The declining rate of ethanol production during batch fermentation by Saccharomyces cerevisiae

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The declining rate of ethanol production during batch fermentation by Saccharomyces cerevisiae
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Dombek, Kenneth Michael, 1959-
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Fermentation   ( lcsh )
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Microbiology and Cell Science thesis Ph.D
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Thesis:
Thesis (Ph. D.)--University of Florida, 1987.
Bibliography:
Bibliography: leaves 152-165.
Statement of Responsibility:
by Kenneth Michael Dombek.
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Typescript.
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Vita.

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Full Text












THE DECLINING RATE OF ETHANOL PRODUCTION
DURING BATCH FERMENTATION BY SACCHAROMYCES CEREVISIAE










BY

KENNETH MICHAEL DOMBEK


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN
PARTIAL FULFILLMENT OF THE REQUIREMENTS
FOR THE DEGREE OF DOCTOR OF PHILOSOPHY


UNIVERSITY OF FLORIDA


1987















ACKNOWLEDGMENTS

The ideas presented in these studies could not have

been developed without the encouragement and patience of my

major advisor, Dr. Neal Ingram. I am greatly indebted to

him for sharing with me his knowledge and expertise. I also

would like to express my gratitude to the other members of

my committee, Dr. Allen, Dr. Farrah, Dr. Gander and Dr.

Preston, for their contributions during preparation and

review of this manuscript. Similarly, thanks are due to my

colleague and friend, Dr. Yehia Osman, for his many

suggestions which were helpful in completing this work and

to the rest of the Microbiology and Cell Science Department

for their part in my graduate education. Finally, I would

like to thank my parents for their love and support while I

pursued this study.
















TABLE OF CONTENTS

Page

ACKNOWLEDGMENTS............... ......................... ii

LIST OF TABLES.......................................... v

LIST OF FIGURES...................................... .. vi

ABSTRACT...................................................... viii

CHAPTERS

I GENERAL INTRODUCTION............................ 1

II CHARACTERIZATION OF THE DECLINING RATES OF
GROWTH AND ETHANOL PRODUCTION DURING BATCH
FERMENTATION BY S. CEREVISIAE KD2............... 12

Introduction..................... ............. 12
Materials and Methods........................... 14
Results........................................... 18
Discussion...................................... 27

III NUTRIENT LIMITATION AS A BASIS FOR THE
APPARENT TOXICITY OF LOW LEVELS OF ETHANOL
DURING BATCH FERMENTATION....................... 30

Introduction.............. ........ ............. 30
Materials and Methods........................... 30
Results................................... ......36
Discussion............... ....................... 50

IV MAGNESIUM LIMITATION AND ITS ROLE IN THE
APPARENT TOXICITY OF ETHANOL DURING YEAST
FERMENTATION....... ................ ............ 56

Introduction. .................................. 56
Materials and Methods........................... 59
Results............................ ..... ....... 62
Discussion.... .............. ................ 80

V GLYCOLYTIC ENZYMES AND INTERNAL pH.............. 84

Introduction................................... 84

iii










Materials and Methods........................... 86
Results........................................... 89
Discussion........................... ........... 103

VI PHOSPHORYLATED GLYCOLYTIC INTERMEDIATES AND
NUCLEOTIDES............... ..................... 113

Introduction................ ................. 113
Materials and Methods........................... 116
Results ......................................... 125
Discussion...................................... 139

VII SUMMARY AND FUTURE DIRECTIONS................... 146

BIBLIOGRAPHY................... ........................ 152

BIOGRAPHICAL SKETCH.. .................................. 166















LIST OF TABLES


Page


Table 1.


Table 2.


Table 3.


Table 4.


Table 5.


Effects of ethanol and fermentation medium
composition on fermentation rate..............

Intracellular and extracellular ethanol
concentrations under various conditions.......

Effect of growth in broths of different
composition on fermentation rate.............

Effect of nutrient supplementation on growth
of S. cerevisiae KD2..........................

Specific activities of glycolytic enzymes at
the peak of fermentative activity (12 h) and
after a 50% decline (24 h) ....................















LIST OF FIGURES


Page


Figure 1.




Figure 2.


Figure 3.


Figure 4.


Figure 5.


Figure 6.


Figure 7.


Figure 8.


Figure 9.


Figure 10.



Figure 11.



Figure 12.


Growth and ethanol production by S.
cerevisiae KD2 during a typical batch
fermentation in YEPD medium containing 20%
glucose.......................................

Growth rate of S. cerevisiae KD2 in the
presence of ethanol..........................

Rate of fermentation in the presence of
ethanol .................................. ...

Determination of intracellular ethanol
concentration................................

Inhibition of fermentation rate of 12-h and
24-h cells by added ethanol..................

Dose-response of cell growth to added
magnesium....................................

Magnesium levels in broth and cells during
the course of batch fermentation.............

Effect of magnesium addition on cell growth
and fermentation..............................

Effect of added magnesium on the rate of
fermentation.................................

Effect of magnesium supplementation on the
inhibition of fermentation rate by added
ethanol ................................. ...

Effect of ethanol removal on the
fermentative activity of cells grown in YEPD
medium containing 0.5 mM MgSO4..............

Effects of ethanol exposure on the
fermentative activities of 12- and 24-h
cells.......................................









Figure 13.



Figure 14.



Figure 15.

Figure 16.


Figure 17.


Figure 18.


Figure 19.


Figure 20.


Changes in the levels of glycolytic and
alcohologenic enzymes during batch
fermentation with 20% glucose................

Changes in intracellular pH and membrane
energization during batch fermentation of
20% glucose..................................


97



101


Effects of added ethanol on A pH........... 104


A typical thin layer chromatogram of a
fermentation sample extract.................

Changes in [32P]-labelled cellular
metabolites during batch fermentation.......


120


126


Comparison of nucleotide levels found in the
cells with those found in the fermentation
broth ....................................... 131


Intracellular concentration of nicotinamide
nucleotides during batch fermentation.......

Intracellular concentration of adenine
nucleotides and energy charge during batch
fermentation................................


133



136


vii















Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy


THE DECLINING RATE OF ETHANOL PRODUCTION
DURING BATCH FERMENTATION BY SACCHAROMYCES CEREVISIAE

By

Kenneth Michael Dombek

August 1987

Chairman: Lonnie O. Ingram
Major Department: Microbiology and Cell Science

As Saccharomyces cerevisiae ferments 20% glucose to

ethanol in batch culture, the rate of this conversion

declines. Previous studies assumed that ethanol caused this

inhibition because added ethanol repressed both yeast growth

and alcohol production. However, produced ethanol appeared

to be a more potent inhibitor than added ethanol. Also,

removal of ethanol from fermenting yeast cells did not fully

alleviate this inhibition. The yeast cell envelope was

freely permeable to ethanol and, thus, the intracellular

accumulation of ethanol to levels high enough to inhibit

glycolytic enzymes appeared unlikely. A nutrient limitation

for magnesium in complex fermentation medium was identified

as being partially responsible for declining rates of

alcohol production. Supplementation of broth with magnesium

prevented much of the decline but did not entirely eliminate

viii









it. Since the levels of glycolytic and alcohologenic

enzymes remained high and internal pH was maintained near

neutrality, inactivation of enzymes or lowered levels of in

vivo activity due to acidification of the cell cytoplasm

appear unlikely. As yeast cells produced ethanol, they did

change physiologically, however, becoming more resistant to

inhibition of fermentation by added ethanol and to ethanol-

induced decreases in ApH. Initially, the intracellular

levels of phosphorylated glycolytic intermediates decreased

as fermentation rate was declining. These results suggested

that the rates of glucose uptake and/or phosphorylation were

slowing relative to carbon flux through the rest of the

pathway. Declining glycolytic intermediate levels probably

were not due to inhibition of glycolytic enzymes by

declining levels of nicotinamide nucleotides. Initially

during fermentation, ATP levels decreased by 60%, while AMP

increased by 900%. One possible explanation for the decline

in glycolytic intermediates and the corresponding decrease

in fermentation rate is that the increased level of AMP

inhibits glucose phosphorylation which may slow the rate of

glucose uptake. In this study, the roles of inhibition by

ethanol, nutrient limitation, and physiological changes in

decreasing the rate of fermentation have been defined and

characterized. Each of these factors appears to be

partially responsible for the decline in ethanol production

by S. cerevisiae during batch fermentation.















CHAPTER I
GENERAL INTRODUCTION


Even before 2000 BC, Saccharomyces cerevisiae was used

to ferment malted barley and wheat into a type of beer-bread

in Mesopotamia (Corran, 1975). The Babylonians and

Egyptians adapted this fermentation process to the

production of a high alcohol beer from many different sugar

sources. In Greece, grape juice was a popular sugar source

for fermentation to make wine. By the middle ages, wine and

ale were among the only beverages available that were not

contaminated with disease-causing agents. The ethanol

produced during their fermentation acted as a preservative.

Even after the advent of refrigeration and sterile packaging

procedures, liquors derived from yeast fermentation remained

as popular beverages.

Recently, uses of ethanol other than for beverages have

received much attention. Because of the growing concern

about protecting the environment from pollution and the

increasing dependence of the United States economy on

imported oil, ethanol has been adopted as both a gasoline

additive and an alternative energy source. Almost 95% of

the global ethanol produced is the product of sugar

fermented by S. cerevisiae.











Despite the obvious importance of ethanol production by

S. cerevisiae, the physiological constraints which limit the

rate of ethanol production are not fully understood.

Identification of these constraints represents an important

step toward the development of improved organisms and

process conditions for more rapid ethanol production. Such

improvements could increase the ethanol-production capacity

of existing fermentation plants and reduce the cost of

future facilities.

Some of the initial research on alcoholic fermentation

took place in the 1830s. Independently, Cagniard-Latour,

Schawn and Kutzing described the microscopic structure of

yeast cells and initiated studies on the role of yeast in

fermentation (Schlenk, 1985). Twenty years later, Pasteur

expanded these findings and performed the first decisive

experiments showing that yeast cells were the living

entities responsible for fermenting sugar to ethanol

(Pasteur, 1860). He then went on to describe the effect of

oxygen on yeast fermentation, an effect which has since

become known as the "Pasteur effect" (Pasteur, 1861).

Biochemistry had its beginnings with the work of Buchner who

showed that the proteinaceous material in yeast juice was

capable of converting sugar to ethanol (Buchner, 1897). In

the decades that followed this discovery, the chemical

nature of alcoholic fermentation and the enzymes responsible











for the conversion which Pasteur had described 50 years

earlier were characterized (Fraenkel, 1982).

Even before these early studies on alcoholic

fermentation, it had been observed that yeast stopped

growing and fermenting before all of the sugar in the

fermentation broth had been utilized. In one of the

earliest investigations of this phenomenon, Brown examined

the influence of various environmental conditions on the

rate of growth of S. cerevisiae (Brown, 1905). The addition

of ethanol to growth medium, indeed, did inhibit yeast

reproduction. However, inhibition occurred only at a much

higher ethanol concentration than was observed to have

accumulated at the point during the fermentation when yeast

growth had ceased. This was the first indication that the

presence of ethanol may not be the only factor involved in

the premature termination of carbohydrate fermentation by

yeast.

The studies of Brown were complemented by the work of

Richards (1928) who showed that removing ethanol produced

during growth and maintaining a constant nutrient supply

allowed yeast cell multiplication to continue almost

indefinitely. Recently, the vacuum fermentation experiments

conducted by Cysewski and Wilke (1977) and Maiorella et al.

(1983) have corroborated and extended this finding. Boiling

off the ethanol as it was produced from a continuous culture

under reduced pressure, increased achievable cell densities











and fermentation rates. Because such investigations have

shown that ethanol does inhibit yeast growth and

fermentation, many studies have dealt with characterizing

these inhibitory effects (Brown et al., 1981; Hoppe and

Hansford, 1982; Jones and Greenfield, 1985; Lafon-Lafourcade

and Ribereau-Gayon, 1984; Vega et al., 1987).

The inhibitory effect of ethanol on the rate of sugar

conversion to ethanol was first quantitated by Rahn (1929).

He demonstrated an inverse relationship between the amount

of fermentation product, including ethanol, added to the

medium and the rate of its production by determining the

amount of heat evolved when sucrose was converted to

ethanol. Also, as ethanol accumulated in the medium, the

rate of fermentation declined. This represents some of the

original evidence that ethanol inhibits alcohol production

by yeast.

Recent studies also have attempted to quantitatively

describe the inhibitory effect of ethanol on product

formation during yeast fermentation (Luong, 1985; van Uden,

1985). Holtzberg et al. (1967) examined grape juice

fermentation by S. cerevisiae var. elipsoideus and

calculated an inverse linear relationship between the rate

of alcohol production and the amount of ethanol in the

juice. Similar findings were made by Ghose and Tyagi (1979)

for the batch fermentation of cellulose hydrolysate by S.

cerevisiae NRL y-132, however, the constants in the equation











relating the rate of alcohol production to the amount of

ethanol in the fermentation broth were slightly different.

A glucose-limited continuous culture of a respiratory-

deficient baker's yeast was found by Aiba et al. (1968) to

exhibit an exponential decrease in rate of alcohol formation

as increasing concentrations of ethanol were added to the

fermentation broth. They also demonstrated that ethanol

acted as a non-competitive inhibitor of alcohol formation.

Both batch and continuous culture fermentations of S.

cerevisiae ATCC No. 4126 in a synthetic medium were shown by

Bazua and Wilke (1977) to exhibit kinetics of ethanol

inhibition entirely different than had been described

previously. In each case, similar models were constructed

for growth in the presence of ethanol. The large variety of

kinetic models suggests that many factors, such as strain

variations, environmental conditions and nutritional state,

also may have important roles in the inhibition of growth

and alcohol production during yeast fermentation.

Strains of yeast able to grow and ferment in the

presence of higher concentrations of ethanol may be capable

of producing larger amounts of alcohol at faster rates.

Thus, many investigations have examined the mechanism of

ethanol tolerance in yeast (Casey and Ingledew, 1986; Ingram

and Buttke, 1984; Ingram et al., 1986; Moulin et al., 1984).

Initial studies on the alcohol tolerance of yeast were

performed by Gray (1941). Various ethanol producing species











of yeast, including several different strains of S.

cerevisiae, were grouped according to their ability to

utilize glucose in the presence of a series of ethanol

concentrations. The ethanol tolerance trait was not

characteristic of any specific genus or species since

different strains of the same species varied in their

tolerance. Yeast strains with differing ethanol tolerances

also had different cellular compositions (Gray, 1948).

Strains of lower alcohol tolerance contained higher amounts

of carbohydrate and lipid than did the more tolerant ones.

The studies of Troyer (1953) confirmed the results of

Gray and further examined the relationship between growth

and glucose utilization. Alcohol tolerant strains of yeast

exhibited increased growth in parallel with increased rates

of glucose utilization over less tolerant strains. During

yeast fermentation, the initial effect of ethanol added to

the medium was to decrease the total number of cells formed

followed by a corresponding decrease in glucose utilization.

The manner in which ethanol inhibits glucose

utilization by S. cerevisiae was examined by Gray and Sova

(1956). The ability of ethanol to inhibit glucose

utilization was not a specific property of this fermentation

product but rather a property shared by a class of

substances, short chain aliphatic alcohols. The potency of

normal alcohols as inhibitors of glucose utilization was

related to their chain length and primary alcohols were











found to be more inhibitory than secondary or tertiary

alcohols. This suggested that the mode of action of ethanol

may involve a hydrophobic site. Numerous investigations

have described the detrimental effects of a variety of

hydrophobic compounds, including alcohols, on the function

of many different types of cells and membranes (Barondes et

al., 1979; Eaton et al., 1982; Hayashida and Ohta, 1978;

Leao and van Uden, 1982b; Lenaz et al., 1978; Seeman, 1972).

The inhibitory potential of an alcohol on D-xylose transport

of S. cerevisiae was correlated directly with the lipid-

buffer partition coefficient of that alcohol by Leao and van

Uden. Hayashida and Ohta reported that ethanol promoted

leakage of ultraviolet-absorbing material from yeast cells,

an ethanol induced compromise of membrane barrier function.

Since ethanol alters the physical state of both artificial

and biological membranes (Chin and Goldstein, 1977; Dombek

and Ingram, 1984; Janoff and Miller, 1982; Rowe, 1983;

Vanderkooi et al., 1977), one site of ethanol action may be

the cell cytoplasmic membrane.

The cytoplasmic membrane of S. cerevisiae contains

about 50% protein and 40% lipid by dry weight (Hunter and

Rose, 1971). It has a unique lipid composition, containing

phospholipids with no polyunsaturated fatty acids like those

found in prokaryotes and large proportions of

phosphatidylcholine and sterols like those found in

eukaryotic cells (Ingram and Buttke, 1984). As much as 80%











of the fatty-acyl residues are unsaturated and as much as 6%

of the membrane dry weight consists of sterols, mainly

ergosterol (Hunter and Rose, 1971). The synthesis of

unsaturated fatty acids from saturated acids requires an

oxygen-dependent desaturase enzyme and the synthesis of

ergosterol from squalene requires oxygen-dependent

peroxidation and demethylation reactions (Henry, 1982).

When cultured under anaerobic conditions, S. cerevisiae

exhibits a requirement for both unsaturated fatty acids and

ergosterol in its nutrient supply for growth (Andreasen and

Stier, 1953; Andreasen and Stier, 1954).

Taking advantage of this anaerobically induced

nutritional requirement, the laboratory group of Rose

selectively enriched plasma membranes of S. cerevisiae by up

to 60% with an individual unsaturated fatty-acid or by up to

70% with a particular sterol (Hossack and Rose, 1976).

Using yeast with altered plasma membrane lipid composition,

they showed that cells enriched in ergosterol and linoleyl

residues remained viable for a longer period of time than

cells enriched in other sterols and oleyl residues when

exposed to ethanol (Thomas et al., 1978). Ethanol was also

less inhibitory to growth and solute accumulation when

plasma membrane lipids were enriched similarly (Thomas and

Rose, 1979). Because unsaturated fatty acids and

ergosterol appear to protect yeast cells from the inhibitory

effects of ethanol, it is not surprising that S. cerevisiae











has been shown to alter its plasma membrane lipid

composition when grown in the presence of ethanol (Beaven et

al., 1982). The amount of unsaturated fatty-acyl chains in

the phospholipids rose with increasing amounts of added

ethanol. The proportion of oleyl residues increased by 100%

with a corresponding decrease in the proportion of palmitic

residues in the presence of 1.5 M ethanol. In light of the

previous studies on yeast cells containing plasma membranes

enriched in unsaturated fatty-acyl residues, this change in

lipid composition may be an adaptive response to growth in

the presence of ethanol. As ethanol accumulated during

normal growth, however, the lipid fatty-acyl composition

became more saturated, probably the result of decreasing

oxygen tension in the medium.

Since enrichment of the plasma membrane with

unsaturated fatty-acyl residues and ergosterol protects

yeast cells from ethanol inhibition and the plasma membrane

fatty-acyl composition becomes more saturated during growth,

addition of unsaturated lipid supplements to the

fermentation broth might be expected to enhance the

fermentation rate and final ethanol yield from fermentable

substrates. Studies of sake fermentation by Hayashida have

shown that addition of proteolipid containing linoleyl

fatty-acyl residues to synthetic medium promotes the

formation of over 20% ethanol (Hayashida et al., 1974).

Sake yeast normally only produce this high amount of ethanol











under the very specialized conditions of the sake

fermentation. Koji mold proteolipid, found in sake mash,

also enhanced yeast growth, survival and fermentative

activity (Hayashida et al., 1975). This proteolipid,

isolated from Aspergillus oryzae, was shown to contain a

high percentage of phosphatidylcholine with linoleic acid

comprising the major portion of its fatty-acyl residues

(Hayashida et al., 1976). Supplementation with proteolipid

actually increased the proportion of phosphatidylcholine and

linoleyl fatty-acyl residues in the sake yeast plasma

membrane (Hayashida and Ohta, 1978). The

phosphatidylcholine promoted yeast growth and fermentative

activity, while addition of ergosterol-oleate increased

survivability in the presence of ethanol (Hayashida and

Ohta, 1980). Many confirming studies have shown that

addition of unsaturated lipids improves the fermentative

productivity of yeast (Damiano and Wang, 1985; Janssens et

al., 1983; Lafon-Lafourcade et al., 1979; Ohta and

Hayashida, 1983; Watson, 1982).

Although much is known about the effects of ethanol on

various aspects of yeast fermentation and about the

involvement of the plasma membrane in mediating many of

these effects, very little is known about the factors which

cause the rate of fermentation to decline as ethanol

accumulates in the medium. The following studies have

examined this phenomenon in more detail to determine the









11

role of ethanol produced during fermentation in causing this

decline in ethanol production rate. Other possible causes

of the decreasing rate of ethanol production during batch

fermentation, such as nutrient limitation, also were

studied. Finally, the physiological changes which accompany

the declining fermentation rate were characterized in order

to better understand the constraints that limit the rate at

which S. cerevisiae produces ethanol.















CHAPTER II
CHARACTERIZATION OF THE DECLINING RATES OF GROWTH AND
ETHANOL PRODUCTION DURING BATCH FERMENTATION BY
SACCHAROMYCES CEREVISIAE KD2


Introduction

Yeast metabolize sugar via Embden-Meyerhof glycolysis

to produce ethanol as the major reduced product of

fermentation. As ethanol accumulates in the fermentation

broth, both the rate of growth and alcohol production

declines (Ingram and Buttke, 1984; Luong, 1985; van Uden,

1985). The potency of ethanol as an inhibitor of yeast

growth and fermentation, however, differs in various species

(Gray, 1941). Not all strains of Saccharomyces cerevisiae

have equal abilities to grow and ferment in the presence of

added ethanol.

Environmental factors, such as temperature, also have a

key role in determining the potency of ethanol as an

inhibitor (Casey and Ingledew, 1986; Jones et al., 1981; van

Uden, 1985). Elevating the temperature reduced the maximum

ethanol yield of wine fermentations (Hohl and Cruess, 1936).

Both high and low temperatures decreased the ability of S.

cerevisiae to grow in the presence of ethanol (Loureiro and

van Uden, 1982; Sa-Correia and van Uden, 1983). Sa-Correia

and van Uden (1983) have shown that the temperature of











maximum ethanol tolerance for growth of S. cerevisiae is

between 280C and 300C. Similarly, the ability to survive in

the presence of ethanol decreased with increasing

temperature (Casey and Ingledew, 1986; Leao and van Uden,

1982a; Nagodawithana et al., 1974). Ethanol also enhanced

the thermal death rate of S. cerevisiae (van Uden and da

Cruz Duarte, 1981). In contrast, the rate of sugar

conversion became more resistant to ethanol inhibition as

the fermentation temperature was raised to 450C (Brown and

Oliver, 1982). Alcohol production proceeds at an

accelerated pace at the higher temperatures. When

optimizing the conditions for carrying out a fermentation

process, a compromise between these competing factors must

be reached.

Other environmental factors which affect the ability of

yeast to tolerate ethanol include the sugar and oxygen

concentrations in the fermentation broth (Jones et al.,

1981). Glucose concentrations above 14% decreased the

ability of S. cerevisiae to convert the sugar to ethanol

(Gray, 1945). This inhibition occurred as the cells began

to undergo plasmolysis and probably was caused by the

osmotic effects of these high amounts of glucose on the

yeast. Osmotic pressure also has an adverse effect on yeast

cell viability during fermentation (Panchal and Stewart,

1980). These effects of high sugar concentrations appear to

be synergistic with the inhibitory effects of ethanol











(Kunkee and Amerine, 1968; Moulin et al., 1980). A small

amount of oxygen increases the ethanol tolerance of S.

cerevisiae because it is required for the synthesis of

unsaturated lipids and ergosterol (Henry, 1982). These

lipids protect anaerobically grown yeast cells from ethanol-

induced growth inhibition (Ingram and Buttke, 1984). Under

continuous culture conditions, trace amounts of oxygen were

shown to decrease the ethanol inhibition of growth without

significantly affecting the ethanol yield per amount of

substrate consumed (Hoppe and Hansford, 1984).

Because strain differences and environmental factors

influence the ability of yeast to grow and ferment in the

presence of ethanol, it was necessary to characterize these

processes in the organism chosen for this study. This

organism was a genetically undefined petite brewery yeast,

S. cerevisiae KD2. Decreasing growth and alcohol production

rates as ethanol accumulated during batch fermentations were

characterized. Also, the effect of ethanol added to the

growth medium on the rates of growth and alcohol production

was examined.

Materials and Methods

Yeast Strains

The principal organism used in these studies was S.

cerevisiae KD2, a petite mutant of strain CC3 (G.G. Stewart,

Labatts Brewery, London, Canada). It was derived from the

parent strain by selection for the inability to form











colonies on lactate agar plates (Ogur and St. John, 1956)

after growth for 3 days in broth containing 6 mM MnCl2

(Putrament et al., 1973). It has been speculated that

manganese induces mutations by interacting with the

manganese-sensitive mitochondrial DNA polymerase causing

error-prone replication of the mitochondrial DNA. Strain

KD2 did not grow on glycerol containing medium or reduce

2,3,5-triphenyltetrazoleum chloride (Ogur et al., 1957). It

also lacked the cytochrome a+a3 absorbance bands at 600 nm

and 440 nm and the cytochrome b bands at 560 nm and 530 nm.

A petite strain was chosen for this study because growth and

fermentation have been shown to be almost identical both

anaerobically and aerobically, eliminating the need to

perform experiments under anaerobic conditions (Loureiro-

Dias and Arrabaca, 1982). In some studies, S.cerevisiae CC3

and S. cerevisiae A10 p (NRRL Y-12707) were used for

comparison. The latter strain was provided generously by

N.J. Alexander (Northern Regional Research Center, U.S.

Department of Agriculture, Peoria, Ill.).

Growth Conditions

All organisms were grown on YEPD medium which contained

5 g/liter yeast extract, 10 g/liter peptone and 200 g/liter

glucose as described by Leao and van Uden (1982a). The

medium was adjusted to pH 5.0 with 2.0 N HCl prior to

autoclaving. Solid medium for culture maintenance consisted

of YEPD broth containing 1.5% agar.











Batch fermentations were carried out in 250-mi tissue

culture spinner bottles (Bellco Glass, Inc., Vineland,

N.J.), immersed in a 300C water bath and agitated at

150 rpm. Culture bottles were fitted with water-trapped

exit ports for the escape of carbon dioxide and sampling

ports for the removal of culture by syringe. Growth was

allowed to proceed under conditions of self-induced

anaerobiosis. Inocula were prepared by transferring cells

from a slant to a test tube containing 10 ml of YEPD broth.

Cells were incubated at 30C for 36 h without agitation and

diluted 1:40 into 300 ml of fresh YEPD in a spinner bottle.

This culture was incubated for approximately 12 h until an

optical density at 550 nm of 3.5 (1.3 mg of cell protein per

ml) was reached. Fermentations were started by diluting the

12-h culture 1:100 into 300 ml of growth medium.

Preparation of Fermentation Samples for Analysis

Fermentation samples were centrifuged at 10,000 x g for

0.5 min. The supernatant was removed and saved by freezing

at -200C. Cells were washed once in 50 mM KH2PO4 buffer (pH

5.0), and the pellets were saved for further analysis by

freezing at -200C.

Analytical Methods

Ethanol was measured by gas-liquid chromatography as

described by Goel and Pamment (1984) with 2% (vol/vol)

acetone as an internal standard. Glucose was initially

determined with the glucose oxidase procedure (Raabo and











Terkildsen, 1960) using the Glucostat reagents supplied by

the Sigma Chemical Company (St. Louis, Mo.). In later

experiments, glucose was measured with a YSI model 27

glucose analyzer (YSI, Yellow Springs, Oh.) Cell mass was

measured as optical density at 550 nm with a Bausch and Lomb

Spectronic 70 spectrophotometer and as total cell protein by

the method of Lowry et al. (1951) as described by Layne

(1957).

Respirometry Measurements

Samples were pipetted into Warburg flasks and

equilibrated for 10 min at 300C. During the first 5 min of

the equilibration period, the flasks were flushed with

nitrogen gas. Rates of CO, production were measured with a

differential respirometer (Gilson, Middleton, Wis.). These

values were used to calculate fermentation rates as pmoles

of CO2 evolved per mg of cell protein. The rate of CO,

evolution was independent of sample volume, up to 4 ml, and

linearly increased with cell concentration, up to 5 mg cell

protein per ml.

Chemicals

Yeast extract, peptone and agar were obtained from

Difco Laboratories, Detroit, Mich. Glucose and other

biochemicals were obtained from Sigma Chemical Co. Acetone

and inorganic salts were purchased from Fisher Scientific

Company, Orlando, Fla. Absolute ethanol was supplied by

AAPER Alcohol and Chemical Co., Shelbyville, Ky. Gas











chromatography supplies were obtained from Supelco,

Bellefonte, Pa.

Results

Batch Fermentation by S. cerevisiae KD2

A typical batch fermentation profile of S. cerevisiae

KD2 in YEPD medium is shown in figure 1. Glucose conversion

essentially was completed after 60 h under these conditions

with the production of between 12 and 13% (vol/vol) ethanol.

Cell protein stopped increasing after 24 h at 2.4 mg per ml

medium, although the optical density at 550 nm of this

culture continued to rise for an additional 12 h period

(data not shown). Nearly identical profiles were obtained

in medium supplemented with Tween 80 (5 g/liter), linoleate

(45 mg/liter) and ergosterol (30 mg/liter). Similar

profiles also were obtained by the addition of small amounts

of 10 N KOH during the course of fermentation using a pH

stat to maintain the pH of the growth medium at 5.0.

Likewise, batch fermentations of S. cerevisiae CC3, the

parental grande strain, were indistinguishable from those of

strain KD2.

Inhibition of Growth Rate by Ethanol

Although it is not obvious from figure 1, the growth

rate of S. cerevisiae KD2 decreases as ethanol accumulates

in the fermentation broth (Fig. 2). Using the data from

batch fermentations, rates of growth were calculated as the

increase in cell protein over a 1.6 h period at the various































Figure 1. Growth and ethanol production by S.
cerevisiae KD2 during a typical batch
fermentation in YEPD medium containing
20% glucose. Symbols: O cell
protein (mg/ml culture); ,
glucose; Q ethanol.
























I0.0

5.0


I a I I I I 1


0 10 20 30
TIME


40
(h)


50 60


I0.0

5.0
0


I-
r

1.0
0
-4
0.5 m
z

o.
3

3
O. I"

0.05


- 0.01
70


1.0

J 0.5
U)
0
0
-J

* 0.I

0.05


0.01


























Figure 2.


Growth rate of S. cerevisiae KD2 in
the presence of ethanol. Rates were
measured from batch culture
experiments as the increase in cell
protein during a 1.6 h interval around
the time point sampled. These are
plotted as a function of ethanol
accumulated in the medium during
fermentation. Growth rates in YEPD
medium containing different
concentrations of ethanol were
calculated as the exponential increase
in cell mass per h as measured by
optical density at 550 nm. These are
plotted as a function of ethanol added
to the medium. Symbols: 0, growth
rate during batch fermentation; U ,
effect of added ethanol on growth
rate.


















































ETHANOL (% V/V)


0.40


0.30 1


0.201


0. 10


0.00


I I 2 I I I I 7
0 I 2 3 4 5 6 7











times sampled during the fermentations. Growth rate

decreased exponentially as ethanol accumulated in the

medium. The concentration of ethanol that had been produced

at half the maximum observed growth rate was

1.08% (vol/vol).

The addition of ethanol to fermentation broth also

decreased growth rate (Fig. 2). Cells inoculated into broth

containing increasing concentrations of ethanol were

inhibited in a dose-dependent manner. This inhibition was

linear above concentrations of 2% (vol/vol). The amount of

ethanol required to decrease the growth rate by 50% was

4.66% (vol/vol). Thus, four times more ethanol was required

to decrease growth rate by one-half than was produced when

growth rate had declined by this same fraction.

Inhibition of Fermentation Rate by Ethanol

The rate of alcohol production per mg cell protein was

calculated in a fashion analogous to the growth rate data.

These fermentation rates are shown as a function of average

ethanol accumulated in the medium in figure 3. Fermentation

rates also were determined by manometry using samples from

batch fermentations with excellent agreement for samples

taken 12 h and later. Identical plots were obtained with S.

cerevisiae CC3, the grande parent strain of KD2. The trends

observed were similar for cells grown with and without lipid

supplements and for cells grown in a pH stat where the pH of

the medium was held at 5.0. The fermentative activity of






















Figure 3. Rate of fermentation in the presence
of ethanol. Fermentative activity was
determined from batch culture
experiments as the increase in ethanol
concentration over 1.6-h time
intervals divided by the average
cellular protein concentration in the
medium during that time interval. It
is plotted as a function of ethanol
accumulated in the growth medium and
is expressed as umoles ethanol
produced per h per mg cell protein.
The effect of added ethanol on the
activity of cells at their highest
measured rate of fermentation, 12 h
after inoculation, is included for
comparison. Ethanol was added
directly to fermentation samples and
fermentation rates were measured by
respirometry. These are expressed as
umoles CO, produced per h per mg cell
protein and are plotted as a function
of total ethanol in the medium.
Symbols: *, fermentation rate
during batch fermentation; ,
effect of added ethanol on the
fermentation rate of 12-h cells.

































> 40-
I-\


< 30

> *
I-
S20
w


10 -



0
0 2 4 6 8 10 12
ETHANOL (% V/V)











cells exhibited a biphasic decline as a function of

accumulated ethanol. An initial decline in fermentation

rate occurred during the accumulation of 3.7% (vol/vol)

ethanol with a 50% loss of activity. This was followed by a

more gradual decline in fermentation rate with approximately

20% of the original activity remaining after the production

12% (vol/vol) ethanol. The fermentation rate of S.

cerevisiae AI0 po, a respiratory-deficient haploid

laboratory yeast strain, also declined as ethanol

accumulated in the medium (data not shown). As with S.

cerevisiae KD2, a 50% decrease in fermentation rate was

observed after the accumulation of 3.5% (vol/vol) ethanol.

However, the maximum rate of fermentation was lower,

40 pmoles CO2 per h per mg protein compared to 50 for strain

KD2 and greater than 90% of the maximum fermentation rate

was lost by the time 6.5% (vol/vol) ethanol had accumulated.

Unlike ethanol accumulated during fermentation, the

addition of low concentrations of ethanol to rapidly

fermenting cells 12 h after inoculation did not result in a

large decline in fermentative activity (Fig. 3). Ethanol

caused a dose-dependent linear decline in activity.

Fermentation was inhibited only 12% by the addition of 3.7%

(vol/vol) ethanol and 8.5% (vol/vol) added ethanol was

required to cause 50% inhibition.











Discussion

Because the ability of ethanol to inhibit alcohol

production varies with the yeast strain and fermentation

conditions employed (Jones et al., 1981), the effect of

ethanol on these processes in the yeast strain chosen for

these studies of alcohol production was characterized. The

observed decline in both growth and fermentation rates was

similar under a variety of culture conditions for S.

cerevisiae KD2. This decline also was seen with other

strains of S. cerevisiae, but strain KD2 maintained equal or

greater rates and accumulated more ethanol during

fermentation. Identical fermentation profiles of strain KD2

and its grande parent strain suggest that the manganese

treatment used to obtain respiratory-deficient cells, in

itself, was not responsible for the observed decline in rate

of growth and alcohol production. Induction of respiratory-

deficiency has been observed to decrease ethanol tolerance

in some petite strains of yeast (Aguilera and Benitez, 1985;

Esser et al., 1982) while increasing the fermentation rate

and ethanol tolerance of other strains (Bacilia et al.,

1978; Moulin et al., 1981). Differences between wild-type

strains, the harshness of the mutagenic treatments and the

limited numbers of mutants screened may be responsible for

these contradictory reports.

The brewery yeast strain used in most of these studies

exhibited an exponential decrease in growth rate (Fig. 2)











and a biphasic decrease in rate of alcohol production

(Fig. 3) as ethanol accumulated in the fermentation broth.

The exponentially falling growth rate was similar to that

observed by Aiba et al. (1968). However, the biphasic

decrease in ethanol production fits both an exponential

model and a linear model describing a combination of two

events. In batch fermentations by strain KD2, one cellular

site may be much more sensitive to the accumulation of

ethanol than another. Thus, a biphasic inhibition profile

occurs with the more sensitive site being characterized by

the line with the steepest slope. Fermentation rate

declined by 50% after the accumulation of about three and

one-half times more ethanol than was accumulated when growth

rate decreased by a similar amount (Fig. 2 and Fig. 3).

Clearly, ethanol added to the fermentation broth

linearly decreased both the rate of cell growth and alcohol

production as previously described (Brown et al., 1981;

Moulin et al., 1984). This contrasts with the results of

Luong (1985) who reported that S. cerevisiae ATCC 4126

exhibited non-linear decreases in growth and fermentation

rates under anaerobic conditions. As with the accumulation

of ethanol, growth rate was inhibited more than fermentation

rate, but was only about two-fold more sensitive.

Growth rate decreased faster than fermentation rate as

ethanol accumulated in the medium and as increasing amounts

of ethanol were added exogenously to the medium. Thus, the











actual alcohol production machinery is more resistant to

ethanol inhibition than is cell growth (Brown et al., 1981;

Luong, 1985). The magnitude of the observed inhibition,

however, was greater for endogenously produced ethanol than

for exogenously added ethanol for both processes in

agreement with previous reports (Moulin et al., 1984; Novak

et al., 1981). These results suggest that the mere presence

of ethanol may not be entirely sufficient to account for the

observed decline in fermentation rates. In the following

studies, the causal role of ethanol in the decreasing

fermentation rate was chosen for more detailed examination.

The role of factors other than ethanol also will be studied

in order to account more fully for this observed decrease in

rate of ethanol production.















CHAPTER III
NUTRIENT LIMITATION AS A BASIS FOR THE APPARENT TOXICITY OF
LOW LEVELS OF ETHANOL DURING BATCH FERMENTATION


Introduction

As has already been described for Saccharomyces

cerevisiae KD2, the rate of alcohol production per unit cell

mass decreases substantially during batch fermentations as

ethanol accumulates in the medium (Fig. 3). This decrease

has been attributed to the inhibitory effects of ethanol by

most researchers (Aiba et al., 1968; Bazua and Wilke, 1977;

Ghose and Tyagi, 1979; Luong, 1985; Millar et al., 1982;

Moulin et al., 1984; Rahn, 1929). However, recent studies

by Casey et al. (1983, 1984) have provided evidence that

nutrient limitation, in addition to ethanol accumulation, is

also an important factor limiting the rate of fermentation

during high-gravity brewing. The following studies examine

the role of ethanol in limiting the rate of alcohol

production and provide evidence that nutrient limitation is

an additional factor which contributes to the initial

decline in fermentative activity.

Materials and Methods

Organism and Growth Conditions

The organism used in these studies was Saccharomyces

cerevisiae KD2, a petite derivative of strain CC3 (G.G.

30











Stewart, Labatts Brewery, London, Canada). This organism

was grown in YEPD medium as described in chapter II.

Fermentations were carried out at 300C in spinner bottles

designed for tissue culture, also as described in

chapter II. "Conditioned broth" refers to medium in which

cells have been allowed to grow for 12 or 24 h and have been

removed by centrifugation. This broth was sterilized by

filtration.

Analytical Methods

Cell mass was monitored by measuring optical density at

550 nm using a Bausch and Lomb Spectronic 70

spectrophotometer. Total cell protein was determined using

the method of Layne (1957). Cell viability was measured

with the methylene blue dye exclusion procedure of Trevors

et al. (1983). Ethanol was determined using gas

chromatography as described by Goel and Pamment (1984).

Rates of fermentation were measured as the rate of CO2

production at 300C under a nitrogen atmosphere using

respirometry as described in chapter II.

Measurement of the Intracellular Ethanol Concentration

The procedure used to determine the intracellular

concentration of ethanol in actively fermenting yeast cells

is illustrated in figure 4. Cells from batch fermentations

were concentrated by centrifugation (10,000 x g, 2 min,

ambient temperature) and resuspended in the same medium to a

density of 50 mg of cell protein per ml. [14C]sorbitol












a)


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(specific activity, 50 pCi/mmole) was added to a 1-ml

suspension at a final activity of 42 nCi/ml. The suspension

was mixed for 10 sec using a vortex mixer, and 0.1 ml

samples were transferred to Whatman 3MM filter paper disks

(3 cm) for sorbitol measurements and to sample vials

containing 0.1 ml perchloric acid for subsequent alcohol

determinations. The remaining suspension was centrifuged

immediately at 10,000 x g for 30 sec in a microcentrifuge.

Supernatant samples of 0.1 ml then were transferred to 3MM

filter paper disks and to sample vials containing 0.1 ml of

0.58 M perchloric acid for the measurement of ethanol.

Filter disks were air dried at 800C before the addition of

scintillation fluid. The radioactivity of these samples was

measured using a Beckman model 8000 scintillation

spectrometer. Total cell volume was estimated as the

difference in ["4C]sorbitol counts between the suspension

and the supernatant.

A correction was made for the volume of total cell

solids included in the sorbitol-based estimate of cell

volume (Fig. 4b). This was done in a separate experiment to

ensure sufficient time for equilibration of tritiated water.

Control experiments were performed to confirm that tritiated

water had reached equilibrium after 5 min and that the

sorbitol did not leak into the cells during this period.

Tritiated water and [1C]sorbitol (specific activity,

50 pCi/mmole) were added to concentrated cell suspensions











(50 to 100 mg of cell protein per ml) at a final activity of

2 pCi/ml and 84 nCi/ml, respectively. After 5 min, 0.1-ml

samples were pipetted directly into scintillation fluid for

aqueous samples. The cell solid volume was calculated as

the difference between the tritiated water counts in the

suspension and the supernatant samples. The fraction of the

total cell volume occupied by solids was computed as

follows:


1 (3Hsu8/SHup)
V. = (1)
1 (14C-u.14CB U)


where V, is the fraction of solid volume, sus is the

suspension and sup is the supernatant fraction. Typically,

the cell solid volume represented 20 to 25% of the sorbitol-

excluded volume. The intracellular water content decreased

from 2.23 ul per mg cell protein 12 h after inoculation to

0.83 pl per mg cell protein by 48 h.

The intracellular concentration of ethanol was computed

based on the aqueous cell volume, i.e., the sorbitol-

excluded volume minus the solid volume. This was calculated

by assuming that the amount of ethanol in the suspension is

equal to the intracellular concentration of ethanol times

the aqueous cell volume plus the concentration of ethanol in












the supernatant times the supernatant volume as follows:

14C 14Cu
8US BUS
EsuS = Ecell (1 ) (1 V) + Eup (---- (2)
14CS14U
1 sup 1 sup


where Eus is the ethanol concentration in the suspension,

Eup is the ethanol concentration in the supernatant and

Ecell is the ethanol concentration within the aqueous cell

volume.

Chemicals

Complex medium components and agar were purchased from

Difco Laboratories, Detroit, Mich. Glucose and other

biochemicals were obtained from Sigma Chemical Co., St.

Louis, Mo. Inorganic salts were purchased from Fisher

Scientific Company, Orlando, Fla. Absolute ethanol was

supplied by AAPER Alcohol and Chemical Co., Shelbyville, Ky.

Radioactive compounds were purchased from New England

Nuclear, Boston, Mass. Gas chromatography supplies were

obtained from Supelco, Bellefonte, Pa.

Results

Effect of Ethanol Removal on Fermentation Rate

These studies have focused on two time points during

batch fermentation, 12-h and 24-h, to investigate the

possible reasons for the initial drop in fermentative

activity. To minimize possible variability arising from

inoculum differences, autoclaving, etc., 12-h cells have

been operationally defined as those which have increased in











cell mass 100-fold after inoculation. Typically, these

samples contain 1.2 to 1.3% (vol/vol) ethanol and 1.3 mg

cell protein per ml culture medium. Cells which have

produced 5.0 to 5.6% (vol/vol) ethanol, in addition to any

ethanol that may have been present in the original medium,

were operationally defined as 24-h cells. Typically, these

samples contained 2.6 mg cell protein per ml culture medium.

Table 1 shows the effects of ethanol removal on the

fermentation rates of 12-h and 24-h cells. This activity of

12-h cells was much higher than that of 24-h cells. Ethanol

removal by suspension in fresh broth had little effect on

the activity of 12-h cells and did not result in a

significant increase in the fermentation rate of 24-h cells.

Similarly, suspension in conditioned broth from 12-h

fermentations, containing 1.1% (vol/vol) ethanol, did not

affect fermentation rate. Suspension of cells in the 24-h

conditioned broth, containing 5.6% (vol/vol) ethanol,

reduced the fermentative activity of 12-h cells but had less

effect on the activity of 24-h cells. Removal of volatile

medium components from the 24-h conditioned broth eliminated

its inhibitory effect on the fermentation rate of 12-h cells

but did not result in a significant increase in activity of

the 24-h cells. The addition of ethanol to the 24-h

conditioned broth restored its ability to repress the

fermentation rate of 12-h cells, indicating that ethanol was
















Table 1. Effects of ethanol and fermentation medium
composition on fermentation rate


Rate of Fermentationa
(pmoles CO2 per h per mg protein (SD))
Assay medium

12-h cells 24-h cells


Original broth 36.3 (2.4) 16.5 (2.6)

Fresh broth 39.5 (2.3) 20.3 (2.5)

Conditioned broth 38.9 (1.9) 22.7 (5.7)
(12-h, 1.1% (vol/vol)
ethanol)

Conditioned broth 22.8 (0.6) 16.1 (2.0)
(24-h, 5.6% (vol/vol)
ethanol)

Conditioned broth 34.9 (0.8) 17.6 (2.0)
(24-h, volatiles removed
under vacuum)

Conditioned broth 21.9 (0.2) 15.7 (1.0)
(24-h, volatiles removed
under vacuum,
reconstituted to give
5.6% (vol/vol) ethanol)


a Cells from 12-h and 24-h batch fermentations were
harvested by centrifugation at ambient temperature and
suspended to their original volume in various broths. Where
indicated, volatiles were removed from conditioned broth by
vacuum distillation at 550C, reducing the volume by two-
thirds. The broth then was reconstituted with distilled
water or distilled water plus ethanol. Fermentation rates
were measured by respirometry. Averages and standard
deviations (SD) represent the results from three separate
batch fermentations.











the principle volatile component responsible for this

inhibition.

Cell Viability and Overcrowding Effects on Fermentation Rate

A trivial possibility for the failure of 24-h cells to

recover activity after suspension in broth lacking ethanol

would be the presence of large numbers of dead cells.

However, based on methylene blue dye exclusion, over 90% of

the yeast cells appeared active and intact at 24 h. Another

trivial possibility for the failure of 24-h cells to recover

activity after suspension in fresh medium is that by 24 h,

the cells are so crowded that they can no longer efficiently

take up nutrients and glucose for conversion to ethanol.

This possibility was addressed by suspending 24-h cells in

fresh medium at different cell concentrations and measuring

the rate of CO, production by respirometry. The rate of CO,

production increased linearly with increasing cell

concentrations. At a cell concentration comparable to that

of 12-h cells, the fermentation rate of 24-h cells was only

half that of 12-h cells (data not shown).

Intracellular Ethanol Concentration

The failure of 24-h cells to recover activity after

suspension in fresh medium could be caused by the failure of

the suspension procedure to effectively remove the

intracellular ethanol or by the accumulation of large

amounts of intracellular ethanol that could permanently

damage the fermentative capacity of the cells. To explore











these possibilities, the intracellular and extracellular

concentrations of ethanol were measured at 12 h and 24 h

during batch fermentations (Table 2). The external ethanol

concentrations in these suspensions at the time of sampling

were 1.2% (vol/vol) and 5.0% (vol/vol), respectively, before

concentrating the cells. The level of extracellular ethanol

measured in the concentrated cell suspension was slightly

higher than the starting culture reflecting the rapid

metabolism of cells during the less than 3 min period of

cell concentration and sampling. In all cases, the

calculated intracellular ethanol concentration was lower

than or equivalent to the extracellular ethanol

concentration.

To confirm that the higher amounts of ethanol in the

concentrated cell suspension resulted from rapid metabolism,

a potent inhibitor of enolase (Warburg and Christian, 1941)

and of fermentation, potassium fluoride, was added before

cell concentration. Previously, 50 mM potassium fluoride

was determined to cause immediate cessation of CO, evolution

(data not shown). In both 12-h and 24-h fermentation

samples, the addition of fluoride prevented the increase in

extracellular ethanol during cell concentration and sampling

(Table 2).

The removal of ethanol by suspension of cell pellets in

fresh medium lacking ethanol substantially decreased the

intracellular ethanol concentration (Table 2). Regardless






















Table 2. Intracellular and extracellular ethanol
concentrations under various conditions


Ethanol concentration (% vol/vol) (SD) in
different media


Fresh
Sample Native Native Fresh + 10%
+ KF ethanol


12b
Int 1.9 (0.4) 1.4 (0.2) 0.6 (0.1) 6.9 (1.5)

Ext 1.7 (0.1) 1.3 (0.1) 0.4 (0.1) 10.7 (1.1)

24b
Int 3.4 (0.7) 3.7 (0.7) 0.7 (0.1) 8.1 (1.3)

Ext 5.5 (0.1) 5.1 (0.1) 0.5 (0.1) 9.7 (0.1)


a Three or more independent determinations. Native refers
to the broth in the batch fermentation with or without added
KF (50 mM). Fresh refers to sterile, unused medium with or
without added ethanol (10% (vol/vol)).

b Age of batch fermentation. Int and Ext refer to the
intracellular and extracellular ethanol concentrations,
respectively.











of the cell age and original ethanol concentration, the

intracellular ethanol concentration was found to be 0.6 to

0.7% (vol/vol) after ethanol removal. These values were

somewhat higher than anticipated and appeared to be due to

ethanol production by continued metabolism during suspension

and sampling. The inclusion of potassium fluoride during

harvesting and suspension in fresh medium resulted in a very

low internal and external ethanol concentration (0.06%

(vol/vol)), consistent with dilution of the cell pellet

volume with fresh medium.

In an analogous fashion, the failure of exogenously

supplied ethanol to raise the internal ethanol concentration

of 24-h cells to a level equivalent with that of cells

during fermentative alcohol production could provide an

explanation for the apparent resistance of 24-h cells to the

inhibitory effects of added ethanol (Table 1). Samples

taken after 48 h and processed to determine the

intracellular ethanol concentration contained approximately

11.2% (vol/vol) (SD 1.0) ethanol in the fermentation broth.

The intracellular ethanol concentration of these cells was

8.2% (vol/vol) (SD 1.7). Suspension of 24-h cells in broth

containing 10% (vol/vol) ethanol resulted in an increase in

the intracellular ethanol concentration to 8.1% (vol/vol)

(SD 1.3). These values indicate that the addition of

ethanol to 24-h cells increased the intracellular ethanol

concentration to the level found in cells during batch











fermentation. Thus, on the time scale of the ethanol

removal experiments, 24-h cells appear to be freely

permeable to ethanol added to the fermentation broth.

Effect of Added Ethanol on the Fermentation Rate of 12-h and
24-h Cells

The sensitivity of 12-h and 24-h cells to inhibition of

fermentative activity by added ethanol is illustrated in

figure 5. The fermentation rate of 24-h cells was

approximately one-half that of 12-h cells when assayed in

fresh broth lacking ethanol. Both types of cells were

insensitive to ethanol concentrations up to 2% (vol/vol)

after which they exhibited a dose-dependent linear decline

in activity up to between 12 and 14% (vol/vol) ethanol.

When plotted as a percentage of maximal rate, 24-h cells

appeared slightly more resistant, 50% inhibition at 8.3%

(vol/vol) ethanol as compared with 7.4% for 12-h cells.

Effect of Medium Composition During Growth on the
Fermentation Rate of 12-h and 24-h Cells

The slight differences in sensitivity to inhibition by

ethanol and the failure of ethanol removal to increase

fermentation rates suggest that the reduced activity of 24-h

cells may be primarily due to physiological changes in the

cells rather than to the immediate presence of ethanol.

Several experiments were performed to identify possible

causes of the physiological changes which may be involved

(Table 3). In these experiments, cells were grown under a

variety of conditions, harvested by centrifugation at











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ambient temperature and suspended in fresh medium lacking

ethanol to measure the rate of fermentation under standard

conditions. In all experiments, the fermentation rate of

cells used as inoculum to start these batch fermentations

were included as controls.

Experiment 1 examined the possibility that the

physiological changes in 12-h cells to produce 24-h cells

were due to growth in the presence of ethanol. Batch

fermentations in which 5% (vol/vol) ethanol was added prior

to inoculation were allowed to grow to the same cell mass as

12-h control cells, 1.2 mg cell protein per ml fermentation

broth. The fermentation rate of these cells grown in the

presence of added ethanol was only slightly lower than that

of control cells grown for 12-h in the absence of ethanol.

This indicated that exposure to 5% (vol/vol) ethanol during

growth was not sufficient to account for most of the

observed reduction in fermentation rate.

The possibility that growth in the presence of ethanol

and other fermentation products may be responsible for the

reduction in fermentation rate was examined in experiment 2.

Cultures were inoculated into bottles containing filter-

sterilized conditioned broth which had been supplemented

with 5 g/L yeast extract and enough glucose to increase the

concentration in the broth back to 20%. Conditioned broth

from 12-h cultures contained 1.2% (vol/vol) ethanol and

broth from 24-h cultures contained 4.5% (vol/vol) ethanol.











These fermentations were allowed to proceed until the

culture cell density reached the state defined as 12-h

cells, approximately 1.3 mg protein per ml broth. Cells

grown in the supplemented 12-h conditioned broth fermented

at rates equal to those of control cells. The fermentation

rate of cells grown in the supplemented 24-h broth was

lower, but was at least twice that of the 24-h control.

After allowing these fermentations to continue until 5%

(vol/vol) ethanol had been produced in addition to that

present at the time of inoculation, the fermentation rate of

both types of "24-h" cells were similar to that of the

control cells. Thus, the decline in rate of fermentation

observed after the production of 5% (vol/vol) ethanol is not

due entirely to the accumulation of ethanol and/or other

stable inhibitors in the fermentation broth.

The last possibility examined in order to understand

the reasons for the decline in fermentation rate of cells

after 24 h was the effect of nutrient limitation. Neither

12-h conditioned broth nor 24-h conditioned broth

supplemented with glucose supported vigorous growth of

strain KD2 following reinoculation (Table 3, experiment 3).

In experiment 2, the addition of yeast extract restored the

ability of conditioned broth to support growth, promoting

fermentation rates equivalent to the control. Cells grown

in broth containing 25 g/L yeast extract, 5-fold greater

than that of control broth, exhibited fermentation rates











equivalent to control cells after 12 h. These were twice as

high as control cells after the production of 5% (vol/vol)

ethanol in approximately 24 h (Table 3, experiment 4).

Discussion

Previous studies have shown that the rate of alcohol

production by yeast per unit cell mass decreases as ethanol

accumulates during fermentation (Holtzberg et al., 1967;

Navarro and Durand, 1978; Strehaiano and Goma, 1983). Most

of these studies have attributed this reduction in

fermentation rate to adverse effects of ethanol (Ingram and

Buttke, 1984; Maiorella et al., 1983; Millar et al., 1982;

Moulin et al., 1984). In fact, the possible accumulation of

high intracellular concentrations of ethanol in S.

cerevisiae and its involvement in the inhibition of growth

and fermentation have been the subject of considerable

controversy. Previous studies have shown that the growth

and fermentation rates of S. cerevisiae are much less

sensitive to inhibition by added ethanol than is inferred by

the decrease in alcohol production and growth rates which

accompany the accumulation of ethanol during fermentation

(Ingram and Buttke, 1984; Moulin et al., 1984; Nagodawithana

and Steinkraus, 1976; Navarro and Durand, 1978; Novak et

al., 1981). To explain this anomaly, it has been proposed

that the leakage of ethanol from yeast cells is, in some

way, limited by the permeability of the plasma membrane.

This would result in the accumulation of high cytosolic











levels of ethanol during rapid fermentation. Addition of

exogenous ethanol would not readily duplicate this

condition. However, the ethanol retention hypothesis is not

supported by direct measurements of intracellular ethanol

concentrations during fermentation.

A series of attempts to measure the intracellular

concentration of ethanol have resulted in conflicting data.

Problems in experimental design associated with the

measurement of a small, rapidly produced metabolite, such as

ethanol, contribute to these differences. The conflicting

reports result from two basic problems. First, measurements

of ethanol concentrations in the pellets of rapidly

fermenting cells result in calculated intracellular

concentrations of ethanol which are often several-fold

higher than those of the surrounding medium (Navarro and

Durand, 1978; Novak et al., 1981; Panchal and Stewart,

1980). This was, in large part, due to the continued

production of ethanol by the cells in pellets during

centrifugation and processing (Dasari et al., 1984). The

acuteness of this problem also would be expected to decrease

as fermentation rate and substrate levels declined during

batch fermentations. The reduction in apparent

intracellular/extracellular ratios of ethanol observed by

Beaven et al. (1982) during the latter stages of

fermentation supports this idea. Dasari et al. (1984)

demonstrated that precooling the culture significantly











reduced the error introduced by continued ethanol production

during cell harvesting. However, cooling may introduce

other potential problems associated with temperature-

induced changes in the organization of and permeability

properties of the plasma membrane.

The second type of experimental problem associated with

measurements of internal ethanol involves washing of the

yeast cells. Experimental designs which included washing of

cells (Nagodawithana and Steinkraus, 1976; Panchal and

Stewart, 1980) before estimation of ethanol resulted in

lower apparent intracellular ethanol concentrations than do

unwashed samples (Beaven et al., 1982; Dasari et al., 1984).

Beaven et al. (1982) clearly showed that even minimal

washing leaches most of the intracellular ethanol from the

cells. These two experimental designs, measurement of

ethanol in a metabolically active cell pellet and washing,

each introduce errors which change the calculated values of

intracellular ethanol in opposite ways.

Recent studies by Guijarro and Lagunas (1984) have

employed a procedure which eliminated these two basic

problems in experimental design by using glass fiber filters

to rapidly harvest cells. With this method, extracellularly

added ["4C]ethanol rapidly equilibrated with the

intracellular environment, indicating that the plasma

membrane is freely permeable to ethanol. However, this

still does not answer the question of the true intracellular











ethanol concentration in yeast cells during active

fermentation and ethanol production.

In the studies presented in this chapter, the

intracellular ethanol concentration was estimated by an

independent method using cells that were actively producing

ethanol in suspension culture. The results obtained using

this method confirm the reports by Beaven et al. (1982) and

Guijarro and Lagunas (1984) which indicated that yeast cells

are freely permeable to ethanol. In addition, these results

provide direct evidence that the intracellular concentration

of ethanol produced during fermentation is not several-fold

higher than that of the surrounding medium as proposed

previously (Beaven et al., 1982; Nagodawithana and

Steinkraus, 1976; Novak et al., 1981; Strehaiano and Goma,

1983). Identical conclusions were reached by Dasari et al.

(1985) using high cell density fermentations which allowed

rapid processing of the cells for analysis. There does not

appear to be any problem associated with the efficient

diffusion of ethanol from yeast cells into the environment

during fermentation. Thus, it is unlikely that the

retention of unusually high intracellular ethanol

concentrations contributes toward the decrease in

fermentative activity of S. cerevisiae during fermentation.

Recently, Casey et al. (1983, 1984) have shown that

yeast nutritional requirements limit fermentative activity

in high gravity brewing. Supplementing worts with yeast











extract and lipids substantially improved fermentation rates

and reduced the time required to complete the fermentation.

The studies reported in this chapter using a yeast

extract/peptone-based fermentation broth also illustrate

this point and provide further support for the hypothesis

that nutritional deficiencies, in addition to accumulated

ethanol, also are responsible for the initial decline in

fermentation activity during the accumulation of low levels

of ethanol.

The reduced fermentation rate of cells after the

production of approximately 5% (vol/vol) ethanol appears to

result from the combination of a small inhibitory effect of

ethanol and physiological changes in the cells. These

physiological changes were not induced by growth in the

presence of 5% (vol/vol) added ethanol or by growth in the

presence of ethanol along with other natural fermentation

products. Conditioned broth was deficient in nutrients

provided by yeast extract and supported very little growth.

The addition of 5 g/L of yeast extract restored the ability

of this spent broth to support vigorous growth and

fermentation. By further increasing the concentration of

yeast extract to 25 g/L in the growth medium, the decline in

fermentative activity associated with the initial production

of 5% (vol/vol) ethanol was partially prevented. These

results support the hypothesis that physiological changes in

the cells caused by nutrient limitation are major factors in









55

the initial 50% decline in fermentative activity. Further

studies will include identification of this limiting

nutrient and, upon supplementation, characterization of its

effect on growth and fermentation.















CHAPTER IV
MAGNESIUM LIMITATION AND ITS ROLE IN THE APPARENT TOXICITY
OF ETHANOL DURING YEAST FERMENTATION


Introduction

The rate of ethanol production by Saccharomyces spp.

decreases in batch fermentations as alcohol accumulates in

the medium (Moulin et al., 1984; Rahn, 1929; Strehaiano and

Goma, 1983). The onset of this decline in fermentative

activity occurs at very low ethanol concentrations, often

less than 3% (vol/vol). Since ethanol has been shown to

inhibit fermentation (Brown et al., 1981; Cysewski and

Wilke, 1977; Gray, 1941), it generally has been accepted

that this accumulation of ethanol is responsible for the

progressive decline in fermentative activity (Bazua and

Wilke, 1977; Ghose and Tyagi, 1979; Luong, 1985). However,

the extent of inhibition by exogenously added ethanol is

less than would be predicted by the decline in fermentation

rate which normally occurs during the fermentative

accumulation of ethanol (Fig. 3).

Further studies have attempted to define the

mechanisms) of ethanol inhibition of fermentation and to

reconcile the failure of added ethanol to inhibit

fermentation to the extent observed during the fermentative

accumulation of ethanol. Early experiments provided

56











evidence that the intracellular concentration of ethanol was

much higher than that of the surrounding medium during

fermentation (Nagodawithana and Steinkraus, 1976; Navarro

and Durand, 1978; Panchal and Stewart, 1980), a condition

not readily duplicated by exogenously added ethanol.

However, these early data can be explained by problems in

the measurement of internal ethanol concentrations (Dasari

et al., 1984). Several research groups have developed

independent methods which demonstrated that ethanol is

freely permeable in Saccharomyces spp. and that the

intracellular concentration of this metabolic product is the

same as that in the surrounding fermentation broth (Dasari

et al., 1985; Guijarro and Lagunas, 1984; Table 2).

Additional studies have investigated the sensitivity of

glycolytic enzymes and alcohologenic enzymes to in vitro

inhibition by ethanol. Millar et al. (1982) have shown that

these enzymes are stable in ethanol concentrations higher

than 20% (vol/vol). The two enzymes most sensitive to

inhibition by ethanol were pyruvate decarboxylase and

phosphoglycerate kinase. Both, however, retained, 50% of

maximal activity in the presence of over 12% (vol/vol)

ethanol, the final alcohol concentration achieved by the

complete fermentation of 200 g of glucose per L of broth.

Similarly, Larue et al. (1984) concluded that the cessation

of alcohol production during stuck fermentations was not due











to ethanol inhibition of alcohol dehydrogenase and

hexokinase activities.

Casey et al. (1984) have reported that nutrient

limitation is a major factor restricting the ethanol

productivity of high-gravity fermentations. Anaerobically

cultured yeasts are known to have a nutritional requirement

for ergosterol and unsaturated lipids (Hossack and Rose,

1976; Nes et al., 1978; Proudlock et al., 1968).

Unsaturated lipids have been shown to increase biomass,

alcohol production and ethanol durability of yeast cells

during anaerobic fermentation (Ingram and Buttke, 1984;

Janssens et al., 1983; Lafon-Lafourcade et al., 1979; Thomas

et al., 1978). A variety of lipid-protein complexes and

nutrient supplements, ranging from albumin-ergosterol-

monoolein to soy flour and yeast extract, also have been

shown to yield increased rates of alcohol production and

higher final ethanol concentrations (Damiano and Wang, 1985;

Hayashida et al., 1976; Lafon-Lafourcade et al., 1979; Ohta

and Hayashida, 1983).

The studies described in chapter III suggest that the

initial decline in fermentative activity during batch

fermentation of 20% glucose is not caused by the presence of

ethanol or by growth in the presence of 5% (vol/vol)

ethanol. These studies indicated that a components) of

yeast extract was limiting cell growth and that this

limitation contributed to the early loss of fermentative











activity. The results presented in this chapter identify

magnesium as the limiting component of yeast extract and

demonstrate that when this nutrient limitation is relieved,

a dramatic decrease in the time required for total

conversion of glucose to ethanol is achieved. This decrease

in time required for the completion of fermentation resulted

from a delay in the onset of stationary phase which

increased the total cell number during that part of

fermentation in which over 90% of the ethanol is produced.

Materials and Methods

Organisms and Growth Conditions

The principal organism used in these studies was

Saccharomyces cerevisiae KD2, described in chapter II. In

addition, S. cerevisiae CC3, S. cerevisiae A10 (NRRL Y-

12707) and S. sake (NRRL Y-11572) were used for comparison

in some experiments. The latter two strains generously were

provided by N.J. Alexander (Northern Regional Research

Center, U.S. Department of Agriculture, Peoria, Ill.). All

organisms were grown in YEPD broth and maintained on YEPD

agar, as stated in chapter II. Batch fermentations also

were carried out as described in chapter II.

Preparation of Fermentation Samples for Analysis

Fermentation samples were centrifuged at 10,000 x g for

0.5 min. The supernatant was removed and saved by freezing

at -200C. Cells were washed once in 50 mM KH2PO4 buffer (pH











5.0) and the pellets were saved for further analysis by

freezing at -200C.

Preparation of Glucose-Reconstituted Medium for Growth
Experiments

Batch fermentations were allowed to reach an optical

density at 550 nm of 3.5. Cells were removed by

centrifugation in a Sorvall RC-2B centrifuge at 10,000 x g

for 2 min. The amount of ethanol in the supernatant was

determined and used to estimate the amount of glucose needed

to reconstitute the medium to a concentration of 20%. This

glucose-reconstituted medium was sterilized by vacuum

filtration with 0.45 pm Metricel membrane filters (Gelman

Sciences Inc., Ann Arbor, Mich.).

Preparation of Ashed Medium Components

Yeast extract (20 g) and peptone (30 g) were burned

over a gas burner for 5 h in a porcelain crucible. After

being transferred to a muffle furnace, the medium components

were ashed at 6000C for 72 h. The yeast extract ash was

suspended in 40 ml of deionized water and the peptone ash

was suspended in 30 ml of deionized water. These aqueous

suspensions of ash were adjusted to pH 5.0 with concentrated

HC1 and sterilized by autoclaving.

Nutrient Supplementation Growth Experiments

Nutrient supplements were added to culture tubes

containing 5 ml fresh YEPD medium or glucose-reconstituted

medium and a 1% by volume inoculum (initial optical density

at 550 nm of 0.035). Culture tubes were incubated at 30C











and agitated (30 rpm) in a Rototorque culture rotator (Cole-

Parmer, Chicago, Ill.).

Medium Analyses

Ethanol and glucose were measured as described in

chapter II. The magnesium concentration of the medium was

measured with the 60 Second Magnesium reagents purchased

from American Monitor Corporation, Indianapolis, Ind., as

described by Osman and Ingram (1985).

Cellular Analyses and Respirometry Measurements

Cell mass and total cell protein were measured as

described in chapter II. To determine the amount of

intracellular magnesium, yeast cells were washed once in

50 mM KH2PO4 buffer (pH 5.0) and the cell pellets were

stored frozen at -200C until analyzed. These yeast pellets

contained 1 to 3 mg of cell protein and were permeabilized

by incubation in a boiling-water bath for 1.5 min. The

resulting debris was suspended in 1 ml of 50 mM KH2PO4

buffer (pH 5.0) and then pelleted. The supernatant was

analyzed for magnesium as described above. Respirometry

measurements were made as described in chapter II and

fermentation rates were calculated from these values as

pmoles of CO2 produced per h per mg cell protein.

Viable-Cell Determinations

Cell numbers were determined microscopically with a

Petroff-Hausser counting chamber. Viable-cell counts were











determined by the methylene blue staining procedure of Mills

(1941).

Chemicals

Yeast extract, peptone and agar were obtained from

Difco Laboratories, Detroit, Mich. Glucose and other

biochemicals were obtained from Sigma Chemical Co., St.

Louis, Mo. Magnesium sulfate and other inorganic salts were

purchased from Fisher Scientific Company, Orlando, Fla.

Absolute ethanol was supplied by AAPER Alcohol and Chemical

Co., Shelbyville, Ky. Gas chromatography supplies were

obtained from Supelco, Bellefonte, Pa.

Results

Effect of Nutrient Supplements on Growth in Glucose-
Reconstituted Medium

As shown in Tables 3 and 4, fermentation broth in which

S. cerevisiae KD2 had grown for 12 h supported very little

further growth and limited the fermentative activity of

strain KD2 even after supplementation with glucose (glucose-

reconstituted medium). At this stage of fermentation (1.2%

(vol/vol) accumulated ethanol), ethanol production was at

its maximum rate (50 pmoles/h per mg protein). This time

point also marked the end of exponential growth (Fig. 1),

indicating either a nutrient limited state or the presence

of an inhibitor.

The addition of yeast extract and peptone at the

original medium concentration restored the ability of the

used medium to support growth at 71% and 54% of the control





















Table 4. Effect of nutrient supplementation
S. cerevisiae KD2


on growth of


Optical Density % of
Medium Supplement at 550 nm after control (SD)
48 h (SD)


YEPD None 13.8 (1.5) 100

12-ha None 1.25 (0.48) 9.1 (4.0)

12-ha Yeast extract (5 g/L) 9.77 (0.75) 71 (9)
12-ha Peptone (10 g/L) 7.43 (0.35) 54 (6)
12-ha Ashed yeast extract 9.70 (0.40) 70 (8)
12-ha Ashed peptoneb 3.47 (0.20) 25 (3)

12-ha Trace minerals 1.88 (0.20) 14 (2)
12-ha KH2PO4 (7.3 mM) 1.49 (0.17) 11 (2)
12-ha (NH4)2SO4 (7.6 mM) 1.60 (0.35) 12 (3)
12-ha MgSO4 (2 mM) 12.7 (0.1) 92 (10)
12-ha MgCl2 (2 mM) 12.1 (0.5) 88 (10)
12-ha CaC12 (2 mM) 1.55 (0.14) 11 (2)
12-ha Na2SO4 (2 mM) 1.62 (0.20) 12 (2)


a Medium isolated from a batch fermentation after 12 h of
yeast growth and supplemented to 20% with glucose.

b An amount of ashed yeast extract equivalent to 5 g of
whole yeast extract per L or an amount of ashed peptone
equivalent to 10 g of whole peptone per L.


c As described by Wickersham (1951).











level, respectively (Table 4). These results indicate that

nutrient limitation rather than the presence of an inhibitor

was responsible for the inability of the used medium to

support further yeast growth. Vitamin supplements also were

tested and did not promote growth in this glucose-

reconstituted medium (data not shown).

The organic components of yeast extract and peptone are

both diverse and complex. Before embarking on a

fractionation of these, the inorganic constituents were

tested after ashing. Supplementation with ashed yeast

extract was as effective as with whole yeast extract, while

ashed peptone was only half as effective as whole peptone.

These results suggested that an inorganic component of YEPD

medium was the principle factor limiting growth.

The inorganic constituents of a mineral-based minimal

medium were tested to determine which ions were limiting

(Table 4). The addition of potassium, ammonium, sodium,

calcium, phosphate, sulfate and a trace mineral mixture

described by Wickersham (1951) did not promote growth in

glucose-reconstituted medium. Only magnesium salts were

effective as nutrient supplements, allowing growth

equivalent to 90% of the control in fresh YEPD medium.

The dose-response of growth to added magnesium, yeast

extract and ashed yeast extract is shown in figure 6. Yeast

extract contained 27 moles of magnesium per g. This value

was used to calculate the appropriate amount of whole and




























Figure 6.


Dose-response of cell growth to added
magnesium. Magnesium values represent
the amount of magnesium contained in
the added nutrient supplement. Error
bars represent the average standard
deviation for each experiment.
Symbols; U whole yeast extract
added to glucose-reconstituted, used
medium; 0 ashed yeast extract
added to glucose-reconstituted, used
medium; 0 MgSO4 added to
glucose-reconstituted, used medium;
0 MgSO4 added to fresh YEPD broth.
















18

16


12I


0 0.2 0.4
MAGNESIUM


1.0


0.6 0.8
(mMolar)











ashed yeast extract to be added. Whole yeast extract, ashed

yeast extract and MgSO4 gave similar dose-responses.

However, at concentrations below 0.2 mM, MgSO4 appeared to

be a better supplement. Magnesium sulfate-supplemented

fresh YEPD medium also was plotted for comparison. Maximum

growth occurred at added magnesium concentrations above

0.2 mM. A MgSO4 concentration of 0.5 mM was chosen for

subsequent fermentation studies because growth at this

concentration was no longer limited by an inadequate supply

of magnesium.

To confirm that magnesium indeed was limiting in YEPD

medium, the magnesium content of cells and the surrounding

broth was determined at various times during batch

fermentation (Fig. 7). The magnesium content of the cells

reached a maximum of 130 nmoles/mg cell protein at 12 h,

rapidly declining to 48 nmoles/mg cell protein by 24 h and

remaining at this lower level throughout the final period of

fermentation. In the medium, the magnesium content fell to

less than 0.05 mM by 24 h and remained constant until

fermentation had been completed. Thus, the decline in

magnesium content per mg cell protein observed after 12 h

appears to result from continued cell growth after near

depletion of the magnesium in the surrounding broth.

Supplementing the broth with 0.5 mM magnesium resulted in

the peak accumulation of higher levels of magnesium

(200 nmoles/mg cell protein) at 12 h, followed by a decline












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to about 130 nmoles of magnesium per mg cell protein after

24 h. Magnesium-supplemented cultures maintained a higher

level of cellular magnesium throughout fermentation than

cultures grown in unsupplemented YEPD medium.

Magnesium-Limited Growth of Other Yeast Strains

Three other yeast strains were investigated to

determine whether magnesium also limited their growth: S.

cerevisiae CC3 (parent organism), S. cerevisiae A10 and S.

sake. Glucose-reconstituted medium was prepared for each of

these strains. Cultures were inoculated into their

respective glucose-reconstituted medium with and without

added magnesium (0.5 mM) and incubated for 48 h on a

rotator. S. cerevisiae CC3 and S. sake exhibited magnesium-

dependent growth almost identical to that reported for

strain KD2. The optical density at 550 nm was 8.3 to 9.4

after 48 h with added magnesium and 0.5 without added

magnesium. S. cerevisiae A10 grew poorly in its glucose-

reconstituted medium, with an optical density at 550 nm of

1.0 to 1.2 after 48 h for both control and supplemented

cultures. All three strains reached a similar cell density

in fresh YEPD medium (optical density at 550 nm of 14.2 to

15.2). These results indicate that the magnesium limitation

observed in strain KD2 was not caused by the petite mutation

and was not limited to strain CC3 and its derivatives.

However, additional factors were clearly involved with S.

cerevisiae A10.











Effect of Magnesium Supplementation of Batch Fermentation

The effects of supplementing YEPD medium with 0.5 mM

MgSO4 on batch fermentation are illustrated in figure 8.

The production of cell mass as measured by cellular protein

is shown in figure 8A. Supplementation with magnesium

prolonged the exponential rise in cellular protein, allowing

a 53% increase in cell mass over that of the control within

18 h after inoculation. The addition of magnesium also

increased the rate at which glucose was consumed and ethanol

was produced (Fig. 8B and 8C). After 30 h of incubation,

magnesium-supplemented cultures had produced one-third more

ethanol than the controls. The conversion of glucose to

ethanol was complete after 48 h in magnesium-supplemented

cultures, but required 72 h in control YEPD broth. The

final yield of ethanol was essentially identical for both

magnesium-supplemented and control cultures, 12.7% (vol/vol)

(98% of theoretical maximum yield).

Effect of Magnesium Supplementation on Rate of Fermentation

Samples were removed from magnesium-supplemented and

control fermentations at various times during batch

fermentation. Ethanol concentration, cell protein and CO,

evolution of unwashed cells were measured. Figure 9 shows

the fermentation rate as a function of accumulated ethanol.

Both control and magnesium-supplemented cultures exhibited

the same maximum rate of fermentation at 1% (vol/vol)

ethanol. However, magnesium-supplemented cultures











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Figure 9.


Effect of added magnesium on the rate
of fermentation. Fermentation rates
were measured by respirometry of
unwashed cells immediately after
sampling and are plotted as a function
of accumulated ethanol for four
separate batch fermentations. Closed
symbols represent cultures
supplemented with 0.5 mM MgSO4 and
open symbols represent control
fermentations in YEPD broth alone.
















60-



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x 40
Z E



z- O
Wo


E
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0 '

0 2 4 6 8 10 12
ETHANOL (%v/v)











maintained a higher rate of fermentation as ethanol

accumulated during the completion of the batch fermentation.

This rate was 40% higher than that of control cells after

the accumulation of 8% (vol/vol) ethanol. The fermentation

rate of both supplemented and unsupplemented cultures fell

precipitously at about 12.5% (vol/vol) ethanol, coincident

with exhaustion of glucose.

Effect of Magnesium Addition on Cell Viability

The percentage of viable cells in both magnesium-

supplemented and unsupplemented batches remained greater

than 90% for the first 48 h of the fermentation. Glucose

was exhausted at this time in supplemented cultures and the

percentage of viable cells began to decrease, reaching 58%

by 72 h. The unsupplemented batches consumed glucose more

slowly and maintained high viability (>90%) until between 60

and 72 h, the time at which glucose was exhausted.

Effect of Ethanol on the Fermentation Rate of Magnesium-
Supplemented Cultures

Two points during fermentation were chosen at which to

compare cells grown with and without added magnesium. These

were the same two points described in chapter III as 12-h

and 24-h cells. Cells at 12 h were still undergoing

exponential growth and cells at 24 h were in early

stationary phase.

Figure 10A shows the dose-response of fermentative

activity of the younger cells plotted as a function of total

ethanol concentration endogenouss plus added). Magnesium-











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supplemented and unsupplemented cells had a similar initial

fermentation rate, about 57 moles of CO2 produced per h per

mg cell protein and exhibited identical dose-response

curves. A concentration of 7.6% (vol/vol) ethanol resulted

in 50% inhibition of fermentative activity.

Figure 10B shows the effect of ethanol on the

fermentation rate of the older cells. Supplemented cells

had an initial fermentation rate of 27.4 pmoles of CO2

produced per h per mg cell protein with an SD of 0.5, while

unsupplemented cells exhibited a significantly lower rate,

22.9 pmoles of CO2 produced per h per mg cell protein with

an SD of 1.2. The fermentation rate of cells from

supplemented cultures was always higher than that of control

cells. The amount of ethanol present in the supplemented

culture was 3.0% (vol/vol) higher than in the unsupplemented

culture when compared at equal fermentation rates. The

fermentation rate of magnesium-supplemented and control

fermentations exhibited linear dose-responses to ethanol

addition. A measure of the sensitivity of fermentation rate

to ethanol is the slope of the dose-response curve. The

slope for supplemented batches was -0.17 with an SD of 0.02,

while that of the controls was -0.14 with an SD of 0.01.

The differences in these slopes (Fig. 10B) are suggestive,

but do not conclusively demonstrate that supplementation

with magnesium reduced the sensitivity of fermentation in

older cells to inhibition by ethanol.











Discussion

The decline in fermentation rate that begins at low

alcohol concentrations does not seem to be exclusively

caused by the immediate presence of ethanol, by growth in

the presence of ethanol or by cell death (Chapter III).

This decline appears to be related in part to a magnesium

deficiency, although other factors are involved also. In

yeast extract-peptone-based medium, a magnesium deficiency

is developed which limits cellular growth and the rate of

carbohydrate conversion into ethanol. The addition of

magnesium to batch fermentations prolonged exponential

growth, allowing greater accumulation of cell mass without

affecting cell viability. In addition, cells in magnesium-

supplemented cultures maintained a higher fermentation rate

as ethanol accumulated. These two factors, increased cell

mass plus higher fermentation rate, combined to reduce the

time required for the conversion of 20% glucose to ethanol

by one-third in magnesium-supplemented cultures.

During batch fermentation, yeast cells concentrated

magnesium from the medium. In unsupplemented cultures,

magnesium uptake stopped at the end of exponential growth.

At this point, the concentration of magnesium in the medium

was 48 pM, within the range of Km values reported for

magnesium transport by microorganisms (Jasper and Silver,

1977). The end of exponential growth also coincided with

the beginning of the decline in fermentative activity. In











magnesium-supplemented cultures, higher levels of

intracellular magnesium were achieved early in fermentation

and decreased to a lesser extent than observed in

unsupplemented cultures. Magnesium-supplemented cultures

had 6.5 times more magnesium in the medium at the end of

exponential growth than did unsupplemented cultures. Thus,

the magnesium supply of the supplemented culture appears to

be adequate for growth and other factors are limiting the

fermentative ability of the yeasts under these conditions.

The ubiquitous role of magnesium in cellular processes

is well documented (Jasper and Silver, 1977). Magnesium

constitutes a major portion of the cellular cations, mostly

bound in structures such as ribosomes and the cell envelope.

The free cation concentration, however, may play a more

direct role in regulating overall cellular metabolism and

cell division (Walker and Duffus, 1980). Many of the

enzymes that function in DNA replication, transcription and

translation require magnesium for activity. In fermentation

pathways, magnesium is a required cofactor and nucleotide

counter-ion in many reactions. Magnesium levels typically

are maintained at mmolar intracellular concentrations and it

is not surprising that this cation is a limiting nutrient

during high-gravity fermentations.

Previous studies have demonstrated that the inhibition

of fermentation by added ethanol in Zymomonas mobilis is

primarily due to ethanol-induced leakage, particularly of











magnesium (Osman and Ingram, 1985). The addition of

magnesium salts at 0.5 mM substantially reversed the

inhibitory effects of up to 13% (vol/vol) ethanol. Although

analogous studies have not been performed with S.

cerevisiae, it is likely that ethanol also increases the

leakage of small molecules in this organism.

Casey et al. (1984) also have reported that nutrient

limitation is an important factor in limiting the

productivity of a fermentation. Supplementation of high-

gravity brewing wort (containing up to 31% dissolved solids)

with yeast extract, ergosterol and oleic acid allowed the

production of 16.2% (vol/vol) ethanol by brewers' yeast.

Higher rates of alcohol production primarily resulted from

an increase in cell mass associated with nutrient-

supplemented fermentations and did not appear to include an

increase in the resistance of fermentation rate to ethanol.

Addition of nutrients in the form of soy flour to

fermentation broth has been shown to increase the

fermentative productivity of both S. cerevisiae (Damiano and

Wang, 1985) and Z. mobilis (Ju et al., 1983). Viegas et al.

(1985) also reported that soy flour addition to a yeast

extract-based medium containing 30 to 40% glucose enhanced

the rate of ethanol production by S. bayanus. Again,

supplementation led to an increase in cell concentration.

It was further demonstrated that the aqueous fraction of soy

flour, rather than the lipid fraction, contained the











components beneficial for fermentation. This aqueous

fraction would have included inorganic ions, such as

magnesium. Indeed, it is possible that many of the complex

nutrient additives used to increase ethanol production also

are correcting an inorganic ion deficiency.

The causes of the progressive decline in fermentative

activity which is observed as ethanol accumulates during

batch fermentation appear to be much more complicated than

expected. The results presented in this chapter indicate

that direct ethanol inhibition is only partially

responsible. A nutrient limitation for magnesium also

appears to be partially responsible. With abundant

magnesium, only a 50% further increase in cell mass was

observed, indicating that another factors) becomes limiting

for growth and fermentation at this point. Indeed, a

complete understanding of the biochemical basis for the

decline in fermentation rate in yeasts may require

determination of the factors responsible for the termination

of exponential growth and the associated physiological and

enzymatic changes.















CHAPTER V
GLYCOLYTIC ENZYMES AND INTERNAL pH


Introduction

Saccharomyces cerevisiae is capable of very rapid rates

of glycolysis and ethanol production under optimal

conditions, producing over 50 pmoles of ethanol per h per mg

of cell protein (Fig. 9). However, this high rate is

maintained for only a brief period during batch fermentation

and declines progressively as ethanol accumulates in the

surrounding broth (Casey and Ingledew, 1986; Ingram and

Buttke, 1984; Moulin et al., 1984). Earlier studies have

identified a requirement for lipids (Beaven et al., 1982;

Casey et al., 1984; Thomas et al., 1978) or molecular oxygen

for lipid biosynthesis (Andreasen and Stier, 1954; Buttke et

al., 1980; Buttke and Pyle, 1982) in many fermentation

broths as being essential for the maintenance of high

fermentative activity. Magnesium is an essential cofactor

for many of the glycolytic enzymes and has been identified

also as a limiting nutrient in fermentation broth containing

peptone and yeast extract (Chapter IV). Supplying these

nutritional needs reduces but does not eliminate the decline

in fermentative activity during batch fermentation (Fig. 9).











The basis for the decline in fermentation rate is not

fully understood. Since the addition of ethanol to cells in

batch cultures and in chemostats causes a dose-dependent

inhibition of ethanol production (Casey and Ingledew, 1986;

Fig. 10), most investigations have focused on ethanol as the

inhibitory agent (Casey and Ingledew, 1986; Ingram and

Buttke, 1984; Millar et al., 1982). Ethanol is known to

alter membrane permeability and disrupt membrane function in

a variety of biological systems (Casey and Ingledew, 1986;

Ingram and Buttke, 1984). In yeast, ethanol causes an

increase in hydrogen ion flux across the plasma membrane of

cells suspended in water (Cartwright et al., 1986). This

increased hydrogen ion flux has been proposed as being

responsible for the ethanol-induced decline in transport

rates observed under similar conditions (Beaven et al.,

1982; Leao and van Uden, 1982b, 1984a, 1984b).

Evidence has been accumulating which indicates that the

presence of ethanol may not be the only factor responsible

for the decline in fermentative activity. The replacement

of fermentative broth containing ethanol with fresh medium

lacking ethanol did not immediately restore fermentative

activity (Table 1). In a comprehensive study, Millar et al.

(1982) demonstrated that ethanol concentrations below

12% (vol/vol) do not denature glycolytic enzymes or cause

appreciable inhibition of activity in vitro under substrate-

saturating conditions. Since ethanol does not accumulate











within yeast cells, but rapidly diffuses across the cell

membrane (Dasari et al., 1985; Guijarro and Lagunas, 1984;

Table 2), direct inhibition of glycolytic enzymes by

intracellular ethanol is unlikely during fermentations which

produce 12% (vol/vol) ethanol or less.

In this chapter, changes in the amounts of glycolytic

and alcohologenic enzymes, and internal pH and membrane

energization have been examined as possible physiological

causes for the decline in fermentative activity during batch

fermentations of 20% glucose in a yeast extract-peptone-

based medium supplemented with magnesium.

Materials and Methods

Organism and Growth Conditions

The petite yeast strain characterized in chapter II, S.

cerevisiae KD2, was used in the studies presented in this

chapter. This yeast was grown in the complex medium also

described in chapter II, except that the medium was

supplemented with 0.5 mM MgSO4, as described in chapter IV.

Batch fermentations were carried out as outlined in

chapter II.

Analytical Methods

Cell mass, glucose, ethanol and fermentation rates all

were determined by the methods discussed in chapter II.

Cell protein was measured as described by Lowry et al.

(1951). The protein content of cell extracts was

quantitated using the method of Bradford (1976). Bovine











serum albumin served as the protein standard for both

methods.

Enzyme Analyses

Activities of glycolytic and alcohologenic enzymes were

determined in 2-ml samples removed at various times during

batch fermentation. Cells were harvested by centrifugation

at 10,000 x g for 30 sec at 40C and washed in an equal

volume of 50 mM potassium phosphate buffer (pH 7.4). All

subsequent steps were carried out at 40C. The pellet was

suspended in the same buffer containing 2 mM mercaptoethanol

and 2 mM EDTA, and disrupted with 0.1-mm glass beads using a

Mini-Bead Beater (Biospec Products, Bartlesville, Okla.).

Five 1-min periods of disruption, each were followed by 5-

min periods of cooling on ice. Cell debris was removed by

centrifugation at 10,000 x g for 5 min, and the supernatant

was assayed immediately for enzymatic activities. Only two

enzymes at a time were assayed in each batch fermentation

experiment to avoid problems which could result from storage

of cells or extracts.

Pyruvate decarboxylase and all glycolytic enzymes were

assayed spectrophotometrically by the methods of Maitra and

Lobo (1971) as modified by Clifton et al. (1978). All

enzymes were assayed under substrate-saturating conditions

except triose phosphate isomerase, which was assayed with

1 mM substrate. The amounts of coupling enzymes were

adjusted as needed to ensure a linear reaction rate.











Alcohol dehydrogenase was assayed by measuring the oxidation

of ethanol as described by Maitra and Lobo (1971), but using

a buffer at pH 8.7 containing 75 mM sodium pyrophosphate,

75 mM semicarbazide hydrochloride and 21 mM glycine (Bernt

and Gutman, 1971).

Determination of Internal pH and Membrane Energization

The measurements of internal pH and A were

performed using 7-[14C]benzoic acid and [3H-

phenyl]tetraphenylphosphonium bromide, respectively.

Protocols were similar to those described by Cartwright et

al. (1986) except that cells were incubated in their native

growth medium rather than distilled water and 0.4-pm pore

size polycarbonate filters were used instead of mixed

cellulose ester filters. Cell volumes were determined as

described in chapter III. As a control for adventitious

binding of radioactive compounds, cells were permeabilized

with a combination of ethanol, toluene and Triton X-100 as

described by Salmon (1984), washed with 50 mM phosphate

buffer, resuspended in native broth and processed. This

treatment resulted in a complete collapse of A pH and loss

of membrane potential. Calculations were performed as

described by Rottenberg (1979).

Materials

Yeast extract, peptone and agar were obtained from

Difco Laboratories, Detroit, Mich. Glucose, coupling

enzymes, coenzymes and substrates were purchased from Sigma











Chemical Co., St. Louis, Mo. Inorganic salts were obtained

from Fisher Scientific Co., Orlando, Fla. Absolute ethanol

was supplied by AAPER Alcohol and Chemical Co., Shelbyville,

Ky. Radioactive compounds were purchased from New England

Nuclear Corp., Boston, Mass.

Results

Reversibility of the Decline in Fermentative Activity

In chapter IV, the ability of magnesium supplementation

to partially relieve the decline in fermentative activity

associated with the accumulation of ethanol was

demonstrated. After supplementation, immediately reversible

inhibition by accumulated ethanol may be responsible for the

remaining decrease in fermentation rate. To investigate

this possibility, cells were removed at various times during

batch fermentation and the rate of ethanol production per mg

cell protein was determined (Fig. 11). Cells were most

active at the earliest times measured, 12 h, and

fermentation rate declined by 50% when 6.5% (vol/vol)

ethanol had accumulated after 24 h. Approximately 40% of

the fermentative activity was retained after the

accumulation of 10% (vol/vol) ethanol with 30 g glucose

per L remaining in the fermentation broth. The abrupt,

final decline in activity reflects the near-complete

exhaustion of glucose. Removal of ethanol from cells by

washing and suspending in fresh medium resulted in only a

modest increase in fermentative activity in all but the




























Figure 11. Effect of ethanol removal on the
fermentative activity of cells grown
in YEPD medium containing 0.5 mM
MgSO4. Cells were sampled during
batch fermentation and were either
untreated or washed once and then
suspended in fresh medium containing
20% glucose. The fermentation rate of
these samples were measured
immediately by respirometry. Symbols:
0 activity measured in native
broth; 0 activity measured after
cells were suspended in fresh medium.
























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