We here explain step by step the implementation of gas chromatography coupled with tandem mass spectrometry for the quantitative analysis of intracellular metabolites from the tricarboxylic acid (TCA) cycle such as citrate, isocitrate, alpha-ketoglutarate, succinate, malate, and fumarate. Isotope dilution is used to correct for potential metabolite losses during sample processing, matrix effects, incomplete derivatization, and liner contamination. All measurements are performed in selected reaction monitoring (SRM) mode. Standards and samples are first diluted with a fixed volume of a mixture of fully ¹³C-labeled internal standards and then derivatized to give trimethylsilyl-methoxylamine derivatives prior GC-MS/MS analysis.

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In its natural environment, the filamentous fungus Aspergillus niger grows on decaying fruits and plant material, thereby enzymatically degrading the lignocellulosic constituents (lignin, cellulose, hemicellulose, and pectin) into a mixture of mono- and oligosaccharides. To investigate the kinetics and stoichiometry of growth of this fungus on lignocellulosic sugars, we carried out batch cultivations on six representative monosaccharides (glucose, xylose, mannose, rhamnose, arabinose, and galacturonic acid) and a mixture of these. Growth on these substrates was characterized in terms of biomass yields, oxygen/biomass ratios, and specific conversion rates. Interestingly, in combination, some of the carbon sources were consumed simultaneously and some sequentially. With a previously developed protocol, a sequential chemostat cultivation experiment was performed on a feed mixture of the six substrates. We found that the uptake of glucose, xylose, and mannose could be described with a Michaelis–Menten-type kinetics; however, these carbon sources seem to be competing for the same transport systems, while the uptake of arabinose, galacturonic acid, and rhamnose appeared to be repressed by the presence of other substrates.

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Poly(3-hydroxybutyrate) (PHB) is an interesting biopolymer for replacing petroleum-based plastics, its biological production is performed in natural and engineered microorganisms. Current metabolic engineering approaches rely on high-throughput strain construction and screening. Analytical procedures have to be compatible with the small scale and speed of these approaches. Here, we present a method based on isotope dilution mass spectrometry (IDMS) and propanolysis extraction of poly(3-hydroxybutyrate) from an Escherichia coli strain engineered for PHB production. As internal standard (IS), we applied an uniformly labeled 13C-cell suspension, of an E. coli PHB producing strain, grown on U-13C-glucose as C-source. This internal 13C-PHB standard enables to quantify low concentrations of PHB (LOD of 0.01 µg/gCDW) from several micrograms of biomass. With this method, a technical reproducibility of about 1.8% relative standard deviation is achieved. Furthermore, the internal standard is robust towards different sample backgrounds and dilutions. The early addition of the internal standard also enables higher reproducibility and increases sensitivity and throughput by simplified sample preparation steps@en

Ammonium (NH4+) is the most common N-source for yeast fermentations, and N-limitation is frequently applied to reduce growth and increase product yields. While there is significant molecular knowledge on NH4 + transport and assimilation, there have been few attempts to measure the in vivo concentration of this metabolite. In this article, we present a sensitive and accurate analytical method to quantify the in vivo intracellular ammonium concentration in
Saccharomyces cerevisiae based on standard rapid sampling and metabolomics techniques. The method validation experiments required the development of a proper sample processing protocol to minimize ammonium production/consumption during biomass extraction by assessing the impact of amino acid degradation—an element that is often overlooked. The resulting cold chloroform metabolite extraction method, together with quantification using ultra high performance liquid chromatography-isotope dilution mass spectrometry (UHPLC-IDMS), was not only more sensitive than most of the existing methods but also more accurate than methods that use electrodes, enzymatic reactions, or boiling water or boiling ethanol biomass extraction because it minimized ammonium consumption/production during sampling processing and interference from other metabolites in the quantification of intracellular ammonium. Finally, our validation experiments showed that other
metabolites such as pyruvate or 2-oxoglutarate (KG) need to be extracted with cold chloroform to avoid measurements being biased by the degradation of other metabolites (e.g., amino acids).@en

Ammonium is the most common N source for yeast fermentations. Although its transport and assimilation mechanisms are well documented, there have been only a few attempts to measure the in vivo intracellular concentration of ammonium and assess its impact on gene expression. Using an isotope dilution mass spectrometry (IDMS)-based method, we were able to measure the intracellular ammonium concentration in N-limited aerobic chemostat cultivations using three different N sources (ammonium, urea, and glutamate) at the same growth rate (0.05 h^-1). The experimental results suggest that, at this growth rate, a similar concentration of intracellular (IC) ammonium, about 3.6 mmol NH₄ ⁺/liter_IC, is required to supply the reactions in the central N metabolism, independent of the N source. Based on the experimental results and different assumptions, the vacuolar and cytosolic ammonium concentrations were estimated. Furthermore, we identified a futile cycle caused by NH₃ leakage into the extracellular space, which can cost up to 30% of the ATP production of the cell under N-limited conditions, and a futile redox cycle between Gdh1 and Gdh2 reactions. Finally, using shotgun proteomics with protein expression determined relative to a labeled reference, differences between the various environmental conditions were identified and correlated with previously identified N compound-sensing mechanisms.

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Eukaryotic metabolism consists of a complex network of enzymatic reactions and transport processes which are distributed over different subcellular compartments. Currently, available metabolite measurement protocols allow to measure metabolite whole cell amounts which hinder progress to describe the in vivo dynamics in different compartments, which are driven by compartment specific concentrations. Phosphate (Pi) is an essential component for: (1) the metabolic balance of upper and lower glycolytic flux; (2) Together with ATP and ADP determines the phosphorylation energy. Especially, the cytosolic Pi has a critical role in disregulation of glycolysis in tps1 knockout. Here we developed a method that enables us to monitor the cytosolic Pi concentration in S. cerevisiae using an equilibrium sensor reaction: maltose+Pi glucose+glucose-1-phosphate. The required enzyme, maltose phosphorylase from L. sanfranciscensis was overexpressed in S. cerevisiae. With this reaction in place, the cytosolic Pi concentration was obtained from intracellular glucose, G1P and maltose concentrations. The cytosolic Pi concentration was determined in batch and chemostat (D=0.1 h^-1) conditions, which was 17.88μmol/gDW and 25.02μmol/gDW, respectively under Pi-excess conditions. Under Pi-limited steady state (D=0.1 h^-1) conditions, the cytosolic Pi concentration dropped to only 17.7% of the cytosolic Pi in Pi-excess condition (4.42μmol/gDW vs. 25.02μmol/gDW). In response to a Pi pulse, the cytosolic Pi increased very rapidly, together with the concentration of sugar phosphates. Main sources of the rapid Pi increase are vacuolar Pi (and not the polyPi), as well as Pi uptake from the extracellular space. The temporal increase of cytosolic Pi increases the driving force of GAPDH reaction of the lower glycolytic reactions. The novel cytosol specific Pi concentration measurements provide new insight into the thermodynamic driving force for ATP hydrolysis, GAPDH reaction, and Pi transport over the plasma and vacuolar membranes.

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Fluxomics and metabolomics are crucial tools for metabolic engineering and biomedical analysis to determine the in vivo cellular state. Especially, the application of 13C isotopes allows comprehensive insights into the functional operation of cellular metabolism. Compared to single MS, tandem mass spectrometry (MS/MS) provides more detailed and accurate measurements of the metabolite enrichment patterns (tandem mass isotopomers), increasing the accuracy of metabolite concentration measurements and metabolic flux estimation. MS-type data from isotope labeling experiments is biased by naturally occurring stable isotopes (C, H, N, O, etc.). In particular, GC-MS(/MS) requires derivatization for the usually non-volatile intracellular metabolites introducing additional natural isotopes leading to measurements that do not directly represent the carbon labeling distribution. To make full useof LC- and GC-MS/MS mass isotopomer measurements, the influence of natural isotopes has to be eliminated (corrected). Our correction approach is analyzed for the two most common applications; 13C fluxomics and isotope dilution mass spectrometry (IDMS) based metabolomics. Natural isotopes can have an impact on the calculated flux distribution which strongly depends on the substrate labeling and the actual flux distribution. Second, we show that in IDMS based metabolomics natural isotopes lead to underestimated concentrations that can and should be corrected with a nonlinear calibration. Our simulations indicate that the correction for natural abundance in isotope based fluxomics and quantitative metabolomics is essential for correct data interpretation. Biotechnol.@en