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RESEARCH ARTICLE
Anaerobic homolactate fermentation with Saccharomyces cerevisiae
results in depletion of ATP and impaired metabolic activity
Derek A. Abbott1,2, Joost van den Brink1,2, Inge M.K. Minneboo1,2, Jack T. Pronk1,2 & Antonius J.A. van
Maris1,2
1
Department of Biotechnology, Delft University of Technology, Delft, The Netherlands; and 2Kluyver Centre for Genomics of Industrial Fermentation,
Delft, The Netherlands
Correspondence: Antonius J.A. van Maris,
Department of Biotechnology, Delft University
of Technology, Julianalaan 67, 2628BC Delft,
The Netherlands. Tel.: 131 15 278 1616;
fax: 131 15 278 2355; e-mail:
[email protected]
Received 7 January 2009; revised 12 February
2009; accepted 13 February 2009.
First published online 30 March 2009.
DOI:10.1111/j.1567-1364.2009.00506.x
Editor: Monique Bolotin-Fukuhara
Keywords
lactate; lactic acid; energetics; enzyme activity;
intracellular metabolite.
Abstract
Conversion of glucose to lactic acid is stoichiometrically equivalent to ethanol
formation with respect to ATP formation from substrate-level phosphorylation,
redox equivalents and product yield. However, anaerobic growth cannot be
sustained in homolactate fermenting Saccharomyces cerevisiae. ATP-dependent
export of the lactate anion and/or proton, resulting in net zero ATP formation, is
suspected as the underlying cause. In an effort to understand the mechanisms
behind the decreased lactic acid production rate in anaerobic homolactate cultures
of S. cerevisiae, aerobic carbon-limited chemostats were performed and subjected to
anaerobic perturbations in the presence of high glucose concentrations. Intracellular measurements of adenosine phosphates confirmed ATP depletion and
decreased energy charge immediately upon anaerobicity. Unexpectedly, readily
available sources of carbon and energy, trehalose and glycogen, were not activated
in homolactate strains as they were in reference strains that produce ethanol. Finally,
the anticipated increase in maximal velocity (Vmax) of glycolytic enzymes was not
observed in homolactate fermentation suggesting the absence of protein synthesis
that may be attributed to decreased energy availability. Essentially, anaerobic
homolactate fermentation results in energy depletion, which, in turn, hinders
protein synthesis, central carbon metabolism and subsequent energy generation.
Introduction
Lactic acid, used for food preservation, production of
cosmetics and pharmaceuticals and traditionally produced
using various species of lactobacilli (Benninga, 1990), can
also be used for the production of the bio-based biodegradable polymer polylactic acid (Benninga, 1990; Datta et al.,
1995). The pH sensitivity and limited ability to synthesize
B-vitamins and amino acids of these lactobacilli, increases
the cost of lactic acid production due to the requirement of
complex nutrients and the formation of large amounts of
gypsum as a byproduct (Benninga, 1990; Chopin, 1993).
Because Saccharomyces cerevisiae is relatively acid tolerant
and can grow anaerobically in synthetic media, it is an
interesting candidate to produce lactic acid (Dequin &
Barre, 1994; Porro et al., 1995).
The deletion of one or more of the functional genes
encoding pyruvate decarboxylase in combination with the
expression of a heterologous lactate dehydrogenase results in
FEMS Yeast Res 9 (2009) 349–357
S. cerevisiae strains capable of producing lactic acid with the
limitation or elimination of ethanol formation (Adachi
et al., 1998; Porro et al., 1999; van Maris et al., 2004b; Saitoh
et al., 2005; Ishida et al., 2006). Considering the intracellular
stoichiometry, production of lactic acid is equivalent to
ethanol formation with respect to energy and redox metabolism. As with ethanol formation, the conversion of 1 mol
of glucose to 2 mol of lactic acid via glycolysis results in the
formation of 2 moles of ATP. Similarly, NADH generated in
glycolysis is oxidized to NAD1 during both ethanol and
lactic acid production via the activity of alcohol dehydrogenase or lactate dehydrogenase, respectively. As such,
anaerobic lactic acid production by S. cerevisiae was expected to be equivalent to alcoholic fermentation. Indeed,
the exposure of homofermentative lactate-producing
S. cerevisiae strains, grown in aerobic carbon-limited chemostat conditions, to an excess of glucose resulted in the
immediate secretion of lactate as the major fermentation
product. However, under anaerobic conditions the strain
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350
was incapable of sustaining growth, and the lactate production rate rapidly decreased (van Maris et al., 2004b). Subsequently, oxygen-limited chemostat cultivation showed that
lactic acid production did not result in the (net) formation
of ATP in lactic acid-producing S. cerevisiae. Moreover, the
positive influence of oxygenation on the lactate production
was observed with a 2.5-fold increase of the production rate
in aerobic conditions (van Maris et al., 2004b).
It has been suggested that, whereas ethanol leaves the cell
via passive diffusion (Guijarro & Lagunas, 1984), energydependent efflux of the proton and/or the lactate anion
results in a net ATP requirement for lactic acid export (van
Maris et al., 2004b). The high intracellular pH ensures that
the majority of lactic acid is present as the lactate anion,
which is incapable of diffusing across the plasma membrane.
Therefore, analogous to the export of other weak organic
acid anions (Piper et al., 1998; Fernandes et al., 2005), it is
likely that ATP-dependent export of the lactate anion is
required. At the very least, ATP is required to export the
dissociated proton in order to maintain intracellular pH.
Although not in agreement with the above mentioned
observations under oxygen limitation, in the worst case
scenario ATP-dependent mechanisms may be involved in
both proton and anion export.
These intriguing observations make one wonder about
the intracellular processes that occur during anaerobic
homolactic fermentation. Perturbations in oxygen availability and especially the glucose concentrations are known to
affect the intracellular concentrations of the adenosine
phosphates in wild-type strains (Kresnowati et al., 2006).
The drastic differences between homolactic and alcoholic
fermentation might therefore either originate from, or
cause, differences in the energy charge of the cells. Not only
these conserved moieties, but also the activities of the
glycolytic enzymes themselves are known to respond to
perturbations in the glucose concentration (van den Brink
et al., 2008a). Additionally, in wild-type S. cerevisiae cells,
the storage carbohydrates glycogen and trehalose are mobilized when carbon-limited cells are exposed to glucose excess
(van den Brink et al., 2008b). Conversion of glycogen to
glucose-6-phosphate via glucose-1-phosphate uses inorganic phosphate (Pi) and is therefore ATP neutral (François &
Parrou, 2001). Consequently, glycogen accumulated during
aerobic carbon-limited chemostat cultivation may initially
provide a source of free energy to drive lactic acid production and cellular maintenance under anaerobic conditions.
In this study, aerobic carbon-limited chemostats were
used as a reproducible starting point for comparison of the
physiological response of homolactic (RWB 850-2) and
alcoholic-fermenting (CEN.PK 113-7D) S. cerevisiae strains
to anaerobic glucose-pulse experiments. To investigate
the role of ATP and storage carbohydrates in homolactate
fermentation, intracellular metabolites (including ATP, ADP
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D.A. Abbott et al.
and AMP) and glycogen and trehalose were measured.
Finally, the glycolytic pathway was measured for changes in
maximal velocity (Vmax) to evaluate the effects of anaerobic
homolactate fermentation on central carbon metabolism.
Materials and methods
Strains and media
The S. cerevisiae reference strain used in this study was a
prototrophic haploid reference strain CEN.PK113-7D
(MATa) (van Dijken et al., 2000). The lactate-producing
strain, RWB 850-2, is derived from the reference strain with
disruptions in PDC1, PDC5 and PDC6 and high-level
expression of Lactobacillus casei LDH gene (van Maris et al.,
2004b). Cultures were grown at 30 1C in shake flasks
containing 100 mL of synthetic medium (Verduyn et al.,
1992) with 2% (w/v) glucose for the reference strain or 2%
(v/v) ethanol, providing cytosolic acetyl-CoA and avoiding
excessive acidification, for the lactate-producing strain.
Chemostat cultivation
Synthetic medium was prepared as described previously
(Verduyn et al., 1992). Glucose (6.0 g L1) was added to the
synthetic medium after separate heat sterilization at 110 1C
and 1.15 g L1 of absolute ethanol was added separately.
Each strain was grown at 30 1C in 2.0-L bioreactors (Applikon) with a working volume of 1.5 L via an electrical level
sensor. The dilution rate was set at 0.10 h1, and the pH was
measured online and maintained at 5.0 by the automatic
addition of 2 M KOH using an Applikon ADI 1030 Biocontroller. A stirrer speed of 800 r.p.m. and air flow of
0.75 L min1 were applied to maintain the dissolved oxygen
concentration, as measured with an oxygen electrode, above
60% of air saturation in all chemostat cultivations performed. Steady-state samples were withdrawn after c. 10
volume changes to avoid strain adaptation due to long-term
cultivation (Ferea et al., 1999; Jansen et al., 2004).
Analytical methods
Chemostat cultures were assumed to be in steady state when,
after at least five volume changes, the culture dry weight and
specific carbon dioxide production rate changed by o 2%
over 2 volume changes. Culture dry weights were determined in duplicate via filtration onto dry, preweighed
nitrocellulose membranes. Samples were dried in a microwave oven for 20 min at 360 W. Culture supernatants were
obtained after centrifugation of chemostat broth or by a
rapid sampling method using precooled ( 20 1C) steel
beads (Mashego et al., 2003). Extracellular metabolite concentrations were analyzed via HPLC using an AMINEX
HPX-87H ion exchange column with 5 mM H2SO4 as the
FEMS Yeast Res 9 (2009) 349–357
351
Energetic constraints in yeast homolactate fermentation
mobile phase. Off-gas was first cooled with a condenser
(2 1C) and then dried with a Perma Pure dryer (PD-62512P). CO2 and O2 concentrations in the off-gas were
measured with an NGA 2000 Rosemount gas analyzer.
Perturbation experiments
Anaerobic glucose-pulse experiments were started by sparging a steady-state carbon-limited aerobic chemostat culture
with 0.75 L min1 of nitrogen gas (Hoek-Loos, Schiedam,
o 5 p.p.m. O2). NorpreneTM tubing and butyl rubber septa
were used to minimize oxygen diffusion into the anaerobic
cultures (Visser et al., 1990). Two minutes after nitrogen
sparging started and just before adding the glucose, the
medium and effluent pumps were switched off. The 200 mM
(added as 60 g of glucose monohydrate and 60 mL water)
glucose pulse was injected aseptically through a rubber
septum, and samples were removed periodically for analysis.
Viability
Viability of RWB 850-2 was determined by plating appropriate dilutions (prepared in sterile 0.1% peptone water) on
synthetic medium containing ethanol as the sole carbon
source (as described above) and determining the CFUs.
Triplicate plates containing between 20 and 100 colonies
were counted for each of the duplicate fermentations. The
number of CFUs present during the chemostat phase (t = 0)
was taken as 100% viability and successive samples were
reported as a percentage of that value.
duplicates for two different concentrations of cell extracts.
Each in vitro enzyme assay for the glycolytic pathway was
performed as described previously (Jansen et al., 2005).
Protein concentrations in cell-free extracts were determined
by the Lowry method (Lowry et al., 1951) with dried bovine
serum albumin (fatty acid free, Sigma) as a standard.
Determination of intracellular storage
carbohydrates and adenosine phosphates
Samples for all other intracellular metabolite determinations
were obtained by withdrawing 1 mL of broth from the
fermenter by a rapid sampling setup (Lange et al., 2001)
into 5 mL of 60% (v/v) methanol/water at 40 1C to
immediately quench the metabolic activities. The sample
was then processed according to the intracellular sampling
processing method described by Wu et al. (2005) to give
about 500 mL of intracellular metabolite solution.
Intracellular ATP concentrations were determined using
the ATP Bioluminescence Assay Kit CLS II (Roche Diagnostics GmbH, No. 1699 695) according to the manufacturer’s
instructions in black Costar 96-well microtiter plates. Luminescence was read on Mediators PhL plate reader (Mediators
Diagnostics GmbH, Vienna, Austria). Concentrations of
intracellular ADP and AMP were determined enzymatically
according to Mashego et al. (2005) based on myokinase,
pyruvate kinase and lactate dehydrogenase. The energy
charge (EC) (Atkinson, 1968) was calculated as shown in
Eqn. (1). An EC of 1 indicates that all adenosine phosphate
is present as ATP, whereas an EC of 0 would indicate that all
adenosine phosphate is present as AMP.
In vitro enzymatic assays
Frozen cell-free extracts ( 20 1C) were thawed at room
temperature, washed and resuspended in 2 mL ice-cold
sonication buffer (100 mM KPB, 2 mM MgCl2, pH 7.5) with
1 mM 1,4-dithiothreitol final concentration. One milliliter
of cell suspension was added to safelock tubes with 0.75 g of
cold glass beads (Sigma, G8772). The tubes were placed in a
FastPrep120A machine (Thermo Scientific) and shaken in
four bursts of 20 s, at speed 6 (6.0 m s1), for efficient
breakdown of the cell membrane. Samples were placed on
ice between bursts. The unbroken cells and debris were
removed by 20 min centrifugation at 47 000 g, 4 1C. The
supernatant was stored on ice and used for determination of
enzyme activities.
Enzyme activities in freshly prepared cell-free extracts
were analyzed via spectrophotometric enzyme-linked assays,
using a TECAN GENios Pro microtiter plate reader. All
determinations were performed at 30 1C and 340 nm
(eNAD(P)H at 340 nm = 6.33 mM1). Samples were prepared
manually in microtiter plates (transparent flat bottom
Costarplate, 96 wells) with a total volume of 300 mL per
well. To assure reproducibility, all assays were performed in
FEMS Yeast Res 9 (2009) 349–357
EC ¼
½ATP þ 0:5½ADP
½ATP þ ½ADP þ ½AMP
ð1Þ
Trehalose and glycogen measurements were performed as
described previously (Parrou & Francois, 1997) in duplicate
measurements on two independent replicate cultures. Glucose was determined using the UV-method based on the
BioControl EnzyPlusTM kit (No. EZS781).
Results
Dynamic responses to anaerobic glucose excess
Aerobic steady-state carbon-limited chemostat cultures of
the reference strain, CEN.PK 113-7D, and the isogenic
lactate-producing strain, RWB 850-2, were used to obtain
well defined, reproducible and comparable starting conditions for the study of the dynamic response to anaerobic
glucose excess. Consistent with previous work (van Maris
et al., 2004b), growth of both strains on the mixture of 80%
glucose and 20% ethanol (on carbon basis) was fully
respiratory at the dilution rate of 0.10 h1 as indicated by
the respiratory quotient of 0.88 0.04, the lack of
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352
D.A. Abbott et al.
lactate-producing strain was 91 13% and never dropped
below 75% during the first 6 h after the addition of glucose.
Viability was not measured for the reference strain.
fermentation products and the high biomass yields of c.
14.7 g biomass mol1 carbon. The culture dry weights in
these steady-state cultures were identical with values of
3.68 0.05 g L1 for RWB 850-2 and 3.64 0.03 g L1 for
CEN.PK 113-7D.
In contrast to the almost identical behavior of both
strains in aerobic chemostat cultures, large differences
became apparent shortly after introduction of anaerobicity
and glucose excess (200 mM). The growth of the reference
strain (as measured by OD660 nm) began c. 1–2 h after the
pulse and continued for several hours (Fig. 1). In sharp
contrast, the biomass concentration of RWB 850-2 had not
increased after 8 h. In agreement with previous research (van
Maris et al., 2004b), growth of the reference strain was
accompanied by formation of ethanol, carbon dioxide and
glycerol with complete glucose consumption occurring after
c. 4 h. Although lactic acid production (excluding export) is
theoretically equal to ethanol formation with respect to ATP
generation and redox metabolism, the glucose consumption
rate in the lactic acid-producing strain was lower and even
more importantly, the rate of lactate production continually
decreased over time (Fig. 1). As a result, only 5% of the
glucose was consumed within 8 h. The average viability (as
measured by CFUs) of duplicate fermentations with the
Changes in adenosine phosphates
To study this dramatic difference in response to anaerobic
glucose excess and to determine the possible impact of
lactate export on the cellular energetics, intracellular levels
of ATP, ADP and AMP were measured at the aerobic steady
state and throughout the dynamic anaerobic phase (Fig. 2).
During aerobic, carbon-limited growth, the levels of intracellular ATP were identical for each strain at
7.0 mmol g1 biomass (Fig. 2, t = 0). In these steady states,
the energy charge of both strains had a value of c. 0.8, which
is normal for these conditions (Ball & Atkinson, 1975). Five
minutes after the shift to glucose-excess conditions both
strains showed a drastic decrease in the ATP concentration
to 3.6 mmol g1 biomass for the reference strain and an even
lower 2.4 mmol g1 biomass for the lactic acid-producing
strain. However, the ATP concentration in the reference
strain quickly recovered to a new pseudo-steady state, while
ATP levels in the RWB 850-2 strain decreased for the first
30 min of the pulse experiment, subsequently leveled of at
Fig. 1. Lactate (m) or ethanol (’) production and glucose consumption ( ; left panel) for aerobically grown chemostat (t = 0) cultures of RWB 850-2
(top row) and CEN.PK 113-7D (bottom row) after exposure to 200 mM glucose and anaerobicity. The specific rate of product formation (&) and the
OD660 nm ( ) are also indicated (right panel). The average data of two independent fermentations is presented.
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FEMS Yeast Res 9 (2009) 349–357
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Energetic constraints in yeast homolactate fermentation
Fig. 2. Intracellular concentrations of adenosine
phosphates (ATP, ADP and AMP), sum of
nucleotide concentrations (AXP) and energy
charge of the reference (closed circles) and RWB
850-2 (open circles) strain after the shift to
anaerobic glucose-excess conditions. Steadystate concentrations for the chemostat phase are
indicated as t = 0. Error bars of the adenosine
concentrations represent SDs between duplicate
analyses of two independent culture samples.
the low concentration of 1.0 mmol g1 biomass and did not
recover within 2 h (Fig. 2).
When ATP concentrations decrease, the concentrations of
the other two forms of the adenosine phosphate conserved
moieties are expected to increase. Indeed, although the
intracellular concentrations of ADP were almost constant
for both strains, the response of the AMP concentrations to
the glucose pulse was strikingly different in both strains.
Whereas the AMP level for the reference strain remained
constant, the AMP concentration in the RWB 850-2 strain
increased 1.5-fold over the first 15 min after the shift and
remained at this elevated level. The fluctuations of the ATP
concentration on both the reference strain and RWB 850-2
were not totally accounted for by the change in ADP and
AMP, as shown by the changes in total adenosine phosphates
(AXP, Fig. 2). This decrease in AXP has been described
previously (Kresnowati et al., 2006). As a result of these
concentration changes, the energy charge for the reference
strain decreased the first 5 min after the shift before it
recovered to the initial steady-state value of 0.8. In contrast,
the energy charge of the RWB 850-2 strain mirrored the
drop in ATP concentration and decreased to the extremely
FEMS Yeast Res 9 (2009) 349–357
low value of 0.4, thereby demonstrating energetic differences
between alcoholic fermentation and homolactic fermentation in S. cerevisiae.
Storage carbohydrates
The levels of storage carbohydrates are not only known to
vary strongly with cultivation conditions (Parrou et al.,
1997; François & Parrou, 2001; Guillou et al., 2004), but
their consumption can, in theory, also provide the lactic
acid-producing cells with (additional) ATP if lactic acid
production indeed fails to replenish the ATP pools due to
export energetics. As expected, the levels of the storage
carbohydrates, trehalose and glycogen, were approximately
equal in aerobic chemostats for both the reference strain and
the lactic acid-producing strain. However, the dynamic
responses to the anaerobic glucose excess conditions were
dramatically different. As previously observed (van den
Brink et al., 2008b), both the glycogen and trehalose
concentrations in the reference strain decreased quickly
from 1.4% and 2.4% of the dry weight (w/w) in the steady
state to c. 0.25% of the dry weight (w/w) within the first 2 h
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354
(Fig. 3). Surprisingly, these storage carbohydrates were not
mobilized in RWB 850-2 throughout the experiment (Fig.
3). Therefore, the initially higher rate of lactic acid production was not explained by additional ATP provided by
consumption of glycogen.
Glycolytic enzyme activities
The activity of several glycolytic enzymes was measured for
changes in maximal velocity (Vmax) to determine the
influence of anaerobic lactate production on central carbon
metabolism. Predictably, pyruvate decarboxylase activity
was absent in RWB 850-2 and lactate dehydrogenase was
absent in the reference strain. In the reference strain,
substantial increases in activity were observed for the
majority of the measured glycolytic enzymes, including the
activities of phosphoglucose isomerase, fructose-bisphosphate aldolase, triose-phosphate isomerase, glyceraldehyde-3-phosphate dehydrogenase, phosphoglycerate kinase,
phosphoglycerate mutase and pyruvate kinase (Fig. 4). The
increased activity of these glycolytic enzymes has been well
documented and is directly correlated to changes in transcript levels (van den Brink et al., 2008a, b). In contrast to
both the observations and documented response of wildtype S. cerevisiae strains, the enzyme activities were unchanged in RWB 850-2 (Fig. 4). The stable activity of these
enzymes in anaerobic lactate-producing cultures suggests a
problem with de novo protein synthesis or alternatively a
drastic change in protein turnover.
Interestingly, activities of hexokinase (HXK), phosphofructokinase (PFK) and alcohol dehydrogenase (ADH),
remained unchanged or only decreased slightly in both
Fig. 3. Comparison of glycogen (circles) and trehalose (squares) levels in
RWB 850-2 (open symbols) and CEN.PK 113-7D (closed symbols) in
response to an anaerobic pulse of 200 mM glucose to aerobic chemostat
cultivations (t = 0). Concentrations of each storage carbohydrate are
expressed as percentage of total dry weight (w/w). Error bars represent
SDs between duplicate analyses of two independent culture samples.
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D.A. Abbott et al.
RWB 850-2 and the reference strain. For PFK, notorious
for its sensitivity to many allosteric effectors, such as
fructose-2,6-biphosphate and ATP (Bartrons et al., 1982),
relating in vitro activities to in vivo fluxes is precarious. The
activity of both HXK and ADH was already lower during the
steady states, even though the physiological parameters
under steady-state conditions were similar between the
strains. This might indicate different ratios of the HXK and
ADH iso-enzymes in each strain even under nonfermentative conditions (van den Brink et al., 2008a).
Discussion
The observed rapid decrease of the intracellular ATP concentration within the first few minutes after the anaerobic
glucose pulse to homolactic S. cerevisiae, combined with
sustained high cellular viabilities ( 4 75%) throughout the
experiment support the hypothesis that lactic acid production does not yield ATP (van Maris et al., 2004a, b). As
transcription and especially translation are energetically
expensive processes (Warner, 1999), the depletion of the
ATP concentration may also hinder protein synthesis. This
conclusion is supported by the absence of increased glycolytic enzyme activities in RWB 850-2, and resulted in reduced
metabolic rates, which further amplified the energetic constraints in these lactate-producing cultures. These observations are consistent with previous publications, which
hypothesized that lactic-acid production in this engineered
S. cerevisiae strain does not yield (net) ATP, due to an ATP
requirement for export of lactic acid (van Maris et al.,
2004a).
Alternative explanations for the decreased lactic-acid
production rate include cytosolic acidification (Valli et al.,
2006), intracellular accumulation of organic acid anions
(Pampulha & Loureiro-Dias, 1990) or extensive remodeling
of the plasma membrane and cell wall upon alteration of
culture aeration (Nurminen et al., 1975; Klis et al., 2002;
Kwast et al., 2002; Snoek & Steensma, 2007). However, the
increased rates of lactic acid production in aerobic conditions, where respiration can provide ATP required for
growth and maintenance, and the short time scale (minutes)
of the experiments appear to contradict these hypotheses.
Storage carbohydrates are known as easily accessible
forms of carbon and energy for use during shifts in
metabolism (Silljé et al., 1999; Enjalbert et al., 2000; Guillou
et al., 2004). As expected from the post-transcriptional
activation of trehalose and glycogen phosphorylases by the
Protein Kinase A pathway upon sensing of the high glucose
levels (as reviewed by François & Parrou, 2001), the reference strain rapidly consumed these compounds after the
perturbation. It was therefore surprising, especially in view
of glycogen as a possible additional source of ATP, that the
reserve carbohydrates were not mobilized by the lactic acidFEMS Yeast Res 9 (2009) 349–357
355
Energetic constraints in yeast homolactate fermentation
Fig. 4. In vitro glycolytic enzyme activity in
CEN.PK 113-7D (closed symbols) and RWB 850-2
(open symbols) in response to an anaerobic
glucose pulse. Steady-state chemostat values are
indicated as t = 0. Error bars represent SDs
between duplicate analyses of two independent
culture samples.
producing strain. Putatively, the rapid decrease of the ATPlevels in the cell prevented activation and/or synthesis of the
enzymes required for the mobilization. Alternatively, the
protective role of glycogen and trehalose (Parrou et al.,
FEMS Yeast Res 9 (2009) 349–357
1997) might inhibit their utilization in order to maintain
stores of these protective compounds.
Although these experiments produced irrefutable
evidence that anaerobic homolactate fermentation in
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356
S. cerevisiae RWB 850-2 results in ATP depletion, the exact
underlying cause of this ‘energy limitation’ has not been
established. Detailed studies of the transport mechanism of
lactic acid, intracellular pH, proton-motive force and/or the
intracellular total lactic acid concentrations, which are
difficult to measure due to high background concentrations
in the broth, are required to solve the enigma of anaerobic
homolactic fermentation by S. cerevisiae.
Acknowledgements
The authors acknowledge Marijke Luttik for excellent technical assistance and thank Tate & Lyle Ingredients Americas
Inc. for financial support. The Kluyver Center for Genomics
of Industrial Fermentation is supported by the Netherlands
Genomics Initiative.
Authors’contribution
D.A.A. and J.v.d.B. contributed equally to this study.
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