1
Synthesis and use of stable isotope labelled internal standards for quantification
of phosphorylated metabolites by LC-MS/MS
Stéphanie Arrivault1*, Manuela Guenther1, Stephen C. Fry2, Maximilian M.F.F. Fuenfgeld1, Daniel
Veyel1, Tabea Mettler-Altmann1,3, Mark Stitt1 and John E. Lunn1
1Max
Planck Institute of Molecular Plant Physiology, Am Muehlenberg 1, 14476 Potsdam-Golm,
Germany
2The
Edinburgh Cell Wall Group, Institute of Molecular Plant Sciences, School of Biological
Sciences, The University of Edinburgh, Edinburgh, UK
3Present
address: Cluster of Excellence on Plant Sciences (CEPLAS) and Institute of Plant
Biochemistry, Heinrich-Heine-Universität Düsseldorf, Universitätsstraße 1, 40225 Düsseldorf,
Germany
* To whom correspondence should be addressed: Stéphanie Arrivault, Max Planck Institute of
Molecular Plant Physiology, Am Muehlenberg 1, 14476 Potsdam-Golm, Germany; telephone
+493315678114; [email protected]
2
ABSTRACT
Liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS) is a highly specific
and sensitive technique for measuring metabolites. However, co-eluting components in tissue
extracts can interfere with ionization at the interface of the LC and MS/MS phases, causing underor over-estimation of metabolite concentrations. Spiking of samples with known amounts of
stable isotope labelled internal standards (SIL-IS) allows measurements of the corresponding
metabolites to be corrected for such matrix effects. We describe criteria for selection of suitable
SIL-IS, and report the enzymatic synthesis and purification of nine SIL-IS for hexose-, pentoseand triose-phosphates, UDP-glucose and adenosine monophosphate (AMP). Along with
commercially available SIL-IS for seven other metabolites, these were validated by LC-MS/MS
analyses of extracts from leaves, non-photosynthetic plant tissues, mouse liver, and cells of
Chlamydomonas reinhardtii, Escherichia coli and baker’s yeast (Saccharomyces cerevisiae). With
only a few exceptions, spiking with SIL-IS significantly improved the reproducibility of LC-MS/MSbased metabolite measurements across a wide range of extract dilutions, indicating effective
correction for matrix effects by this approach. With use of SIL-IS to correct for matrix effects, LCMS/MS offers unprecedented scope for reliable determination of photosynthetic and respiratory
intermediates in a diverse range of organisms.
3
INTRODUCTION
Liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS) has become the
method of choice for measurement of plant metabolites1 including phosphorylated
intermediates2,3,4, lipids5 and secondary compounds6,7,8, and is also widely used for metabolomics
in other organisms including bacteria9 and yeast10.
Like other MS-based analytical techniques, LC-MS/MS is susceptible to matrix effects that can
reduce the accuracy and reliability of the analysis11. Matrix effects occur when sample
components co-elute with the analyte of interest and interfere with the ionization process. The
most common matrix effect is ionization suppression, which leads to under-estimation of the
affected analyte, but enhancement of ionization, leading to over-estimation, can also occur.
While the exact mechanisms are still not fully understood, it has been postulated that co-eluting
compounds interfere with the charging or desolvation of analytes at the interface between the
LC and the MS system (the ion source)12,13. Electrospray ionization (ESI), the most widely used
ion source, is more susceptible to matrix effects than atmospheric pressure photoionization
(APPI) and atmospheric pressure chemical ionization (APCI)13, 14. The degree of interference also
depends on the chemical nature of the analytes12, and on the biological source of the sample15.
Phospholipids contribute to ion suppression during analysis of blood plasma samples13,16, and
chloride ions from the culture medium caused ion suppression during analysis of
Chlamydomonas reinhardtii cell extracts15. However, the source of interference is usually
unknown.
Matrix effects can be overcome by avoiding or minimizing the interference, and by using methods
to quantify and correct for the interference. To reduce interference (reviewed in 17), changes can
be made to either the source of the biological material (e.g. by using a different cell culture
medium15) or the analytical technique itself. Potential analytical modifications include: (i)
decreasing the volume or concentration of the extract injected; (ii) partial purification of the
sample prior to analysis (e.g. by selective precipitation of interfering compounds, two-phase
(liquid-liquid or liquid-solid) extraction16, or membrane filtration2); (iii) reducing the flow rate
during the LC phase18; (iv) changing the type of column matrix, eluent composition or eluent
gradient to separate the analyte of interest from the interfering compound(s) 16; (v) combining
two different types of chromatographic separation in series (LC-LC)19; and (vi) use of a different
ion source20. Some of these options might not be suitable for low abundance metabolites, due
to poor recoveries from pre-analytical cleanup or inadequate machine sensitivity, and most of
these approaches increase the technical complexity and often the cost of the analysis.
Despite efforts to minimize matrix effects, it is often not possible to eliminate them altogether.
Therefore, methods to quantify and compensate for the interference are required for metabolite
measurements to be accurate and reproducible21,22,15. Standard addition23 and matrix-matched
calibration24 attempt to compensate for the interference by mixing standards with extract or
interfering compounds to generate sample-specific calibration curves. However, both of these
approaches increase the workload and time needed for analysis. Furthermore, unless applied to
each individual sample, correction using these methods is still prone to error arising from
variation in matrix effects between different biological samples.
4
These techniques have largely been superseded by use of stable isotope labelled internal
standards (SIL-IS), which allow correction for matrix effects in each individual sample with
minimal addition to the workload and analysis time23. A SIL-IS is an isotopically-labelled form of
the metabolite of interest, in which two or more atoms have been substituted by non-radioactive
isotopes. The most commonly used isotopes are 2H, 13C and, to a lesser extent, 15N and 17O. The
inclusion of stable isotopes has no significant effect on most of the physicochemical properties
of the molecule, so both the labelled and unlabelled forms behave in the same way during sample
extraction and chemical derivatization. Labelled and unlabelled forms usually co-elute during
chromatography, although heavily deuterated isotopomers sometimes show a shift in retention
time25. The unlabelled and labelled isotopomers have essentially identical ionization
characteristics. However, following ionization, the resulting ions have different m/z values and
so can be readily distinguished by MS. To correct for matrix effects, standard mixes and samples
are spiked with a known amount of the SIL-IS before analysis. The labelled and unlabelled forms
of the metabolite of interest co-elute from the LC and are exposed to identical ionization
conditions, including any interference by other molecules, before entering the MS. There are
several ways to use SIL-IS to correct for matrix effects. A simple approach is to compare the
signals from a known amount of the SIL-IS analyzed on its own (i.e. no matrix effects) and when
spiked into a sample. The ratio between the signals can then used as a correction factor to
normalize the signal from the unlabelled compound in the sample. Some types of software used
for analysis of LC-MS/MS chromatograms allow more automated approaches. For example,
mixtures of unlabelled standards can be spiked with a known amount of the SIL-IS to set up
calibration curves, and then the ratio of the signals from the labelled and unlabelled metabolite
in an SIL-IS-spiked sample is compared with the calibration standards to calculate the amount of
unlabelled metabolite in the sample. Ideally, SIL-IS should be used for each measured metabolite
but, with few exceptions26, this goal is rarely achieved, or even approached, due to the limited
commercial availability of suitable labelled isotopomers. In fact in many studies, the issue of
matrix effects is simply, but ill-advisedly, ignored. For molecules of particular importance, effort
may be made to synthesize a suitable labelled isotopomer for use as an IS2.
Arrivault and colleagues3 developed a reverse-phase LC-MS/MS protocol to measure 27
metabolites involved in photosynthetic or respiratory metabolism in plants. Commercially
available isotopomers of six of these metabolites were used as SIL-IS: [2,3,3-2H3]glutamate,
[2,3,3-2H3]aspartate, [2,3,3-2H3]glycerate, [U-13C]succinate, [2,3,3-2H3]malate and [1,2,3,4-13C4]2oxoglutarate, and a deuterated form of glucose 6-phosphate, [6,6-2H2]G6P, was enzymatically
synthesized from commercially available [6,6-2H2]glucose. Here we discuss what factors need to
be considered when selecting or designing an SIL-IS, and report the enzymatic synthesis of
isotopomers of eight further phosphorylated metabolites and their validation for use as SIL-IS in
LC-MS/MS analysis. SIL-IS can also be used to assess the recovery of metabolites during
extraction, but here we focus on their use to correct for matrix effects in LC-MS/MS analysis.
EXPERIMENTAL SECTION
Materials (chemicals)
5
[2,3,3-2H3]Aspartic acid, [2,3,3-2H3]glutamic acid and [1,2,3,4-13C4]2-oxoglutarate (2-OG) were
obtained from Cambridge Isotope Laboratories (http://www.isotope.com). [2,3,3-2H3]malic acid,
[U-13C]succinic acid, [2,3,3-2H3]glyceric acid, [6,6-2H2]glucose, [1,6-13C2]fructose, [13C10,14N5]ATP,
[13C10,14N5]UTP and ATP were obtained from Sigma-Aldrich (http://www.sigmaaldrich.com). [U13C]FBP (fructose 1,6-bisphosphate), [U-13C]sucrose and [2,3,4,5-13C ]ribose were from Campro
4
(http://www.campro.eu). All enzymes were from Sigma-Aldrich with the exception of
hexokinase, inorganic pyrophosphatase and aldolase (Roche Diagnostics GmbH, Mannheim,
Germany, http://www.roche.de) and E. coli ribokinase (kindly provided by Dr Roman Esipov,
Shemyakin–Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of Sciences,
Moscow, Russia). The sources of plant, bacterial and animal material and sampling procedures
are described in the Supplementary Material and Methods.
Synthesis of isotopically labelled compounds
Unless stated otherwise, reaction mixtures (1 ml final volume) were incubated at 30 °C for 3 h
and then the reaction stopped by heating at 99°C for 2 min. Precipitated proteins were removed
by centrifugation (18,000×g, 2 min) and the labelled product purified by high voltage paper
electrophoresis (HVPE) of the supernatant. The composition of specific labelling reactions was as
follows.
[6,6-2H2]G6P and [1,6-13C2]F6P (fructose 6-phosphate): 10 µmol [6,6-2H2]glucose or 10 µmol [1,613C ]fructose, respectively, 12 μmol ATP, 5 U hexokinase (EC 2.7.7.1), 5 mM Tris-HCl (pH 7.5) and
2
1 mM MgCl2.
[U-13C]G1P (glucose 1-phosphate): 10 µmol of [U-13C]sucrose, 10 U sucrose phosphorylase (EC
2.4.1.7), 50 µmol K2HPO4-KOH (pH 7.0), 5 mM Tris-HCl (pH 7.0) and 1 mM MgCl2. As the supplied
sucrose phosphorylase contained sucrose as stabilizer, 125 U of the enzyme was dissolved in 1.5
ml of 20 mM MES-NaOH (pH 7.0) and desalted using a 20-ml centrifugal concentrator as
according the manufacturer recommendations (10 kDa cut-off; Amicon Ultra, Merck Millipore,
http://merckmillipore.com).
[13C10,14N5]UDPG (UDP glucose): 12 µmol [13C10,14N5]UTP, 10 µmol G1P, 5 U UGPase (EC 2.7.7.9),
5 U inorganic pyrophosphatase (EC 3.6.1.1), 5 mM Tris-HCl (pH 7.5) and 1 mM MgCl2.
[2,3,4,5-13C4]R5P (ribose 5-phosphate): 10 µmol [2,3,4,5-13C4]ribose, 12 µmol ATP, with 5 U
ribokinase (EC 2.7.1.15), 5 mM Tris-HCl (pH 7.9), 1mM MgCl2 and 30 mM KCl (the presence of
potassium and magnesium ions increase ribokinase activity27), and was incubated at 37°C for 4
h. After stopping the reaction by heating at 100°C for 2 min, ATP and AMP were quantitatively
hydrolysed to AMP by addition of 5 U nucleoside-triphosphate phosphatase (apyrase1; EC
3.6.1.19) and 5 U nucleoside-diphosphate phosphatase (apyrase2; EC 3 6.1.6) and incubation at
37°C for 2 h. The apyrase reactions were stopped by heating at 100°C for 2 min.
[2,3,4,5-13C4]Ru5P (ribulose 5-phosphate) and [2,3,4,5-13C4]Xu5P (xylulose 5-phosphate): 10 µmol
[2,3,4,5-13C4]ribose, 12 µmol ATP, with 5 U ribokinase, 5 U ribose-5-phosphate isomerase (EC
5.3.1.6), 5 U ribulose-phosphate 3-epimerase (EC 5.1.3.1), 5 mM Tris-HCl (pH 7.9), 1mM MgCl2
and 30 mM KCl. After incubation at 37°C for 2 h, a further 5 U of ribokinase were added and the
6
reaction incubated at 37°C for another 2 h. After stopping the reaction by heating at 100°C for 2
min, ATP and ADP were removed by treatment with apyrase 1 and apyrase 2 as described above.
[U-13C]DHAP (dihydroxy-acetone-phosphate): 10 µmol [U-13C]FBP, 2 U aldolase (EC 4.1.2.13), 3 U
TPI (EC 5.3.1.1), 5 mM Tris-HCl (pH 7.9)and 1 mM MgCl2.
[13C10,15N5]AMP (adenosine monophosphate): 12 µmol [13C10,15N5]ATP, 5 U apyrase1, 5 U
apyrase2, 5 mM MES-NaOH (pH 6.0) and 1 mM CaCl2.
Separation by HVPE
Preparative HVPE was performed as described in28,29. Standards and reaction mixtures were
applied to Whatman No. 1 paper (42 × 57 cm), along an origin line 9 cm from the edge of the
paper. Authentic standards were loaded at 50 µg per spot except DHAP (25 µg), and Pi and PPi
(15 µg each). Five µg of the visible mobile marker orange G was mixed with the sample or loaded
as a series of spots alternating with the samples. After sample loading, the paper was wetted
with a volatile electrophoresis buffer at pH 3.5 – acetic acid/pyridine/H2O (10:1:189 by volume);
or pH 6.5 – acetic acid/pyridine/H2O (1:33:300 by volume). In preliminary trials to identify the
optimal electrophoresis conditions for separation, a volatile electrophoresis buffer at pH 2.0
(formic acid/acetic acid/H2O; 1:4:45 by volume) was also tested. The paper was suspended in the
HVPE apparatus, consisting of a glass tank fitted with platinum electrodes at top and bottom. The
edge of the paper closest to the origin was placed uppermost, dipping into a glass trough
containing electrophoresis buffer (cathode), while the lower end of the paper dipped into a layer
of electrophoresis buffer at the bottom (anode) of the tank. The remainder of the tank was filled
with a non-polar solvent (white spirit for pH 2.0 and 3.5, toluene/pyridine 20:1 by volume for pH
6.5) that was cooled by circulation of cold tap-water through cooling coils suspended from the
lid of the apparatus. The paper was electrophoresed at 4.5 kV for 20–40 min (as indicated for
individual experiments). Following electrophoresis and drying to remove the volatile buffers and
coolant, standards were visualized by UV absorbance (nucleotides, including UDPG) or staining
with molybdate reagent or AgNO329. Using the appropriate standard as a guide, a strip containing
the labelled compound of interest was excised from the preparative zone, and the compound
eluted with water using the syringe-barrel method29. The eluted compound was lyophilized and
redissolved in 250 or 500 µl of deionized water (Millipore, http://www.merckmillipore.com)
before use.
Quantification of metabolites
Metabolite analysis was performed by ion pair reverse-phase chromatography coupled to
tandem mass spectrometry using a Thermo triple quadrupole MS as previously described3, with
slight modifications of the LC gradient30. Aliquots of frozen tissue powder from Arabidopsis,
maize, wheat and tobacco leaves (15 mg FW), green and red tomato fruits, potato tuber and
mouse liver (20 mg FW) were extracted with chloroform-methanol as described3.
Chlamydomonas reinhardtii cells were extracted as described31,30. Yeast and E. coli were
extracted by addition of 1 ml of 90 % (v/v) methanol cooled to -20 °C to the frozen cells on liquid
nitrogen, thawed on ice, refrozen and sonicated in a bath of iced-water for 10 min. The extract
was centrifuged at 21,000 ×g (0°C) for 10 min. The supernatant (900 µl) was transferred to a fresh
7
tube and 260 µl of chloroform was added to each methanolic extract, with subsequent phase
partitioning and lyophilization as previously described3. After lyophilization, extracts of mouse
liver, Chlamydomonas reinhardtii, yeast and E. coli cells were dissolved in 0.5 vol. (150 µl) of water
before analysis. Synthesis reaction mixtures and purified SIL-IS were diluted 1/150 with deionized
water prior to analysis. Fragmentation products for SIL-IS are listed in Table S1. A concentrated
SIL-IS mixture was prepared and 5 µl, containing the amounts shown in Table S1, were added to
each sample prior to measurement. ATP, fructose and sucrose were measured enzymatically
using a Sigma-22 dual-wavelength photometer32.
RESULTS AND DISCUSSION
Criteria for selection of SIL-IS
The stable isotopes most commonly used for labelling of biological molecules also occur
naturally. The most abundant is 13C, which comprises 1.1 % of total C on a global scale. Other
stable isotopes are less abundant: 2H, 0.0156 %; 17O, 0.0373 % and 15N 0.364 %, but can be higher
(or lower) in biological materials due to environmental variation or biological discrimination33.
The presence of stable isotopes in molecules of biological origin can interfere with the signal of
the corresponding SIL-IS in LC-MS/MS assays if the m/z value of the selected SIL-IS overlaps with
naturally occurring isotopomers in the sample.
We used the ChemCalc (http://www.chemcalc.org/main)34 tool to calculate the abundance of
naturally occurring isotopomers of the metabolites of interest. Calculations were based on the
monovalent anionic form of metabolites of interest ([M-H]-) as our established LC-MS/MS
protocols2,3 operate in negative ESI mode. Table 1 lists metabolites of interest with estimates of
the natural abundance of isotopomers with m/z values of +1 to +6 above the most abundant
naturally occurring form, and expressed as a percentage of the latter. It should be noted that
these calculations might overstate the observed isotopic interference because they do not take
into account the fragmentation used in MS/MS35. Nevertheless, this is a simple approach to avoid
selection of inappropriate SIL-IS that might be subject to isotopic interference. For example,
putative UDPG SIL-IS labelled with one or two 2H or 13C atoms would have non-negligible overlap
with naturally occurring isotopomers with m/z vales of M+1 (18 % abundance) and M+2 (5 %
abundance), respectively. Therefore, for UDPG, an SIL-IS with an m/z value of M+3 or higher
would be preferred. Following this principle, we selected the following commercially available
isotopomers for use as SIL-IS: [1,2,3,4-13C4]2-OG, [2,3,3-2H3]malate, [2,3,3-2H3]glycerate,
[U13C]succinate, [2,3,3-2H3]glutamate and [2,3,3-2H3]aspartate. The estimated signal contribution
from M+3 and M+4 isotopomers of these compounds was <0.06 % (Table 1). Of two commercially
available isotopomers of fructose 1,6-bisphosphate (FBP), [1/2/6-13C1]FBP and [U-13C]FBP, we
chose the latter, as the potential signal overlap is negligible compared to that of the M+1
isotopomer (>7 %). Previously we had synthesized [2,2-2H2]G6P, from [2,2-2H2]glucose using
hexokinase (Figure 1A), for use as a SIL-IS for this metabolite3. M+2 isotopomers were predicted
to have a natural abundance of 2 % (Table 1). The potential error from isotopic interference by
G6P (M+2) in G6P measurements would be in the same range as pipetting error, so use of [2,22H ]G6P as a SIL-IS was judged to be acceptable. No retention time shift was observed for this
2
deuterated SIL-IS.
8
There were no commercially available stable isotopomers for the other metabolites of interest.
Therefore, we designed strategies for synthesizing suitable SIL-IS enzymatically, based on the
availability of potential precursors and enzymes, and using the data in Table 1 to decide the
minimum degree of labelling necessary to avoid isotopic interference.
Synthesis of 13C-labelled hexose-phosphates
Hexose-phosphates are respiratory intermediates in most organisms, and in plants are also
important as intermediates of the Calvin-Benson cycle (F6P), and sucrose and starch synthesis.
We synthesized [1,6-13C2]F6P from [1,6-13C2]fructose in a one-step reaction catalyzed by
hexokinase (Figure 1B), with a stoichiometric excess of ATP to ensure quantitative conversion of
[1,6-13C2]fructose to [1,6-13C2]F6P. ADP is the major by-product of the reaction, although AMP
can also be produced if the hexokinase is contaminated with adenylate kinase (myokinase). To
avoid interference with measurements of ADP and AMP in samples, it was essential to remove
these by-products from the preparation of [1,6-13C2]F6P to be used as a SIL-IS. [U-13C]G1P was
synthesized by phosphorolytic cleavage of [U-13C]sucrose using sucrose phosphorylase (SPase;
Figure 1C) and a stoichiometric excess of orthophosphate (Pi), with fructose as a by-product36.
Two methods offered the necessary resolution and capacity for purification of the synthesized
compounds: high-performance liquid chromatography (HPLC) and HVPE. However, typical HPLC
eluents (e.g. sodium hydroxide) would be incompatible with LC-MS/MS analysis, whereas HVPE
uses volatile solvents (formic + acetic acids for electrophoresis at pH 2.0, and pyridinium acetate
buffers for electrophoresis at pH 3.5 or 6.529) that can easily be removed by drying the paper
before elution of the labelled compound. Therefore, HVPE was chosen for purification of the SILIS. Low concentrations of buffering agents and cofactors (e.g. MgCl2) were used for the enzymatic
synthesis reactions, to minimize the ionic load in the HVPE purification step. The optimal pH for
separation of the labelled compound from potentially interfering contaminants was selected,
based on previous knowledge or preliminary analyses of authentic standards.
Figure 2A shows an HVPE electrophoretogram at pH 6.5, of hexose-phosphates and adenine
nucleotide standards, showing clear separation of G6P, F6P and G1P from AMP, ADP and ATP.
Results obtained at pH 2.0 and 3.5 are shown in Figure S1. At pH 2 hexose-phosphates were not
separated from ATP and at pH 3.5 they partially overlapped with ATP and ADP. Therefore, [2,22H ]G6P and [1,6-13C ]F6P were purified by HVPE at pH 6.5. Neutral sugars (sucrose, glucose and
2
2
fructose) remain at or near the origin under such conditions29. HVPE at pH 3.5 gave a good
resolution of G1P from Pi, so [U-13C]G1P was purified by electrophoresis at this pH, where sucrose
and fructose also remained at or near the origin (Figure 2B). Quantification by enzymatic or LCMS/MS analysis of the HVPE-purified isotopomers indicated final yields of 87 % for [2,2-2H2]G6P
and >95 % for [1,6-13C2]F6P and [U-13C]G1P (Table S2). In biological samples, naturally occurring
(M+2) isotopomers will represent approx. 2% of the total F6P (Table 1), therefore, isotopic
interference with the [1,6-13C2]F6P SIL-IS will be in a similar range to pipetting error and, as
discussed above for G6P, this was judged to be acceptable. Isotopic interference from G1P (M+6)
should be negligible (Table 1).
Synthesis of [13C11, 15N2]UDPG
9
UDPG is the glucosyl donor for synthesis of sucrose and cellulose in plants, and the precursor of
glycogen in animals. As noted above, potential for isotopic interference was particularly high for
UDPG, making it desirable to synthesize an isotopomer with an m/z of at least M+3 (<1 % natural
abundance; Table 1). In vivo, UDPG is synthesized primarily by UDP-glucose pyrophosphorylase
(UGPase), using UTP and G1P as substrates37, and unlike other UDPG-metabolizing enzymes (e.g.
sucrose synthase), UGPase is commercially available. Labelled UDPG could potentially have been
synthesized from the [U-13C]G1P, produced as described above, but use of commercially available
[13C9,15N2]UTP avoided the need to pre-synthesize labelled G1P as a substrate. For the reaction,
[13C9,15N2]UTP was incubated with a stoichiometric excess of unlabelled G1P and UGPase to
produce [13C9,15N2]UDPG and inorganic pyrophosphate (PPi) (Figure 1D). Alkaline
pyrophosphatase was also included to drive the UGPase reaction to completion by irreversibly
hydrolyzing the PPi by-product to Pi. [13C9,15N2]UDPG was purified by HVPE at pH 3.5, which gave
a clear separation from G1P, UTP, UDP and Pi (Figure 2C). The final yield of [13C9,15N2]UDPG after
HVPE purification was 86 % (Table S2). Isotopic interference from naturally occurring M+11
isotopomers of UDPG in biological samples should be negligible (Table 1).
Synthesis of 13C-labelled pentose-phosphates
Pentose-phosphates are intermediates of the oxidative pentose-phosphate pathway and
ribonucleotide synthesis in almost all organisms, and of the Calvin-Benson cycle in plants. A
reaction scheme was designed to synthesize several pentose-phosphates, starting from
commercially available [2,3,4,5-13C4]ribose (Figure 1E). The first reaction in the sequence was the
synthesis of [2,3,4,5-13C4]R5P by phosphorylation of [2,3,4,5-13C4]ribose by ribokinase38, with a
stoichiometric excess of ATP. Recombinant E. coli ribokinase27 was kindly provided by Dr. Roman
S. Esipov (Shemyakin–Ovchinnikov Institute of Bioorganic Chemistry, Russian Academy of
Sciences), as there were no commercially available forms of the enzyme. Ru5P and Xu5P are
epimers that differ only in the conformation of a hydroxyl group on C-2, and are poorly resolved
by most chromatographic systems, including our reverse-phase LC-MS/MS platform3. Therefore,
we considered that a mixture of labelled Ru5P and Xu5P would be suitable for use as a combined
SIL-IS. Following phosphorylation of [2,3,4,5-13C4]ribose with ribokinase, as described above,
[2,3,4,5-13C4]R5P was converted to [2,3,4,5-13C4]Ru5P and then [2,3,4,5-13C4]Xu5P using
commercially available ribose-5-phosphate isomerase39 and ribulose-phosphate-3-epimerase40
(Figure 1E).
The pentose-phosphates could be separated from ADP, the by-product of the initial ribokinase
reaction by HVPE at pH 3.5, but not from unreacted ATP (Figure 2D). Therefore, the reaction
mixtures were incubated with nucleoside-triphosphate phosphatase (apyrase 1) and nucleosidediphosphate phosphatase (apyrase 2) to hydrolyse unreacted ATP and the ADP by-product to
AMP and Pi, which were well resolved from the pentose-phosphates by HVPE at pH 3.5 (Figure
2D). As an example, a preparative electrophoretogram for purification of [2,3,4,5-13C4]R5P is
shown in Figure S2. Although the initial ribokinase reaction gave 79 % conversion of [2,3,4,513C ]ribose to [2,3,4,5-13C ]R5P, after HVPE purification the final yield of the latter was only 24 %,
4
4
while the final yield of the mixed epimers, [2,3,4,5-13C4]Ru5P and [2,3,4,5-13C4]Xu5P, was 53 %
(Table S2). All of the purified pentose-phosphate isotopomers are four mass units greater than
10
the unlabelled forms, so isotopic interference from the naturally isotopomers is expected to be
negligible (<0.02 %; Table 1).
Synthesis of [U-13C]dihydroxyacetone phosphate (DHAP)
DHAP is a glycolytic intermediate and precursor of glycerol 3-phosphate in bacteria, animals and
plants, as well as being an intermediate of the Calvin-Benson cycle and sucrose synthesis in
plants. Commercially available [U-13C]FBP was incubated with aldolase to generate an equimolar
mixture of [U-13C]DHAP and [U-13C]glyceraldehyde 3-phosphate (GAP) (Figure 1F). In the
presence of triose phosphate isomerase (TPI), this was converted to an equilibrium mixture
containing >95 % [U-13C]DHAP 41, which was separated from unreacted FBP and GAP by HVPE at
pH 6.5 (Figure 2E). The final yield of [U-13C]DHAP after HVPE purification was 37 %, mainly
reflecting the inefficiency of the aldolase reaction (Table S2). Isotopic interference from the
naturally occuring M+3 isotopomer will be negligible (<0.05 %; Table 1).
Synthesis of [13C10, 15N2]AMP
AMP is present in all organisms as a precursor of ATP and often is a metabolite signal of energy
status. Commercially available [13C10,15N5]ATP was successively hydrolyzed to [13C10,15N5]ADP and
then [13C10,15N5]AMP by incubation with apyrase 1 and apyrase 242 (Figure 1G). AMP was well
resolved from ADP, ATP, Pi and PPi by HVPE at pH 6.5 (Figure 2F). The final yield of [13C10,15N5]AMP
after HVPE purification was 68%, essentially reflecting the efficiency of the apyrase reactions (67
%; Table S2). Isotopic interference from the naturally occuring M+15 isotopomer will be negligible
(Table 1).
Signal overlap between natural isotopomers and SIL-IS
To validate the commercially available and synthesized SIL-ISs for routine use, equal amounts of
each SIL-IS and its corresponding unlabelled isotopomer were analyzed by LC-MS/MS in selected
reaction monitoring (SRM) mode. With the exceptions of G6P, F6P, FBP and aspartate, the
unlabelled isotopomer made no detectable contribution to the ion intensity signal from the
corresponding SIL-IS (Table S3). The naturally occurring isotopomers of G6P and F6P contributed
1.41 % of the total signal for the M+2 isotopomers ([6,6-2H2]G6P and [1,6-13C2]F6P) selected as
SIL-IS (Table S3). This is somewhat less than the predicted interference of 2.06 % (Table 1), which
was calculated without taking fragmentation into consideration. The 0.1 % contribution from
aspartate to the M+3 ([2,3,3-2H3]aspartate) isotopomer signal, and 0.4 % contribution of FBP to
the M+6 ([U-13C]FBP) isotopomer signal, were both slightly higher than the predicted values
(0.04% and <0.002%, respectively; Table 1), but too low to have any significant impact on
measurements of these metabolites.
In the reciprocal direction, the SIL-IS contributed <0.2 % (in most cases zero) of the ion intensity
signal for the corresponding unlabelled metabolite (Table S3). The only exception was FBP, with
3 % of the signal originating from the [U-13C]FBP, presumably reflecting contamination of the
commercially supplied isotopomer with unlabelled FBP. Despite the higher than expected cross
contamination, the [U-13C]FBP should not significantly interfere with FBP measurements as long
as the SIL-IS is not added in large excess of the amount of FBP in the sample. Chromatogram
11
analysis software often includes a function to correct automatically for known levels of cross
contamination, for example, the “correction for isotope contribution” function in the LCquanTM
software from Thermo-Scientific (www-thermoscientific.com). Otherwise data can be corrected
manually using a nonlinear calibration function43.
Correction of ion suppression or enhancement using SIL-IS
Ion suppression or enhancement is usually dependent on the concentration of interfering
molecules in biological extracts. Therefore, the effectiveness of the SIL-IS for quantification and
correction of matrix effects was assessed in extracts from diverse biological sources over a wide
range of dilutions. Extracts were analyzed from leaves of eudicotyledonous (Arabidopsis and
tobacco) and monocotyledonous (maize and wheat) plant species, green and red tomato fruits,
potato tubers and Chlamydomonas reinhardtii cells. Extracts from E. coli and Baker´s yeast
(Saccharomyces cerevisiae) cells and mouse liver were also tested, to assess the broader utility
of the SIL-IS for metabolite analysis in non-plant samples. Several dilutions were prepared from
each extract and spiked with the same amounts of SIL-IS prior to measurement. Quantification
of metabolites was performed with and without correction by SIL-IS in each sample. Data for G6P
are shown in Figure 3 as an example of the analysis, with equivalent data for all the other
measured metabolites presented in Figure S3 and Table S4. Information on the LC-MS/MS
settings and amount of SIL-IS used to spike the samples is given in Table S1.
In almost all of the biological extracts tested, there was a clear trend for the measured amount
of G6P in the non-spiked extract to be lower in the less-diluted extract (Figure 3), suggesting a
higher degree of ion suppression. The difference between the level of G6P measured in 100-fold
diluted extract compared to undiluted extract (or the lowest dilution measured) ranged from
about 15 % for Chlamydomonas reinhardtii to almost 60 % for Arabidopsis leaf extract, with most
of the other tissues being near the upper end of this range. In maize and tobacco leaf, tomato
fruit and Chlamydomonas reinhardtii extracts, it was not possible to quantify G6P in the undiluted
extract due to excessive distortion of the G6P peak in the LC-MS/MS chromatograms, indicating
an even stronger disturbance by matrix effects.
In contrast, the corrected measurements from extracts spiked with the [2,2-2H2]G6P SIL-IS were
much less variable, generally differing by less than 10 % across the whole range of dilutions. In
all of the samples there was a significant (P<0.05) difference between the measurements in the
spiked and non-spiked extracts, except at the highest dilutions, where the measurements tended
to converge (Figure 3). Chlamydomonas reinhardtii cell extracts were a notable exception. Here
a consistently higher level of G6P was measured in the spiked samples, even at the highest
dilution. The cells were harvested directly from the bio-fermenter into cold (-70°C) methanol
without separation from the culture medium, to ensure rapid quenching of metabolism so that
metabolite levels accurately reflected those in the living, illuminated cells15,30. Thus, the samples
contained cell extract effectively diluted with culture medium, mostly inorganic salts including
chloride, which is known to interfere with LC-MS/MS analysis15. Yeast cell extracts presented the
smallest differences between spiked and non-spiked extracts for the G6P measurements,
indicating only a marginal benefit from spiking with the SIL-IS. Broadly similar patterns were
observed for most of the other measured metabolites (Figure S3). Of special note were a few
12
examples of signal enhancement, where the measured amount of the metabolite in non-spiked
extract was apparently higher than in the SIL-IS-spiked extract. This phenomenon was observed
for malate, glycerate and succinate in yeast and E. coli cell extracts (Figures S3J-L). Ion
enhancement, i.e. greater efficiency of ionisation in the extract than in the standards used for
calibration, could be a contributory factor. Ionisation-independent matrix effects related to the
performance of the LC column, especially its binding capacity for these rather abundant organic
acids, might also be a factor.
In summary, these results clearly demonstrate the utility of using SIL-IS for correction of matrix
effects when measuring metabolites by LC-MS/MS from a diverse range of biological materials,
including photosynthetic and non-photosynthetic tissues from plants, animal tissue and bacterial
and fungal cells.
CONCLUSIONS
Spiking of biological extracts with SIL-IS is the method of choice for correction of matrix effects
when measuring metabolites by LC-MS/MS. The main criteria for selection of suitable
isotopomers to use as SIL-IS are: (i) minimal overlap (preferably none) with the naturally
occurring isotopomers expected to be present in biological extracts and (ii) no significant
contamination of the SIL-IS by unlabelled isotopomer of the corresponding metabolite, or indeed
of other metabolite being measured. Isotopomers that meet these criteria are commercially
available for some metabolites, but not for most phosphorylated intermediates. SIL-IS for a range
of hexose-, pentose-, and triose-phosphates, and AMP were synthesized enzymatically, purified
by HVPE, and validated by LC-MS/MS measurements in a wide variety of plant and non-plant
tissue or cell extracts. This suite of SIL-IS greatly expands the coverage of photosynthetic and
respiratory intermediates, allowing more accurate and reliable LC-MS/MS-based measurements
of metabolites, not only in plants, but also in samples of bacterial, fungal or animal origin.
SUPPORTING INFORMATION
Detailed description of plant and cell culture growth conditions, validation of plant sampling for
accurate metabolite quantification, additional tables and figures. This material is available free
of charge via the Internet at http://pubs.acs.org/.
AUTHOR CONTRIBUTIONS
SA, JEL and MS designed the experiments; SA performed the isotopomer abundance analysis; SA
and MG synthesized isotopically labelled compounds, which were purified by SCF. MMFFF, DV
and TMA grew and harvested plant material and microbial cells; SA performed the LC-MS/MS
analysis. SA and JEL drafted the manuscript and all authors have given approval to the final
version of the manuscript.
ACKNOWLEDGEMENTS
We thank Roman Esipov, Saleh Alseekh, Joerg Schenk and Carlos Figueroa for generously
providing us with recombinant E. coli ribokinase, tomato fruits, mouse liver and yeast cells,
respectively.
13
FUNDING
SA was supported by the European Commission FP7 collaborative project 3to4 (contract No.
289582 to MS), TM was supported by the German Federal Ministry of Education and Science
GoFORSYS project (contract No. 0313924 to MS), MMFFF was supported by a PhD studentship
from the International Max Planck Research School “Primary Metabolism and Plant Growth”, and
JL was supported by the Max Planck Gesellschaft.
The authors declare no competing financial interest.
REFERENCES
1.
Grebe, S. K., Singh, R. J., Clin. Biochem. Rev. 2011, 32, 5-31.
2.
Lunn, J. E.; Feil, R.; Hendriks, J. H.; Gibon, Y.; Morcuende, R.; Osuna, D.; Scheible, W. R.;
Carillo, P.; Hajirezaei, M. R., Stitt, M., Biochem. J. 2006, 397, 139-148.
3.
Arrivault, S.; Guenther, M.; Ivakov, A.; Feil, R.; Vosloh, D.; van Dongen, J. T.; Sulpice, R.,
Stitt, M., Plant J. 2009, 59, 824-839.
4.
Buescher, J. M.; Moco, S.; Sauer, U., Zamboni, N., Anal. Chem. 2010, 82, 4403-4412.
5.
Burgos, A.; Szymanski, J.; Seiwert, B.; Degenkolbe, T.; Hannah, M. A.; Giavalisco, P.,
Willmitzer, L., Plant J. 2011, 66, 656-668.
6.
Broeckling, C. D.; Huhman, D. V.; Farag, M. A.; Smith, J. T.; May, G. D.; Mendes, P.; Dixon,
R. A., Sumner, L. W., J. Exp. Bot. 2005, 56, 323-336.
7.
Yonekura-Sakakibara, K.; Tohge, T.; Matsuda, F.; Nakabayashi, R.; Takayama, H.; Niida, R.;
Watanabe-Takahashi, A.; Inoue, E., Saito, K., Plant Cell 2008, 20, 2160-2176.
8.
Giavalisco, P.; Li, Y.; Matthes, A.; Eckhardt, A.; Hubberten, H.-M.; Hesse, H.; Segu, S.;
Hummel, J.; Köhl, K., Willmitzer, L., Plant J. 2011, 68, 364-376.
9.
Liebeke, M.; Dörries, K.; Meyer, H., Lalk, M., in Functional Genomics, ed. M. Kaufmann, C.
Klinger. Springer New York, 2012, vol. 815, pp 377-398.
10.
Crutchfield, C. A.; Lu, W.; Melamud, E., Rabinowitz, J. D., in Methods in Enzymology, ed.
W. Jonathan, G. Christine, R. F. Gerald. Academic Press, 2010, vol. Volume 470, pp 393-426.
11.
(a) Kebarle, P., Tang, L., Anal. Chem. 1993, 65, 972A-986A; (b) Tang, L., Kebarle, P., Anal.
Chem. 1993, 65, 3654-3668.
12.
Bonfiglio, R.; King, R. C.; Olah, T. V., Merkle, K., Rapid Commun. Mass Spectrom. 1999, 13,
1175-1185.
13.
King, R.; Bonfiglio, R.; Fernandez-Metzler, C.; Miller-Stein, C., Olah, T., J. Am. Soc. Mass
Spectrom. 2000, 11, 942-950.
14.
Trufelli, H.; Palma, P.; Famiglini, G., Cappiello, A., Mass Spectrom. Rev. 2011, 30, 491-509.
15.
Tohge, T.; Mettler, T.; Arrivault, S.; Carroll, A. J.; Stitt, M., Fernie, A. R., Front. Plant Sci.
2011, 2, article 61.
16.
Chambers, E.; Wagrowski-Diehl, D. M.; Lu, Z., Mazzeo, J. R., J. Chromatogr. B Analyt.
Technol. Biomed. Life Sci. 2007, 852, 22-34.
17.
Hall, T. G.; Smukste, I.; Bresciano, K. R.; Wang, Y.; McKearn, D., Savage, R. E., in Tandem
Mass Spectrometry - Applications and Principles, ed. J. K. Prasain. 2012.
18.
Chen, J.; Yang, L.; Kapron, J. T.; Ma, L.; Pace, E.; Van Pelt, C. K., Rudewicz, P. J., J.
Chromatogr. B Analyt. Technol. Biomed. Life Sci. 2004, 809, 205-210.
14
19.
Sancho, J. V.; Pozo, O. J.; Lopez, F. J., Hernandez, F., Rapid Commun. Mass Spectrom. 2002,
16, 639-645.
20.
Schuhmacher, J.; Zimmer, D.; Tesche, F., Pickard, V., Rapid Commun. Mass Spectrom.
2003, 17, 1950-1957.
21.
Buhrman, D. L.; Price, P. I., Rudewiczcor, P. J., J. Am. Soc. Mass Spectrom. 1996, 7, 10991105.
22.
Matuszewski, B. K.; Constanzer, M. L., Chavez-Eng, C. M., Anal. Chem. 2003, 75, 30193030.
23.
Xu, R. N.; Fan, L.; Rieser, M. J., El-Shourbagy, T. A., J. Pharm. Biomed. Anal. 2007, 44, 342355.
24.
Cuadros-Rodríguez, L.; Bagur-González, M. G.; Sánchez-Viñas, M.; González-Casado, A.,
Gómez-Sáez, A. M., J. Chromatogra. A 2007, 1158, 33-46.
25.
Berg, T.; Karlsen, M.; Øiestad, Å. M. L.; Johansen, J. E.; Liu, H., Strand, D. H., J. Chromatogr.
A 2014, 1344, 83-90.
26.
Olsson, A. O.; Baker, S. E.; Nguyen, J. V.; Romanoff, L. C.; Udunka, S. O.; Walker, R. D.;
Flemmen, K. L., Barr, D. B., Anal. Chem. 2004, 76, 2453-2461.
27.
Chuvikovsky, D. V.; Esipov, R. S.; Skoblov, Y. S.; Chupova, L. A.; Muravyova, T. I.;
Miroshnikov, A. I.; Lapinjoki, S., Mikhailopulo, I. A., Bioorg. Med. Chem. 2006, 14, 6327-6332.
28.
Fry, S. C., The Growing Plant Cell Wall: Chemical and Metabolic Analysis. 2000.
29.
Fry, S. C., in The Plant Cell Wall: Methods and Protocols, Methods in Molecular Biology,
ed. Z. A. Popper. 2011, vol. 715, pp 55-80.
30.
Mettler, T.; Muhlhaus, T.; Hemme, D.; Schottler, M. A.; Rupprecht, J.; Idoine, A.; Veyel, D.;
Pal, S. K.; Yaneva-Roder, L.; Winck, F. V.; Sommer, F.; Vosloh, D.; Seiwert, B.; Erban, A.; Burgos,
A.; Arvidsson, S.; Schonfelder, S.; Arnold, A.; Gunther, M.; Krause, U.; Lohse, M.; Kopka, J.;
Nikoloski, Z.; Mueller-Roeber, B.; Willmitzer, L.; Bock, R.; Schroda, M., Stitt, M., Plant Cell 2014,
26, 2310-2350.
31.
Blaby, I. K.; Glaesener, A. G.; Mettler, T.; Fitz-Gibbon, S. T.; Gallaher, S. D.; Liu, B.; Boyle,
N. R.; Kropat, J.; Stitt, M.; Johnson, S.; Benning, C.; Pellegrini, M.; Casero, D., Merchant, S. S., Plant
Cell 2013, 25, 4305-4323.
32.
Merlo, L.; Geigenberger, P.; Hajirezaei, M., Stitt, M., J. Plant Physiol. 1993, 142, 392-402.
33.
Rosman, K. J. R., Taylor, P. D. P., in Pure Appl. Chem. 1998, vol. 70, pp 217-235.
34.
Patiny, L., Borel, A., J. Chem. Inf. Model. 2013, 53, 1223-1228.
35.
Gu, H.; Wang, J.; Aubry, A. F.; Jiang, H.; Zeng, J.; Easter, J.; Wang, J. S.; Dockens, R.; Bifano,
M.; Burrell, R., Arnold, M. E., Anal. Chem. 2012, 84, 4844-4850.
36.
Mieyal, J. J., Abeles, R. H., in The Enzymes, ed. P. D. Boyer. Academic Press: New York,
1972, pp 515-532.
37.
Kim, H.; Choi, J.; Kim, T.; Lokanath, N. K.; Ha, S. C.; Suh, S. W.; Hwang, H. Y., Kim, K. K., Mol.
Cells 2010, 29, 397-405.
38.
Agranoff, B. W., Brady, R. O., J. Biol. Chem. 1956, 219, 221-229.
39.
Zhang, R.; Andersson, C. E.; Savchenko, A.; Skarina, T.; Evdokimova, E.; Beasley, S.;
Arrowsmith, C. H.; Edwards, A. M.; Joachimiak, A., Mowbray, S. L., Structure 2003, 11, 31-42.
40.
Lee, L. V.; Vu, M. V., Cleland, W. W., Biochemistry (Mosc). 2000, 39, 4808-4820.
41.
Veech, R. L.; Raijman, L.; Dalziel, K., Krebs, H. A., Biochem. J. 1969, 115, 837-842.
15
42.
Kettlun, A. M.; Uribe, L.; Calvo, V.; Silva, S.; Rivera, J.; Mancilla, M.; Antonieta, M.;
Valenzuela, Traverso-Cori, A., Phytochemistry 1982, 21, 551-558.
43.
Rule, G. S.; Clark, Z. D.; Yue, B., Rockwood, A. L., Anal. Chem. 2013, 85, 3879-3885.
Figure 1: Synthesis of SIL-IS. (A) [6,6-2H2]G6P, (B) [1,6-13C2]F6P, (C) [U-13C]G1P, (D)
[13C11,15N2]UDPG, (E) [2,3,4,5-13C4]R5P/Xu5P/Ru5P, (F) [U-13C]DHAP and (G) [13C10,15N5]AMP. The
isotopically-labelled elements are indicated in red. Structures were made with ACD/ChemSketch
(http://www.acdlabs.com/resources/freeware/chemsketch/).
16
Figure 2: Preparative HVPE for substrates, products and for enzymatic reaction mixtures.
Separation of compounds involved in the synthesis of [6,6-2H2]G6P, [1,6-13C2]F6P (A), [U-13C]G1P
(B), [13C11,15N2]UDPG (C), [2,3,4,5-13C4]R5P (D), [U-13C]DHAP (E) and [13C10,15N5]AMP (F).
Compounds run are listed along the origin of each electrophoretogram. HVPE were performed
at pH 3.5 for B, C and D and at pH 6.5 for A, E and F. Electrophoresis was conducted at 4.5 kV for
17
30 min (pH 3.5) or 35 min (pH 6.5). The coloured marker Orange G (orange dots) was added into
each sample before electrophoresis except in A where it was loaded alternatively with the
samples. Phosphate-containing compounds were stained with molybdate reagent. Sucrose and
fructose in B were stained with AgNO3. Some of the fainter spots were manually outlined with
dotted lines, with the synthesized IS shown in red. The origin of each electrophoretogram is
indicated by a solid grey line.
Fig.
A
B
C
D
E
F
pH
6.5
3.5
3.5
3.5
6.5
6.5
kV
4.5
4.5
4.5
4.5
4.5
4.5
minutes
30
30
30
30
40
30
18
Figure 3: Quantification of G6P with (filled circles) or without (open circles) SIL-IS in various
biological materials. Measurements were performed in extracts from leaves of Arabidopsis,
maize, tobacco and wheat diluted 100-, 50-, 20-, 10-fold (indicated on the x-axis as 0.01, 0.02,
0.05 and 0.1, respectively) and undiluted (1). Green and red tomato fruits and potato tuber
extracts were additionally diluted 5-fold (0.2). Extracts from mouse liver, Chlamydomonas
reinhardtii, yeast and E. coli were measured after 50-, 20-, 10-, 5- and 2-fold dilutions (indicated
on the x-axis as 0.02, 0.05, 0.1, 0.2 and 0.5, respectively), undiluted (1) and in 2-fold (2)
concentrated extracts. Data shown are means ± SD (n = 3, except for undiluted potato tuber
extracts where n = 2). The original data are presented in Supplementary Table S4. Significant
differences (according to Student’s t test) from values obtained with signal corrected by SIL-IS
are indicated by asterisks (*P < 0.05, **P < 0.01 and ***P < 0.001).
19
Table 1: Isotope distribution for parent ions. Abundances were obtained using an online
calculator (34, http://www.chemcalc.org/main).
Compounds
Parent ion
([M-H]-)
formula
Mass
(M)
M
M+1
Abundance (%)
M+2
M+3
M+4
G6P/F6P/G1P
C6H12O9P -
259
100
6.9703
2.0562
0.1307
UDPG
C15H23N2O17P2 -
565
100
17.8664
4.9997
R5P/Ru5P/Xu5P
C5H10O8P -
229
100
5.8276
DHAP
C3H6O6P -
169
100
AMP
C10H13N5O7P -
346.1
FBP
C6H13O12P2 -
2-OG
M+5
M+6
0.0184
ˉ
ˉ
0.7009
0.1114
0.0113
ˉ
1.7838
0.0965
0.0137
ˉ
ˉ
3.5423
1.2778
0.0424
0.0063
ˉ
ˉ
100
13.0585
2.2285
0.2150
0.0193
ˉ
ˉ
339
100
7.0961
2.6812
0.1766
0.0322
0.0018
ˉ
C4CH5O5 -
145
100
5.6558
1.1580
0.0584
0.0054
ˉ
ˉ
Malate
C4H5O5 -
133
100
4.5743
1.1085
0.0460
0.0042
ˉ
ˉ
Glycerate
C3H5O4 -
105
100
3.4546
0.8639
0.0267
0.0025
ˉ
ˉ
Succinate
C4H5O4 -
117
100
4.5362
0.9013
0.0356
0.0025
ˉ
ˉ
Glutamate
C5H8NO4 -
146
100
6.0176
0.9720
0.0487
0.0025
ˉ
ˉ
100
4.9130
0.9176
0.0386
0.0025
ˉ
ˉ
Aspartate
C4H6NO4
-
132
20
For TOC only