OUP user menu

Anaerobic homolactate fermentation with Saccharomyces cerevisiae results in depletion of ATP and impaired metabolic activity

Derek A. Abbott, Joost van den Brink, Inge M.K. Minneboo, Jack T. Pronk, Antonius J.A. van Maris
DOI: http://dx.doi.org/10.1111/j.1567-1364.2009.00506.x 349-357 First published online: 1 May 2009


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.

  • lactate
  • lactic acid
  • energetics
  • enzyme activity
  • intracellular metabolite


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, 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, 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 S. cerevisiae strains capable of producing lactic acid with the limitation or elimination of ethanol formation (Adachi, 1998; Porro, 1999; van Maris, 2004b; Saitoh, 2005; Ishida, 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 NAD+ 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 was incapable of sustaining growth, and the lactate production rate rapidly decreased (van Maris, 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, 2004b).

It has been suggested that, whereas ethanol leaves the cell via passive diffusion (Guijarro & Lagunas, 1984), energy-dependent efflux of the proton and/or the lactate anion results in a net ATP requirement for lactic acid export (van Maris, 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, 1998; Fernandes, 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, 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, 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, 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 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, 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, 2004b). Cultures were grown at 30 °C in shake flasks containing 100 mL of synthetic medium (Verduyn, 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, 1992). Glucose (6.0 g L−1) was added to the synthetic medium after separate heat sterilization at 110 °C and 1.15 g L−1 of absolute ethanol was added separately. Each strain was grown at 30 °C 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 h−1, 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 min−1 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, 1999; Jansen, 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 <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 °C) steel beads (Mashego, 2003). Extracellular metabolite concentrations were analyzed via HPLC using an AMINEX HPX-87H ion exchange column with 5 mM H2SO4 as the mobile phase. Off-gas was first cooled with a condenser (2 °C) and then dried with a Perma Pure dryer (PD-625-12P). 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 min−1 of nitrogen gas (Hoek-Loos, Schiedam, <5 p.p.m. O2). Norprene tubing and butyl rubber septa were used to minimize oxygen diffusion into the anaerobic cultures (Visser, 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 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.

In vitro enzymatic assays

Frozen cell-free extracts (−20 °C) 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 s−1), 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 °C. 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 °C and 340 nm (ɛNAD(P)H at 340 nm=6.33 mM−1). Samples were prepared manually in microtiter plates (transparent flat bottom Costarplate, 96 wells) with a total volume of 300 μL per well. To assure reproducibility, all assays were performed in duplicates for two different concentrations of cell extracts. Each in vitro enzyme assay for the glycolytic pathway was performed as described previously (Jansen, 2005). Protein concentrations in cell-free extracts were determined by the Lowry method (Lowry, 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, 2001) into 5 mL of 60% (v/v) methanol/water at −40 °C to immediately quench the metabolic activities. The sample was then processed according to the intracellular sampling processing method described by Wu (2005) to give about 500 μL 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 (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.

Embedded Image 1

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 EnzyPlus kit (No. EZS781).


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, 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 h−1 as indicated by the respiratory quotient of 0.88±0.04, the lack of fermentation products and the high biomass yields of c. 14.7 g biomass mol−1 carbon. The culture dry weights in these steady-state cultures were identical with values of 3.68±0.05 g L−1 for RWB 850-2 and 3.64±0.03 g L−1 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, 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 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.

Figure 1

Lactate (▲) 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.

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 μmol g−1 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 μmol g−1 biomass for the reference strain and an even lower 2.4 μmol g−1 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 the low concentration of 1.0 μmol g−1 biomass and did not recover within 2 h (Fig. 2).

Figure 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. Steady-state 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.

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, 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 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, 1997; François & Parrou, 2001; Guillou, 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, 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 (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.

Figure 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.

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, 2008a, b). In contrast to both the observations and documented response of wild-type 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.

Figure 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.

Interestingly, activities of hexokinase (HXK), phosphofructokinase (PFK) and alcohol dehydrogenase (ADH), remained unchanged or only decreased slightly in both 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, 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, 2008a).


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 (>75%) throughout the experiment support the hypothesis that lactic acid production does not yield ATP (van Maris, 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, 2004a).

Alternative explanations for the decreased lactic-acid production rate include cytosolic acidification (Valli, 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, 1975; Klis, 2002; Kwast, 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, 1999; Enjalbert, 2000; Guillou, 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 acid-producing strain. Putatively, the rapid decrease of the ATP-levels in the cell prevented activation and/or synthesis of the enzymes required for the mobilization. Alternatively, the protective role of glycogen and trehalose (Parrou, 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 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.


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.


  • Editor: Monique Bolotin-Fukuhara


View Abstract