ethanol and thermotolerance in hte bioconversion of … · starch and fermentation of the resulting...

Ethanol and Thermotolerance in the Bioconversionof Xylose by Yeasts

THOMAS W. JEFFRIES

Institute for Microbial and Biochemical TechnologyForest Service, Forest Products LaboratoryUnited States Department of Agriculture

Madison, Wisconsin 53705-2366and

Department of Bacteriology, University of Wisconsin, MadisonMadison, Wisconsin 53706-1527

YONG-SU JIN

Department of Food ScienceUniversity of Wisconsin, MadisonMadison, Wisconsin 53706-1527

I. IntroductionII. Lignocellulose

A. PretreatmentB. Simultaneous Saccharification and Fermentation

III. Xylose-Fermenting MicrobesA. BacteriaB. Xylose-Fermenting Yeasts and FungiC. Genetic Studies with P. stipitis and P. tannophilusD. Expression of Pichia Genes in Saccharomyces

IV. Critical Parameters for Yeast Xylose FermentationA. Carbon Source

B. TemperatureC. pHD. AerationE. Nutrient Uptake

V. Factors Affecting Thermo- and Ethanol ToleranceA. Membrane LipidsB. Plasma Membrane H+-ATPaseC. Mitochondrial StabilityD. TrehaloseE. Heat-Shock Proteins

VI. summaryReferences

221ADVANCES IN APPLIED MICROBIOLOGY, VOLUME 47

Copyright © 2000 by Academic Press

222 THOMAS W. JEFFRIES and YONG-SU JIN

I. Introduction

Most ethanol produced in the world today is derived from starch orsucrose (Gong et al., 1999). These storage carbohydrates are readilyhydrolyzed by enzymes, and Saccharomyces cerevisiae easily fermentsthe resulting sugars-glucose and fructose-to high concentrations ofethanol (e.g., van Hoek et al., 1998). Ethanol fermentations are tradi-tionally carried out for wine or beer production, but ethanol for trans-portation is a large and growing use. In 1997, about 13,000 tons ofethanol were produced for fuel worldwide (Wilke, 1999). Ethanol isclean burning. Its oxygen content decreases emissions when combustedwith gasoline, and because ethanol is derived ultimately from plantmatter, its use as a fuel does not contribute to the net accumulation ofcarbon dioxide in the atmosphere (Costello and Chum, 1998). Starchesand sugars are abundant in many crops, but expansion of ethanolproduction as an automotive fuel in the new millennium will requirefeedstocks that do not compete for food or fiber (Wheals et al., 1999).Such feedstocks include lignocellulosic byproduct residues from agri-culture and silviculture (Saddler, 1993). Their utilization will requirenew technologies for efficient, inexpensive bioprocessing.

Lignocellulose is a generic term for plant matter derived from woodand agricultural residues. It is composed mainly of lignin and cellulose,but the lignocellulosic phytomass also contains significant amounts ofhemicellulose. Xylose is the principal component of hemicellulosefound in angiosperm agricultural and silvicultural residues. It is ob-tained by acid or enzymatic hydrolysis of xylan. The cellulosic fibercomponent of wood and many agricultural residues is in demand forfiber production, but the hemicellulose and lignin components areavailable for recovery through biorefining and bioconversion.

Biorefining is analogous to petrochemical refining in which a crudefeedstock is separated into its higher-value components. While petro-chemical refining is carried out at high temperatures on a complexmixture of relatively low-molecular-weight monomers, biorefining iscarried out at moderate temperatures on mixtures of polymeric andmonomeric materials. Corn wet milling best represents biorefining asit is practiced today. In this large-scale industrial process, corn is sepa-rated into its starch, oil, gluten, fiber, and nitrogenous liquid compo-nents. Subsequent bioconversion involves enzymatic hydrolysis of thestarch and fermentation of the resulting glucose to ethanol, citrate,lactate, and various other products such as antibiotics. Biorefining oflignocellulose to useful products is more difficult, because wood andagricultural residues are composed of structural polymers rather than

THERMO- AND ETHANOL TOLERANCE 223

the storage polymers found in grains. Moreover, the various extractivesreleased and degradation products formed during pretreatment oftenmake lignocellulosic hydrolysates harder to use.

Lignocellulose contains five major sugars, the abundance of whichvaries with the feedstock (Pettersen, 1984). They are the hexoses D-glu-cose, D-mannose, and D-galactose, and the pentoses D-xylose andL-arabinose. Fructose is not normally found in lignocellulose. Commer-cial bioconversion of lignocellulose to ethanol requires efficient fer-mentation of sugar mixtures-including xylose (Hinman et al., 1989).Otherwise, product yields are low and waste disposal costs excessive.

The fermentation of glucose and fructose has been establishedthrough thousands of years of practice. The prevailing yeast strainsused for producing wine, beer, and bread have been isolated from manydifferent sources. They belong to S. cerevisiae and a few other taxo-nomic groups (Vaughn-Martini and Martini, 1995). In contrast, theobjective of producing ethanol from pentose sugars has arisen relativelyrecently, and, despite much effort in several laboratories around theworld, it remains problematic. Even though anhydrides of xylose areabundant, xylose itself does not usually occur as a free sugar. Moreover,the five-carbon structure of xylose does not lend itself readily to fer-mentations in which ethanol is the sole product. Therefore, little natu-ral selection for xylose-fermenting species has taken place. This reviewfocuses on achieving high ethanol concentrations at elevated tempera-tures-conditions that would be appropriate for enzymatic saccharifi-cation and cofermentation of lignocellulosic feedstocks.

I I. Lignocellulose

Lignocellulosic materials are complex matrices of lignin, cellulose,hemicellulose, various extractives, and inorganic components. Thecompositions vary widely with plant species, age, time of harvest, andcondition or stage of growth (Higuchi 1997). Analysis is challenging(Puls, 1993). About 45% of the total dry weight of wood is cellulose,the hydrolysis of which yields glucose. In agricultural residues, cellu-lose comprises 8 to 35% of the total dry weight. Glucose is also presentin hemicellulosic sugars. Overall, glucose averages 30% of the total dryweight in these materials (Pettersen and Schwandt, 1991; Pettersen,1984; Krull and Inglett, 1980). The prevalence of glucose in starch andother storage carbohydrates such as sucrose makes D-glucose the mostabundant carbohydrate in terrestrial plants. Xylose is the second mostabundant sugar. It is especially prevalent in angiosperms (floweringplants). In woody angiosperms (hardwoods), D-xylose averages about


17% of the total dry weight, but in herbaceous angiosperms, such asresidues from agricultural crops, it can comprise up to 31%. Othersugars- s u c h as mannose, galactose, and arabinose-are found in theglucomannan, arabinoxylan, and glucuronoxylan hemicellulosic com-ponents. Glucomannan is the main hemicellulose of gymnosperms(softwoods). In these plants, mannose comprises about 10% of the totaldry weight. The lignin content is slightly higher in softwoods than inhardwoods, and because the lignin is more crosslinked in softwoods itis harder to remove. Although the glucose and mannose present insoftwoods can be fermented readily, timber and pulp manufactureplaces a high value on straight trunks and long fibers. Therefore, agri-cultural residues and fast-growing hardwood species are most com-.monly considered for fuel ethanol production. The high content ofxylose in these materials requires that it be used efficiently.

A. PRETREATMENT

Lignocellulosic materials must be treated with physical, chemical, orthermal processes in order to release fermentable sugars or increasetheir susceptibility to enzymatic hydrolysis. Several pretreatments arepresently under investigation. They include lime (Chang et al., 1997),ammonia (Dale et al., 1999), high-temperature dilute acid (Lee et al.,1999), and concentrated acid (Goldstein et al., 1989). These and otherpretreatments have been reviewed more recently (Szczodrak andFiedurek, 1996).

Acid hydrolysis is one of the oldest and most established technolo-gies for converting lignocellulose into fermentable sugars. There are twoprincipal approaches: dilute sulfuric acid and concentrated-acid hy-drolysis. Dilute-acid hydrolysis is often carried out in two stages. In thefirst stage, a relatively mild hydrolysis is used to recover the hemicel-lulosic sugars. Depending on the substrate and the conditions used,between 80 and 95% of the hemicellulosic sugars can be recovered fromthe lignocellulosic feedstock (Torget and Hsu, 1994; Torget et al., 1996;Katzen and Fowler, 1994). In the second stage, a higher concentrationof acid and a higher temperature hydrolyze the cellulose to glucose.The first- and second-stage hydrolysates can be recovered and fer-mented separately by different organisms or combined and fermented.Other variations include a mild acid pretreatment stage combined withsubsequent enzymatic saccharification and fermentation (see §I.B).

One of the difficulties with dilute-acid hydrolysis is that it degradesthe lignin into a nonreactive form, and it generates large amountsof toxic byproducts that inhibit the growth of fermentative microbes


(Larsson et al., 1999a; Mandenius et al., 1999). A variation on dilute-acid hydrolysis uses continuous countercurrent high-temperature di-lute acid to reduce formation of degradation products in the secondstage (Lee et al., 1999). While dilute-acid hydrolysis itself is inexpen-sive and can be used with a wide variety of feedstocks, the detoxifica-tion steps add complexity and cost (Larsson et al., 1999b). The high-temperature countercurrent continuous approach results in a higherproduct yield, but generates a much more dilute sugar stream. Withbatch-wise dilute-acid hydrolysis, only about 50 to 55% of the cellulosein wood can be converted to sugar. The balance of the material is eitherleft as residual cellulose or is degraded. Therefore, while the technologyis inexpensive, it is not sufficiently effective for commercial develop-ment unless the feedstock is very cheap. Advanced forms of high-tem-perature countercurrent hydrolysis or dilute-acid pretreatment com-bined with enzymatic saccharification and cofermentation may proveto be cost-effective.

Concentrated-acid hydrolysis is carried out at lower temperaturesand generates fewer degradation byproducts, so fermentation of theresulting sugars is much less problematic. However, concentrated sul-furic or hydrochloric acid is difficult to work with, and essentially allof the acid must be recovered and reconcentrated in order for theprocess to be economical. Electrodialysis (Goldstein, 1989) and ion-ex-change chromatography have been investigated as technological ap-proaches to acid recovery, and are presently being pursued for commer-cialization.

B. SIMULTANEOUS SACCHARIFICATION AND FERMENTATION

Simultaneous saccharification and fermentation (SSF) is the most effi-cient way to convert pretreated lignocellulose to ethanol (McMillan etal., 1999; Banat et al., 1998; Wyman, 1994; Philippidis et al., 1993). Itis often effective when combined with dilute-acid or high-temperaturehot-water pretreatment (Sreenath et al., 1999). In SSF, cellulases andxylanases convert the carbohydrate polymers to fermentable sugars.These enzymes are notoriously susceptible to feedback inhibition bythe products-glucose, xylose, cellobiose, and other oligosaccharides.But the efficiency of enzymatic saccharification increases if the result-ing sugars are converted to ethanol. Because cellulases function well atrelatively high temperatures (50 to 70°C), the limiting factor is fermen-tation. The efficiency of product formation increases with increasingethanol concentration up to about 5% on a w/w basis, so fermentation


at high temperatures (>40°C) and at or above 5% ethanol are prioritiesfor commercialization of this technology.

A few yeasts will ferment glucose at temperatures up to 45°C. Thebest studied of these is Kluyveromyces marxianus IMB3 (Fleming et al.,1993; Singh et al., 1998). Strains of this yeast have been adapted bycontinuous culture in the presence of ethanol to ferment glucose to 44g/liter ethanol and at temperatures as high as 45°C (Hack and Marchant,1998). However, growth is slight and the trait is not stable. Researchershave sought thermotolerant yeasts to use in SSF processes for almost20 years (McCracken and Gong, 1983), but this effort has been largelylimited to empirical screening studies. Accumulated knowledge aboutthe basis for ethanol and thermotolerance will provide new opportuni-ties for developing better strains through molecular genetics.

III. Xylose-Fermenting Microbe’s

A. BACTERIA

Bacteria have been known to ferment pentoses since the studies oflactobacilli by Fred et al. (1920). Today, xylose-fermenting bacteriainclude both native and genetically engineered organisms, and manyhave characteristics useful for simultaneous saccharification and fer-mentation (Table I). Bacterial fermentations of xylose for ethanol pro-duction are being commercialized, but yeasts have ‘several perceivedadvantages. With a few exceptions, such as lactic acid fermentation(Anuradha et al., 1999), bacteria produce a wide mixture of metabolicproducts and exhibit much lower ethanol tolerance. This makes prod-uct recovery more difficult. Yeasts have larger cells and thicker cellwalls than bacteria, which makes cell harvest and recycle easier. Per-haps most importantly, yeast fermentations are not as susceptible tocontamination by bacteria or viruses. For these reasons many industrialethanol processors retain an interest in xylose-fermenting yeasts. As-pects of xylose fermentation by bacteria have been reviewed previously(Jeffries, 1983).

B. XYLOSE-FERMENTING YEASTS AND FUNGI

Karczewska (1959) first reported direct conversion of xylose to ethanolby yeast. Even though many yeasts were known to assimilate xylose,this discovery did not enter the review literature (e.g., Lodder, 1971),and was not cited for over 20 years. Various laboratories began toreexamine yeast xylose fermentation after Wang et al. (1980) reportedthat S. cerevisiae and Schizosaccharomyces pombe could ferment xy-lulose, a keto pentulose, to ethanol. Soon afterward, Schneider et al.


TABLE INATIVE AND E NGINEERED BACTERIAL SPECIES CAPABLE OF FERMENTING XYLOSE TO ETHANOL

Species Characteristics References

Clostridiumacetobuti-licum

Clostridiumthermocellum

Escherichiacoli

Klebsiellaoxytoca

Lactobacilluspentoaceticus

Lactobacilluscasei

Lactobacillusxylosus

Lactobacilluspentosus

Lactobacillusplantarum

Zymomonas Normally ferments glucose andmobilis fructose; engineered to ferment xylose.

Useful in fermentation of xyloseto acetone and butanol; ethanolproduced in low yield.

Capable of converting cellulosedirectly to ethanol and acetic acid:ethanol concentrations are generallyless than 5 g/liter.

Native strains ferment xylose to amixture of ethanol, succininc, andacetic acids but lack ethanoltolerance; genetically engineeredstrains predominantly produceethanol.

Native strains rapidly ferment xyloseand cellobiose; engineered to fermentcellulose and produce ethanolpredominantly.

Consumes xylose and arabinose.Slowly uses glucose and cellobiose.Acetic acid is produced along withlactic in 1:1 ratio.

Ferments lactose very well; particu-larly useful for bioconversion of whey.

Uses cellobiose if nutrients aresupplied: uses n-glucose, D-xylose,and L-arabinose.

Homolactic fermentation. Somestrains produce lactic acid fromsulfite waste liquors.

Consumes cellobiose more rapidlythan glucose, xylose, or arabinose.Appears to depolymerize pectins;produces lactic acid from agriculturalresidues.

El Kanouni et al. (1998)

Herrero and Gomez(1980)

Lindsay et al. (1995),Yamano et al. (1998)

Ingram et al. (1999)

Chaillou et al. (1998),Sreenath et al. (1999)

Chaillou et al. (1999),Roukas and Kotzekidou(1998)

Sreenath, personal com-munication (1999)

Chaillou et al. (1999)

Chaillou et al. (1999).Sreenath et al. (1999)

Zhang et al. (1995)


could ferment xylose to ethanol. Gong et al. (1981) reported a mutantCandida sp. that would produce ethanol from xylose, and about thatsame time, Jeffries (1981) reported that Candida tropicalis requiredaeration to convert xylose to ethanol. The early progress in this fieldhas been reviewed by several authors (e.g., Jeffries and Kurtzman, 1994;Hahn-Hägerdal et al., 1994). Other information on metabolic engineer-ing and regulation of yeast xylose fermentation has been reviewed(Jeffries and Shi, 1999). The current review focuses on attaining ele-vated levels of ethanol at temperatures that are normally consideredextreme for yeast growth.

(1981) and Slininger et al. (1982) reported that Pachysolen tannophilus

At least 22 yeast strains have been shown to produce some ethanolfrom D-xylose (Toivola et al., 1984; du Preez and van der Walt,1983; Schneider et al., 1981) (Table II). However, only six of these

TABLE IINATIVE OR ENGINEERED YEAST AND FUNGAL SPECIES CAPABLE

OF FERMENTING XYLOSE TO ETHANOL

Species Characteristics References

Candidashehatae

Has both active and passive transportsystems for xylose uptake; producesmoderate amounts of xylitol; does notgrow anaerobically: requires biotinand thiamine.

Candidaboidinii

Produces large amounts of xylitol;oxidizes methanol.

Pichiastipitis

Ferments all sugars found in wood;some strains ferment xylan.

Fusariumoxysporum;Fusariumoxysporumvar. lini

Ferments 20 different carbon sources,including xylitol; does not use xylanor cellulose; converts xylose toethanol, CO2, and acetic acid.

Mucor sp. Ferments pentoses and alditols toethanol.

Pachysolentannophilus

Ferments xylose, glucose andglycerol; metabolizes xyloseanaerobically; produces largeamounts of xylitol.

du Preez and van derWalt (1983)

Vandeska et al. (1996)

Lee et al. (1986)

White and Williams(1928), Gibbs et al.(1954), Suihko et al.(1991), Suihko (1983)

Ueng and Gong, 1982

Schneider et al. (1981),Slininger et al. (1982)


(Brettanomyces naardenensis, Candida shehatae, Candida tenuis,P. tannophilus, Pichia segobiensis, and Pichia stipitis) produce signifi-cant amounts of ethanol, and of these only three (C. shehatae, P. tanno-philus, and P. stipitis) have been studied extensively. Several researchgroups have developed genetic transformation systems for P. tannophi-lus (Wedlock and Thornton, 1989; Reiser et al., 1990; Hayman andBolen, 1993) and for P. stipitis (Ho et al., 1991; Morosoli et al., 1993;Yang et al., 1994; Lu et al., 1998a; Piontek et al., 1998). It is possible totransform auxotrophic strains of C. shehatae with selectable markersand vectors designed for P. stipitis, but the absence of a sexual matingsystem in C. shehatae makes it less useful for fundamental studies.

C. GENETIC STUDIES WITH P. STIPITIS AND P. TANNOPHILUS

Cloning and disruption of genes for the critical enzymes involved inxylose metabolism or their overexpression in S. cerevisiae have led togreatly improved understanding of the rate-limiting steps in yeast xy-lose metabolism. At the end of 1999, GenBank listed approximately 26entries for genes cloned from P. stipitis or P. tannophilus–in additionto ribosomal genes used for taxonomic classification studies (Kurtzman,1994).

Stevis and Ho (1987) did some of the earliest research on the genesfor xylose metabolism by creating a xylulokinase mutant of Escherichiacoli (Stevis et al., 1987), then using it to clone the xylulokinase genefrom P. tannophilus. Subsequently, Ho and Chang (1989) used as similarapproach to clone a gene for xylulokinase from S. cerevisiae. Kötter etal. (1990) first reported cloning the gene for xylitol dehydrogenase,Xyl2, from P. stipitis, and, more recently, Shi et al. (2000) sequenced amore complete clone from another strain. Takuma et al. (1991) andHallborn et al. (1991) independently cloned the gene for aldose reduc-tase (Xyl1) from P. stipitis. Billard et al. (1995) cloned a gene with 62%identity to P. stipitis Xyl1, and many other related genes are on depositin GenBank. Hallborn et al. (1995) isolated a short-chain dehydrogenasegene from P. stipitis that has D-arabinitol dehydrogenase activity. Thebiochemical differences among these and other dehydrogenases havebeen reviewed previously (Jeffries and Shi, 1999). Walfridsson et al.(1995) isolated the P. stipitis Tkl1 and Tal1 genes for transketolase andtransaldolase, and Weierstall et al. (1999) isolated the genes for theglucose transporters of P. stipitis. Other genes cloned from P. stipitis include the XynA gene for xylanase (Lee et al., 1986; Basaran et al.,1999), the Cyc1 gene for cytochrome c (Shi et al., 1999), the selectablemarkers LEU2 (Lu et al., 1998a) and Ura3 (Yang et al., 1994), an ARS2


sequence that confers autonomous replication (Yang et al., 1994), thePdhe1α component for pyruvate dehydrogenase (Davis and Jeffries,1997), two genes for pyruvate decarboxylase (Pdc1, Pdc2) (Lu et al.,1998b), and two genes for alcohol dehydrogenase (Adh1, Adh2) (Choand Jeffries, 1998; Passoth et al., 1998).

Fewer genes have been cloned from P. tannophilus. They includegenes for aldose reductase (Bolen et al., 1996), ornithine carbamoyl-transferase (Skrzypek et al., 1990), UDP galactose-4-epimerase(Skrzypek and Maleszka, 1994), cytochrome c (Clark-Walker, 1999a),cytochrome c oxidase subunit 2 (Clark-Walker, 1999b), and OMP decar-boxylase (Ura3) (Clark-Walker, 1998).

Functions for most of these genes can be surmised through compara-tive genomics, but when two or more isomers are present or when thegene products are active in a poorly defined pathway, their functionscannot be determined until physiological effects have been demon-strated. For example, Cho and Jeffries (1998) showed that disruption ofthe PsAdh1 gene resulted in accumulation of xylitol in P. stipitis,whereas disruption of PsAdh2 had no significant effect on cells growingon xylose under oxygen limited conditions. Xylitol dehydrogenase is amember of the alcohol dehydrogenase family, but the principal sub-strate that it acts on is 15 biochemical steps away from ethanol. Xylitoldehydrogenase and alcohol dehydrogenase both use NAD(H) as a co-factor, so deletion of PsAdh1 is thought to cause accumulation of xylitolby increasing the internal concentration of this cofactor. These findingsare in accord with the observed expression of PsAdh1 under fermenta-tive conditions (Passoth et al., 1998; Cho and Jeffries, 1999). ThePsAdh1 gene therefore appears to keep intracellular NADH levels lowand to be responsible for ethanol formation. Pyruvate decarboxylase(PsPdc1) has unusual kinetic (Passoth et al., 1996) and structural (Luet al., 1998b) features that could be important in supplying acetalde-hyde as an electron sink. Some evidence suggests that the PsAdh2 genemight be involved in ethanol oxidation.

Disruption of the Cyc1 gene in P. stipitis has the interesting effect ofgreatly reducing both respiration and cell growth, thereby divertingreductant into ethanol production (Shi and Jeffries, 1999). The cells areable to survive apparently because they possess an alternative oxidase(Jeppsson et al., 1995).

P. stipitis has not been used as extensively as S. cerevisiae for het-erologous expression, but its unique physiology has allowed for a fewinteresting experiments. The heterologous expression of a Cryptococcusxylanase enables P. stipitis to ferment xylan (Morosoli et al., 1993), and


the heterologous expression of S. cerevisiae Ura1, which codes for adihydroorotate dehydrogenase that uses fumarate as an alternative elec-tron acceptor, enables P. stipitis to grow anaerobically on glucose (Shiand Jeffries, 1998).

D. EXPRESSION OF PICHIA GENES IN SACCHAROMYCES

Even though S. cerevisiae does not ferment D-xylose, it does convertD-xylulose to ethanol under microaerobic conditions (Maleszka andSchneider, 1984). It possesses a gene for aldose reductase (Garay-Arroyoand Covar-Rubias, 1999), and it will produce xylitol from xylose. It hasa gene that is very similar to the xylitol dehydrogenase of P. stipitis(Richard et al., 1999), and it also possesses and expresses a gene forD-xylulokinase activity (Deng and Ho, 1990). Therefore, the question is“Why does S. cerevisiae NOT assimilate and produce ethanol on xy-lose?” Researchers have addressed this by examining the effects ofheterologous gene expression of various genes from P. stipitis.

The metabolic engineering of xylose fermentation in S. cerevisiae hasbeen progressively more successful. Genes for P. stipitis xylose reduc-tase (Xyl1) (Amore et al., 1991; Hallborn et al., 1991), Xyl1 plus xylitoldehydrogenase (Xyl2) (Kötter et al., 1990; Tantirungkij et al., 1993,1994a), transketolase (Tkt) plus Xyl1 and Xyl2 (Metzger et al., 1994), orTkt plus transaldolase (TaI) and Xyl1 and Xyl2 (Walfridsson et al., 1995)have been expressed in S. cerevisiae in order to impart xylose fermen-tation.

The introduction of Xyl1 from P. stipitis did not enable S. cerevisiaeto grow on or produce ethanol from xylose. However, heterologousexpression of Xyl1 does enable it to make xylitol from xylose, as longas a supplemental carbon source is provided. Galactose is particularlyuseful because it does not compete with xylose for transport (Kötter andCiriacy, 1993). The presence of both Xyl1 and Xyl2 enables S. cerevisiaeto grow on xylose (Tantirungkij et al., 1994a,b; Meinander et al., 1996),but it is necessary to also overexpress the gene for xylulokinase (Xks1)in order to obtain significant growth or ethanol production on xylose.An S. cerevisiae fusion strain containing PsXyl1, PsXyl2, and the Sac-charomyces gene for xylulokinase (Xks1) shows higher fermentativecapacity on glucose and xylose (Chang and Ho, 1988; Ho et al., 1998).Presumably, overexpression of the native Saccharomyces Xks1 im-proves xylose metabolism in this strain. Rodriguez-Peña et al. (1998)examined the function of Xks1 in well-defined laboratory strains of S.cerevisiae, and they reported that deletion of Xks1 blocked growth onxylulose, but overexpression also had a negative effect on growth. This


suggests that the role for xylulokinase might be rather complex, andthat its expression needs to be closely regulated in S. cerevisiae. Ex-pression of P. stipitis Tkt and Tkl genes does not show significant effectsin recombinant S. cerevisiae strains, which indicates that the nonoxi-dative portion of the pentose phosphate pathway is not limiting in thisyeast.

It is particularly difficult to engineer S. cerevisiae for the fermenta-tion of mixtures of glucose and xylose because glucose interferes withxylose transport in S. cerevisiae, and in the absence of glucose theenzymes responsible for fermentation are not induced. To better under-stand the problem of xylose uptake, van Zyl et al. (1999) expressed P.stipitis Xyl1 and Xyl2 on a multicopy vector in a uracil phosphoribo-syltransferase-deficient (fur1) strain of S. cerevisiae. This enabled cul-tivation of the transformants in a rich medium with autoselection forgrowth on xylose. The researchers used either glucose or raffinose, aslowly metabolized carbon source, as co-metabolizable carbon sources.By using a complex medium rather than a minimal defined medium,they increased xylose utilization twofold. Addition of glucose or raffi-nose as a cosubstrate increased xylose utilization another threefold. Inrich medium with raffinose as the cosubstrate, the transformants con-sumed 50 g/liter of xylose and produced about 5 g/liter each of xylitoland ethanol after 80 hr. Overexpression of Xks1 in this genetic back-ground and with these nutritional conditions would probably increaseethanol production further.

It is unclear from the current literature whether the conditions forethanol production from xylose have actually been optimized withrecombinant S. cerevisiae strains-particularly with respect to aeration.In most instances, the amounts of ethanol produced are well belowwhat would be considered toxic levels for this organism, so the limitingfactors are still not well understood. The fermentation characteristicsof various microbes are shown on Table III.

IV. Critical Parameters for Yeast Xylose Fermentation

Carbon and nitrogen sources, aeration, pH, and temperature are impor-tant for cell growth and product formation. Aeration plays a criticalrole. Oxygen limitation induces fermentation in P. stipitis and C. she-hatae (Alexander et al., 1988; Alexander and Jeffries, 1990; Cho andJeffries, 1998; Passoth et al., 1998). At the same time, these yeastsrequire oxygen for growth and maximal ethanol production (Neirincket al 1984; Rizzi et al., 1989). Aeration is also important for pentose

TABL

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metabolism in S. cerevisiae (Maleszka and Schneider, 1984). Attainingand maintaining optimum aeration rates present special challenges.

A. CARBON SOURCE

Du Preez et al. (1989) found that the maximum ethanol concentrationattained by P. stipitis and C. shehatae was not affected by using D-glu-cose rather than D-Xylose as a substrate. However, Meyrial et al. (1995a)found that the ethanol concentration resulting in growth inhibitiondepended on the sugar consumed. In the case of xylose, growth inhibi-tion occurred at 30 g/liter, but with glucose, cells continued to grow upto 34 g ethanol liter–1. The higher ethanol tolerance observed withglucose as a carbon source correlated with higher plasma membraneH+-ATPase activity (see below, §V.B). Moreover, the carbon source alsodetermined whether or not the ATPase activity was stimulated by ad-dition of 10 g/liter ethanol (Meyrial et al., 1995b). P. stipitis producedmore ethanol in the studies reported by du Preez et al. (1989) than itdid in the studies by Meyrial et al. (1995a). Probably, du Preez attainedthe higher concentrations by using fed-batch fermentation becausesugar concentrations above 50 g/liter adversely affect both P. stipitis andC. shehatae. The fed-batch fermentation might also account for thedifferent effects observed of sugars on ethanol tolerance.

The ethanol tolerance of P. tannophilus changes with the carbonsource used for growth (Jeffries et al., 1985). When cultivated on xyloseas a sole carbon source, this yeast produces only about 20 g/liter ofethanol, but it produces up to 55 g/liter when cultivated on glucose. P.tannophilus also produces ethanol much more rapidly on glucose thanon xylose. Addition of glucose to xylose fermentations by P. tannophi-lus increases ethanol production from the former sugar. Jeffries et al.(1985) hypothesized that this might be attributable to repression ofethanol respiration, but glucose could have affected ethanol toleranceas well.

B. TEMPERATURE

The effects of temperature on growth and ethanol concentration havebeen well studied in the xylose-fermenting yeasts. Prior et al. (1989),Hahn-Hägerdal et al. (1994), du Preez (1994), and McMillan (1994) havereviewed the literature on optimum conditions for yeast xylose fermen-tation in detail.

Temperatures that provide for optimum biomass and ethanol produc-tivities do not necessarily enable maximum ethanol accumulation


(Slininger et al., 1990). This implies that ethanol toxicity affects pro-duction. In P. stipitis, xylitol and residual xylose concentrations in-crease with temperature. Maximum biomass and ethanol productivityby P. stipitis occurred at 26 to 35%. Maximum ethanol selectivity wasachieved at 25 to 26°C. The optimum pH range for growth and fermen-tation on xylose was 4-7 at 25%. In contrast to the low-temperatureoptimum for xylose, ethanol productivity and accumulation for glucosewere optimal at 34°C. Slininger et al. (1985,1990) obtained a maximumof 57 g/liter ethanol with P. stipitis cultivated at 25°C. Du Preez et al.(1986a,b) observed that the limit for ethanol production with P. stipitisincreased from 33 g/liter at 30°C to 43 g/liter at 25°C.

Du Preez et al. (1989) reported that ethanol inhibits growth of P.stipitis and C. shehatae at lower levels than it inhibits ethanol produc-tion. This difference in inhibitory concentrations has also been ob-served with S. cerevisiae (Brown et al., 1981; Lee et al., 1980), Kluy-veromyces fragilis (Rosa et al., 1986), and the bacterium Zymomonasmobilis (Jöbses and Roels, 1986). These observations are probably re-lated. Microbes rely on proton gradients for active transport, and ifplasma membrane ATPase activities are disrupted by ethanol, cellgrowth will stop as nutrient uptake ceases. Facilitated diffusion couldcontinue to support sugar uptake and fermentation, but growth will beimpaired in the absence of mechanisms for nitrogen uptake. Somestrains of xylose-fermenting yeasts produce up to 47 g ethanol · liter–1

at 30°C (Table IV).Du Preez et al. (1989) found that in P. stipitis and C. shehatae ethanol

production was mainly growth associated and that the volumetric rateof ethanol production increased linearly with the volumetric growthrate over a three- to fivefold range. They could not increase the maxi-mum ethanol concentration on D-xylose by increasing the initial celldensity, and they concluded that the low ethanol tolerance of thesexylose-fermenting yeast strains is not a consequence of the metabolicpathway used during pentose fermentation. These results suggest to thisreviewer that the inhibitory effect of ethanol could result from impairedactive uptake systems. Because ethanol production is growth related,any loss of viability or decrease in growth rate will reduce ethanolformation.

Lucas and van Uden (1985) found that ethanol enhances thermaldeath of C. shehatae. The specific growth rate did not vary significantlyfrom its maximum (~31°C) down to 20°C. Maximum ethanol tolerance(6% v/v) occurred over a temperature plateau (10 to 17.5°C). Ethanoldepressed the maximum temperature for growth from 31 to 17.5°C andincreased the minimum temperature for growth from 2.5 to 10°C.


TABLE IVCONCENTRATIONS OF E THANOL I NHIBITORY TO G ROWTH AND E THANOL P RODUCTION BY PICHIA

STIPITIS AND CANDIDA SHEHATAE IN F ED-BATCH F ERMENTATIONS ON X YLOSE AND G LUCOSEa

Carbon Growth FermentationYeast strain source (g/liter) (g/liter)

C. shehatae Y492 Xylose 32.3 44C. shehatae Y798 Xylose 30.5 38.9C. shehatae Y981 Xylose 31.2 45.4C. shehatae Y981 Glucose 34.9 44.8P. stipitis Y663 Xylose 35.1 47.1P. stipitis Y663 Glucose 34.9 43.8

a Data from du Preez et al. (1989).

Acetic acid also shifts growth and thermal death profiles to lowertemperatures (Rodrigues-Alves et al., 1992). Forced cycling of pH 0.5units above and below the optimum of 4.5 decreased the fermentationrate but did not affect ethanol yield (Ryding et al., 1993). Ethanoltolerance of C. shehatae decreases when inhibitory compounds arepresent. This is particularly conspicuous with acid hydrolysates ofwood (Hahn-Hägerdal et al., 1991). C. shehatae was able to tolerate upto 0.4% (v/v) acetic acid at pH 4.5, but its presence in the mediumreduced the temperature growth range from 5 to 34°C to between 21and 27°C. Acetic acid decreased the cell yield by 64% and tolerance toadded ethanol from 5% (v/v) to 2% (Rodrigues-Alves et al., 1992). It ishypothesized that un-ionized acetic acid disrupts proton gradients bydiffusing from the acidified external medium into the cells and disso-ciating in the more neutral environment of the cytoplasm. With loss ofthe proton gradient to drive nutrient uptake, ethanol tolerance declines.

C. pH

Sanchez et al. (1997) found that the best initial pH for ethanol produc-tion from D-xylose by C. shehatae in batch fermentation was 4.5. Underthese conditions, the maximum specific growth rate (µmax) was 0.329hr–1 and the specific ethanol production rate (qE) was 0.72 kg · kg–1 hr–1.The average xylitol yield was 0.078 kg · kg–1, and the overall ethanolyield was 0.41 kg · kg–1. Both the specific substrate uptake rate (qS) andqE diminished once the exponential growth phase was over. A maxi-


mum qE of 0.72 kg · kg–1 hr–1 equates to about 15.6 mmol ethanol g–1

hr–1. By comparison, van Hoek et al. (1998) showed that the fermenta-tive capacity of S. cerevisiae increased with the specific growth rate andranges between 10 and 22 mmol of ethanol g–1 hr–1. Thus, under opti-mal conditions the fermentative capacities of P. stipitis on xylose andS. cerevisiae on glucose do not differ greatly. The performance charac-teristics of several xylose-fermenting yeasts are listed in Table V.

The pH and temperature optima for biomass accumulation by P.tannophilus on xylose were 3.7 and 31.5, respectively, at an initialxylose concentration of 50 g/liter (Roebuck et al., 1995). As in the casefor P. stipitis and C. shehatae, P. tannophilus attains maximum ethanolproductivity under microaerobic conditions (Kruse and Schugerl,

TABLE VPERFORMANCE OF XYLOSE-FERMENTING YEASTS ON GLUCOSE AND XYLOSE

a

strains

Carbonsource(g/liter) Aeration

Eth- Produc- Xy- Bio-anol tivity litol massyield (g/g · yield yield(g/g) 1 · hr) (g/g) (g/g)

Pichia stipitis Xylose (40) AerobicCBS7126 Oxygen limited

Anaerobic

Glucose (40) Aerobic 0.26 0.17 0.23Oxygen limited 0.38 0.28 0.14Anaerobic 0.33 0.13 0.10

AnaerobicPachysolen Xylose (40) Aerobic

tannophilus Oxygen limitedNRRL Y-2460 Anaerobic

Glucose (40) AerobicOxygen limitedAnaerobic

a Data from Ligthelm et al. (1988).

AerobicOxygen limitedAnaerobic

AerobicOxygen limited

Xylose (40)

Glucose (40)

CandidashehataeCBS 2779

0.18 0.17 0 0.390.47 0.20 0.06 0.050.40 0.02 0 0.03

0.22 0.21 0.04 0.330.37 0.32 0.13 0.010.41 0.15 0.18 0.01

0.33 0.35 0.210.42 0.51 0.030.44 0.29 0.02

0.10 0.04 0.17 0.250.28 0.10 0.30 0.010.26 0.07 0.30 0.01

0.31 0.38 0.140.43 0.49 0.060.42 0.18 0.04


1996). However, P. tannophilus xylitol yields tend to be much greater.With a detoxified hemicellulose hydrolysate at pH between 6.0 and 7.5,P. tannophilus converted 90% of the available xylose into xylitol. AtpH values outside this range, cells respired up to 30% of the xylose(Converti et al., 1999). Kavinaugh and Whittaker (1994) reported thatrecycling cells of P. tannophilus NCYC 614 from batch fermentationsover a period of 31 days increased ethanol tolerance in this strain. Thiseffect did not apply to all instances, because they were not able to seea significant increase in ethanol tolerance with P. tannophilus CBS4045.

D. AERATION

The dissolved oxygen tension (DOT) is particularly critical in attainingmaximal ethanol production with xylose-fermenting yeasts. P. stipitisand C. shehatae require aeration for maximal ethanol production (duPreez et al., 1986a,b). Under anoxic conditions, the specific ethanolproductivity of P. stipitis and C. shehatae decreased, and especially inthe case of C. shehatae, xylitol production increased (du Preez et al.,1989). This requirement is not unique to P. stipitis or C. shehatae,because native S. cerevisiae also requires oxygen to metabolize xylulose(Maleszka and Schneider, 1984), and recombinant S. cerevisiae requiresoxygen to produce ethanol from xylose (Kotter et al., 1990). EvenFusarium oxysporum requires oxygen for the fermentation of D-Xyloseand D-glucose (Singh et al., 1992). The oxygen requirement for ethanolproduction was considered novel when first reported (Jeffries, 1981),but it is apparent that oxygen plays various roles in the metabolism ofxylose by eukaryotes. It is important for a xylose-fermenting yeast topossess an aldose reductase that is active with both NADH and NADPHin order to maintain redox balances during xylose assimilation (Ver-duyn et al., 1985), but oxygen enters into xylose metabolism in otherways as well.

One of the factors limiting ethanol production is its simultaneousassimilation. With P. stipitis at ethanol concentrations in excess of 28g/liter, ethanol assimilation exceeds production, even when the dis-solved oxygen tension is kept to 0.2% of saturation. In the absence ofaeration, ethanol accumulation continues, but at a much lower rate, andxylitol production increases (du Preez et al., 1989). Reduced respirationcapacity could be the reason that P. stipitis cyc∆ strains (Shi et al., 1999)exhibit higher specific ethanol production rates. du Preez et al. (1989)attained 47 g · liter–1 ethanol at 30°C with the DOT controlled at 0.2%


of air saturation. Jeffries and Alexander (1990) attained 56 g/liter etha-nol within 38 hr (1.53 g · liter–1 · hr–1) with C. shehatae in fed-batchfermentation after the cell inoculum had been cultivated at a highdilution rate and shifted to oxygen limited conditions.

Candida shehatae requires oxygen to maintain viability. Kastner etal. (1999) showed that oxygen starvation induces cell death in C. she-hatae when it is grown on D-xylose, but not when it is cultivated onD-glucose. Growth of C. shehatae was limited to one division or lesswhen cells cultivated aerobically on either glucose or xylose are shiftedfrom aerobic to anaerobic conditions. Cell viability rapidly declinedwith cells cultivated on xylose, but cells cultivated on glucose remainedviable nine times longer. Shi and Jeffries (1998) also observed differ-ences’ between glucose and xylose during anaerobic cultivation of P.stipitis. The basis for this difference is not fully understood, but it couldreflect differences in the cells to produce metabolic energy on glucoseand xylose under anaerobic conditions or the ability of glucose-growncells to maintain nutrient transport systems. For example, Meyrial etal. (1995a) reported that cultivation of P. stipitis on glucose increasesthe activity of plasma membrane ATPase threefold in comparison to theactivity obtained when cells are grown on xylose. The pH and tempera-ture optima did not shift, and the enzymatic activities showed similaraffinity for ATP However, the glucose-activated enzyme was less sen-sitive to ethanol. These results show that plasma membrane ATPaseactivity, which is critical for transport, correlates with ethanol toleranceand the inhibitory effect of ethanol on growth. Plasma membrane AT-Pase is essential for maintaining the proton gradient that is responsiblefor uptake of nutrients.

Biosynthesis of ergosterol, cardiolipin, and unsaturated fatty acidsrequires oxygen (Hossack and Rose, 1976; Mandal et al., 1978), andexogenous supplies are necessary for the anaerobic growth of S. cere-visiae (Hossack et al., 1977). However, these lipids are not sufficient forthe anaerobic growth of P. tannophilus (Neirinck et al., 1984) or P.stipitis (Shi and Jeffries, 1998). These yeasts, like most other eukaryotes,require active electron transport for the synthesis of uracil, and hencecannot make mRNA under anaerobic conditions. The critical enzymestep that imposes this limitation is dihydroorotate dehydrogenase(DHODase). In most eukaryotes, it is located in the mitochondria, andregeneration of its cofactor requires active electron transport coupledwith respiration. S. cerevisiae possesses an unusual DHODase (ScUra1)that resides in the cytoplasm, which couples the reduction of fumarateto succinate with the regeneration of its cofactor. Expression of ScUra1


in P. stipitis enables this yeast to grow anaerobically on glucose, but noton xylose, so some other factor such as sugar transport or energeticsprobably limits anaerobic xylose metabolism in this organism.

E. NUTRIENT UPTAKE

Membrane transport is mediated by two different systems in yeasts:facilitated diffusion and active proton symport. Facilitated diffusion isenergy independent and functions well at elevated sugar concentra-tions. Proton symport requires generation of a proton gradient but isuseful during growth at low extracellular sugar concentrations. In S.cerevisiae, the transport of glucose into the cells plays a direct role insensing glucose and in signal transduction. S. cerevisiae uses facilitateddiffusion systems to take up hexoses but uses proton symport systemsto take up disaccharides. S. cerevisiae can handle wide ranges of sugarconcentrations up to 1.5 M by developing a group of hexose transport(Hxt) proteins. The presence and the concentration of appropriate sub-strates tightly regulate expression of these enzymes.

In S. cerevisiae, hexose uptake is mediated by a large number ofrelated transporter proteins. Six out of 20 genes for hexose transportmediate the uptake of glucose, fructose, and mannose at metabolicallyrelevant rates (Boles and Hollenberg, 1997). Two others catalyze thetransport of only small amounts of these sugars. One protein is agalactose transporter but is also able to transport glucose. Hexose-trans-port-deficient mutants (hxt) have no clearly detectable phenotypes.Expression of Hxt1, 2, 3, 4, 6, or 7 is sufficient to allow various degreesof glucose utilization (Reifenberger et al., 1997)

In yeasts that utilize both xylose and glucose, these sugars share thesame transporter systems (Boles and Hollenberg, 1997). Glucose caninhibit xylose uptake by competing with the xylose transporters. Even0.05 mM glucose can compete with xylose uptake, which significantlyreduces xylose transport. Xylose transport in P. stipitis is mediated bylow- and high-affinity proton symporters (Weierstall et al., 1999). Bothtransporters are constitutively expressed with low Vmax values. Thelow-affinity system takes up glucose in the range of 0.3-1 mM. More-over, inhibitor studies indicate that uptake of xylose requires aerobicrespiration, which suggests that both systems involve proton symport(Loureiro-Dias and Santos, 1990).

A putative xylose transporter gene from P. stipitis, PsStu1, has beencloned recently (Weierstall et al., 1999). This gene can confer high-af-finity uptake of glucose and growth to a S. cerevisiae hxt1-7 strain(Boles and Hollenberg, 1997). The deduced amino-acid sequence shows


54% identity to the Hxt glucose transporters of S. cerevisiae. WhenPsStu1 is introduced into S. cerevisiae, it can transport xylose but witha considerably lower affinity than what is achieved in P. stipitis. Thissuggests that the transport system in I? stipitis is coupled to otherelements. Two other Stu-related genes have been identified from P.stipitis by crosshybridization with S. cerevisiae transporter genes asprobes. Uptake of xylose in S. cerevisiae is mediated nonspecificallyand with low affinity by the hexose transporters, so other factors wouldbe required to increase xylose metabolism in this yeast.

In S. cerevisiae genetically engineered for xylose uptake, glucose,mannose, and fructose inhibited xylose conversion by 99, 77, and 78%,respectively. These sugars are transported with by the same high-affin-ity transport system as xylose, and the results are thought to reflectcompetitive inhibition of xylose transport (Meinander and Hahn-Häger-dal, 1997). Galactose is less inhibitory to xylose transport than is glu-cose and was therefore a better co-metabolizable carbon source forxylitol production. Membrane transport plays an important role in theutilization of xylose and other sugars in lignocellulose hydrolysates,and it can limit utilization of sugars (Spencer-Martins, 1994).

V. Factors Affecting Thermo- and Ethanol Tolerance

Ethanol tolerance is very important in brewing, wine making, andespecially in the biosynthesis of industrial ethanol. Because it is socritical, it has been studied extensively (e.g., Mishra and Singh, 1993).The mechanisms underlying ethanol resistance are very complex. Theydiffer from one yeast to another (Alexandre et al., 1994) and with theconditions for cultivation. Many genes appear to be involved (D’Amoreel al., 1990), and the exact basis for ethanol tolerance is not fullyunderstood (Chi et al., 1999). Factors that affect ethanol tolerance in-clude the proportion of ergosterol in the cellular membranes, phos-pholipid biosynthesis, the degree of unsaturation of membrane fattyacids (Alexandre et al., 1994), temperature, the activity of plasma mem-brane ATPase, superoxide dismutase, and the capacity of a strain toproduce trehalose.

Sublethal heat and ethanol exposure induce essentially identicalstress responses in yeast (Piper, 1995). Factors affecting the capacity ofyeast to survive at high temperatures include the presence of stress-re-sponse pathways to signal induction of appropriate heat-shock pro-teins. One induced protein, Hsp104, contributes to both thermotoler-ance and ethanol tolerance. Heat and ethanol stress cause similarchanges to plasma membrane protein composition, reducing the levels


of plasma membrane H+-ATPase protein and inducing the plasma mem-brane-associated Hsp30.

A. MEMBRANE LIPIDS

Temperature is a critical variable that affects both growth and ethanoltolerance. Decreasing temperature decreases membrane fluidity; in-creasing temperature increases membrane fluidity. According to theprinciple of homeoviscous or homeophasic adaptation, the cell mustcompensate for environmental changes by altering its composition tomaintain fluidity at the new temperature (Vigh et al., 1998). The mannerin which this adaptation occurs, however, can vary with cell type andthe conditions. Organisms commonly adapt to low temperatures byincreasing the proportion of cis-unsaturated fatty-acyl groups in theirmembrane lipids. Physical principles suggest that fluidity would de-crease as the ratio of saturated to unsaturated fatty acids increasesbecause desaturation introduces a bend in the fatty acid chain. How-ever, the bulk of fatty acids in the membranes of S. cerevisiae areunsaturated (Fig. 1), so other factors may be more important.

For example, when S. cerevisiae cells are grown for an extendedperiod at 37°C, they adapt to the higher temperature by increasing theircontent of unsaturated fatty acids. The cells adapted to 37°C require ahigher temperature in order to induce heat-shock proteins (Chatterjeeet al., 1997). Guerzoni et al. (1997) found that the unsaturation level ofS. cerevisiae cellular fatty acids increases at both sublethal or supraop-timal temperatures. The adaptation can be reversed by cultivation at25°C. They hypothesized that a high content of unsaturated fatty acidsis not in itself a prerequisite for withstanding temperature stress, butrather results from activation of oxygen-consuming desaturase activity.

Membrane fluidity is affected by the ratios of cell lipids and proteins.These vary with the yeast strain and the conditions under which it iscultivated. Alexandre et al. (1994) found that the average protein, lipid,phospholipid, and sterol contents of two S. cerevisiae strains wereabout 30, 15, 3.5, and 1.5%, respectively. When cultivated with 4 or10% added ethanol, the lipid, phospholipid, and sterol concentrationsall decreased. S. cerevisiae FY 169—the strain used in the EuropeanUnion Yeast Genome Sequencing Program—is atypical in that it hassignificantly higher palmitic (C16:0) and lower amounts of oleic (C18:1)fatty acids as compared to other wild-type yeasts. Therefore, the datashown in Figure 1 would be even more biased toward unsaturated fattyacids in a typical strain of S. cerevisiae (Daum et al., 1999).


FIG. 1. Fatty acid composition of subcellular membranes from late logarithmic phasecells of Saccharomyces cerevisiae FY 169 cultivated on 2% glucose or 2% ethanol at30°C under aerobic conditions (Tuller et al., 1999). PM = plasma membrane, Mito =mitochondria, Mic = microsomes (endoplasmic reticulum), Vac = vacuoles; C numbersindicate chain length and degree of unsaturation.

Decreases in the sterol:protein and sterol:phospholipid ratios, and anincrease in the unsaturation index all increase membrane fluidity. How-ever, fluidity indirectly deduced from the sterol:protein, sterol:phos-pholipid, or unsaturated:saturated fatty acid ratios does not alwaysreflect the real state of the plasma membrane (Alexandre et al., 1994).It is necessary to measure membrane fluidity directly. Swan and Watson(1999) substituted various unsaturated fatty acids in a ∆9-desaturase(olei1) mutant and found that the most heat- and ethanol-tolerant cellshad membranes enriched in oleic acid (C18:1). S. cerevisiae does notproduce the polyunsaturated linoleic (18:2) and linolenic (18:3) fattyacids, which are present in other plants and fungi. To investigate the


possible roles of such lipids in ethanol resistance, Kajiwara et al. (1996)examined the heterologous expression of an Arabidopsis thaliana ∆1 2

desaturase in S. cerevisiae. They showed that the transformed yeast iscapable of accumulating hexadecadienoyl- (16:2) and lineloeoyl- (18:2)substituted membrane lipids and that the cells had greater resistanceto ethanol than the controls.

Ethanol also affects membrane fluidity, but through different mecha-nisms. Alcohols lower the temperature required for maximal activationof heat-shock genes, and the concentration of alcohol needed decreaseswith alcohol chain length (Vigh et al., 1998). Ethanol is thought to altermembrane organization and permeability by entering the hydrophobicinterior and increasing the polarity of this region (Alexandre et al.,1993). This would weaken hydrophobic interactions and increase thediffusion of polar molecules through the membrane. Ethanol in the cellmembrane is said to weaken the water lattice structure, decrease thestrength of lipophilic interactions, promote membrane leakage, anddecrease the integrity of the semipermeable barrier (Sajbidor and Grego,1992; Sajbidor, 1997). Such changes would put extraordinary demandson active transport systems for nutrient uptake. Jones and Greenfeld(1987) found that, beyond a critical threshold concentration, the fluid-ity of the yeast plasma membrane increases exponentially. Cells grownunder anaerobiosis have a lower level of fatty acids than cells grownaerobically, but the anaerobically grown cells exhibit higher membranefluidity. Sensitivity to both heat and oxidative stress depends on mem-brane lipid composition. In the case of anaerobically grown cells, themost stress resistant have membranes enriched in saturated fatty acids(Steels et al., 1994).

An increase in fatty acid unsaturation in cellular membranes in-creases ethanol tolerance (Alexandre et al., 1994). Sterols, especiallyergosterol, promote growth and ethanol tolerance by providing rigidityto the cell membrane (Zinser et al., 1991). The plasma membrane isparticularly rich in ergosterol, and the fraction increases with growthon ethanol (Fig. 2). Total sterol content of cells increases with cultiva-tion under anaerobic conditions and decreases under aerobic condi-tions (Sajbidor et al., 1995). Ergosterol plays an important role in yeaststress tolerance. It stabilizes cell membranes independently of heat-shock proteins or trehalose. Swan and Watson (1999) concluded thatmembrane lipid composition plays a more consistent role in stresstolerance than trehalose, heat-shock proteins, or ergosterol. The S. cere-visiae FY 169 genome type strain also has a relatively low total sterolcontent and tends to accumulate lanosterol, the sterol precursor (Daumet al., 1999).


FIG. 2. Sterol composition of subcellular membranes from late logarithmic phase cellsof Saccharomyces cerevisiae FY 169 cultivated on 2% glucose or 2% ethanol at 30°Cunder aerobic conditions (Tuller et al., 1999). PM = plasma membrane, Mito = mitochon-dria, Mic = microsomes (endoplasmic reticulum).

Phospholipids have not been examined as extensively as fatty acidsand ergosterol for their effects on ethanol tolerance, but recent studiesby Chi et al. (1999) suggest that phosphatidylinositol (PI) may be criticalin determining ethanol tolerance in yeast strains that are capable of highethanol production. PI is among the most abundant phospholipids ofyeast membranes (Fig. 3), and addition of inositol increases phospho-lipid synthesis. Inositol can become the factor determining ethanoltolerance in cells that have sufficient fatty acids and ergosterol. Syn-thesis of phosphatidylcholine (PC) from phosphatidylethanolamine(PE) is induced by inositol (Greenberg and Lopes, 1996). Early work byMishra and Prasad (1988) showed that cells enriched with phospha-tidylserine had greater tolerance to ethanol, and they concluded that


FIG. 3. Phospholipid composition of subcellular membranes from late logarithmicphase cells of Saccharomyces cerevisiae FY 169 cultivated on 2% glucose or 2% ethanolat 30°C under aerobic conditions (Tuller et al., 1999). PM = plasma membrane = Mito =mitochondria, Mic = microsomes (endoplasmic reticulum), Vac = vacuoles, PtdCho =phosphatidylcholine, PtdEtn = phosphatidylethanolamine, PtdIns = phosphatidylinosi-tol, PtdSer = phosphatidylserine, PtdDMEtn = phosphatidylmethylethanolamine, CL =cardiolipin, PA = phosphatidic acid, Lyso-PL = lysophospholipids, Lyso-PtdEtn =lysophosphatidylethanolamine.

this resulted from an alteration in the charge of the membrane phos-pholipids rather than changes in membrane fluidity. Even so, ergosterolappears to be more important than phospholipids. For example, Chi etal. (1999) found that a more ethanol-tolerant strain of S. cerevisiae hada higher ergosterol:phospholipid ratio, a higher incorporation of long-chain fatty acids in total phospholipids, and a slightly higher propor-tion of unsaturated fatty acids in total phospholipids than a less etha-nol-tolerant strain. Likewise, el Dein (1997) found that in Candida


lambica, a yeast with only moderate ethanol tolerance, growth in up to8% (v/v) ethanol was accompanied by an increase in total fatty acidsbut a decrease in phospholipids. Murakami et al. (1996) did not observea significant correlation between phospholipid content and toleranceto freezing, but it is possible that these and other researchers haveneglected inositol as a limiting factor in yeast phospholipid biosynthe-sis.

Neirinck et al. (1984) and Shi and Jeffries (1998) have examined theeffects of lipids on anaerobic growth of pentose-fermenting yeasts.Neirinck et al. (1984) were not able to obtain anaerobic growth of P.tannophilus even after incorporating essential lipids into the medium.Shi and Jeffries (1998) did obtain anaerobic growth of P. stipitis trans-formed with ScUra1 by adding ergosterol, Tween 80, and linoleic acidto the medium (Jessens et al., 1983) along with fumarate as a terminalelectron acceptor for the DHODase activity (Nagy et al., 1992). However,the transformants grew on glucose but not on xylose. Ethanol yieldswere comparable to those attained by S. cerevisiae control cultures. Toour knowledge, the effect of ergosterol, unsaturated fatty acids, inositol,or phospholipids on ethanol or thermotolerance has not been investi-gated with xylose-fermenting yeasts.

Eubacteriales do not produce ergosterol, and, for the most part, theydo not tolerate high ethanol concentrations either. Zymomonas mobilisproduces large amounts of ethanol rapidly. It has cardiolipin, phospha-tidylethanolamine, phosphatidylglycerol, and phosphatidylcholine asits major phospholipids (Carey and Ingram, 1983). In yeasts, cardiolipinis found almost exclusively in the mitochondria (Fig. 3), and its con-centration increases when cells are cultivated in the presence of etha-nol. Z. mobilis also produces large amounts of pentacycline triterpa-noid lipids known as hopanoids (Moreau et al., 1997). The structuresof these compounds superficially resemble sterols. In the absence ofethanol, hopanoid content can be as high as 30% of total cellular lipids.In the presence of ethanol, complex changes occur in the levels ofhopanoids and other membrane constituents. Various researchers havehypothesized that either hopanoid or cis-vaccenic acid in the phos-pholipids of the bacterial membrane could account for the ethanoltolerance of Z. mobilis, but such relationships have not been estab-lished unambiguously. The heterologous expression of genes for ho-panoid biosynthesis (Perzl et al., 1998) could help establish whetherthese lipids play a major role in bacterial ethanol tolerance. Hobley andPamment (1994) showed that one of the reasons Z. mobilis is able totolerate high concentrations of ethanol is that it adapts rapidly tochanges in extracellular ethanol without significant viability loss.


The biochemical, molecular, genetic (Daum et al., 1998), and envi-ronmental (Sajbidor, 1997) factors affecting lipid synthesis in S. cere-visiae and other organisms have been reviewed. Lipid composition byitself probably does not affect ethanol tolerance. Rather, the physiologi-cal function that the various lipids play in maintaining membraneintegrity is the critical factor, and the most important function of themembrane is nutrient transport and accumulation,

B. PLASMA MEMBRANE H+-ATPASE

The plasma membrane (PM) proton pump (H+-ATPase) of yeast couplesATP hydrolysis to proton extrusion, thereby providing the means forsolute uptake by secondary transporters and for regulating cytoplasmicpH. By pumping protons out of the cytoplasm, the H+-ATPase acidifiesthe external medium, and makes the cytoplasm relatively alkaline. ThePM H+-ATPase is a highly abundant, essential enzyme in S. cerevisiae.It belongs to the family of P-type ATPases, a class of enzymes thatincludes the Na+,K+-ATPase and the gastric H+,K+-ATPase (Monk et al.,1995).

S. cerevisiae possesses two plasma membrane H+-ATPase isoforms:Pma1 and Pma2. They are 89% identical at the protein level (Supplyet al., 1995), but they exhibit different activation, kinetic, and regula-tory properties, which suggests that they have different functions. Themajor yeast plasma membrane H+-ATPase is encoded by the essentialPma1 gene. The Pma2 gene encodes an H+-ATPase that is functionallyinterchangeable with the one encoded by Pma1, but it is expressed ata much lower level than Pma1 and it is not essential (Fernandes andSá-Correia, 1999). Pma1 is primarily responsible for proton transloca-tion in S. cerevisiae. The expression of Pma1 is regulated by glucoseand by the Tuf/Rap1/Grf1 transcription factor in S. cerevisiae (Rao etal., 1993). Deletion of Apa1 causes defective regulation of Pma1 expres-sion by glucose but has no noticeable effect on expression of otherTuf-regulated genes (Garcia-Arranz et al., 1994). Pma1 is activated byglucose (Serrano, 1983) through a mechanism that involves the Cdc25gene product (Portillo and Mazon, 1986). Cdc25 and Ras are two pro-teins required for cAMP signaling in S. cerevisiae. Cdc25 is the proto-type guanine nucleotide exchange protein that activates Ras. Ras, inturn, activates adenylyl cyclase (Mintzer and Field, 1999). The N-ter-minal portion of Cdc25 is important in signal processing (Gross et al.,1999).

The specific activity of Pma1 increases with growth temperature.However, this activation does not result from increased Pma1 synthesis,


Expression levels actually decrease along with the amount of enzymepresent in the membrane (Viegas et al., 1995). Ethanol also activatesATPase in S. cerevisiae (Monteiro and Sá-Correia, 1998). Ethanol-in-duced stimulation of plasma membrane H+-ATPase increases the Vm a x .The affinity for Mg+ATP decreases. Sublethal concentrations of ethanolenhance expression of Pma2 while reducing expression of Pma1. Theinhibition of Pma1 expression by ethanol correlates with a decline inthe content of plasma membrane ATPase as measured by immunoassays(Monteiro et al., 1994). Therefore, the observed increase in activityfollowing stress is not attributable to synthesis of new protein, butrather to activation of the existing enzyme.

Piper et al. (1997) reported that in S. cerevisiae, Hsp30 is a stress-in-ducible regulator of ATPase activity. Hsp30 is induced by heat shock,ethanol exposure, severe osmostress, weak organic acid exposure, andglucose limitation. Hsp30 induction downregulates stimulation of H+-ATPase caused by stress. Plasma membrane H+-ATPase consumes asubstantial fraction of the ATP generated by the cell. Hsp30 mighttherefore play an energy conservation role during prolonged stressexposure or glucose limitation (Piper et al., 1997).

Barbosa and Lee (1991) were the first to study the effects of alcoholson PM ATPase activity in a xylose-fermenting yeast. In-vitro studiesshowed that the amount of alcohol required to inhibit ATPase activity50% decreased with increasing chain length. Moreover, the concentra-tion required to inhibit ATPase activity corresponded approximatelywith the concentration required to inhibit growth.

Meyrial et al. (1995a,b, 1997) carried out extensive studies of PMATPase activity in P. stipitis. The enzyme from this yeast attained itshighest activity between pH 7.3 and 7.5 at 35°C. This value is very farfrom the ATPase activities in S. cerevisiae or K. marxianus (Rosa andSá-Correia, 1992), and Meyrial et al. (1995a) concluded from a compari-son of the pH optima of ATPases reported from different yeasts that themost ethanol-tolerant have ATPases with low pH optima. They alsonoted that the P. stipitis and K. marxianus Km values for ATP are about2- to 10-fold higher than the values reported for S. cerevisiae. A higheraffinity for ATP should enable the ATPase to maintain proton gradientseven under conditions where energy generation is low, such as duringanaerobic growth. They found that glucose stimulates the activity of P.stipitis PM ATPase threefold in comparison to the activity measuredwhen xylose is used as the carbon source (Meyrial et al., 1995a). Thestimulation occurs mainly through doubling of Vmax. The Km value forATP remains similar in both glucose- and xylose-grown cells. Thestimulation of ATPase activity correlates with an increase in the maxi-


mum amount of ethanol produced by glucose-grown cells. It is possiblethat glucose activation of ATPase is related to the observation by Shiand Jeffries (1998) that P. stipitis transformed with ScUra1 will growanaerobically on glucose but not on xylose. The concentration of etha-nol that is required to cause a 50% inhibition of ATPase activity is 34.5and 41.4 g/liter in xylose- and glucose-grown cells, respectively. Forboth glucose- and xylose-grown cells, the concentration of ethanolrequired to inhibit growth reduced ATPase activity of control cells(grown without ethanol) by 60%.

Ethanol also stimulates ATPase activity in P. stipitis (Meyrial et al.,1995b); unlike S. cerevisiae, however, the extent of stimulation dependson the carbon source used for growth. With glucose, addition of 1%ethanol doubles ATPase activity. No significant increase in ATPaseactivity is observed with xylose-grown cells. Moreover, the stimulatoryeffects of glucose and ethanol are additive. Similar activation of ATPaseactivity has been reported for S. cerevisiae and K. marxianus (Rosa andSá-Correia, 1991, 1992). It is unclear whether ATPase activity in P.stipitis involves one protein or two, as in the case of S. cerevisiae.

Physiological concentrations of ethanol do not affect passive entry ofprotons into either glucose- or xylose-grown P. stipitis (Meyrial et al.,1997). In contrast, active proton extrusion is affected differently byethanol, depending on the carbon source used for cell growth. Low,physiological concentrations of ethanol decrease the proton extrusionrate in glucose-grown cells, but only nonphysiological ethanol concen-trations reduce proton extrusion in xylose-grown cells. By comparingthe rates of proton efflux with ATPase activity, Meyrial et al. (1997)concluded that glucose activates both ATP hydrolysis and the proton-pumping activities of H+-ATPase, whereas ethanol causes an uncou-pling between ATP hydrolysis and proton extrusion. Heat shock andethanol stress cause similar changes in the protein composition of thePM. Heat shock and ethanol stress each reduce the levels of plasmamembrane H+-ATPase while simultaneously increasing the specific ac-tivity of the remaining enzyme. Proton efflux places an energy demandbut may help to counteract the adverse effects on membrane permeabil-ity that result from stress (Piper, 1995).

Plasma membrane ATPase activity is essential for basal heat resis-tance. Moreover, thermotolerance is enhanced by prior exposure tostress. Prestressed cells are able to protect the proton gradient longerthan cells that have not adapted to heat or ethanol (Coote et al., 1994).Stress resistance is largely conferred by a mechanism independent ofthe ATPase. A mutant strain with reduced expression of plasma mem-brane ATPase showed significantly less resistance to lethal heat than


the wild-type parent. However, prior exposure to sublethal tempera-tures induced levels of thermotolerance similar to the parent strain,suggesting that the mechanism of sublethal heat-induced thermotoler-ance is independent of ATPase activity (Coote et al., 1994).

C. MITOCHONDRIAL STABILITY

Mitochondria are essential for ethanol tolerance in S. cerevisiae (Aguil-era and Benítez, 1985). This presents a difficulty because ethanol alsotends to induce the formation of petite mutants (Jiménez et al., 1988).In fact, the mitochondria in populations of flor yeasts isolated from

sherry wine show an unusual level of mitochondrial DNA polymor-phism (Ibeas and Jiménez, 1997). Treatment with ethanol did not causean increase in the spontaneous mutation rate of nuclear genes, butpetite mutants increased from a low spontaneous level (about 1%) tonearly half the survivors at 24% ethanol (Bandas and Zakharov, 1980).

Chi and Arneborg (1999) demonstrated that ethanol tolerance corre-lated with a lower frequency of ethanol-induced respiratory-deficientmutants, a higher ergosterol:phospholipid ratio, a higher proportion ofphosphatidylcholine, a lower proportion of phosphatidylethanolamine,a higher incorporation of long-chain fatty acids in total phospholipids,and a slightly higher proportion of unsaturated fatty acids in totalphospholipids (Chi and Arneborg, 1999).

Manganese superoxide dismutase (MnSOD), which is found in mito-chondria, is essential for ethanol tolerance in S. cerevisiae (Costa et al.,1993). In E. coli, MnSOD is associated with DNA (Steinman et al.,1994), and in yeasts it might serve to prevent damage to mitochondrialDNA. Ethanol toxicity correlates with the production of reactive oxygenspecies in the mitochondria, as evidenced from results obtained withrespiration-deficient mutants (Costa et al., 1997). The freezing-and-thawing process also generates free radicals and oxidative stress, whichcause major injury to cells during aerobic freezing and thawing. Thesuperoxide radical is mainly generated on the matrix side of the innermitochondrial membrane of yeast cells (Balzan et al., 1999). Superoxideradicals from the mitochondrial electron transport chain initiate dam-age in the cytoplasm (Park et al., 1998). Normally such radicals aredispersed by cytoplasmic Cu and Zn superoxide dismutase. It is alsopossible to protect the cells against oxidative stress by heterologousexpression of Escherichia coli iron superoxide dismutase (FeSOD)when it is targeted to the mitochondrial matrix (Balzan et al., 1995).Nedeva et al. (1993) used thermotolerance as a screening mechanismto identify yeasts that produce heat-stable superoxide dismutase. After


screening 10 thermotolerant yeasts from the genera Saccharomyces,Kluyveromyces, and Pichia, they found that higher cell yields at 45°Cwere obtained with two thermotolerant strains of K. lactis, which alsoproduced thermostable superoxide dismutase. Because the free radicalsgenerated during respiration are so destructive, they might be the rootcause for the frequency of petites formed at high temperatures andelevated ethanol level.

D. TREHALOSE

Tolerance to heat, freezing, oxidation, and desiccation overlap to someextent with ethanol tolerance, and all but ethanol tolerance correlatewith trehalose accumulation (Piper, 1995; Eleutherio et al., 1995; Lewiset al., 1997). Roles for trehalose in preventing protein denaturationduring desiccation or at elevated temperature are fairly well estab-lished, but its role in ethanol tolerance is less clear. Trehalose is barelydetectable in log-phase cells grown on glucose, but it accumulates upto 20% of the total dry weight of the cell in stationary phase cells aswell as in exponential-phase cells growing at elevated temperature.Cells growing on nonfermentable carbon sources have high levels oftrehalose in both log-phase and stationary-phase cells (Nwaka andHolzer, 1998).

Trehalose is a nonreducing disaccharide of glucose that is present inyeasts and higher organisms. It enables cells to survive desiccation andosmotic and thermal stress (Singer and Lindquist, 1998). Trehaloseaccumulates in the cytosol of yeast during heat shock, where it providesprotection to proteins. Trehalose is synthesized from UDP-glucose andglucose-6-phosphate in a two-step reaction catalyzed by the trehalose-6-phosphate synthase (Tps1) and trehalose-6-phosphate phosphatasecomplex (Piper, 1993). Ribeiro et al. (1997) used a tps1 mutant of thefission yeast Schizosaccharomyces pombe in a genetic approach todetermine the specific role of trehalose in a heat-induced cell. Theyshowed that the tps1 mutant has a serious defect in heat-shock-inducedacquisition of thermotolerance at 40 to 42.5°C. Earlier studies that usedcycloheximide to block protein synthesis found that heat shock did notinduce S. pombe to produce trehalose (DeVirgilio et al., 1990). Thediscrepancy between these two studies might be attributable to thetoxicity of cycloheximide. When cells are heat shocked, trehalose im-mediately starts to accumulate, even before the induction of enzymesfor trehalose metabolism (Neves and Francois, 1992). This shows thatrapid increase in trehalose is related not to induction of enzymes fortrehalose synthesis but to activation of preexisting enzymes. Trehalose


stabilizes enzymes and also greatly reduces their activities. Trehalosedegradation is therefore necessary in order for the cell to resume activ-ity once the stress condition has passed. This is carried out by trehalase.In vitro, the activity of trehalase decreases as the activity of the treha-lose-6-phosphate synthase-phosphate complex increases (Neves andFrancois, 1992). Hexokinase also appears to act synergistically withtrehalose synthesis because it provides glucose-6-phosphate for treha-lose synthesis. After incubation in D-xylose, an inhibitor of hexokinase,trehalose levels in these cells dropped almost in 90%, confirming theinvolvement of both hexokinases in the accumulation of trehalose(Peixoto and Panek, 1999).

Probably trehalose will be found to play a role in thermotolerance,osmotolerance, and resistance to ethanol because the mechanism is sowidely distributed among yeasts, fungi, and plants. For example, tre-halose synthesis is an important factor in thermotolerance of the fissionyeast S. pombe. On induction of trehalose-6P synthase, the elevatedlevels of intracellular trehalose correlated not only with increased tol-erance to heat shock but also with resistance to freezing and thawing,dehydration, osmostress, and toxic levels of ethanol (Soto et al., 1999).The broad nature of this stress response suggests that trehalose may bethe metabolite underlying the overlap in induced tolerance.

An exhaustive search of the current literature did not uncover anyreports about trehalose synthesis in xylose-fermenting yeasts.

E. HEAT-SHOCK PROTEINS

When yeasts confront extreme temperatures (50°C), synthesis of heat-shock proteins (Hsps) is induced (Parsell and Lindquist, 1993). Theseproteins play important roles in protecting cells from extreme tempera-ture. In S. cerevisiae, Hsp90 and Hsp70 are expressed even at normaltemperature and are only modestly induced by heat (Craig and Jacob-sen, 1984). Hsp104 is expressed at a very low level at normal tempera-tures and is very strongly induced by heat. Hsp104 expression is criticalfor survival following heat shock in yeast (Lindquist and Kim, 1996).Recent research indicates that Hsp104 promotes survival by resolubi-lizing heat-damaged aggregated proteins (Parsell et al., 1994; Schirmeret al., 1998). The function of Hsp104 is to repair damage after stressrather than to prevent damage from stress, as in the case of trehalose.

Hsp30, the integral plasma membrane heat-shock protein of S. cere-visiae, is a stress-inducible regulator of plasma membrane H+-ATPase(Piper et al., 1997). Cells lacking Hsp30 showed greater plasma mem-brane H+-ATPase activity than cells with normal levels of Hsp30. This


indicates that Hsp30 reduces stimulation of H+-ATPase. Hsp30 reducesthe Vmax of H+-ATPase in heat-shocked cells. The action of Hsp30 is lostwith deletion of the C-terminal 11 amino acids of H+-ATPase, a deletionthat does not abolish stress stimulation of this enzyme (Braley andPiper, 1997). This suggests that Hsp30 modulates ATPase activity byinteracting with this portion of the enzyme.

VI. Summary

The mechanisms underlying ethanol and heat tolerance are complex.Many different genes are involved, and the exact basis is not fullyunderstood. The integrity of cytoplasmic and mitochondrial mem-branes is critical to maintain proton gradients for metabolic energy andnutrient uptake. Heat and ethanol stress adversely affect membraneintegrity. These factors are particularly detrimental to xylose-ferment-ing yeasts because they require oxygen for biosynthesis of essential cellmembrane and nucleic acid constituents, and they depend on respira-tion for the generation of ATP. Physiological responses to ethanol andheat shock have been studied most extensively in S. cerevisiae. How-ever, comparative biochemical studies with other organisms suggestthat similar mechanisms will be important in xylose-fermenting yeasts.The composition of a cell’s membrane lipids shifts with temperature,ethanol concentration, and stage of cultivation. Levels of unsaturatedfatty acids and ergosterol increase in response to temperature andethanol stress. Inositol is involved in phospholipid biosynthesis, and itcan increase ethanol tolerance when provided as a supplement.

Membrane integrity determines the cell’s ability to maintain protongradients for nutrient uptake. Plasma membrane ATPase generates theproton gradient, and the biochemical characteristics of this enzymecontribute to ethanol tolerance. Organisms with higher ethanol toler-ance have ATPase activities with low pH optima and high affinity forATP. Likewise, organisms with ATPase activities that resist ethanolinhibition also function better at high ethanol concentrations. ATPaseconsumes a significant fraction of the total cellular ATP, and understress conditions when membrane gradients are compromised the ac-tivity of ATPase is regulated. In xylose-fermenting yeasts, the carbonsource used for growth affects both ATPase activity and ethanol toler-ance. Cells can adapt to heat and ethanol stress by synthesizing treha-lose and heat-shock proteins, which stabilize and repair denaturedproteins. The capacity of cells to produce trehalose and induce HSPscorrelate with their thermotolerance. Both heat and ethanol increasethe frequency of petite mutations and kill cells. This might be attribut-


able to membrane effects, but it could also arise from oxidative damage.Cytoplasmic and mitochondrial superoxide dismutases can destroy oxi-dative radicals and thereby maintain cell viability. Improved knowl-edge of the mechanisms underlying ethanol and thermotolerance in S.cerevisiae should enable the genetic engineering of these traits in xy-lose-fermenting yeasts.

Acknowledgments

The authors acknowledge H. K. Sreenath for the use of unpublisheddata and N. Q. Shi for a critical reading of the manuscript. Research forthis review was supported in part by NREL contract No. XXL-9-29034-02 to T. W. J.

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