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1 Kluyver Centre for Genomics of Industrial Fermentation and Department of Biotechnology, Delft University of Technology, Julianalaan 67, 2628 BC Delft, The Netherlands
2 DSM Food Specialties, PO Box 1, 2600 MA Delft, The Netherlands
Correspondence
P. Daran-Lapujade
p.a.s.daran-lapujade{at}tudelft.nl
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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In the case of galactose, a four-enzyme metabolic route, the Leloir pathway, has to be expressed to enable its catabolism. The structural genes for galactose permease (GAL2), galactokinase (GAL1), galactose-1-phosphate uridylyltransferase (GAL7) and uridine-diphosphoglucose 4-epimerase (GAL10) all belong to the GAL regulon, which is subject to tight transcriptional regulation. Glucose causes a nearly complete transcriptional repression of the GAL genes, mediated by the non-phosphorylated form of Mig1, and thereby effectively shuts down galactose utilization (Johnston et al., 1994
). Induction of the GAL genes is initiated by interaction of galactose and ATP with Gal3, which then forms a complex with the negative regulator Gal80 (Platt & Reece, 1998). This releases the positive transcriptional regulator Gal4 from Gal80 control and allows it to activate transcription of the GAL1, GAL2, GAL7 and GAL10 genes, which contain upstream activation sequences (UASGAL) in their promoter regions (Leuther & Johnston, 1992; Wu et al., 1996). Consequently, S. cerevisiae cells grown in batch cultures on glucose/galactose mixtures show diauxic utilization of these sugars (Ostergaard et al., 2001b; Raamsdonk et al., 2001).
To investigate the adaptive mechanisms involved in the transition between glucose and galactose metabolism, we used well-controlled, glucose-limited chemostat cultures that were switched to galactose-excess conditions. Unexpectedly, the first results showed that, under anaerobic conditions, S. cerevisiae could not switch from glucose-limited growth to galactose utilization. Still, S. cerevisiae does not exhibit a Kluyver effect for galactose and is therefore able to ferment galactose in the absence of oxygen (Fukuhara, 2003) [the Kluyver effect has been defined as the inability to grow on and ferment sugars under anaerobic conditions that do support aerobic, respiratory growth (Barnett et al., 1990)]. This unexpected behaviour could reflect different metabolic problems: (i) lack of transcriptional induction of the GAL regulon, (ii) absence of Gal protein synthesis, (iii) accumulation or depletion of low-molecular-mass metabolites and effectors that affect in vivo activity of the Leloir pathway. For example, the glucose-to-galactose switch might cause an impaired flux through the Leloir pathway by insufficient substrate levels for galactokinase (e.g. ATP) (Ostergaard et al., 2001a) or through metabolite toxicity (e.g. galactose 1-phosphate) (de Jongh et al., 2008).
The goal of the present study was to elucidate the cause of the observed impaired galactose consumption after a switch from anaerobic, glucose-limited chemostat cultivation to anaerobic, galactose-excess conditions. To dissect the involvement of different levels of cellular information and regulation, we analysed the dynamics of glucose–galactose transitions via a multi-level approach.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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), a wild-type strain CBS8066 (van Dijken et al., 2000), a hxk2 : : KanMX4 mutant (Diderich et al., 2002), a reg1 : : KanMX4 mutant (Raghevendran et al., 2004) and a rip1 : : KanMX4 mutant (this study). All three mutant strains were derived from the CEN.PK113 background. Stock cultures were grown at 30 °C in shake flasks containing 100 ml synthetic medium with 20 g glucose per litre. The synthetic medium contained, per litre of demineralized water, 5 g (NH4)2SO4, 3 g KH2PO4, 0.5 g MgSO4.7H2O, 0.15 ml silicon antifoam (BDH) and trace element concentrations according to Verduyn et al. (1992). After heat sterilization of the medium for 30 min at 120 °C, a filter-sterilized vitamin solution (Verduyn et al., 1992) was added. Ergosterol (10 mg l–1) and Tween 80 (420 mg l–1) for the anaerobic cultivation were dissolved in pure ethanol and steamed at 80 °C before addition to the medium. The concentration of glucose in the reservoir medium was 7.5 g l–1 or 25 g l–1, respectively, for the aerobic and anaerobic cultivation. Glucose was added to the synthetic medium after separate heat sterilization at 110 °C.
Chemostat cultivation and perturbation experiment.
The S. cerevisiae strains were grown at 30 °C in 2 litre bioreactors (Applikon) with a working volume of 1.5 litres that was controlled via an electrical level sensor. Removal of effluent from the centre of the culture ensured that biomass concentrations in the effluent line differed by less than 1 % from those in the culture (van den Berg et al., 1996). The dilution rate was set at 0.10 h–1. The pH was measured online and kept constant 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 keep the dissolved oxygen concentration, as measured with an oxygen electrode, above 60 % of air saturation in all aerobic chemostat cultivations. In the anaerobic cultivation, the bioreactor and medium vessel were sparged with pure nitrogen gas (<5 p.p.m. O2; Linde Gas Benelux) and Norprene tubing was used to minimize oxygen diffusion into the fermenters. Steady-state samples were taken after 
10 volume changes to avoid strain adaptation due to long-term cultivation (Ferea et al., 1999; Jansen et al., 2004; Mashego et al., 2005). Biomass dry weight, extracellular metabolite, dissolved oxygen and gas profiles were constant over at least three volume changes.
Galactose-pulse experiments were started by simultaneously switching the medium pump off and injecting the galactose. The pulse consisted of 60 g galactose in 60 ml demineralized water and was injected aseptically through a septum. The galactose concentration immediately after the pulse was approximately 200 mM.
Analytical methods.
The exhaust gas was cooled by a condenser connected to a cryostat set at 2 °C and dried with a Perma Pure dryer (Inacom Instruments) before analysis of the O2 and CO2 concentrations with a Rosemount NGA 2000 analyser. The gas flow rate was determined with an Ion Science Saga digital flow meter. Culture dry weights were determined as described by Postma et al. (1989). Ethanol, galactose, glycerol and glucose concentrations in supernatants were determined by HPLC analysis with a Bio-Rad Aminex HPX-87H column at 60 °C. The column was eluted with 5 mM sulfuric acid at a flow rate of 0.6 ml min–1. All relevant metabolites were detected by a Waters 2410 refractive index detector.
Intracellular metabolites.
Samples for intracellular metabolite analysis were obtained by withdrawing 1 ml broth from the fermenter via a rapid sampling set-up (Lange et al., 2001) into 5 ml 60 % (v/v) methanol/water at –40 °C to immediately quench all metabolic activities. The samples were then processed according to a previously published intracellular sampling processing method (Wu et al., 2005) to give 0.5 ml intracellular metabolite solution ready for further analysis.
Intracellular concentrations of galactose 1-phosphate, glucose 1-phosphate and glucose 6-phosphate were measured enzymically according to Bergmeyer & Michal (1974). The assay mixture contained 100 mM glycylglycine (pH 8.7), 10 mM MgCl2, 0.01 mM glucose 1,6-biphosphate, 0.2 mM NADP+, and sample. The reaction to measure glucose 6-phosphate was started with 17 mU glucose-6-phosphate dehydrogenase. The reaction to measure glucose-1-phosphate was started with 7 mU phosphoglucomutase. Finally, the reaction to measure galactose 1-phosphate was started with 0.45 mU galactose-1-phosphate uridylyltransferase.
Intracellular concentrations of UDP-glucose and UDP-galactose were measured enzymically according to Bergmeyer & Michal (1974). The assay mixture contained 100 mM glycylglycine (pH 8.7), 10 mM MgSO4.7H2O and 0.25 mM NAD+. The reaction to measure UDP-glucose was started with UDP-glucose-6-phosphate dehydrogenase (0.01 U ml–1). The reaction to measure UDP-galactose was started with galactose-1-phosphate uridylyltransferase (0.1 U ml–1, Sigma).
The assays were performed in black flat-bottomed Costar 96-well microtitre plates (Corning) by following NADH or NADPH fluorescence in a Tecan Genios microtitre plate reader. The excitation wavelength was 340 nm and the emission wavelength was 465 nm.
Quantitative real-time PCR.
Sampling of cells from the bioreactors and total RNA extraction were performed as described previously (Piper et al., 2002). Isolated RNA was DNase I treated according to the Qiagen RNeasy mini kit protocol for RNA cleanup. First-strand cDNA synthesis was carried out using 2 µg DNase-treated RNA, random hexamer primers and Moloney murine leukaemia virus reverse transcriptase (Invitrogen). Quantitative real-time PCR was run on a DNA Engine Opticon I system (Bio-Rad) with the following settings: 94 °C for 15 min, 94 °C for 10 s, 66 °C for 10 s, 72 °C for 10 s, and plate reading. The denaturation, annealing, elongation and reading steps were repeated for 39 cycles. A melting curve from 50 to 94 °C was generated at the end of the reaction. The 20 µl reaction mixture consisted of 10 µl Absolute QPCR SYBR green mix (Westburg; cat. no. AB-1158), 0.2 µM forward primer, 0.2 µM reverse primer, and cDNA (20, 2, 0.2 or 0.02 ng per PCR). The primers used are listed in Table 1. The cycle threshold value was calculated with the Opticon Monitor software version 1.08 (Bio-Rad). Each PCR was carried out in triplicate. The relative expression of GAL1, GAL2, GAL7, GAL10 and PGM2 was quantified by comparing (
) with ACT1 as the reference gene. The results for the steady state and each time point after the perturbation (5, 10, 15, 30 and 120 min) were derived from two independent replicate cultures.
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). Enzyme activities were assayed with freshly prepared cell extracts in spectrophotometric enzyme-linked assays, using a Tecan Genios microtitre plate reader. All determinations were performed at 30 °C and 340 nm (
NAD(P)H at 340 nm=6.33 mM–1). Samples were prepared manually in microtitre plates (transparent flat-bottom 96-well plates; Corning) in a 12-assay run with a total volume of 300 µl per well. All assays were performed with two concentrations of cell extracts; data are presented as U (mg protein)–1 (1 U=1 µmol min–1). Protein concentrations in cell-free extracts were determined by the Lowry method. Dried BSA (fatty-acid-free, Sigma) was used as a standard.
Galactokinase (EC 2.7.1.6) was assayed according to Platt et al. (2000) with minor modifications. The assay mixture contained 100 mM potassium phosphate buffer (pH 7.5), 100 mM NaCl, 5 mM MgCl2, 0.5 mM phosphoenolpyruvate, 1 mM DTT, 0.15 mM NADH, 5 mM ATP, 2.2 units pyruvate kinase, 3.0 units lactate dehydrogenase (PK/LDH; Sigma) and cell extract. The reaction was started with 5 mM D-galactose.
Galactose-1-phosphate uridylyltransferase (EC 2.7.7.12) was assayed according to Colowick & Kaplan (1967) with minor modifications. The assay mixture contained 100 mM glycine/KOH (pH 8.5), 10 mM L-cysteine, 5 mM MgCl2, 0.4 mM NADP+, 40 mM galactose 1-phosphate, 0.01 mM glucose 1,6-biphosphate, 1.6 units phosphoglucomutase, 1.8 units glucose-6-phosphate dehydrogenase and cell extract. The reaction was started with 2 mM UDP-D-glucose.
UDP-galactose 4-epimerase (EC 5.1.3.2) was assayed according to Majumdar et al. (2004) with minor modifications. The assay mixture contained 100 mM glycylglycine (pH 8.0), 0.40 mM NAD, 0.16 units UDP-glucose-6-dehydrogenase and cell extract. The reaction was started with 0.4 mM UDP-D-galactose.
Phosphoglucomutase (EC 5.4.2.2) was assayed according to Masuda et al. (2001) with minor modifications. The assay mixture contained 50 mM Tris/HCl (pH 7.5), 5 mM MgCl2, 1 mM NADP+, 0.01 mM glucose 1,6-biphosphate, 3.6 units glucose-6-dehydrogenase and cell extract. The reaction was started with 4 mM glucose 1-phosphate.
Proteomics.
Samples were taken 180 min after the galactose pulse from two independent cultures under aerobic and anaerobic conditions. An equal volume of 20 % trichloroacetic acid was added to 4 ml culture sample (10–14 mg cells). After 30 min incubation on ice, the samples were centrifuged at 4 °C and washed with –20 °C acetone. The samples were centrifuged for 10 min (5 min at 4600 g, 4 °C) and the remaining acetone was evaporated. The washed pellets were suspended in 500 µl 100 mM ammonium bicarbonate pH 7.8 and 20 µl 250 µg ml–1 TPCK-treated trypsin in 0.025 % HCl (pH 3). Proteins were digested overnight by incubation at 37 °C in a Thermomixer. Then 6 µl 100 mM 1,4-DTT was added to the digests and incubated for 30 min at room temperature. Samples were acidified with 5 % formic acid (FA) and any insolubles were removed by centrifugation (5 min at 4600 g, 4 °C). The supernatant was analysed by LC-MS/MS.
Peptides were separated and analysed using an Accela high-performance liquid chromatograph coupled to an LTQ linear iontrap mass spectrometer. LC-MS/MS was performed using a gradient of buffer A (LC-MS-grade water containing 0.1 % FA) and buffer B (LC-MS-grade acetonitrile containing 0.1 % FA). A 5 µl sample of the peptide solution was trapped on an NS-MP-10-Robust pre-column (OD/ID 360/100 mm, Poros 10R2 and C18 5 µm particles 120 Å pores; NanoSeparations) at 5 % B with a flow rate of 15 µl min–1 for 5 min. Peptides were eluted from the pre-column onto an analytical column (OD/ID 360/75 µm, Biosphere C18 3 µm particles 120 Å pores, length 23 cm; NanoSeparations) and to the MS using a switching valve (Meiring et al., 2002). The gradient was run at 200 nl min–1 from (i) 5.01 min 15 % B to 60 min 25 % B, (ii) 60 min 25 % B to 155 min 35 % B.
An Nth-order triple-play MS-method was used in the LTQ. This method consisted of three scan events: (i) an MS scan from m/z 300 to 2000, (ii) data-dependent zoom scans on the top three peaks from the first scan event, (iii) data-dependent MS/MS scans on the top three peaks from the first scan event. The acquired data were searched against the Saccharomyces Genome Database protein database (http://www.yeastgenome.org/), using Sequest in the Bioworks software (version 3.3.1; Thermo Fisher Scientific). Identified peptides were filtered for SF final score >0.85 and Peptide Probability <0.001. Peptide and protein areas were calculated using PepQuan in the Bioworks software.
Adenosine phosphates.
Intracellular ATP was determined using the ATP Bioluminescence Assay kit CLS II (Roche Diagnostics; cat. no. 1699 695), according to the manufacturer's instructions, in black Costar 96-well microtitre plates. Luminescence was read on a Mediators PhL plate reader (Mediators Diagnostics). Intracellular ADP and AMP were determined enzymically according to Mashego et al. (2005) based on myokinase, pyruvate kinase and lactate dehydrogenase (Bergmeyer et al., 1985).
The energy charge (EC) (Atkinson, 1968) was calculated as follows:
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Trehalose and glycogen.
Trehalose and glycogen concentration measurements were performed as described by Parrou & Francois (1997) in duplicate measurements on two independent replicate cultures. Glucose was determined using the UV method based on Roche kit no. 0716251.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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; Martinez-Pastor & Estruch, 1996; Nilsson et al., 2001). This processing, during which biomass is temporarily starved and potentially deprived of (or exposed to) oxygen, may significantly affect experimental reproducibility and responses of the cells to substrate transitions. In the present study, S. cerevisiae CEN.PK113-7D was grown in glucose-limited chemostat cultures. Subsequently, excess galactose (200 mM) was injected into the cultures while concomitantly stopping the glucose feed. This set-up is excellently suited for studying substrate transitions, because a different carbon source can be added directly into the fermenter without prior processing of the biomass. Another advantage of this set-up is the alleviation of glucose repression. Glucose exerts a strong catabolite repression on the utilization of alternative carbon sources (e.g. maltose, ethanol and galactose) (Gancedo, 1998). During shifts starting with glucose excess the transcriptional repression of the genes involved in alternative carbon source utilization first has to be relieved. In glucose-limited chemostats, the low residual glucose concentration (below 0.17 mM at D=0.10 h–1 in aerobic cultures) causes glucose repression to be alleviated already at the onset of the experiment.
When S. cerevisiae was grown under glucose limitation in the presence of oxygen, galactose metabolism was induced as expected and was detectable 4 h after galactose addition (Fig. 1). The length of this adaptation phase was consistent with previously reported data for aerobic batch cultures grown on glucose/galactose mixtures (Ostergaard et al., 2001b; Raamsdonk et al., 2001). The high initial galactose concentration made it difficult to measure the exact time point at which galactose consumption set in. Measurements of intracellular intermediates of the Leloir pathway demonstrated that galactose metabolism started much earlier than the 4 h suggested by the extracellular galactose measurements. Concentrations of galactose 1-phosphate, glucose 1-phosphate and UDP-galactose were already increased 60 min after galactose addition (Fig. 2). Conversely, intracellular concentrations of glucose 6-phosphate and UDP-glucose rapidly decreased within the first 15 min after the shift from glucose limitation to galactose excess. The decrease of glucose 6-phosphate and UDP-glucose concentration may reflect the role of UDP-glucose in storage carbohydrate metabolism and that of glucose 6-phosphate is as an intermediate of the glycolytic pathway, which was temporarily interrupted after the shift. After this initial decrease, the intracellular glucose 6-phosphate and UDP-glucose concentrations increased and followed the increase in flux through the Leloir pathway.
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), even several days after the shift (data not shown). The inability of S. cerevisiae to consume galactose under these conditions was also observed with other laboratory strains (e.g. CBS8066), confirming that the phenomenon was not strain-specific (data not shown). To investigate whether low rates of galactose consumption might have gone unnoticed due to the relative insensitivity of the extracellular galactose measurements, intracellular concentrations of the intermediates of the Leloir pathway were measured (Fig. 2). The intracellular concentrations of UDP-glucose and glucose 6-phosphate, metabolites already present when glucose is the carbon source, rapidly decreased after glucose exhaustion and were undetectable 2 h after the shift from glucose to galactose. Glucose 1-phosphate concentration did not show significant changes and remained close to the detection limit. Although the noise of the enzymic assays was largely due to the low concentrations, galactose 1-phosphate concentration initially increased, possibly reflecting non-specific kinase activity. UDP-galactose, one of the two galactose-specific intermediates, could not be detected after galactose addition, suggesting that this metabolite was not synthesized under anaerobiosis. The apparent inability to induce galactose metabolism in the anaerobic experiments was fully reversible, since sparging the cultures with air resulted in galactose consumption (Fig. 1). Galactose consumption upon oxygenation of the anaerobic cultures was presumably enabled by the ATP generated via ethanol consumption.
The absence of galactose consumption under anaerobiosis does not result from impaired transcription of the GAL genes
To investigate whether the absence of galactose consumption under anaerobic conditions could result from a deficiency in galactose signalling and induction of the GAL genes (Ostergaard et al., 2000), transcript levels of the five structural steps of the Leloir pathway were monitored by quantitative PCR (Fig. 3). In both the aerobic and the anaerobic glucose–galactose transitions, transcription of the four genes of the GAL regulon (GAL1, GAL2, GAL7 and GAL10) was immediately induced after the addition of galactose and increased in the 15 min following galactose addition before stabilizing. Under both conditions, the transcript levels of the GAL genes upon galactose induction were high (Fig. 3). However, they were about tenfold lower under anaerobiosis than under aerobiosis. Phosphoglucomutase, encoded by PGM2 (also called GAL5), catalyses the last step in the Leloir pathway, but is also involved in other processes unrelated to galactose metabolism (Francois & Parrou, 2001). As a result, PGM2 is not under the control of the GAL regulon (Bevan & Douglas, 1969) and, accordingly, its transcript profiles differed from those of the other GAL genes.
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The enzymes involved in galactose utilization are not synthesized upon a sudden shift from glucose to galactose in the absence of oxygen
The presence of the Leloir-pathway transcripts does not necessarily imply the presence of a functional pathway. Therefore, the Vmax (maximum enzyme capacity) of the Leloir-pathway enzymes was assayed in vitro (Fig. 4). In good correlation with its corresponding transcript, the enzyme activity of Pgm2 was already present during the glucose-limited cultivations and remained throughout the aerobic and anaerobic glucose–galactose transitions. Aerobically, in vitro enzyme activities for all Leloir-pathway enzymes under control of the GAL system (Gal1, Gal7 and Gal10) were undetectable during glucose-limited growth and slowly increased after the galactose pulse. Conversely, in the absence of oxygen the enzyme activities for Gal1, Gal7 and Gal10 remained undetectable throughout the experiment.
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Lack of GAL protein does not result from active translation inhibition
This absence of protein synthesis following shifts of growing cells from glucose to alternative carbon sources has previously been ascribed to an active, though temporary, inhibition of translation (Ashe et al., 2000; Kuhn et al., 2001; Martinez-Pastor & Estruch, 1996; Uesono et al., 2004). The resistance to translational inhibition upon glucose depletion of reg1
, hxk2 and snf3rgt2 mutants showed that several glucose signalling pathways were involved (Ashe et al., 2000). Our experimental set-up differed significantly from that used in these previous studies. Although glucose was the initial carbon source, its concentration in our study was too low to exert significant carbon catabolite repression. Furthermore, previous reports of glucose–raffinose switches involving translational inhibition showed very long lag phases of 20 h or more before S. cerevisiae started to consume raffinose (Martinez-Pastor & Estruch, 1996). Our intracellular metabolite measurements revealed that in aerobic cultures, galactose utilization started after 15 min of exposure to galactose-excess conditions. This time frame was consistent with reported transcription and translation rates and suggested that translation inhibition was not involved. Moreover, galactose consumption did not occur faster in experiments where the glucose feed was not switched off upon galactose addition (data not shown). These results suggested that translation inhibition did not occur in our experimental set-up when oxygen was supplied. In glucose-limited chemostat cultures, the residual glucose concentration, although very small, is typically slightly higher in the absence than in the presence of oxygen [0.17 mM aerobically versus 0.43 mM anaerobically (van Hoek et al., 2000)], which might result in a somewhat stronger catabolite repression anaerobically. Since, therefore, occurrence of active translational inhibition under anaerobic conditions could not be ruled out, reg1 and hxk2 mutants were grown in an anaerobic glucose-limited chemostat and then exposed to galactose excess. Neither reg1 nor hxk2 grew after the switch from glucose to galactose, arguing against involvement of active translational inhibition by glucose depletion.
The inability of S. cerevisiae to switch from glucose to galactose utilization anaerobically is caused by ATP shortage
The mode of sugar dissimilation in S. cerevisiae and, consequently, its ATP yield is strongly affected by the availability of oxygen. In the presence of oxygen, ATP can be formed via both substrate-level and oxidative phosphorylation, leading to the production of up to 16 moles of ATP per mole of hexose sugar [with a P/O ratio of 1.0 (Verduyn et al., 1991)]. In the absence of oxygen, substrate-level phosphorylation is the only mode of ATP generation, resulting in only 2 ATP per sugar molecule. Compared to aerobic respiration, the anaerobic fermentative metabolism therefore has a considerably lower energy yield per sugar molecule. To assess the energetic state of the cells the intracellular concentrations of adenosine phosphates were measured (Fig. 5). In the aerobic experiments, the ATP concentration initially decreased by 30 % (from 7.0 to 5.0 µmol gdry weight–1) and then stabilized. Under anaerobic conditions, the ATP concentration continued to decrease and reached undetectable levels 1 h after the glucose-to-galactose switch. The concentrations of ADP and AMP slightly increased in the aerobic experiments while, under anaerobic conditions, the concentration of ADP transiently increased and the AMP concentration increased threefold after the glucose-to-galactose switch. Consistently, the energy charge did not change significantly during the aerobic switch experiment but decreased dramatically in the anaerobic experiment.
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The mode of glucose dissimilation is not the only difference between aerobically and anaerobically grown cells. For instance, it is well documented that lipid metabolism, membrane and cell wall composition differ between aerobic and anaerobic cultures (Snoek & Steensma, 2006). To assess the importance of energetic status and other oxygen-related effects, a respiration-deficient strain was constructed and grown in the same experimental set-up as the congenic reference strain. rip1 strains, which carry a deletion in the structural gene for the Rieske Fe–S protein of the mitochondrial cytochrome bc1 complex, lack a functional respiratory chain (Beckmann et al., 1987). Even in the presence of oxygen, rip1 mutants rely solely on substrate-level phosphorylation for ATP synthesis, thus mimicking sugar dissimilation in anaerobically grown wild-type strains. The rip1 strain indeed failed to grow with non-fermentative carbon sources and displayed a marginal oxygen consumption rate with glucose as carbon source (details of the general physiology of the rip1 mutant are available as supplementary data with the online version of this paper). The rip1 strain was incapable of consuming galactose in aerobic substrate-switch experiments (Fig. 6). Moreover, the intracellular ATP concentration in the rip1 strain drastically decreased after the aerobic glucose-to-galactose switch (Fig. 6). These observations strongly support the notion that a shortage of ATP is the primary cause of the inability of S. cerevisiae to switch on galactose metabolism in the anaerobic experiments.
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). Especially the difference in trehalose concentration was striking: 30 mg gdry weight–1 under aerobiosis and 1 mg gdry weight–1 under anaerobiosis. However, neither glycogen nor trehalose was mobilized after the glucose-to-galactose switch under either aerobic or anaerobic conditions (Fig. 7). This observation indicates that other mechanisms play a role in maintaining the ATP level after the glucose-to-galactose switch under aerobic conditions. For example, consumption of minute amounts of galactose may be enough to synthesize a few Gal proteins and thereby start up galactose consumption. The absence of reserve carbohydrate consumption after the anaerobic glucose-to-galactose switch demonstrates that S. cerevisiae is ill equipped to handle sudden switches in carbon sources under anaerobic, fermentative conditions.
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) and the characteristic time of transcription is much shorter than that of translation (Stephanopoulos et al., 1998). Transcript levels of the GAL genes reached upon full induction were, however, 10-fold lower in the anaerobic substrate-switch experiments than in the aerobic experiments. These observations indicate that the collapse of the energy charge in the anaerobic cultures did not allow the same transcription rates as in the anaerobic cultures and completely prevented translation of the GAL mRNAs.
Catabolism of the storage carbohydrates present in the anaerobic, glucose-limited cultures could, in principle, yield ATP to energize the synthesis of Leloir-pathway enzymes. However, no such mobilization of glycogen occurred during the glucose–galactose transition. Apparently, the well-established mobilization of storage carbohydrates during the start-up of glycolysis (Thomsson et al., 2005; van den Brink et al., 2008) cannot be extrapolated to the activation of the Leloir pathway. The fact that, through evolution, S. cerevisiae did not acquire the ability to switch from anaerobic, glucose-limited growth to galactose utilization suggests that it seldom encounters this type of condition in its natural environments.
This study identifies the intracellular energy status of S. cerevisiae as a key factor in its metabolic flexibility, i.e. its ability to rapidly and functionally express novel pathways required for the utilization of alternative substrates upon the depletion of favoured substrates. In nature, energy shortage may occur not only as a result of a sudden depletion of the growth-limiting preferred substrate, as demonstrated in the chemostat set-up used in this study, but also by the presence of uncoupling agents under substrate-excess conditions. For example, it can be envisaged that weak organic acids have a similar negative impact on substrate transitions, since these uncouple the plasma membrane pH gradient of bakers' yeast and thereby negatively affect cellular energetics (Verduyn et al., 1991). This would be of direct relevance to the application of S. cerevisiae for the production of fuels and chemicals from lignocellulosic hydrolysates. These feedstocks contain complex mixtures of sugars (including galactose) but also high concentrations of acetic acid and other weak organic acids (Klinke et al., 2002; Palmqvist et al., 1999; Rabelo et al., 2008; Sun & Cheng, 2002). Research into the energetic aspects of metabolic flexibility and strategies to buffer cellular energy status [e.g. via engineering of ATP-buffering systems (Sauer & Schlattner, 2004)] is therefore not only of fundamental but also of applied significance.
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ACKNOWLEDGEMENTS |
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mutant and we thank Rintze Zelle, Nuno Tenazinha and Iris van Hoorn, who contributed to this work as part of their studies. This project was financially supported by the IOP Genomics programme of Senter Novem, The Netherlands. Edited by: M. Molina
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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Received 4 November 2008;
revised 17 December 2008;
accepted 19 December 2008.
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) and ethanol (
) after a galactose pulse to aerobic (left panel) and anaerobic (right panel) glucose-limited chemostat cultures of S. cerevisiae. The anaerobic cultures were shifted to aerobicity 10 h after the galactose pulse. Data points represent the average and mean-deviation of values from at least two independent cultures.
), GAL7 (
) are shown in the top panels and PGM2 (
) and AMP (
