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Critical role of RPI1 in the stress tolerance of yeast during ethanolic fermentation

Rekha Puria, M. Amin-ul Mannan, Rohini Chopra-Dewasthaly, Kaliannan Ganesan
DOI: http://dx.doi.org/10.1111/j.1567-1364.2009.00549.x 1161-1171 First published online: 1 December 2009


Stress tolerance of yeast Saccharomyces cerevisiae during ethanolic fermentation is poorly understood due to the lack of genetic screens and conventional plate assays for studying this phenotype. We screened a genomic expression library of yeast to identify gene(s) that, upon overexpression, would prolong the survival of yeast cells during fermentation, with the view to understand the stress response better and to use the identified gene(s) in strain improvement. The yeast RPI1 (Ras-cAMP pathway inhibitor 1) gene was identified in such a screen performed at 38 °C; introducing an additional copy of RPI1 with its native promoter helped the cells to retain their viability by over 50-fold better than the wild type (WT) parent strain, after 36 h of fermentation at 38 °C. Disruption of RPI1 resulted in a drastic reduction in viability during fermentation, but not during normal growth, further confirming the role of this gene in fermentation stress tolerance. This gene seems to improve viability by fortifying the yeast cell wall, because RPI1 overexpression strain is highly resistant to cell lytic enzyme zymolyase, compared with the WT strain. As the RPI1 overexpression strain substantially retains cell viability at the end of fermentation, the cells can be reused in the subsequent round of fermentation, which is likely to facilitate economical production of ethanol.

  • Saccharomyces cerevisiae
  • fermentation stress tolerance (FST)
  • ethanolic fermentation
  • overexpression screen
  • RPI1
  • cell wall integrity


The yeast Saccharomyces cerevisiae is the backbone of the ethanolic fermentation industry owing to its capability for rapid and efficient conversion of sugars into ethanol. During fermentation, it is subjected to multiple stress conditions that include a high initial substrate concentration, accumulation of a high concentration of ethanol, accumulation of toxic byproducts and a decrease in pH of the medium (Pampulha & Loureiro, 1989; Attfield, 1997). It is also subjected to higher temperatures during fermentation, particularly in tropical countries. These conditions, especially ethanol and high temperature (Piper, 1995), have synergistic effects on the viability and efficiency of yeast to produce ethanol. We refer to the ability of yeast cells to withstand these conditions as ‘fermentation stress tolerance’ (FST), as it is not always possible to clearly separate the effect of one stress condition from another prevailing during fermentation.

FST is poorly understood due to the lack of a plate screen that can simulate the conditions encountered by yeast within the liquid fermentation broth. Thus, isolation of mutants impaired in FST and identification of corresponding genes are quite difficult. To circumvent these limitations, earlier we have reported a method that can simultaneously monitor the fitness of individual mutants of yeast present as a mixed population in liquid broth, and used the same for studying the role of several genes critical for FST (Sharma, 2001). Several other genes important for the growth of yeast under ethanol stress were identified by screening yeast transposon insertion mutants (Takahashi, 2001; van Voorst, 2006) and deletion mutants (Kubota, 2004) that became impaired in ethanol tolerance. In contrast, ethanol tolerance could be enhanced by disruption of URA7 or GAL6 genes (Yazawa, 2007) or by mutation in the global transcription machinery Spt15p (Alper, 2006). An alternative to mutation-based approaches is screens based on overexpression of genes. For example, overproduction of ubiquitin has been shown to enhance ethanol and osmotic stress tolerance (Chen & Piper, 1995), and overexpression of HAL1 (Gaxiola, 1992) increases tolerance to high salt. The possible involvement of tryptophan biosynthesis and tryptophan permease genes in ethanol tolerance was deduced from DNA microarray data and from ethanol sensitivity of corresponding knockout strains. Subsequent overexpression of these genes was shown to improve ethanol tolerance (Hirasawa, 2007).

In this study, instead of overexpressing individual genes, we screened a genomic expression library to identify genes that, upon overexpression, would enhance FST. The selection was set up such that gene(s) that prolong the survival of yeast during ethanolic fermentation, particularly at a high temperature, would be identified. We envisaged that such gene(s) could be used to engineer yeast strains that will retain their viability at a much higher level toward the end of fermentation. Such strains would allow reuse of cells during repeated rounds of fermentation, which is likely to make ethanol production more economical.

Materials and methods

Yeast strains and media

Saccharomyces cerevisiae FY3 (MATa ura3-52) (Winston, 1995) was used as a parent strain for all the strains constructed in this study. Standard molecular biology techniques were used as described previously (Ausubel, 2002) or as otherwise described below. A Ura+ control strain was made by transforming FY3 with pRS306 (Sikorski & Hieter, 1989) linearized within its URA3 region by StuI, thereby targeting it to the ura3-52 locus of FY3. A G418R control strain, for use in mixed culture experiments, was made by cloning a 1.5-kb (HindIII+BamHI) fragment of the kanMX module (Wach, 1994) in pRS306 and integrating this construct at the ura3-52 locus of the FY3 strain. Media used were SD (0.17% yeast nitrogen base without amino acids and ammonium sulfate, 0.5% ammonium sulfate and 2% glucose), YPD (1% yeast extract, 2% peptone and 2% glucose), YPD20 (1% yeast extract, 2% peptone and 20% glucose) and YPD25 (1% yeast extract, 2% peptone and 25% glucose). YPD20 and YPD25 were used in fermentation experiments carried out at 38 and 30 °C, respectively. All the media components were from Difco (BD); chemicals and reagents were procured either from Sigma or USB.


Fermentation was set up in 30 mL of YPD20, in 100-mL flasks closed with corks having syringe needle vents, at 38 °C, with shaking at 200 r.p.m. YPD25 was used instead of YPD20 for fermentation at 30 °C. The selected yeast strains were first outgrown for 24 h at 30 °C in 100 mL SD medium. The cells were harvested and made into a 50% cell suspension using sterile water; 300 μL of this suspension was transferred to 30 mL YPD20, i.e. to a starting cell density of 0.5% (w/v) wet weight of cells. The progress of fermentation was monitored in near real-time from measurements of weight loss due to CO2 evolution because ethanol and CO2 are produced at an equimolar ratio (Inlow, 1988; Varma, 1999).

Reuse of cells in fermentation

The first round of fermentation was set up with wild type (WT), RPI1 (Ras-cAMP pathway inhibitor 1) overexpression and delete strains in 30 mL YPD20 at 38 °C, as described above. After 36 h, when the fermentation had gone to completion, cells were harvested from the entire broth, and reused (without washing) in the second round of fermentation in 30 mL of fresh YPD20, at 38 °C. The progress of fermentation was monitored as described above.

Mixed culture fermentation

G418R control strain and G418S test strain were grown separately for 24 h at 30 °C in SD medium. Equal OD600 nm of these cultures were mixed, harvested and suspended in water to make a 50% cell suspension. YPD20 medium was inoculated with this suspension at a starting cell density of 0.5% (w/v) wet weight, and incubated at 38 or 30 °C. Various dilutions of fermentation broth were plated on YPD plates at different time points to determine the viability of cells. Colonies from these plates were then replicated onto YPD-G418 (200 μg mL−1) and YPD plates to determine the proportion of G418R control strains and G418S test strains surviving in the fermentation broth.

Ethanol estimation

Ethanol was enzymatically estimated from fermentation broth according to the manufacturer's instructions (Sigma). The cell-free supernatant (10 μL) of fermentation broth, collected at different time points, was diluted 100- or 1000-fold in sodium pyrophosphate buffer and used for ethanol estimation. The reaction was set up in microcentrifuge tubes, containing 50 mM sodium pyrophosphate buffer (pH 8.8), 7.5 mM β-NAD, 0.3 mM sodium phosphate, 0.003% bovine serum albumin, 100 μL of diluted samples and 4.5 U alcohol dehydrogenase in a final reaction volume of 1.1 mL, at 25 °C. The NADH reduced during the reaction was measured using a spectrophotometer at 340 nm, and the ethanol was calculated using a standard graph prepared with ethanol solutions of known concentrations.

Overexpression library screening

A yeast genomic expression library made in λADH (alcohol dehydrogenase) was obtained from ATCC (# 87288; λADH S. cerevisiae genomic library). λADH is a phage vector with excisable centromeric plasmid pADH, having a URA3 selection marker and a constitutive ADH1 promoter. The phage λ clones were converted to plasmid clones as described (Elledge, 1991), transformed into yeast FY3 strain and plated on five 22.5 × 22.5 cm plates. About 105 transformants were pooled together and outgrown for a day at 30 °C in 100 mL SD medium and fermentation was set up at 38 °C, as described above. After about 36 h of fermentation, by which time the viability of the population declines about 10-fold, the surviving cells from 3 mL of the broth were inoculated into 30 mL of fresh YPD20 medium. This was repeated for a total of five rounds of fermentation, after which 3 mL each of the two duplicate fermentation broths was plated onto SD plates to select for cells retaining the overexpression plasmids. Similarly, the unselected parent population of transformants was also plated on SD plates. The cells were pooled from these plates and inoculated separately into 100 mL SD medium to a starting OD600 nm of 0.5. After 24 h of growth at 30 °C, the cells were harvested and inoculated at 0.5% (w/v) wet weight of cells into YPD20 for fermentation at 38 °C to compare the selected population with the parent population. Total genomic DNA along with the plasmids was isolated from the selected population and transformed into the Escherichia coli strain. Plasmids from 90 independent clones were analyzed individually by restriction digestion with enzymes EcoRV, XbaI and XhoI. Twenty-five plasmids apparently having unique inserts were retransformed into FY3 strain; a strain transformed with pADH vector was used as the control. The individual transformants were checked for their survival during fermentation at 38 °C. The identity of DNA inserts in these plasmids was determined by DNA sequencing using vector-specific primers ADH-F (5′-TCTGCACAATATTTCAAGCT) and LacP (5′-CTTTATGCTTCCGGCTCGTATG).

Construction of overexpression and disruption strains

A 2.6-kb insert from one of the RPI1 clones (# 38) having the entire RPI1-coding region along with 0.6 kb of upstream and 0.8 kb of downstream regions was excised out from the plasmid with XhoI enzyme and cloned in pRS306 (Sikorski & Hieter, 1989) linearized with the same enzyme. This recombinant plasmid harboring the RPI1 gene (pRS-RPI1) was linearized within the URA3 gene by StuI digestion and integrated at the ura3-52 locus of the FY3 strain. To create disruption mutants, the same insert was also subcloned in vector pHSS8, and subjected to insertional mutagenesis using mTn-3xHA/lacZ, as described (Ross-Macdonald, 1997). Positive clones with transposon insertions were screened initially by NotI digestion, followed by sequencing using a transposon-specific primer (Tn3-S2; 5′-GGATCCTATCCATATGACGTTCCAG). An insertion mutant with integration at position 428 bp within ORF was used for further studies. The insert, along with the transposon insertion, was released from pHSS8 by NotI digestion and transformed into the FY3 strain.

MIG1 deletion

MIG1 deletion was performed by overlap extension PCR. Regions upstream and downstream of the MIG1-coding region were amplified using primers AGT-MIG1-S (5′-AGTGCCACTAACCTACATTAATCG) and MIG1-F-A (5′-GTCAGCGTAGGCTATGGTAGTATGTCGTC) for the upstream region, and MIG1-F-S (5′-ACCATAGCCTACGCTGACAAGTTTTTGG) and GAT-MIG1-A (5′-GATACCTGGTTCCATGAATGAGAG) for the downstream region. Fusion PCR was performed with upstream and downstream PCR products as a template and AGT-MIG1-S and GAT-MIG1-A as primers. This fusion product, which is deleted from the MIG1-coding region, was cloned in pRS306 at XhoI–XbaI sites using the DISEC/TRISEC strategy (Dietmaier, 1993). The recombinant plasmid was linearized at the SpeI site present within the downstream flank of MIG1 to target the corresponding region of the FY3 strain. Transformants were initially selected on SD plates, and then patched on YPD plates, followed by patching on 5-fluoroorotic acid (FOA) plates for selection of cells that have lost the integrated plasmid. Transformants growing on FOA plates were confirmed for deletion of the MIG1-coding region by PCR using primers AGT-MIG1-S and GAT-MIG1-A. Deletion of MIG1 was further confirmed by the ability of the delete strain to grow on raffinose in the presence of 2-deoxyglucose (Nehlin & Ronne, 1990). To construct an RPI1 overexpression strain with MIG1 deletion, the pRS-RPI1 plasmid was integrated at the ura3-52 locus of the MIG1-deleted strain.

RNA isolation and reverse transcription- quantitative PCR (RT-qPCR)

Yeast total RNA was isolated using the RNeasy mini kit, as per the manufacturer's instructions (Qiagen) or by a modified hot phenol method (Mannan, 2009). RNA was quantified using a Nanodrop spectrophotometer (ND-1000) by A260 nm and RNA integrity was assessed using a running denaturing agarose gel. For the Northern experiment, the blot was hybridized with an [α-32P] dATP-labeled RPI1 gene-specific probe, and rRNA was used as a loading control. RT-qPCR was performed with 100 ng of RNA samples using the one-step SYBR-I green kit (Invitrogen) using primers specific for RPI1 (5′-CAATGCCTATACAGCTACCGC and 5′-TGAGTCTGAAGTTGACATATTATGC), and IPP1 (5′-TACTTCCCAGGTCTGTTGAGG and 5′-AACAGAACCGGAGATGAAG). The PCR reaction (20 μL) was set up in 96-well plates (Axygen) in duplicate, and was analyzed using i-cycler iQ real-time detection system (Bio-Rad Laboratories). The specificity of the PCR product was checked by melt-curve analysis and further confirmed by running the samples on gel. All data analyses, including normalization and data pretreatment were performed with genex software (version 4.3.5, MultiD Analyses).

Zymolyase sensitivity assay

The zymolyase sensitivity of strains was tested in a microtiter plate format, as described (Ovalle, 1999), with some modifications. Yeast cells, equivalent to 0.8 OD600 nm, were harvested from broth after 24 h fermentation at 38 °C. The cells were pelleted, washed with water and resuspended in 250 μL zymolyase assay buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 5 mM EDTA and 4% PEG 8000) with 4 U (200 μg) of zymolyase 20T (Seikagaku Corporation, Japan). The plates were incubated at 30 °C with shaking (300 r.p.m.) in Thermomixer comfort (Eppendorf). The decrease in OD600 nm was measured periodically in a microplate reader and data were analyzed using kc junior software (Bio-Tek Instruments).

Calcofluor white sensitivity assay

Yeast cells, equivalent to 0.5 OD600 nm, were harvested from fermentation broth, as described for the zymolyase sensitivity assay, washed once with sterile water and resuspended in 200 μL water. This suspension was 10-fold serially diluted and 5 μL each of the dilutions was spotted on YPD plates containing different concentrations of calcofluor white (Sigma, catalog number F-3543). The growth of yeast cells was assessed after incubation at 30 °C for 2 days.

Results and discussion

Overexpression library screening

An overexpression strategy was followed to directly identify genes that would prolong the viability of yeast cells under fermentation conditions. Fermentation was carried out at 38 °C, which is much higher than what is optimal for yeast (30 °C). These conditions were chosen to identify genes that would improve the viability of yeast cells in the presence of ethanol and at a high temperature, and that are known to have synergistic deleterious effects on yeast (Piper, 1995). Sufficient glucose was provided in the fermentation broth such that at least 10% (v/v) ethanol is produced. Based on preliminary experiments, the duration of fermentation was chosen such that the fermentation goes to completion and the viability of the population as a whole would decline at least 10-fold toward the end, which was typically 36 h. The rate of fermentation was monitored by periodic measurements of weight loss due to CO2 evolution. As reported by others (Inlow, 1988; Varma, 1999), we have also observed a stoichiometric relationship between CO2 and ethanol production; from enzymatic estimations of ethanol produced by different strains at multiple time points, we found that the relation between ethanol production and weight loss is 0.9 g ethanol g−1 weight loss (Supporting Information, Fig. S1). Thus, periodic weighing of flasks to monitor weight loss served as a rapid method to follow the rate of fermentation. We anticipated that if the average viability of the population of transformants declines due to fermentation stress and if a plasmid-borne gene improves survival, then that particular transformant would become enriched with respect to other transformants. Thus, to select such transformants, the cells surviving at the end of fermentation were reused in the next round, and this was repeated for another five rounds, so at the end the selected transformants will comprise mostly those bearing genes conferring better viability. To check whether the selection has worked, the selected pool of transformants and the starting pool were grown under identical conditions, and subjected to fermentation. The results showed that while the viability of both the populations started to decline after 24 h of fermentation, the viability of the selected population declined much slower than the unselected population. The viability of the selected population was about 40-fold better than that of the unselected population at around 40 h, and about 700-fold better at around 50 h of fermentation, indicating that the selected population is enriched with yeast transformants that can withstand the harsh fermentation conditions much better than the parent strain (Fig. S2). However, the better survival seen could be due to plasmid-borne genes or chance chromosomal mutations. To check this, plasmids recovered from the selected population were retransformed into yeast, and tested under fermentation conditions. The retransformed yeast cells also showed much higher viability than the starting pool of cells during fermentation, confirming that the plasmid-borne genes indeed confer the phenotype. To identify the genes carried by these plasmids, plasmid DNA isolated from 90 independent E. coli clones were characterized by restriction enzyme digestion as described in the Materials and methods. Twenty-five plasmids that appeared unique on the basis of the digestion pattern were retransformed into yeast, and the transformants were individually monitored for rate of ethanol production and viability along with a control strain transformed with vector alone. Ten plasmids that conferred at least 20-fold higher viability compared with vector control after 40 h of fermentation at 38 °C were further characterized. The genomic DNA regions present in these plasmids were identified by sequencing and blast homology search.

Role of RPI1 in FST

All the inserts of the plasmids carried the RPI1/YIL119ccoding region, along with varying lengths of flanking regions (Fig. 1). The RPI1 gene was present in both orientations with respect to the ADH1 promoter of the vector, indicating that it is being expressed with its native promoter. Because the expression vector used was a centromeric plasmid, it appeared that a single additional copy of RPI1 was sufficient to improve survival. To confirm this further, RPI1 overexpression strain was made by integrating an additional copy of RPI1 gene along with its native promoter in the FY3 strain. A mutant disrupted in the RPI1 gene was also constructed by transposon mutagenesis, as described in Materials and methods. These strains were tested under fermentation conditions at 38 °C along with a WT control strain. After 36 h of fermentation, the strain with an additional copy of RPI1 showed >50-fold better viability, while the disruptant survived 100-fold less viability when compared with the control strain (Fig. 2a). However, the rate of fermentation of the RPI1 overexpression strain was slightly slower than that of the control strain and RPI1 disruptant (Fig. 2b). To test whether the effect of RPI1 is linked to a high temperature or fermentation per se, additional fermentation experiments were performed at 30 °C. The results are comparable to that of 38 °C (Fig. S3). Moreover, overexpression or disruption of RPI1 did not have any effect on the growth or the survival of strains in a standard YPD medium (data not shown). These results indicate that RPI1 is critical for the survival of yeast cells during ethanolic fermentation under both optimal and high-temperature conditions.


Schematic representation of RPI1 clones. The DNA fragments carried by various clones with respect to the RPI1 region of the genome are shown. The ADH1 promoter of the vector is shown (as arrowheads) to indicate the orientation of the RPI1-coding region with respect to this promoter.


RPI1 prolongs cell survival during fermentation. (a) Survival and (b) rate of ethanol production of WT (○), RPI1 overexpression (△) and disruption (▿) strains at 38°C. The results shown are means±SEMs of triplicate samples of one of two independent experiments.

The better survival of the RPI1 overexpression strain might be a consequence of a slower rate of fermentation, because this strain would experience less ethanol stress compared with other strains during fermentation. Thus, mixed culture experiments were performed to eliminate the complication of differences in the rate of fermentation and ethanol stress experienced. To distinguish between the strains in the mixed pool, a control WT strain of yeast was constructed by integrating a plasmid bearing kanMX4 module conferring G418 resistance (Wach, 1994) at the ura3-52 locus. The G418 resistance marker is known to exert only a small effect on growth rate (Baganz, 1997). We found that the survival of WT strains, with and without the G418R marker, when mixed together, was comparable during fermentation (Fig. 3), indicating that this marker does not influence the fitness of strains under these conditions. To test the relative fitness of RPI1 overexpression and disruption strains, these were independently mixed in an equal ratio with the G418R WT control strain and subjected to fermentation. The rate of fermentation was comparable in all the mixed culture fermentations (Fig. S4). In the mixed culture fermentation performed with the RPI1 overexpression strain and the G418R control strain, >95% of the surviving cells at the end of fermentation corresponded to the RPI1 overexpression strain, at both 30 and 38 °C (Fig. 3). The abundance of RPI1 disruption strain became <3% at the end of mixed culture fermentation with the WT strain. These results confirm that the higher survival conferred by RPI1 is not because of any difference in the rate of fermentation, but due to overexpression of the gene per se. The poor survival of the RPI1 disruption strain in individual (Fig. 2) and mixed fermentations (Fig. 3) confirms that this gene is critical for survival of cells during ethanolic fermentation.


RPI1 prolongs cell survival during mixed culture fermentation. Relative percentage abundance of WT, RPI1 overexpression (RPI1↑) and RPI1 disruption (rpi1Δ) strains during fermentation at (a) 38°C and (b) 30°C. Competitive fermentation experiment was performed by mixing each test strain with a WT control strain tagged with a G418 resistance marker. The abundance of the test strain, at different time points, is shown as a percent of the total CFU in the pool. Mean values of data from two independent experiments are shown.

Because the viability of the RPI1 overexpression strain at the end of fermentation was as much as that at the start of fermentation (Fig. 2a), we then checked whether the cells could be reused in the next round of fermentation. The cells of all three strains were harvested at the end of the first round of fermentation and reused in the next round, as described in Materials and methods. While the RPI1 overexpression strain started fermentation almost immediately, the WT strain took about 12 h and the RPI1 delete strain about 26 h to initiate fermentation (Fig. 4). Thus, strains engineered for overexpression of RPI1 are likely to be advantageous for the economical production of ethanol. However, improved economics of ethanol production can be properly assessed only in fermenters after considering the residence time for each round of fermentation, and the cost advantages accruing from reusing cells compared with preparing fresh inocula.


Reuse of cells in batch fermentation. Yeast cells recovered at the end of the first round of fermentation (36 h) were used for the next round of fermentation in a fresh YPD20 medium at 38°C. Data represent the rate of fermentation of WT (○), RPI1 overexpression (△) and disruption (▿) strains. Results are means±SEMs of two independent transformants, carried out in three replicates each.

RPI1 was initially reported as a dosage suppressor of the Ras cAMP pathway (Kim & Powers, 1991). It was also identified as a multicopy suppressor of glucose-induced loss of heat resistance in hsp104Δ-hxk2Δ-tps1Δ triple mutant of yeast (Versele & Thevelein, 2001). Later, it was identified as a high-copy-number suppressor of the cell lysis defect associated with loss of function of the MPK1 of the cell wall integrity pathway (Sobering, 2002). Rpi1p is a putative transcription factor, because it resides in the nucleus, possesses a C-terminal transcriptional activation domain and enhances the mRNA levels of a subset of genes involved in cell wall metabolism (Sobering, 2002). Although RPI1 was reported to aid the cells in preparation for the stationary phase, this is only seen in the mpk1 deletion mutant. While incubation of cultures for 14 h beyond the logarithmic growth phase resulted in a viability loss of 4 log units in the mpk1Δrpi1Δ double mutant and 1 log unit in the mpk1Δ mutant, no viability loss was observed in the rpi1Δ mutant even after 48 h in culture (Sobering, 2002). On the other hand, the rpi1Δ mutant rapidly loses viability under fermentation conditions, and it is about 100-fold less viable compared with the WT, after 36 h of fermentation at 38 °C (Fig. 2). Thus, our results indicate a novel role for RPI1 in FST. The role of RPI1 during fermentation is more definitive than in the stationary phase, because the survival of the strain mutated only in RPI1 is highly impaired in viability during ethanolic fermentation, but is comparable to WT in the stationary phase.

Improving the fermentation efficiency of RPI1 overexpression strain

A faster rate of fermentation is a desired feature of yeast strains used for ethanol production. MIG1 is known to mediate glucose repression, and its disruption results in derepression of a large number of genes, including glucose transporters HXT2 and HXT4 and glucose sensor SNF3 (Klein, 1998; Johnston, 1999). Thus, we speculated that disruption of MIG1 might increase the fermentation rate. Moreover, disruption of MIG1 is also known to aid in simultaneous utilization of several sugars present in complex media such as molasses (Klein, 1998). To test our hypothesis, we deleted MIG1 in the WT and RPI1 overexpression strains. Fermentation studies at 38 °C showed that MIG1 deletion increased the rate of fermentation in the RPI1 overexpression strain to nearly WT level (Fig. 5a). However, at the end of fermentation (38 h), the viability of the RPI1 overexpression strain with MIG1 deletion was fivefold less than that of the RPI1 overexpression strain, although it was still 20-fold better than that of the WT strain (Fig. 5b). Deletion of MIG1 in the WT strain did not affect its fermentation rate (Fig. 5a), but reduced its viability by about 13-fold (Fig. 5b). Compiled data in the YEASTRACT database (http://www.yeastract.com/; Teixeira, 2006) show that 170 genes are regulated by MIG1. Using SGD gene ontology [go] slim mapper (http://www.yeastgenome.org/cgi-bin/GO/goSlimMapper.pl), these genes could be assigned to over 35 biological processes (Yeast GO:Slim-process). These include 32 genes that are annotated to the GO term ‘transport’ and 25 genes that are annotated to the term ‘stress response.’ Thus, deletion of MIG1 is expected to have pleiotropic effects on yeast, and the increase in the fermentation rate and the decline in survival could be two independent consequences of MIG1 deletion in the RPI1 overexpression strain. As the survival of this strain is much less than that of the RPI1 overexpression strain at the end of fermentation, it was not used in further studies.


Effect of MIG1 deletion on fermentation efficiency and survival. (a) Fermentation rate and (b) survival of WT (○), RPI1 overexpression (△), MIG1 deletion (▿) and RPI1 overexpression–MIG1 deletion (◊) strains at 38°C. Mean values of data from two independent experiments are shown.

Expression level of the RPI1 gene

It was surprising that introducing an additional copy of RPI1 dramatically prolonged the survival of strains during fermentation. To gain further insight into this, we checked the expression level of RPI1 gene by a Northern blot. The RPI1 transcript level was always higher in the overexpression strain compared with the control WT strain, for all the time points tested during fermentation (results not shown). The hybridization signal for WT at earlier time points was close to the background, and hence quantification of the relative levels of expression was difficult. Thus, reverse transcription qPCR was performed to accurately determine the relative RPI1 transcript levels. The results showed that RPI1 transcripts increased by about 2.5-fold during the course of fermentation in both the WT and the overexpression strain, which is consistent with its role in FST. Moreover, for all the time points tested, the RPI1 transcript was at least threefold higher in the overexpression strain compared with the WT strain (Fig. 6). RPI1 shares its 1875-bp upstream region with the divergently transcribed RHO3 gene (Saccharomyces Genome Database; http://www.yeastgenome.org/). Moreover, the regulation of RPI1 appears to be complex, because at least 31 different transcription factors have been documented to bind within the 1-kb region upstream of RPI1 using the ‘Search Transcription Factors’ tool of the YEASTRACT database (http://www.yeastract.com/; Teixeira, 2006). However, only a 599-bp upstream region is retained in the 2.6-kb region of the RPI1 gene that was subcloned in pRS306, and integrated at the URA3 locus in the overexpression strain. We examined the binding sites of Nrg1p, in particular, as it is a known repressor of RPI1 expression (Berkey, 2004; Vyas, 2005). It has binding sites at positions −1571, −1086 and −731, but none within the 599-bp region (Berkey, 2004). Thus, the lack of binding sites for the Nrg1p repressor could perhaps explain the slightly higher level of RPI1 expression than that predicted by the copy number. This observation is further corroborated by RT-qPCR analysis of the RPI1 expression level in an nrg1 delete strain, which was found to be 2.4-fold higher than that in the WT strain (results not shown).


Expression level of RPI1 during fermentation. Transcript levels of the RPI1 gene were measured in WT and RPI1 overexpression strains by RT-qPCR normalized with IPP1 levels and shown as fold change. Results are means±SEMs of two independent experiments, performed with two replicates each.

RPI1 mediates its phenotype through cell wall modification

Stationary-phase cells of yeast are resistant to various environmental stresses such as heat, ethanol and cell wall lytic enzymes (Werner-Washburne, 1993), which could partly be due to their thicker and less porous cell wall compared with that of cells in the exponential phase (de Nobel, 1990). Sobering (2002) have suggested that RPI1 prepares the cells for the stationary phase by modulating the expression of cell wall biosynthetic genes. Thus, it is possible that even during ethanolic fermentation, RPI1 modulates cell wall integrity and thereby enhances the survival of yeast cells. To examine this, we recovered cells from the 24-h fermentation broth and tested them for sensitivity to widely used cell wall-perturbing agents calcofluor white and zymolyase. While the RPI1 overexpression strain showed a slight increase in resistance to calcofluor white, the resistance of the disruption strain was comparable to that of the WT strain (Fig. 7a and b). However, the RPI1 overexpression strain was highly resistant to zymolyase treatment compared with the WT strain, while the RPI1 disruption mutant was more sensitive than the WT strain (Fig. 7c). These results indicate that RPI1 mediates FST, at least partly, through cell wall modification. It appears that some stress tolerance pathways important for resistance during the stationary phase (Herman, 2002) are also critical for survival of cells during ethanolic fermentation.


RPI1 fortifies the cell wall during fermentation. Calcofluor white sensitivity (a and b) of WT, RPI1 overexpression (RPI1↑) and disruption (rpi1Δ) strains. Yeast cells harvested after 24 h of fermentation at 38°C were washed with sterile water, and spotted on YPD agar plates without (a) or with (b) 7.5 μg of calcofluor white. Images were captured after incubating the plates at 30°C for 2 days. One representative result from three independent experiments is shown. (c) Zymolyase sensitivity of WT (○), RPI1 overexpression (△) and disruption (▿) strains. Results are means±SEMs of three independent experiments, performed with two replicates each.

Earlier, several genes were reported as critical for growth in an ethanol-containing medium from a screen of transposon disruption mutants or deletion mutants (Takahashi, 2001; van Voorst, 2006). A recent high-resolution quantitative analysis of the growth behavior of yeast deletion mutants under ethanol stress has revealed that hundreds of genes are critical for ethanol tolerance (Yoshikawa, 2009). However, RPI1 was not found in these screens. Besides, overexpression or disruption of RPI1 did not affect ethanol tolerance (results not shown), when tested as described (Takahashi, 2001; van Voorst, 2006). RPI1 is also not required for growth/survival at 38 °C (results not shown). Thus, it is likely that the role of RPI1 becomes evident only under fermentation conditions when cells experience a combination of stress conditions. Another possibility is that RPI1 mediates tolerance to a specific, as yet unexplored, stress encountered by yeast cells during ethanolic fermentation.

In summary, we report the identification of RPI1 from an overexpression screen performed to discover genes that would prolong the survival of cells during ethanolic fermentation at 38 °C. Our finding confers a novel biological role to RPI1 in stress tolerance during ethanolic fermentation. This gene is much more important for FST than for the stationary phase, because the rpi1 mutant rapidly loses viability during fermentation, but not in the stationary phase. Overexpression of this gene improves viability, at least partly, by modulating cell wall integrity. Because overexpression of RPI1 prolongs the survival of yeast cells during fermentation, cells can be reused in the subsequent round of fermentation, which is likely to make ethanol production more economical.

Authors' contribution

R.P., M.A.M. and R.C.-D. contributed equally to this work.

Supporting Information

Fig. S1. Correlation between ethanolestimation by weight-loss and enzymatic methods.

Fig. S2. Overexpression library screening.

Fig. S3. Essential role of RPI1 during fermentation at 30°C.

Fig. S4. Fermentation rate during mixed culture fermentation.

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We thank P. Ross-Macdonald and M. Snyder for mTn-3xHA/lacZ mutagenesis strains, A. Wach for plasmid pFA6-KanMX4, F. Winston for yeast strain FY3 and Anand K. Bachhawat for helpful discussions during the course of this work. R.P., M.A.M. and R.C.-D. acknowledge the Council of Scientific and Industrial Research, New Delhi, for fellowships. This work was supported by an intramural grant from the Institute of Microbial Technology.


  • Editor: Isak Pretorius


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