Quantification and Mass Isotopomer Profiling of Î±-Keto Acids in

Article pubs.acs.org/ac

Quantification and Mass Isotopomer Profiling of α‑Keto Acids in Central Carbon Metabolism Michael Zimmermann, Uwe Sauer, and Nicola Zamboni* Institute of Molecular Systems Biology, ETH Zurich, Zurich 8093, Switzerland S Supporting Information *

ABSTRACT: Mass spectrometry has been established as a powerful and versatile technique for studying cellular metabolism. Applications range from profiling of metabolites to accurate quantification and tracing of stable isotopes through the biochemical reaction network. Despite broad coverage of central carbon metabolism, most methods fail to provide accurate assessments of the α-keto acids oxaloacetic acid, pyruvate, and glyoxylate because these compounds are highly reactive and degraded during sample processing and mass spectrometric measurement. We present a derivatization procedure to chemically stabilize these compounds readily during quenching of cellular metabolism. Stable derivatives were analyzed by ultrahigh pressure liquid chromatography coupled tandem mass spectrometry to accurately quantify the abundance of α-keto acids in biological matrices. Eventually, we demonstrated that the developed protocol is suited to measure mass isotopomers of these α-keto acids in tracer studies with stable isotopes. In conclusion, the here described method fills one of the last technical gaps for metabolomics investigations of central carbon metabolism. We owe many of the analytical approaches to quantify α-keto acids to clinical research, which focused on the analysis of αketocaproate, α-ketoisovalerate, α-ketobutyrate, and glyoxylate in body fluids.6−10 Only a limited number of methods have been reported for the analysis of different α-keto acids.11−18 Studies aiming at sampling and quantifying intracellular α-keto acids are even more sparse.2,13,15,18,19 The simplest approach for quantification of α-keto acids relies on enzymatic assays with purified NAD(P)H-dependent dehydrogenases that are specific to individual α-keto acids and can be monitored with a spectrophotometer.20−22 Additionally, gas chromatography analysis after derivatization with ophenylenediamine or hydrazine species has been widely used.4,6,23−25 Similar derivatization protocols have also been applied prior to the separation of α-keto acids by high-pressure liquid chromatography to enable their photometric or fluorescent detection.5,8,16,26,27 Mass spectrometry-based detection from cells was reported for native α-ketoglutarate or α-

he α-keto acids are chemically reactive compounds, which in nature undergo decarboxylation, carboxylation, condensation, and amination reactions. The broad spectrum of products to which α-keto acids can be converted places them at pivotal positions in metabolism, both in health and disease. For instance, the two α-keto acids pyruvate and oxaloacetic acid, together with phosphoenolpyruvate, are at metabolic crossroads for flux rerouting between glycolysis and gluconeogenesis in central carbon metabolism.1 α-Ketoglutarate plays a crucial regulatory role at the intersection between the cellular carbon and nitrogen household.2,3 Glyoxylate is the most important physiological precursor molecule of oxalate, which upon crystallization leads to kidney stones.4 Accumulation of branched chain α-keto acids in urine, plasma, and cerebrospinal fluid was observed in patients with maple syrup urine disease, caused by impaired degradation of branched chain amino acids.5 Whereas the high reactivity of the α-keto acids is the chemical basis for their multiple metabolic functions, it causes inherent instability that poses an analytical challenge for their accurate quantification in biological matrices. In fact, measurements of intracellular concentrations of pyruvate, glyoxylate, and oxaloacetate are rare and often confounded by large errors.

T

© 2014 American Chemical Society

Received: February 3, 2014 Accepted: February 17, 2014 Published: February 17, 2014 3232

dx.doi.org/10.1021/ac500472c | Anal. Chem. 2014, 86, 3232−3237

Analytical Chemistry

Article

at final concentrations ranging from 25 to 6400 μM and prechilled at −20 °C. A solution containing glyoxylate, pyruvate, oxaloacetic acid, and α-ketoglutarate at a concentration of 100 μM was prepared in a fully 13C-labeled metabolite extract of E. coli corresponding to 100 μL of extract per 2 mL of culture volume at a density of OD600 = 1.0.29 Eight 3-fold dilutions of the α-keto acids mixture were prepared in the same 13C-labeled metabolic matrix. 100 μL of each dilution was added to the derivatization solution concomitantly with 100 μL of internal standard solution containing 5 μM [13C3]pyruvate and 2-keto-4-(methyl-d3)-pentanoic acid. Derivatization was performed at −20 °C for 1 h, and samples were stored at −80 °C before further processing. Samples were dried under vacuum (0.14 mbar) and resuspended in 100 μL of H2O, and 10 μL was injected for analysis by UHPLC-MS/MS. Analysis of Bacterial Cultures. E. coli BW25113 was from internal laboratory stocks. Bacteria were cultured on Luria Agar to pick colonies for preculturing in Luria Broth at 37 °C, 300 rpm. Exponentially growing cultures were diluted 1:1000 in M9 minimal medium with either glucose or acetic acid (5 g/L) as a single carbon source: Na2HPO4 (42.2 mM), KH2PO4 (22 mM), NaCl (8.56 mM), (NH4)2SO4 (11.34 mM), ZnSO4 (6.3 μM), CuCl2 (7 μM), MnSO4 (7.1 μM), CoCl2 (7.66 μM), thiamine (2.8 μM), MgSO4 (1 mM), CaCl2 (0.1 mM), and FeCl3 (0.6 mM). The pH was adjusted to 7.0 using NaOH. Bacteria were cultured to midexponential growth phase on either glucose or acetate as carbon source. A culture volume equivalent to 2 mL at OD600 = 1.0 was filtered through a 0.22 μm Durapore membrane (Millipore, Billerica, MA, USA). The bacteria on the filter were washed with 500 μL of fresh medium, and the filter was transferred to 3 mL of precooled (−20 °C) extraction solution (acetonitrile/methanol/water; 2:2:1 supplemented with 25 μM freshly prepared phenylhydrazine). 100 μL of internal standard solution containing [13C3]pyruvate and 2keto-4-(methyl-d3)-pentanoic acid (5 μM each) was added to the extraction solution. After incubation at −20 °C for 1 h, the samples were dried under vacuum at 30 °C and resuspended in 100 μL of water. Ten μL was injected for UHPLC-MS/MS analysis. Absolute concentrations were calculated on the basis of the total intracellular volume of the sampled biomass using 3.6 μL·mL−1·OD−1 and 4.0 μL·mL−1·OD−1 for glucose and acetate conditions, respectively.30 Measurement of Mass Isotopomer Distribution. E. coli was cultured as described above using a mixture of 40% [13C6]glucose and 60% [1-13C1]glucose. Following the same sample preparation as described above, the m/z of precursor ions of the MRM-assays was adjusted to all possible mass isotopomers of the hydrazone products. Analysis of 13Clabeling patterns of proteinogenic amino acids was performed by gas chromatography-coupled mass spectrometry (GC/MS) after hydrolysis of proteins and derivatization of amino acids with N-tertbutyldimethylsilyl-N-methyltrifluoroacetamide.31 Data Analysis. UHPLC-MS/MS and GC/MS traces were processed and analyzed by custom software using Matlab (Mathworks, Natick, MA United States).

ketoisovalerate15,28 or upon derivatization of carboxylic group. 18 However, these methods are not capable of quantifying intracellular concentrations of the chemically most unstable α-keto acids, such as glyoxylate or oxaloacetate. We present a method for quantifying intracellular α-keto acids with a focus on the four compounds belonging to central carbon metabolism: glyoxylate, pyruvate, oxaloacetate, and αketoglutarate. α-Keto acids are stabilized immediately during sampling by derivatization with phenylhydrazine. The conjugated system of the resulting hydrazone compounds not only stabilizes the α-keto acids but also provides an excellent molecular substructure for selective and sensitive detection by electrospray-tandem mass spectrometry. Using selected reaction monitoring (SRM), we demonstrate that our ultrahigh pressure liquid chromatography coupled tandem mass spectrometry (UHPLC-MS/MS) method can also be applied for the determination of the mass isotopomer distribution of αketo acids.

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EXPERIMENTAL SECTION Chemicals. All chemicals were purchased from SigmaAldrich (Schnelldorf, Switzerland). Solvents purchased were LC-MS grade, and chemicals were used at the highest purity available. Water was generated with a NANOpure purification unit (Barnstead, Dubuque, United States). 13C-labeled glucose was purchased from Cambridge Isotope Laboratories (Andover, MA). SRM Assays. A Thermo TSQ Quantum Ultra triple quadrupole instrument (Thermo Fisher Scientific, Waltham, MA, United States) with a heated electrospray ionization source (Thermo Fisher Scientific, Waltham, MA, United States) was used. Electrospray ionization parameters were optimized as described previously:15 spray voltage 2500 V, sheath gas pressure 80 arbitrary units, auxiliary gas pressure 50 arbitrary units, ion sweep gas pressure 5 arbitrary units, capillary temperature 380 °C, and spray temperature 400 °C. Ion optics were set to 0.5 amu Q1 and Q3 resolution, 0.01 amu scan width, and 10 ms dwell time. SRM parameters were optimized following the manufacturer’s recommendations using direct infusion. To estimate SRM parameters for phenylhydrazine derivatives, 1 mM aqueous solutions of each α-keto acid were incubated with 4 mM freshly prepared phenylhydrazine and incubated at ambient temperature (22 °C) for 1 h. Chromatographic Separation. Ultrahigh pressure chromatography was performed using a Waters Acquity UPLC (Waters Corporation, Milford, MA, United States) with a Waters Acquity HSS T3 column with dimensions 150 mm × 2.1 mm × 1.8 μm (Waters Corporation, Milford, MA, United States) thermostatted at 40 °C. A gradient of mobile phases A (10 mM tributylamine, 15 mM acetic acid, 5% (v/v) methanol) and B (2-propanol) was applied. Starting conditions: 0% B, 0.4 mL/min; 0.5 min: 0% B, 0.4 mL/min; 1.5 min: 12% B, 0.4 mL/ min; 5 min: 27.5% B, 0.25 mL/min; 10 min: 90% B, 0.15 mL/ min; 13 min: 90% B, 0.15 mL/min; 16 min: 0% B; 0.15 mL/ min; 20 min: 0% B; 0.4 mL/min. The flow rate was adjusted to keep the system pressure below 14 500 psi throughout the run. Ten μL was injected with full loop injection. Nonderivatized compounds were analyzed following the protocol described previously.15 Detection Limits and Linearity in Cellular Extracts. Three mL of metabolic extraction and derivatization solution was prepared for each sample consisting of a mixture of acetonitrile, methanol, and water (2:2:1) with phenylhydrazine

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RESULTS AND DISCUSSION MRM Assays and Chromatographic Separation. Measurement of α-keto acids is biased by their degradation during sample processing before detection. Therefore, we set out to chemically stabilize them in situ as early as sample preparation starts. We chose a derivatization protocol suitable for conditions applied to the quenching of metabolism being 3233



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the first step in sampling cells for metabolomic analyses. Derivatization of reactive α-keto acids with phenylhydrazine forms stable hydrazones, which can withstand extraction and analysis by mass spectrometry. The reaction can be carried quantitatively, at low temperatures, and in the presence of water. Therefore, it can be performed during metabolic quenching, that is the very first step of metabolome processing and aims at blocking endogenous catalytic activity. Phenylhydrazones are also beneficial for LC-MS/MS. The addition of an aromatic ring by derivatization facilitates chromatographic separation of the polar compounds by ion-pairing reveredphased chromatography. Furthermore, the favorable distribution of negative charge in the conjugated system of electrons upon collision-induced fragmentation amplifies the signal for detection by mass spectrometers based on SRM. To define the optimal parameters for LC-MS/MS analysis, 1 mM solutions of glyoxylate, pyruvate, oxaloacetate, and αketoglutarate in water were mixed with a 4-fold excess of freshly prepared phenylhydrazine to synthesize the corresponding hydrazones (Figure 1A). Derivatization was verified and followed by UV spectrometry at 324 nm.16 The hydrazones were used for optimization of compound-specific acquisition settings on the triple quadrupole instrument operated in negative mode ionization15 (Table 1). As expected from the chemical structure, the most prominent product ions upon collision-induced fragmentation were the aniline ion, the aniline radical, and the precursor ion after neutral loss of CO2 (Figure 1A). Separation of hydrazones was achieved by applying an ultrahigh pressure reversed-phase liquid chromatography system using water and isopropanol as aqueous and organic phase, respectively, plus tributylamine for ion pairing.15,32 A chromatogram demonstrating the separation of the four hydrazones in a biological matrix is shown in Figure 1B, together with the chromatogram of the stable isotopes [13C3]pyruvate and 2-keto-4-(methyl-d3)-pentanoate serving as internal standards. A massive increase in sensitivity was found when comparing the chromatograms obtained on the same LC-MS/MS platform for equivalent amounts of derivatized and native α-keto acids (Figure 2). Detection Limits and Linearity in Cellular Extracts. A mixture of glyoxylate, pyruvate oxaloacetate, and α-ketoglutarate was used to optimize derivatization conditions in a complex biological matrix. A dilution series of the four α-keto acids ranging from 0.01 to 100 μM was prepared with the background of a constant metabolome extract obtained from fully 13C-labeled E. coli cultures. 100 μL aliquots of each dilution were mixed with a prechilled metabolite extraction solution containing freshly prepared phenylhydrazine. The optimal concentration of phenylhydrazine had to be evaluated experimentally because high concentrations promote rapid and complete derivatization but also interfere with separation, provoke a loss of sensitivity in LC-MS/MS, and can lead to undesired side products derived from less reactive metabolites. Therefore, we assessed the detection limit and the range of linearity of the four analytes of interest for several phenylhydrazine concentrations between 25 and 6400 μM (Figure 3). All tested phenylhydrazine concentrations led to a comparable signal for the highest concentration of α-keto acids, indicating that the derivatization time of 1 h at −20 °C was generally sufficient. This also showed that phenylhydrazine was not limiting in a complex biological matrix, even at the highest concentrations of α-keto acids tested. However, the detection of low concentrations of α-keto acids and the range of linearity

Figure 1. Derivatization, MRM assay, and chromatographic separation. (A) Derivatization of the α-keto group leads to hydrazones. Collisioninduced fragmentation in the triple-quadrupole mass spectrometer leads to aniline ion, the aniline radical, and the precursor ion after neutral loss of CO2. (B) Chromatographic separation of hydrazone derivatives by UHPLC-MS/MS in a complex biological matrix. GOX, glyoxylate; PYR, pyruvate; OAA, oxaloacetate; AKG, α-ketoglutarate; 13C-PYR, [13C3]pyruvate; D3-KIV, 2-keto-4-(methyl-d3)-pentanoic acid.

were clearly impaired at phenylhydrazine concentrations higher than 400 μM (Figure 3). We used 25 μM phenylhydrazine in all further experiments to keep the amount of the derivatization agent minimal but sufficient. At high phenylhydrazine concentration, we also observed that the measurement of oxaloacetate in biological samples could be biased by a very slow derivatization of fumarate (Figure S1, Supporting Information). At a concentration of 25 μM phenylhydrazine, 3234



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Table 1. Phenylhydrazine Derivatives Detected monoisotopic mass [m/z]

Q1 mass of hydrazone [m/z]

tube lens [V]

glyoxylate pyruvate oxaloacetic acid

73.0 87.0 131.0

163.1 177.1 221.1

34 53 48

α-ketoglutarate [13C3]pyruvate 2-keto-4-(methyl-d3)pentanoate

145.0 90.0 132.1

235.1 180.1 222.1

40 53 53

compound

Q3 massesa [m/z] 92; 119; 92; 133; 92; 177; 91 191; 92; 92; 133; 92; 178;

91 91 133; 98 91 91

CE [AU]b 18; 14; 21 15; 14; 21 18; 177; 133; 91 12; 24; 16 15; 14; 21 20; 16; 91

RT [min]

detection limit [pmol]c

linear range [log3]d

5.48 5.26 7.33

0.12 0.041 0.041

4 5 5

6.85 5.26 8.52

0.013

6

a

Ions upon collision induced fragmentation in the order of their relative intensities. bCollision energy corresponding to the product ions of the previous column. cAmounts are indicated per injection on the column. dLinear range based on linear regression with R2 ≥ 0.99.

for a concentration range spanning 4 log3-units or more in the biological matrix (Table 1, Figure 4). The aniline fragment with

Figure 2. Comparison of peak shape and peak intensity between phenylhydrazine derivatized and nonderivatized GOX and PYR separated by ultrahigh pressure reversed-phase ion-pairing chromatography. MRM parameters and chromatographic settings for hydrazones and nonderivatized compounds were as described above and previously reported, respectively. Two pmol of each compound was injected for both methods. Figure 4. Linearity of peak integral normalized by internal standard signals. Measured ions resulting from collision-induced fragmentation in the triple-quadrupole mass spectrometer: ■ aniline ion; ● aniline radical; ▲ decarboxylation of the precursor ion. Average values of three independent dilutions are shown, and standard deviations are indicated by error bars.

m/z 92 could be used for sensitive quantification of all four derivatized α-keto acids. Instead, the aniline radical was used only for the quantification of glyoxylate and pyruvate derivatives, whereas the decarboxylated fragment was advantageous for the quantification of oxaloacetate and α-ketoglutarate (Figure 4). Determination of Intracellular Concentrations. To test the method and quantify α-keto acids in cells, we grew the bacterium Escherichia coli in batch cultures with either glucose or acetate as the sole carbon source. In the midexponential growth phase, cells were harvested by fast-filtration and immerged in the acetonitrile/methanol/water solution (2:2:1) with 25 μM phenylhydrazine at −20 °C to quench metabolism, extract metabolites, and form hydrazone. As a control, the same procedure was performed in the absence of the derivatization agent phenylhydrazine. Samples were dried, resuspended, and immediately analyzed with the here presented LC-MS/MS method devised for hydrazones, and a previously reported ion-

Figure 3. Optimization of the phenylhydrazine concentrations for the derivatization of α-keto acids. Dilution series of a mixture of glyoxylate, pyruvate, oxaloacetate, and α-ketoglutarate were derivatized in the presence of different concentrations of phenylhydrazine (25 to 6400 μM) and measured by LC-MS/MS. Representative results are shown for α-ketoglutarate, whose signal was normalized by the 2-keto-4(methyl-d3)-pentanoic acid signal.

the detection limit was below 0.12 pmol per injection for each of the four compounds tested. A linear response was obtained 3235



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Table 2. Intracellular Concentrations of α-Keto Acids in E. coli glucose acetate a

glyoxylate [μM]

pyruvate [μM]

oxaloacetate [μM]

α-ketoglutarate [μM]

α-ketoglutaratea [μM]

3.8 ± 1.7 17.6 ± 3.0

247 ± 78.8 57.6 ± 9.62

4.1 ± 0.61 1.5 ± 0.75

121 ± 23.9 35.8 ± 5.55

109 ± 11.9 39.7 ± 7.90

Quantified without derivatization following the protocol by Buescher et al.15

pairing method was optimized for nonderivatized intermediates of central carbon metabolism.15 13C-labeled internal standards were used for absolute quantification15,29,33 (Table 2). Among the four α-keto acids tested, α-ketoglutarate is the most stable and can be quantified also without derivatization. For growth on glucose, we determined intracellular concentrations of 121 ± 24 and 109 ± 12 μM α-ketoglutarate based on four independent cultures with and without phenylhydrazine derivatization, respectively. Although the intracellular concentration of α-ketoglutarate in E. coli was significantly lower during growth on acetate, the results derived from the two methods were equivalent (Table 2). From these results, we conclude that the protocol for sampling, quenching, and derivatizing α-keto acids is appropriate for their intracellular quantification. Hence, the developed procedure provides the possibility to also quantify the chemically unstable α-keto acids that are not quantitatively measurable without stabilizing derivatization. For instance, the glyoxylate concentration was about 4.6-fold increased under acetic acid compared to glucose conditions, consistent with increased metabolic flux through the glyoxylate shunt, which is essential for growth on acetic acid. Measurement of Mass Isotopomer Distribution. Stable isotope tracing is a technique widely applied for qualitative and quantitative analysis of metabolic fluxes through cells.34 Measuring labeling patterns of intermediates within central carbon metabolism improves the precision and resolution of fluxes.35 Specifically, the labeling patterns of the four α-keto acids tested in our study are of outmost relevance to investigate the fluxes in, and connecting, lower glycolysis and the TCA cycle. With the exception of α-ketoglutarate, the intensity of αketo acids in nonderivatized samples is generally too low for quantification of labeling patterns. To verify accurate measurement of mass isotopomer distributions in hydrazone derivatives, we cultured E. coli in minimal medium containing 40% [13C6]glucose and 60% [1-13C1]glucose and collected samples as described above. Mass isotopomer distributions of the hydrazone derivatives were determined by UHPLC-MS/MS as previously reported for other intermediates of central carbon metabolism.35 The intense peaks obtained through derivatization allowed us to precisely measure the relative abundances. We verified the measurement accuracy by comparing the mass isotopomer distributions obtained for the α-keto acids with that of the corresponding amino acids that in biosynthesis result from transamination of the α-keto group. Given that glucose was the unique carbon source for the synthesis of amino acids and samples were collected at isotopic steady state, the mass isotopomer distributions are expected to be identical for matching pairs. We determined the mass isotopomer fractions in proteinogenic amino acids by gas chromatography−mass spectrometry.31 The fractional labeling results from both methods were corrected for natural occurring isotopes introduced by the derivatization procedure or not pertinent to carbon.36 The results confirm that our method provides direct quantification of the labeling pattern of α-keto acids (Figure 5).

Figure 5. Comparison of mass isotopomer distribution of α-keto acids and according proteinogenic amino acids corrected for the natural abundance of 13C. Error bars indicate standard deviations of four independent biological replicates.

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CONCLUDING REMARKS We presented a protocol for metabolic quenching, derivatization, and sensitive quantification of α-keto acids in central carbon metabolism by ultrahigh pressure reversed-phase liquid chromatography coupled to tandem mass spectrometry. We demonstrated optimization of the derivatization procedure to produce hydrazones; we identified the linear range of measurements and determined the detection limit to the subpicomol range per injection for all compounds tested. The focus of our study was on the development of a protocol allowing simultaneous quenching of metabolism and derivatization of α-ketoacids in bacterial cells, which have been shown to alter their metabolite levels within seconds after environmental perturbations or long sampling procedure.37 Owing to 3236



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simultaneous quenching and derivatization performed at −20 °C, the presented method was even capable of quantifying the chemically very reactive glyoxylate and oxaloacetate in cellular extracts. In conclusion, the presented protocol closes one of the remaining gaps for comprehensive, quantitative metabolomics of central carbon metabolism allowing analysis of α-keto acids. Access to absolute intracellular concentrations of all intracellular compounds of central carbon metabolism in combination with the ability to measure their mass isotopomers will be a valuable tool for future investigations exploiting stoichiometric models for a better understanding of central carbon metabolism’s regulation.

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(15) Buescher, J. M.; Moco, S.; Sauer, U.; Zamboni, N. Anal. Chem. 2010, 82, 4403−4412. (16) Lange, M.; Mályusz, M. J. Chromatogr., B: Biomed. Sci. Appl. 1994, 662, 97−102. (17) Lee, S. H.; Kim, S. O.; Chung, B. C. J. Chromatogr., B: Biomed. Sci. Appl. 1998, 719, 1−7. (18) Han, J.; Gagnon, S.; Eckle, T.; Borchers, C. H. Electrophoresis 2013, 34, 2891−2900. (19) Mühling, J.; Tussing, F.; Nickolaus, K. A.; Matejec, R.; Henrich, M.; Harbach, H.; Wolff, M.; Weismüller, K.; Engel, J.; Welters, I. D.; Langefeld, T. W.; Fuchs, M.; Weigand, M. A.; Heidt, M. C. Amino Acids 2009, 38, 167−177. (20) Goodwin, G. W.; Kuntz, M. J.; Paxton, R.; Harris, R. A. Anal. Biochem. 1987, 162, 536−539. (21) Livesey, G.; Lund, P. Biochem. J. 1980, 188, 705. (22) Carpenter, S. E.; Merkler, D. J. Anal. Biochem. 2003, 323, 242− 246. (23) Cree, T. C.; Hutson, S. M.; Harper, A. E. Anal. Biochem. 1979, 92, 156−163. (24) Ohmori, S.; Tanaka, Y.; Ikeda, M.; Hirota, K. Anal. Biochem. 1981, 112, 204−212. (25) Kallio, H.; Linko, R. R. J. Chromatogr. 1973, 76, 229−232. (26) Hayashi, T.; Todoriki, H.; Naruse, H. J. Chromatogr., B: Biomed. Sci. Appl. 1981, 224, 197−204. (27) Fuchs, M.; Engel, J.; Campos, M.; Matejec, R.; Henrich, M.; Harbach, H.; Wolff, M.; Weismüller, K.; Menges, T.; Heidt, M. C.; Welters, I. D.; Krüll, M.; Hempelmann, G.; Mühling, J. Amino Acids 2008, 36, 1−11. (28) Bennett, B. D.; Kimball, E. H.; Gao, M.; Osterhout, R.; Van Dien, S. J.; Rabinowitz, J. D. Nat. Chem. Biol. 2009, 5, 593−599. (29) Mashego, M. R.; Wu, L.; Van Dam, J. C.; Ras, C.; Vinke, J. L.; Van Winden, W. A.; Van Gulik, W. M.; Heijnen, J. J. Biotechnol. Bioeng. 2004, 85, 620−628. (30) Volkmer, B.; Heinemann, M. PLoS One 2011, 6, No. e23126. (31) Zamboni, N.; Fendt, S.-M.; Rühl, M.; Sauer, U. Nat. Protoc. 2009, 4, 878−892. (32) Zimmermann, M.; Thormann, V.; Sauer, U.; Zamboni, N. Anal. Chem. 2013, 85, 8284−8290. (33) Ewald, J. C.; Heux, S.; Zamboni, N. Anal. Chem. 2009, 81, 3623−3629. (34) Sauer, U. Mol. Syst. Biol. 2006, 2, 62. (35) Rühl, M.; Rupp, B.; Nöh, K.; Wiechert, W.; Sauer, U.; Zamboni, N. Biotechnol. Bioeng. 2011, 109, 763−771. (36) Yuan, J.; Bennett, B. D.; Rabinowitz, J. D. Nat. Protoc. 2008, 3, 1328−1340. (37) Link, H.; Kochanowski, K.; Sauer, U. Nat. Biotechnol. 2013, 31, 357−361.

ASSOCIATED CONTENT

S Supporting Information *

Kinetics of hydrazone formation of oxaloacetate, malate, and fumarate with phenylhydrazine. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank Juri Demmer and Flavia Tschan for their assistance during sampling of bacteria and the anonymous reviewer for highlighting an important inconsistency. This research was supported by funding of the EU FP7 project SysteMTb.

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REFERENCES

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Quantification and Mass Isotopomer Profiling of Î±-Keto Acids in

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