Pdr3 is required for DNA damage induction of MAG1 and DDI1 via a bi-directional promoter element (original) (raw)

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Department of Microbiology and Immunology, University of Saskatchewan, 107 Wiggins Road, Saskatoon, SK S7N 5E5, Canada

*To whom correspondence should be addressed. Tel: +1 306 966 4308; Fax: +1 306 966 4311; Email: wei.xiao@usask.ca Present address: Yu Zhu, Division of Experimental Pathology, Department of Laboratory Medicine and Pathology, Mayo Foundation, Rochester, MN 55905, USA

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Department of Microbiology and Immunology, University of Saskatchewan, 107 Wiggins Road, Saskatoon, SK S7N 5E5, Canada

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01 January 2004

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Yu Zhu, Wei Xiao, Pdr3 is required for DNA damage induction of MAG1 and DDI1 via a bi-directional promoter element, Nucleic Acids Research, Volume 32, Issue 17, 1 September 2004, Pages 5066–5075, https://doi.org/10.1093/nar/gkh838
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Abstract

In order to understand how gene regulation is achieved in eukaryotes in response to DNA damage, we used budding yeast as a model lower eukaryotic organism and investigated the molecular events leading to the expression of two closely clustered damage-inducible genes, MAG1 and DDI1. MAG1 and DDI1 are co-activated by a shared 8 bp repeat sequence, UAS_DM_. In this study, we screened a yeast genomic library, identified Pdr3 as the transcriptional activator and demonstrated in vivo and in vitro that Pdr3 binds UAS_DM_. Pdr3 is required for the activation of a number of genes encoding membrane efflux pumps and deletion of PDR3 results in reduced basal-level expression and loss of DNA damage induction of MAG1 and DDI1. Interestingly, Pdr1, another transcriptional activator homologous to Pdr3 that is also required for the activation of multidrug-resistance genes, is not involved in the regulation of MAG1 and DDI1 expression, although it may also bind to UAS_DM_. Deletion of PDR3 does not affect the expression of other well-documented DNA damage-inducible genes; hence, yeast DNA damage-inducible genes appear to have distinct effectors although to a certain extent they share a common regulatory pathway mediated by DNA damage checkpoints.

Received May 31, 2004; Revised and Accepted September 2, 2004

INTRODUCTION

DNA-damaging agents are the major source of environmental genotoxic pollution. In order to maintain genome integrity, all living organisms have developed a comprehensive set of DNA repair genes. Many of these genes are induced upon DNA damage through a signal transduction cascade. The SOS regulatory network of Escherichia coli is one of the damage response mechanisms that is understood in considerable detail (1,2). However, the regulatory network governing DNA damage response appears to be rather different in eukaryotes. The budding yeast Saccharomyces cerevisiae has been extensively used as a model organism to explore whether eukaryotes possess a regulatory cascade analogous to the prokaryotic SOS system. There are a large number of DNA metabolism-related genes that are expressed at a low level and their expression is activated upon exposure to DNA damage (2–4). The expression of many of these genes appears to be affected by cell-cycle checkpoints through a central kinase cascade represented by Mec1–Rad53–Dun1 (4–6), and it has been suggested that this checkpoint pathway may be analogous to the E.coli SOS response (7,8). Surprisingly, there does not appear to be a common downstream transactivator that controls the expression of a large number of genes. For example, RNR genes including RNR1, RNR2, RNR3 and RNR4, which encode the large and small subunits of ribonucleotide reductase, can be induced by a variety of DNA-damaging and DNA replication-blocking agents (9,10). The induction is regulated by the Mec1–Rad53–Dun1 kinases and accomplished by release of a transcriptional repressor, Crt1, from the X-box found in the promoter of RNR genes (8). The DNA damage induction of the PHR1 gene, which encodes the apoenzyme for the DNA repair enzyme photolyase (11), requires Rad53 but neither Dun1 nor Chk1 (12). It is immediately regulated by two repressors Rph1 and Gis1 (13). These observations suggest that eukaryotic cells may possess multiple pathways to respond to DNA damage.

MAG1 encodes a 3-methyladenine (3MeA) DNA glycosylase (14), which acts in the first step of a multistage base excision repair pathway for the removal of lethal lesions such as 3MeA, and protects yeast cells from killing by DNA-alkylating agents, such as methyl methanesulfonate (MMS) (15). MAG1 is inducible by a variety of DNA-damaging agents, regardless of whether Mag1 is required for the repair of the type of damage (14,16,17). DDI1 encodes an ubiquitin-related protein and is involved in a DNA-damage cell-cycle checkpoint (18). It is located immediately upstream of MAG1, transcribed in an opposite direction, and is co-induced with MAG1 (19). Among a number of upstream activating sequences (UASs) and upstream repressing sequences (URSs) defined within the promoter region of MAG1 and DDI1 (17,19,20), only an 8 bp tandem repeat designated as UAS_DM_ is required for the regulation of both MAG1 and DDI1 (19).

Gene regulation is often achieved by interaction between cis_-acting elements and sequence-specific trans_-acting regulatory proteins. Indeed, previous studies revealed that UAS_DM binds to a protein from yeast crude cell extract (19), suggesting that UAS_DM is a classical bi-directional UAS element that interacts with a transcriptional activator and directs the downstream gene expression. Hence, it is important to identify the regulatory protein(s) in order to gain a full understanding of how MAG1 and DDI1 are co-regulated in response to DNA damage. In this study, a yeast one-hybrid screen was employed to identify the trans_-acting transcriptional factor(s) that binds to UAS_DM. Our results showed that Pdr3, a transcriptional activator previously characterized as a multidrug-resistance gene product, is required for the basal-level and DNA damage induction of both MAG1 and DDI1. Further investigations into the pathways involved in the above regulation suggest a novel stress-responsive network in eukaryotic cells that coordinately regulates the responses to different environmental stresses.

MATERIALS AND METHODS

Yeast strains and plasmids

Yeast strains used in this study are listed in Table 1. Wild-type FY1679-28c and its pdr1 and pdr3 deletion mutants were originally from Dr A. Delahodde (CNRS Paris, France). Wild-type JD52 and its _rpn4_Δ mutant (21) were received from Dr A. Varshavsky (California Institute of Technology, CA, USA).

Table 1.

Saccharomyces cerevisiae strains

Table 1.

Saccharomyces cerevisiae strains

Yeast cells were grown at 30°C in either complete YPD or SD (synthetic dextrose) medium supplemented with the appropriate nutrients (22). Plasmids were transformed into yeast cells by a lithium acetate protocol (23). The transformants were streaked onto a fresh selective plate before being utilized for further analysis.

To delete the PDR3 gene from the yeast genome, a PDR3 disruption plasmid was constructed as follows. Plasmid pGEX–PDR3 was obtained from Dr Delahodde and contains the 2.9 kb PDR3 open reading frame as a BamHI fragment cloned into pGEX-2T (24). A 1.4 kb SnaBI–HindIII fragment within the PDR3 open reading frame was deleted and replaced with a BglII linker to form pdr3ΔBg. A 3.8 kb BglII–BamHI fragment containing hisG–URA3–hisG was isolated from plasmid pNKY51 (25) and cloned into pdr3ΔBg to form pdr3Δ::hisG–URA3–hisG. The pdr3Δ::hisG–URA3–hisG disruption cassette was released by BsrGI and EcoRI digestion prior to yeast transformation. The pdr3 deletion strains were confirmed by Southern hybridization, and the URA3 pop-out strains were subsequently selected on plates containing 5-fluoroorotic acid.

Plasmids used for the detection of β-galactosidase (β-gal) expression, YEp_MAG1_–lacZ, YEp_DDI1_–lacZ, YEp_MAG1_ΔDR–lacZ, YEp_DDI1_ΔDR–lacZ, pWX1254 and pWX1813 have been described previously (17,19,26). Plasmid pZZ18 (YEp, HIS3, RNR2–lacZ) was received from Dr S. Elledge (Baylor College of Medicine, Houston) and pGBS116 (YEp, URA3, PHR1–lacZ) from Dr G. B. Sancar (University of North Carolina at Chapel Hill).

One-hybrid library screening

The host strain YM4271 (Table 1) and vector plasmids pHISi-1 and pLacZi used in the one-hybrid screen were purchased from Clontech (Matchmaker One-hybrid System, Palo Alto, CA, USA) and used as instructed. Two oligonucleotides, DR-1 (5′-TCGACGGTGGCGATGAATTTACAGGGCGGGGTGGCGAC-3′) and DR-2 (5′-TCGAGTCGCCACCCGCCCTGTAAATTCATCGCCACCG-3′) containing the UAS_DM_ sequence (8 bp tandem sequence underlined) were annealed, the 5′ protruding ends were filled by the Klenow fragment of E.coli DNA polymerase I, and the blunted fragment cloned into the SmaI site of pHISi-1 and pLacZi. The resulting positive clones were screened by sequencing for those containing three copies of the UAS_DM_ inserts in the same orientation, which were named pHISi–3UASDM and pLacZi–3UASDM, respectively. Both plasmids were integrated into the genome of YM4271 to make YM4271DM. The GAL4AD fusion yeast genomic library (27), a gift from Dr D. Gietz (University of Manitoba, Canada), was used for screening genes encoding proteins that bind to UAS_DM_. The library was a mixture of three separate libraries differing in their vector GAL4AD reading frame. Hence, the random yeast genomic DNA fragments may be fused to GAL4AD in all three possible reading frames. The library DNA was transformed into YM4271DM; transformants were screened for growth in SD medium without Ura, Leu or His but containing 30 mM 1,2,4-amino triazole (3-AT). After confirmation of 3-AT resistance, the yeast transformants were subjected to a β-gal filter assay (28). Plasmids from the positive clones were recovered in E.coli DH10B cells and used to re-transform YM4271DM cells to confirm the positive phenotypes and to quantitatively measure the β-gal activity.

RNA isolation and northern hybridization

One milliliter overnight culture was used to inoculate 3 ml of fresh medium, and cells were incubated for another 2 h. For MMS treatment, MMS was added to the indicated final concentrations and the incubation was continued for another 30 min. RNA was isolated by a glass-bead method (29), separated by gel electrophoresis, blotted on a GeneScreen Plus membrane (NEN, Boston) and hybridized with α-32P-labeled DNA probe as instructed. After an overnight hybridization, the membrane was washed and exposed to a phosphorimager screen. The mRNA band intensity was measured by Molecular Imager FX (Bio-Rad, Hercules, CA) supported by the Quality One 4.2.1 software. The source and preparation of MAG1, DDI1 and ACT1 probes have been described previously (26).

β-galactosidase assay

β-gal activity was measured as described previously (30) to determine the levels of MAG1–lacZ and DDI1–lacZ gene expression. Briefly, a 0.5 ml fresh overnight culture of the yeast transformants was used to inoculate 2.5 ml of minimal medium without Leu or without Ura, depending on the selectable marker of the lacZ fusion plasmid. Two sets of cells were allowed to grow for another 2 h. MMS was added to one set of cells at the desired concentration, and both sets were incubated at 30°C for 4 h. One milliliter of cells was used to determine cell titer by optical density at 600 nm. The remaining 2 ml of cells were collected and permeabilized in 1 ml of Buffer Z, and β-gal expression was determined. The specific activity is expressed in Miller units. All the results presented are the average of at least three independent experiments with standard deviations less than 30% of the mean values.

Preparation of crude cell extracts

Yeast cultures were grown to mid-log phase in YPD, and were harvested by centrifugation at 4°C. All of the following steps were carried out at 4°C or on ice. Cells were washed once with buffer A [25 mM HEPES (pH 7.5), 5 mM MgCl2, 0.1 mM dithiothreitol, 10% glycerol, 50 mM KCl] and resuspended in 1 ml of buffer A containing 0.3 M (NH4)2SO4 and protease inhibitors (1 mM phenylmethylsulfonyl fluoride, 5 μg/ml antipain, 1 μg/ml leupeptin and 1.4 μg/ml pepstatin). The cells were disrupted by vortexing with glass-beads (0.4–0.5 mm; acid-washed); after centrifugation at 8000 r.p.m. for 20 min, aliquots of the supernatant were immediately frozen and stored at −70°C.

To make bacterial crude cell extracts, BL21(DE3)–RIL cells (Stratagene, La Jolla, CA) transformed with pGEX–PDR3 or vector alone were incubated, induced with 0.4 mM IPTG for 2 h and crude cell extracts were prepared in the presence of protease inhibitors as previously described (31). The expression of target protein was confirmed by western analysis using an anti-glutathione _S_-transferase (anti-GST) antibody (Amersham, Piscataway, NJ).

Electrophoretic mobility shift assay (EMSA)

Double-stranded oligonucleotides were made by mixing equal amounts of complementary oligonucleotides at a concentration of 1 μg/μl, heating to 70°C for 10 min and allowing to slowly cool down to room temperature. The UAS_DM_ oligonucleotide probe was prepared by an end-filling method in the presence of [α-32P]dCTP, dATP, dGTP, dTTP and Klenow fragment, and unincorporated nucleotides were removed by repeated ethanol precipitation. The UAS_DM_ competitor was made by incubating the double-stranded DR-1/DR-2 oligonucleotides with the Klenow fragment and dNTPs. Protein–DNA interaction was assayed in buffer A plus 1 mM phenylmethylsulfonyl fluoride. Standard reaction mixtures (20 μl) contained 1 μg of poly(dI–dC), 2 μg of BSA, 5 ng of probe, and various amounts of cell extract with or without competitors. After incubation at room temperature for 30 min, the reaction mixture was separated by electrophoresis in a 6% polyacrylamide gel containing 2.5% glycerol. The gel was dried and exposed to X-ray film.

Chromatin immunoprecipitation (ChIP) assay

The random hemagglutinin epitope tag (HAT) inserted diploid strains were created by means of transposon insertion containing the HAT sequence (32). We received two such strains from Dr M. Snyder (Yale University) containing the in-frame HAT insertion in the coding region of PDR3, among which TN7-87A9 (Table 1) was found to be capable of supporting MAG1–lacZ and DDI1–lacZ DNA damage induction like its wild-type cells, and the expected Pdr3::HA band was identified by western analysis using an anti-HA polyclonal antibody (Upstate, Charlottesville, VA). After growing 50 ml TN7-87A9 cells to log phase, 1.4 ml of 37% formaldehyde solution was added and the incubation continued for 15 min at room temperature to crosslink proteins to the chromatin. To quench crosslinking, 2.5 M glycine was added to the above culture to a final concentration of 125 mM and the incubation continued for 5 min at room temperature. Chromatin-containing whole-cell extract was then prepared from the above culture as described in (33). Approximately 2 × 108 cell equivalents of the whole-cell extracts with an average size of 0.5–1 kb chromatin fragments were incubated with 10 μl of polyclonal anti-HA antibody in a final volume of 0.2 ml for 2 h. An aliquot of 10 μl Protein A Sepharose beads (Sigma, St Louis) was added and the mixture was incubated overnight at 4°C. The immunoprecipitates were washed stringently and the recovered chromatin (precipitates) as well as the total chromatin (input) were incubated with 0.5 μg/μl proteinase K at 37°C for 2 h to remove proteins from the chromatin and phenol–chloroform purified genomic DNA was then used as templates for PCR analysis using primer sets of MAG1-1 (5′-GGCAGTGGCCAATTCTC-3′)/MAG1-8 (5′-CTGCAGTAATGCTATTAG-3′), which amplify a 0.3 kb MAG1–DDI1 promoter region containing the UAS_DM_, and SSU1-2 (5′-GCCGAATTCATGGTTGCCAATTGGGTA-3′)/SSU1-3 (5′-GCCCTCGAGTGCCAATTATGTACGTAT-3′), which amplify a 1.4 kb SSU1 coding region. The PCR products were then subjected to agarose gel electrophoretic analysis.

RESULTS

Isolation of PDR3 by library screen

UAS_DM_ was previously found to control the expression of both MAG1 and DDI1 (19). In order to identify the trans_-acting protein that interacts with UAS_DM, we performed a one-hybrid screen using the 32 bp UAS_DM_ as bait (Figure 1A) against a Gal4AD-fusion yeast genomic library. In order to enhance the screening stringency and eliminate false-positive clones, three tandem copies of the UAS_DM_ elements were inserted upstream of two separate reporter genes, HIS3 and lacZ, which lack their own promoters (Figure 1A). Furthermore, both reporter genes were integrated into the yeast genome to reduce potential background expression associated with multicopy plasmids. A total of 72 independent transformants formed colonies on the selective plates containing 30 mM 3-AT, among which 18 were also positive in a β-gal filter assay. Upon plasmid recovery and re-transformation into the reporter strain, 12 clones maintained 3-ATR and β-gal activity, as shown by a representative clone (Figure 1B, PDR3) and thus were considered positive clones. DNA sequencing of the 5′ and 3′ ends of genomic DNA inserts from each of the positive clones revealed that 9 out of 12 clones contain the full-length PDR3 gene. No other genes adjacent to PDR3 are intact in all nine clones. Hence, Pdr3 appears to be the primary candidate that binds to UAS_DM_ in the one-hybrid assay. Furthermore, DNA sequencing revealed that none of the clones contain proper gene fusion that would produce in-frame Gal4AD–Pdr3. Hence, we conclude that Pdr3 activates UAS_DM_ through its own DNA binding and activation domains, that the library simply serves as a yeast multicopy genomic library and that results shown in Figure 1B served as a promoter–reporter gene assay.

Figure 1.

Pdr3 binds and activates UAS_DM_ in a one-hybrid assay. (A) A diagram of UAS_DM_–lacZ and UAS_DM_–HIS3 reporter constructs. Three copies of UAS_DM_, each containing an 8 bp tandem repeat, were inserted into the promoter region of the one-hybrid reporter genes lacZ and HIS3. Both reporter genes were integrated into the genome of host strain YM4271 to create YM4271DM. (B) Multicopy PDR3, but not RPN4, is able to transactivate UAS_DM_–lacZ and UAS_DM_–HIS3. YEp-based plasmid carrying either PDR3, RPN4 or the vector alone (mock) was transformed into YM4271DM. The transformants were replica-plated onto SD minimal selective medium or SD+ 30 mM 3-AT and incubated at 30°C for 2 days (for UAS_DM_–HIS3 expression). Cells from the SD medium were used for an X-gal filter assay and incubated at 30°C for 2 h (for UAS_DM_–lacZ expression) before taking photograph.

Pdr3 is an in vivo transactivator of MAG1 and DDI1

PDR3 encodes a 976 amino acid, 112.5 kDa protein that is an important transcriptional activator involved in multidrug resistance (34,35). PDR3 regulates the expression of several multidrug-resistance genes encoding ABC transporters serving as membrane efflux pumps; the pdr3 null mutant displays enhanced sensitivity to rhodamine 6G and diazaborine but not to cycloheximide (24,36). Our observation that Pdr3 binds and activates reporter gene expression through UAS_DM_ in a dual-promoter assay suggests, but does not prove, that PDR3 is involved in the regulation of MAG1–DDI1 expression in vivo. In order to confirm the effect of PDR3 on the damage induction of MAG1 and DDI1, we examined the expression of MAG1 and DDI1 in wild-type and its isogenic pdr3 null mutant by both northern hybridization (Figure 2A) and using lacZ fusion constructs (Figure 2B). Compared with isogenic wild-type cells, the pdr3 mutation reduced basal-level expression of MAG1 and DDI1 by 2.5–3-fold and DNA damage induction by up to 8-fold. This result is comparable with our previous observation of the deletion of, and site-specific mutagenesis within, UAS_DM_ (19). In order to further confirm that Pdr3 is acting on UAS_DM_, we determined the effects of pdr3 on MAG1 and DDI1 promoters deleted for UAS_DM_. As seen in Figure 3, deletion of PDR3 does not further affect MAG1 and DDI1 expression if the UAS_DM_ is deleted, indicating that UAS_DM_ is indeed the target of Pdr3 activation. The results are thus consistent with our hypothesis that Pdr3 acts as a transcriptional activator of MAG1 and DDI1 through the bi-directional UAS_DM_ element shared by these two promoters. Furthermore, the good correlation between results of northern hybridization and lacZ reporters lends support to our use of the lacZ fusion constructs for future quantitative analyses.

Figure 2.

Deletion of PDR3 results in decreased basal-level expression and loss of DNA damage induction of both MAG1 and DDI1. (A) Northern analysis. Total RNA was isolated from FY1679-28C (wt) and naΔ3 (_pdr3_Δ) after 0, 0.01, 0.05 and 0.1% MMS treatments (indicated on the top panel for 30 min.) Each lane contains about 15 μg of total RNA. The blot was sequentially hybridized and stripped with MAG1, DDI1 and ACT1 probes. (B) MAG1–lacZ and DDI1–lacZ induction by 0.05% MMS treatment for 4 h in the wild-type (FY1679-28C) and _pdr3_Δ (naΔ3) mutant transformed with either MAG1–lacZ or DDI1–lacZ. The results are the average of 3–6 independent experiments. β-gal activity is given in Miller units.

Figure 3.

UAS_DM_ deletion is epistatic to pdr3_Δ. JD52 (wt) and its pdr3_Δ derivative WXY1090 were transformed with either YEp_MAG1_ΔDR–_lacZ (A) or YEp_DDI1_ΔDR–_lacZ (B) and the β-gal activity was determined as described. The results are the average of three independent experiments.

Pdr1 contributes to in vitro binding of UAS_DM_, but does not play a role in MAG1–DDI1 activation

PDR1 is a structural and functional homolog of PDR3 (37). Pdr1 and Pdr3 have overlapping functions in the regulation of multidrug-resistance genes, and both transcriptional factors bind to the same consensus palindrome sequence, 5′-TCCGCGGA-3′, called pleiotropic drug resistance element (PDRE), which is identified in the promoter region of most of their target multidrug-resistance genes. The UAS_DM_ tandem repeat sequence is rather different from PDRE, and no other PDRE sequence is found in the internal region flanked by the UAS_DM_ tandem repeats (Figure 1A). In order to establish that Pdr3 and/or Pdr1 indeed bind to UAS_DM_, an EMSA was performed by using radioactively labeled UAS_DM_ as a probe against total yeast cell extracts. As previously reported (19) and shown in Figure 4, a cell extract from wild-type cells forms two protein–DNA complexes with UAS_DM_ in a dose-dependent manner. Increased concentration of cell extract appears to favor the higher molecular weight complex formation over the lower molecular weight complex. The formation of both complexes was abolished by proteinase K treatment (lane 1), indicating that they are indeed protein–DNA complexes. The complexes are deemed DNA sequence-specific, since their formation was inhibited by excess unlabeled UAS_DM_ (lane 2), but not by excess non-specific DNA such as poly(dI–dC) (in all reactions) and other double-stranded DNA (data not shown). The band intensity, especially that of the higher molecular weight band, decreased dramatically with a cell extract from the pdr3 mutant, but did not show a significant alteration with a cell extract from the pdr1 mutant. In contrast, the lower molecular weight complex appears to decrease with increasing wild-type cell extract, whereas this complex formation in pdr1 and pdr3 mutants is correlated with amount of cell extracts. It is interesting to note that Pdr1 and Pdr3 can form a heterodimer to bind PDRE (38). Nevertheless, the formation of both UAS_DM_–protein complexes was completely abolished in the pdr1 pdr3 double mutant. This result indicates that Pdr3 is the major UAS_DM_-binding protein in yeast cells, whereas Pdr1 may also be able to bind UAS_DM_ in the absence of Pdr3.

Figure 4.

EMSA using the UAS_DM_ probe. Each reaction contained 1 μg of poly(dI–dC), 2 μg of BSA, 5 ng of labeled UAS_DM_ probe and various amount of yeast cell extract in buffer A. The source of yeast crude extract used in each reaction is indicated. Yeast strains used in this study: FY1679-28C (wt), yYA14 (_pdr1_Δ), naΔ3 (_pdr3_Δ) and naΔ1Δ3 (_pdr1_Δ pdr3_Δ). All strains are isogenic to FY1679-28C. Cell extracts with increasing amounts are 5, 10 and 20 μg. Lane 0, no protein control. Lanes 1 and 2 contain 20 μg wild-type cell extract plus 10 μg proteinase K (lane 1) or 1 μg UAS_DM competitor (lane 2).

The contribution of Pdr1 in the in vitro binding of UAS_DM_ in the above EMSA appears to imply that PDR1 plays a backup or overlap role in the in vivo regulation of MAG1–DDI1 expression, as seen in the regulation of PDR genes (35). To address this possibility, we measured the β-gal activity of MAG1–lacZ and DDI1–lacZ transformants in the isogenic pdr1 and pdr3 single mutants and the pdr1 pdr3 double mutant. Results as shown in Figure 5 clearly indicate that the pdr1 mutation does not alter the basal-level or DNA damage-induced MAG1 and DDI1 expression in either wild-type or pdr3 backgrounds. Hence, we conclude that although Pdr1 appears to be able to bind UAS_DM_ in vitro, it is not involved in the transcriptional regulation of MAG1 and DDI1.

Figure 5.

DNA damage induction of MAG1–lacZ and DDI1–lacZ in the pdr1 and pdr3 mutants. (A) MMS-induced MAG1–lacZ expression. (B) MMS-induced DDI1–lacZ expression. Yeast strains used in this study are the same as in Figure 4. The results are the average of at least three independent experiments.

Pdr3 binds UASDM both in vivo and in vitro

The results obtained from the EMSA using yeast cell extracts suggest that Pdr3 is required for UAS_DM_–protein interaction, but do not clarify whether Pdr3 is directly involved in UAS_DM_ promoter binding in vivo. A ChIP assay is considered a direct assessment of protein–DNA interaction in vivo. Such a result as shown in Figure 6A provides evidence that indeed Pdr3 is associated with the UAS_DM_ promoter region in vivo, as in a strain expressing a native level of HA-tagged Pdr3, antibodies against the HA tag are specifically associated with DNA fragments containing UAS_DM_, but not unrelated genomic DNA fragments.

Figure 6.

In vivo and in vitro binding of Pdr3 to UAS_DM_. (A) Interaction of Pdr3 and UAS_DM_ by a ChIP assay. The ChIP experiment was performed as described using TN7-87A9 cells expressing Pdr3-HA. After HA immunoprecipitation and extensive washing, the chromatin-containing samples were used as templates for 30-cycle PCR using UAS_DM_-specific (MAG1-1/MAG1-8, 0.3 kb) and non-specific (SSU1-2/SSU1-3, 1.4 kb) primer pairs. Templates used: lane 1, DNA after immunoprecipitation; lane 2, DNA before immunoprecipitation (input DNA); lanes 3 and 4, total yeast genomic DNA. Primers used in the PCR reaction: lanes 1–3, MAG1-1/MAG1-8 plus SSU1-2/SSU1-3; lane 4, SSU1-2/SSU1-3. Lane 5 contains molecular size marker (in kb). Lane 1 contains the ChIP result and lanes 2–4 serve as various controls. (B) Physical interaction of bacterially produced GST–Pdr3 with UAS_DM_ in EMSA. All reactions contained 5 ng of labeled UAS_DM_ probe. Lane 1, no protein control; lanes 2–4, 1, 2 and 4 μg cell extracts, respectively, from the pGEX–PDR3 (GST–Pdr3) transformant; lane 5, 2 μg cell extract from the pGEX (GST) vector transformant; lane 6, same as in lane 4 but the sample was incubated with 10 μg proteinase K for 10 min prior to loading.

Although the above ChIP result demonstrates the assembly of Pdr3 to the MAG1–DDI1 dual promoter in vivo, it does not rule out the possibility that Pdr3 is assembled to this promoter through other DNA-binding factors. To further address whether this interaction is a result of direct contact between Pdr3 and UAS_DM_, we performed an EMSA using bacterial cell extracts (Figure 6B) and found that cells expressing GST–Pdr3 were able to form a specific Pdr3–UAS_DM_ complex in a dose-dependent manner (lanes 2–4), whereas the extract from cells expressing GST alone (lane 5) was unable to form such a complex. The complex is abolished by proteinase K treatment (lane 6), indicating that it is indeed a DNA–protein complex. Hence, Pdr3 alone appears to be able to interact with UAS_DM_ both in vivo and in vitro.

Deletion of PDR3 does not affect RNR or PHR1 expression

Transcriptional regulators of RNR (8) and PHR1 (13) genes have been previously identified. In both cases, the immediate regulators are repressors and the DNA damage induction is achieved through derepression. We have previously shown that genes involved in the regulation of _RNR_s (CRT1, SSN6 and TUP1) and PHR1 (RPH1 and GIS1) are not involved in the regulation of MAG1 and DDI1 (39). In this study, we found that deletion of PDR3 has no apparent effect on basal-level and DNA damage-induced expression of PHR1 (Figure 7A), RNR2 (Figure 7B) and RNR3 (data not shown). Hence, we conclude that all three sets of well-studied yeast damage-inducible genes (_RNR_s, PHR1 and MAG1–DDI1) have distinct regulators and that the regulatory mechanisms differ from each other.

Figure 7.

Deletion of PDR3 and RPN4 does not affect PHR1 and RNR2 expression. (A) PHR1–lacZ expression. (B) RNR2–lacZ expression. Yeast strains used in this study are: JD52 (wt), EJY140 (_rpn4_Δ), WXY1090 (_pdr3_Δ) and WXY1091 (_rpn4_Δ _pdr3_Δ).

Rpn4 is required for both MAG1 and DDI1 expression but does not appear to bind UASDM

Rpn4 was initially identified as a transcriptional activator through its interaction with proteasome-associated control element (PACE, 5′-GGTGGCAAA-3′), a nonamer box found in the promoters of several 26S proteasome genes (40,41). The rpn4 mutation inhibits the activity of ubiquitin/proteasome-dependent N-end rule and ubiquitin-fusion degradation pathways (42). Recently, it was shown that the rpn4 mutation abolished MMS-induced MAG1 expression (43). We examined MAG1 and DDI1 expression in the rpn4 mutant by northern hybridization (data not shown) and the β-gal assay (Figure 8), and found that deletion of RPN4 reduced both basal-level expression and DNA damage induction in a manner as seen in the pdr3 mutant. Furthermore, our observation that both MAG1 and DDI1 expression is affected by rpn4 (Figure 8), that the effect is comparable to the UAS_DM_ deletion/mutation (19) and that UAS_DM_ is the only known cis_-acting regulatory element shared by both MAG1 and DDI1, together strongly indicate that Rpn4 directly or indirectly acts on UAS_DM.

Figure 8.

DNA damage induction of MAG1–lacZ and DDI1–lacZ in the pdr3 and rpn4 mutants. (A) MMS-induced MAG1–lacZ expression. (B) MMS-induced DDI1–lacZ expression. The results are the average of at least three independent experiments. Yeast strains used in this study are the same as in Figure 7. The results are the average of at least three independent experiments.

As noted before (43), the 8 bp tandem repeat sequence in UAS_DM_ is very similar to PACE, differing in only two nucleotides, suggesting that Rpn4 may directly bind UAS_DM_. To determine whether Rpn4 is directly involved in UAS_DM_ binding in vivo, we performed the yeast reporter gene assay with a multicopy plasmid containing the RPN4 gene. Like Pdr3, Rpn4 contains an activation domain and is able to transactivate RPN genes (41); hence it does not need to be fused to GalAD for the assay. However, as shown in Figure 1B, under the conditions used to isolate and positively identify PDR3, overexpression of RPN4 in YM4271DM did not activate UAS_DM_–lacZ or UAS_DM_–HIS3. This observation is consistent with our previous one-hybrid library screen, which relies on direct physical contact between the DNA-binding protein and the target UAS_DM_. Despite repeated isolation of PDR3, none of the positive clones contains the RPN4 gene. Furthermore, deletion of RPN4 does not alter MAG1 and DDI1 expression in the pdr3 mutant cells (Figure 8), nor does it affect basal and DNA damage-induced expression of PHR1 (Figure 7A) or RNR2 (Figure 7B). We conclude from the above results that Rpn4 is required for MAG1 and DDI1 basal level and DNA damage induction, although it does not physically interact with its putative target UAS_DM_.

DISCUSSION

We report here the isolation and functional characterization of the PDR3 gene as a novel transcriptional activator of the bi-directionally expressed MAG1–DDI1 through a dual-promoter element UAS_DM_. We also demonstrate that another transcriptional activator, RPN4, is required for MAG1–DDI1 expression and that PDR3 and RPN4 appear to function in the same signal transduction pathway. PDR3 and RPN4 were previously identified to be involved in two distinct stress-response pathways. This is the first report that the two seemingly unrelated regulatory pathways contribute to DNA damage response by acting on a _cis_-acting element different from their respective consensus binding sequences.

PDR3 was repeatedly isolated through one-hybrid screening as the sole candidate that binds to and activates the UAS_DM_ tandem repeat. Indeed, deletion of PDR3 results in reduced basal level and DNA damage induction of MAG1 and DDI1. This is consistent with the phenotypes observed when UAS_DM_ is deleted or mutated in the above promoters (19), and provides direct evidence that Pdr3 is an in vivo transcriptional activator of UAS_DM_. Given the structural homology and functional overlap of Pdr1 with Pdr3 in the multidrug-resistance pathway, it was anticipated that Pdr1 may also play a role in the regulation of MAG1 and DDI1. However, deletion of PDR1 in either wild-type or in the pdr3 mutant does not alter MAG1–DDI1 basal or DNA damage-induced expression, although in the absence of Pdr3, Pdr1 appears to bind UAS_DM_ in an in vitro EMSA assay. The differential effect of PDR1 and PDR3 on the target gene is not unprecedented. For example, among ABC transporter genes regulated by PDR1/PDR3, the expression of PDR5 is significantly inhibited in the pdr1 single mutant, while SNQ2 transcription is only abolished in the pdr1 pdr3 double mutant (35). It was also shown that the activation of PDR5 by mitochondrial defects required the presence of PDR3 but not PDR1 (44). A recent microarray analysis (45) suggests that many yeast genes are differentially regulated by PDR1 and PDR3. Furthermore, PDR1 and PDR3 may act as either positive or negative regulators on different target genes.

Pdr1 and Pdr3 belong to a family of bi-nuclear Gal4-like Zn(II)2Cys6 transcription factors and their target PDRE sequence has been well defined (24,46). However, to the best of our knowledge, this is the first report that Pdr3, and possibly Pdr1, also recognizes promoter sequences other than PDRE and that Pdr3 is involved in the regulation of genes other than ABC transporter genes. The direct recognition of UAS_DM_ by Pdr3 is demonstrated by EMSA using yeast cell extracts as well as recombinant GST–Pdr3 produced in bacterial cells, and this result is further supported in vivo by the ChIP assay. This is reminiscent of the microarray study (45), which found that only 15 out of the 26 genes positively regulated by PDR1/PDR3 contain PDRE elements in their promoter regions. It remains to be determined whether there are other Pdr1 and/or Pdr3 binding consensus sequence than PDRE and UAS_DM_, or whether these genes are activated by a third transcriptional factor subject to regulation by Pdr1/Pdr3. Recently, it was shown that Pdr1 and/or Pdr3 can activate transcription through a downstream target gene YRR1, which also encodes a transcriptional factor (47).

Rpn4 functions as a transcriptional activator of genes encoding subunits of the 26S proteasome and several other components of the ubiquitination system (21,41). Rpn4 binds PACE, a consensus sequence found in the promoter region of 26 out of the 32 proteasomal genes (41). The rpn4 mutation was recently found to affect MAG1 expression and the effect was attributed to the sequence similarity between PACE and the 8 bp tandem repeats found in the promoter of MAG1 (43). We examined the roles of RPN4 in the basal-level and damage-induced expression of MAG1 and DDI1, and found that indeed RPN4 is required for the expression of both genes. Since UAS_DM_ is the only cis_-acting element known to function bi-directionally, our observations support the notion that RPN4 acts via the 8 bp tandem repeat. However, our results do not support UAS_DM as the direct binding site of Rpn4. First, RPN4 was not identified in a yeast library screen despite repeated isolation of PDR3, and overexpression of RPN4 does not activate UAS_DM_ reporter genes. Second, although the RNR2 promoter also contains a sequence with a 7/9 match to PACE and UAS_DM_, its expression was not affected by deletion of RPN4.

In summary, our observation that two distinct stress-responsive pathways (i.e. multidrug resistance and ubiquitination) are involved in DNA damage response suggests that eukaryotic cells may possess a central coordinating system in response to environmental stresses. The notion of cross-talk between stress-responsive pathways is supported by recent reports of interactions between Pdr1/3 and Rpn4 (48). This type of general control system appears to differ from bacteria, where a given environmental signal may trigger only a particular regulon with a specific set of genes solely responsible for the stress. It is generally believed that the difference between eukaryotes and prokaryotes in response to DNA damage is due to their distinct cell cycles. Indeed, the cell-cycle checkpoint genes are involved in the DNA damage induction of almost all the target genes analyzed in budding yeast (4,5,26,49), and the phenomenon has been referred to as a eukaryotic SOS response. Interestingly, although these DNA damage-inducible genes share a common regulatory pathway mediated by cell-cycle checkpoints, the effectors are apparently different and strikingly diverse. Among three well-studied cases, the four RNR genes are regulated by the repressor Crt1 and two co-repressors, Tup1 and Ssn6; inactivation of any one of these three proteins results in constitutive derepression (8). In contrast, PHR1 is regulated by two repressors, Rph1 and Gis1, and either is sufficient for repression (13). We have shown previously that inactivation of any of the above repressors has no effect on MAG1–DDI1 expression (39). Accordingly, this study demonstrates that two transactivators, Pdr3 and Rpn4, are required for the positive regulation of the dually expressed MAG1–DDI1 genes, but are not involved in the regulation of RNR2 and PHR1. A diagram summarizing our current understanding of DNA damage induction in budding yeast is shown in Figure 9. The fact that all three sets of genes studied so far are coordinately regulated by checkpoints, but each has its own effectors, testifies to the complexity of the signal transduction cascade in this simple eukaryote. Why eukaryotic cells evolved this complicated monitoring and regulatory system remains to be illustrated.

Figure 9.

Signal transduction cascade of DNA damage induction of selected genes in S.cerevisiae. Data are assembled from various literatures. Only three sets of target genes, whose promoters have been dissected and binding proteins identified, are presented. ‘P’, promoter region with defined target sequences. ‘X’, an unknown protein is expected to act as the target for Pds1 phosphorylation and mediate the regulation of MAG1–DDI1 expression. ‘?’, while Pdr3 is defined as an effecter, how Rpn4 relates to Pdr3 in the regulation of MAG1 and DDI1 is currently unclear.

The authors wish to thank Dr Gietz for the yeast two-hybrid library, Drs Delahodde, Elledge, Sancar, Snyder and Varshavsky for yeast strains and plasmids, Yu Fu for technical assistance and Michelle Hanna for proofreading the manuscript. This work was supported by the Natural Sciences and Engineering Research Council of Canada operating grant OGP0138338 to W.X.

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