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Identification of RCN1 and RSA3 as ethanol-tolerant genes in Saccharomyces cerevisiae using a high copy barcoded library

Michael J. Anderson, Sarah L. Barker, Charlie Boone, Vivien Measday
DOI: http://dx.doi.org/10.1111/j.1567-1364.2011.00762.x 48-60 First published online: 1 February 2012


Saccharomyces cerevisiae (S. cerevisiae) encounters a multitude of stresses during industrial processes such as wine fermentation including ethanol toxicity. High levels of ethanol reduce the viability of yeast and may prevent completion of fermentation. The identification of ethanol-tolerant genes is important for creating stress-resistant industrial yeast, and S. cerevisiae genomic resources have been utilized for this purpose. We have employed a molecular barcoded yeast open reading frame (MoBY-ORF) high copy plasmid library to identify ethanol-tolerant genes in both the S. cerevisiae S288C laboratory and M2 wine strains. We find that increased dosage of either RCN1 or RSA3 improves tolerance of S288C and M2 to toxic levels of ethanol. RCN1 is a regulator of calcineurin, whereas RSA3 has a role in ribosome maturation. Additional fitness advantages conferred upon overproduction of RCN1 and RSA3 include increased resistance to cell wall degradation, heat, osmotic and oxidative stress. We find that the M2 wine yeast strain is generally more tolerant of stress than S288C with the exception of translation inhibition, which affects M2 growth more severely than S288C. We conclude that regulation of ribosome biogenesis and ultimately translation is a critical factor for S. cerevisiae survival during industrial-related environmental stress.

  • ethanol tolerance
  • environmental stress
  • wine yeast
  • ribosome biogenesis
  • calcineurin


The budding yeast Saccharomyces cerevisiae has evolved to survive in a variety of niches and thus has specialized functions within each of these environments. For example, S. cerevisiae is commonly used both as a model organism for basic research of biological processes in eukaryotic cells and as an industrial microorganism for the production of biofuel, food and beverage products. Whilst laboratory strains of S. cerevisiae have been selected for ease of genetic manipulation and consistent growth in nutrient-rich media, industrial yeast strains have been selected for their ability to perform well in a stressful fermentation environment (Attfield, 1997; Bauer & Pretorius, 2000).

Throughout commercial production of active dry strains, S. cerevisiae is exposed to osmotic and oxidative stress, nutrient limitation and desiccation. After rehydration, wine yeasts are exposed to osmotic and organic acid stress upon inoculation into grape must, which may also contain antimicrobial agents such as sulphur dioxide or residual agrochemicals (Zuzuarregui & del Olmo, 2004). Towards completion of fermentation, essential nutrients such as glucose, nitrogen and amino acids are often depleted and the concentration of ethanol can reach toxic levels of 16% weight per total volume (w/v) (Zuzuarregui & del Olmo, 2004; Marks et al., 2008). Ethanol toxicity in S. cerevisiae largely targets the plasma membrane, increasing its fluidity and causing influx of protons. Proton influx disrupts the electrochemical gradient across the plasma membrane resulting in intracellular acidification and ultimately a decrease in growth rate and cell viability (Casey & Ingledew, 1986; Piper, 1995). Ethanol stress also denatures proteins, inhibits glycolysis and amino acid transport pathways and generates reactive oxygen species that inhibit mitochondrial function and cause oxidative damage to lipids, proteins and DNA (Leao & van Uden, 1982, 1984; Costa et al., 1997).

By studying natural variation within S. cerevisiae, we have gained insight into the genetic loci and biological and chemical processes that give rise to desirable fermentative properties (Dunn et al., 2005; Borneman et al., 2008). Although most industrial yeasts are diploid, the presence of chromosome length variation, aneuploidy or even polyploidy occurs in some strains (Barre et al., 1993; Codon et al., 1998). One theory explaining the presence of extra genetic material in industrial yeast is that increased dosage of genes is necessary for survival in stressful environments such as fermentation (Bakalinsky & Snow, 1990). Several examples of increased gene dosage or gene expression corresponding to specific industrial environments exist. Duplication of alcohol dehydrogenase (ADH) genes, hexose transporters and other glycolysis genes improves the ability of S. cerevisiae to survive the rapid production and consumption of ethanol as a strategy to outcompete microorganisms (Thomson et al., 2005; Gordon et al., 2009). Moreover, flor yeast, employed in wine making for their ability to oxidize ethanol to pyruvate, have multiple copies of ADH2 and ADH3 genes necessary for this function (Guijo et al., 1997). Industrial baker's and distiller's yeast have widespread copies of SUC family genes, encoding related invertase genes necessary for sucrose utilization, which confers a faster growth rate on molasses compared with laboratory strains (Codon et al., 1998). Lastly, a chromosomal rearrangement that led to increased expression of the sulphite tolerance gene SSU1 was identified in the sulphite resistant wine yeast T73, suggesting that alterations in gene expression can have a similar effect to increasing gene dosage (Perez-Ortin et al., 2002).

Although differences between gene copy number and ploidy levels exist, more recent studies suggest a lesser prevalence of aneuploidy amongst S. cerevisiae wine strains than previously thought. Genetic analysis of 45 commercial yeast strains found that only five were actually aneuploid (Bradbury et al., 2006). Similarly, karyotyping of four commercial S. cerevisiae wine strains found no major aneuploidies or chromosomal rearrangements although variations in gene copy number were noted when compared with S288C. Although the variations were moderate, correlation between gene copy and drug sensitivity was observed even amongst intrastrain isolates (Dunn et al., 2005). Sequencing of the commonly used commercial wine yeast EC1118 demonstrated that 99% of its predicted genes are in common with S288C, and three distinct chromosomal regions are of non-Saccharomyces origin. Located within these distinct regions are 34 genes, many of which have key functions related to wine making such as sugar utilization and nitrogen-related functions (Novo et al., 2009). A cluster of five conserved ORFs of non-Saccharomyces origin was also recently identified in four industrial wine S. cerevisiae strains, suggesting that these ORFs are beneficial under industrial conditions (Novo et al., 2009; Borneman et al., 2011). Proteome comparison of the wine yeast strain AWRI1631 and S288C found that despite nearly 70 000 single-nucleotide changes, their predicted proteomes contain over 99% amino acid identity (Borneman et al., 2008). Taken all together, we can conclude that small changes in genetic material can account for differences in fermentative properties between industrial and laboratory yeasts and because of high gene and proteome similarity between commercial wine yeasts and S288C, we can utilize resources developed for the latter to investigate gene function for industrial applications.

A plethora of studies have utilized the yeast S288C deletion set to screen for gene deletions that result in ethanol sensitivity; however, the high variability of genes identified in these screens implies that the ethanol response may be highly dependent upon experimental conditions and ethanol concentration (Kubota et al., 2004; van Voorst et al., 2006; Kumar et al., 2008). Because most of these studies were conducted at low (5–8%) ethanol concentrations, atypical of industrial fermentations, we suspect that further genes involved in ethanol tolerance remain to be identified. Recently, the MoBY-ORF plasmid library was generated where all S. cerevisiae barcoded genes (including their native promoter and terminator sequences) have been cloned into a centromere-based plasmid (Ho et al., 2009). The barcodes enable quantification of the plasmids using microarray technology. A high copy (2μm based) plasmid library has been derived from the MoBY-ORF centromere base library (MoBY-ORF 2.0) such that dosage suppression studies can be conducted and plasmids conferring a growth advantage can be identified by quantification (Magtanong et al., 2011). In this study, we have introduced the MoBY-ORF 2.0 library into two strains of S. cerevisiae, S288C and the commercial wine yeast Enoferm M2 (Bradbury, 2006), and subjected both strains to 12% ethanol stress to identify genes that confer a growth advantage. We have identified two genes, RSA3 and RCN1, that when present in high copy number enable S. cerevisiae to tolerate the high ethanol concentrations typical of a wine fermentation.

Materials and methods

Strains and DNA manipulation

The S. cerevisiae strains used were as follows: YPH499 (S288C) (MATa his3∆200 ade2-101 ura3-52 lys2-801 leu2∆1 trp1-∆63) and M2/WYM31 (MATa ho∆::hphMX4 leu2∆::natMX4), an industrial wine yeast derived from a parental strain obtained as a gift from Richard C. Gardner (Bradbury et al., 2006). To enable selection of the 2μm MoBY-ORF 2.0 library in the parental M2 strain, a kanMX6 marker cassette was PCR amplified with primers containing 70 bp upstream of the LEU2 start codon and 70 bp downstream of the LEU2 stop codon (Longtine et al., 1998). Amplified PCR products were transformed via standard yeast lithium acetate methods, and integration of the kanMX6 cassette into the LEU2 locus was confirmed by PCR. The kanMX6 marker was then switched to natMX4 as described (Tong et al., 2001). Mating types were determined by PCR and a haploid M2 isolate, WYM31 (MATa ho∆::hphMX4 leu2∆::natMX4) was selected for further utilization in this study (Huxley et al., 1990). The two strains YPH499 and WYM31 are referred to as S288C and M2 for the purpose of this study. For primer sequences utilized in this study, refer to Supporting Information, Table S1.

Media and growth conditions

Using standard yeast methods, strains of S. cerevisiae were cultured and maintained with yeast peptone dextrose (YPD) broth and transformants were selected and maintained with synthetic complete media supplemented with 2% glucose and all necessary amino acids except for leucine (SC−Leu) (Guthrie & Fink, 1991). Cultures were aerobically grown at a temperature of 25 °C unless otherwise stated. For studies where cells were grown in 96-well microtitre plates, an Infinite® Pro microplate reader (Magellan v7.0 Software; Tecan Group Ltd.) was used as follows. Cultures were diluted from log phase to an optical density (OD600 nm) of 106 cells mL−1 (OD600 nm 0.1), and a 200-μL volume was added to each well with at least three technical replicates. The plate was sealed with a transparent film (Beckman Coulter®) to minimize evaporation. Air exchange was facilitated by puncturing the film near the edge of each well with a 27.5 gauge hypodermic needle. OD600 nm was read after kinetic cycles of 20 min with equal amounts of orbital and linear shaking at an amplitude of 2.5 mm. The temperature of the growth chamber was maintained at 25 °C except where otherwise noted.

MoBY-ORF 2.0 transformations

The 2μm MoBY-ORF 2.0 library was amplified in Escherichia coli by growth overnight at 37 °C in 2YT media (pH 7.0) containing 1.6% Bacto tryptone, 1% Bacto yeast extract, 0.5% NaCl and supplemented with 0.4% glucose, 200 μg mL−1 carbenicillin and 100 μg mL−1 kanamycin. Aliquots of 1.5 mL of cells (OD600 nm c.20.0) were collected by centrifugation and resuspended in 50 μL of 2YT containing 15% glycerol and stored at −80 °C until needed. Plasmids from one aliquot were extracted with a Genejet Plasmid Miniprep kit (Fermentas) following manufacturer's instructions and transformed using a LiAc method as described (Butcher & Schreiber, 2006) into YPH499/S288C. For transformation of the 2μm MoBY-ORF 2.0 library into M2, electroporation was used to obtain sufficiently high yields of transformants. Fifty millilitres of log phase M2 cells (OD600 nm 1.0) was harvested by centrifugation at 4 °C and resuspended in 10 mL of TE/LiAc buffer (pH 7.5). After 45-min incubation at 30 °C, 250 μL of 1 M DTT was added and further incubated for 15 min at 30 °C. The yeast suspension was then diluted to 50 mL with dH2O and washed and concentrated three times (4000 g, 4 °C) with 25 mL dH2O, 3 mL 1 M sorbitol and 0.5 mL 1 M sorbitol, respectively, after which 40 μL of concentrated yeast (OD600 nm c.50.0) was mixed with a single aliquot of plasmid DNA and transferred to a 0.2-cm gap electroporation cuvette. The cuvette was pulsed at 1.5 kV, 25 μF, 200 Ω, after which cells were recovered in YPD for 2 h at 25 °C.

In both transformation protocols, cells were spread onto SC−Leu plates and grown for 2–3 days at 25 °C until single colonies were obtained. Approximately 40 × 103 colonies were scraped off the plates and pooled into 20 mL YPD containing 15% glycerol and 100 ug mL−1 G418 and frozen in aliquots at OD600 nm c.20 at −80 °C. To transform individual MoBY-ORF 2μm plasmids, amplification and extraction of plasmids were conducted as described previously, and the same standard LiAc protocol was utilized for both S288C and M2 with selection on SC−Leu.

MoBY-ORF 2.0 plasmid screen

To screen for genes that improve ethanol tolerance when their dosage is increased, we used the following approach for both the M2 and S288C MoBY-ORF 2.0 (2μm) plasmid libraries. Overnight cultures of the plasmid pool were diluted and grown to log phase (OD600 nm c.1.0) in 20 mL SC−Leu. The pool was then split into two 10-mL cultures at an OD600 nm c.0.2, one containing SC−Leu (to select for the plasmid) with 12% v/v ethanol (stress condition) and one with SC−Leu containing no ethanol (control). Both the stress and control cultures were grown aerobically for 48 h at 25 °C. The control pool was maintained in log phase by dilution every 12 h in fresh SC−Leu media. To prevent exhaustion of glucose and nutrients over 48 h in the ethanol stress pool, cells were also harvested and resuspended in fresh SC−Leu 12% ethanol media at 24 h. After 48 h, cells from both the stress and control conditions were harvested and plasmids were extracted as described (Butcher & Schreiber, 2006) using a Genejet Plasmid Miniprep kit (Fermentas) and following the manufacturer's directions.

A competitive hybridization was performed for each screen as outlined previously (Magtanong et al., 2011). Briefly, barcodes from the 12% ethanol condition were amplified with biotinylated universal TAG4 primers, whereas nonbiotinylated universal TAG4 primers were used to amplify barcodes from the 0% ethanol control condition. Each hybridization mixture contained 3 : 1 (v/v) nonbiotinylated:biotinylated PCR-amplified barcodes. Barcodes were hybridized to a TAG4 microarray as previously described (Pierce et al., 2006). Barcode values for each ORF were ranked in descending order, and cut-off values of 800 for S288C and 600 for M2 were established to identify candidate genes that confer ethanol tolerance.

Ethanol tolerance assay

To test strains carrying individually transformed MoBY-ORF 2μm plasmids for improved tolerance to ethanol, 5 mL log phase cultures (OD600 nm c.0.6–0.8) were diluted to an OD600 nm 0.1 in SC−Leu media containing 16% v/v ethanol and incubated for 3 h at 25 °C. After incubation, cells were collected by centrifugation and resuspended in SC−Leu media containing no ethanol. Recovery of MoBY-ORF 2μm strains after 16% ethanol shock was monitored in 96-well microtitre plates, and growth was measured by recording optical density as described previously.

Zymolyase cell wall digest

Differences in cell wall constitution were tested using a zymolyase cell wall digestion assay adapted to a 96-well plate microtitre format as described (Ovalle et al., 1999) with exception to the utilization of 20T zymolyase in this study rather than 100T. Briefly, overnight cultures were grown to log phase (OD600 nm c.0.8), collected by centrifugation and resuspended in Tris–EDTA (TE) buffer with 5% polyethylene glycol 8000 (PEG 8000) at OD600 nm c.2.0. In a 96-well plate, 200 uL aliquots of each strain were added to individual wells in quadruplicate and 50 μL of a 20T zymolyase solution was added. Different concentrations of 20T zymolyase were used for each strain background (200 μg mL−1 for S288C and 500 μg mL−1 for M2) because of the intrinsic differences in cell wall strength between S288C and M2. Changes in OD600 nm were measured every minute for 60 min. The OD600 nm/Embedded Image was calculated for each replicate, and mean values were determined for each time point. Log-transformed values were plotted vs. time, and the maximum lysis rate (MLR) and lag time for each replicate were also calculated as previously described (Ovalle et al., 1999). A Student's t-test was also performed to test whether mean values for MLR and lag time were significantly different from the vector control strain.

Stress response growth curves

Growth in the presence of a variety of stress media was monitored by OD600 nm in a 96-well microtitre plate as described previously. For heat stress, log phase cultures (OD600 nm c.0.6–0.8) grown at 25 °C in SC−Leu were added to a 96-well plate and immediately transferred to the growth chamber preheated to 37 °C for S288C and 42 °C for M2. For osmotic stress, log phase cells were grown directly in SC−Leu supplemented with 0.4 M NaCl (S288C) or 1.0 M NaCl (M2). For oxidative stress, growth in 0.75 mM H2O2 (S288C) or 1.0 mM H2O2 (M2) SC−Leu media was utilized. To test sensitivity to translation inhibition, log phase cells were resuspended in SC−Leu with 100 ng mL−1 cycloheximide (S288C) and 25 ng mL−1 cycloheximide (M2), which we had determined to be sublethal concentrations of cycloheximide that conferred growth inhibition to each strain.

Growth curve statistical analysis

For analysis of growth curves from the automated plate reader, the OD600 nm of each time point was averaged amongst replicates and the mean values were plotted vs. time. To determine statistical significance of growth curves, we analysed three distinct growth phases for individual replicates: the lag phase, the logarithmic growth rate and the maximum OD600 nm (ODmax). The logarithmic growth rate was determined to be the slope of a linear trend line fit to the exponential portion of each curve. Where the exponential growth phase was less distinct, the growth rate was taken to be the slope of a linear trend line fit to the maximal growth increase consisting of at least 12 time points (4 h). Minimum OD600 nm values (ODmin) were calculated by averaging the first six time points of each curve, and the lag phase was determined to be time of intersection between ODmin values and an extrapolated tangent line from the exponential growth phase. Finally, ODmax values were calculated by averaging the last six time points of each curve. For each of the parameters measured, a Student's t-test was used to determine if the mean values for strains carrying 2μm MoBY-ORF plasmids were significantly different from a control strain carrying an empty 2μ vector.


Increased dosage of RCN1 and RSA3 confer ethanol tolerance

To identify genes involved in ethanol tolerance in yeast, we independently transformed two strains of S. cerevisiae – our laboratory S288C strain and the M2 industrial wine yeast strain – with the 2μm MoBY-ORF 2.0 plasmid library and grew each set of transformants in 12% ethanol (v/v) as the stress condition and no ethanol as the control condition. After 48 h of incubation, plasmid barcodes from the 12% ethanol stress pool were quantified relative to the control pool as described in the . Quantified raw average values for all uptag and downtag barcodes identified by microarray from our S288C (Fig. 1a) and M2 (Fig. 1b) screen are shown.

Figure 1

Quantified abundance of MoBY-ORF barcode tags after prolonged ethanol stress (12% v/v) in Saccharomyces cerevisiae. Descending order raw average values of all barcode tags are shown for parallel screens with (a) S288 and (b) M2. Dashed box indicates genes above cut-off values for each screen. (c) Genes associated with barcode values above 600 in M2 (black) and 800 in S288C (white) are presented.

To validate the results of our screen, we identified genes with barcodes that had a raw hybridization value significantly above background (boxed area Fig. 1a and b). With this criterion, we identified 14 plasmids from the M2 screen and 11 plasmids from the S288C screen that were more abundant in 12% ethanol vs. control cultures (Fig. 1c, Table S2). Five of the plasmids from the M2 and S288C screen had genes in common, which also happened to be the top-ranked genes from the S288C screen. These genes are as follows: RCN1, MAP2, EMI5, RSA3 and YAT1 (Fig. 1c). We tested whether strains carrying high copy plasmids carrying these genes had improved growth following exposure to toxic levels of ethanol. Both M2 and S288C strains were individually transformed with RCN1, MAP2, RSA3, EMI5 and YAT1 plasmids in addition to an empty vector control plasmid. We exposed cells to 16% ethanol (v/v) in minimal media for 3 h, then resuspended cells in SC−Leu media and monitored growth curves using an automated 96-well plate reader. Compared with conditions used in our primary ethanol tolerance screen (12% ethanol for 48 h), we found that exposure to 16% ethanol for 3 h yielded higher reproducibility in our growth curves, likely due to the rapid ethanol shock over a shorter period of time. Growth curve analysis consisted of measuring the lag phase, logarithmic growth rate and the ODmax following ethanol shock. Mean values for growth curve parameters of each strain and plasmid combination following ethanol shock are shown in Table 1. In S288C, increased dosage of RCN1, RSA3 and MAP2 resulted in a significantly faster growth rate and a higher ODmax compared with the vector control strain (Fig. 2a, Table 1). Introduction of EMI5 or YAT1 on a high copy plasmid did not significantly improve growth following 16% ethanol shock in S288C (Fig. S1).

Figure 2

S288C and M2 strains carrying high copy RCN1 and RSA3 plasmids exhibit improved recovery following 16% ethanol shock. 2μm MoBY-ORF 2.0 vector (blue), RCN1 (red), RSA3 (green) and MAP2 (purple) plasmids were individually transformed into (a) S288C and (b) M2 and incubated in 16% ethanol minimal media for 3 h. Cells were then resuspended in SC−Leu media, and growth of strains was monitored in quadruplicate using an automated 96-well plate reader by measuring OD600 nm every 20 min. Mean optical density (OD600 nm) is plotted over a 24-h period.

View this table:

Growth curve analysis of S288C and M2 strains carrying 2μm high copy plasmids after 16% ethanol shock

Strain + plasmidMean lag time (h)Mean growth rate (105 cells mL−1 h−1)Mean ODmax (107 cells mL−1)
S288C + vector6.842.070.388
S288C + RCN18.873.560.571
S288C + MAP210.05.180.722
S288C + EMI56.941.930.410
S288C + RSA38.345.740.765
S288C + YAT18.811.430.335
M2 + vector14.412.11.41
M2 + RCN110.512.61.46
M2 + MAP212.78.411.17
M2 + EMI514.110.71.24
M2 + RSA311.511.41.39
M2 + YAT113.713.11.45
  • Analysis was conducted with a Student's t-test comparing mean values for lag phase, logarithmic growth rate and ODmax values for each 2μm plasmid with the vector control strain.

  • Significance of P < 0.05,

  • significance of P < 0.01,

  • significance of P < 0.001.

In the M2 strain background, increased dosage of RCN1, RSA3 and MAP2 significantly improved recovery from 16% ethanol shock by reducing the lag phase prior to logarithmic growth (Fig. 2b, Table 1). As the length of lag phase can be affected by the physiological competence of cells damaged from prior events, the shorter lag phases observed with the presence of RCN1, RSA3 and MAP2 on 2μm plasmids may be due to less cellular damage induced by 16% ethanol exposure (McMeekin et al., 2002). Compared with the vector, the logarithmic growth rate and ODmax did not significantly differ upon increased dosage of RCN1 or RSA3 in M2 (Fig. 2b, Table 1). Contrary to the improved growth rate in S288C, MAP2 reduced the growth rate and ODmax when present in high copy number in M2, suggesting that there may be intrinsic differences between strains regarding the function of MAP2 in ethanol tolerance. The presence of YAT1 and EMI5 on high copy plasmids did not significantly affect growth of M2 following 16% ethanol shock, similar to the results obtained in S288C (Fig. S1a and b). Based on these data, we continued our analysis with RCN1 and RSA3, which were the only genes that improved the growth of both S288C and M2 after ethanol stress.

RCN1 and RSA3 dosage influence cell wall assembly

When grown in the presence of cell membrane or cell wall perturbing agents, S. cerevisiae alters the constituents of its cell wall by increasing membrane proteins, chitin and mannoproteins bound to chitin through β1,3 and β1,6-glucans (de Groot et al., 2001). As genes involved in cell wall maintenance are important for ethanol tolerance, we determined whether increased dosage of RCN1 and RSA3 influences cell wall assembly (Teixeira et al., 2009). Strains were grown to log phase in SC−Leu and resuspended in a TE buffer with zymolyase, an enzyme mixture comprised of proteinases and β1,3-glucanase that digests cell wall constituents and subsequently causes cell lysis. The decrease in OD600 nm because of cell wall lysis was monitored using an automated microplate reader, and semi-log plots of normalized OD600 nm values were graphed for analysis. The MLR and lag time prior to lysis were used as parameters for quantifying zymolyase resistance or sensitivity as described in the . A 10% or greater difference in MLR was deemed a significant difference between strains, as previously described (Ovalle et al., 1999). S288C carrying high copy RCN1 and RSA3 increased the lag time by 5.5 and 4.0 min respectively, both of which were statistically different from the vector control (Fig. 3a, Table 2). Following the lag time, curves were characterized by a steady decline in optical density. Increased dosage of RCN1 decreased the MLR by over 10% compared with the vector control strain, whereas increased dosage of RSA3 had no effect (Table 2). These data suggest that RCN1, when present in high copy number, can improve resistance to zymolyase by influencing cell wall assembly in S288C. RSA3, when present in high copy number in M2, significantly decreased the MLR by nearly 18%, from 5.88 × 104 log cells mL−1 min−1 (vector control) to 4.83 × 104 log cells mL−1 min−1, whereas high copy numbers of RCN1 did not decrease the MLR by a significant margin (Fig. 3b, Table 2). The lag time of strains carrying high copy plasmids of RCN1 and RSA3 was again increased, albeit to a lesser extent than in S288C, from 18.4 min (vector control) to 19.8 min for RCN1 and 20.4 min for RSA3 (Table 2). Statistical analysis found that the lag time upon increased RSA3 dosage was significantly different from the vector control; however, the lag time upon increased RCN1 dosage was not (Table 2). Whereas increased dosage of RCN1 resulted in greater resistance to zymolyase than increased dosage of RSA3 in S288C, it is interesting that the reverse holds true for the M2 strain, and thus, each gene appears to influence cell wall assembly in a strain-specific manner.

Figure 3

Increased dosage of RCN1 in S288C and RSA3 in M2 decreases the cell lysis rate during zymolyase digestion. Log OD600 nm values of (a) S288C and (b) M2 strains carrying the 2μm vector (blue), RCN1 (red) and RSA3 (green) plasmids are shown as zymolyase digestion progresses. Because of inherent cell wall differences, 200 and 500 μg mL−1 zymolyase for S288C and M2, respectively, were added to individual wells (in quadruplicate) of a 96-well plate, and OD600 nm was automatically measured every minute for up to 1 h.

View this table:

Zymolyase cell wall digestion analysis of S288C and M2 strains carrying 2μm RCN1 and RSA3 plasmids

Strain + plasmidMean lag time (min)MLR (log 104cells mL−1 min−1)% MLRvector
S288C + vector23.51.78
S288C + RCN129.01.6089.9
S288C + RSA327.51.7397.1
M2 + vector18.45.88
M2 + RCN119.85.6896.6
M2 + RSA320.44.8382.1
  • Statistical analysis was conducted with a Student's t-test comparing lag time and MLR values for each plasmid to the vector control strain.

  • strains with a MLR <90% of the vector control have significantly higher resistance to zymolyase.

  • S288C strains were digested with 200 μg mL−1 zymolyase.

  • M2 strains were digested with 500 μg mL−1 zymolyase.

  • Significance of P < 0.05.

Increased dosage of RSA3 improves growth during environmental stress

It has been suggested that genes that respond to thermal, osmotic and oxidative stress also play a role in ethanol tolerance; thus, we analysed the growth rates of M2 and S288C strains carrying high copy plasmids of RCN1 and RSA3 in the presence of these stressors (Piper, 1995; Costa et al., 1997; Yoshikawa et al., 2009). We monitored the growth of S288C carrying 2μm RCN1 and RSA3 plasmids in SC−Leu media with a range of inhibitory, but sublethal, levels of thermal stress (37 °C), osmotic stress (0.4 M NaCl) and oxidative stress (0.75 mM H2O2) according to previously published data (Izawa et al., 1995; Rep et al., 1999; Gasch et al., 2000). We found that M2 is more tolerant to stress; therefore, a higher range of heat (42 °C), osmotic (1.0 M NaCl) and oxidative (1.0 mM H2O2) stressors were used to challenge M2 carrying 2μm RCN1 and RSA3 plasmids. Prior to stress exposure, growth curves of both M2 and S288C carrying each high copy plasmid were compared with the vector control in no stress conditions (SC−Leu media at 25 °C) with the finding that growth rates and ODmax did not differ significantly (data not shown). Over all, increased dosage of RCN1 and RSA3 did not significantly change the lag phase period during exposure to any of the environmental stressors tested, and thus, only the growth rate and ODmax are reported. When subjected to heat stress at 37 °C, increased dosage of either RCN1 or RSA3 in S288C conferred a significantly faster growth rate and a higher ODmax compared with the vector control strain suggesting that high copies of both genes can improve thermal tolerance in S288C (Fig. 4a, Table 3). The presence of high copy plasmids of RCN1 and RSA3 in M2 during 42 °C heat stress resulted in statistically higher growth rates and ODmax values (Fig. 4b, Table 4). Whereas the improvement in growth was more subtle with increased dosage of RCN1, increased dosage of RSA3 caused a drastic improvement of growth under heat stress in the M2 strain.

Figure 4

The presence of the 2μm RSA3 and the 2μm RCN1 plasmid improves growth rate at elevated temperatures in S288C and M2. (a) S288C and (b) M2 strains carrying vector (blue), RCN1 (red) and RSA3 (green) 2μm plasmids were grown in minimal media at 37 °C (S288C) or at 42 °C (M2). OD600 nm was recorded in an automated 96-well plate reader, and mean values for each strain shown for up to 24 h.

View this table:

Comparison of growth rate and ODmax values during environmental stress exposure with S288C carrying RCN1 and RSA3 2μm plasmids

Strain + 2μm plasmidHeat stressOsmotic stressOxidative stress
Growth rateODmaxGrowth rateODmaxGrowth rateODmax
S288C + vector6.941.115.150.6635.110.841
S288C + RCN17.191.176.640.7545.280.866
S288C + RSA37.241.24***7.220.7275.930.981
  • Statistical analysis was performed with a Student's t-test to compare values from each 2μm plasmid strain with values from the vector control strain.

  • Growth rate is given in 105 cells mL−1 h−1.

  • Growth in SC−Leu at 37 °C.

  • Growth in SC−Leu + 0.4 M NaCl.

  • Growth in SC−Leu + 0.75 mM H2O2.

  • Significance of P < 0.05,

  • significance of P < 0.01.

View this table:

Comparison of growth rate and ODmax values during environmental stress exposure with M2 carrying RCN1 and RSA3 2μm plasmids

Strain + 2μm plasmidHeat stressOsmotic stressOxidative stress
Growth rateODmaxGrowth rateODmaxGrowth rateODmax
M2 + vector0.9020.3222.940.6897.811.29
M2 + RCN11.390.3643.160.6998.201.28
M2 + RSA34.230.6503.620.7869.951.39
  • Statistical analysis was performed with a Student's t-test to compare growth rate and ODmax values from each 2μm plasmid strain with values from the vector control strain.

  • Growth rate is given in 105 cells mL−1 h−1.

  • Growth in SC−Leu at 42 °C.

  • Growth in SC−Leu + 1.0 M NaCl.

  • Growth in SC−Leu + 1.0 mM H2O2.

  • Significance of P < 0.05,

  • significance of P < 0.01,

  • significance of P < 0.001

When exposed to osmotic salt stress, S288C carrying 2μm RCN1 and RSA3 plasmids displayed increased growth rates compared with vector control, but M2 carrying 2μm RCN1 and RSA3 plasmids did not significantly alter any of the growth curve parameters (Fig. 5a and b Tables 3 and 4). The mean values for growth rate and ODmax upon increased dosage of RSA3 in M2 during osmotic stress were notably higher; however, high variability in growth in 1.0 M NaCl media did not allow for statistical significance (Table 4). Growth at lower NaCl concentrations also yielded no significant differences between strains (data not shown). As the M2 strain has adapted to a high-osmolarity environment during wine fermentation, inherent osmotic tolerance is naturally high, and thus, increased dosage of a single gene may have a lesser effect in M2 than S288C.

Figure 5

Increased dosage of RCN1 and RSA3 improves osmotolerance in S288C but not in M2. (a) S288C and (b) M2 strains carrying vector (blue), RCN1 (red) and RSA3 (green) 2μm plasmids were grown in minimal media containing 0.4 M NaCl (S288C) or 1.0 M NaCl (M2) in triplicate. OD600 nm was recorded in an automated 96-well plate reader, and mean values for each strain are shown for up to 35 h.

The final ethanol-related stress that we tested was sensitivity to oxidative stress by exposing cells to hydrogen peroxide. Increasing the copy number of RCN1 in either S288C or M2 did not significantly improve growth during incubation with hydrogen peroxide, suggesting that RCN1 does not have a significant role in response to oxidative stress (Fig. 6, Tables 3 and 4). To the contrary, increasing the copy number of RSA3 in S288C resulted in a higher maximal OD600 and increasing the copy number of RSA3 in M2 resulted in a faster growth rate (Fig. 6, Tables 3 and 4). Therefore, increasing the dosage of RSA3 improves tolerance to oxidative stress.

Figure 6

S288C and M2 carrying high copy RSA3 but not RCN1 improve oxidative stress tolerance. (a) S288C and (b) M2 strains carrying vector (blue), RCN1 (red) and RSA3 (green) 2μm plasmids were grown in minimal media containing 0.75 mM H2O2 (S288C) or 1.0 mM H2O2 (M2) in triplicate. OD600 nm was recorded in an automated 96-well plate reader, and mean values for each strain are shown for up to 40 h.

The M2 wine yeast is hypersensitive to cycloheximide

Increasing RSA3 production imparts a growth advantage to S288C and M2 strains under ethanol, heat and osmotic and oxidative stress, with exception of the M2 strain during osmotic stress. Given that Rsa3p has a role in 60S ribosomal subunit assembly, we supposed that increased dosage of RSA3 may improve resistance of cells to a variety of stressors by affecting protein translation (Rosado et al., 2007). To test this hypothesis, we began by determining sublethal concentrations of cycloheximide, a drug that inhibits translation in S. cerevisiae, for M2 and S288C strains. By analysing growth of S288C and M2 vector control strains in the presence of a range of cycloheximide concentrations (Fig. 7a–c), we noticed that the M2 strain is more sensitive to cycloheximide than S288C. The hypersensitivity of M2 to cycloheximide was surprising because M2 is an industrial strain and our analysis here demonstrates that M2 is inherently more tolerant to other stresses than the S288C laboratory strain. Both strains showed a concentration-dependent reduction of growth in cycloheximide, and M2 growth was inhibited at 50 and 100 ng mL−1 of cycloheximide, whereas S288C was still able to grow (Fig. 7b and c). We then analysed the growth of both S288C and M2 strains carrying the high copy RSA3 plasmid upon exposure to cycloheximide. The growth rate of strains carrying the 2μm RSA3 plasmid did not significantly differ from the vector control when grown in a range of 100–150 ng mL−1 cycloheximide for S288C and 25–50 ng mL−1 for M2 (data not shown). Although increased dosage of RSA3 may improve tolerance to ethanol and other stressful environments by influencing translation, increasing the copy number of a single trans-acting ribosome assembly factor was not sufficient to improve growth of either S288C or M2 strains in the presence of a translation inhibitor.

Figure 7

The Saccharomyces cerevisiae wine yeast M2 is more sensitive to translation inhibition than S288C. S288C (purple) and M2 (yellow) vector control strains were grown with (a) 25 ng mL−1, (b) 50 ng mL−1 and (c) 100 ng mL−1 cycloheximide in minimal media. Strains were grown in an automated 96-well plate reader in quadruplicate, and mean OD600 nm values are shown over 40 h.


We have conducted the first genome-wide screen using the 2μm MoBY-ORF 2.0 plasmid library for genes, which when present in high copy, confer ethanol tolerance to the M2 industrial wine yeast and the S288C laboratory yeast strain. By setting stringent cut-offs for quantified barcode values, we identified a small group of genes to further analyse. The top five ranked genes in S288C also exceeded the cut-off in M2 and thus were likely candidate genes for conferring ethanol tolerance when present on high copy plasmids. Increased dosage of two of these genes, RCN1 and RSA3, improved growth in both strain backgrounds upon recovery from 16% ethanol shock. We also found that RCN1 and RSA3 high copy plasmids conferred tolerance of S888C and M2 strains to cell wall, heat, osmotic and oxidative stress to varying degrees. The variability detected in these assays could be due to expression differences of RCN1 and RSA3 under each stress condition as each gene is expressed from its native promoter and may be subjected to transcriptional regulation in a specific stress condition. Nonetheless, RCN1 and RSA3 emerge from our studies as genes important for ethanol and general stress tolerance in S. cerevisiae.

The role of RCN1 in ethanol tolerance

The calcineurin signalling pathway is induced during heat, osmotic and cell wall stress. RCN1 is a regulator of calcineurin-, a calcium- and calmodulin-dependent protein phosphatase that responds to changes in intracellular calcium levels (Kingsbury & Cunningham, 2000). In our study, RCN1 was initially identified in a screen for ethanol tolerance genes. An overlap between calcineurin signalling and osmotic and ethanol stress is not surprising as high ethanol can induce osmotic shock by binding free water, which effectively reduces the water activity of the extracellular environment (Hallsworth, 1998). This overlap was confirmed by our data showing that S288C strains carrying RCN1 in high copy number improve resistance to ethanol, saline osmotic shock and zymolyase. The role of RCN1 in response to these individual stresses was less clearly defined in M2.

Recently, it was shown that the Crz1p transcription factor translocates into the nucleus in a calcineurin-dependent manner when exposed to 8% ethanol stress (Araki et al., 2009). In addition, the calcineurin/Crz1 pathway is required for adaptive tolerance to 18% ethanol (Araki et al., 2009). We find that increased dosage of RCN1 improves recovery of both M2 and S288C strains after a 3-h 16% ethanol shock, which further suggests that the calcineurin pathway is important for adaptation to ethanol stress (Fig. 2). Both high copy and deletion of RCN1 diminish calcineurin signalling in S. cerevisiae; therefore, a biphasic model of calcineurin regulation by RCN1 has been proposed (Hilioti et al., 2004). When phosphorylated by the Mck1p protein kinase, Rcn1p stimulates calcineurin signalling, which activates Crz1p and induces transcription of RCN1. In turn, high levels of Rcn1p bind and inhibit calcineurin by preventing the access of Ca2+ and calmodulin to the calcineurin active site (Hilioti et al., 2004). Whether increased dosage of RCN1 improves stress resistance under our assay conditions because of inhibition or stimulation of calcineurin signalling remains to be determined.

The role of RSA3 in ethanol tolerance

In S. cerevisiae, relatively little is known about the molecular function of Rsa3p. RSA3 is a nonessential gene that was first identified in a synthetic lethal interaction with dpb6 (Kressler et al., 1999; de la Cruz et al., 2004). Dbp6p is a putative ATP-dependent RNA helicase that is required for assembly of the 60S ribosomal subunit (Kressler et al., 1998). Likewise, deletion of RSA3 results in a deficit of 60S ribosomal subunits and Dbp6p and Rsa3p cosediment with 60S ribosomal subunits and interact in vivo (de la Cruz et al., 2004). Both Dpb6p and Rsa3p have been identified in a low-molecular mass complex with three other proteins, Npa1p, Npa2p and Nop8p, all of which function in early 60S ribosomal biogenesis (Rosado et al., 2007). In our study, however, neither DBP6, NOP8, NPA1 nor NPA2 was highly ranked in either of our screens. RSA3 may be a more critical factor for 60S ribosomal assembly than its molecular complex partners, or RSA3, when present in high copy, may have alternative roles during ethanol stress.

A slow growth phenotype at 37 °C has been observed in rsa3 deletion mutants, which concurs with our data that increased dosage of RSA3 improves growth during thermal stress (de la Cruz et al., 2004). These data also suggest that increased dosage of RSA3 does not confer a dominant negative phenotype, at least with respect to heat stress, as the rsa3 null phenotype is opposite to the phenotype upon increased RSA3 dosage. Given that rsa3 mutants lack normal levels of 60S ribosomal subunits, one hypothesis is that increasing the copy number of RSA3, a trans-acting ribosome assembly factor, increases ribosome levels and subsequently, translation rates. We were surprised to discover that the M2 industrial yeast strain is highly sensitive to the translation stalling agent cycloheximide compared with the S288C laboratory strain because M2 is more tolerant of other stresses (Fig. 7). We did not observe a significantly improved growth rate when RSA3 dosage was increased in cycloheximide-treated cultures of M2 or S288C (data not shown). Therefore, overproduction of RSA3 cannot prevent translation stalling by cycloheximide but could still have a subtle impact on translation rates.

As increasing RSA3 or RCN1 dosage improved the resistance of yeast to a variety of stresses, we asked whether M2 carrying the RSA3 or RCN1 2μm plasmids could outperform the M2 vector control strain during a wine fermentation. We inoculated the M2 strains into 32% equimolar glucose:fructose synthetic grape juice media and carried out a 24-day fermentation (refer to Appendix S1 for details). We detected a very minor, yet statistically significant, increase in ethanol production in strains carrying high copy RSA3, but not RCN1, when compared with strains carrying vector alone (Fig. S2). Monitoring ethanol production may not strictly reflect the robustness of the yeast; however, yeast deletion set fitness profiling studies from our laboratory have found that heterozygous diploid mutants of ribosomal protein genes, including RSA3, actually have a fitness advantage during a 14-day wine fermentation (Piggott et al., 2011). Therefore, in the high osmolarity, low pH and gradually increasing ethanol conditions of a wine fermentation, decreasing ribosomal production is beneficial to the cell.

Ribosome biosynthesis consumes a majority of the cell's resources in growing yeast cells, and a number of environmental conditions can rapidly induce or repress transcription of ribosomal RNA and ribosomal protein genes (Warner, 1999). Therefore, it is not surprising that both induction and repression of ribosomal genes have been reported in response to different stresses. During nitrogen-limited wine fermentation, microarray analysis revealed an increase in ribosome biogenesis and assembly genes; a similar response has been observed under cold and high-salinity environments (Mendes-Ferreira et al., 2007). Protein synthesis genes are reported to be downregulated in response to short-term ethanol stress (7% ethanol for 30 min) but induced in response to long-term ethanol stress (7% ethanol for c.20 h) (Alexandre et al., 2001; Li et al., 2010). Moreover, a S. cerevisiae strain previously adapted to 10% ethanol, induced expression of genes related to ribosomes and protein synthesis during long-term ethanol stress (10% ethanol for c.150 h), whereas its unadapted parent strain largely downregulated these genes (Dinh et al., 2009). These results suggest that after initial transcriptome changes to conserve energy, survival during prolonged ethanol exposure requires induction of ribosomal genes to enable growth and metabolic function in ethanol-stressed environments. In our 3-h 16% ethanol shock assay, high copy numbers of RSA3 could have aided the recovery of both S288C and M2 yeast strains by elevating ribosome levels or translational activity prior to or following ethanol exposure allowing for fast repair of damaged cells. Similarly, the decrease in cellular lysis with increased dosage of RSA3 during zymolyase digestion in M2 could be a product of increased de novo synthesis of cell wall repair proteins. Finally, the hypersensitivity of the M2 wine yeast strain to cycloheximide compared with S288C suggests that S. cerevisiae strains may have optimal growth at inherently different translation rates and that perturbation of ribosome biogenesis and ultimately translation will be an important avenue to explore in the creation of stress-resistant industrial yeast strains.

Supporting Information

Fig. S1. Growth of S. cerevisiae S288C and M2 strains carrying high copy EMI5 and YAT1 plasmids following 16% ethanol shock.

Fig. S2. Fermentations conducted with the M2 wine yeast carrying 2µm RCN1 and RSA3 plasmids.

Table S1. Primers utilized in this study

Table S2. Genes identified that improve ethanol tolerance when present in high copy number in Saccharomyces cerevisiae strains S288C or M2

Appendix S1. Materials and methods.

Please note: Wiley-Blackwell is not responsible for the content or functionality of any supporting materials supplied by the authors. Any queries (other than missing material) should be directed to the corresponding author for the article.


The authors would like to acknowledge Dr Richard C. Gardner for his generous donation of the M2 parental strain to the UBC Wine Research Centre. We would also like to thank Dr H.J.J. van Vuuren and Dr L. Madilao for their assistance in HPLC quantification of wine fermentation samples. M.J.A. was supported by the American Wine Society, the American Society for Enology and Viticulture and the Canadian Vintners association. C.B. was supported by the Natural Sciences and Engineering Research Council of Canada (NSERC, RGPIN 204899-06) and the Canadian Institutes of Health Research (MOP-57830). V.M. was supported by the CIHR (MOP-84242) and NSERC (RGPIN 326924-06). V.M. is a Canada Research Chair in Enology and Yeast Genomics.


  • Editor: Isak Pretorius


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