Accurate determination of intracellular metabolite levels requires well-validated procedures for sampling and sample treatment. Several methods exist for metabolite extraction, but the literature is contradictory regarding the adequacy and performance of each technique. Using a strictly quantitative approach, we have re-evaluated five methods (hot water, HW; boiling ethanol, BE; chloroform-methanol, CM; freezing-thawing in methanol, FTM; acidic acetonitrile-methanol, AANM) for the extraction of 44 intracellular metabolites (phosphorylated intermediates, amino acids, organic acids, nucleotides) from S. cerevisiae cells. Two culture modes were investigated (batch and chemostat) to check for growth condition dependency, and three targeted platforms were employed (two LC-MS and one GC/MS) to exclude analytical bias. Additionally, for the determination of metabolite recoveries, we applied a novel approach based on addition of (13)C-labeled internal standards at different stages of sample processing. We found that the choice of extraction method can drastically affect measured metabolite levels, to an extent that for some metabolites even the direction of changes between growth conditions can be inverted. The best performances, in terms of efficacy and metabolite recoveries, were achieved with BE and CM, which yielded nearly identical levels for the metabolites analyzed. According to our results, AANM performs poorly in yeast and FTM cannot be considered adequate as an extraction method, as it does not ensure inactivation of enzymatic activity.

Accurate determination of intracellular metabolite levels requires reliable, reproducible techniques for sampling and sample treatment. Quenching in 60% (v/v) methanol at -40°C is currently the standard method for sub-second arrest of metabolic activity in microbial metabolomics but there have been contradictory reports in the literature on whether leakage of metabolites from the cells occurs. We have re-evaluated this method in S. cerevisiae using a comprehensive, strictly quantitative approach. By determining the levels of a large range of metabolites in different sample fractions and establishing mass balances we could trace their fate during the quenching procedure and confirm that leakage of metabolites from yeast cells does occur during conventional cold methanol quenching, to such an extent that the levels of most metabolites have been previously underestimated by at least twofold. In addition, we found that the extent of leakage depends on the time of exposure, the temperature and the properties of the methanol solutions. Using the mass balance approach we could study the effect of different quenching conditions and demonstrate that leakage can be entirely prevented by quenching in pure methanol at B-40°C, which we propose as a new improved method. Making use of improved data on intracellular metabolite levels we also re-evaluated the need of sub-second quenching of metabolic activity and of removing the extracellular medium. Our findings have serious implications for quantitative metabolomics-based fields such as non-stationary 13 C flux analysis, in vivo kinetic modeling and thermodynamic network analysis.

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Realistic quantitative models require data from many laboratories. Therefore, standardization of experimental systems and assay conditions is crucial. Moreover, standards should be representative of the in vivo conditions. However, most often, enzyme-kinetic parameters are measured under assay conditions that yield the maximum activity of each enzyme. In practice, this means that the kinetic parameters of different enzymes are measured in different buffers, at different pH values, with different ionic strengths, etc. In a joint effort of the Dutch Vertical Genomics Consortium, the European Yeast Systems Biology Network and the Standards for Reporting Enzymology Data Commission, we have developed a single assay medium for determining enzyme-kinetic parameters in yeast. The medium is as close as possible to the in vivo situation for the yeast Saccharomyces cerevisiae, and at the same time is experimentally feasible. The in vivo conditions were estimated for S. cerevisiae strain CEN.PK113-7D grown in aerobic glucose-limited chemostat cultures at an extracellular pH of 5.0 and a specific growth rate of 0.1 h(-1). The cytosolic pH and concentrations of calcium, sodium, potassium, phosphorus, sulfur and magnesium were determined. On the basis of these data and literature data, we propose a defined in vivo-like medium containing 300 mM potassium, 50 mM phosphate, 245 mM glutamate, 20 mM sodium, 2 mM free magnesium and 0.5 mM calcium, at a pH of 6.8. The V(max) values of the glycolytic and fermentative enzymes of S. cerevisiae were measured in the new medium. For some enzymes, the results deviated conspicuously from those of assays done under enzyme-specific, optimal conditions.

The coenzyme NAD plays a major role in metabolism as a key redox carrier and signaling molecule but current measurement techniques cannot distinguish between different compartment pools, between free and protein-bound forms and/or between NAD(H) and NADP(H). Local free NAD/NADH ratios can be determined from product/substrate ratios of suitable near-equilibrium redox reactions but the application of this principle is often precluded by uncertainties regarding enzyme activity, localization and coenzyme specificity of dehydrogenases. In Saccharomyces cerevisiae, we circumvented these issues by expressing a bacterial mannitol-1-phosphate 5-dehydrogenase and determining the cytosolic free NAD/NADH ratio from the measured [fructose-6-phosphate]/[mannitol-1-phosphate] ratio. Under aerobic glucose-limited conditions we estimated a cytosolic free NAD/NADH ratio between 101(+/-14) and 320(+/-45), assuming the cytosolic pH is between 7.0 and 6.5, respectively. These values are more than 10-fold higher than the measured whole-cell total NAD/NADH ratio of 7.5(+/-2.5). Using a thermodynamic analysis of central glycolysis we demonstrate that the former are thermodynamically feasible, while the latter is not. Furthermore, we applied this novel system to study the short-term metabolic responses to perturbations. We found that the cytosolic free NAD-NADH couple became more reduced rapidly (timescale of seconds) upon a pulse of glucose (electron-donor) and that this could be reversed by the addition of acetaldehyde (electron-acceptor). In addition, these dynamics occurred without significant changes in whole-cell total NAD and NADH. This approach provides a new experimental tool for quantitative physiology and opens new possibilities in the study of energy and redox metabolism in S. cerevisiae. The same strategy should also be applicable to other microorganisms.

A sensitive, selective, and comprehensive method for the quantitative determination of tryptophan and 18 of its key metabolites in serum, urine, and cell culture supernatants was developed. The analytes were separated on a C18 silica column by reversed-phase liquid chromatography and detected by electrospray ionization tandem mass spectrometry in positive ion multiple reaction monitoring (MRM) mode, except for indoxyl sulfate which was measured in negative ion MRM mode in a separate run. The limits of detection and lower limits of quantification were in the range of 0.1-50 and 0.5-100 nM, respectively. Fully 13 C isotope-labeled and deuterated internal standards were used to achieve accurate quantification. The applicability of the method to analyze serum, urine, and cell culture supernatants was demonstrated by recovery experiments and the evaluation of matrix effects. Precision for the analysis of serum, urine, and cell culture supernatants ranged between 1.3% and 16.0%, 1.5% and 13.5%, and 1.0% and 17.4%, respectively. The method was applied to analyze changes in tryptophan metabolism in cell culture supernatants from IFN-γ-treated monocytes and immature or mature dendritic cells.

A major challenge in systems biology lies in the integration of processes occurring at different levels, such as transcription, translation, and metabolism, to understand the functioning of a living cell in its environment. We studied the high temperatureinduced glycolytic flux increase in Saccharomyces cerevisiae and investigated the regulatory mechanisms underlying this increase. We used glucose-limited chemostat cultures to separate regulatory effects of temperature from effects on growth rate. Growth at increased temperature (38°C versus 30°C) resulted in a strongly increased glycolytic flux, accompanied by a switch from respiration to a partially fermentative metabolism. We observed an increased flux through all enzymes, ranging from 5-to 10-fold. We quantified the contributions of direct temperature effects on enzyme activities, the gene expression cascade and shifts in the metabolic network, to the increased flux through each enzyme. To do this we adapted flux regulation analysis. We show that the direct effect of temperature on enzyme kinetics can be included as a separate term. Together with hierarchical regulation and metabolic regulation, this term explains the total flux change between two steady states. Surprisingly, the effect of the cultivation temperature on enzyme catalytic capacity, both directly through the Arrhenius effect and indirectly through adapted gene expression, is only a moderate contribution to the increased glycolytic flux for most enzymes. The changes in flux are therefore largely caused by changes in the interaction of the enzymes with substrates, products, and effectors.Microorganisms encounter environmental changes, which they have to withstand and adapt to in order to survive. Changes in ambient temperature are common to almost every ecological niche. Temperature influences the structural and functional properties of cellular components, both physically and chemically. Physically, temperature affects membrane fluidity (1, 2) and diffusion rates, as well as protein folding and stability (3). Chemically, temperature directly affects reaction rates in the cell. This study focuses on the adaptation of cells to temperatures higher than that optimal for growth.Microbes adapt to high temperature by altering their cellular make-up such as lipid composition, membrane fluidity, and the induction of large numbers of heat shock genes (3-11), which have a wide variety of functions. Many encode protein chaperones involved in protein (un)folding (12, 13) or degradation of damaged proteins (14). Others are involved in the synthesis of the thermoprotecting disaccharide trehalose, which is known to be involved in stabilization of membranes and proteins (15, 16) as well as in storage of free energy (17). Many of these adaptive responses put a significant additional energy burden on the cells (18).There still is little clarity on the actual mechanisms by which cells maintain a balance between the energy needs for adaptive responses to stress survival and those for processes indispensable for grow...

The field of systems biology is often held back by difficulties in obtaining comprehensive, highquality, quantitative data sets. In this paper, we undertook an interlaboratory effort to generate such a data set for a very large number of cellular components in the yeast Saccharomyces cerevisiae, a widely used model organism that is also used in the production of fuels, chemicals, food ingredients and pharmaceuticals. With the current focus on biofuels and sustainability, there is much interest in harnessing this species as a general cell factory. In this study, we characterized two yeast strains, under two standard growth conditions. We ensured the high quality of the experimental data by evaluating a wide range of sampling and analytical techniques. Here we show significant differences in the maximum specific growth rate and biomass yield between the two strains. on the basis of the integrated analysis of the highthroughput data, we hypothesize that differences in phenotype are due to differences in protein metabolism.