Download Central carbon metabolism of Saccharomyces

Eur. J. Biochem. 268, 2464±2479 (2001) q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae explored by
biosynthetic fractional 13C labeling of common amino acids
Hannu Maaheimo1,2,*, Jocelyne Fiaux3,*, Z. Petek CËakar4,², James E. Bailey 4, Uwe Sauer 4 and
Thomas Szyperski1
1
Department of Chemistry, University at Buffalo, The State University of New York, NY, USA; 2VTT Biotechnology, Espoo, Finland;
Institut fuÈr Molekularbiologie und Biophysik and 4Institut fuÈr Biotechnologie, EidgenoÈssische Technische Hochschule HoÈnggerberg,
ZuÈrich, Switzerland
3
Aerobic and anaerobic central metabolism of Saccharomyces cerevisiae cells was explored in batch cultures on a
minimal medium containing glucose as the sole carbon
source, using biosynthetic fractional 13C labeling of
proteinogenic amino acids. This allowed, firstly, unravelling of the network of active central pathways in cytosol
and mitochondria, secondly, determination of flux ratios
characterizing glycolysis, pentose phosphate cycle, tricarboxylic acid cycle and C1-metabolism, and thirdly, assessment of intercompartmental transport fluxes of pyruvate,
acetyl-CoA, oxaloacetate and glycine. The data also revealed
that alanine aminotransferase is located in the mitochondria,
and that amino acids are synthesized according to documented pathways. In both the aerobic and the anaerobic
regime: (a) the mitochondrial glycine cleavage pathway is
active, and efflux of glycine into the cytosol is observed; (b)
the pentose phosphate pathways serve for biosynthesis only,
i.e. phosphoenolpyruvate is entirely generated via glycolysis; (c) the majority of the cytosolic oxaloacetate is
synthesized via anaplerotic carboxylation of pyruvate; (d)
the malic enzyme plays a key role for mitochondrial
pyruvate metabolism; (e) the transfer of oxaloacetate from
the cytosol to the mitochondria is largely unidirectional,
and the activity of the malate±aspartate shuttle and the
succinate-fumarate carrier is low; (e) a large fraction of the
mitochondrial pyruvate is imported from the cytosol; and
(f ) the glyoxylate cycle is inactive. In the aerobic regime,
75% of mitochondrial oxaloacetate arises from anaplerotic
carboxylation of pyruvate, while in the anaerobic regime,
the tricarboxylic acid cycle is operating in a branched
fashion to fulfill biosynthetic demands only. The present
study shows that fractional 13C labeling of amino acids
represents a powerful approach to study compartmented
eukaryotic systems.
Biosynthetically directed fractional (BDF) 13C labeling of
proteinogenic amino acids combined with 2D 13C-1H
correlation NMR spectroscopy (COSY) for analysis of
13
C± 13C scalar coupling fine structures and balancing of
contiguous carbon fragments within metabolic networks,
has been used as a powerful tool to investigate intermediary
metabolism of prokaryotic cells [1]. The desired fractional
13
C labeling is achieved by feeding a mixture of uniformly
13
C labeled and unlabeled compounds as the sole carbon
source for cellular growth [2±4]. An outstanding strength of
this approach is its ability to enable both a comprehensive
characterization of the network of active biosynthetic
pathways [1,5±7] and the accurate determination of
metabolic flux ratios [1,6,8±11] in a single experiment
(reviewed in [12]). This led us to name the approach
metabolic flux ratio (METAFoR) analysis by NMR [9,14].
Additional attractive features with regard to routine
applications are an inherently small demand for 13C labeled
source compounds, the high sensitivity of 2D 13C-1H
COSY, and the availability of a user-friendly program,
fcal, for rapid data analysis [14].
Keywords: Saccharomyces cerevisiae; central metabolism;
13
C NMR; METAFoR; metabolic engineering.
Correspondence to T. Szyperski, Department of Chemistry, University at Buffalo, The State University of New York, Buffalo, USA.
Fax:. 1 1716 6457338, Tel.: 1 1716 6456800 ext. 2245, E-mail: [email protected]
Abbreviations: cyt, cytosolic; mt, mitochondrial; BDF, biosynthetically directed fractional; METAFoR, metabolic flux ratio; TCA,
tricarboxylic acid; OxAc, oxaloacetate; OxGlt, 2-oxoglutarate; Prv, pyruvate; AcCoA, acetyl-coenzyme A; GriP, 3-phosphoglycerate; PPrv,
phosphoenolpyruvate; Ri5P, ribose 5-phosphate; E4P, erythrose 4-phosphate; PenPp, pentose phosphate pathway; SHMT, serine
hydroxymethyltransferase; GCV, glycine cleavage pathway; MDH, malate dehydrogenase.
Enzymes: acetolactate synthase (EC 4.1.3.18); alanine aminotransferase (EC 2.6.1.2); carnitine O-acetyltransferase (EC 2.3.1.7); fumarase
(EC 4.2.1.2.); glycine aminotransferase (EC 2.6.1.4); glycine cleavage system (EC 2.1.2.10); homocitrate synthase (EC 4.1.3.21); a-isopropylmalate
synthase (EC 4.1.3.12); malate dehydrogenase (EC 1.1.1.37); malic enzyme (EC 1.1.1.39/1.1.1.40); 2-oxoglutarate dehydrogenase (EC 1.2.4.2);
pyruvate carboxylase (EC 6.4.1.1); pyruvate decarboxylase (EC 4.1.1.1); pyruvate dehydrogenase (EC 1.2.4.1); pyruvate formate-lyase
(EC 2.3.1.54); serine hydroxymethyltransferase (EC 2.1.2.1); threonine aldolase (EC 4.1.2.5); transaldolase (EC 2.2.1.2); transketolase (EC 2.2.1.1).
*Note: these authors contributed equally to this paper
²Present address: Department of Biochemistry, Biozentrum, University of Basel, CH-4056 Basel, Switzerland
(Received 10 January 2001, accepted 26 February 2001)
q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2465
Previous applications of the BDF 13C labeling protocol
included its combination with metabolic flux balancing in
order to derive net fluxes in Bacillus subtilis [6], the
exploration of amino-acid biosynthesis in the halophilic
archaeon Haloarcula hispanica [7], the investigation of
Escherichia coli central carbon metabolism in microaerobic
bioprocesses [8], and the larger scale evaluation of the
impact of genetic (or environmental) modulations [9] and
the introduction of D2O into the growth medium [11] on
E. coli central carbon metabolism. These studies neatly
exemplified the value of BDF 13C labeling experiments
for studying prokaryotic central metabolism in both
fundamental and applied biotechnological research.
In the present publication, we describe the extension of
the BDF 13C labeling protocol for investigating eukaryotic
systems. The key challenge is quite evidently the compartmentation of the eukaryotic cell, which leads to a dissection
of central carbon metabolism in subnetworks localized in
either the cytosol or in organelles, e.g. mitochondria or
peroxisomes [15]. Yeasts, such as Saccharomyces cerevisiae, are eukaryotic model organisms that are pivotal for
many areas of biology, biomedicine and biotechnology.
Moreover, an outstanding body of experimental data with
respect to central and amino-acid metabolism is available
[16±18], and the genome of S. cerevisiae has been completely sequenced [19±21]. Hence, we decided to explore
central carbon metabolism of the yeast S. cerevisiae when
grown in batch cultures under glucose repression, in both
aerobic or anaerobic regimes.
M AT E R I A L S A N D M E T H O D S
Biosynthetically directed fractional
13
C labeling
S. cerevisiae strain CEN.PK 113±7D (MATa, URA3, HIS3,
LEU2, TRP1, MAL2±8c, SUC2) was obtained from
P. KoÈtter (Institute of Microbiology, Johann Wolfgang
Goethe-University, Frankfurt, Germany). The cells were
grown on a yeast minimal medium, containing 6.7 g´L21 of
yeast nitrogen base lacking amino acids (Difco), and 0.5%
(w/v) glucose consisting of a mixture of 10% (w/w) U-13C
labeled (Martek Biosciences Corporation, Columbia, USA)
and 90% (w/w) unlabeled glucose. Aerobic cultivations
were performed using a rotary shaker at 30 8C and
300 r.p.m., in 1-L baffled shake flasks containing 100 mL
medium. Anaerobic cultivations were carried out under
moderate shaking (140 r.p.m.) at 30 8C in 125 mL flasks
with tight rubber caps and 125 mL medium. The flasks
were flushed with nitrogen prior to cultivation.
Cell growth was monitored spectrophotometrically by
measuring the optical density of the cultures at 600 nm
(D600). The cells were harvested during the mid-exponential
phase at D600 ˆ 0.8±1.0 by centrifugation at 3000 g for
10 min using a benchtop centrifuge (Beckman, GS-6R).
The resulting pellets were resuspended in Tris/HCl (20 mm,
pH 7.6) and then centrifuged again at the previous settings,
after which the supernatant was discarded. The pellets were
vacuum-dried (Hetovac, VR-1) overnight to constant mass
and the amount of dried biomass was determined. The
biomass was then resuspended in 3 mL Tris/HCl (20 mm,
pH 7.6). Upon addition of 6 mL 6 m HCl, hydrolysis was
performed in sealed glass tubes at 110 8C for 24 h, after
which the solutions were filtered through 0.2-mm filters
(Millipore, Millex-GP). Finally, the hydrolysed biomass
was lyophilized. Approximately 60±70 mg of dried
hydrolysate were obtained from each cultivation.
NMR spectroscopy
The dried hydrolysates were dissolved in 0.1 m DCl in
D2O, and 2D 13C-1H COSY spectra [22,23] were acquired
for both aliphatic and aromatic resonances as described
[1,5,6,8] at 40 8C on a Bruker DRX 500 spectrometer. The
measurement time was < 8 h for each 2D spectrum, and
the spectra were processed using the program prosa [24].
The 13C abundance was determined from the resolved 13C
satellites in 1D 1H NMR spectra [1], and an overall degree
of 13C labeling of 10% was obtained for both the aerobic
and the anaerobic preparation. The degree of 13C labeling
was in close agreement with the composition of the
minimal medium, and was confirmed by analysis of the
13
C scalar coupling fine structure of Leu-b [1].
Data analysis
The program fcal [14] (R. W. Glaser; fcal v2.3.0) was
used for data analysis including both the integration of 13C
scalar fine structures and the calculation of relative
abundances, f, of intact carbon fragments arising from a
single source molecule of glucose [1]. Notably, the use of
fcal allowed integration of the fine structure of Lys-a,
which largely overlaps with the signal of Arg-a [1]. In
contrast to prokaryotic cells [1,12], the fine structure of
Lys-a provides unique information: only this carbon
encodes the 13C labeling pattern of cytosolic AcCoA (see
below). The equations relating the 13C fine multiplet
intensities to the f values allow derivation of `fragment
balancing' equations, which can provide accurate flux
information largely independent of the overall degree of
13
C incorporation [1,12].
The nomenclature used here to identify such fragments
has been described previously [1,5]. Briefly, f (1) represents
the fraction of molecules in which the observed carbon
atom and its neighbouring carbons originate from different
source molecules of glucose, and f (2) the fraction of
molecules in which the observed carbon atom and at least
one neighbouring carbon originate from the same source
molecule. For a central carbon in a C3 fragment that
exhibits different 13C± 13C scalar coupling constants with
the two attached carbons, f (2) represents the fraction of
molecules for which the central carbon and the carbon with
the smaller coupling come from the same source molecule,
while f (2*) is used if the carbon with the larger coupling
comes from the same source molecule as the observed
carbon. f (3) denotes the fraction of molecules in which the
observed carbon atom and both neighbours in the C3
fragment originate from the same glucose molecule.
Previous experiments with prokaryotic cells have provided ample support for the suggestion that central carbon
metabolism operates in a quasi steady-state manner during
the exponential growth phase [7,9,11,14]. Consequently, the
quasi steady-state assumption was adopted for the present
study.
2466 H. Maaheimo et al. (Eur. J. Biochem. 268)
q FEBS 2001
Biochemical reaction network for S. cerevisiae
At the outset of this study, we compiled the currently
available knowledge about central carbon metabolism of
S. cerevisiae from literature [15±18] and databases (e.g.
http://www.expasy.ch/cgi-bin/search-biochem-index; http://
wit.mcs.anl.gov/WIT2/CGI/index.cgi; http://www.genome.
ad.jp/kegg/metabolism.html; http://www.rz.uni-frankfurt.
de/FB/fb16/mikro/euroscarf/index.html) in order to construct a biochemical reaction network suited for the
presently chosen growth conditions where glucose served
as the sole carbon source (Fig. 1). The salient feature of
eukaryotic central carbon metabolism is apparently its
dissection into cytosolic and mitochondrial subnetworks,
connected by intercompartmental transport of metabolites
[15±18]. Glycolysis and the pentose phosphate pathway
(PenPp) are located in the cytosol, and the tricarboxylic
acid (TCA) cycle operates in the mitochondria where
respiration takes place [25±27]. Oxaloacetate (OxAc),
pyruvate (Prv) and acetyl-CoA (AcCoA) are present in
both compartments, and systems for their transport across
the mitochondrial membrane have been identified [27±30].
Hence, cytosolic (cyt) and mitochondrial (mt) pools are
distinguished throughout for these three key intermediates.
Oxaloacetate, pyruvate and acetyl-CoA
In S. cerevisiae, OxAc is produced both as an intermediate
of the TCA cycle in the mitochondrial matrix from malate,
thus yielding mt-OxAc [25,26], and from cyt-Prv by the
action of pyruvate carboxylase in the cytosol [31,32]
yielding cyt-OxAc (Fig. 1). To enable the anaplerotic
supply of the TCA cycle, cyt-OxAc is transferred across
the mitochondrial membranes by the oxaloacetate carrier
protein [30] in order to refill the mt-OxAc pool. The
transport is driven by the proton motive force at the inner
mitochondrial membrane, i.e. oxaloacetate is actively transferred across the membrane in symport with protons
(Fig. 1). This mode of operation indicates directional
transfer from the cytosol to the mitochondria.
cyt-Prv is produced in the cytosol by glycolysis [26] and
a proportion of it is transported into the mitochondria [27]
yielding mt-Prv, which may also be synthesized from
malate by the malic enzyme. In S. cerevisiae, this enzyme
can involve either NADH or NADPH as a cofactor [33].
The gene encoding the malic enzyme has recently been
identified, and the enzyme was shown to be located
exclusively in mitochondria [34]. Pyruvate is transported
from the cytosol into the mitochondria by the pyruvate
(monocarboxylate) carrier protein [35]. The transport is
actively driven by the mitochondrial proton motive force
(Fig. 1), suggesting a largely unidirectional transport from
the cytosol into the mitochondria.
mt-AcCoA and cyt-AcCoA can be derived from mt-Prv
and cyt-Prv, respectively, either by the pyruvate dehydrogenase complex in the mitochondria [36], or via a cytosolic
`by-pass pathway', the first reaction of which is catalyzed
by pyruvate decarboxylase [37]. When S. cerevisiae cells
have access to glucose, AcCoA is expected to originate
exclusively from Prv [25]. AcCoA can cross the inner
mitochondrial membrane via the `carnitine shuttle' [28,29].
This shuttle consists of the carnitine O-acetyltransferase
[38,39] catalyzing the exchange of acetyl groups between
Fig. 1. Network of active biochemical pathways constructed for
S. cerevisiae cells grown with glucose as the sole carbon source. The
network was inferred from current literature and data deposited in
databases (see text). The dissection of central carbon metabolism into
two subnetworks is apparent: glycolysis and the pentose phosphate
pathway take place in the cytosol, while the TCA cycle takes place in
the mitochondrial matrix. Irreversible reactions, represented by
unidirectional arrows, have been inferred from (a) estimations of
in vivo metabolite concentrations [86, 87] and in vitro free enthalpies
(http://wwwbmcd.nist.gov:8080/enzyme), and (b) the experimental
NMR data presented in this publication. 13C labeling patterns of
metabolites shown in ellipses can be assessed through analysis
of proteinogenic amino acids (Fig. 2). Carrier systems for transport
of metabolites between cytosol and mitochondrial matrix are
represented by grey circles. For the Prv and OxAc carrier systems,
active transport by the mitochondrial proton motive force is indicated,
and the corresponding back-fluxes are represented by dashed arrows.
The AcCoA carrier system (AT) operates by facilitated diffusion.
Abbreviations: AcCoA, Acetyl-Coenzyme A; AcH, acetaldehyde;
AcOH, acetic acid; EtOH, ethanol; E4P, erythrose 4-phosphate; F6P,
fructose 6-phosphate; G6P, glucose 6-phosphate; Fum, fumarate; Gly,
glycine; G3P, glyceraldehyde 3-phosphate; GriP, 3-phosphoglycerate;
Mal, malate; OxGlt, 2-oxoglutarate; Prv, pyruvate; PPrv, phosphoenolpyruvate; Ri5P, ribose-5-phosphate; Ru5P, ribulose 5-phosphate;
S7P, seduheptulose 7-phosphate; Ser, serine; Suc, succinate; Xu5P,
xylulose 5-phosphate. For Prv, AcCoA and OxAc, cytosolic (cyt) and
mitochondrial (mt) pools are indicated separately.
carnitine and AcCoA (which itself cannot pass the
mitochondrial membrane), and the acetyl-translocase. The
acetyl-translocase represents a carnitine/acetylcarnitine
antiport system which serves to balance the cytosolic and
mitochondrial AcCoA pools by facilitated diffusion.
Glyoxylate pathway
When S. cerevisiae cells are grown on a medium containing glucose as the sole carbon source, the cytosolic
q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2467
enzymes constituting the glyoxylate pathway [40,41] are
not expressed [40,42]. Notably, both cytosolic and peroxisomal malate dehydrogenase isozymes (MDH2 and
MDH3, respectively), which solely serve to establish the
glyoxylate cycle, are glucose-repressed along with the other
glyoxylate cycle enzymes [43].
C1 metabolism
The major one-carbon donor in S. cerevisiae is serine,
which is reversibly cleaved into glycine and a C1 unit by
serine hydroxymethyl transferase (SHMT). Mitochondrial
and cytoplasmic SHMT isozymes exist [44,45], and their
action is complemented by the mitochondrial glycine
cleavage pathway (GCV) which interconverts glycine into
a C1 unit and CO2 [46].
Biosynthesis of proteinogenic amino acids in
S. cerevisiae
In order to recruit proteinogenic amino acids as probes to
study central carbon metabolism [1,4], their biosynthetic
pathways must be available. When applying this approach
to eukaryotic cells, it is necessary to identify which of the
cytosolic or mitochondrial pools of Prv, AcCoA or OxAc
are used for the biosynthesis of which amino acids.
Moreover, the amount of protein synthesized within the
mitochondria is negligible when compared with the amount
generated in the cytosol. In yeast, the mitochondrially
synthesized proteins have been estimated to represent only
5±10% of the total mitochondrial protein [47], which itself
represents only a small fraction of the total cellular protein.
Hence, potentially different 13C labeling patterns for mitochondrial and cytosolic amino-acid pools cannot be
detected, and f values refer, to a very good approximation,
to cytosolic amino-acid pools only.
Except for Lys and Gly, the common amino acids are
synthesized in S. cerevisiae from the same precursors
(Fig. 2A) as in E. coli [15]. Specifically, Lys is synthesized
via the a-aminoadipate pathway from OxGlt and cyt-AcCoA,
while Gly is generated through both Ser cleavage via
SHMT [15] and Thr cleavage via threonine aldolase [48], as
well as the mitochondrial GCV (see above). During routine
acidic hydrolysis of biomass, tryptophan and cysteine are
lost due to oxidation. Hence, these amino acids are not
considered in the framework of the present study.
Identification of the subcellular location of intermediates
involved in amino-acid biosynthesis (Fig. 2B) requires
knowledge about the location of the enzymes catalysing
their incorporation into the amino-acid carbon skeleton
[15]. Ser, Gly, Asp, Met, Thr, Tyr, Phe and His are
exclusively derived from metabolite pools located in the
cytosol. Ala, Val, Leu and Ile synthesis requires Prv. For
Ala, the first step is the amination of pyruvate catalysed by
alanine aminotransferase. For Val and Leu, the condensation of two molecules of Prv to 2-acetolactate catalysed by
acetolactate synthase, and for Ile, the condensation of Prv
and 2-oxobutyrate (arising from Thr) to 2-aceto-2-hydroxybutyrate (Ile) catalysed by acetolactate synthase, serve as
the first steps. While the subcellular localization of the
alanine aminotransferase has not yet been identified, acetolactate synthase is located in the mitochondria [49,50].
Moreover, the incorporation of AcCoA into Leu is catalysed
by mitochondrial a-isopropylmalate synthase [49]. Glu, Pro
and Arg are synthesized from OxGlt, which is exclusively
generated in the mitochondria. Lys is synthesized from
OxGlt and mt-AcCoA via the cytosolic homocitrate synthase [51]. (It should be added here that Ala can also be
synthesized from glyoxylate via glycine aminotransferase
[52]. However, this pathway is not active when S. cerevisiae is grown on glucose [52], and was thus not further
considered for the present study.)
Figure 2 summarizes the biochemical data outlined in
this paragraph and shows a mapping of the carbon skeletons
of the proteinogenic amino acids to the intermediates of
central carbon metabolism involved in their biosynthesis.
This mapping thus assigns the 13C fine structure observed
for a given carbon in the amino acids to carbon atoms in
the intermediates. For the sake of simplicity, and when
appropriate, we may refer f values experimentally determined for the amino acids directly to the corresponding
carbon of the metabolic intermediates. Figure 2 provides a
guide to make this reasoning readily transparent to the
reader.
Analysis of 13C scalar coupling fine structures in
S. cerevisiae
In order to cope with cellular compartmentation as
encountered in eukaryotic organism S. cerevisiae (Fig. 1),
the formalism derived to analyse prokaryotic metabolism
[1,5±9] needs to be extended accordingly. An established
but key assumption is that the metabolic network is
operating in a quasi steady-state (see above). Moreover,
fluxes of glycolysis and PenPp, both of which are entirely
located in the cytosol, can be treated as described for
prokaryotic cells [1,6±9,12] (see above).
Anaplerosis of TCA cycle
The contribution of the anaplerotic interconversion of
cytosolic PPrv into mt-OxAc, which is participating in the
TCA cycle, shall be calculated. The synthesis of OxAc
from PPrv via the joint action of pyruvate kinase and
pyruvate carboxylase is restricted to the cytosol in
S. cerevisiae [31,32], and cyt-OxAc thus needs to be
imported into the mitochondria to warrant anaplerosis.
However, only cyt-OxAc, but not mt-OxAc, can be assessed
via Asp (Fig. 2), and the required f values of mt-OxAc must
be inferred from OxGlt. For growth of S. cerevisiae on
glucose, it can be safely assumed that the condensation of
mt-AcCoA, assessed via Leu-a (Fig. 2), and mt-OxAc is
irreversible [15]. Hence, the fraction of intact C2-C3
connectivities in mt-OxAc, which is equal to the fraction of
mt-OxAc stemming from anaplerotic carboxylation [1],
denoted X ana ˆ X(mt-OxAc à PPrv), can be derived from
OxGlt and PPrv and is given by Eqn (1):
X ana ˆ X…mt-OxAc à cyt-PPrv†
ˆ ‰ f …2† 1 f …3† Š{Glu-a}/‰ f …2† 1 f …3† Š{Phe; Tyr-a}
…1†
The f values of Glu-a and Phe/ Tyr-a represent C2 of
OxGlt and PPrv, respectively (Fig. 2).
2468 H. Maaheimo et al. (Eur. J. Biochem. 268)
q FEBS 2001
Fig. 2. Carbon fragments originating from a single intermediate molecule present in proteinogenic amino acids, provided that only anabolic
pathways are active in S. cerevisiae cells [15±18]. (A) Representation of the carbon skeleton of the intermediates of glycolysis, TCA and the
pentose phosphate pathway with circles, squares and triangles, respectively. Acetyl-Coenzyme A (AcCoA) is also represented with circles, and
phosphoryl groups are denoted by `P'. Intermediates in the solid box are generated in the cytosol, and intermediates in the dashed box are
synthesized in the mitochondria. Hence, pyruvate, acetyl-CoA and oxaloacetate are generated in both compartments, and the exchange flux between
the mitochondrial (mt) and cytosolic (cyt) pools need to be considered for proper data interpretation. (B) Representation of the carbon skeletons of
amino acids, as well as the origin of their carbon atoms with respect to the metabolic intermediates displayed in (A) and the compartment in which
the intermediate is recruited for amino-acid biosynthesis (dashed and solid boxes indicate mitochondrial and cytosolic origin, respectively). Ala is
marked with an asterisk (*) because the mitochondrial origin has been inferred from the present study (see text). Thin lines denote carbon bonds that
are formed between fragments arising from different intermediate molecules, while thick lines indicate carbon±carbon connectivities in intact
fragments arising from a single intermediate molecule. Dashed lines connect fragments arising from the same intermediate molecule that are not
directly attached in the amino-acid carbon skeleton. Unlabeled carbon atoms in the amino acids are C 0 carbons. It is indicated that in S. cerevisiae
Gly can be synthesized either via serine or threonine cleavage. In (B), those amino acids that are lost due to oxidation (Cys, Trp) or that are
deamidated (Asn, Gln) during routine hydrolysis of cellular protein are not considered. Note that Ser and Gly are affected by C1 metabolism so that
their 13C fine structures usually do not reflect the isotopomeric composition of GriP. The superscript `x' indicates that the two carbons d1 and d2, and
:1 and :2, respectively, of Tyr and Phe give rise to only one 13C fine structure each [1]. The nomenclature of the carbon atoms follows IUPAC-IUB
recommendations [88].
q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2469
Exchange flux between OxAc and fumarate
Intact C1-C2-C3 fragments originating from a single source
molecule of glucose in OxGlt must arise from mt-OxAc
that was converted to mt-fumarate or mt-succinate and
subsequently reacted back to mt-OxAc [1]. Hence, the
fraction of mt-OxAc molecules that were at least once
reversibly interconverted to mt-fumarate, denoted mt-X exch,
is given by Eqn (2):
arise from free rotation in solution, is not considered.
Assuming full symmetrization, i.e. neglecting any metabolite channeling, the contributions multiplied by the
(1 2 X ana) term in Eqns (4) are averaged between C2
and C3 of OxAc, yielding Eqns (5):
f
…i†
{mt-OxAc-C2} ˆ X ana ´‰ f
f …2 *† {Asp-a}; f
mt-X exch ˆ X…mt-OxAc $ mt-fumarate†
…3†
ˆ ‰2 f
…2†
Š{Glu-a}/‰ f
…3†
1f
cyt-X
f
…i†
…3†
Š{Asp-b}/‰ f
…2†
1f
…3†
f …2 *† {Asp-b}; f
Š{Asp-b}
…3†
´‰ f
exch
In the case that (a) cyt-X
is nonzero and (b) the transfer
from OxAc into the mitochondria is unidirectional, Eqn (2)
can readily be extended to:
mt-X exch ˆ ‰2 f
…3†
Š{Glu-a}/‰ f
…2†
1f
…3†
Š{Glu-a}
2 cyt-X exch
…2b†
exch
Calculation of X
according to Eqns (2) and (3) assumes
complete symmetrization of 13C labeling patterns about the
C2-C3 bond of succinate or fumarate [1] arising from
unrestricted rotation in solution. In cases where metabolite
channeling [53±57], which may restrict rotational reorientation, had to be invoked for the exchange reaction, the
value for X exch would represent a lower bound (see below).
f
…i†
{mt-OxAc-C2} ˆ X ana ´‰ f
f …2 *† {Asp-a}; f
´‰ f
f
…i†
…1†
…3†
…2†
{Glu-b}; 0Š;
{mt-OxAc-C3} ˆ X ana ´‰ f
´‰ f
…1†
…3†
…2†
{Asp-a};
{Asp-a}Š 1 …1 2 X ana †
{Glu-b}; 0; f
f …2 *† {Asp-b}; f
{Asp-a}; f
…1†
{Asp-b}; f
…4a†
…2†
f
…i†
where i ˆ 1, 2, 2*, or 3 in these and the following equations.
Eqns (4) were derived in the limit of complete metabolite
channeling [53±57], i.e. symmetrization of 13C labeling
patterns about the C2-C3 bond of succinate or fumarate that
…5a†
…1†
…3†
{Glu-b} 1 f
…1†
{Asp-b}; f
…2†
{Asp-b};
{Asp-b}Š 1 …1 2 X ana †´0:5
…1†
{Glu-g}; 0; f
…2†
{Glu-b}
…5b†
{mt-OxAc-C2} ˆ X ana ´…1 2 0:5´mt-X exch †
´‰ f
…1†
{Asp-a}; f
…2†
{Asp-a}; f …2 *† {Asp-a};
f
…3†
{Asp-a}Š 1 X ana ´0:5´mt-X exch ´‰ f
f
…2†
{Asp-b}; f …2 *† {Asp-b}; f
1 …1 2 X ana †´0:5´‰ f
f
…i†
…2†
…1†
…3†
…1†
{Asp-b};
{Asp-b}Š
{Glu-b} 1 f
…1†
{Glu-g}; 0;
{Glu-b} 1 f …2 *† {Glu-g}; 0Š;
…6a†
{mt-OxAc-C3} ˆ X ana ´…1 2 0:5´mt-X exch †
´‰ f
…1†
{Asp-b}; f
…2†
{Asp-b}; f …2 *† {Asp-b};
f
…3†
{Asp-b}Š 1 X ana ´0:5´mt-X exch ´‰ f
f
…2†
{Asp-a}; f …2 *† {Asp-a}; f
1 …1 2 X ana †´0:5´‰ f
f
…2†
…1†
…3†
…1†
{Asp-a};
{Asp-a}Š
{Glu-b} 1 f
…1†
{Glu-g}; 0;
{Glu-b} 1 f …2 *† {Glu-g}; 0Š:
…6b†
Provided that Ala is synthesized from mt-Prv, Eqns (7) and
(8) yield the aforementioned upper, X ub, and lower bounds,
X lb, assuming that malate is entirely derived from either
mt-OxAc or OxGlt, respectively. With f (2*){mt-OxAc-C2}
from Eqn (6a) this yields:
X ub -I …mt-Prv à mt-malate† ˆ ‰ f …2 *† {Ala-a}
{Asp-b};
…4b†
{Glu-b}
Finally, the symmetrization of C labeling patterns arising
from reversible interconversion from OxAc to fumarate,
characterized by mt-X exch, is considered, yielding Eqns (6):
2 f …2 *† {Phe-a}Š/‰ f …2 *† {mt-OxAc-C2}
{Asp-b}Š 1 …1 2 X ana †
{Glu-g}; 0; f …2 *† {Glu-g}; 0Š;
…2†
13
f
…1†
{Asp-a};
1 f …2 *† {Glu-g}; 0Š:
Flux via the malic enzyme
The interconversion of mt-malate to mt-Prv via the malic
enzyme is manifested by the flux of excess intact C1-C2
fragments (represented by f (2*)) and C3-C2 fragments
(represented by f (2)) into the mt-Prv pool when compared
with the PPrv pool [6]. Upper and lower bounds for the
fraction of mt-Prv from mt-malate can be derived under the
assumptions that malate is entirely derived from either
mt-OxAc or OxGlt, respectively [6]. The calculation of the
upper bound requires the f values for mt-OxAc (see below).
If X ana and X exch are known from Eqns (1) and (2), the f
values can be calculated from Asp and Glu, representing
cyt-OxAc and OxGlt, respectively:
{Glu-g}; 0; f
{mt-OxAc-C3} ˆ X ana ´‰ f
X…cyt-OxAc $ cyt-fumarate†
ˆ ‰2 f
…2†
{Asp-a}Š 1 …1 2 X ana †´0:5
…1†
{Glu-b} 1 f
{Asp-a}; f
1 f …2 *† {Glu-g}; 0Š;
Š{Glu-a} …2†
The same fraction can be calculated for the cytosolic pool
[1] using Eqn (3):
exch
…1†
´‰ f
…3†
…1†
2 f …2 *† {Phe-a}Š;
…7†
and
X lb -I …mt-Prv à mt-malate† ˆ ‰ f …2 *† {Ala-a}
2 f …2 *† {Phe-a}Š/‰1 2 f …2 *† {Phe-a}Š:
…8†
2470 H. Maaheimo et al. (Eur. J. Biochem. 268)
q FEBS 2001
Fig. 3, X mt
1 corresponds to X(mt-Prv à mt-malate) introduced in Eqns (7±9) to estimate the involvement of the
malic enzyme to synthesize mt-Prv.
The cytosolic PPrv pool is experimentally assessible via
Phe or Tyr (Fig. 2), i.e. f (i) {PPrv-C2} ˆ f (i) {Phe-a,
Tyr-a}. Assuming that the mt-Prv pool is observed at
Ala, i.e. that f (i) {mt-Prv-C2} ˆ f (i) {Ala-a}, one obtains
cyt
cyt
out
with X cyt
1 ˆ n1 /[nPrv 1 n1 ] Eqn (11):
f
…i†
{cyt-Prv-C2} ˆ X cyt
1 ´f
1 …1 2 X cyt
1 †´f
from which the flux ratio
Fig. 3. Sub-network for analysis of fluxes at the interface between
glycolysis and TCA cycle considering also the compartmentation in
S. cerevisiae cells (Fig. 1). Metabolic fluxes, n , and flux ratios, X, are
labeled with `mt' for mitochondrial and with `cyt' for cytosolic
depending on the compartment in which the biochemical reactions
takes place. The flux ratios which involve the exchange fluxes (see
text) are depicted in grey. Abbreviations for metabolites are as given in
the legend of Fig. 1. Single-headed arrows represent reactions that are
assumed to be unidirectional under the present growth conditions (see
legend of Fig. 1). Metabolites which are assessible by BDF 13C
labeling of proteinogenic amino acids are encircled.
Considering that f (2){mt-malate-C2} # f (2){mt-OxAcC2}, a second lower bound can be derived with f (2){mtOxAc-C2} from Eqn (6a) when introduced into balancing
Eqn (9):
lb
X -II …mt-Prv à mt-malate† ˆ ‰ f
2f
…2†
{Phe-a}Š/‰ f
…2†
…2†
{Ala-a}
{mt-OxAc-C2} 2 f
{Phe-a}Š:
…9†
Intercompartmental exchange fluxes
Balancing equations for relative abundances of intact
carbon fragments (i.e. f values), which characterize the
cytosolic and mitochondrial pools of Prv, AcCoA and
OxAc, can provide estimations of metabolic flux ratios
involving intercompartmental fluxes. Figure 3 displays the
selected subnetwork as well as metabolic flux definitions
chosen to derive the desired equations. Reactions that were
considered to be irreversible are indicated by unidirectional
arrows (see legend of Fig. 1).
1. Prv. Assuming that the mt-Prv pool is experimentally
assessible via Ala (Fig. 2), i.e.
f
…i†
one obtains with
f
…i†
{mt-Prv-C2} ˆ f
X mt
1
ˆ
in
nmt
1 /[nPrv
{mt-Prv-C2} ˆ X mt
1 ´f
1 …1 2 X mt
1 †´f
…i†
…i†
{Ala-a};
1 nmt
1 ] Eqn (10):
…i†
{mt-malate-C2}
{cyt-Prv-C2};
f
…1†
{mt-AcCoA-C1} ˆ X mt
2 ´‰ f
1 …1 2 X mt
2 †´f
f
…2†
and estimations of f (i) {mt-malate-C2} and f (i) {cyt-PrvC2} yield the flux ratio X mt
1 . For the bioreaction network of
…1†
{mt-Prv-C2};
…11†
can be calculated.
1 …1 2 X mt
2 †´f
…2†
…1†
1 f …2 *† Š{mt-Prv-C2}
{cyt-AcCoA-C1};
{mt-AcCoA-C1} ˆ X mt
2 ´‰ f
…2†
1f
…3†
…12a†
Š{mt-Prv-C2}
{cyt-AcCoA-C1};
…12b†
X mt:
2
yielding the flux ratio
Analogous equations are valid for the cyt-AcCoA pool,
cyt
cyt
out
and with X cyt
2 ˆ n2 /[nAcCoA 1 n2 ] Eqn (13) is obtained:
f
…2†
{cyt-AcCoA-C1} ˆ X cyt
2 ´‰ f
…2†
…2†
1f
…3†
Š{cyt-Prv-C2}
{mt-AcCoA-C1}:
…13†
The f values for cyt-Prv-C2 need to be derived from PPrv
(i.e. Phe-a, Tyr-a) or cyt-OxAc (i.e. Asp-a).
3. OxAc. The mt-OxAc pool has been considered for
assessing the anaplerosis of the TCA cycle: a flux ratio
invoking nin
OxAc can be derived in a straightforward fashion
when considering that anaplerosis is solely due to the
transport of cyt-OxAc into the mitochondria (see above).
in
mt
in
X mt
3 ˆ nOxAc /‰n3;net 1 nOxAc Š;
…14a†
where nmt
3;net represents the net flux providing mt-OxAc
from OxGlt through operation of the TCA cycle. For
unidirectional interconversion of PPrv into cyt-OxAc
(Fig. 1), one obtains with X ana from Eqn (1) and the
bioreaction network of Fig. 3 that
ana
X mt
:
3 ˆX
…14b†
The cyt-OxAc pool is experimentally assessible via
Asp (Fig. 2), i.e. f (i){cyt-OxAc-C2} ˆ f (i) {Asp-a}.
cyt
cyt
out
With X cyt
3 ˆ n3 /[nOxAc 1 n3 ] one arrives at the balancing
Eqns (15):
f
…10†
X cyt
1
{PPrv-C2}
2. AcCoA. The mt-AcCoA pool is experimentally assessible via Leu-a (Fig. 2), i.e. f (2) {mt-AcCoA-C2} ˆ
f (2) {mt-AcCoA-C1} ˆ f (2*) {Leu-a} (Fig. 2), and the
cytosolic values f (i) {cyt-AcCoA-C2} are obtained correspondingly from analysis of Lys-a. One then obtains with
mt
in
mt
X mt
2 ˆ n2 /[nAcCoA 1 n2 ] Eqns (12):
1 …1 2 X cyt
2 †´f
…2†
…i†
…i†
…i†
{cyt-OxAc-C2} ˆ X cyt
3 ´f
1 …1 2 X cyt
3 †´f
…i†
…i†
{cyt-Prv-C2}
{mt-OxAc-C2}:
…15†
Hence, estimations of f (i) {cyt-Prv-C2} (see above) and the
f (i) {mt-OxAc-C2} values yield the flux ratio X cyt
3 .
q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2471
During hydrolysis of the biomass, Cys and Trp were
oxidized and could thus not be evaluated, Asn and Gln were
deamidated to Asp and Glu, and the ring carbons of
phenylalanine were not considered because of strong
13
C± 13C scalar coupling effects [1]. A survey of cross
sections taken along v1(13C) from the 2D spectra is
afforded by Fig. 4.
R E S U LT S A N D D I S C U S S I O N
Biosynthesis of proteinogenic amino acids in S. cerevisiae
The 2D 13C-1H COSY spectra were analysed as described
previously [1,14] yielding the desired f values (Table 1).
Table 1. Relative abundances of intact C2 and C3 fragments in proteinogenic amino acids. The first column indicates the carbon position for
which the 13C fine structure was observed. The f values were calculated as described [1], and are given for the aerobic and the anaerobic preparation
in columns 2±5 and 6±9, respectively. Note, firstly, that for terminal carbons f (2*) and f (3) are not defined, and, secondly, that for Tyr the two
carbons d1 and d2, and :1 and :2, respectively, give rise to only one 13C fine structure each [1].
Relative abundance of intact carbon fragments
Aerobic
Carbon
atom
f
Ala-Ca
Ala-Cb
Arg-Cb
Arg-Cd
Asp-Ca
Asp-Cb
Glu-Ca
Glu-Cb
Glu-Cg
Gly-Ca
His-Ca
His-Cb
His-Cd2
Ile-Ca
Ile-Cg1
Ile-Cg2
Ile-Cd1
Leu-Ca
Leu-Cb
Leu-Cd1
Leu-Cd2
Lys-Ca
Lys-Cb
Lys-Cg
Lys-Cd
Lys-C:
Met-Ca
Phe-Ca
Phe-Cb
Pro-Ca
Pro-Cb
Pro-Cg
Pro-Cd
Ser-Ca
Ser-Cb
Thr-Ca
Thr-Cb
Thr-Cg2
Val-Ca
Val-Cg1
Val-Cg2
Tyr-Ca
Tyr-Cb
Tyr-Cdx
Tyr-C:x
0Š.03
0Š.09
0Š.26
0Š.05
0Š.02
0Š.03
0Š.05
0Š.25
0Š.03
0Š.07
0Š.02
0Š.06
0Š.11
0Š.05
0Š.98
0Š.12
1
0Š.03
0Š.99
0Š.18
1
0Š.04
0Š.26
0Š.26
0Š.04
0Š.05
0Š.03
0Š.02
0Š.02
0Š.07
0Š.26
0Š.02
0Š.04
0Š.01
0Š.21
0Š.02
0Š.01
1
0Š.09
0Š.10
0Š.93
0Š.03
0Š.03
0Š.04
0Š.31
(1)
Anaerobic
(2)
f (2*)
f
0Š.05
0Š.91
0Š.00
0Š.95
0Š.02
0Š.96
0Š.61
0Š.75
0Š.00
0Š.93
0Š.00
0Š.62
0Š.89
0Š.00
0Š.00
0Š.88
0Š.00
0Š.00
0Š.00
0Š.82
0Š.00
0Š.00
0Š.74
0Š.74
0Š.95
0Š.95
0Š.05
0Š.01
0Š.98
0Š.63
0Š.74
0Š.00
0Š.96
0Š.01
0Š.79
0Š.02
0Š.99
0Š.00
0Š.01
0Š.90
0Š.07
0Š.01
0Š.97
0Š.96
0Š.02
0Š.05
±Š
0Š.70
±
0Š.02
0Š.00
0Š.21
0Š.00
0Š.97
±
0Š.00
0Š.00
±
0Š.94
0Š.02
±
±
0Š.96
0Š.00
±
±
0Š.96
0Š.00
0Š.00
0Š.00
±
0Š.05
0Š.00
0Š.00
0Š.22
0Š.00
0Š.93
±
0Š.19
±
0Š.03
0Š.00
±
0Š.89
±
±
0Š.00
0Š.00
±
0Š.25
0Š.87
±
0Š.04
±
0Š.94
0Š.00
0Š.13
0Š.00
0Š.00
±
0Š.98
0Š.32
±
0Š.01
0Š.00
±
±
0Š.01
0Š.01
±
±
0Š.00
0Š.00
0Š.00
0Š.01
±
0Š.87
0Š.97
0Š.00
0Š.08
0Š.00
0Š.05
±
0Š.79
±
0Š.93
0Š.00
±
0Š.01
±
±
0Š.96
0Š.00
0Š.00
0Š.42
f
(3)
f
(1)
0Š.03
0Š.04
0Š.12
0Š.07
0Š.03
0Š.04
0Š.03
0Š.04
0Š.03
0Š.23
0Š.03
0Š.07
0Š.11
0Š.08
0Š.98
0Š.07
1Š.00
0Š.04
1Š.00
0Š.12
1Š.00
0Š.03
0Š.04
0Š.07
0Š.04
0Š.06
0Š.03
0Š.04
0Š.03
0Š.06
0Š.05
0Š.03
0Š.05
0Š.04
0Š.22
0Š.03
0Š.04
1Š.00
0Š.14
0Š.05
0Š.93
0Š.04
0Š.04
0Š.06
0Š.32
(2)
f (2*)
f
0Š.08
0Š.96
0Š.00
0Š.93
0Š.04
0Š.95
0Š.86
0Š.53
0Š.00
0Š.77
0Š.00
0Š.67
0Š.89
0Š.00
0Š.00
0Š.93
0Š.00
0Š.00
0Š.00
0Š.88
0Š.00
0Š.00
0Š.96
0Š.93
0Š.95
0Š.94
0Š.06
0Š.00
0Š.97
0Š.85
0Š.91
0Š.00
0Š.95
0Š.01
0Š.78
0Š.04
0Š.96
0Š.00
0Š.00
0Š.95
0Š.07
0Š.01
0Š.96
0Š.94
0Š.01
0Š.00
±
0Š.86
±
0Š.01
0Š.00
0Š.03
0Š.43
0Š.97
±
0Š.00
0Š.00
±
0Š.92
0Š.02
±
±
0Š.96
0Š.00
±
±
0Š.97
0Š.00
0Š.00
0Š.00
±
0Š.05
0Š.00
0Š.00
0Š.03
0Š.00
0Š.92
±
0Š.17
±
0Š.02
0Š.00
±
0Š.85
±
±
0Š.00
0Š.00
±
0Š.26
0Š.89
±
0Š.02
±
0Š.92
0Š.01
0Š.08
0Š.00
0Š.00
±
0Š.97
0Š.26
±
0Š.00
0Š.00
±
±
0Š.00
0Š.00
±
±
0Š.00
0Š.00
0Š.00
0Š.01
±
0Š.86
0Š.96
0Š.00
0Š.06
0Š.04
0Š.05
±
0Š.78
±
0Š.91
0Š.00
±
0Š.01
±
±
0Š.95
0Š.00
0Š.00
0Š.41
f
(3)
2472 H. Maaheimo et al. (Eur. J. Biochem. 268)
Analysis of the f values [1,7] obtained in both the aerobic
and anaerobic regimes (Table 1) provided strong evidence
for the hypothesis that the proteinogenic amino acids in
S. cerevisiae are synthesized according to known pathways
[15±18] (Fig. 2). Except for a yet unexplained slight
deviation of the f values of Leu-d1, which was previously
also observed for prokaryotes [1], inconsistencies could not
be detected: f values predicted [1,7] from (a) the bioreaction
network (Fig. 1), (b) the location of intermediates serving
for amino acid biosynthesis (Fig. 2), and (c) the known
biosynthetic routes [15±18] (Fig. 2) were in excellent
agreement with those determined experimentally. As (a)
f (i){Lys-d} ˆ f (i){Glu-g} ˆ f (i){mt-OxAc-C4},
(b)
f (i){Lys-g} ˆ f (i){Glu-b} ˆ f (i){mt-OxAc-C3} and (c)
[ f (2) 1 f (3)] {Lys-a} ˆ 0 (Fig. 2; Table 1), Lys synthesis
must occur from homocitrate and AcCoA, i.e. via the aaminoadipate pathway.
When grown on glucose, S. cerevisiae cells are using
both Thr and Ser cleavage to generate Gly [44,48].
However, as Ser is derived from GriP, which can be
assumed to exhibit virtually the same f values as cyt-Prv [1]
(from which Thr is derived via carboxylation to cyt-OxAc),
we have that [ f (2*) 1 f (3)] {Ser-a} ˆ [ f (2*) 1 f (3)]
{Thr-a} (Table 1). Cleavage of the Ca-Cb bond of either
Ser or Thr would thus lead to the same f (2) {Gly-a} values,
i.e. the relative contribution of the two cleavage pathways
cannot be determined using the presently employed 13C
labeling protocol. In contrast, we find that f (1){Serb} 2 f (1){Phe-b} ˆ 0.19 in both the anaerobic and the
aerobic regime. This provides evidence for reversible
cleavage of 19% of the Ser molecules into Gly and a C1
unit [1], while f (1){Thr-b} < 0 shows that Thr cleavage is,
if present, not reversible. Hence, we are left with evidence
for the operation of SHMT, but we cannot judge on the
relative importance of Thr cleavage for Gly biosynthesis.
Considering that Thr cleavage has, in principle, been shown
to be of importance when S. cerevisiae grows on glucose
[44,48], we tentatively conclude that this reaction is
irreversible under the currently growth conditions. Consistently, DG8 0 < 10 kJ´mol21 (http://wwwbmcd.nist.gov:
8080/enzyme/ename.html) has been measured for the Thr
cleavage, while DG8 0 < 20 kJ´mol21 was obtained for the
Ser cleavage reaction. This indicates that the Ser cleavage
reaction tends to be more on the side of the intact amino
acid than the Thr cleavage.
Gly can also be synthesized from a C1 unit and CO2 via
mitochondrial GCV. This reaction generates increased
f (1) {Gly-a} values [1], provided that mt-Gly is transported
into the cytosol where the bulk part of the cellular protein is
generated. As f (1){Gly-a} 2 f (1){Phe-a} ˆ 0.05 and 0.19
for the aerobic and anaerobic preparation, respectively
(Table 2; Fig. 2), we have evidence for (a) the reversible
action of mitochondrial GCV in both the aerobic and the
anaerobic regime, and for (b) a significant flux of mt-Gly
into the cytosolic pool (we also find that f (1) {Ser-a} ˆ
f (1) {Gly-a} [ f (1){Ser-b} 2 f (1){Phe-b}], which is consistent with the view that cytosolic Ser and Gly are used for
the generation of the bulk part of the cellular protein). We
cannot determine the fraction of cyt-Gly arising from mtGly because the 13C labeling patterns of the latter are not
available. However, the value of f (1){Gly-a} 2 f (1){Phe-a}
constitutes a lower bound for cyt-Gly arising from mt-Gly,
which would be reached only if all mt-Gly molecules were
q FEBS 2001
reversibly cleaved by the GCV. Considering that GCV
probably cleaves only a minor fraction of the mitochondrial
Gly pool, lower bounds of 5 and 19% in the aerobic and
anaerobic regime, respectively, indicate a significant flux of
Gly from the mitochondria into the cytosol (Table 2). This
finding is consistent with the earlier observation that
mammalian cells with active cytosolic SHMT but inactive
mitochondrial SHMT are glycine auxotrophs [58], i.e. the
cytosolic Gly production is not sufficient to meet cellular
demands.
Novel insight from the 13C labeling experiment was also
obtained with respect to the location of the pyruvate pool
serving for Ala biosynthesis: the f values of Ala-a (Fig. 2)
reflect the action of the malic enzyme (see below; Table 1)
which itself has been shown to be exclusively localized in
the mitochondria [34]. Our data thus show that alanine
aminotransferase is located in the mitochondria.
Overall, the biosynthetically directed fractional 13C
labeling experiment yielded the assignment of amino-acid
13
C-fine structures to carbon atoms in intermediates as well
as their subcellular locations as shown in Fig. 2. This
allows an investigation of the central carbon metabolism of
S. cerevisiae, and in particular the determination of
metabolic flux ratios.
Cytosol: pentose phosphate pathway and glycolysis
Intermediates of the pentose phosphate pathway (PenPp)
and glycolysis are entirely located in the cytosol, and the
biosynthetic origin of these intermediates (Table 2) was
determined as previously described for prokaryotic cells
[1,6±9,11]. Firstly, the data show that the flux ratios of the
PenPp are astonishingly similar when comparing the
aerobic (glucose repressed), and the anaerobic growth
regime. Secondly, the employment of PenPp for generation
of PPrv is beyond detection (, 4%), i.e. PPrv stems from
glycolysis only, suggesting an exclusively anabolic role of
the PenPp when S. cerevisiae cells are grown under the
current conditions. In fact, this is consistent with a previous
study [59] estimating that S. cerevisiae catabolizes less
than 1% of the glucose via PenPp to PPrv. Notably, the
inability of glycolytic enzyme knock-out mutants to employ
the PenPp for glucose degradation is apparently not due to
an insufficient capacity of the pathway ensuring appropriate
metabolic fluxes for glucose catabolism [60], but rather is a
result of the lack of a transhydrogenase in S. cerevisiae [61]
for equilibration of the NADH and NADPH pools. Thirdly,
PenPp is fed via both the oxidative and nonoxidative
branches. The activity of the oxidative branch [1,6,8] could
be directly assessed from registering intact C5 fragments in
Ri5P (Fig. 2,4) which yields lower bounds around 30% for
pentose synthesis by glucose oxidation (Table 2). The
operation of the nonoxidative branch is implied by a
significant flux from fructose 6-phosphate to erythrose 4phosphate (< 15%) catalysed by transketolase and
exchange fluxes catalysed by transaldolase and transketolase combined with the notion that PPrv generation through
PenPp is not detected.
Cytosol: glyoxylate cycle and pyruvate carboxylation/
decarboxylation
The exchange fluxes between cyt-OxAc and cyt-fumarate,
cyt-X exch, calculated with Eqn (3), are zero in both the
q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2473
Fig. 4. 13C scalar coupling fine structures observed for BDF 13C labeled amino acids obtained during growth of the S. cerevisiae cultures on a
minimal medium containing glucose as the sole carbon source. In the left-most column the carbon position is listed for which the 13C fine
structure was observed. The right-most column indicates the carbon positions for which closely similar 13C fine structures were observed (for a
quantitative comparison see Table 1). The columns two and three display the v1(13C) cross sections as observed in the 2D 13C-1H COSY spectrum
recorded for the aerobic and anaerobic preparation, respectively. The carbon atoms are grouped according to the metabolic intermediates from
which they are derived (Fig. 2). The section on the left comprises intermediates generated in the cytosol, while the section on the right contains
those synthesized in the mitochondria. The doublet of doublets arising from histidine 13Cg-13Cb-13Ca is further split by the two-bond coupling
constant 2JCbCd, indicating the presence of intact C5 fragments in histidine. Note that due to the influence of 13C isotope effects on the 13C chemical
shifts the doublet, as well as the doublet of doublets is shifted by < 1±2 Hz to higher field relative to the singlet line [89]. Hence, the superposition
of the multiplets arising from different isotopomers is not symmetrical with respect to the singlet line.
2474 H. Maaheimo et al. (Eur. J. Biochem. 268)
q FEBS 2001
Table 2. Origin of intermediates in S. cerevisiae central carbon metabolism inferred from analysis of BDF 13C labeled amino acids. The
fraction of the total pool of the metabolite indicated on the left is given. Values obtained for the cytosol, the mitochondria and C1 metabolism are
listed separately. The abbreviations of the metabolites are given in the legend of Fig. 1, and the Eqns employed to calculate the fractions are given in
the corresponding footnotes. Entries containing NA could not be obtained for the present system due to degeneracy of f values (see text; Table 1).
Experimental errors have been estimated from analysis of redundant 13C scalar coupling fine structures [1] and the signal-to-noise ratio of the cross
peaks in the 2D 13C-1H COSY spectra (Fig. 4). Ranges of fractions were derived either from bounds, or from multiple determinations employing the
balancing Eqns (11), (14) and (15) for different f (i) values (or linear combinations thereof ). For the definition of the flux ratios, X, see Fig. 3.
cyt
cyt
According to these definitions, the fractions …1 2 X cyt
1 † …1 2 X 2 † and …1 2 X 3 †, which can readily be derived from the corresponding X values
given in this table, represent `cyt-Prv from mt-Prv', `cyt-AcCoA from mt-AcCoA' and `cyt-OxAc from mt-OxAc', respectively. (1 2 X mt
1 )
mt
(1 2 X mt
2 ) and (1 2 X 3 ) represent `mt-Prv from cyt-Prv', `mt-AcCoA from cyt-AcCoA' and `mt-OxAc from mt-OxGlt', respectively.
Fraction of total pool (%)
Metabolites
Aerobic
Cytosol
P-Prv from pentoses (upper bound)
Ri5P from glucose (lower bound)
Ri5P from G3P and S7P (transketolase reaction)
Ri5P from E4P (transketolase and transaldolase reactions)
E4P from fructose (lower bound)
cyt-Prv from a P-Prv (X cyt
1 )
cyt-AcCoA from b cyt-Prv (X cyt
2 )
cyt-OxAc from c cyt-Prv (X cyt
3 )
cyt-OxAc reversibly converted d to fumarate (cyt-Xexch)
0±4
32 Š^ 2
68 ^
Š 2
10 ^
Š 2
15 Š^ 4
70±100
NA
88±100
0±4
Mitochondria
mt-Prv from malate (X1mt)
(upper e bound I, Xub-I)
(lower f bound I, Xlb-I)
(lower g bound II, Xlb-II)
25±30
< 25
<5
< 30
mt-AcCoA from h mt-Prv (X2mt)
mt-OxAc from k PPrv (Xana/X3mt)
mt-OxAc reversibly l converted to fumarate (mt-Xexch)
C1 metabolism
Ser from Gly and C1 unit
Gly from CO2 and C1 unit
Cyt-Gly from mt-Gly
a
Eqn (11);
b
Eqn (13); c Eqn (15);
d
Eqn (3); e Eqn (7); f Eqn (8);
g
aerobic and anaerobic regime (Table 2). This exchange
reaction is catalyzed by enzymes constituting the glyoxylate cycle. Hence, our data strongly support the previously
published finding that the glyoxylate pathway enzymes
[40±42], including the cytosolic malate dehydrogenase
[43], are not expressed when S. cerevisiae cells grow under
glucose repression.
f (3) {Asp-a} . 0.9 and f (2*) {Lys-a} . 0.9 show that
the majority of the cyt-OxAc and cyt-AcCoA pools arise,
respectively, from carboxylation and decarboxylation of
cyt-Prv in both the aerobic and the anaerobic regime.
Mitochondria: TCA cycle and malic enzyme activity
The relative supply of the TCA cycle with either AcCoA for
generation of reduction equivalents or OxAc to fulfill
biosynthetic demands is a pivotal parameter characterizing
the regulatory state of central metabolism. Employing
Eqn (1), we obtained X ana ˆ 0.76 and 1.00 for the aerobic
and anaerobic regime, respectively (Table 2). In the
anaerobic regime, the reactions constituting the TCA
cycle are evidently operating in a branched fashion as it
Eqn (9);
h
Eqn (12);
Anaerobic
0±4
25 Š^ 2
75 Š^ 2
11 Š^ 2
17 Š^ 4
50±100
NA
71±100
0±4
70±100
NA
NA
70±100
NA
76 Š^ 4
35 Š^ 4
NA
92±100
17 Š^ 4
19 Š^ 2
4 Š^ 3
.4
19 Š^ 2
19 Š^ 3
. 19
k
Eqns (1) and (14); and l Eqn (2b)
is typically observed for prokaryotic cells [1]. Hence, the
flux from OxGlt to mt-OxAc is zero. Consistently, OxGlt
dehydrogenase activity cannot be detected under anaerobic
conditions [62,63]. The aerobic value of X ana ˆ 0.76 is
quite high when compared with that of prokaryotic cells
[1,6,8,9,11,12,14], and reflects that S. cerevisiae cells
grown under glucose repression generate a large fraction
of the cellular ATP through fermentative metabolism. This
implies a diminished role of ATP synthesis by respiration,
so that the TCA cycle functions primarily for biosynthesis.
Consistently, Polakis & Bratley [64] found reduced TCA
enzyme activities for S. cerevisiae cells grown in the
presence of 0.9% glucose, and Wales et al. [62] obtained
similar results for Saccharomyces carlsbergensis.
A second parameter that can be readily obtained from
BDF 13C labeling of amino acids is the exchange flux
between mt-OxAc and mt-fumarate. As cyt-X exch ˆ 0 (see
above), we obtained mt-X exch ˆ 0.35 and 0.17 for the
aerobic and anaerobic regime, respectively, using Eqn (2).
In order to assess the activity of the malic enzyme, the f
values of mt-OxAc were calculated with Eqns (4±6)
(Table 3). Due to the prevalence of the anaplerotic
q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2475
Table 3. f values calculated for mitochondrial oxaloacetate (mt-OxAc). The values are calculated from f values observed for Glu (Table 1). X ana
and mt-X exch were calculated using Eqns (1) and (2b), respectively. Calculations for Eqns (4), (5) and (6) were based on conditions (a), (b) and (c),
respectively. (a) Assuming complete metabolite channeling during synthesis of mt-OxAc from OxGlt and neglecting reversible exchange from mtOxAc to mt-fumarate, (b) neglecting both metabolite channeling and the reversible exchange, and (c) neglecting metabolite channeling and
considering the reversible exchange.
Aerobic Xana ˆ 0.76a mt-Xexch ˆ 0.35
(2)
f (2*)
f
0Š.08
0Š.04
0Š.02
0Š.73
0Š.19
0Š.23
0Š.71
0
Eqns (5)
C2
C3
0Š.05
0Š.06
0Š.02
0Š.73
0Š.22
0Š.21
Eqns (6)
C2
C3
0Š.05
0Š.06
0Š.14
0Š.60
0Š.22
0Š.21
Carbon
f
Eqns (4)
C2
C3
(1)
Anaerobic Xana ˆ 1.00 mt-Xexch ˆ 0.17
f
(3)
(2)
f (2*)
f
0Š.03
0Š.04
0Š.04
0Š.96
0Š.01
0
0Š.92
0
0Š.71
0
0Š.03
0Š.04
0Š.04
0Š.96
0Š.01
0
0Š.92
0
0Š.59
0Š.13
0Š.03
0Š.04
0Š.12
0Š.87
0Š.01
0
0Š.84
0Š.09
generation of mt-OxAc, potential channeling of metabolites
during interconversion of OxGlt to mt-OxAc [53±57]
would have little impact (compare f values in Eqns 4±6
in Table 3), i.e. the following results are largely independent of the actual degree of metabolite channeling in the
mitochondria. Employing Eqns (7), (8) and (9), we obtain
X ub-I(mt-Prv à mt-malate) ˆ 0.25, X lb-I(mt-Prv à mtmalate) ˆ 0.05 and X lb-II(mt-Prv à mt-malate) ˆ 0.30
for the aerobic regime. As the first upper and the second
lower bound are nearly equally large, i.e. X ub-I(mtPrv à mt-malate) < X lb-II(mt-Prv à mt-malate), we
conclude that 25±30% of the mt-Prv pool is synthesized
via the malic enzyme, i.e. X(mt-Prv à mt-malate) < 0.25±
0.30. In turn, this indicates that the f values of mt-malate
and mt-OxAc are very similar, if not identical. Consistently,
DG8 0 < 230 kJ´mol21 (http://wwwbmcd.nist.gov:8080/
enzyme/ename.html) has been measured for the interconversion of mt-OxAc into mt-malate, i.e. the production
of malate is strongly favoured in this equilibrium. Under
the anaerobic regime, Eqns (7) and (8) do not apply as
f (2*) {mt-OxAc-C2} ˆ 0. However, Eqn (9) yields
X lb-II(mt-Prv à mt-malate) ˆ 0.70. This rather high
value is consistent with the observation that malic enzyme
gene expression under anaerobic growth is about threefold
increased at the transcriptional and about fourfold increased
translational level when compared with the aerobic regime
[34]. Obviously, the malic enzyme plays a pivotal role for
mt-Prv metabolism in both the aerobic and anaerobic
regime.
Intercompartmental fluxes: pyruvate, acetyl-CoA and
oxaloacetate
Fig. 3 shows the metabolic subnetwork that was selected to
analyse the exchange fluxes of Prv, AcCoA and OxAc
connecting cytosolic and mitochondrial pools (Table 2).
For this network we have that X mt
1 of Eqn (10) is equal to
X(mt-Prv à mt-malate) < 25±30% in the aerobic and
X(mt-Prv à mt-malate) . 70% in the anaerobic regime
(see above).
f
(1)
f
(3)
X cyt
1 , i.e. the fraction of cyt-Prv arising from PPrv, can be
estimated using Eqn (11). Assuming that f (3) {cyt-PrvC2} ˆ f (3){cyt-OxAc-C2} one obtains X cyt
1 . < 70% for
the aerobic and X cyt
.
<
50%
for
the
anaerobic
regime. As
1
f (3){cyt-Prv-C2} might be somewhat reduced by the
combination of the flux from mt-OxAc to cyt-OxAc
followed by the decarboxylation of cyt-OxAc (see below),
these values constitute, in principle, lower bounds (Table 2).
Although a more accurate determination of X cyt
is
1
prevented for the current system by rather similar f (3)
values, the data provide evidence that a large fraction of
cyt-Prv is derived from PPrv in both regimes.
Recruiting Leu-a and Glu-g to obtain f (2){mt-AcCoAC1}, Lys-a to derive f (2){cyt-AcCoA-C1}, and Ala-a to
get [ f (2) 1 f (2*)]{mt-Prv-C2}, employment of Eqn (12)
shows that, for the present growth conditions, X mt
2 cannot be
obtained because of degeneracy of f values. Similarly, with
f (2) {Lys-a} ˆ f (2) {cyt-AcCoA-C1} ˆ f (2) {Leu-a} ˆ
f (2) {mt-AcCoA-C1}, degeneracy of f values prevents from
calculating X cyt
using Eqn (13). The identity of the 13C
2
labeling labeling patterns indicates that the two AcCoA
pools are either fully equilibrated, or that one is derived
from the other. Future studies need to identify whether the
cytosolic and mitochondrial AcCoA pools are possibly
quite generally in rapid exchange due to the facilitated
diffusion catalysed by the carnitine shuttle.
The mt-OxAc pool is characterized by the flux ratio
X ana, which provides the fraction of mt-OxAc arising from
PPrv via Prv. As the carboxylation occurs in the cytosol,
X ana also establishes a ratio involving the transport flux of
cyt-OxAc into the mitochondria (Eqns 14). The f values
calculated for mt-OxAc (Table 3) allow determination of
X cyt
using Eqn (15). Selecting f (3), which exhibits the
3
largest difference when comparing Asp-a (Table 1), and
mt-OxAc-C2 (Table 3) for calculation of X cyt
3 , we obtain
cyt
that X cyt
3 . 0.88 for the aerobic and X 3 . 0.71 in the
anaerobic regime, i.e. the transfer of mt-OxAc into the
cytosol is of minor importance for the generation of
cyt-OxAc (Table 2). Hence, the transport of OxAc from the
cytosol to the mitochondria appears to be largely unidirectional, and this finding is in agreement with the view that
2476 H. Maaheimo et al. (Eur. J. Biochem. 268)
the OxAc carrier protein is actively transporting OxAc into
the mitochondria by symport of protons. Moreover, the data
provide evidence that the employment of the malate±
aspartate shuttle [65±67], which serves for transfer of
cytosolic NADH into the mitochondria, is small as it would
be expected to export mt-OxAc, which is not detected
(Table 1). The same holds for the succinate±fumarate
carrier [68,69]: as the glyoxylate cycle is switched off,
transport of succinate into the mitochondria (as well as
concomitant export of mitochondrial fumarate into the
cytosol) is apparently not required to balance cytosolic and
mitochondrial demands of succinate.
Finally, we have not invoked the import of citrate (or
isocitrate) from the cytosol into the mitochondria for data
interpretation, although S. cerevisiae can express a mitochondrial citrate transfer protein [70]. However, as
described for mammalian mitochondria [71], this protein
primarily serves for catalysing citrate efflux into the cytosol
in order to fulfill biosynthetic demands. As we find no
indication of a substantial transfer of citrate that was
generated in the cytosol into the mitochondria, our data
support the above indicated functional assignment of the
citrate transfer protein.
Comparison with previous analyses of E. coli cells
Previously reported METAFoR analyses of E. coli [1,9]
were obtained under very similar growth conditions as for
the S. cerevisiae cultivations described here, i.e. aerobic
and anaerobic shake flask cultures and glucosesupplemented minimal media were used to study central
carbon metabolism under exponential growth. Hence, these
studies provide an attractive framework to compare the
salient features of E. coli and S. cerevisiae central carbon
metabolism when glucose is supplied as the sole carbon
source.
Most strikingly, and in sharp contrast to E. coli, the
PenPp in S. cerevisiae operates exclusively for generating
NADPH and pentoses, i.e. virtually no PPrv is synthesized
through PenPp. Moreover, S. cerevisiae exhibits a significantly higher fraction of mt-Prv originating from malate
than E. coli, suggesting a higher in vivo activity of the
malic enzyme. In both organisms, OxAc is entirely
generated via anaplerosis in the anaerobic regime, illustrating a branched mode of operation for the TCA cycle. In the
aerobic regime, however, the contributions of anaplerosis
are rather different: about 75% and 30±50% contribution
for OxAc synthesis were observed in S. cerevisiae and
E. coli, respectively. This observation reflects the phenomenon of `glucose repression' of the TCA cycle in
S. cerevisiae, which is not observed for the E. coli.
Furthermore, Prv cleavage through pyruvate±formate
lyase, a characteristic feature of anaerobic E. coli metabolism, is not detected, simply because S. cerevisiae does not
possess such a lyase. Finally, we found that the glyoxylate
shunt is generally not active for both E. coli and
S. cerevisiae when glucose is provided as the sole carbon
source.
Conclusions
We have extended to application of BDF 13C labeling of
proteinogenic amino acids, an efficient analytical tool to
q FEBS 2001
study intermediary metabolism [1], to the compartmented
eukaryotic organism S. cerevisiae. This allowed (a)
unravelling the network of biosynthetic pathways activated
under glucose repression in both cytosol and mitochondria;
(b) determination of a large number of metabolic flux ratios
in the two compartments, and (c) characterization of
intercompartmental transport fluxes. The wealth of new
insights into compartmented yeast metabolism (Table 2)
obtained from the present investigation complements
previous 13C-NMR studies conducted for S. cerevisiae
[72±76], and enhances our understanding of yeast
metabolism in general [77].
In a manner previously demonstrated for prokaryotes
[1,5±12,14], the current methodology paves the way for
investigating the metabolic response of unicellular eukaryotes such as yeasts to genetic and environmental modulations. Future applications may include: (a) high-throughput
analyses in the framework of metabolic engineering in
biotechnology research [12,78,79]); (b) the joint employment of BDF 13C labeling with metabolic flux balancing
(reviewed in [12]) in order to obtain reliable estimates for
net fluxes [6]; and (c) the synergistic application of mass
spectrometry for analysis of the 13C labeling patterns
[12,80±82].
The rapid assessment of active biosynthetic pathways on
the level of the metabolic intermediates afforded by BDF
13
C labeling [1] is clearly of keen interest for future
metabolic engineering efforts. The fact that this approach
yields quantitative information about the in vivo malic
enzyme activity in S. cerevisiae is a telling example.
Engineered expression of malic enzyme may serve to
improve food quality [83], and heterologous malic enzymes
using both NADH and NADPH as cofactors may effectively
act as transhydrogenases, thus expanding the enzymatic
portfolio of S. cerevisiae. Evidently, such metabolic
engineering projects would greatly profit from monitoring
malic enzyme activity via determination of metabolic flux
ratios.
Overall, `metabolic profiling' using BDF 13C labeling of
proteinogenic amino acids [1] may well be an outstandingly
potent complement to both the `proteome approach',
aiming at the construction of complete protein maps [84],
and transcriptome analysis for genomewide exploration of
gene expression [85].
ACKNOWLEDGEMENT
We thank Prof. Kurt WuÈthrich for continuous support and again Dr M.
Hochuli for helpful discussions. The work was supported by a start-up
fund of the State University of New York to T. S., a postdoctoral
fellowship of VTT Biotechnology and the National Technology
Agency to H. M., and the Swiss Priority Program in Biotechnology.
REFERENCES
1. Szyperski, T. (1995) Biosynthetically directed fractional
13
C-labeling of proteinogenic amino acids. An efficient analytical
tool to investigate intermediary metabolism. Eur. J. Biochem. 232,
433±448.
2. Senn, H., Werner, B., Messerle, B.A., Weber, C., Traber, R. &
WuÈthrich, K. (1989) Stereospecific assignment of the methyl 1H
NMR lines of valine and leucine in polypeptides by nonrandom
13
C labeling. FEBS Lett. 249, 113±118.
q FEBS 2001
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2477
3. Neri, D., Szyperski, T., Otting, G., Senn, H. & WuÈthrich, K. (1989)
Stereospecific nuclear magnetic resonance assignments of the
methyl groups of valine and leucine in the DNA-binding domain
of the 434 repressor by biosynthetically directed fractional 13C
labeling. Biochemistry 28, 7510±7516.
4. WuÈthrich, K., Szyperski, T., Leiting, B. & Otting, G. (1992)
Biosynthetic pathways of the common proteinogenic amino acids
investigated by fractional 13C labeling and NMR spectroscopy. In
Frontiers and New Horizons in Amino Acid Research (Takai, K.,
ed.), pp. 41±48. Elsevier, Amsterdam.
5. Szyperski, T., Bailey, J.E. & WuÈthrich, K. (1996) Detecting and
dissecting metabolic fluxes using biosynthetic fractional 13C
labeling and two-dimensional NMR spectroscopy. Trends Biotechnol. 14, 453±459.
6. Sauer, U., Hatzimanikatis, V., Bailey, J.E., Hochuli, M., Szyperski,
T. & WuÈthrich, K. (1997) Metabolic fluxes in riboflavin-producing
Bacillus subtilis. Nat. Biotechnol. 15, 448±452.
7. Hochuli, M., Patzelt, H., Oesterhelt, D., WuÈthrich, K. & Szyperski,
T. (1999) Amino acid biosynthesis in the halophilic archaeon
Haloarcula hispanica. J. Bacteriol. 181, 3226±3237.
8. Fiaux, J., Andersson, C.I.J., Holmberg, N., BuÈlow, L., Kallio, P.T.,
Szyperski, T., Bailey, J.E. & WuÈthrich, K. (1999) 13C-NMR flux
ratio analysis of Escherichia coli central carbon metabolism in
microaerobic bioprocesses. J. Am. Chem. Soc. 121, 1407±1408.
9. Sauer, U., Lasko, D.R., Fiaux, J., Hochuli, M., Glaser, R., Szyperski,
T., WuÈthrich, K. & Bailey, J.E. (1999) Metabolic flux ratio
analysis of genetic and environmental modulations of Escherichia
coli central carbon metabolism. J. Bacteriol. 181, 6679±6688.
10. Schmidt, K., Nùrregaard, L.C., Pedersen, B., Meissner, A., Dues,
J.é., Nielsen, J.O. & Villadsen, J. (1999) Quantification of
intracellular metabolic fluxes from fractional enrichment and
13
C-13C coupling constraints on the isotopomer distribution in
labeled biomass components. Metabolic Eng. 1, 166±179.
11. Hochuli, M., Szyperski, T. & WuÈthrich, K. (2000) Deuterium
isotope effects on the central carbon metabolism of Escherichia
coli cells grown on a D2O-containing minimal medium. J. Biomol.
NMR 17, 33±42.
12. Szyperski, T. (1998) 13C-NMR, MS and metabolic flux balancing
in biotechnology research. Q. Rev. Biophys. 31, 41±106.
13. Reference withdrawn.
14. Szyperski, T., Glaser, R.W., Hochuli, M., Fiaux, J., Sauer, U.,
Bailey, J.E. & WuÈthrich, K. (1999) Bioreaction network topology
and metabolic flux ratio analysis by biosynthetic fractional 13C
labeling and two-dimensional NMR spectroscopy. Metabolic Eng.
1, 189±197.
15. Michal, G. (1998). Biochemical Pathways: an Atlas of Biochemistry and Molecular Biology. Wiley, New York.
16. Rose, A.H. & Harrison, J.S., eds (1989) The Yeasts: Metabolism
and Physiology. Academic Press, New York.
17. Strathern, J.N., Jones, E.W. & Broach, J.R., eds (1982) Molecular
Biology of the Yeast Saccharomyces. Metabolism and Gene
Expression. Cold Spring Harbor Laboratory Press, Plainview,
New York.
18. Zimmermann, F.K. & Entian, K.D. (1997) Yeast Sugar Metabolism: Biochemistry, Genetics, Biotechnology and Applications.
Technomic Publishing. Lancaster.
19. Mewes, H.W., Albermann, K., BaÈhr, M., Frishman, D., Gleissner,
A., Hani, J., Heumann, K., Kleine, K., Maierl, A., Oliver, S.G.,
Pfeiffer, F. & Zollner, A. (1997) Overview of the yeast genome.
Nature 387 (Suppl.), 7±65.
20. Hieter, P., Basset, D.E.J. & Valle, D. (1996) The yeast genome ± a
common currency. Nat. Genet. 13, 253±255.
21. Johnston, M. (1996) Genome sequencing: the complete code for a
eukaryotic cell. Curr. Biol. 6, 500±503.
22. Bodenhausen, G. & Ruben, D. (1980) Natural abundance
nitrogen-15 NMR by enhanced heteronuclear spectroscopy.
Chem. Phys. Lett. 69, 185±188.
23. Szyperski, T., Neri, D., Leiting, B., Otting, G. & WuÈthrich, K.
(1992) Support of 1H NMR assignments in proteins by
biosynthetically directed fractional 13C-labeling. J. Biomol. NMR
2, 323±334.
24. GuÈntert, P., DoÈtsch, V., Wider, G. & WuÈthrich, K. (1992)
Processing of multi-dimensional NMR data with the new software
PROSA. J. Biomol. NMR 2, 619±629.
25. Fraenkel, D.G. (1982) Carbohydrate metabolism. In Molecular
Biology of the Yeast Saccharomyces. Metabolism and Gene
Expression (Strathern, J.N., Jones, E.W. & Broach, J.R., eds),
pp. 1±37. Cold Spring Harbor Laboratory Press, Plainview, New
York.
26. Gancedo, C. & Serrano, R. (1989) Energy-yielding metabolism. In
The Yeasts, Vol. 3, Metabolism and Physiology of Yeasts (Rose,
A.H. & Harrison, J.S., eds), pp. 205±251. Academic Press, New
York.
27. Pronk, J.T., Steensma, H.Y. & van Dijken, J.P. (1996) Pyruvate
metabolism in Saccharomyces cerevisiae. Yeast 12, 1607±1633.
28. Kispal, G., Sumegi, B., Dietmeier, K., Bock, I., Gajdos, G.,
Tomcsanyi, T. & Sandor, A. (1993) Cloning and sequencing of a
cDNA encoding Saccharomyces cerevisiae carnitine acetyltransferase. J. Biol. Chem. 268, 1824±1829.
29. van Roermund, C.W.T., Hettema, E.H., van der Berg, M., Tabak,
H.F. & Wanders, R.J.A. (1999) Molecular characterization of
carnitine-dependent transport of acetyl-CoA from peroxisomes to
mitochondria in Saccharomyces cerevisiae and identification of a
plasma membrane carnitine transporter, Agp2p. EMBO J. 18,
5843±5852.
30. Palmieri, L., Vozza, A., Agrimi, G., De Marco, V., Runswick, M.J.,
Palmieri, F. & Walkers, J.E. (1999) Identification of the yeast
mitochondrial transporter for oxaloacetate and sulfate. J. Biol.
Chem. 274, 22184±22190.
31. Rohde, M., Lim, F. & Wallace, J.C. (1991) Electron microscopic
localization of pyruvate carboxylase in rat liver and Saccharomyces cerevisiae by immunogold procedures. Arch. Biochem.
Biophys. 290, 197±201.
32. van Urk, H., Schipper, D., Breedveld, G.J., Mak, P.R., Scheffers,
W.A. & van Dijken, J.P. (1989) Localization and kinetics of
pyruvate-metabolizing enzymes in relation to aerobic alcoholic
fermentation in Saccharomyces cerevisiae CBS 8066 and Candida
utilis CBS 621. Biochim. Biophys. Acta 992, 78±86.
È pfelsaÈurestoffwechsel bei
33. Fuck, E., StaÈrk, G. & Radler, F. (1973) A
Saccharomyces. II. Anreicherung und Eigenschaften eines Malatenzyms. Arch. Microbiol. 89, 223±231.
34. Boles, E., de Jong-Gubbels, P. & Pronk, J.T. (1998) Identification
and characterization of MAE1, the Saccharomyces cerevisiae
structural gene encoding mitochondrial malic enzyme. J. Bacteriol.
180, 2875±2882.
35. Nalecz, M.J., Nalecz, K.A. & Azzi, A. (1991) Purification and
functional characterisation of the pyruvate (monocarboxylate)
carrier from baker's yeast mitochondria (Saccharomyces cerevisiae). Biochim. Biophys. Acta 1079, 87±95.
36. Haarasilta, S. & Taskinen, R. (1977) Location of three key
enzymes of gluconeogenesis in baker's yeast. Arch. Microbiol.
113, 159±161.
37. Steensma, H.Y. (1997) From pyruvate to acetyl-coenzyme A and
oxaloacetate. In Yeast Sugar Metabolism. Biochemistry, Genetics,
Biotechnology and Applications (Zimmermann, F.K. & Entian,
K.-D., eds), pp. 339±357. Technomic Publishing, Lancaster, UK.
38. Atomi, H., Ueda, M., Suzuki, J., Kamada, Y. & Tanaka, A. (1993)
Presence of carnitine acetyltransferase in peroxisomes and in
mitochondria of oleic acid-grown Saccharomyces cerevisiae.
FEMS Microbiol. Lett. 112, 31±34.
39. Indiveri, C., Iacobazzi, V., Giangregorio, N. & Palmieri, F. (1998)
Bacterial overexpression, purification, and reconstitution of
the carnitine/acylcarnitine carrier from rat liver mitochondria.
Biochem. Biophys. Res. Commun. 249, 589±594.
2478 H. Maaheimo et al. (Eur. J. Biochem. 268)
40. Duntze, W., Neumann, D., Gancedo, J.M., Atzpodien, W. &
Holzer, H. (1969) Studies on the localization of the glyoxylate
cycle enzymes in Saccharomyces cerevisiae. Eur. J. Biochem. 10,
83±89.
41. McCammon, M.T., Veenhuis, M., Trapp, S.B. & Goodman, J.M.
(1990) Association of glyoxylate and beta-oxidation enzymes with
peroxisomes of Saccharomyces cerevisiae. J. Bacteriol. 172,
5816±5827.
42. Entian, K.-D. & SchuÈller, H.-J. (1997) Glucose repression (carbon
catabolite repression) in yeast. In Yeast Sugar Metabolism.
Biochemistry, Genetics, Biotechnology, and Applications (Zimmermann, F.K. & Entian, K.-D., eds). Technomic Publishing,
Lancaster, UK.
43. Minard, K.I. & McAlister-Henn, L. (1992) Glucose-induced
degradation of the MDH2 isozyme of malate dehydrogenase in
yeast. J. Biol. Chem. 267, 17458±17464.
44. McNeil, J.B., McIntosh, E.M., Taylor, B.V., Zhang, F.-R., Tang, S.
& Bognar, A.L. (1994) Cloning and molecular characterization of
three genes, including two genes encoding serine hydroxymethyltransferases, whose inactivation is required to render yeast
auxotrophic for glycine. J. Biol. Chem. 269, 9155±9165.
45. McNeil, J.B., Bognar, A.L. & Pearlman, R.E. (1996) In vivo
analysis of folate coenzymes and their compartmentation in
Saccharomyces cerevisiae. Genetics 142, 371±381.
46. Ogur, M., Liu, T.N., Cheung, I., Paulavicius, I., Wales, W.,
Mehnert, D. & Blaise, D. (1977) `Active' one-carbon generation in
Saccharomyces cerevisiae. J. Bacteriol. 129, 926±933.
47. Tzagoloff, A. (1982) Mitochondria. Plenum Press, New York.
48. Liu, J.Q., Nagata, S., Dairi, T., Misosno, H., Shimizu, S. &
Yamada, H. (1997) The GLY1 gene of Saccharomyces cerevisiae
encodes a low-specific l-threonine aldolase that catalyzes
cleavage of l-allo-threonine and l-threonine to glycine ±
expression of the gene in Escherichia coli and purification and
characterization of the enzyme. Eur. J. Biochem. 245, 289±293.
49. Ryan, E.D., Tracy, J.W. & Kohlhaw, G.B. (1973) Subcellular
localization of the leucine biosynthetic enzymes in yeast.
J. Bacteriol. 116, 222±225.
50. Ryan, E.D. & Kohlhaw, G.B. (1974) Subcellular localization of
isoleucine-valine biosynthetic enzymes in yeast. J. Bacteriol. 120,
631±637.
51. Chen, S., Brockenbrough, J.S., Dove, J.E. & Aris, J.P. (1997)
Homocitrate synthase is located in the nucleus in the yeast
Saccharomyces cerevisiae. J. Biol. Chem. 272, 10839±10846.
52. Monschau, N., Stahmann, K.P., Sahm, H., McNeil, J.B. & Bognar,
A.L. (1997) Identification of Saccharomyces cerevisiae GLY1 as a
threonine aldolase: a key enzyme in glycine biosynthesis. FEMS
Microbiol. Lett. 150, 55±60.
53. Hradzina, G. & Jensen, R.A. (1992) Spatial organization of
enzymes in plant metabolic pathways. Annu. Rev. Plant Physiol.
Plant Mol. Biol. 43, 241±267.
54. Sherry, A.D., Sumegi, B., Miller, B., Cottam, G.L., Gavva, S.,
Jones, J.G. & Malloy, C.R. (1994) Orientation-conserved transfer
of symmetric Krebs cycle intermediates in mammalian tissue.
Biochemistry 33, 6268±6275.
55. Ovadi, J. & Srere, P.A. (1992) Channel your energies. Trends
Biochem. Sci. 17, 445±447.
56. Sherry, A.D. & Malloy, C.R. (1996) Isotopic methods for probing
organization of cellular metabolism. Cell Biochem. Func. 14,
259±268.
57. Kholodenko, B.N., Westerhoff, H.V. & Cascante, M. (1996) Effect
of channeling on the concentration of bulk-phase intermediates as
cytosolic proteins become more concentrated. Biochem. J. 313,
921±926.
58. Chasin, L.A., Feldman, A., Konstam, M. & Urlaub, G. (1974)
Isolation of a chinese hamster cell mutants deficient in
dihydrofolate reductase activity. Proc. Natl Acad. Sci. USA 71,
718±722.
q FEBS 2001
59. Gancedo, J.M. & Lagunas, R. (1973) Contribution of the pentosephosphate pathway to glucose metabolism in Saccharomyces
cerevisiae: a critical analysis on the use of labelled glucose. Plant
Sci. Lett. 1, 193±200.
60. Boles, E., Lehnert, W. & Zimmermann, F.K. (1993) The role of the
NAD-dependent glutamate dehydrogenase in restoring growth on
glucose of a Saccharomyces cerevisiae phosphoglucose isomerase
mutant. Eur. J. Biochem. 217, 469±477.
61. Bruinenberg, P.M., Jonker, R., van Dijken, J.P. & Scheffers, W.A.
(1985) Utilization of formate as an additional energy source by
glucose-limited chemostat cultures of Candida utilis CBS621 and
Saccharomyces cerevisiae CBS8066. Evidence for the absence
of transhydrogenase activity in yeasts. Arch. Microbiol. 142,
302±306.
62. Wales, D.S., Cartledge, T.G. & Lloyd, D. (1980) Effects of glucose
repression and anaerobiosis on the activities and subcellular
distribution of tricarboxylic acid cycle enzymes in Saccharomyces
carlsbergensis. J. Gen. Microbiol. 116, 93±98.
63. Zitomer, R.S. & Lowry, C.V. (1992) Regulation of gene
expression by oxygen in Saccharomyces cerevisiae. Microbiol.
Rev. 56, 1±11.
64. Polakis, E.S. & Bartley, W. (1965) Changes in the enzyme
activities of Saccharomyces cerevisiae during aerobic growth on
different carbon sources. Biochem. J. 97, 284±297.
65. Lancar-Benda, J., Foucher, B. & Saint-Macary, M. (1996)
Characterization, purification and properties of the yeast mitochondrial dicarboxylate carrier (Saccharomyces cerevisiae).
Biochimie 78, 195±200.
66. Palmieri, L., Palmieri, F., Runswick, M.J. & Walker, J.E. (1996)
Identification by bacterial expression and functional reconstitution
of the yeast genomic sequence encoding the mitochondrial
dicarboxylate carrier protein. FEBS Lett. 399, 299±302.
67. Palmieri, L., Vozza, A., HoÈnlinger, A., Dietmeier, K., Palmisano,
A., Zara, V. & Palmieri, F. (1999) The mitochondrial dicarboxylate
carrier is essential for the growth of Saccharomyces cerevisiae on
ethanol or acetate as the sole carbon source. Mol. Microbiol. 31,
569±577.
68. Bojunga, N., KoÈtter, P. & Entian, K.-D. (1998) The succinate/
fumarate transporter Acr1p of Saccharomyces cerevisiae is part of
the gluconeogenetic pathway and its expression is regulated by
Cat8p. Mol. Gen. Genet. 260, 453±461.
69. Palmieri, L., Lasorsa, F.M., De Palma, A., Palmieri, F., Runswick,
M.J. & Walker, J.E. (1997) Identification of the yeast ACR1 gene
product as a succinate-fumarate transporter essential for growth on
ethanol or acetate. FEBS Lett. 417, 114±118.
70. Kaplan, R.S., Mayor, J.A., Gremse, D.A. & Wood, D.A. (1995)
High level expression and characterization of the mitochondrial
transport protein from the yeast Saccharomyces cerevisiae. J. Biol.
Chem. 270, 4108±4114.
71. Conover, T.E. (1987) Does citrate transport supply both acetyl
groups and NADPH for cytoplasmic fatty acid synthesis? Trends
Biochem. Sci. 12, 88±89.
72. Pasternack, L.B., Laude, D.A.J. & Appling, D.R. (1992) 13C NMR
detection of folate-mediated serine and glycine synthesis in vivo in
Saccharomyces cerevisiae. Biochemistry 31, 8713±8719.
73. Pasternack, L.B., Laude, D.A.J. & Appling, D.R. (1994) Wholecell detection by 13C NMR of metabolic flux through the
C1-tetrahydrofolate synthase/serine hydroxymethyltransferase
enzyme system and effect of antifolate exposure in Saccharomyces
cerevisiae. Biochemistry 33, 7166±7173.
74. Pasternack, L.B., Littlepage, L.E., Laude, D.A.J. & Appling, D.R.
(1996) 13C NMR analysis of the use of alternative donors to the
tetrahydrofolate-dependent one-carbon pools in Saccharomyces
cerevisiae. Arch. Biochem. Biophys. 326, 158±165.
75. Tran-Dinh, S., Beganton, F., Nguyen, T.-T., Bouet, F. & Herve, M.
(1996) Mathematical model for evaluating the Krebs cycle flux
with non-constant glutamate-pool size by 13C-NMR spectroscopy.
q FEBS 2001
76.
77.
78.
79.
80.
81.
82.
83.
Central carbon metabolism of Saccharomyces cerevisiae (Eur. J. Biochem. 268) 2479
Evidence for the existence of two types of Krebs cycles in cells.
Eur. J. Biochem. 242, 220±227.
Tran-Dinh, S., Bouet, F., Huynh, Q.-T. & Herve, M. (1996)
Mathematical models for determining metabolic fluxes through
the citric acid and glyoxylate cycles in Saccharomyces cerevisiae
by 13C-NMR spectroscopy. Eur. J. Biochem. 242, 770±778.
Flores, C., Rodriguez, C., Petit, T. & Gancedo, C. (2000)
Carbohydrate and energy-yielding metabolism in non-conventional
yeasts. FEMS Microbiol. Rev. 24, 507±529.
Bailey, J.E. (1991) Toward a science of metabolic engineering.
Science 252, 1668±1675.
Stephanopoulos, G. & Sinskey, A.J. (1993) Metabolic engineering
± methodologies and future prospects. Trends Biotech. Sci. 11,
392±396.
Christensen, B. & Nielsen, J. (1999) Isotopomer analysis using
GC-MS. Metabolic Eng. 1, 282±290.
Wittmann, C. & Heinzle, C. (1999) Mass spectrometry for
metabolic flux analysis. Biotechnol. Bioeng. 62, 739±750.
Dauner, M. & Sauer, U. (2000) GC-MS analysis of amino acids
rapidly provides rich information for isotopomer balancing.
Biotechnol. Prog. 16, 642±649.
Volschenk, H., Viljoen, M., Grobler, J., Petzold, B., Bauer, F.,
84.
85.
86.
87.
88.
89.
Subden, R., Young, R., Lonvaud, A., Denayrolles, M. & van
Vuuren, H. (1997) Engineering pathways for malate degradation in
Saccharomyces cerevisiae. Nat. Biotechnol. 15, 253±257.
James, P. (1997) Protein identification in the post-genome era: the
rapid rise of proteomics. Quart. Rev. Biophys. 30, 279±331.
Gerhold, D., Rushmore, T. & Caskey, C.T. (1999) DNA chips:
promising toys have become powerful tools. Trends Biochem. Sci.
24, 168±173.
Theobald, U., Mailinger, W., Baltes, M., Rizzi, M. & Reuss, M.
(1997) In vivo analysis of metabolic dynamics in Saccharomyces
cerevisiae: I. Experimental observations. Biotechnol. Bioeng. 55,
305±316.
Mailinger, W., Baumeister, A., Reuss, M. & Rizzi, M. (1998)
Rapid and highly automated determination of adenine and pyridine
nucleotides in extracts of Saccharomyces cerevisiae using a micro
robotic sample preparation-HPLC system. J. Biotechnol. 63,
155±157.
IUPAC-IUB Commission on Biochemical Nomenclature (1970)
Abbreviations and symbols for the description of the conformation
of polypeptide chains. Biochemistry 9, 3471±3479.
Hansen, P.E. (1988) Isotope effects in nuclear shielding. Prog.
NMR Spectrosc. 20, 207±255.