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The regulatory inputs controlling pleiotropic drug resistance and hypoxic response in yeast converge at the promoter of the aminocholesterol resistance gene RTA1

Anna Kołaczkowska, Myriam Manente, Marcin Kołaczkowski, Justyna Laba, Michel Ghislain, Donata Wawrzycka
DOI: http://dx.doi.org/10.1111/j.1567-1364.2011.00768.x 279-292 First published online: 1 May 2012


Aminosterols possessing potent fungicidal activity are attractive alternatives to currently available antifungals. Although their precise mechanism of action is not fully understood, the effect of 7-aminocholesterol (7-ACH) involves a partial block of Δ8-Δ7 isomerase and C-14 reductase. The function of RTA1 encoding the 7-transmembrane helix protein, cloned as the multicopy suppressor of 7-ACH toxicity in yeast, remains unclear. In this report, we show that Rta1p is localized in the plasma membrane and has a high rate of metabolic turnover, as revealed by fluorescence microscopy, cell fractionation and pulse-chase experiments. Analysis of the RTA1lacZ reporter activity and deletion mapping of the promoter allowed the identification of the regions responsible for negative regulation by Tup1 and the two synergistically acting repressors of hypoxic genes, Rox1p and Mot3p. This was in line with increased RTA1-mediated resistance to 7-ACH under hypoxic conditions, associated with increased Rta1p level. Overexpression of RTA1 also affected the response to the signalling sphingolipid precursor phytosphingosine. Positive inputs of two transcriptional activators Pdr1p and Upc2p were also detected, indicating a regulatory link common to sterol biosynthetic genes as well as those involved in pleiotropic drug resistance and sphingolipid metabolism.

  • yeast
  • antifungal resistance
  • 7-aminocholesterol
  • phytosphingosine
  • transcriptional repressor
  • stress response


Sphingolipids and sterols are structural lipids present in the membranes of eukaryotic cells. In yeast, the metabolic precursor of sphingolipids is ceramide, which consists of a fatty acyl group linked to the C-2 amino group of dihydrosphingosine (DHS) or phytosphingosine (PHS) (Dickson & Lester, 1999). The most common sterol component of fungal membranes is ergosterol, which has a C7–C8 double bond (Gaber et al., 1989). When growing aerobically, Saccharomyces cerevisiae strains satisfy their sterol requirement by ergosterol biosynthesis. One of the enzymes involved in this process, Δ8-Δ7-sterol isomerase, is the target for the morpholine and piperidine derivatives that are used as antifungal compounds (Rahier et al., 1990). Triazole derivatives constitute a second family of sterol biosynthesis inhibitors (Sanati et al., 1997). Aminocholesterol derivatives have also been shown to impair sterol biosynthesis, and the sterol pattern of treated yeast is consistent with a reduction in Δ8-Δ7-sterol isomerase activity and C-14 sterol reductase activity (Elkihel et al., 1994). Thus, 7-aminocholesterol (7-ACH) and morpholines have the same target enzymes in the ergosterol pathway. However, cell growth is inhibited by 7-ACH without complete inhibition of ergosterol formation, indicating a different mechanism of toxicity from that of morpholines (Elkihel et al., 1994).

The RTA1 gene from S. cerevisiae was identified during screening for genes that confer resistance to 7-ACH, but not to azoles and morpholine derivatives, when cloned on a high copy number plasmid (Soustre et al., 1996). The encoded polypeptide, Rta1p, has seven predicted membrane-spanning segments and has high sequence similarity with homologous membrane proteins, such as Rtm1p (resistance to molasses) (Ness & Aigle, 1995), Rsb1p (resistance to sphingoid bases) (Kihara & Igarashi, 2002), Pug1p (protoporphyrin uptake gene) (Protchenko et al., 2008) and YLR046, an open reading frame of unknown function (Kihara & Igarashi, 2002). The addition of 7-ACH to cells containing or lacking the RTA1 on plasmid results in a comparable reduction in ergosterol levels, indicating that Rta1p is not involved in the efflux of 7-ACH (Soustre et al., 1996). Although the maintenance of normal ergosterol levels is crucial for S. cerevisiae, a null rta1Δ mutant is viable under normal growth conditions (Soustre et al., 1996), indicating that the RTA1 gene is not essential.

Large-scale mRNA expression profiling indicated that transcription of RTA1 is induced by 8-methoxypsoralen and UVA (Dardalhon et al., 2007), adverse conditions such as high hydrostatic pressure (Iwahashi et al., 2005) or the presence of the mycotoxin citrinin (Iwahashi et al., 2007), indicating its possible role in response to stress stimuli. The molecular mechanisms underlying its regulatory circuitry are not known. Independent genome microarray analyses using a hyperactive mutant version of the Zn(II)-Cys6 transcription factor Pdr1p (DeRisi et al., 2000) or a chimeric fusion protein (Devaux et al., 2001; Onda et al., 2004) suggested that RTA1 and RSB1 are among multiple transcriptional targets of this major activator of the pleiotropic drug resistance network of genes. These include the major multidrug transporters of the ATP-binding cassette (ABC) superfamily, along with many genes involved in multiple cellular functions (Bauer et al., 1999; Kolaczkowska & Goffeau, 1999; Shahi & Moye-Rowley, 2009). Strikingly, the functional binding sites for Pdr1p and its close homologue Pdr3p, the so-called pleiotropic drug-response elements (PDREs), were found in the promoter region of several genes involved in lipid metabolism, such as IPT1, LAC1, SUR2 or PDR16 (van den Hazel et al., 1999; Hallstrom et al., 2001; Kolaczkowski et al., 2004). In particular, multiple steps in the sphingolipid biosynthetic pathway were found to respond to Pdr1p/Pdr3p regulation (Hallstrom et al., 2001; Kolaczkowski et al., 2004). These observations suggest that RTA1 is a member of the PDR regulatory system, closely associated with the properties of plasma membranes. The regulation of expression of the RTA1 gene combined with subcellular localization and turnover of its protein product was investigated here in detail to shed more light on its cellular function.

Materials and methods

Strains, media and β-galactosidase assay

The S. cerevisiae strains used in this study are listed in Table 1. The JD52 strain (Ghislain et al., 1996) was used for Rta1p purification. Strains were grown at 28 °C in YPD (2% yeast extract, 2% peptone and 2% glucose), YGal (2% yeast extract, 2% peptone and 2% galactose) or minimal medium (0.7% yeast nitrogen base and 0.1% casamino acids) and either 2% glucose (SD) or 2% galactose (SGal) supplemented with appropriate amino acids as required. Solid media contained 2% (w/v) Bacto agar. Yeast strains were transformed by the lithium acetate method (Chen et al., 1992). β-Galactosidase assays on lysates were performed as described previously with o-nitrophenyl-β-d-galactopyranoside (ONPG) as a substrate (Rose et al., 1981). More sensitive measurements using permeabilized cells and 4-methylumbelliferyl β-d-galactopyranoside (MUG) (Sigma Aldrich) were taken using the Fluoroscan Ascent FL microplate spectrofluorometer (Thermo Electron Corporation) according to previously published procedures (Guarente, 1983; Gee et al., 1999). All assays were carried out with at least four independent transformants, and β-galactosidase activity was expressed in relative fluorescence units (RFU) normalized to optical density of cell's suspensions used in the reaction.

View this table:

Saccharomyces cerevisiae strains used in this study

StrainGenotypeReference or source
FY1679-28CMATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 GAL2 +C. Fairhead
FY1679-28C/TDECMATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 GAL2+ pdr1Δ::TRP1 pdr3Δ::HIS3Delaveau et al. (1994)
FY1-3MATaura3-52 his3Δ200 leu2Δ1 trp1-Δ63 GAL2+ PDR1-3Kolaczkowski et al. (1998)
FYΔ125MATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 GAL2+ pdr5-Δ1::hisG snq2-Δ1::hisG yor1-Δ1::hisGKolaczkowski et al. (1998)
FYΔrta1MATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 GAL2+ RTA1::kanMXThis study
FY1-3Δ125MATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 GAL2+ PDR1-3 pdr5-Δ1::hisG snq2-Δ1::hisG yor1-Δ1::hisGKolaczkowski et al. (2009)
FY1-3 Δ125 Δrta1MATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 GAL2+ PDR1-3 pdr5-Δ1::hisG snq2-Δ1::hisG yor1-Δ1::hisG RTA1::kanMXThis study
FY1-3 Δ125 Δrta1
PDR5-RTA1-His10MATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 PDR1-3 GAL2+ pdr5-Δ1::hisG snq2-Δ1::hisG yor1-Δ1::hisG RTA1::kanMX PPDR5::RTA1This study
FY1-3 RTA1-HAMATaura3-52 his3Δ200 leu2Δ1 trp1-Δ63 GAL2+ PDR1-3 RTA1-HAThis study
FY1-3 Δ125 Δrta1
RT-HAMATaura3-52 his3-Δ200 leu2-Δ1 trp1-Δ63 GAL2+ PDR1-3 pdr5-Δ1::hisG snq2-Δ1::hisG yor1-Δ1::hisG RTA1-HAThis study
BY 4741MATa his3Δ1 leu2Δ0 met15 Δ0 ura3Δ0Brachmann et al. (1998)
Δ mot3 BYMATa his3Δ1 leu2Δ0 met15 Δ0 ura3Δ0, MOT3::kanMX EUROSCARF
Δ rox1 BYMATa his3Δ1 leu2Δ0 met15 Δ0 ura3Δ0 ROX1::kanMX EUROSCARF
Δ mot3Δ rox1 BYMATa his3Δ1 leu2Δ0 met15 Δ0 ura3Δ0 ROX1::kanMX MOT3::natMXThis study
Δ rsb1 BYMATa his3Δ1 leu2Δ0 met15 Δ0 ura3Δ0 RSB1::kanMXEUROSCARF
Δ tup1 BYMATa his3Δ1 leu2Δ0 met15 Δ0 ura3Δ0 TUP1::kanMXEUROSCARF
Δ upc2 BYMATa his3Δ1 leu2Δ0 met15 Δ0 ura3Δ0 UPC2::kanMXEUROSCARF
JD52MATaura3-52 his3-Δ200 leu2-3, 112 trp1-Δ63 lys2-801Ghislain et al. (1996)
BY4742MATα his3Δ1 ura3Δ0 leu2Δ0 lys2Δ0Brachmann et al. (1998)
  • EUROSCARF (Frankfurt am Main, Germany).

To generate the rta1::kanMX6 allele, the plasmid pFA6a-kanMX6 was used as a template to amplify the kanMX6 cassette (Wach, 1996). The forward primer consisted of 40 nucleotides of the RTA1 sequence upstream of the translation initiating AUG codon, followed by 20 nucleotides from the kanMX6 cassette upstream of the KanR coding region, while the reverse primer consisted of 40 nucleotides of the RTA1 sequence downstream of the translation stop codon, followed by 20 nucleotides from the kanMX6 cassette downstream of the KanR coding region. This cassette was transformed into S. cerevisiae. Transformants were selected on YPD plates containing 300 μg mL−1 of geneticin G-418 sulphate (GIBCO) and screened by PCR analysis.

The Rta1-HA derivative containing the RTA1 gene tagged with the 3HA epitope was constructed by the integration of the PCR-amplified tagging cassette into the genome. The pFA6a-3HA-HisMX6 plasmid was used as a template along with forward primer consisted of 42 nucleotides upstream of the STOP codon, followed by 20 nucleotides from the 3HA-HisMX6 cassette and reverse primer containing 40 nucleotides downstream of the last coding codon of RTA1 followed by 20 nucleotides of the 3HA-HisMX6 cassette. His+ transformants were screened by PCR.

Escherichia coli strains DH5α, MC1061 or TOP 10 (Invitrogen) were used for plasmid amplification. All plasmids and constructs generated in this study were verified by DNA sequencing.

Growth inhibition assays

Resistance to drugs was determined by spotting cell suspensions on plates containing increasing drug concentrations (Balzi et al., 1994; Dherbomez et al., 1995), and PHS resistance was assessed on plates containing varying concentrations of the compound (Sigma) as well as 0.05% Tergitol (type NP-40, Sigma). Liquid cultures were incubated in the presence of 20 μM PHS for 5 h before the cells were processed for β-galactosidase assays. Resistance to 7-ACH (Department of Biochemistry, University of Environmental and Life Science, Wrocław, Poland and Laboratoire de Synthèse et Etude de Substances Naturelles à Activité Biologique, IMRN INRA 1111, Université d'Aix-Marseille, France) was tested on YPD plates containing 20 μg mL−1 of ergosterol, 0.5% Tween 80 and varying concentration of the drug. Plates were incubated at 30 °C for 2 days (normoxia) or in a hypoxic chamber (BBL™ GasPak™Anaerobic Systems) at 30 °C for 4 days. Alternatively, microdilution assays in 96-well plates were performed as described earlier (Kolaczkowski et al., 2009).


The RTA1–lacZ gene fusion contains PCR-amplified promoter sequences (from −969 to +3 relative to the ATG codon) of RTA1 fused to E. coli lacZ on the low copy number plasmid pSEYC102MK (Kolaczkowski et al., 2009). The following primers were used for PCR amplification: 5′primer: ctatctcccgggcagcaggccac and 3′primer gacgggatcctttgccatgttcgaagtg (restriction enzyme sites are in boldface). Deletion derivatives of the RTA1 promoter were constructed as follows: A4 variant (−702 to +3) was generated by the removal of EcoRI fragment from the RTA1 promoter region. To obtain B4 (−503 to +3), RTA1–lacZ was digested with SmaI and MssI, and the shortened RTA1-pSEY102MK backbone was then religated (fragment encompassing from −969 bp to −502 bp was removed). To construct F8 (−418 to +3), E4 (−307 to +3) and D5 (−265 to +3), PCR-amplified RTA1 promoter sequences were digested with SspI/BamHI (F8), Bsh1236I/BamHI (E4) or HincII/BamHI (D5) and inserted into SmaI/BamHI sites of pSEYC102MK vector. The PDR3–lacZ fusion gene was described previously (Hallstrom & Moye-Rowley, 2000). The 0.8-kb RSB1 promoter fragment was amplified from genomic DNA using primers BERSB1 (ccagaattcaaaccaacaaaactctcc) and 2BBRSB1 (ggaggatccttggacattttgaatttc). The EcoRI/BamHI-digested fragment was cloned into pSEYC102 to give the RSB1–lacZ plasmid.

Overlap extension PCR mutagenesis was used to alter the PDRE in the RTA1 promoter region (−500 to +8). The first PCR was performed with wild-type genomic DNA and the mutagenic primer 1mPDRTA1 (ttgctaaagatgagttaagttgatctttggtgaaaaga, mutant bases in boldface) and the BBRTA1 primer. The second reaction was performed with the mutagenic primer 2mPDRTA1 (tcttttcaccaaagatcaacttaactcatctttagcaa) and the BERTA1 primer. The PCR products were purified, mixed and subjected to a second amplification using the two outside primers BBRTA1 and BERTA1. The resulting PCR product was then cloned in pSEYC102 (Emr et al., 1986) as a BamHI/EcoRI fragment, yielding the mPDRERTA1*–lacZ fusion gene. The wild-type RTA1 promoter region was similarly cloned in pSEYC102 to give the RTA1*–lacZ fusion gene.

All RTA1 expression plasmids were generated by PCR amplification using FY wt genomic DNA as a template. The multicopy plasmid pwRTA1, with the RTA1 coding region under its own promoter, was obtained by ligating an EcoRI/XhoI-digested 1.4-kb PCR product amplified with primers BERTA1 (ccagaattctttacaagccacaataag) and XRTA1 (ccactcgagttattcgctcttgaagcgag) and the PRS424 plasmid containing the TRP1 gene as a selectable marker (Christianson et al., 1992). The same RTA1 fragment was also cloned into the URA3-marked pRS426 plasmid (Christianson et al., 1992) to give pRTA1. In order to place RTA1 under the control of the GAL1 promoter, its coding region was PCR-amplified with BRTA1 (ccaggatccatggcaaaagacggcttcgag) and XRTA1 primers. The resulting PCR product was cut with BamHI and XhoI and cloned into the pRS316GAL1 vector (Mumberg et al., 1994) yielding pGALRTA1. A 2-kb DNA fragment containing the RSB1 promoter and coding region was amplified by PCR using primers SpeIRSB1 (ccaactagtaaaccaacaaaactctcc) and XhoRSB1 (ccactcgagctaaagtttagccttcttttt). The SpeI/XhoI-digested PCR fragment was inserted into pRS424 to give the pRSB1 recombinant plasmid. These constructs were used for drug resistance assays.

C-terminally His6-tagged versions of Rta1p were generated using primers BERTA1 and XHRTA1 (ccactcgagttatccatctgtatcgtgatggtgaccttcgctctttgaagcgagatcttc). The EcoRI/XhoI-digested PCR fragment was cloned into EcoRI and XhoI-cut pRS426 yielding URA3-marked puRta1-His6. The EcoRI/XhoI RTA1 fragment from puRta1-His6 was recloned into pRS424 to give TRP1-marked pwRta1-His6. The PDRE in the RTA1 promoter region from puRta1-His6 was altered by PCR mutagenesis using the mutagenic 1mPDRTA1 and 2mPDRTA1 primers. The resulting plasmid was called pumRta1-His6. To express a His6-tagged version of Rta1p under the control of the GAL1 promoter, the PCR product amplified with primers BRTA1 (ccaggatccatggcaaaagacggcttcgag) and XHRTA1 was cut with BamHI and XhoI and cloned into the GAL1 promoter–containing plasmid pRS316GAL1, resulting in puGALRta1-His6. To express a FLAG-tagged version of Rta1p under the control of the GAL1 promoter, a PCR fragment amplified with primers BRTA1 and RTA1flagXho was digested with BamHI and XhoI and inserted into pRS316GAL1 to give pGALRta1-FLAG. Plasmid pRta1-FLAG expressed the tagged form under the control of the RTA1 promoter. For the construction of the C-terminal GFP fusion, the RTA1 coding region was amplified using primers BRTA1 and RTA1Cla1 (ccaatcgatttcgctctttgaagcgag). The BamHI/ClaI-digested amplicon was then cloned into the pUG35 vector (Niedenthal et al., 1996) yielding pRta1–GFP.

Fluorescence microscopy

The FY wt strain was transformed with plasmids expressing the Rta1–GFP protein fusion from the MET25 promoter. Exponentially growing cells were visualized for GFP fluorescence using a DMR (Leica) microscope equipped with a mercury HBO 103 w/2 lamp (OSRAM) and a GFP filter (excitation 470/40 nm and emission 525/50 nm, Leica). Images were captured using a DC200 camera and analysed by the Leica IM1000 software. Nomarski optics was used to visualize the cells with the transmitted light.

Preparation of plasma membranes and protein analysis

Exponentially growing JD52 cells expressing Rta1-His6 under the control of the GAL1 promoter (300 OD600) were harvested by centrifugation at 4 °C, washed with water and resuspended in an equal volume of MBS buffer (25 mM Tris, 2 mM MgCl2 and 250 mM sorbitol, pH 7.5) supplemented with 1 mM phenylmethylsulphonyl fluoride (PMSF) and 2 μg mL−1 each of leupeptin, aprotinin, antipain, pepstatin and chymostatin. Plasma membranes were prepared as previously described (Dufour et al., 1988).

Protein concentration was determined by the Bradford method (Bradford, 1976). Proteins were separated on 10% or 12% SDS-polyacrylamide gels and stained with colloidal Coomassie Blue (Serva Blue G-250 no35050) or subjected to immunological analysis. For Western blotting, proteins were separated by SDS-PAGE, transferred to nitrocellulose and probed with anti-GFP antibodies (1:5000, Roche) or anti-HA antibodies (1:1000, Sigma). The membrane was stripped with 0.5 M NaOH and reprobed with antibodies against Pma1p (1 : 15 000) (Capieaux et al., 1993) or Cdc48p (1 : 10 000) (Decottignies et al., 2004). Protein–antibody complexes were detected using an enhanced chemiluminescence system (Roche).

For Western blot analysis of whole-cell extracts, 5 OD600 units of cells were harvested by centrifugation at 4 °C, washed with water and resuspended in 200 μL of lysis buffer (25 mM Tris pH7.5 and 5 mM EDTA) supplemented with 1 mM PMSF and 2 μg mL−1 each of leupeptin, aprotinin, antipain, pepstatin and chymostatin. All subsequent steps were performed at 4 °C. Glass beads (200 μL) were added and the cells were broken by agitation for 4 min with a 15-s pause on ice every 30 s, and then 100 μL of 3× Laemmli buffer was added; the mixture was incubated for 15 min at 56 °C and centrifuged to remove the cell debris and the glass beads.

Pulse-chase experiments

Ten millilitres of exponentially growing cells (OD600 of 0.5–1) in SGal medium was harvested, resuspended in 400 μL of SGal medium and labelled for 10 min at 28 °C with 15–20 μL of Pro-Mix L-[35S] labelling mix (Amersham Pharmacia Biotech). Then, the cells were centrifuged, resuspended in 400 μL of growth medium containing 0.2 mg mL−1 of cycloheximide and 10 mM methionine and incubated at 28 °C; 100-μL samples were withdrawn at different time points, and then glass beads were added along with 400 μL of lysis buffer (50 mM Hepes and 5 mM EDTA, pH 7.5) supplemented with 1 mM PMSF and 2 μg mL−1 each of leupeptin, aprotinin, antipain, pepstatin and chymostatin. Cells were disrupted by agitation for 3 min, with intermittent cooling on ice, and then 20 μL of 10% SDS was added, and the mixture was further agitated for 30–40 s before centrifugation. The soluble material was added to four volumes (1.6 mL) of lysis buffer containing 1% Triton X-100 and 150 mM NaCl and incubated with 5 μL of anti-FLAG agarose beads for 2 h at 4 °C with agitation. The beads were then washed three times with lysis buffer containing 1% Triton X-100, 150 mM NaCl and 0.1% SDS, resuspended in 20 μL of 2× Laemmli buffer and heated to 56 °C for 15 min. After centrifugation at 20 000 g for 5 min, the solubilized material was subjected to 12% SDS-PAGE. The half-life of Rta1p was determined from a plot of the number of pixels in each band as a function of time (Ghislain et al., 1996) by phosphorimage scanning of dried gels (Pharos FX Molecular Imager System; Bio-Rad).


Rta1p is a plasma membrane protein

A low copy plasmid expressing a C-terminally His6-tagged derivative of Rta1p under the control of the GAL1 promoter (plasmid pGALRta1-His6) was introduced into yeast cells (wild-type strain JD52), and whole-cell extracts from appropriate transformants were analysed by immunoblotting with anti-His5 antibodies. As shown in Fig. 1a, no Rta1-His6 band was detected in cells grown in glucose-containing media, whereas a 33-kDa band corresponding to the expected size of Rta1-His6 was detected in cells grown in galactose-containing media. The Rta1-His6 form was functional as determined by its ability to support the cell growth at 10 μM 7-ACH as compared to the control strain transformed with an empty vector, which failed to grow at that concentration (Fig. 2a).


Rta1p is a plasma membrane protein. (a) Detection of the Rta1–His6 fusion protein expressed from the GAL1 promoter. Wild-type JD52 cells were transformed with low copy plasmids lacking (−) or containing a C-terminally tagged form of Rta1p under the control of the GAL1 promoter. Whole-cell extracts (WCE) from transformants grown in glucose- or galactose-containing media were fractionated by SDS-PAGE and analysed by immunoblotting with anti-His5 antibodies. The asterisk indicates an 120-kDa band that cross-reacted with the anti-His5 serum. (b) Rta1-His6 is enriched in a plasma membrane fraction. Lysates from JD52 cells expressing Rta1-His6 were fractionated by a series of centrifugation steps into a cytosolic fraction (S15/40, lane 3) and pellet fractions P6 (lane 1) and P15/40 (lane 2). The P15/40 fraction was processed to separate mitochondria (lane 4) from plasma membranes (lane 5). Various fractions were analysed by Western blotting using anti-His5 antibodies. (c) Fluorescence microscopy of cells expressing Rta1-GFP. FY1679-28C cells expressing Rta1–GFP or GFP were examined by fluorescence microscopy (top panels) and light microscopy (bottom panels). The bars correspond to 5 μm. (d) Detection of the Rta1–GFP fusion protein expressed from the MET25 promoter Whole-cell extracts from FY wt cells expressing GFP alone or Rta1–GFP were analysed by Western blotting using antibodies against GFP or the plasma membrane H+-ATPase (Pma1p) as a control for equal protein loading.


Mutation of the PDRE in the RTA1 promoter region impairs resistance to 7-ACH. Growth assay of FY wt cells expressing various Rta1p forms from the GAL1 promoter on a low copy plasmid (a), the MET25 promoter on a high copy plasmid (b), the RTA1 promoter or a PDRE mutated (mPDRE) form on a high copy plasmid (c). Cells transformed with an empty plasmid (–) were used as a control. The cells were serially diluted and spotted on glucose (Glc)- and galactose (Gal)-containing media supplemented with the indicated 7-ACH concentrations. The plates were photographed after 3 days of incubation at 28 °C. Note slower cell growth on galactose media.

To determine the subcellular localization of Rta1-His6, cell lysates were fractionated by centrifugation into a nuclei-enriched (P6) pellet, a supernatant (S15/40) consisting of microsomes and cytosolic proteins and a pellet (P15/40) enriched in plasma membranes, in which the presence of Rta1-His6 was revealed by Western blot analysis (Fig. 1b, lane 2). Further purification of plasma membrane proteins by removing mitochondrial contaminants resulted in a fourfold increase in the Rta1-His6 band intensity (lane 5). The weak signal associated with mitochondrial fraction (lane 4) results from the loss of some plasma membrane proteins in that fraction during purification.

The plasma membrane localization of Rta1p was further confirmed using a C-terminal GFP fusion and fluorescence microscopy. A low copy plasmid expressing the Rta1–GFP protein fusion under the control of the MET25 promoter was introduced into wild-type FY1679-28C (FY wt) cells, and appropriate transformants were selected on the basis of increased 7-ACH resistance (Fig. 2b). As shown in Fig. 1c, Rta1-GFP localized to a single compartment corresponding to the plasma membrane. Whole-cell extracts were analysed by Western blotting with anti-GFP antibodies. Antibodies against the plasma membrane H+-ATPase (Pma1p) were also used to confirm that equal amounts of total protein were loaded. As shown in the left panel of Fig. 1d, the cells expressing Rta1-GFP gave a 63-kDa band (lane 3), the size of which corresponds to the calculated size of the fusion protein. The 27-kDa band detected in the extract from cells containing an empty plasmid (pUG35) corresponds to GFP alone (lane 1).

Rta1p is a short-lived protein

Metabolic stability of Rta1p was analysed in galactose-depleted JD52 cells expressing the Rta1-His6 fusion from the GAL1 promoter. Cells grown in the presence of galactose were split and further cultivated in galactose- or glucose-containing media. At 75-min intervals, an equal amount of cells were lysed and a crude plasma membrane (P15/40) fraction was purified from the lysate and separated by SDS-PAGE, and the relative amount of Rta1-His6 was analysed by immunoblotting. As shown in Fig. 3a, the Rta1-His6 band decreased after 1 h 30 min of incubation in glucose and was undetectable after 4 h. In contrast, the amount of Rta1p-His6 remained at the same level during the 4-h incubation in the presence of galactose. From this experiment, we conclude that Rta1-His6 is a moderately short-lived protein, with a half-life of ~ 60 min.


Rta1p is a short-lived protein. (a) Analysis of the Rta1-His6 turnover. The wild-type JD52 strain was transformed with the low copy pGALRta1-His6 plasmid expressing Rta1-His6 under the control of the GAL1 promoter and grown in Sgal medium. The cells were collected and transferred to minimal medium containing glucose or galactose. At the indicated time points, samples were withdrawn for the preparation of plasma membrane–enriched fractions (P15/40). Samples were fractionated by SDS-PAGE and analysed by immunoblotting with anti-His5 antibodies. Fifty micrograms of proteins isolated from the cells grown in galactose-containing medium was loaded. In order to correct for cell division and arrest of Rta1-His6 synthesis, the following amount of proteins isolated from cells grown in the presence of glucose were loaded: 50 μg (0 min), 75 μg (90 min), 125 μg (180 min) and 175 μg (240 min). (b) Pulse-chase analysis of Rta1-FLAG degradation. BY wt cells expressing Rta1-FLAG under the control of the GAL1 promoter on a low copy plasmid were labelled with 35S-methionine for 10 min (Sgal medium), then, after 0, 20, 40 or 60 min of incubation in the presence of an excess of unlabelled methionine and cycloheximide, the cells were lysed, processed for immunoprecipitation with anti-FLAG agarose beads and fractionated by SDS-PAGE, and band intensity was quantified by phosphorimaging. The anti-FLAG antibodies cross-reacted with an unknown low molecular weight band (marked with an asterisk).

The metabolic instability of Rta1p was confirmed by pulse-chase analysis of wild-type BY4742 (BY wt) cells expressing a C-terminally FLAG-tagged form under the control of the GAL1 promoter. As shown in Fig. 3b, Rta1-FLAG is rapidly degraded (half-life of 30 min). The difference with the previously estimated half-life of 60 min is explained by the fact that protein synthesis was not inhibited by cycloheximide in the galactose-depletion experiments.

The RTA1 promoter responds to the status of the major repressors of hypoxic genes and activators of pleiotropic drug resistance

Genomewide analysis suggested RTA1 as a target for the hyperactive form of master regulator of the PDR network, PDR1-3 (DeRisi et al., 2000). However, no other regulators of RTA1 expression were identified. Therefore, we searched for candidate transcription factors using the YEAst Search for Transcriptional Regulators And Consensus Tracking (YEASTRACT), (http://www.yeastract.com) (Monteiro et al., 2008), as well as Mining Yeast Binding Sites databases (mybshttp://cg1.iis.sinica.edu.tw/~mybs/index.php). The 5′ UTR region encompassing 969 nucleotides upstream of the ATG codon was examined (Fig. 4). This analysis revealed a number of potential regulatory elements for transcription factors involved in yeast response to different stress stimuli (Yap1p, Mns2/4p, Yrr1p, Gcn4p, Gln3p, Pdr1/3p, Mot3p and Rox1p), as well as in the regulation of flocculation, pseudohyphal and invasive growth (Flo8p, Nrg1p and Tec1p).


Putative regulatory elements in the RTA1 promoter. Position of binding motifs relative to the translation start site is shown. Predicted regulatory elements were identified by the use of yeastract (http://www.yeastract.com) and mybs (http://cg1.iis.sinica.edu.tw/~mybs/index.php) databases.

The evolutionary conservation indicating functional importance of some of these sites was observed in a multiple sequence alignment of promoter regions of RTA1 orthologs from related yeast species (S. paradoxus, S. mikatae and S. bayanus). Strikingly among these cross-species conserved sites, there were multiple varying consensus binding sites for Mot3p spread along the entire promoter and two sites for Rox1p binding. The close proximity of the putative Rox1p binding sites to the Mot3p sites (Fig. 4) was in line with the synergistic effect of these global repressors of hypoxic genes observed previously by others (Sertil et al., 2003; Klinkenberg et al., 2005). Additionally, one conserved binding motif for the transcriptional activator Upc2p (Uptake Control) was identified. Upc2p was previously shown to control basal and induced expression of genes encoding enzymes of the ergosterol biosynthesis pathway (Vik & Rine, 2001; Davies & Rine, 2006) and other genes inducible by hypoxia (Abramova et al., 2001; Kwast et al., 2002).

The response of the RTA1 promoter to the loss of the transcription factors identified by the bioinformatic approach was analysed by measurements of lacZ reporter activity expressed from the RTA1 promoter on a low copy plasmid introduced in a set of isogenic yeast strains (derivatives of BY4741). Activation of the RTA1 promoter was observed in cells deleted for MOT3 or ROX1 (Fig. 5a), and a higher ninefold induction was observed in a double Δmot3 Δrox1 mutated strain. The deletion of the MOT3 and ROX1 genes had no effect on the expression of the PDR3–lacZ gene fusion used as a control (Fig. 5b). Because Rox1p exerts its inhibitory action by recruitment of the general repressor Tup1–Ssn6 complex (Balasubramanian et al., 1993), a tup1Δ strain was included in the analysis. A 10-fold increase in the RTA1-driven β-galactosidase activity was observed in the null mutant strain (Fig. 5a). These results indicate that Mot3p and Rox1p act in concert to repress RTA1 expression under standard laboratory conditions with the involvement of Tup1p. On the contrary, a reduction in the RTA1 promoter activity was observed upon the deletion of UPC2 (Fig. 5a).


The RTA1 promoter is responsive to different regulators associated with hypoxic conditions. (a) A series of BY strains bearing deletions in the MOT3, TUP1, ROX1 and UPC2 genes were transformed with a low copy plasmid containing the RTA1 promoter fused to the lacZ coding region. β-Galactosidase activity was determined on at least four independent transformants, and the mean values including the standard deviations are indicated. (b) Mot3p and Rox1p act in a synergistic manner on the RTA1 promoter. RTA1–lacZ reporter constructs were introduced into isogenic BY wild-type or mot3Δrox1Δ cells, and the β-galactosidase activity was measured. The unresponsive PDR3-lacZ was used as a negative control. (c) The RTA1 promoter is responsive to the major regulators and effectors of the pleiotropic drug resistance network. FY derivatives carrying the PDR1-3 hyperactive mutation or triple deletion in the major multidrug ABC transporter genes, PDR5, SNQ2 and YOR1, were transformed with the RTA1–lacZ reporter. The β-galactosidase activity determined on at least four independent transformants is expressed as RFU normalized to optical density of cell suspensions used.

Examination of the RTA1 promoter region revealed the presence of a single conserved B-type PDRE (TCCGTGGA), located 194 nucleotides upstream of the initiator ATG codon. Therefore, the response of the RTA1 promoter to the hyperactive PDR1-3 allele was verified using the lacZ–RTA1 fusion gene. As expected, the constitutive PDR1 activation led to a fivefold increase in the activity of the reporter β-galactosidase (Fig. 5c). On the basis of previous mutagenesis analyses of PDRE determinants (Katzmann et al., 1994, 1999; Delahodde et al., 1995; Hallstrom & Moye-Rowley, 1998), the PDRE sequence in the RTA1 promoter was modified to TAAGTTGA. The loss of the PDRE resulted in reduced 7-ACH resistance as shown with the isogenic set of FY wt cells expressing Rta1-His6 under the control of endogenous RTA1 promoter or its mutated PDRE version (Fig. 2c).

The activation of Pdr1p in response to deletions of its major transcriptional target genes encoding PDR5, SNQ2 and YOR1 has recently been observed (Kolaczkowska et al., 2008). Interestingly, this triple deletion led to a similar level of RTA1 promoter induction as the hyperactive PDR1-3 mutant (Fig. 5c), suggesting that RTA1 activity is modulated by the status of the PDR network.

We did not find significant responses of the RTA1–lacZ reporter to the loss of genes encoding other candidate regulators under the conditions tested (data not shown).

Mapping of the RTA1 promoter

To better understand the transcriptional regulation of RTA1 and locate promoter regions important for its activity, a series of 5′ progressive deletions in the RTA1 promoter fused to the lacZ gene were assayed for the reporter activity. TUP1 (BY wt) and tup1Δ cells were transformed with low copy plasmid–based truncation mutants, and β-galactosidase activity was measured. RTA1lacZ fusion containing 969 bp of 5′-UTR directed the production of 196 RFU (of β-galactosidase activity per optical density unit) and 1235 RFU in tup1Δ (Fig. 6a). Progressive 5′ deletions up to −418 (variants A4, B4 and F8) led to a threefold increase in RTA1 promoter activity (537 RFU/OD, wt cells). In this region (−969 to −418), putative binding sites for repressors Mot3p and Nrg1p were found (see Fig. 4). Higher promoter activity was consistently observed in tup1Δ cells transformed with variants A4 and F8, both of them lacking Mot3p binding sites. Deletion of the next 85 nucleotides (−418 to −302, variant E4) decreased RTA1 promoter activity in TUP1 wild-type and tup1Δ mutant cells. One potential regulatory element for Upc2p activator was identified in this region (see Fig. 4). Further 5′ UTR truncation elevated promoter activity nearly thirteen-fold (construct D5, wild-type strain). A similar high level of activation was observed under de-repressed conditions in the tup1Δ cells.


Deletion mapping of the RTA1 promoter. (a) Schematic representations of the 5′ progressive deletions of the RTA1 promoter are shown. Deletion endpoint is marked with a number. ATG, start site of the RTA1 open reading frame. The RTA1-driven β-galactosidase activity was determined for each deletion variant in BY wild-type and tup1Δ cells. (b) BY wild-type and mot3Δ cells were transformed with RTA1lacZ or its deletion derivatives. The β-galactosidase activity was determined on at least four independent transformants, and the mean values including the standard deviations are indicated.

In summary, 5′ progressive deletions mapped a region of the RTA1 promoter extending up to 265 bp upstream of the transcription start site that conserved the activity of the proximal promoter under normal and de-repressed conditions (wt vs. tup1Δ) and regions responsive to repression I (−969 to −702), II (−503 to −418) and III (−302 to −265). Fragments I and II contain multiple Mot3p binding sites in the close proximity to putative Rox1p binding motifs (−382, −175). One of the mapped regions (−418 to −302) is likely to be involved in Upc2p-dependent activation.

The RTA1 promoter deletion variants were also introduced into a mot3Δ mutant strain. As shown in Fig. 6b, a region most responsive to Mot3p lies between −503 and −418 upstream of the transcription start site. This is in agreement with deletion mapping data and presence of the Mot3p binding site in this region.

An increased Rta1p activity associated with hypoxic condition

Activation of the RTA1 promoter in response to elimination of the global repressors of hypoxic genes indicated an elevated expression and activity under oxygen depletion. Therefore, we compared the level of RTA1-mediated 7-ACH resistance under normoxic and hypoxic conditions. Figure 7a illustrates that wild-type FY1679-28C (FY wt) cells were able to tolerate 7-ACH concentrations up to 2 μg mL−1 upon growth under oxygen-deficient conditions. This concentration is toxic to the wild-type cells grown in normoxia. Moreover, this effect was strictly dependent on the presence of RTA1, as its deletion led to marked hypersensitivity to 7-ACH in both the PDR1 wild-type and the PDR1-3 mutant strain backgrounds. Cells overexpressing Rta1p from the PDR5 promoter (Table 1) were able to grow at the highest 7-ACH concentration tested.


Resistance to 7-ACH is dependent on RTA1 under oxygen depletion. (a) Set of isogenic strains FY wt, FYΔrta1, FY1-3Δ125Δrta1 or FY1-3Δ125Δrta1 containing integrated version of the RTA1-his10 tag expressed from the PDR5 (PDR5-RTA1-his10) promoter were grown to mid-log phase in YPD media, and then serially diluted cells were spotted onto plates containing 20 μg mL−1 of ergosterol, 0.5% Tween 80 and 7-ACH. As a control plates without drug were used. Plates were incubated for 2 days at 30 °C under standard laboratory conditions (normoxia, right panel) or 4 days at 30 °C in a hypoxic chamber (hypoxia, left panel). (b) Rta1-HA protein level is elevated under hypoxia. Isogenic strains FY1-3RTA1-HA and FY1-3Δ125RTA1-HA (two independent clones) expressing RTA1 from the single genomic allele marked with the integrated 3xHA tag or the control FYΔRta1 were cultivated under normoxic or hypoxic conditions. Cell lysates prepared from the indicated strains were fractionated by SDS-PAGE (normoxia, 200 μg of proteins loaded; hypoxia, 100 μg proteins loaded) and analysed by Western blotting using anti-HA antibodies.

Importantly, oxygen depletion led to an increased production of Rta1 protein (Fig. 7b). RTA1 was expressed from its own promoter and a single genomic allele, marked with the integrated 3xHA tag. Detection of Rta1p, however, required de-repression of its endogenous promoter by hypoxia, combined with the presence of a hyperactivating PDR1-3 mutation. An even higher Rta1p level was observed when in addition the three major ABC transporters encoding genes PDR5, SNQ2 and YOR1 were deleted (FY1-3Δ125). Despite loading a double amount of proteins, the HA-tagged Rta1p was barely detectable in the extracts of cells grown under normoxic conditions (Fig. 7b); however, upon further fractionation, a strong signal was observed solely in the purified plasma membrane fraction (data not shown).

RTA1 overexpression increases cell tolerance to phytosphingosine

Rsb1p, a close homologue of Rta1p, has been shown to confer tolerance to phytosphingosine (PHS), a precursor of sphingolipids and mediator of stress-dependent growth arrest (Kihara & Igarashi, 2004; Panwar & Moye-Rowley, 2006). To verify whether the sequence similarity between Rta1p and Rsb1p reflects a functional overlap, we assessed the possible involvement of Rta1p in resistance to PHS. Trp FY wt cells were transformed with the URA3-marked plasmids pRTA1 (RTA1 promoter) and pGALRta1-His6 (GAL1 promoter), and appropriate transformants were examined for PHS sensitivity. The growth of tryptophan (Trp) auxotrophic strains is more strongly inhibited by PHS than the growth of Trp+ strains, as PHS reduces tryptophan uptake and starves cells for this amino acid (Skrzypek et al., 1998; Johnson et al., 2010). An increase in the RTA1 copy number was not sufficient to elevate PHS tolerance (data not shown). However, we found that cells overproducing Rta1-His6 from the GAL1 promoter tolerated PHS concentrations up to 80 μM on galactose-containing medium (Fig. 8). The wild-type strain transformed with a high copy plasmid containing RSB1 was unable to grow in the presence of 20 μM 7-ACH (data not shown), indicating that RTA1 and RSB1 are not functionally interchangeable.


Overexpression of RTA1 results in increased tolerance to PHS. FY wt strain was transformed with empty p316GAL vector or pGALRta1-His6 derivative expressing GAL1-driven His6-tagged version of Rta1p. The cells grown overnight in galactose-containing medium (SGal) were serially diluted in water and spotted on SGal plates supplemented with the indicated PHS concentrations. The plates were photographed after 4 days of incubation at 28 °C.

To determine whether an induction of the RTA1 promoter occurs upon PHS exposure, the PDR1 (FY wt) and PDR1-3 (FY1-3) strains were transformed with the RTA1*–lacZ fusion gene or the mutated mPDRERTA1*lacZ variant lacking the PDRE. We also used the RSB1–lacZ fusion gene to compare the expression levels between RTA1 and RSB1. β-Galactosidase activity was determined from transformants that were incubated in the presence or absence of 20 μM PHS for 5 h before preparation of the cell lysates. RTA1*–lacZ-dependent β-galactosidase activity of the PDR1 strain showed a fourfold increase in the presence of PHS, which was not impaired by the PDRE mutation (Table 2). The activity levels seen with the PDR1-3 strain increased twofold in the presence of PHS, and this stimulation was not affected by the PDRE mutation. By comparison, the RSB1–lacZ fusion gene was slightly responsive to PHS (Table 2). These results indicate that RTA1 transcription is induced in the presence of PHS and this activation seems to be independent of Pdr1p and the PDRE.

View this table:

PHS induction of RTA1-dependent β-galactosidase activity

Gene fusionStrainβ-Galactosidase activity(nmol min−1 mg−1)
0 μM PHS20 μM PHS
RTA1*–lacZPDR10.43 ± 0.051.78 ± 0.20
RTA1*–lacZPDR1-33.52 ± 0.186.3 ± 0.38
mPDRERTA1*–lacZPDR10.24 ± 0.030.64 ± 0.08
mPDRERTA1*–lacZPDR1-30.37 ± 0.080.52 ± 0.05
RSB1–lacZPDR16.6 ± 0.838.83 ± 1.93
RSB1–lacZPDR1-3109.08 ± 14.45120.68 ± 18.34
  • The lacZ fusion genes were introduced into PDR1 (FY wt) and PDR1-3 (FY1-3) cells. RTA1* contains the wild-type PDRE sequence, TCCGTGGA; mPDRERTA1*, the mutated TAAGTTGA sequence.

  • The β-galactosidase values represent the average for three independent clones ± standard deviation. Comparable results were obtained with clones from 3 (RSB1–lacZ) and 4 (RTA1*–lacZ and mPDRERTA1*–lacZ) independent transformations.


The RTA1 gene was isolated as a sole multicopy suppressor of the toxic effects of 7-ACH (Soustre et al., 1996). Neither overexpression nor deletion of other members of the Rta1-like family, RSB1 and RTM1, had an effect on 7-ACH tolerance (Ness & Aigle, 1995). RTA1 overexpression has no effect on porphyrin uptake, in contrast to the PUG1 gene, which is expressed under haeme- and oxygen-deficient conditions (Protchenko et al., 2008). These studies indicated that the Rta1-like family consists of members that are not functionally interchangeable, consistent with the moderate level of sequence conservation. Interestingly, we found that overexpression of RTA1 may result in increased cell tolerance to PHS, as shown for Rsb1p (Kihara & Igarashi, 2002; Panwar & Moye-Rowley, 2006), and that this sphingolipid precursor activated RTA1 transcription. PHS has been shown to mediate the heat stress–induced ubiquitin-dependent degradation of membrane proteins (Chung et al., 2000). We showed that Rta1p was a short-lived plasma membrane protein and that the rate of Rta1p degradation was not affected by PHS and in a sphingolipid synthesis–defective mutant strain (data not shown).

Several studies of sterol efflux in mammals have identified the involvement of ABC transporters (Bodzioch et al., 1999; Brooks-Wilson et al., 1999; Rust et al., 1999; Berge et al., 2000). Although no yeast homologues have yet been identified, it is unlikely that Rta1p is a transporter involved in 7-ACH efflux, as the sterol profiles of aminocholesterol-treated strains containing or lacking RTA1 on a multicopy plasmid are comparable (Soustre et al., 1996). The lack of an ATP-binding motif is also an argument against Rta1p being an efflux pump. Predicted topology of Rta1p resembles rather those of 7-transmembrane receptor family (RbDe program, http://icb.med.cornell.edu/crt/RbDe/RbDe.html; Griffin, http://griffin.cbrc/jp), acting in concert with G proteins. Thus, similar to Rsb1p (Johnson et al., 2010), Rta1p could be a potential member of a family of G-protein-coupled receptors, but functioning under defined stress conditions.

To get more insight into its transcriptional regulation, we investigated in detail the RTA1 promoter sequence. Previous studies as well as our data indicate that the RTA1 promoter responds to different stress conditions, including high hydrostatic pressure (Iwahashi et al., 2005) and cytotoxic stress (Iwahashi et al., 2007). We found that two global repressors of hypoxic genes, Mot3p and Rox1p, control RTA1 promoter activity, and we have mapped regions responsive to repression. Both regulators were shown to repress their targets during aerobic growth (Abramova et al., 2001; Cohen et al., 2001). They could act in combination leading to a synergistic repression of the hypoxic genes (Abramova et al., 2001; Klinkenberg et al., 2005). In this study, double deletion of MOT3 and ROX1 led to higher RTA1 promoter induction than removal of only MOT3, indicating synergistic effect of these single loci on the regulation they exert on RTA1. Along with RTA1 promoter analysis, we have shown clear dependence of overproduced Rta1p-mediated resistance to 7-ACH under hypoxic conditions. These results are in line and largely extend the previous observation that levels of RTA1 transcript increased upon oxygen depletion (Protchenko et al., 2008).

DNA-binding protein Rox1p has been shown to recruit the Tup1p–Ssn6p general repressor complex (Balasubramanian et al., 1993) to exert its function. If Rox1p acted in concert with Tup1p–Ssn6p, loss of either the repressor or one of the components of Tup1p–Ssn6p would result in de-repression and activation of the RTA1 promoter. As we show, the deletion of TUP1 led to a 10-fold increase in the RTA1 promoter activity. In line with these data, we observed elevated activity of almost all deletion mutants (apart E4 variant) in tup1Δ mutant cells.

Deletion of most of the 5′UTR up to 265 upstream of the ATG codon (variant E4) led to a high increase in RTA1 promoter activity irrespective of the genetic background tested (wild type, mot3Δ or tup1Δ). Sequence analysis of this region identified a putative Rox1p binding site (−267). Its deletion could result in the loss of Tup1p-dependent repression and constitutive induction similar to that obtained by removal of TUP1. However, one cannot rule out the possibility that deletion of most of the promoter sequences containing multiple binding sites for repressors might lead to its Rox1p, Tup1p-independent activation and state of de-repression.

Apart from Rox1p and Mo3p, the RTA1 promoter was found to contain an UPC2 responsive element. The UPC2 gene encodes a Zn(II)2Cys6 transcription factor that binds to sterol regulatory elements in the promoter region of sterol biosynthetic genes as well as to ABC transporter genes PDR11 and AUS1, which mediate sterol uptake when sterol biosynthesis is compromised, for example, by anaerobic growth or by mutations impairing haeme synthesis (Wilcox et al., 2002). In this study, we show that the RTA1 promoter was responsive to UPC2 as revealed by reduced RTA1-driven β-galactosidase activity in upc2Δ cells. Inspection of the RTA1 promoter sequence together with 5′ progressive deletions identified a region (−418 to −302) responsive to activation and containing phylogenetically conserved Upc2p regulatory element (TCGTTTAG at position −333; see Fig. 4).

Examination of the RTA1 promoter region revealed the presence of a binding site for Pdr1p and Pdr3p and its mutation resulted in decreased 7-ACH resistance. Increased levels of RTA1 mRNA were found in mutant cells containing the PDR1-3 gain-of-function allele (DeRisi et al., 2000), and deletion of PDR5, SNQ2 and YOR1 led to increased RTA1 promoter activity. The loss of these plasma membrane ABC transporters induces a compensatory activation of Pdr1p target genes, via unknown regulatory mechanisms (Kolaczkowska et al., 2008). These are likely to be related to proper membrane function as lipid derivatives and lipid-like molecules were identified among substrates of Pdr5p (Kean et al., 1997; Decottignies et al., 1998; Kolaczkowski et al., 1998). These signals have previously been shown to induce Pdr1p-mediated induction of RSB1 expression and PHS resistance (Kihara & Igarashi, 2002).

In conclusion, hypoxic-mediated de-repression from Rox1p, Mot3p, Tup1p inhibition, Upc2 activation of sterol biosynthesis genes and the Pdr1p-dependent response to cytotoxic drugs induce regulatory signals converging at the promoter of the RTA1 gene. A common function for these complex gene networks is the prompt regulation of the permeability barrier to sustain defined stress conditions. Further studies are required to obtain a better understanding of the mechanism of action of these regulators on Rta1p expression and to elucidate the biochemical activity of this 7-transmembrane segments protein.


This work was supported by the Polish Ministry of Science and Higher Education (N N303 026537) and the Belgian National Found for Scientific Research (FSR-FNRS). Myriam Manente was a recipient of a FSR (Université catholique de Louvain) doctoral fellowship, and Donata Wawrzycka was a recipient of a FSR-FNRS (Belgium) postdoctoral fellowship. We thank Marie Lorfèvre (Université catholique de Louvain, Louvain-la-Neuve, Belgium) for help in generating gene constructs; Prof. Y. Letourneux, J.-M. Brunel (Université d'Aix-Marseille, France) and Dr M. Anioł (Department of Biochemistry, University of Environmental and Life Science, Wrocław) for synthesizing 7-ACH; and Tadeusz Lezniak for the construction of the B4 deletion variant.

Author's contribution

A.K. and M.M. contributed equally to this work.


  • Editor: Guenther Daum


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