealing buffer (250 mm KCl, 10 mm Tris-Cl (pH 8.2), and 1 mm EDTA (pH 8.0)) and heated to 65 °C for 45 min. This mixture was cooled to and allowed to anneal at 37 °C for 6 h. Five units of avian myeloblastosis virus reverse transcriptase, 5 μgof actinomycin D, 50 mm Tris-HCl (pH 8.3), 50 mm KCl, 10 mm MgCl2,10 mm dithiothreitol, 0.5 mm spermidine, 5 mm each dNTP, and 40 units of RNasin were mixed and brought to a final volume of 100 μl. Reverse transcription took place at 37 °C for 1 h. The DNA·RNA complexes were treated with 30 μg of RNase A for 30 min at 37 °C. The DNA was then precipitated, resuspended in stop buffer (95% formamide, 20 mm EDTA, 0.05% bromphenol blue, and 0.05% xylene cyanol), heated to 90 °C, chilled briefly, and then run at 20 watts on a 6% denaturing acrylamide gel. Sequencing reactions were performed with the Sequenase DNA sequencing kit (U. S. Biochemical Corp.) according to the manufacturer's instructions. Chromatin Immunoprecipitation—Cells were grown in liquid medium until an A600 nm of 0.8–1.0 was reached. Formaldehyde was directly added to the culture to a final concentration of 2% to form cross-linked protein·DNA complexes. Cells were incubated at room temperature for 15 min with occasional swirling. Formaldehyde cross-links were quenched by addition of 2.5 m glycine to a final concentration of 250 mm. Cells were then lysed in chromatin immunoprecipitation (ChIP) lysis buffer (50 mm HEPES (pH 7.5), 140 mm NaCl, 1% Triton X-100, 0.1% sodium deoxycholate, and protease inhibitors), and the chromatin was sheared by sonication (Fisher Model 550 sonic dismembrator). The sheared chromatin was then immunoprecipitated using anti-tandem affinity purification (TAP) antibody (Open Biosystems) and protein A-agarose beads (Santa Cruz Biotechnology, Inc.). The beads were then washed with ChIP wash buffer (10 mm Tris (pH 8.0), 250 mm LiCl, 0.5% Nonidet P-40, 0.5% sodium deoxycholate, and 1 mm EDTA), and precipitates were eluted. The eluted precipitates were incubated overnight at 65 °C to reverse cross-links. DNA was then precipitated and subjected to PCR analysis. Co-immunoprecipitation—All immunoprecipitation assays were performed using lysed spheroplasts. In brief, 100 ml of cells were harvested at A600 nm = 0.8–1.0 and resuspended in Tris-HCl (pH 8) and 10 mm dithiothreitol. Cells were incubated at room temperature for 10 min, centrifuged, and resuspended in 10 ml of spheroplast buffer (1 m sorbitol in 0.5× YPD medium and 50 mm KPO4) with oxylyticase and incubated at 30 °C for 30 min on a shaker. After chilling on ice, cell suspensions were overlaid on a sucrose cushion (20 mm HEPES, 1.2 m sucrose, and 0.02% sodium azide) and pelleted by centrifugation at 3000 × g for 25 min at 4 °C. The resulting spheroplasts (treated and untreated with 1 mm hydrogen peroxide at 30 °C for 40 min) were suspended in intracellular (lysis) buffer (100 mm potassium acetate, 50 mm KCl, 20 mm PIPES, and 200 mm sorbitol (pH 6.8)). Protein lysates were prepared by grinding the spheroplasts in the presence of glass beads, followed by centrifugation at 10,000 × g with recovery of the supernatant. These lysates were precleared by rotation with protein A-agarose mixture (100 μl/ml of lysate) at 4 °C for 1 h, followed by centrifugation. Precleared lysates were rotated with 4 μg of anti-TAP antibody for 3 h at 4 °C. For immunoprecipitation, 50 μl of protein A-agarose were added to samples and further rotated for 1 h at 4 °C. The beads were then spun down at 10,000 × g for 2 min and washed twice with intracellular buffer. The immunoprecipitated proteins were recovered by adding Twirl buffer (8 m urea, 5% SDS, 10% glycerol, and 50 mm Tris (pH 6.8)). These precipitates were then loaded onto 12% polyacrylamide gel and probed with anti-Yap1p antibody (14Wemmie J.A. Steggerda S.M. Moye-Rowley W.S. J. Biol. Chem. 1997; 272: 7908-7914Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar) to detect Yap1p or with peroxidase-conjugated anti-peroxidase antibody to detect TAP-tagged Rox3p as described (27Puig O. Caspary F. Rigaut G. Rutz B. Bouveret E. Bragado-Nilsson E. Wilm M. Seraphin B. Methods (Orlando). 2001; 24: 218-229Google Scholar). Aliquots of the protein lysates prior to immunoprecipitation were also electrophoresed and blotted for Yap1p to ensure that equivalent levels of Yap1p were used in the co-immunoprecipitation assays. Differential Role of the Yap1p C Terminus in Oxidative Stress Induction—The importance of the C terminus of Yap1p in the oxidant-regulated nuclear localization of this factor has been well documented (10Kuge S. Jones N. Nomoto A. EMBO J. 1997; 16: 1710-1720Crossref PubMed Scopus (344) Google Scholar, 11Yan C. Lee L.H. Davis L.I. EMBO J. 1998; 17: 7416-7429Crossref PubMed Scopus (203) Google Scholar, 13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar, 16Delaunay A. Isnard A.D. Toledano M.B. EMBO J. 2000; 19: 5157-5166Crossref PubMed Scopus (414) Google Scholar). Previously, we demonstrated that normal function of the C-terminal region of Yap1p is crucial for H2O2-induced activation of the thioredoxin-encoding TRX2 gene but not an artificial Yap1p-responsive reporter gene (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar). A C-terminal mutant form of Yap1p lacking a cysteine residue in the C-CRD (C629A Yap1p) is constitutively localized to the nucleus and fails to normally induce TRX2 expression upon H2O2 challenge (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar). We compared the ability of this mutant Yap1p derivative to control expression of the oxidant-inducible GSH1 gene (28Stephen D.W. Rivers S.L. Jamieson D.J. Mol. Microbiol. 1995; 16: 415-423Crossref PubMed Scopus (176) Google Scholar), encoding γ-glutamylcysteine synthetase, the first step in glutathione biosynthesis (29Ohtake Y. Yabuuchi S. Yeast. 1991; 7: 953-961Crossref PubMed Scopus (112) Google Scholar). GSH1 transcription has been previously shown to be oxidant-responsive in a Yap1p-dependent fashion (8Wu A.-L. Moye-Rowley W.S. Mol. Cell. Biol. 1994; 14: 5832-5839Crossref PubMed Google Scholar, 28Stephen D.W. Rivers S.L. Jamieson D.J. Mol. Microbiol. 1995; 16: 415-423Crossref PubMed Scopus (176) Google Scholar). A yap1Δ mutant strain was transformed with low-copy-number plasmids expressing either wild-type or C629A Yap1p along with lacZ gene fusions to either TRX2 or GSH1. Appropriate transformants were then tested for β-galactosidase expression in the presence or absence of oxidative stress (Fig. 1). GSH1-lacZ expression was elevated by ∼2- and 3.5-fold upon oxidative stress induced by H2O2 or diamide, respectively, in the presence of wild-type Yap1p, consistent with the findings of other groups (28Stephen D.W. Rivers S.L. Jamieson D.J. Mol. Microbiol. 1995; 16: 415-423Crossref PubMed Scopus (176) Google Scholar, 30Sugiyama K. Izawa S. Inoue Y. J. Biol. Chem. 2000; 275: 15535-15540Abstract Full Text Full Text PDF PubMed Scopus (119) Google Scholar). Expression of C629A Yap1p increased GSH1 expression by 2-fold in the absence of oxidant, and this expression failed to increase in the face of H2O2 exposure and was only modestly enhanced during diamide stress. Notably, expression of GSH1 during H2O2 treatment was identical in the presence of either wild-type or C629A Yap1p. This behavior contrasts with a TRX2-lacZ fusion gene that showed a striking expression defect in the presence of C629A Yap1p during H2O2 stress. C629A Yap1p was unable to significantly induce TRX2 upon H2O2 challenge. We believe that this selective inability to drive TRX2 expression in the presence of H2O2 is at the heart of the failure of C-terminal mutant forms of Yap1p to support normal resistance to this oxidant (13Coleman S.T. Epping E.A. Steggerda S.M. Moye-Rowley W.S. Mol. Cell. Biol. 1999; 19: 8302-8313Crossref PubMed Scopus (127) Google Scholar, 14Wemmie J.A. Steggerda S.M. Moye-Rowley W.S. J. Biol. Chem. 1997; 272: 7908-7914Abstract Full Text Full Text PDF PubMed Scopus (85) Google Scholar, 16Delaunay A. Isnard A.D. Toledano M.B. EMBO J. 2000; 19: 5157-5166Crossref PubMed Scopus (414) Google Scholar, 31Takeuchi T. Miyahara K. Hirata D. Miyakawa T. FEBS Lett. 1997; 416: 339-343Crossref PubMed Scopus (22) Google Scholar). This analysis demonstrated that the wild-type Yap1p C-CRD is required for significant induction of TRX2 but not GSH1 during H2O2-induced oxidative stress, indicating that TRX2 places a unique requirement on the C terminus of Yap1p that is not shared by all Yap1p target genes. To explore this unique demand on Yap1p at the TRX2 promoter, we characterized the structure of this promoter. YREs in TRX2—Previous work suggested the presence of two YREs in the 5′-noncoding region of TRX2 located at positions -218 and -181 upstream of the ATG codon (7Kuge S. Jones N. EMBO J. 1994; 13: 655-664Crossref PubMed Scopus (383) Google Scholar). A double mutant promoter lacking both of these YREs was unable to respond to Yap1p. To determine the individual contribution of these two putative YREs to Yap1p regulation of TRX2, we constructed a set of mutant TRX2 promoters lacking either one or both of the TRX2 YREs. We carried out DNase I protection analysis using bacterially produced Yap1p to determine whether both YREs were bound by Yap1p in vitro and to ensure that the mutant sites were no longer recognized by the recombinant protein (Fig. 2). The YRE at position -181 was strongly protected from DNase I cleavage by Yap1p. Only very weak interaction was detected at the -218 YRE, and the mutant YREs failed to detectably bind this factor in vitro. To assess the functional contribution of these YREs to TRX2 expression, two analyses were carried out. First, each mutant promoter was fused to lacZ to facilitate measurement of gene expression. Second, each mutant promoter was placed in the context of the wild-type TRX2 gene to evaluate the effect of removal of the YREs on the ability of the resulting constructs to complement the H2O2 sensitivity of a trx2Δ strain. Both the lacZ fusions and the TRX2 clones were returned to yeast on low-copy-number plasmids, and appropriate assays were carried out (Fig. 3). Each TRX2-lacZ fusion plasmid was introduced into either wild-type or yap1Δ cells, and β-g
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