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Yeast Cellular Stress: Impacts on Bioethanol Production

School of Pharmacy and Biomolecular Sciences, University of Brighton, Huxley Building, Lewes Road, Brighton BN2 4GJ, UK
Author to whom correspondence should be addressed.
Fermentation 2020, 6(4), 109;
Submission received: 24 May 2020 / Revised: 8 November 2020 / Accepted: 10 November 2020 / Published: 13 November 2020


Bioethanol is the largest biotechnology product and the most dominant biofuel globally. Saccharomyces cerevisiae is the most favored microorganism employed for its industrial production. However, obtaining maximum yields from an ethanol fermentation remains a technical challenge, since cellular stresses detrimentally impact on the efficiency of yeast cell growth and metabolism. Ethanol fermentation stresses potentially include osmotic, chaotropic, oxidative, and heat stress, as well as shifts in pH. Well-developed stress responses and tolerance mechanisms make S. cerevisiae industrious, with bioprocessing techniques also being deployed at industrial scale for the optimization of fermentation parameters and the effective management of inhibition issues. Overlap exists between yeast responses to different forms of stress. This review outlines yeast fermentation stresses and known mechanisms conferring stress tolerance, with their further elucidation and improvement possessing the potential to improve fermentation efficiency.

1. Introduction

The biotechnological potential of Saccharomyces cerevisiae has been exploited traditionally for the purposes of baking, brewing, and wine making. This species has also been incredibly useful in basic biochemistry and genetics studies [1]. Yeasts can be found thriving in diverse sugar-rich habitats and can be considered to be microbial weed species due to their dominant, biocide producing nature [2]. Its fermentative nature evolved between 125 and 150 million years ago and is shared by many yeast species, but the acquisition of ethanol tolerance by S. cerevisiae is a more recent event occurring after the whole genome duplication event, which is believed to have happened 100 million years ago [3,4]. This ~6000 gene-containing unicellular eukaryote was the first organism to have its genome fully sequenced [5,6]. Today it remains an indispensable model system for experimental studies and a mainstay of biotechnology. The curated Saccharomyces Genome Database (SGD; contains vast amounts of information for this microorganism, obtained through “omics” technologies, traditional biochemistry and molecular biology techniques [7]. Yeast continues to play a pivotal role in the study of fundamental biological processes including aging, cell cycle and stress, as a model organism and in emerging biotechnological areas including drug screening, computational and systems biology—see, for example [8,9,10,11,12,13,14,15,16,17,18,19,20]. The high genetic tractability of this microorganism allows for the manipulation of metabolic pathways; for example biofuel researchers often minimize carbon fluxes to natural metabolic by-products to maximize ethanol productivity (see also Section 3.1) [21,22].
The displacement of conventional transport fuels with biofuels including high octane number bioethanol increases fuel security, whilst also being an effective strategy for reducing transport sector emissions [23]. First-generation bioethanol remains the most dominant biofuel globally, yet policy now favors adoption of second-generation or advanced biofuels due to their higher sustainability rating [24]. First-generation bioethanol production generally involves the pre-processing of food-feedstocks, notably including sugarcane, corn, wheat, for the efficient extraction of contained fermentable sugars like glucose [25,26]. Sugarcane bioethanol production is particularly prevalent in Brazil, a global pioneer for transportation ethanol use [27]. However large-scale bioethanol production previously caused a spike in global food prices, triggering the so-called food-versus-fuel debate [28,29,30]. Non-food sources, including waste lignocellulosic biomass, are utilized for the production of second-generation bioethanol, yielding pentose and hexose sugars whilst possessing a higher sustainability rating. Higher associated cost and energy pre-processing steps currently render commercial production of second-generation bioethanol unviable [31]. Furthermore, pre-processing steps associated with second-generation bioethanol feedstocks may yield highly inhibitory (chaotropic) by-products including vanillin, not found or generated in first-generation bioethanol fermentations [32].
Due to its industrious, high ethanol productivity, high ethanol tolerance and ability to ferment a wide range of sugars, S. cerevisiae is the principal yeast employed for global bioethanol production [21,33]. Obtaining maximum ethanol yields remains a technological challenge however, with contaminant microorganisms, feedstock impurities, undesired side products and cellular stress impacting on yeast ethanol productivity [34,35]. S. cerevisiae is “stressed” sequentially and in concert throughout an ethanol fermentation [32]. On occasion, these inhibitory factors, notably ethanol-induced inhibition, but also contamination, can make an ethanol fermentation become “stuck” [36,37]. The provision of commercially available, higher ethanol tolerance yeasts to “restart” fermentation can be required, demonstrating the necessity for deploying a highly stress tolerant yeast for industrial fuel ethanol production [38].
Here, we provide an overview of the major stresses encountered by yeast cells during fermentations and the responses those cells make to mitigate these stresses. We pay particular attention to the production of ethanol by fermentations in the biofuel industry. Many of the processes are also relevant to other fermentations carried out by yeast, for example the production of other chemicals or the use of yeast as a host for the production of recombinant proteins. We also largely focus on the budding yeast S. cerevisiae. We do this partly because this organism is so well studied, partly due to its significant role in the biotechnology industry and also because studies in this organism tend to be generalizable to other unicellular fungi. The roles and functions of the proteins mentioned in this review are summarized in Table 1.

2. Yeast Stress Responses

Fluctuating environments affect all living organisms. Yeast’s response to fermentation associated stress conditions includes the switching of growth and metabolic strategies, including the synthesis of stress proteins functioning to mitigate cell damage and stress. These responses are is coordinated by cell stress stimulus sensing and signal transduction, triggering downstream transcriptional and post-translational responses [64]. The stress response elements (STRE; consensus CCCCT) are activated during numerous stress conditions including oxidative, heat shock, nitrogen starvation, low external pH, weak lipophilic acids and ethanol [65,66,67,68,69,70]. Yeast when subjected to ethanol stress initially struggles to maintain energy production. This compromised energy production leads to the increased expression of genes associated with energy metabolism [71]. While stress is often perceived as detrimental, which it often is doing industrial fermentations, it should also be remembered that environmental challenges and stresses can improve evolutionary fitness and drive the selection of more robust traits, strains and species [72,73].
The genetics underlying stress tolerance have been extensively investigated in S. cerevisiae and genetic and metabolic improvements having been attempted with success [68,69,70,71,74,75,76,77,78,79,80,81]. It is common for genes found to be up-regulated in the presence of ethanol stress to be targets for genetic engineering approaches [82]. A common experimental approach for the eventual isolation of desirable industrial traits including high stress tolerance is directed evolution. Selective pressures imposed on microorganisms promote the eventual emergence of traits that provide a selective advantage [83,84,85]. Repetitive batch cultivations in the presence of lignocellulosic hydrolysates for example evolved strains displaying increased tolerance to all inhibitors present [86]. Evolutionary methods are essentially “blind”, but they enable tolerance to develop through changes in many genes, including ones whose functions are not fully known. Alternatively, more rational methods might be employed. This would involve identifying systems which are known to be involved with stress tolerance and making specific changes which aim to enhance these functions. For example, ethanol tolerance improvement could be achieved through approaches including enhancing the unfolded protein response (UPR), or engineering the enzymes involved in the biosynthesis of cell membrane lipids [87,88,89,90]. Tools including genome shuffling and global transcription machinery engineering have successfully bred stress tolerant yeast [91,92,93,94,95]. The emergence of powerful technologies including CRISPR-Cas9 should also allow for the better engineering of industrially relevant phenotypic traits [96,97].

2.1. Compatible Solutes or Inert Osmolytes

Many microorganisms produce, or increase production of, small molecules in response to external stresses. These compatible solutes are typically hydrophilic and facilitate the protection of biological macromolecules and cells [98,99]. The main yeast compatible solutes are glycerol, proline and trehalose (Figure 1). With the exception of glycerol, these compounds are kosmotropic and promote the ordering of biological macromolecules and assemblies, see Section 3.3 [100]. The physical biochemistry underpinning the mechanism of stabilization has not been fully elucidated [101]. It is likely to involve both direct and indirect effects [102]. Some compatible solutes bind to, and stabilize, biological macromolecules [103]. Kosmotropes generally act indirectly by increasing the ordering of water molecules, i.e., reducing the entropy of the system [104,105]. This enables them to “neutralize” the effects of chaotropes such as ethanol and to protect biological macromolecules from thermal, acid, or alkali denaturation. In the case of osmotic stress, they increase the osmolarity of the cytoplasm and thus help equalize the osmotic pressure on both sides of the plasma membrane [106].
Glycerol is a three carbon, trihydroxy alcohol. It is a by-product of ethanol fermentation in S. cerevisiae, having a role in maintaining cellular redox balance, with imbalances leading to oxidative stress [107]. Glycerol production also prevents osmotic-induced oxidative stress, caused by the intracellular accumulation of reactive oxygen species (ROS) [108]. Aquaporins and glycerol facilitators allow for the rapid export of glycerol for the conditioning of the external cellular environment. It is important in the adaptation to low osmolarity, due to its stabilizing effects on cellular components [106].
Proline is an imino (secondary amino) acid which is synthesized in yeast from glutamate by the actions of the enzymes Pro1p, Pro2p and Pro3p [55]. Unlike glycerol, proline biosynthesis does not appear to be upregulated in response to stress. Natural levels may contribute to stress resistance and strains engineered to produce more, or consume less, proline, show greater tolerance to a range of stresses [109]. Trehelose is a disaccharide of two glucose subunits joined by an α (1→1) glycosidic bond. It is synthesized in yeast from UDP-glucose and glucose 6-phosphate by the enzymes Tps1p and Tps2p [110]. The genes encoding these enzymes have stress response elements upstream ensuring then they are coregulated [111]. The transcription factors responsible for activating the genes encoding these enzymes, along with others regulated by STREs, are Msn2p and Msn4p [50]. These proteins translocate to the nucleus under stress conditions [74,112]. Trehalose itself assists in the activation of the heat shock response via the transcription factor Hsf1p [113].

2.2. The Unfolded Protein Response (UPR)

The unfolded protein response facilitates generic, cell wide responses to stresses which results in disruptions to “proteostasis” or the accumulation of unfolded proteins in the endoplasmic reticulum [114]. The UPR also affects general processes related to secretory pathway homeostasis and therefore plays a maintenance role in yeast cell wall integrity [115]. Its rapid initiation is ensured by cells constitutively expressing mRNA encoding Hac1p [116]. In the non-stressed state this mRNA is maintained in the unspliced form which is not translationally active. Increased levels of unfolded protein are sensed by the chaperone Kar2p. This causes it to stop binding Ire1p, a protein with both nuclease and kinase activities [117,118,119,120]. Ire1p can also directly detect unfolded proteins and damaged phospholipids [121,122]. HAC1 mRNA is spliced by Ire1p and the RNA ligase Trl1p [116,120,123]. It is then translated to produce the transcription factor Hac1p which then translocates to the nucleus where it binds to the UPR response element. This sequence lies upstream from genes which are regulated by the UPR [124]. Hac1p then recruits RNA polymerase and other proteins required for transcriptional activation and increases the expression of a range of genes which code for proteins such as chaperones, and enzymes for phospholipid biosynthesis, cell wall biogenesis, DNA repair and aerobic metabolism [117,122,125,126,127]. They precise subset of genes upregulated depends partly on the conditions giving rise to stress [122,128,129]. Cell wall stress also activates the broad transcriptional response of the UPR through the mitogen-activated protein (MAP) kinase cascade, with ER stress also activating the CWI pathway, demonstrating that the UPR and CWI are co-ordinately regulated in Saccharomyces cerevisiae in order to protect cells against related stressors [130].

3. Yeast Stresses and Their Mitigation

3.1. Osmotic Stress

High initial fermentable sugar concentrations is particularly pertinent to first-generation bioethanol production, with high initial external osmolarity in an alcoholic fermentation facilitating the passive diffusion of cellular water down the concentration gradient, leading to hyperosmotic stress [106]. Effects include rapid decrease of cell volume and turgor pressure, with the concurrent generation of reactive-oxygen species (ROS), leading to redox state imbalances and oxidative stress. Oxidative stress and osmotic stress differ significantly but display overlapping responses with the robust yeast continuing to proliferate over a range of external water activities (aW) [108]. Intracellular volume and water balance are tightly regulated through cell osmoregulation, for the proper maintenance and functioning of biochemical and biological processes [66].
Interest in the molecular mechanisms of yeast osmoadaptation originated from the need to improve the performance of yeast strains under industrial conditions. Hyperosmotic stress disrupts cytoskeleton structure, causes the remodeling of chromatin and can even lead to cell cycle arrest and apoptosis [131,132,133]. An intracellular shift in metabolism results, mediated by the high-osmolarity-glycerol (HOG) pathway and cell wall integrity (CWI) pathways, producing glycerol as a compatible solute and providing cell surface stability respectively, with these processes being co-coordinatively linked [66,115,134]. This cross talk between the two pathways suggests the evolution of highly coordinated stress responses.
The CWI pathway responds to damage to the cell wall. Extracellular domains of transmembrane receptors sense chemicals which may damage the cell wall [135]. The pathway also responds to osmotic, ethanol and pH stress. How these stresses are sensed is currently unknown [136]. Following sensing of cell wall associated stress, a signaling pathway connects the cell membrane with the MAP kinase Slt2p [136,137]. This kinase then activates two transcription factors, Swi4p/Swi6p and Rlm1p [57,138]. The first of these normally regulates cell cycle progression, but in this case regulates some genes involved in cell wall biosynthesis [139,140]. Rlm1p controls the expression of at least 80 genes including those which code for enzymes involved in the synthesis of cell wall components [141,142]. The HOG pathway also responds to cell surface receptors activating a signaling pathway which culminates in the activation of the MAP kinase Hog1p [46,66,143]. Phosphorylated Hog1p acts both in the nucleus and the cytoplasm. In the nucleus it phosphorylates and activates transcription factors which control the expression of over 600 genes [144,145]. This includes genes responsible for controlling the production of compatible solutes and the pausing of the cell cycle in G2 phase [146]. In the cytoplasm Hog1p directly reduces the export of glycerol thus increasing the cellular concentration of this compatible solute [147,148].

3.2. Heat Stress

Stress due to high temperatures can be encountered in ethanol fermentations, particularly in the early stages or if the external temperature is higher than the optimum for the fermentation [149,150]. Excess temperatures also increase sensitivity to ethanol [151]. A brief discussion of the effects of, and remedies for, heat stress is included here due to their similarity with chaotrope stress, see Section 3.3 [65]. High temperatures represent a general threat to living systems. Proteins, nucleic acids and phospholipid bilayers are denatured by increased temperature. Therefore, cells respond by counteracting this. Chaperone protein expression is upregulated In the so called heat shock response [152]. These proteins assist folding of nascent proteins and those which become partially unfolded as a result of denaturation by temperature and other stresses. Membrane composition is altered. Shorter and unsaturated fatty acids in phospholipids are replaced by longer and less saturated ones [153]. Both longer and less saturated fatty acids have higher melting temperatures and thus increase membrane rigidity. This is achieved by the expression of enzymes which extend the number of carbon atoms in fatty acids, e.g., Elo1p [43]. The amount of ergosterol in the membranes also increases [154]. Ergosterol is the predominant sterol in yeast membranes; increasing the mole fraction of this sterol in the membrane causes it to become more rigid and promotes the formation of lipid rafts [155,156,157]. Some strains which have been artificially evolved to have enhanced long term thermotolerance produce fecosterol in place of ergosterol which further rigidifies the membrane (Figure 2) [158]. In industrial fermentations, heat stress can be mitigated by thermostatic control of the reaction vessels. The use of thermotolerant strains also reduces the negative effects of heat stress [159,160,161]. Given the similarities between heat and chaotrope stress these strains are also likely to have improved tolerance to high ethanol concentrations [65,162,163]. Alternatively, additives to the fermentation mix, such as magnesium sulphate, can improve thermotolerance [164].

3.3. Chaotrope Stress

Chaotropes are compounds which cause the disordering of biological macromolecules such as proteins and nucleic acids, as well as the disruption of biological assemblies held together by non-covalent interactions such as cellular membranes [100,165]. Many chaotropes do not to interact directly with the macromolecules and assemblies. Instead, they disrupt the hydrogen bonding networks in the solvent (water) thus increasing the overall entropy of the system [166,167]. However, the mechanism of chaotropicity is likely to vary between different compounds [165,168]. Since chaotropes disrupt critical cellular macromolecules and assemblies they exert a detrimental effect on biological systems. They affect key features of proteostasis and impose a dosage dependant “fitness cost” due to the “toxicity” of misfolded proteins [169,170]. The products of ethanol fermentations are chaotropic. Thus, the production of alcohol will ultimately be self-limiting as there will come a point where the concentration of the chaotropic compound exceeds the cell’s ability to mitigate it [32]. These mitigations are similar to those seen in high temperatures stress [65]. The ER’s quality control system, involving the unfolded protein response becomes activated [87]. This leads to increased expression of ER-located molecular chaperones whilst also aiding in the coordinated transition from fermentative metabolism to slower mitochondrial respiration (post-diauxic shift) during starvation [171]. The fatty acids in membrane phospholipids become longer and less saturated, catalyzed by Elo1p and Ole1p respectively. Compatible solutes such as glycerol and trehelose are produced [172].
Ethanol, a chaotropic solute, is utilized commercially as a protein denaturant, with chaotropic agents in general entropically disordering macromolecular systems, adversely affecting cell constituents including phospholipid bilayers, proteins, and other hydrated cell components [100]. The chaotropic activity of common fermentation products including ethanol (+5.93 kJ kg−1 mol−1) and some by-products of lignocellulose pre-treatment including vanillin (+174 kJ kg−1 mol−1), have been quantified using the agar gelation-temperature model system [100]. The growth rates susceptibility of S. cerevisiae to these substances has also been determined [173,174,175]. Chaotropes, including the aliphatic alcohols, cross and perturb cellular membranes in the high millimolar to molar range leading to a reduction in cell viability, slower growth/proliferation and lower obtainable ethanol yields [37,76,173].
Mitigation of chaotropicity in industrial settings is difficult to achieve. If a fermentation runs to completion, the alcohol poisons the cells. Furthermore, the production of glycerol, which is miscible with ethanol, presents an additional problem since the ethanol must be purified from this mixture. Kosmotropes are they opposite of chaotropes. They cause ordering and rigidity in biological macromolecules and phospholipid membranes. Typically, they reinforce the hydrogen bonding networks in the solvent, reducing the overall entropy of the system [176]. In theory, kosmotropes could be added to fermentation mixes to alleviate chaotrope stress. Indeed, the compatible solutes trehelose and proline are kosmotropic [100]. Glycerol presents an intriguing paradox. It is a compatible solute produced in response to stress by many microorganisms. However, in the agar gel setting assay it was shown to be mildly chaotropic [100]. Other studies suggest glycerol can enhance the hydrogen bonding network in water—a kosmotropic attribute [177,178].
There are a number of issues with using kosmotropes to neutralise chaotrope stress. First, some kosmotropes are themselves toxic to yeast cells and impair efficient growth and fermentation [175]. Second, there is currently no method of rapidly measuring or predicting the net solution chaotropicity. So, knowing the concentration of kosmotrope required to neutralize a given concentration of ethanol is currently impossible [179]. Third, the addition of chemicals to the fermentation mix will alter they osmolarity of the medium and may cause osmotic stress in the yeast. Fourth, the additional cost of purchasing kosmotropic compounds and, potentially, separating them from the ethanol at the end of the process may render this strategy nonviable.

3.4. Other Stresses

Stresses do not occur in isolation. Almost all instances of compounds which cause one form of stress also induce others [32,101]. For example, ethanol causes both osmotic and chaotropic stress [99]. In the context of ethanol fermentations, oxidative, and pH stress are less commonly encountered or occur as a consequence of other stresses.
Oxidative stress can be caused by chaotropic and osmotic stress during fermentations. Both can cause the buildup of reactive oxygen species. These free radicals cause non-specific damage to biomacromolecules and membrane lipids [180]. Uncorrected damage to DNA can cause hereditable mutations which are likely to be deleterious. Reactive oxygen species can also cause lipid peroxidation which damages cell membranes [181]. Yeast cells have a complex, integrated responses to free radical production [182]. This includes enzymes which repair double and single strand breaks in DNA, which reverse lipid peroxidation and which directly reduce reactive oxygen species [183,184,185,186].
Yeast cells normally condition the growth media to a slightly acidic pH. This requires the export of protons across the cell membrane by the plasma-membrane H+-ATPase, Pma1p [52]. Thus, alkaline conditions represent an external stress for yeast with growth being slow or undetectable above pH 8.2 [187,188]. Multiple systems respond to alkaline stress [188]. One key pathway involves the calcium activated phosphatase calcineurin complex which is comprised of one of two catalytic subunits, Cna1p or Cmp2p, and one regulatory subunit, Cnb1p [40,41,189]. This activates the transcription factor Crz1p which controls the expression of stress related genes [190]. In addition, signaling through protein kinase A is transiently inhibited leading to the nuclear localization of the transcription factors Msn2p and Msn4p which bind to stress response elements, see above [191]. One consequence of alkaline stress is a reduction in mRNA transcription rates and in the stability of mRNA molecules [192]. A key response is the upregulation of the ATP dependent sodium ion transporter Ena1p [193]. This is regulated by both the calcineurin and protein kinase A pathways [194]. It is not immediately obvious why the export of sodium ions into the alkaline medium would mitigate the stress caused by excess hydroxide irons. It is generally considered that, unlike many mammalian sodium transporters, Ena1p does not cotransport protons although some early evidence suggests that it may do so [195]. The export of sodium ions may also assist in maintaining the membrane potential under conditions where protons are likely to be consumed.
Excessively acidic conditions also cause stress in yeast. Weak acids such as acetic acid, citric acid and boric acid can inhibit yeast growth and induce apoptosis [196]. These compounds are transported out of the cell by multidrug resistance transporters [197,198]. Pma1p activity is increased in order to transport excess protons out of the cell [199]. Acidification of the external medium results in a corresponding decrease in the pH of the vacuole suggesting that the cell transports excess protons to this compartment in order to maintain cytoplasmic pH [199].
In fermentations, the pH can be monitored and controlled by inline sensors and the addition of appropriate amounts of acid or base. The amounts can be determined either by feedback control or estimation using the Henderson Hasselbalch equation.

4. Conclusions

S. cerevisiae is considered a robust and versatile microorganism and has become an outstanding model system for the elucidation of Eukaryotic cell stress biology. Responses to each form of stress typically involve complex interactions of signaling pathways and transcription factors. This can affect the expression how many different genes. Although genomic and proteomic studies have revealed many candidate genes and proteins for the mitigation of various forms of stress, unravelling and understanding these data is challenging. In particular, understanding which genes are critical for response compared to those whose expression changes consequentially (e.g., many stress responses require energy and so ATP generating metabolic pathway enzymes are likely to be upregulated as a consequence). Distinguishing between those changes which make a minor contribution to stress responses compared to more significant contributions is critical. Here is should be noted that fold changes in expression levels do not provide a reliable guide as to which changes are the most important. This can only be determined by further biochemical experiments, for example assessing the response of strains with key genes deleted. The major players are likely to be those which act in systems which have been identified by traditional molecular biology approaches to be involved in stress remediation. The heat shock response, for example, produces coordinated sets of actions which mitigate the effects of heat on cellular components. Unsurprisingly, elements of this response are also used in adaptations to other forms of stress.
Understanding these responses is critical to enabling rational efforts to engineer strains of yeast which are better adapted to stressful conditions. Inevitably, there must be limits to the amount and duration of stress which yeast can endure. These limits will ultimately impose maximum yields in ethanol fermentations and determine whether, or not, it will be economically viable to transition away from fossil fuels towards sustainable biofuels.
Cellular stresses incurred by yeast throughout a bioethanol fermentation impact on cell growth and fermentative metabolism, reducing final obtainable ethanol titers. The plethora of general and stress-specific response and tolerance mechanisms makes yeast an industrially applicable microorganism. There are two main strategies for minimalizing the effects of stress on an ethanol fermentation. First, we can minimize the stress caused to the yeast. For example, temperatures and pH values of fermentations can be strictly control. To do this requires an understanding of the situations which cause the yeast cells to be stressed and thus reduce ethanol productivity. Second, we can select or engineer strains which are resistant to stresses. That many stress responses overlap assists us in this process. For example, yeast which are highly resistant to temperature stress are also likely to be able to resist high concentrations of ethanol. Again, this requires a knowledge of the biochemistry of stress sensing, stress responses and of unmitigated stresses. Therefore, it is clear that continued research efforts to understand the biochemistry of stress in yeast is vital for improving the economic and environmental sustainability of the biofuel industry.

Author Contributions

J.E.—initial draft of Section 1, Section 2, Section 2.1, Section 3.1, and Section 3.3; editing of entire manuscript; D.J.T.—initial draft of Section 2.2, Section 3.2, Section 3.4, and Section 4; editing of entire manuscript; figures and table. Both authors have read and agreed to the published version of the manuscript.


J.E. thanks the University of Brighton and the Universities Alliance Doctoral Training Alliance (Energy) for a PhD studentship.


The authors thank MDPI for waiving the Open Access fee for this article. We thank Samantha Banford for her assistance with the revised version of the paper.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Barnett, J.A. Beginnings of microbiology and biochemistry: The contribution of yeast research. Microbiology 2003, 149, 557–567. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Cray, J.A.; Bell, A.N.; Bhaganna, P.; Mswaka, A.Y.; Timson, D.J.; Hallsworth, J.E. The biology of habitat dominance; can microbes behave as weeds? Microb. Biotechnol. 2013, 6, 453–492. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  3. Hagman, A.; Säll, T.; Compagno, C.; Piskur, J. Yeast “make-accumulate-consume” life strategy evolved as a multi-step process that predates the whole genome duplication. PLoS ONE 2013, 8, e68734. [Google Scholar] [CrossRef] [PubMed]
  4. Williams, K.M.; Liu, P.; Fay, J.C. Evolution of ecological dominance of yeast species in high-sugar environments. Evolution 2015, 69, 2079–2093. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  5. Goffeau, A.; Barrell, B.G.; Bussey, H.; Davis, R.W.; Dujon, B.; Feldmann, H.; Galibert, F.; Hoheisel, J.D.; Jacq, C.; Johnston, M.; et al. Life with 6000 genes. Science 1996, 274, 546–567. [Google Scholar] [CrossRef] [Green Version]
  6. Engel, S.R.; Dietrich, F.S.; Fisk, D.G.; Binkley, G.; Balakrishnan, R.; Costanzo, M.C.; Dwight, S.S.; Hitz, B.C.; Karra, K.; Nash, R.S.; et al. The reference genome sequence of Saccharomyces cerevisiae: Then and now. G3 Genes Genomes Genet. 2014, 4, 389–398. [Google Scholar] [CrossRef] [Green Version]
  7. Cherry, J.M.; Hong, E.L.; Amundsen, C.; Balakrishnan, R.; Binkley, G.; Chan, E.T.; Christie, K.R.; Costanzo, M.C.; Dwight, S.S.; Engel, S.R.; et al. Saccharomyces Genome Database: The genomics resource of budding yeast. Nucleic Acids Res. 2012, 40, D700–D705. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  8. Kaeberlein, M.; Burtner, C.R.; Kennedy, B.K. Recent developments in yeast aging. PLoS Genet. 2007, 3, e84. [Google Scholar] [CrossRef] [PubMed]
  9. Barberis, M.; Todd, R.G.; van der Zee, L. Advances and challenges in logical modeling of cell cycle regulation: Perspective for multi-scale, integrative yeast cell models. FEMS Yeast Res. 2017, 17, fow103. [Google Scholar] [CrossRef] [PubMed]
  10. Petranovic, D.; Vemuri, G.N. Impact of yeast systems biology on industrial biotechnology. J. Biotechnol. 2009, 144, 204–211. [Google Scholar] [CrossRef]
  11. Bilsland, E.; Sparkes, A.; Williams, K.; Moss, H.J.; de Clare, M.; Pir, P.; Rowland, J.; Aubrey, W.; Pateman, R.; Young, M.; et al. Yeast-based automated high-throughput screens to identify anti-parasitic lead compounds. Open Biol. 2013, 3, 120158. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  12. Bilsland, E.; Pir, P.; Gutteridge, A.; Johns, A.; King, R.D.; Oliver, S.G. Functional expression of parasite drug targets and their human orthologs in yeast. PLoS Negl. Trop. Dis. 2011, 5, e1320. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  13. Ruetenik, A.; Barrientos, A. Exploiting post-mitotic yeast cultures to model neurodegeneration. Front. Mol. Neurosci. 2018, 11, 400. [Google Scholar] [CrossRef] [PubMed]
  14. Lindström, M.; Liu, B. Yeast as a model to unravel mechanisms behind FUS toxicity in amyotrophic lateral sclerosis. Front. Mol. Neurosci. 2018, 11, 218. [Google Scholar] [CrossRef] [PubMed]
  15. Nielsen, J. Yeast systems biology: Model organism and cell factory. Biotechnol. J. 2019, 14, 1800421. [Google Scholar] [CrossRef] [Green Version]
  16. Seynnaeve, D.; Vecchio, M.D.; Fruhmann, G.; Verelst, J.; Cools, M.; Beckers, J.; Mulvihill, D.P.; Winderickx, J.; Franssens, V. Recent insights on Alzheimer’s disease originating from yeast models. Int. J. Mol. Sci. 2018, 19, 1947. [Google Scholar] [CrossRef] [Green Version]
  17. Raghavan, V.; Aquadro, C.F.; Alani, E. Baker’s Yeast Clinical Isolates Provide a Model for How Pathogenic Yeasts Adapt to Stress. Trends Genet. 2019, 35, 804–817. [Google Scholar] [CrossRef]
  18. Daignan-Fornier, B.; Pinson, B. Yeast to study human purine metabolism diseases. Cells 2019, 8, 67. [Google Scholar] [CrossRef] [Green Version]
  19. Cazzanelli, G.; Pereira, F.; Alves, S.; Francisco, R.; Azevedo, L.; Dias Carvalho, P.; Almeida, A.; Côrte-Real, M.; Oliveira, M.J.; Lucas, C. The yeast Saccharomyces cerevisiae as a model for understanding RAS proteins and their role in human tumorigenesis. Cells 2018, 7, 14. [Google Scholar] [CrossRef] [Green Version]
  20. Di Gregorio, S.E.; Duennwald, M.L. ALS yeast models—Past success stories and new opportunities. Front. Mol. Neurosci. 2018, 11, 394. [Google Scholar] [CrossRef]
  21. Macedo, N.; Brigham, C.J. From beverages to biofuels: The journeys of ethanol-producing microorganisms. Int. J. Biotechnol. Wellness Ind. 2014, 3, 79–87. [Google Scholar]
  22. Naghshbandi, M.P.; Tabatabaei, M.; Aghbashlo, M.; Gupta, V.K.; Sulaiman, A.; Karimi, K.; Moghimi, H.; Maleki, M. Progress toward improving ethanol production through decreased glycerol generation in Saccharomyces cerevisiae by metabolic and genetic engineering approaches. Renew. Sustain. Energy Rev. 2019, 115, 109353. [Google Scholar] [CrossRef]
  23. Timilsina, G.R. Biofuels in the long-run global energy supply mix for transportation. Philos. Trans. R. Soc. A Math. Phys. Eng. Sci. 2014, 372, 20120323. [Google Scholar] [CrossRef] [PubMed]
  24. Gasparatos, A.; Stromberg, P.; Takeuchi, K. Sustainability impacts of first-generation biofuels. Anim. Front. 2013, 3, 12–26. [Google Scholar] [CrossRef]
  25. Lee, R.A.; Lavoie, J.-M. From first-to third-generation biofuels: Challenges of producing a commodity from a biomass of increasing complexity. Anim. Front. 2013, 3, 6–11. [Google Scholar] [CrossRef]
  26. Das Neves, M.A.; Kimura, T.; Shimizu, N.; Nakajima, M. State of the art and future trends of bioethanol production. Dynam. Biochem. Proc. Biotechnol. Mol. Biol. 2007, 1, 1–14. [Google Scholar]
  27. Paulino de Souza, J.; Dias do Prado, C.; Eleutherio, E.C.A.; Bonatto, D.; Malavazi, I.; Ferreira da Cunha, A. Improvement of Brazilian bioethanol production - Challenges and perspectives on the identification and genetic modification of new strains of Saccharomyces cerevisiae yeasts isolated during ethanol process. Fungal Biol. 2018, 122, 583–591. [Google Scholar] [CrossRef]
  28. Banerjee, A. Food, feed, fuel: Transforming the competition for grains. Dev. Chang. 2011, 42, 529–557. [Google Scholar] [CrossRef]
  29. Meyer, P.M.; Rodrigues, P.H.; Millen, D.D. Impact of biofuel production in Brazil on the economy, agriculture, and the environment. Anim. Front. 2013, 3, 28–37. [Google Scholar] [CrossRef] [Green Version]
  30. Ewing, M.; Msangi, S. Biofuels production in developing countries: Assessing tradeoffs in welfare and food security. Environ. Sci. Policy 2009, 12, 520–528. [Google Scholar] [CrossRef]
  31. Eisentraut, A. Sustainable Production of Second-Generation Biofuels; OECD/IEA: Paris, France, 2010.
  32. Cray, J.A.; Stevenson, A.; Ball, P.; Bankar, S.B.; Eleutherio, E.C.; Ezeji, T.C.; Singhal, R.S.; Thevelein, J.M.; Timson, D.J.; Hallsworth, J.E. Chaotropicity: A key factor in product tolerance of biofuel-producing microorganisms. Curr. Opin. Biotechnol. 2015, 33, 228–259. [Google Scholar] [CrossRef] [PubMed]
  33. Mohd Azhar, S.H.; Abdulla, R.; Jambo, S.A.; Marbawi, H.; Gansau, J.A.; Mohd Faik, A.A.; Rodrigues, K.F. Yeasts in sustainable bioethanol production: A review. Biochem. Biophys. Rep. 2017, 10, 52–61. [Google Scholar] [CrossRef] [PubMed]
  34. Jönsson, L.J.; Martín, C. Pretreatment of lignocellulose: Formation of inhibitory by-products and strategies for minimizing their effects. Bioresour. Technol. 2016, 199, 103–112. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Beckner, M.; Ivey, M.L.; Phister, T.G. Microbial contamination of fuel ethanol fermentations. Lett. Appl. Microbiol. 2011, 53, 387–394. [Google Scholar] [CrossRef] [PubMed]
  36. Narendranath, N.; Hynes, S.; Thomas, K.; Ingledew, W. Effects of lactobacilli on yeast-catalyzed ethanol fermentations. Appl. Environ. Microbiol. 1997, 63, 4158–4163. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  37. Gibson, B.R.; Lawrence, S.J.; Leclaire, J.P.; Powell, C.D.; Smart, K.A. Yeast responses to stresses associated with industrial brewery handling. FEMS Microbiol. Rev. 2007, 31, 535–569. [Google Scholar] [CrossRef] [Green Version]
  38. Maisonnave, P.; Sanchez, I.; Moine, V.; Dequin, S.; Galeote, V. Stuck fermentation: Development of a synthetic stuck wine and study of a restart procedure. Int. J. Food Microbiol. 2013, 163, 239–247. [Google Scholar] [CrossRef]
  39. Liu, Y.; Ishii, S.; Tokai, M.; Tsutsumi, H.; Ohki, O.; Akada, R.; Tanaka, K.; Tsuchiya, E.; Fukui, S.; Miyakawa, T. The Saccharomyces cerevisiae genes (CMP1 and CMP2) encoding calmodulin-binding proteins homologous to the catalytic subunit of mammalian protein phosphatase 2B. Mol. Gen. Genet. 1991, 227, 52–59. [Google Scholar] [CrossRef]
  40. Cyert, M.S.; Kunisawa, R.; Kaim, D.; Thorner, J. Yeast has homologs (CNA1 and CNA2 gene products) of mammalian calcineurin, a calmodulin-regulated phosphoprotein phosphatase. Proc. Natl. Acad. Sci. USA 1991, 88, 7376–7380. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  41. Cyert, M.S.; Thorner, J. Regulatory subunit (CNB1 gene product) of yeast Ca2+/calmodulin-dependent phosphoprotein phosphatases is required for adaptation to pheromone. Mol. Cell. Biol. 1992, 12, 3460–3469. [Google Scholar] [CrossRef] [Green Version]
  42. Matheos, D.P.; Kingsbury, T.J.; Ahsan, U.S.; Cunningham, K.W. Tcn1p/Crz1p, a calcineurin-dependent transcription factor that differentially regulates gene expression in Saccharomyces cerevisiae. Genes Dev. 1997, 11, 3445–3458. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  43. Schneiter, R.; Tatzer, V.; Gogg, G.; Leitner, E.; Kohlwein, S.D. Elo1p-dependent carboxy-terminal elongation of C14:1Delta(9) to C16:1Delta(11) fatty acids in Saccharomyces cerevisiae. J. Bacteriol. 2000, 182, 3655–3660. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Haro, R.; Garciadeblas, B.; Rodríguez-Navarro, A. A novel P-type ATPase from yeast involved in sodium transport. FEBS Lett. 1991, 291, 189–191. [Google Scholar] [CrossRef] [Green Version]
  45. Mori, K.; Kawahara, T.; Yoshida, H.; Yanagi, H.; Yura, T. Signalling from endoplasmic reticulum to nucleus: Transcription factor with a basic-leucine zipper motif is required for the unfolded protein-response pathway. Genes Cells 1996, 1, 803–817. [Google Scholar] [CrossRef] [PubMed]
  46. Brewster, J.L.; de Valoir, T.; Dwyer, N.D.; Winter, E.; Gustin, M.C. An osmosensing signal transduction pathway in yeast. Science 1993, 259, 1760–1763. [Google Scholar] [CrossRef]
  47. Wiederrecht, G.; Seto, D.; Parker, C.S. Isolation of the gene encoding the S. cerevisiae heat shock transcription factor. Cell 1988, 54, 841–853. [Google Scholar] [CrossRef]
  48. Nikawa, J.; Yamashita, S. IRE1 encodes a putative protein kinase containing a membrane-spanning domain and is required for inositol phototrophy in Saccharomyces cerevisiae. Mol. Microbiol. 1992, 6, 1441–1446. [Google Scholar] [CrossRef]
  49. Tokunaga, M.; Kawamura, A.; Kohno, K. Purification and characterization of BiP/Kar2 protein from Saccharomyces cerevisiae. J. Biol. Chem. 1992, 267, 17553–17559. [Google Scholar]
  50. Martinez-Pastor, M.T.; Marchler, G.; Schuller, C.; Marchler-Bauer, A.; Ruis, H.; Estruch, F. The Saccharomyces cerevisiae zinc finger proteins Msn2p and Msn4p are required for transcriptional induction through the stress response element (STRE). EMBO J. 1996, 15, 2227–2235. [Google Scholar] [CrossRef]
  51. Stukey, J.E.; McDonough, V.M.; Martin, C.E. The OLE1 gene of Saccharomyces cerevisiae encodes the delta 9 fatty acid desaturase and can be functionally replaced by the rat stearoyl-CoA desaturase gene. J. Biol. Chem. 1990, 265, 20144–20149. [Google Scholar]
  52. Serrano, R.; Kielland-Brandt, M.C.; Fink, G.R. Yeast plasma membrane ATPase is essential for growth and has homology with (Na+ + K+), K+- and Ca2+-ATPases. Nature 1986, 319, 689–693. [Google Scholar] [CrossRef] [PubMed]
  53. Serrano, R. Characterization of the plasma membrane ATPase of Saccharomyces cerevisiae. Mol. Cell. Biochem. 1978, 22, 51–63. [Google Scholar] [CrossRef] [PubMed]
  54. Li, W.; Brandriss, M.C. Proline biosynthesis in Saccharomyces cerevisiae: Molecular analysis of the PRO1 gene, which encodes gamma-glutamyl kinase. J. Bacteriol. 1992, 174, 4148–4156. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Tomenchok, D.M.; Brandriss, M.C. Gene-enzyme relationships in the proline biosynthetic pathway of Saccharomyces cerevisiae. J. Bacteriol. 1987, 169, 5364–5372. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  56. Brandriss, M.C.; Falvey, D.A. Proline biosynthesis in Saccharomyces cerevisiae: Analysis of the PRO3 gene, which encodes delta 1-pyrroline-5-carboxylate reductase. J. Bacteriol. 1992, 174, 3782–3788. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  57. Watanabe, Y.; Takaesu, G.; Hagiwara, M.; Irie, K.; Matsumoto, K. Characterization of a serum response factor-like protein in Saccharomyces cerevisiae, Rlm1, which has transcriptional activity regulated by the Mpk1 (Slt2) mitogen-activated protein kinase pathway. Mol. Cell. Biol. 1997, 17, 2615–2623. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  58. Dodou, E.; Treisman, R. The Saccharomyces cerevisiae MADS-box transcription factor Rlm1 is a target for the Mpk1 mitogen-activated protein kinase pathway. Mol. Cell. Biol. 1997, 17, 1848–1859. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  59. Torres, L.; Martín, H.; García-Saez, M.I.; Arroyo, J.; Molina, M.; Sánchez, M.; Nombela, C. A protein kinase gene complements the lytic phenotype of Saccharomyces cerevisiae lyt2 mutants. Mol. Microbiol. 1991, 5, 2845–2854. [Google Scholar] [CrossRef]
  60. Bell, W.; Klaassen, P.; Ohnacker, M.; Boller, T.; Herweijer, M.; Schoppink, P.; Van der Zee, P.; Wiemken, A. Characterization of the 56-kDa subunit of yeast trehalose-6-phosphate synthase and cloning of its gene reveal its identity with the product of CIF1, a regulator of carbon catabolite inactivation. Eur. J. Biochem. 1992, 209, 951–959. [Google Scholar] [CrossRef]
  61. De Virgilio, C.; Bürckert, N.; Bell, W.; Jenö, P.; Boller, T.; Wiemken, A. Disruption of TPS2, the gene encoding the 100-kDa subunit of the trehalose-6-phosphate synthase/phosphatase complex in Saccharomyces cerevisiae, causes accumulation of trehalose-6-phosphate and loss of trehalose-6-phosphate phosphatase activity. Eur. J. Biochem. 1993, 212, 315–323. [Google Scholar] [CrossRef]
  62. Phizicky, E.M.; Schwartz, R.C.; Abelson, J. Saccharomyces cerevisiae tRNA ligase. Purification of the protein and isolation of the structural gene. J. Biol. Chem. 1986, 261, 2978–2986. [Google Scholar] [PubMed]
  63. Xu, Q.; Teplow, D.; Lee, T.D.; Abelson, J. Domain structure in yeast tRNA ligase. Biochemistry 1990, 29, 6132–6138. [Google Scholar] [CrossRef] [PubMed]
  64. Causton, H.C.; Ren, B.; Koh, S.S.; Harbison, C.T.; Kanin, E.; Jennings, E.G.; Lee, T.I.; True, H.L.; Lander, E.S.; Young, R.A. Remodeling of yeast genome expression in response to environmental changes. Mol. Biol. Cell. 2001, 12, 323–337. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Piper, P.W. The heat shock and ethanol stress responses of yeast exhibit extensive similarity and functional overlap. FEMS Microbiol. Lett. 1995, 134, 121–127. [Google Scholar] [CrossRef] [PubMed]
  66. Schüller, C.; Brewster, J.; Alexander, M.; Gustin, M.; Ruis, H. The HOG pathway controls osmotic regulation of transcription via the stress response element (STRE) of the Saccharomyces cerevisiae CTT1 gene. EMBO J. 1994, 13, 4382–4389. [Google Scholar] [CrossRef] [PubMed]
  67. Moskvina, E.; Imre, E.M.; Ruis, H. Stress factors acting at the level of the plasma membrane induce transcription via the stress response element (STRE) of the yeast Saccharomyces cerevisiae. Mol. Microbiol. 1999, 32, 1263–1272. [Google Scholar] [CrossRef]
  68. Moskvina, E.; Schüller, C.; Maurer, C.; Mager, W.; Ruis, H. A search in the genome of Saccharomyces cerevisiae for genes regulated via stress response elements. Yeast 1998, 14, 1041–1050. [Google Scholar] [CrossRef]
  69. Ding, J.; Huang, X.; Zhang, L.; Zhao, N.; Yang, D.; Zhang, K. Tolerance and stress response to ethanol in the yeast Saccharomyces cerevisiae. Appl. Microbiol. Biotechnol. 2009, 85, 253. [Google Scholar] [CrossRef]
  70. Crawford, R.A.; Pavitt, G.D. Translational regulation in response to stress in Saccharomyces cerevisiae. Yeast 2019, 36, 5–21. [Google Scholar] [CrossRef] [Green Version]
  71. Alexandre, H.; Ansanay-Galeote, V.; Dequin, S.; Blondin, B. Global gene expression during short-term ethanol stress in Saccharomyces cerevisiae. FEBS Lett. 2001, 498, 98–103. [Google Scholar] [CrossRef] [Green Version]
  72. Hallsworth, J.E. Stress-free microbes lack vitality. Fungal Biol. 2018, 122, 379–385. [Google Scholar] [CrossRef] [PubMed]
  73. Aertsen, A.; Michiels, C.W. Diversify or die: Generation of diversity in response to stress. Crit. Rev. Microbiol. 2005, 31, 69–78. [Google Scholar] [CrossRef] [PubMed]
  74. Estruch, F. Stress-controlled transcription factors, stress-induced genes and stress tolerance in budding yeast. FEMS Microbiol. Rev. 2000, 24, 469–486. [Google Scholar] [CrossRef] [PubMed]
  75. Rangel, D.E.N.; Finlay, R.D.; Hallsworth, J.E.; Dadachova, E.; Gadd, G.M. Fungal strategies for dealing with environment- and agriculture-induced stresses. Fungal Biol. 2018, 122, 602–612. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Stanley, D.; Bandara, A.; Fraser, S.; Chambers, P.; Stanley, G.A. The ethanol stress response and ethanol tolerance of Saccharomyces cerevisiae. J. Appl. Microbiol. 2010, 109, 13–24. [Google Scholar] [CrossRef] [PubMed]
  77. Endo, A.; Nakamura, T.; Ando, A.; Tokuyasu, K.; Shima, J. Genome-wide screening of the genes required for tolerance to vanillin, which is a potential inhibitor of bioethanol fermentation, in Saccharomyces cerevisiae. Biotechnol. Biofuels 2008, 1, 3. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  78. Swinnen, S.; Schaerlaekens, K.; Pais, T.; Claesen, J.; Hubmann, G.; Yang, Y.; Demeke, M.; Foulquié-Moreno, M.R.; Goovaerts, A.; Souvereyns, K.; et al. Identification of novel causative genes determining the complex trait of high ethanol tolerance in yeast using pooled-segregant whole-genome sequence analysis. Genome Res. 2012, 22, 975–984. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  79. Hu, X.; Wang, M.; Tan, T.; Li, J.; Yang, H.; Leach, L.; Zhang, R.; Luo, Z. Genetic dissection of ethanol tolerance in the budding yeast Saccharomyces cerevisiae. Genetics 2007, 175, 1479–1487. [Google Scholar] [CrossRef] [Green Version]
  80. Haas, R.; Horev, G.; Lipkin, E.; Kesten, I.; Portnoy, M.; Buhnik-Rosenblau, K.; Soller, M.; Kashi, Y. Mapping Ethanol Tolerance in Budding Yeast Reveals High Genetic Variation in a Wild Isolate. Front. Genet. 2019, 10, 998. [Google Scholar] [CrossRef]
  81. Liu, R.; Liang, L.; Choudhury, A.; Garst, A.D.; Eckert, C.A.; Oh, E.J.; Winkler, J.; Gill, R.T. Multiplex navigation of global regulatory networks (MINR) in yeast for improved ethanol tolerance and production. Metab. Eng. 2019, 51, 50–58. [Google Scholar] [CrossRef]
  82. Kasavi, C.; Eraslan, S.; Arga, K.Y.; Oner, E.T.; Kirdar, B. A system based network approach to ethanol tolerance in Saccharomyces cerevisiae. BMC Syst. Biol. 2014, 8, 90. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  83. Ismail, A.; Ali, A. Selection of high ethanol-yielding Saccharomyces. Folia Microbiol. 1971, 16, 350–354. [Google Scholar] [CrossRef] [PubMed]
  84. Brown, S.; Oliver, S. Isolation of ethanol-tolerant mutants of yeast by continuous selection. Eur. J. Appl. Microbiol. Biotechnol. 1982, 16, 119–122. [Google Scholar] [CrossRef]
  85. Jiménez, J.; Benítez, T. Selection of ethanol-tolerant yeast hybrids in pH-regulated continuous culture. Appl. Environ. Microbiol. 1988, 54, 917–922. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. Madhavan, A.; Sindhu, R.; Arun, K.; Pandey, A.; Binod, P.; Gnansounou, E. Advances in Biofuel Production by Strain Development in Yeast from Lignocellulosic Biomass. Bioprocess. Biomol. Prod. 2019, 289–302. [Google Scholar]
  87. Miyagawa, K.-I.; Ishiwata-Kimata, Y.; Kohno, K.; Kimata, Y. Ethanol stress impairs protein folding in the endoplasmic reticulum and activates Ire1 in Saccharomyces cerevisiae. Biosci. Biotechnol. Biochem. 2014, 78, 1389–1391. [Google Scholar] [CrossRef] [PubMed]
  88. Navarro-Tapia, E.; Nana, R.K.; Querol, A.; Pérez-Torrado, R. Ethanol cellular defense induce unfolded protein response in yeast. Front. Microbiol. 2016, 7, 189. [Google Scholar] [CrossRef]
  89. Alexandre, H.; Rousseaux, I.; Charpentier, C. Relationship between ethanol tolerance, lipid composition and plasma membrane fluidity in Saccharomyces cerevisiae and Kloeckera apiculata. FEMS Microbiol. Lett. 1994, 124, 17–22. [Google Scholar] [CrossRef]
  90. Vanegas, J.M.; Contreras, M.F.; Faller, R.; Longo, M.L. Role of unsaturated lipid and ergosterol in ethanol tolerance of model yeast biomembranes. Biophys. J. 2012, 102, 507–516. [Google Scholar] [CrossRef] [Green Version]
  91. Shi, D.-J.; Wang, C.-L.; Wang, K.-M. Genome shuffling to improve thermotolerance, ethanol tolerance and ethanol productivity of Saccharomyces cerevisiae. J. Ind. Microbiol. Biotechnol. 2009, 36, 139–147. [Google Scholar] [CrossRef]
  92. Alper, H.; Moxley, J.; Nevoigt, E.; Fink, G.R.; Stephanopoulos, G. Engineering yeast transcription machinery for improved ethanol tolerance and production. Science 2006, 314, 1565–1568. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  93. Snoek, T.; Nicolino, M.P.; Van den Bremt, S.; Mertens, S.; Saels, V.; Verplaetse, A.; Steensels, J.; Verstrepen, K.J. Large-scale robot-assisted genome shuffling yields industrial Saccharomyces cerevisiae yeasts with increased ethanol tolerance. Biotechnol. Biofuels 2015, 8, 32. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Xue, T.; Chen, D.; Su, Q.; Yuan, X.; Liu, K.; Huang, L.; Fang, J.; Chen, J.; He, W.; Chen, Y. Improved ethanol tolerance and production of Saccharomyces cerevisiae by global transcription machinery engineering via directed evolution of the SPT8 gene. Food Biotechnol. 2019, 33, 155–173. [Google Scholar] [CrossRef]
  95. Jetti, K.D.; GNS, R.R.; Garlapati, D.; Nammi, S.K. Improved ethanol productivity and ethanol tolerance through genome shuffling of Saccharomyces cerevisiae and Pichia stipitis. Int. Microbiol. 2019, 22, 247–254. [Google Scholar] [CrossRef] [PubMed]
  96. Mitsui, R.; Yamada, R.; Ogino, H. Improved stress tolerance of Saccharomyces cerevisiae by CRISPR-Cas-mediated genome evolution. Appl. Biochem. Biotechnol. 2019, 189, 810–821. [Google Scholar] [CrossRef] [PubMed]
  97. Liu, K.; Yuan, X.; Liang, L.; Fang, J.; Chen, Y.; He, W.; Xue, T. Using CRISPR/Cas9 for multiplex genome engineering to optimize the ethanol metabolic pathway in Saccharomyces cerevisiae. Biochem. Eng. J. 2019, 145, 120–126. [Google Scholar] [CrossRef]
  98. Brown, A.D. Compatible solutes and extreme water stress in eukaryotic micro-organisms. Adv. Microb. Physiol. 1978, 17, 181–242. [Google Scholar]
  99. Hallsworth, J.E.; Prior, B.A.; Nomura, Y.; Iwahara, M.; Timmis, K.N. Compatible solutes protect against chaotrope (ethanol)-induced, nonosmotic water stress. Appl. Environ. Microbiol. 2003, 69, 7032–7034. [Google Scholar] [CrossRef] [Green Version]
  100. Cray, J.A.; Russell, J.T.; Timson, D.J.; Singhal, R.S.; Hallsworth, J.E. A universal measure of chaotropicity and kosmotropicity. Environ. Microbiol. 2013, 15, 287–296. [Google Scholar] [CrossRef]
  101. Timson, D.J. The roles and applications of chaotropes and kosmotropes in industrial fermentation processes. World J. Microbiol. Biotechnol. 2020, 36, 89. [Google Scholar] [CrossRef]
  102. McCammick, E.M.; Gomase, V.S.; McGenity, T.J.; Timson, D.J.; Hallsworth, J.E. Water-Hydrophobic Compound Interactions with the Microbial Cell. In Handbook of Hydrocarbon and Lipid Microbiology; Timmis, K.N., Ed.; Springer: Berlin/Heidelberg, Germany, 2010; pp. 1451–1466. [Google Scholar] [CrossRef]
  103. Singer, M.A.; Lindquist, S. Multiple Effects of Trehalose on Protein Folding In Vitro and In Vivo. Mol. Cell 1998, 1, 639–648. [Google Scholar] [CrossRef]
  104. Lever, M.; Blunt, J.; Maclagan, R. Some ways of looking at compensatory kosmotropes and different water environments. Comp. Biochem. Physiol. Part A Mol. Integr. Physiol. 2001, 130, 471–486. [Google Scholar] [CrossRef]
  105. Moelbert, S.; Normand, B.; De Los Rios, P. Kosmotropes and chaotropes: Modelling preferential exclusion, binding and aggregate stability. Biophys. Chem. 2004, 112, 45–57. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  106. Hohmann, S. Osmotic stress signaling and osmoadaptation in yeasts. Microbiol. Mol. Biol. Rev. 2002, 66, 300–372. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  107. Nevoigt, E.; Stahl, U. Osmoregulation and glycerol metabolism in the yeast Saccharomyces cerevisiae. FEMS Microbiol. Rev. 1997, 21, 231–241. [Google Scholar] [CrossRef] [PubMed]
  108. Mager, W.H.; de Boer, A.H.; Siderius, M.H.; Voss, H.-P. Cellular responses to oxidative and osmotic stress. Cell Stress Chaperones 2000, 5, 73. [Google Scholar] [CrossRef] [Green Version]
  109. Takagi, H. Proline as a stress protectant in yeast: Physiological functions, metabolic regulations, and biotechnological applications. Appl. Microbiol. Biotechnol. 2008, 81, 211–223. [Google Scholar] [CrossRef]
  110. Francois, J.; Parrou, J.L. Reserve carbohydrates metabolism in the yeast Saccharomyces cerevisiae. FEMS Microbiol. Rev. 2001, 25, 125–145. [Google Scholar] [CrossRef] [Green Version]
  111. Winderickx, J.; de Winde, J.H.; Crauwels, M.; Hino, A.; Hohmann, S.; Van Dijck, P.; Thevelein, J.M. Regulation of genes encoding subunits of the trehalose synthase complex in Saccharomyces cerevisiae: Novel variations of STRE-mediated transcription control? Mol. Gen. Genet. 1996, 252, 470–482. [Google Scholar] [CrossRef]
  112. Gorner, W.; Durchschlag, E.; Martinez-Pastor, M.T.; Estruch, F.; Ammerer, G.; Hamilton, B.; Ruis, H.; Schuller, C. Nuclear localization of the C2H2 zinc finger protein Msn2p is regulated by stress and protein kinase A activity. Genes. Dev. 1998, 12, 586–597. [Google Scholar] [CrossRef]
  113. Conlin, L.K.; Nelson, H.C.M. The Natural Osmolyte Trehalose Is a Positive Regulator of the Heat-Induced Activity of Yeast Heat Shock Transcription Factor. Mol. Cell. Biol. 2007, 27, 1505–1515. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  114. Wu, H.; Ng, B.S.; Thibault, G. Endoplasmic reticulum stress response in yeast and humans. Biosci. Rep. 2014, 34. [Google Scholar] [CrossRef] [PubMed]
  115. Levin, D.E. Cell wall integrity signaling in Saccharomyces cerevisiae. Microbiol. Mol. Biol. Rev. 2005, 69, 262–291. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  116. Gonzalez, T.N.; Sidrauski, C.; Dörfler, S.; Walter, P. Mechanism of non-spliceosomal mRNA splicing in the unfolded protein response pathway. EMBO J. 1999, 18, 3119–3132. [Google Scholar] [CrossRef] [PubMed]
  117. Cox, J.S.; Shamu, C.E.; Walter, P. Transcriptional induction of genes encoding endoplasmic reticulum resident proteins requires a transmembrane protein kinase. Cell 1993, 73, 1197–1206. [Google Scholar] [CrossRef]
  118. Kimata, Y.; Kimata, Y.I.; Shimizu, Y.; Abe, H.; Farcasanu, I.C.; Takeuchi, M.; Rose, M.D.; Kohno, K. Genetic evidence for a role of BiP/Kar2 that regulates Ire1 in response to accumulation of unfolded proteins. Mol. Biol. Cell 2003, 14, 2559–2569. [Google Scholar] [CrossRef] [Green Version]
  119. Okamura, K.; Kimata, Y.; Higashio, H.; Tsuru, A.; Kohno, K. Dissociation of Kar2p/BiP from an ER sensory molecule, Ire1p, triggers the unfolded protein response in yeast. Biochem. Biophys. Res. Commun. 2000, 279, 445–450. [Google Scholar] [CrossRef]
  120. Sidrauski, C.; Walter, P. The transmembrane kinase Ire1p is a site-specific endonuclease that initiates mRNA splicing in the unfolded protein response. Cell 1997, 90, 1031–1039. [Google Scholar] [CrossRef] [Green Version]
  121. Gardner, B.M.; Walter, P. Unfolded proteins are Ire1-activating ligands that directly induce the unfolded protein response. Science 2011, 333, 1891–1894. [Google Scholar] [CrossRef] [Green Version]
  122. Ho, N.; Yap, W.S.; Xu, J.; Wu, H.; Koh, J.H.; Goh, W.W.B.; George, B.; Chong, S.C.; Taubert, S.; Thibault, G. Stress sensor Ire1 deploys a divergent transcriptional program in response to lipid bilayer stress. J. Cell Biol. 2020, 219. [Google Scholar] [CrossRef]
  123. Sidrauski, C.; Cox, J.S.; Walter, P. tRNA ligase is required for regulated mRNA splicing in the unfolded protein response. Cell 1996, 87, 405–413. [Google Scholar] [CrossRef] [Green Version]
  124. Mori, K.; Ogawa, N.; Kawahara, T.; Yanagi, H.; Yura, T. Palindrome with spacer of one nucleotide is characteristic of the cis-acting unfolded protein response element in Saccharomyces cerevisiae. J. Biol. Chem. 1998, 273, 9912–9920. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  125. Welihinda, A.A.; Tirasophon, W.; Green, S.R.; Kaufman, R.J. Gene induction in response to unfolded protein in the endoplasmic reticulum is mediated through Ire1p kinase interaction with a transcriptional coactivator complex containing Ada5p. Proc. Natl. Acad. Sci. USA 1997, 94, 4289–4294. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  126. Moir, R.D.; Gross, D.A.; Silver, D.L.; Willis, I.M. SCS3 and YFT2 link transcription of phospholipid biosynthetic genes to ER stress and the UPR. PLoS Genet. 2012, 8, e1002890. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  127. Travers, K.J.; Patil, C.K.; Wodicka, L.; Lockhart, D.J.; Weissman, J.S.; Walter, P. Functional and genomic analyses reveal an essential coordination between the unfolded protein response and ER-associated degradation. Cell 2000, 101, 249–258. [Google Scholar] [CrossRef] [Green Version]
  128. Thibault, G.; Ismail, N.; Ng, D.T. The unfolded protein response supports cellular robustness as a broad-spectrum compensatory pathway. Proc. Natl. Acad. Sci. USA 2011, 108, 20597–20602. [Google Scholar] [CrossRef] [Green Version]
  129. Fun, X.H.; Thibault, G. Lipid bilayer stress and proteotoxic stress-induced unfolded protein response deploy divergent transcriptional and non-transcriptional programmes. Biochim. Biophys. Acta (BBA)-Mol. Cell Biol. Lipids 2020, 1865. [Google Scholar] [CrossRef]
  130. Krysan, D.J. The cell wall and endoplasmic reticulum stress responses are coordinately regulated in Saccharomyces cerevisiae. Commun. Integr. Biol. 2009, 2, 233–235. [Google Scholar] [CrossRef]
  131. Chowdhury, S.; Smith, K.W.; Gustin, M.C. Osmotic stress and the yeast cytoskeleton: Phenotype-specific suppression of an actin mutation. J. Cell Biol. 1992, 118, 561–571. [Google Scholar] [CrossRef]
  132. Mas, G.; De Nadal, E.; Dechant, R.; De La Concepción, M.L.R.; Logie, C.; Jimeno-González, S.; Chávez, S.; Ammerer, G.; Posas, F. Recruitment of a chromatin remodelling complex by the Hog1 MAP kinase to stress genes. EMBO J. 2009, 28, 326–336. [Google Scholar] [CrossRef]
  133. Silva, R.D.; Sotoca, R.; Johansson, B.; Ludovico, P.; Sansonetty, F.; Silva, M.T.; Peinado, J.M.; Côrte-Real, M. Hyperosmotic stress induces metacaspase-and mitochondria-dependent apoptosis in Saccharomyces cerevisiae. Mol. Microbiol. 2005, 58, 824–834. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  134. Rodríguez-Peña, J.M.; García, R.; Nombela, C.; Arroyo, J. The high-osmolarity glycerol (HOG) and cell wall integrity (CWI) signalling pathways interplay: A yeast dialogue between MAPK routes. Yeast 2010, 27, 495–502. [Google Scholar] [CrossRef] [PubMed]
  135. Jendretzki, A.; Wittland, J.; Wilk, S.; Straede, A.; Heinisch, J.J. How do I begin? Sensing extracellular stress to maintain yeast cell wall integrity. Eur. J. Cell Biol. 2011, 90, 740–744. [Google Scholar] [CrossRef] [PubMed]
  136. Sanz, A.B.; García, R.; Rodríguez-Peña, J.M.; Arroyo, J. The CWI pathway: Regulation of the transcriptional adaptive response to cell wall stress in yeast. J. Fungi 2018, 4, 1. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  137. Levin, D.E. Regulation of cell wall biogenesis in Saccharomyces cerevisiae: The cell wall integrity signaling pathway. Genetics 2011, 189, 1145–1175. [Google Scholar] [CrossRef] [Green Version]
  138. Baetz, K.; Moffat, J.; Haynes, J.; Chang, M.; Andrews, B. Transcriptional coregulation by the cell integrity mitogen-activated protein kinase Slt2 and the cell cycle regulator Swi4. Mol. Cell. Biol. 2001, 21, 6515–6528. [Google Scholar] [CrossRef] [Green Version]
  139. Kim, K.-Y.; Truman, A.W.; Levin, D.E. Yeast Mpk1 mitogen-activated protein kinase activates transcription through Swi4/Swi6 by a noncatalytic mechanism that requires upstream signal. Mol. Cell. Biol. 2008, 28, 2579–2589. [Google Scholar] [CrossRef] [Green Version]
  140. Kim, K.-Y.; Truman, A.W.; Caesar, S.; Schlenstedt, G.; Levin, D.E. Yeast Mpk1 cell wall integrity mitogen-activated protein kinase regulates nucleocytoplasmic shuttling of the Swi6 transcriptional regulator. Mol. Biol. Cell 2010, 21, 1609–1619. [Google Scholar] [CrossRef] [Green Version]
  141. Jung, U.S.; Levin, D.E. Genome-wide analysis of gene expression regulated by the yeast cell wall integrity signalling pathway. Mol. Microbiol. 1999, 34, 1049–1057. [Google Scholar] [CrossRef]
  142. Lagorce, A.; Hauser, N.C.; Labourdette, D.; Rodriguez, C.; Martin-Yken, H.; Arroyo, J.; Hoheisel, J.D.; François, J. Genome-wide analysis of the response to cell wall mutations in the yeast Saccharomyces cerevisiae. J. Biol. Chem. 2003, 278, 20345–20357. [Google Scholar] [CrossRef] [Green Version]
  143. O’Rourke, S.M.; Herskowitz, I.; O’Shea, E.K. Yeast go the whole HOG for the hyperosmotic response. Trends Genet. 2002, 18, 405–412. [Google Scholar] [CrossRef]
  144. Posas, F.; Chambers, J.R.; Heyman, J.A.; Hoeffler, J.P.; de Nadal, E.; Ariño, J.n. The transcriptional response of yeast to saline stress. J. Biol. Chem. 2000, 275, 17249–17255. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  145. Rep, M.; Krantz, M.; Thevelein, J.M.; Hohmann, S. The transcriptional response of Saccharomyces cerevisiae to osmotic shock Hot1p and Msn2p/Msn4p are required for the induction of subsets of high osmolarity glycerol pathway-dependent genes. J. Biol. Chem. 2000, 275, 8290–8300. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  146. Alexander, M.R.; Tyers, M.; Perret, M.; Craig, B.M.; Fang, K.S.; Gustin, M.C. Regulation of cell cycle progression by Swe1p and Hog1p following hypertonic stress. Mol. Biol. Cell 2001, 12, 53–62. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  147. Beese, S.E.; Negishi, T.; Levin, D.E. Identification of positive regulators of the yeast fps1 glycerol channel. PLoS Genet 2009, 5, e1000738. [Google Scholar] [CrossRef] [Green Version]
  148. Lee, J.; Reiter, W.; Dohnal, I.; Gregori, C.; Beese-Sims, S.; Kuchler, K.; Ammerer, G.; Levin, D.E. MAPK Hog1 closes the S. cerevisiae glycerol channel Fps1 by phosphorylating and displacing its positive regulators. Genes Dev. 2013, 27, 2590–2601. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  149. Auesukaree, C. Molecular mechanisms of the yeast adaptive response and tolerance to stresses encountered during ethanol fermentation. J. Biosci. Bioeng. 2017, 124, 133–142. [Google Scholar] [CrossRef]
  150. Fleet, G.M. Yeasts-growth during fermentation. In Wine Microbiology & Biotechnology; Harword Academic Publishers: Amsterdam, The Netherlands, 1993; pp. 27–54. [Google Scholar]
  151. Van Uden, N. Effect of alcohols on the temperature relations of growth and death in yeasts. In Alcohol Toxicity in Yeast and Bacteria; CRC Press: Boca Raton, FL, USA, 1989; pp. 77–88. [Google Scholar]
  152. Verghese, J.; Abrams, J.; Wang, Y.; Morano, K.A. Biology of the heat shock response and protein chaperones: Budding yeast (Saccharomyces cerevisiae) as a model system. Microbiol. Mol. Biol. Rev. 2012, 76, 115–158. [Google Scholar] [CrossRef] [Green Version]
  153. Swan, T.M.; Watson, K. Stress tolerance in a yeast lipid mutant: Membrane lipids influence tolerance to heat and ethanol independently of heat shock proteins and trehalose. Can. J. Microbiol. 1999, 45, 472–479. [Google Scholar] [CrossRef]
  154. Swan, T.M.; Watson, K. Stress tolerance in a yeast sterol auxotroph: Role of ergosterol, heat shock proteins and trehalose. FEMS Microbiol. Lett. 1998, 169, 191–197. [Google Scholar] [CrossRef]
  155. Endress, E.; Bayerl, S.; Prechtel, K.; Maier, C.; Merkel, R.; Bayerl, T.M. The effect of cholesterol, lanosterol, and ergosterol on lecithin bilayer mechanical properties at molecular and microscopic dimensions: A solid-state NMR and micropipet study. Langmuir 2002, 18, 3293–3299. [Google Scholar] [CrossRef] [Green Version]
  156. Hsueh, Y.-W.; Gilbert, K.; Trandum, C.; Zuckermann, M.; Thewalt, J. The Effect of Ergosterol on Dipalmitoylphosphatidylcholine Bilayers: A Deuterium NMR and Calorimetric Study. Biophys. J. 2005, 88, 1799–1808. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  157. Xu, X.; Bittman, R.; Duportail, G.; Heissler, D.; Vilcheze, C.; London, E. Effect of the structure of natural sterols and sphingolipids on the formation of ordered sphingolipid/sterol domains (rafts) Comparison of cholesterol to plant, fungal, and disease-associated sterols and comparison of sphingomyelin, cerebrosides, and ceramide. J. Biol. Chem. 2001, 276, 33540–33546. [Google Scholar] [PubMed] [Green Version]
  158. Caspeta, L.; Chen, Y.; Ghiaci, P.; Feizi, A.; Buskov, S.; Hallstrom, B.M.; Petranovic, D.; Nielsen, J. Biofuels. Altered sterol composition renders yeast thermotolerant. Science 2014, 346, 75–78. [Google Scholar] [CrossRef] [PubMed]
  159. Abdel-Fattah, W.; Fadil, M.; Nigam, P.; Banat, I. Isolation of thermotolerant ethanologenic yeasts and use of selected strains in industrial scale fermentation in an Egyptian distillery. Biotechnol. Bioeng. 2000, 68, 531–535. [Google Scholar] [CrossRef]
  160. Rajoka, M.; Ferhan, M.; Khalid, A. Kinetics and thermodynamics of ethanol production by a thermotolerant mutant of Saccharomyces cerevisiae in a microprocessor-controlled bioreactor. Lett. Appl. Microbiol. 2005, 40, 316–321. [Google Scholar] [CrossRef] [PubMed]
  161. Banat, I.; Marchant, R. Characterization and potential industrial applications of five novel, thermotolerant, fermentative, yeast strains. World J. Microbiol. Biotechnol. 1995, 11, 304–306. [Google Scholar] [CrossRef]
  162. Piper, P.W.; Talreja, K.; Panaretou, B.; Moradas-Ferreira, P.; Byrne, K.; Praekelt, U.M.; Meacock, P.; Récnacq, M.; Boucherie, H. Induction of major heat-shock proteins of Saccharomyces cerevisiae, including plasma membrane Hsp30, by ethanol levels above a critical threshold. Microbiology 1994, 140, 3031–3038. [Google Scholar] [CrossRef] [Green Version]
  163. Benjaphokee, S.; Hasegawa, D.; Yokota, D.; Asvarak, T.; Auesukaree, C.; Sugiyama, M.; Kaneko, Y.; Boonchird, C.; Harashima, S. Highly efficient bioethanol production by a Saccharomyces cerevisiae strain with multiple stress tolerance to high temperature, acid and ethanol. New Biotechnol. 2012, 29, 379–386. [Google Scholar] [CrossRef]
  164. Birch, R.M.; Walker, G.M. Influence of magnesium ions on heat shock and ethanol stress responses of Saccharomyces cerevisiae. Enzyme Microb. Technol. 2000, 26, 678–687. [Google Scholar] [CrossRef]
  165. Ball, P.; Hallsworth, J.E. Water structure and chaotropicity: Their uses, abuses and biological implications. Phys. Chem. Chem. Phys. 2015, 17, 8297–8305. [Google Scholar] [CrossRef] [PubMed]
  166. Abu-Hamdiyyah, M. The Effect of Urea on the Structure of Water and Hydrophobic Bonding1. J. Phys. Chem. 1965, 69, 2720–2725. [Google Scholar] [CrossRef]
  167. Sahle, C.J.; Schroer, M.A.; Juurinen, I.; Niskanen, J. Influence of TMAO and urea on the structure of water studied by inelastic X-ray scattering. Phys. Chem. Chem. Phys. 2016, 18, 16518–16526. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  168. Timson, D.J. Four Challenges for Better Biocatalysts. Fermentation 2019, 5, 39. [Google Scholar] [CrossRef] [Green Version]
  169. Geiler-Samerotte, K.A.; Dion, M.F.; Budnik, B.A.; Wang, S.M.; Hartl, D.L.; Drummond, D.A. Misfolded proteins impose a dosage-dependent fitness cost and trigger a cytosolic unfolded protein response in yeast. Proc. Natl. Acad. Sci. USA 2011, 108, 680–685. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  170. Telini, B.P.; Menoncin, M.; Bonatto, D. Does Inter-Organellar Proteostasis Impact Yeast Quality and Performance During Beer Fermentation? Front. Genet. 2020, 11, 2. [Google Scholar] [CrossRef]
  171. Tran, D.M.; Ishiwata-Kimata, Y.; Mai, T.C.; Kubo, M.; Kimata, Y. The unfolded protein response alongside the diauxic shift of yeast cells and its involvement in mitochondria enlargement. Sci. Rep. 2019, 9, 12780. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  172. Hallsworth, J.E. Ethanol-induced water stress in yeast. J. Ferment. Bioeng. 1998, 85, 125–137. [Google Scholar] [CrossRef]
  173. De Klerk, C.; Fosso-Kankeu, E.; Du Plessis, L.; Marx, S. Assessment of the viability of Saccharomyces cerevisiae in response to synergetic inhibition during bioethanol production. Curr. Sci. 2018, 115. [Google Scholar] [CrossRef]
  174. Fosso-Kankeu, E.; Marx, S.; Meyer, A. Simulated inhibitory effects of typical byproducts of biomass pretreatment process on the viability of Saccharomyces cerevisiae and bioethanol production yield. Afr. J. Biotechnol. 2015, 14, 2383–2394. [Google Scholar]
  175. Eardley, J.; Dedi, C.; Dymond, M.; Hallsworth, J.E.; Timson, D.J. Evidence for chaotropicity/kosmotropicity offset in a yeast growth model. Biotechnol. Lett. 2019, 41, 1309–1318. [Google Scholar] [CrossRef] [PubMed]
  176. Arakawa, T.; Timasheff, S.N. The stabilization of proteins by osmolytes. Biophys. J. 1985, 47, 411–414. [Google Scholar] [CrossRef]
  177. Gekko, K.; Timasheff, S.N. Thermodynamic and kinetic examination of protein stabilization by glycerol. Biochemistry 1981, 20, 4677–4686. [Google Scholar] [CrossRef] [PubMed]
  178. Gekko, K. Calorimetric Study on Thermal Denaturation of Lysozyme in Polyol-Water Mixtures. J. Biochem. 1982, 91, 1197–1204. [Google Scholar] [CrossRef] [PubMed]
  179. Timson, D.J.; Eardley, J. Destressing yeast for higher biofuel yields: Can Excess Chaotropicity Be Mitigated? Appl. Biochem. Biotechnol. 2020, 192, 1368–1375. [Google Scholar] [CrossRef] [PubMed]
  180. Jamieson, D.J. Oxidative stress responses of the yeast Saccharomyces cerevisiae. Yeast 1998, 14, 1511–1527. [Google Scholar] [CrossRef]
  181. Garre, E.; Raginel, F.; Palacios, A.; Julien, A.; Matallana, E. Oxidative stress responses and lipid peroxidation damage are induced during dehydration in the production of dry active wine yeasts. Int. J. Food Microbiol. 2010, 136, 295–303. [Google Scholar] [CrossRef] [Green Version]
  182. Ikner, A.; Shiozaki, K. Yeast signaling pathways in the oxidative stress response. Mutat. Res./Fundam. Mol. Mech. Mutagenesis 2005, 569, 13–27. [Google Scholar] [CrossRef]
  183. Moradas-Ferreira, P.; Costa, V.; Piper, P.; Mager, W. The molecular defences against reactive oxygen species in yeast. Mol. Microbiol. 1996, 19, 651–658. [Google Scholar] [CrossRef]
  184. Girard, P.; Boiteux, S. Repair of oxidized DNA bases in the yeast Saccharomyces cerevisiae. Biochimie 1997, 79, 559–566. [Google Scholar] [CrossRef]
  185. Pan, X.; Ye, P.; Yuan, D.S.; Wang, X.; Bader, J.S.; Boeke, J.D. A DNA integrity network in the yeast Saccharomyces cerevisiae. Cell 2006, 124, 1069–1081. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  186. Herrero, E.; Ros, J.; Bellí, G.; Cabiscol, E. Redox control and oxidative stress in yeast cells. Biochim. Biophys. Acta (BBA)-Gen. Subj. 2008, 1780, 1217–1235. [Google Scholar] [CrossRef] [PubMed]
  187. Ariño, J. Integrative responses to high pH stress in S. cerevisiae. Omics 2010, 14, 517–523. [Google Scholar] [CrossRef] [PubMed]
  188. Serra-Cardona, A.; Canadell, D.; Ariño, J. Coordinate responses to alkaline pH stress in budding yeast. Microb. Cell 2015, 2, 182–196. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  189. Viladevall, L.; Serrano, R.; Ruiz, A.; Domenech, G.; Giraldo, J.; Barcelo, A.; Arino, J. Characterization of the calcium-mediated response to alkaline stress in Saccharomyces cerevisiae. J. Biol. Chem. 2004, 279, 43614–43624. [Google Scholar] [CrossRef] [Green Version]
  190. Stathopoulos, A.M.; Cyert, M.S. Calcineurin acts through the CRZ1/TCN1-encoded transcription factor to regulate gene expression in yeast. Genes. Dev. 1997, 11, 3432–3444. [Google Scholar] [CrossRef] [Green Version]
  191. Casado, C.; González, A.; Platara, M.; Ruiz, A.; Ariño, J. The role of the protein kinase A pathway in the response to alkaline pH stress in yeast. Biochem. J. 2011, 438, 523–533. [Google Scholar] [CrossRef] [Green Version]
  192. Canadell, D.; García-Martínez, J.; Alepuz, P.; Pérez-Ortín, J.E.; Ariño, J. Impact of high pH stress on yeast gene expression: A comprehensive analysis of mRNA turnover during stress responses. Biochim. Biophys. Acta (BBA)-Gene Regul. Mech. 2015, 1849, 653–664. [Google Scholar] [CrossRef]
  193. Ariño, J.; Ramos, J.; Sychrova, H. Monovalent cation transporters at the plasma membrane in yeasts. Yeast 2019, 36, 177–193. [Google Scholar] [CrossRef]
  194. Platara, M.; Ruiz, A.; Serrano, R.; Palomino, A.; Moreno, F.; Ariño, J. The Transcriptional Response of the Yeast Na+-ATPase ENA1 Gene to Alkaline Stress Involves Three Main Signaling Pathways. J. Biol. Chem. 2006, 281, 36632–36642. [Google Scholar] [CrossRef] [Green Version]
  195. Benito, B.; Quintero, F.J.; Rodríguez-Navarro, A. Overexpression of the sodium ATPase of Saccharomyces cerevisiae: Conditions for phosphorylation from ATP and Pi. Biochim. Biophys. Acta (BBA)-Biomembr. 1997, 1328, 214–225. [Google Scholar] [CrossRef] [Green Version]
  196. Giannattasio, S.; Guaragnella, N.; Corte-Real, M.; Passarella, S.; Marra, E. Acid stress adaptation protects Saccharomyces cerevisiae from acetic acid-induced programmed cell death. Gene 2005, 354, 93–98. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  197. Tenreiro, S.; Nunes, P.c.A.; Viegas, C.A.; Neves, M.S.; Teixeira, M.C.; Cabral, M.G.; Sá-Correia, I. AQR1 gene (ORF YNL065w) encodes a plasma membrane transporter of the major facilitator superfamily that confers resistance to short-chain monocarboxylic acids and quinidine in Saccharomyces cerevisiae. Biochem. Biophys. Res. Commun. 2002, 292, 741–748. [Google Scholar] [CrossRef] [PubMed]
  198. Tenreiro, S.; Rosa, P.C.; Viegas, C.A.; Sá-Correia, I. Expression of the AZR1 gene (ORF YGR224w), encoding a plasma membrane transporter of the major facilitator superfamily, is required for adaptation to acetic acid and resistance to azoles in Saccharomyces cerevisiae. Yeast 2000, 16, 1469–1481. [Google Scholar] [CrossRef]
  199. Carmelo, V.; Santos, H.; Sá-Correia, I. Effect of extracellular acidification on the activity of plasma membrane ATPase and on the cytosolic and vacuolar pH of Saccharomyces cerevisiae. Biochim. Biophys. Acta (BBA)-Biomembr. 1997, 1325, 63–70. [Google Scholar] [CrossRef] [Green Version]
Figure 1. Structures of the three main compatible solutes deployed by yeast. Note that there is little structural similarity between these molecules, except that they all relatively small and hydrophilic.
Figure 1. Structures of the three main compatible solutes deployed by yeast. Note that there is little structural similarity between these molecules, except that they all relatively small and hydrophilic.
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Figure 2. Structures of sterols found in yeast membranes. Cholesterol, which is commonly found in mammalian cell membranes, is shown for comparison.
Figure 2. Structures of sterols found in yeast membranes. Cholesterol, which is commonly found in mammalian cell membranes, is shown for comparison.
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Table 1. The roles of proteins mentioned in this review.
Table 1. The roles of proteins mentioned in this review.
ProteinSystematic Gene NameFunctionRole in StressReferences
Cmp2pYML057WCalcineurin catalytic subunitInvolved in sensing alkaline stress [39,40]
Cna1pYLR433CCalcineurin catalytic subunitInvolved in sensing alkaline stress [39,40]
Cnb1pYKL190WCalcineurin regulatory subunitInvolved in sensing alkaline stress [41]
Crz1pYNL027WTranscription factor Involved in responding to alkaline stress [42]
Elo1pYJL196CMedium-chain fatty acyl elongaseEnables production of longer fatty acids in heat and chaotrope stress [43]
Ena1pYDR040CSodium ion pump Involved in responding to alkaline stress [44]
Hac1pYFL031WTranscription factor Activates genes involved in the unfolded protein response [45]
Hog1pYLR113WMAP kinaseTerminal kinase of the HOG pathway. Activates genes in response to osmotic and other stresses [46]
Hsf1pYGL073WTranscription factorActivates genes involved in the heat shock response [47]
Ire1pYHR079CProtein kinase and nucleaseCleaves HAC1 RNA making it translationally competent [48]
Kar2pYJL034WChaperoneDetects and responds to the presence of unfolded proteins in the endoplasmic reticulum [49]
Msn2pYMR037CTranscription factorActivates STRE responsive genes [50]
Msn4pYKL062WTranscription factorActivates STRE responsive genes [50]
Ole1pYGL055WΔ9 fatty acid desaturaseEnables production of saturated fatty acids in heat and chaotrope stress [51]
Pma1pYGL008CHydrogen ion pumpPumps protons into the vacuole and extracellular medium in acid stress [52,53]
Pro1pYDR300Cγ-glutamyl kinaseEnables synthesis of proline which protects against heat and chaotrope stress [54]
Pro2pYOR323Cγ-glutamyl phosphate reductaseEnables synthesis of proline which protects against heat and chaotrope stress [55]
Pro3pYER023WΔ1-pyrroline-5-carboxylate reductaseEnables synthesis of proline which protects against heat and chaotrope stress [55,56]
Rlm1pYPL089CTranscription factor Activated by Slt2p. Controls expression of genes involved in cell wall maintenance and strengthening [57,58]
Slt2pYHR030CMAP kinaseTerminal kinase of the CWI pathway. Activates genes in response to cell wall stress[59]
Tps1pYBR126CTrehalose-6-phosphate synthaseEnables synthesis of trehalose in heat and chaotrope stress [60]
Tps2pYDR074WTrehalose-6-phosphate phosphataseEnables synthesis of trehalose in heat and chaotrope stress [61]
Trl1pYJL087CRNA ligaseLigates HAC1 following cleavage by Ire1p[62,63]
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