Molecular & Biochemical Parasitology 149 (2006) 1–9
Review
Energy metabolism of trypanosomatids: Adaptation to
available carbon sources
Frédéric Bringaud ∗ , Loı̈c Rivière, Virginie Coustou
Laboratoire de Génomique Fonctionnelle des Trypanosomatides, Université Victor Segalen Bordeaux 2, UMR-5162 CNRS,
146 rue Léo Saignat, 33076 Bordeaux Cedex, France
Received 24 February 2006; received in revised form 30 March 2006; accepted 31 March 2006
Available online 25 April 2006
Abstract
Some development stages of the trypanosomatid protozoan parasites are well adapted to in vitro culture. They can be maintained in rich medium
containing large excess of glucose and amino acids, which they use as carbon sources for ATP production. Under these growth conditions, carbon
sources are converted into partially oxidized end products by so-called aerobic fermentation. Surprisingly, some species, such as the Trypanosoma
brucei, Trypanosoma cruzi and Crithidia insect stages, prefer consuming glucose to amino acids, although their natural habitat is l-proline-rich.
This review focuses on recent progress in understanding glucose and l-proline metabolism of insect stages, how these metabolic processes are
regulated, and the rationale of the aerobic fermentation strategies developed by these parasites.
© 2006 Elsevier B.V. All rights reserved.
Keywords: Trypanosoma; Leishmania; Energy metabolism; ATP production; Glucose; l-Proline; Aerobic fermentation; Glucose-repression effect
Contents
1.
2.
3.
4.
5.
6.
7.
8.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Overview of glucose and amino acid catabolic pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Redox balances and succinic fermentation pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acetyl-CoA metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Lactate production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
The ATP production dilemma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Why do trypanosomatids use aerobic fermentation? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction
Trypanosomatids are parasitic protozoans, among which
several species cause serious diseases in humans such as
sleeping sickness (Trypanosoma brucei), Chagas disease (Trypanosoma cruzi) and leishmaniasis (Leishmania spp.). The life
cycle of trypanosomatids can be complex, sometimes involving
numerous developmental stages in several hosts. Most of the
∗
Corresponding author. Tel.: +33 557574632; fax: +33 557574803.
E-mail address: [email protected] (F. Bringaud).
0166-6851/$ – see front matter © 2006 Elsevier B.V. All rights reserved.
doi:10.1016/j.molbiopara.2006.03.017
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studied trypanosomatids have developed a digenetic life-style
with one or several vertebrate host(s) and a hematophage insect
vector that allows their transmission between vertebrate hosts.
The life cycle of the human pathogens, which are the most
studied species, is presented in Fig. 1. Recently, the genome
sequencing projects of T. brucei (927 strain) [1], T. cruzi (CL
Brener strain) [2] and Leishmania major (Friedlin strain) [3]
have been completed, providing wonderful tools to determine
their metabolic complexities [1].
Most, if not all, trypanosomatids depend on the available carbon sources present in their hosts for their energy metabolism
2
F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
Fig. 1. Life cycle of human infective trypanosomatids (adapted from [52]). The
circular arrows show the duplicative forms. Abbreviations: A, amastigote; BT,
bloodstream trypomastigote; E, epimastigote; M, metacyclic; P, promastigote;
PT, procyclic trypomastigote; T, trypomastigote.
(see Table 1). First, the trypomastigote forms of T. brucei and T.
cruzi use glucose, which is abundant in the fluids of their vertebrate host(s) [4,5]. Second, hematophage insect vectors obtain
their energy from l-proline and/or l-glutamine, the prominent
constituent of their hemolymph and tissue fluids [6]. Consequently, the insect stages of trypanosomatids rely on amino acid
catabolism, with a preference for l-proline [4,7–12]. It is noteworthy that T. cruzi (but not T. brucei and L. major) has the
capacity to utilize d-proline, in addition to l-proline (expression
of a proline racemase), and l-histidine [13,14]. In addition, the
insect stage of Leishmania spp. (promastigote) also expresses
the enzymatic arsenal to metabolize disaccharides, presumably
as an adaptation to its hematophage insect vectors (sandflies),
which can also feed on nectar and aphid honeydew [1]. Third,
the plant parasites Phytomonas spp. consume diverse carbohydrates produced by plants [15]. Fourth, the carbon sources used
by intracellular stages depend on the specific host compartment
in which they live: T. cruzi amastigotes reside in the cytoplasm of
the host cells with ready access to sugar phosphates and develop
a glucose-based metabolism [4], whereas Leishmania amastigotes proliferate in the lysosomal compartment of macrophages
and prefer fatty acids as energy sources [16,17].
All analysed trypanosomatid adaptive forms, with the exception of Leishmania spp., prefer glucose when grown in richmedium, including those in which glucose is not the natural
carbon source (Table 1). Since glucose-rich media are routinely
used to grow these parasites, glucose metabolism has received
most attention and relatively little is known about their aminoacid and fatty-acid metabolism. This review discusses recent
Table 1
End products of the metabolism of carbon sources by trypanosomatids culturated in glucose-rich media
Parasite
Stagea
Host
Carbon source(s)b
Excreted end products in glucose-rich conditionsc
References
T. brucei
T. brucei
BTd
PT
Vertebrate
Insecte
Glucose
Glucose
l-Prolinef
l-Threonine
Pyr
CO2 , Suc, Ace, l-Lac
CO2 , Suc, Ace
Ace, Glycine
[5]
[21,28]
[21,22]
[25]
T. cruzi
T. cruzi
T. cruzi
T
A
E
Vertebrate
Vertebrate
Insecte
Glucose
Glucose
Glucose
Amino acidsf
CO2 , Suc, Ace
CO2 , Ace, Gly, Pyr, Suc
CO2 , Suc, l-Ala, Ace
nd
[4]
[53]
[4]
[4,8,10]
Leishmania spp.
A
Vertebrate
Fatty acids
Glucose
nd
CO2 , l-Ala, Ace, Suc
[16,17]
[54]
Leishmania spp.
P
Insectg
Amino acids
Glucose
nd
CO2 , Suc, l-Ala, Ace, d-Lac
[7,9]
[54,55]
Crithidia spp.
Ch
Insecte
Glucose
Amino acidsf
CO2 , Suc, Eth, Ace
nd
[9,56]
[9]
Phytomonas spp.
P
Plant
Glucose
CO2 , Ace, Eth
[15]
a
For abbreviation see Fig. 1; Ch, choanomastigote.
In boldface are indicated the preferred carbon source consumed in glucose-rich medium.
c Synthesis of published data. Since some differences are observed depending on the publication and on the sub-species analysed, only the major end products
excreted are mentioned. Abbreviations: Ace, acetate; l-Ala, l-alanine; Eth, ethanol; Gly, glycerol; d-Lac and l-Lac, d- and l-lactate; nd, not determined; Pyr,
pyruvate; Suc, succinate.
d Slender bloodstream form grown under aerobic conditions.
e Hematophage only (tsetse flies, triatomines and mosquitoes, respectively).
f Preferred carbon source in the absence of glucose (l-proline is the carbon source of the corresponding insect vector).
g Hematophage and plant sucking insects (sandflies).
b
F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
data concerning the energy metabolism in trypanosomatids,
highlighting the adaptation processes developed by these parasites in response to the available carbon sources. Most of the
recent data concern the procyclic trypanosomes, since T. brucei
has recently become the favourite model to study metabolism
and other processes shared with other trypanosomatid species.
Indeed, RNA interference (RNAi) has been extensively developed in T. brucei. It is a very powerful reverse genetic tool,
which inhibits specifically the expression of a target gene. This
technique failed to be functional in T. cruzi and Leishmania spp.
[18].
2. Overview of glucose and amino acid catabolic
pathways
Within the trypanosomatids, the slender bloodstream form
of T. brucei has the simplest energy metabolism, which is only
based on glycolysis of glucose present in the blood of the vertebrate hosts. The glycolytic pathway is organised in such a
way that the first seven enzymes converting glucose into 3phosphoglycerate are inside the glycosomes (peroxisome-like
organelles), while the last three enzymes of the pathway reside
in the cytosol (not shown here, see Fig. 2 for individual steps).
Under aerobic conditions, pyruvate is the only excreted end
product and net ATP synthesis occurs in the cytosol, in the reaction catalysed by pyruvate kinase (PYK, step 13), whereas in
the glycosomes the consumption (steps 1 and 3) and production (step 9) of ATP are balanced. Similarly the glycosomal
redox balance is maintained, since, the NADH produced by
glyceraldehyde-3-phosphate dehydrogenase (step 8) is reoxidized via a glycerol 3-phosphate shuttle (steps 6 and 45) and
the alternative oxidase present in the mitochondrion (step 47).
The relatively simple glucose/energy metabolism of this adaptive form has been extensively studied and will not be further
addressed in this review (for recent reviews see [19,20]).
In contrast, glucose catabolism in all other trypanosomatids
(or adaptive forms) analysed so far involves more elaborate
metabolic networks both within the glycosomes and the fully
developed mitochondrion. This is exemplified by the model proposed for the procyclic T. brucei presented in Fig. 2, which shows
three main differences compared with the slender bloodstream
form of T. brucei. First, phosphoglycerate kinase (PGK, step
10) is located in the cytosol and, therefore, 3-phosphoglycerate
is produced in the cytosol. Second, the glycosomes contain
two additional kinases converting phosphoenolpyruvate (PEP)
into malate (PEPCK: PEP carboxykinase, step 14) or pyruvate
(pyruvate phosphate dikinase, step 15). The CO2 fixation step
catalysed by PEPCK is the initial step of a branched pathway
leading to the production of succinate, a major end product
excreted by most trypanosomatids (Table 1). Third, pyruvate
is located at a metabolic branching point leading to several
excreted end products, such as acetate, l-alanine, ethanol and
l-lactate (Fig. 2): acetate is a major end product formed in
the mitochondrion of all these trypanosomatids and excreted
by simple diffusion across the mitochondrial and cytoplasmic
membranes; few genera, such as Phytomonas and Crithidia,
also produce ethanol from pyruvate (steps 52 and 53), and most
3
trypanosomatids also convert a significant part of pyruvate
into l-lactate (step 23) and/or by transamination into l-alanine
(step 24).
In addition, some adaptive forms, especially the insect stages,
use amino acids present in their host for energy production. For
instance, insect stages of T. brucei, T. cruzi, Leishmania spp. and
Crithidia spp. dwell in the l-proline-rich environment found in
the insect fluids, and thus particularly appreciate this carbon
source. Indeed, procyclic trypanosomes convert l-proline into
succinate and to a lesser extent into acetate, when incubated
in regular glucose-rich medium [21,22] (Fig. 2). Surprisingly, T.
brucei procyclics, as well as T. cruzi promastigotes and Crithidia
fasciculata, preferentially consume glucose when both glucose
and amino acids are available [9,12]. Nevertheless, recent reports
confirmed the essential role of l-proline metabolism in energy
production of insect-stage trypanosomes. First, procyclic forms
of several T. brucei strains were successfully adapted to glucosedepleted medium, with no significant effect on growth rate
[12,23,24]. Second, in the absence of glucose, l-proline is the
only carbon source sustaining growth of the parasite [12]. Third,
in glucose-depleted medium, the rate of l-proline consumption
is approximately six-fold increased in insect-stage forms of two
different T. brucei strains analysed, which also implies that glucose exerts a negative control on l-proline metabolism [12].
Interestingly, the ability of these parasites to adapt to glucosedepleted conditions may be altered by long-term in vitro culture
in glucose-rich medium. Indeed, doubling time and morphology
of another T. brucei procyclic strain (TREU927) are affected by
the absence of glucose, probably as a consequence of a relative low increase of the rate of l-proline consumption (two-fold
versus six-fold in the well adapted strains) [22]. In addition
to l-proline, the procyclic trypanosomes also convert important amounts of l-threonine into l-glycine and acetate, but its
metabolic role is not clear yet [25].
3. Redox balances and succinic fermentation pathways
In most trypanosomatids, succinate is a major end product
excreted from glucose metabolism (Table 1), but the pathway
leading to its production has been the topic of a long-standing
debate [26,27]. The controversy concerned the relevance of the
NADH-dependent fumarate reductase (FRD) activity detected
in most trypanosomatids (see references in [28]), whose contribution in succinate production was not clearly demonstrated.
The identification in T. brucei of two FRD isoforms closed
this debate, since succinate production dramatically decreased
in a mutant depleted for both glycosomal and mitochondrial
FRD activities [28,29]. According to the current model, succinate is produced from cytosolic PEP by a branched pathway
(Fig. 2). The glycosomal PEPCK (step 14) and malate dehydrogenase (step 16) convert PEP into malate, which is converted into
fumarate by two fumarase isoforms. Fumarate is finally reduced
into the excreted succinate by the glycosomal and mitochondrial
FRD isoforms (steps 18 and 20). We recently identified in procyclic trypanosomes two class I fumarases respectively located
in the cytoplasm (FHc, step 17) and the mitochondrion (FHm,
step 19) (Coustou et al., unpublished data). To our surprise none
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F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
Fig. 2. Schematic representation of carbon source metabolism in the procyclic form of T. brucei grown in glucose-rich medium. Dark, grey and light grey arrows represent enzymatic steps of d-glucose, l-threonine and l-proline/l-glutamine metabolism, respectively. Excreted end products (acetate, l-alanine, glycerol, l-glycine,
l- or d-lactate, succinate and CO2 ) of d-glucose, l-threonine and l-proline/l-glutamine metabolism are in white characters on a black, grey and light grey background,
respectively. Arrows with different thicknesses tentatively represent the metabolic flux at each enzymatic step. Dashed arrows indicate steps, which are supposed to
occur at a background level or not at all, under standard growth conditions (glucose-rich medium). The glycosomal and mitochondrial compartments, the tricarboxylic
acid (TCA) cycle and the links to the pentose phosphate pathway (PPP) and lipid biosynthesis pathway are indicated. The mitochondrial outer membrane is permeable
to metabolites and is only shown in the vicinity of the schematic electron-transport chain. d-Lactate and ethanol production pathways (dashed boxes) have only been
observed in Leishmania spp. and Phytomonas/Crithidia, respectively. Abbreviations: AA, amino acid; AOB, amino oxobutyrate; 1,3BPGA, 1,3-bisphosphoglycerate;
C, cytochrome c; Cit, citrate; CoASH, coenzyme A; DHAP, dihydroxyacetone phosphate; F-6-P, fructose 6-phosphate; FBP, fructose 1,6-bisphosphate; G-3-P, glyceraldehyde 3-phosphate; G-6-P, glucose 6-phosphate; GLU, glutamate; Gly-3-P, glycerol 3-phosphate; IsoCit, isocitrate; 2Ket, 2-ketoglutarate; OA, 2-oxoacid; Oxac,
oxaloacetate; PEP, phosphoenolpyruvate; 3-PGA, 3-phosphoglycerate; Pi, inorganic phosphate; PPi, inorganic pyrophosphate; ␥SAG, glutamate ␥-semialdehyde;
SucCoA, succinyl-CoA; T[SH]2 , reduced form of trypanothione; UQ, ubiquinone pool. Enzymes are: 1, hexokinase; 2, glucose-6-phosphate isomerase; 3,
phosphofructokinase; 4, aldolase; 5, triose-phosphate isomerase; 6, glycerol-3-phosphate dehydrogenase; 7, glycerol kinase; 8, glyceraldehyde-3-phosphate
dehydrogenase; 9, glycosomal phosphoglycerate kinase; 10, cytosolic phosphoglycerate kinase; 11, phosphoglycerate mutase; 12, enolase; 13, pyruvate kinase;
14, phosphoenolpyruvate carboxykinase; 15, pyruvate phosphate dikinase; 16, glycosomal malate dehydrogenase; 17, cytosolic (and glycosomal) fumarase
(FHc); 18, glycosomal NADH-dependent fumarate reductase; 19, mitochondrial fumarase (FHm); 20, mitochondrial NADH-dependent fumarate reductase; 21,
glycosomal adenylate kinase; 22, malic enzyme; 23, unknown enzyme; 24, alanine aminotransferase; 25, pyruvate dehydrogenase complex; 26, acetate:succinate
F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
of the fumarase isoforms (FHc and FHm) are detectable in glycosomes, whereas this compartment contains the other enzymes
of the glycosomal succinate production pathway. The absence of
a glycosomal fumarase was recently confirmed by a proteomics
analysis of glycosomes [30]. However, the previous detection of
a significant fumarase activity in the glycosomal fraction of procyclic trypanosomes, is puzzling [28]. These contradictory data
may be explained by the presence of a cryptic PTS1 glycosomal targeting motif at the C-terminal extremity of FHc (AKLV),
which is also present at the same relative position in the T. cruzi
(SKLL) and L. major (SKTLA) orthologous proteins. Indeed, we
cannot exclude the possibility that the cryptic PTS1 sequence,
after a form of decrypting dependent on environmental conditions and/or strain specificities, may be responsible for transient
(and partial) glycosomal localization of FHc. We also propose
that the unexpected cytosolic location of FHc is the consequence
of the absolute requirement of fumarate in the cytosol, which is
used as an electron acceptor (instead of NAD+ ) by an essential cytosolic enzyme (dihydroorotate dehydrogenase) of the de
novo pyrimidine biosynthesis pathway [31,32].
The main role of the succinate production pathway (succinic
fermentation) is probably to maintain the glycosomal redox balance, by providing two glycosomal oxidoreductase enzymes
(steps 16 and 18) allowing reoxidation of NADH produced
by glyceraldehyde-3-phosphate dehydrogenase in the glycolytic
pathway (step 8). Compared with the more common lactic
fermentation, which involves a single oxidoreductase enzyme
(lactate dehydrogenase), succinic fermentation offers the significant advantage of requiring only half of the PEP produced to
maintain the NAD+ /NADH balance. The remaining PEP is converted into acetate, l-lactate, l-alanine and/or ethanol depending
on the species. This branched glucose catabolic pathway may
provide an important flexibility for adaptation to rapid environmental changes. Another level of flexibility could be gained
by the involvement of both succinic fermentation and respiration. Indeed, procyclic trypanosomes have a functional glycerol
3-phosphate shuttle, which may also be used to maintain the
glycosomal redox balance, feeding the mitochondrial electron
transport chain, as is found in T. brucei bloodstream forms (D.G.
Guerra and P.A.M. Michels, personal communication). This
metabolic flexibility is illustrated by the analysis of T. brucei procyclic mutants. For example, depletion of the glycosomal FRD
activity does not affect the growth rate and glucose catabolism
[28], suggesting that an alternative way is used in this mutant to
substitute for the glycosomal FRD step. Succinic fermentation
also occurs in the mitochondrion, although its essential role in
maintaining this organelle’s redox balance is debatable, due to
the presence of mitochondrial NADH dehydrogenases linked to
the electron transport chain (complex I and step 46).
5
4. Acetyl-CoA metabolism
Acetyl-CoA is a key intermediary metabolite at a crossroad
between catabolic and anabolic pathways, whose production
is essential for most cells. In the procyclic trypanosomes,
catabolism of each of the major carbon sources (glucose, lproline and l-threonine) leads to acetyl-CoA formation (Fig. 2).
Most of the pyruvate produced from glucose by glycolysis is
decarboxylated to acetyl-CoA by the mitochondrial pyruvate
dehydrogenase complex (PDH, step 25). Procyclic cells in
which, the E1␣ subunit of PDH was depleted by RNAi are
viable, although they show a reduced growth rate [33]. This data
is in agreement with the observation that glucose metabolism
is not essential for procyclic trypanosomes [12,22–24] and that
acetyl-CoA is also produced from l-proline and l-threonine
metabolism [22,34]. Early models proposed that, in most trypanosomatids, acetyl-CoA produced from glucose metabolism
is converted into CO2 through the tricarboxylic acid (TCA)
cycle and into acetate (Table 1). However, the recent analysis
of an aconitase (step 30) knockout mutant revealed that acetylCoA does not fuel the TCA cycle of the procyclic trypanosomes
grown in glucose-rich medium [21]. Consequently, most of
acetyl-CoA (if not all) is converted into the excreted acetate. It is
noteworthy that all the trypanosomatid adaptive forms analysed
so far (except the T. brucei bloodstream forms) produce acetate
from glucose (Table 1), highlighting the importance of this
pathway, probably for ATP production (see Section 6). Van
Hellemond et al. demonstrated that acetate is produced by a
two-enzyme cycle: acetate:succinate CoA-transferase (ASCT,
step 26) transfers the CoA moiety of acetyl-CoA to succinate,
yielding acetate and succinyl-CoA that is subsequently reconverted into succinate by succinyl-CoA synthetase (SCS, step 28)
with concomitant production of ATP [35]. ASCT is encoded by
a newly identified member of the eukaryotic CoA-transferase
gene family [36]. A T. brucei procyclic knockout mutant
depleted for ASCT showed a reduced acetate production,
confirming the role of this enzyme in acetate production [36].
However, ASCT mutants still excrete acetate from glucose
metabolism, implying that ASCT is not the only acetateproducing pathway in this parasite. The only alternative gene
candidate identified so far in the genome of T. brucei encodes an
AMP-forming acetyl-CoA synthetase. In Aspergillus nidulans,
this enzyme produces acetate and ATP from acetyl-CoA
and AMP, however, in most organisms analysed the reverse
reaction occurs [37]. This trypanosomal enzyme, and its
possible role in acetate production, are currently under further
investigation.
As mentioned above, acetyl-CoA sits at the crossroad of intermediary metabolism, including catabolism and anabolism of
CoA-transferase; 27, unknown enzyme; 28, succinyl-CoA synthetase; 29, citrate synthase; 30, aconitase; 31, isocitrate dehydrogenase; 32, 2-ketoglutarate dehydrogenase complex; 33, succinate dehydrogenase (complex II of the respiratory chain); 34, mitochondrial malate dehydrogenase; 35, L-proline dehydrogenase; 36,
pyrroline-5 carboxylate dehydrogenase; 37, l-glutamine deaminase; 38, glutamate aminotransferase; 39, glutamate dehydrogenase; 40, l-threonine dehydrogenase;
41, acetyl-CoA:glycine C-acetyltransferase; 42, citrate lyase; 43, acetyl-l-carnitine transferase; 44, acetyl-l-carnitine transferase; 45, FAD-dependent glycerol-3phosphate dehydrogenase; 46, rotenone-insensitive NADH dehydrogenase; 47, alternative oxidase; 48, F0 F1 -ATP synthase; 49, spontaneous reaction; 50, glyoxalase
I; 51, glyoxalase II; 52, pyruvate decarboxylase; 53, NAD-linked alcohol dehydrogenase; I, II, III and IV, complexes of the respiratory chain.
6
F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
carbohydrates, amino acids and fatty acids. For instance, acetylCoA produced by various catabolic pathways can be used for
lipid biosynthesis, as has also shown to be the case in procyclic
trypanosomes, where glucose and l-threonine were found to
be substrates for fatty acid biosynthesis, through acetyl-CoA
production [22,34]. Since this process occurs inside the mitochondrion, whereas biosynthesis of fatty acids takes place in
the cytosol, acetyl-CoA has to be transferred from the mitochondrion to the cytoplasm. Van Weelden et al. proposed that
acetyl-CoA is exchanged between both compartments via citrate [22]; acetyl-CoA and oxaloacetate are condensed to citrate by citrate synthase (step 29), which is then exchanged for
malate, from the cytosol to be converted back into acetyl-CoA
and oxaloacetate by citrate lyase (step 42). Although the trypanosomatids genome encodes citrate synthase and citrate lyase,
available experimental data suggest an alternative hypothesis.
The citrate/malate (or citrate/oxaloacetate) shuttle is widespread
in eukaryotes, however, it is not universal. For instance, Saccharomyces cerevisiae lacks citrate lyase and uses instead two other
pathways, i.e. the l-carnitine acyl-transferase system, which can
exchange acetyl moieties instead of acyl groups [38] and the
cytosolic acetyl-CoA synthetase, which converts acetate into
acetyl-CoA [39]. Trypanosomatids may also use these alternative pathways. First, T. brucei contains high levels of l-carnitine
acetyl-transferase activity described as a key step in lipid biosynthesis from l-threonine-derived acetyl-CoA [34,40]. Second,
trypanosomatids produce considerable amounts of acetate which
diffuses across membranes and might be converted in acetylCoA by the putative AMP-forming acetyl-CoA synthetase (see
above), which is possibly a cytosolic enzyme because it lacks a
recognisable N-terminal mitochondrial targeting motif. Clearly,
more experimental evidence is required to confirm the role of
each of these possible pathways in acetyl-CoA exchange and
lipid biosynthesis. Although the reasons of this possible redundancy are not clear yet, one may consider that, depending on
growth conditions and/or nature of the available carbon sources,
the parasite may need alternative acetyl-CoA exchange systems
to feed anabolic pathways.
5. Lactate production
Most trypanosomatids produce lactate from glucose,
although often as a minor end product [41] (Table 1 only shows
major end products). The insect forms of T. brucei and Leishmania spp. may excrete larger amounts of lactate [28,41,42],
however, they use different pathways to produce l-lactate and
d-lactate, respectively. The promastigote form of L. braziliensis
contains the glyoxalase system as a detoxification pathway to
protect the cell against damage by methylglyoxal [43], a mutagenic and cytotoxic compound that is mainly formed as a byproduct of glycolysis. In trypanosomatids, such as Leishmania,
this system is composed of two enzymes, glyoxalase I and glyoxalase II, which convert methylglyoxal into d-lactate using trypanothione as a cofactor [44,45]. Leishmania spp. express both
glyoxalases, whereas the T. brucei genome only contains the glyoxalase II gene, suggesting that the glyoxalase pathway is not
functional in T. brucei [44]. This is consistent with the absence of
d-lactate production by T. brucei gambiense procyclics, which
only excrete the l-lactate isomer [43]. It may be assumed that
l-lactate arises from pyruvate by the action of l-lactate dehydrogenase, however, the T. brucei genome does not contain this
gene and no l-lactate dehydrogenase activity is detectable in
T. brucei procyclics. Consequently, how l-lactate results from
glucose metabolism in T. brucei is still an open question. Whatever the l-lactate production pathway is in trypanosomes, we
propose that, in procyclic trypanosomes, l-lactate production is
an overflow pathway. Indeed, it appears that the relative amount
of lactate produced is correlated to the glycolytic flux, since
lactate excretion is considerably reduced in mutants showing a
reduced glucose consumption rate [28,29,46]. This hypothesis
may also explain why lactate is a minor end product in all other
trypanosomatids.
6. The ATP production dilemma
Until recently, it was widely accepted that procyclic trypanosomes (as well as most other trypanosomatids) grown in
rich medium primarily produce their ATP by oxidative phosphorylation [20]. Indeed the full enzymatic machinery for oxidative
metabolism is present in most adaptive forms. This includes
a functional cytochrome-containing respiratory chain capable
of proton gradient generation, as well as two separate terminal
oxidases (the cyanide-sensitive cytochrome oxidase – complex
IV and the salicylhydroxamic-sensitive alternative oxidase –
step 47) (Fig. 2). In this model, the proton gradient generated
by the respiratory chain (complexes I, III and IV) is used by
the F0 F1 -ATP synthase (step 48), considered to be the principal site of ATP generation. The essential role of the F0 F1 -ATP
synthase in energy production, under glucose-rich conditions,
has recently been questioned [12,33,46,47]. Indeed, an excess
of oligomycin, a specific inhibitor of the F0 F1 -ATP synthase,
does not affect the procyclic trypanosome viability and an unrealistically large excess is required to kill the cells in glucose
rich-medium [46]. In contrast, when grown in glucose-depleted
medium the same cells become over 1000-times more sensitive to oligomycin, supporting a view that, in the presence of
glucose, procyclic cells are not dependent on oxidative phosphorylation for ATP production. In the absence of glucose, when
cells switch to an amino acid catabolism, oxidative phosphorylation becomes essential [12]. These data also imply that, in
glucose-rich medium, most ATP is produced by substrate level
phosphorylation, with key roles for the cytosolic PGK (step 10),
PYK (step 13) and the mitochondrial SCS (step 28). Indeed,
considering the rate of glucose and/or l-proline consumption
and the number of ATP molecules produced per molecule consumed, we estimate that the net production of ATP by substrate
level phosphorylation is at least three-times higher in glucoserich medium than in glucose-depleted medium. This hypothesis
has been experimentally confirmed by production of PYK and
SCS mutant cell lines, which failed to grow in glucose-rich
medium [33,46], although the interpretation of this growth phenotype was recently reconsidered by Tielens and co-workers
[20,22]. They proposed that the SCS mutant lethality is a consequence of the overall alterations in glucose and/or l-proline
F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
metabolism, including the production of ATP by oxidative phosphorylation. However, this interpretation does not hold for the
PYK mutant, since glucose flux is not affected by the absence
of PYK activity, probably because of the malic enzyme bypass
(step 22), whereas ATP production is altered [46]. In conclusion,
all recent reports addressing experimentally the ATP production
in procyclic trypanosomes grown in glucose-rich medium provide evidence that the mitochondrial F0 F1 -ATP synthase plays
a minor role, whereas most ATP is synthesized by substrate
level phosphorylation [12,33,46]. How trypanosomes regulate
oxidative phosphorylation is still an open question. Eukaryotic
cells developed several approaches to control oxidative phosphorylation, including regulation by substrate availability such
as oxygen, ADP and reducing equivalents [48]. The latter may
play a role in trypanosomes, since l-proline catabolism produces approximately five-times more reducing equivalents than
glucose catabolism. The degree of coupling between respiration and oxidative phosphorylation can also be under control.
Two distinct regulatory processes have been described in eukaryotes: proton electrochemical potential-dissipating systems represented by uncoupling proteins and redox potential-dissipating
systems represented by alternative oxidases. Only the latter has
been described in trypanosomatids (step 47). Electron transfer
through the cytochrome-mediated pathway (complexes III and
IV) is coupled to ATP production, via the proton gradient generation. In contrast, electron flow from the ubiquinol pool to the
alternative oxidase pathway is not coupled to ATP production.
This dual system may provide enough flexibility to modulate the
proton gradient generation and thus be involved in the regulation
of oxidative phosphorylation.
7. Why do trypanosomatids use aerobic fermentation?
As mentioned above, trypanosomatids grown in rich medium
(which contains high concentrations of glucose and amino acids)
degrade glucose and amino acids into partially oxidized end
products, by so-called aerobic fermentation (fermentation in
the presence of oxygen). Obviously, aerobic fermentation is the
consequence of the absence of a “Pasteur effect” (inhibition of
glucose consumption in the presence of oxygen), however, the
rationale of this metabolic strategy developed by these parasites
is not clear. Numerous organisms usually employ fermentation
in the absence of oxygen, however, the insect-stage trypanosomatids are not adapted to long-term growth under anaerobic
conditions: Leishmania promastigotes have a poor capacity for
anaerobic functioning and go into reversible metabolic arrest
during anoxia [49], T. cruzi epimastigotes are also dependent
on respiration for proliferation and have a reduced glucose
metabolism during hypoxia [50], and T. brucei procyclics stop
dividing and start dying after few days in the absence of oxygen
([21] and our unpublished data). Clearly, aerobic fermentation
is neither a preadaptation to, nor a remnant of an anaerobic life
style that the parasite may need to develop in the insect vector.
It is also noteworthy that insects developed a complex network
of interconnected tubes (the tracheal system) that transport oxygen and other gases throughout the body, including the digestive
tractus [51]. Consequently, the insect forms of trypanosomatids
7
proliferate in aerobic conditions and may not need to be adapted
to long-term absence of oxygen.
Alternatively, aerobic fermentation may be a direct consequence of the capacity for a high glycolytic flux, as exemplified
by the slender bloodstream form of T. brucei. Indeed, the constant high glucose concentration found in the blood (5 mM)
is compatible with a glycolysis-based ATP production. Under
these conditions, oxidative metabolism is not required and this
adaptive form down-regulates expression of the TCA cycle
enzymes and the respiratory chain components, despite the presence of oxygen that remains even important for removing excess
reducing equivalents through the alternative oxidase. As discussed above, the insect-stage of T. brucei dwells in a l-prolinerich natural environment, but prefers glucose to l-proline when
grown in rich medium. We propose that the unrealistically large
excess of glucose in rich medium (6 mM) combined with the
lack of a “Pasteur effect” allows a relative high glycolytic flux,
sufficient for ATP generation by substrate level phosphorylation.
Consequently, procyclic trypanosomes probably need to downregulate the functional oxidative metabolism, including F0 F1 ATP synthase, respiratory chain and TCA cycle. This hypothesis
is consistent with the dramatic decrease of oligomycin sensitivity (over 1000-fold) observed in glucose-rich medium compared
to glucose-depleted medium, interpreted as a down-regulation of
oxidative phosphorylation. Down-regulation of l-proline consumption (six-fold) in glucose-rich medium strengthens this
notion, since l-proline catabolism may stimulate oxidative phosphorylation by generating five-times more reducing equivalents
than glucose catabolism (reducing equivalents produced from
glycolysis are primarily reoxidized by succinic fermentation).
This model based on T. brucei procyclic data, may be applicable to other insect stages, such as T. cruzi epimastigotes and
Crithidia choanomastigotes, which also prefer glucose to amino
acids (Table 1). The mechanism of this glucose-repression effect
is not known yet, and is under investigation for procyclic trypanosomes.
8. Concluding remarks
Since 2002, after the development of RNAi as powerful reverse genetic approach in T. brucei, the knowledge of
the energy metabolism in this parasite increased dramatically.
Another level of improvement was recently achieved by the
completion of the T. brucei, T. cruzi and L. major genome
projects. Indeed, the genome wide transcriptomics, proteomics
and metabolomics approaches will considerably speedup the
analysis of metabolic regulation processes. The combination
of these post-genomic and RNAi tools will be used to address
questions raised concerning energy metabolism, such as the
maintenance of the glycosomal ADP/ATP and redox balances,
the role and origin of l-lactate production, the production of glucoconjugates and nucleobase precursors (via gluconeogenesis)
in glucose-depleted medium, the mechanism of the glucoserepression exerted on oxidative metabolism and l-proline consumption, etc. Because of the RNAi technology, the procyclic
trypanosome is the ideal model to address these questions. Ultimately, this knowledge will constitute a framework to investigate
8
F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
equivalent pathways in the other human infective trypanosomes,
with the hope that an equivalent alternative to the “RNAi tool”
will be developed in T. cruzi and Leishmania spp.
Acknowledgements
We are grateful to Paul A.M. Michels and Derrick R. Robinson for critical reading of the manuscript. The authors are supported by the CNRS, the Conseil Régional d’Aquitaine and the
Ministère de l’Education Nationale de la Recherche et de la
Technologie.
References
[1] Berriman M, Ghedin E, Hertz-Fowler C, et al. The genome of the
African trypanosome Trypanosoma brucei. Science 2005;309:416–22.
[2] El-Sayed NM, Myler PJ, Bartholomeu DC, et al. The genome sequence
of Trypanosoma cruzi, etiologic agent of Chagas disease. Science
2005;309:409–15.
[3] Ivens AC, Peacock CS, Worthey EA, et al. The genome of the kinetoplastid parasite, Leishmania major. Science 2005;309:436–42.
[4] Cannata JJ, Cazzulo JJ. The aerobic fermentation of glucose by Trypanosoma cruzi. Comp Biochem Physiol B 1984;79:297–308.
[5] Fairlamb AH, Opperdoes FR. Carbohydrate metabolism in African trypanosomes, with special reference to the glycosome. In: Morgan MJ,
editor. Carbohydrate metabolism in cultured cells. Plenum Publishing
Corporation; 1986. p. 183–224.
[6] Bursell E. The role of proline in energy metabolism. In: Downer RGH,
editor. Energy metabolism in insects. New York: Plenum Press; 1981.
p. 135–54.
[7] Marr JJ, Berens RL. Regulation of aerobic fermentation in protozoans.
VI. Comparative biochemistry of pathogenic and nonpathogenic protozoans. Acta Trop 1977;34:143–55.
[8] Cazzulo JJ. Protein and amino acid catabolism in Trypanosoma cruzi.
Comp Biochem Physiol B 1984;79:309–20.
[9] Cazzulo JJ, Franke de Cazzulo BM, Engel JC, Cannata JJ. End products
and enzyme levels of aerobic glucose fermentation in trypanosomatids.
Mol Biochem Parasitol 1985;16:329–43.
[10] Cazzulo JJ. Intermediate metabolism in Trypanosoma cruzi. J Bioenerg
Biomembr 1994;26:157–65.
[11] North MJ, Lockwood BC. Amino acid and protein metabolism. In: Marr
J, Muller M, editors. Biochemistry and molecular biology of parasites.
Academic Press Ltd.; 1995. p. 67–88.
[12] Lamour N, Riviere L, Coustou V, Coombs GH, Barrett MP, Bringaud F.
Proline metabolism in procyclic Trypanosoma brucei is down-regulated
in the presence of glucose. J Biol Chem 2005;280:11902–10.
[13] Chamond N, Gregoire C, Coatnoan N, et al. Biochemical characterization of proline racemases from the human protozoan parasite Trypanosoma cruzi and definition of putative protein signatures. J Biol
Chem 2003;278:15484–94.
[14] Atwood 3rd JA, Weatherly DB, Minning TA, et al. The Trypanosoma
cruzi proteome. Science 2005;309:473–6.
[15] Chaumont F, Schanck AN, Blum JJ, Opperdoes FR. Aerobic and anaerobic glucose metabolism of Phytomonas sp. isolated from Euphorbia
characias. Mol Biochem Parasitol 1994;67:321–31.
[16] Hart DT, Coombs GH. Leishmania mexicana: energy metabolism of
amastigotes and promastigotes. Exp Parasitol 1982;54:397–409.
[17] Berman JD, Gallalee JV, Best JM, Hill T. Uptake, distribution, and
oxidation of fatty acids by Leishmania mexicana amastigotes. J Parasitol
1987;73:555–60.
[18] Ullu E, Tschudi C, Chakraborty T. RNA interference in protozoan parasites. Cell Microbiol 2004;6:509–19.
[19] Hannaert V, Bringaud F, Opperdoes FR, Michels PA. Evolution of energy
metabolism and its compartmentation in Kinetoplastida. Kinetoplastid
Biol Dis 2003;2:1–30.
[20] Van Hellemond JJ, Bakker BM, Tielens AG. Energy metabolism and
its compartmentation in Trypanosoma brucei. Adv Microb Physiol
2005;50:199–226.
[21] Van Weelden SW, Fast B, Vogt A, et al. Procyclic Trypanosoma brucei
do not use Krebs cycle activity for energy generation. J Biol Chem
2003;278:12854–63.
[22] Van Weelden SW, Van Hellemond JJ, Opperdoes FR, Tielens AG. New
functions for parts of the Krebs cycle in procyclic Trypanosoma brucei,
a cycle not operating as a cycle. J Biol Chem 2005;280:12451–60.
[23] Furuya T, Kessler P, Jardim A, Schnaufer A, Crudder C, Parsons M.
Glucose is toxic to glycosome-deficient trypanosomes. Proc Natl Acad
Sci USA 2002;99:14177–82.
[24] Morris JC, Wang Z, Drew ME, Englund PT. Glycolysis modulates
trypanosome glycoprotein expression as revealed by an RNAi library.
EMBO J 2002;21:4429–38.
[25] Cross GA, Klein RA, Linstead DJ. Utilization of amino acids by Trypanosoma brucei in culture: l-threonine as a precursor for acetate.
Parasitology 1975;71:311–26.
[26] Turrens J. More differences in energy metabolism between Trypanosomatidae. Parasitol Today 1999;15:346–8.
[27] Tielens AG, Van Hellemond JJ. Reply. Parasitol Today 1999;15:347–8.
[28] Besteiro S, Biran M, Biteau N, et al. Succinate secreted by Trypanosoma
brucei is produced by a novel and unique glycosomal enzyme. NADHdependent fumarate reductase. J Biol Chem 2002;277:38001–12.
[29] Coustou V, Besteiro S, Riviere L, et al. A mitochondrial NADHdependent fumarate reductase involved in the production of succinate excreted by procyclic Trypanosoma brucei. J Biol Chem
2005;280:16559–70.
[30] Colasante C, Ellis M, Ruppert T, Voncken F. Comparative proteomics
of glycosomes from bloodstream form and procyclic culture form Trypanosoma brucei brucei. Proteomics, in press.
[31] Annoura T, Nara T, Makiuchi T, Hashimoto T, Aoki T. The origin
of dihydroorotate dehydrogenase genes of kinetoplastids, with special
reference to their biological significance and adaptation to anaerobic,
parasitic conditions. J Mol Evol 2005;60:113–27.
[32] Takashima E, Inaoka DK, Osanai A, et al. Characterization of the
dihydroorotate dehydrogenase as a soluble fumarate reductase in Trypanosoma cruzi. Mol Biochem Parasitol 2002;122:189–200.
[33] Bochud-Allemann N, Schneider A. Mitochondrial substrate level phosphorylation is essential for growth of procyclic Trypanosoma brucei. J
Biol Chem 2002;277:32849–54.
[34] Gilbert RJ, Klein RA. Pyruvate kinase: a carnitine-regulated site of ATP
production in Trypanosoma brucei brucei. Comp Biochem Physiol B
1984;78:595–9.
[35] Van Hellemond JJ, Opperdoes FR, Tielens AG. Trypanosomatidae produce acetate via a mitochondrial acetate: succinate CoA transferase. Proc
Natl Acad Sci USA 1998;95:3036–41.
[36] Rivière L, Van Weelden SW, Glass P, et al. Acetyl: succinate CoAtransferase in procyclic Trypanosoma brucei. Gene identification and
role in carbohydrate metabolism. J Biol Chem 2004;279:45337–46.
[37] Takasaki K, Shoun H, Yamaguchi M, et al. Fungal ammonia fermentation, a novel metabolic mechanism that couples the dissimilatory and
assimilatory pathways of both nitrate and ethanol. Role of acetyl CoA
synthetase in anaerobic ATP synthesis. J Biol Chem 2004;279:12414–
20.
[38] Schmalix W, Bandlow W. The ethanol-inducible YAT1 gene from yeast
encodes a presumptive mitochondrial outer carnitine acetyltransferase. J
Biol Chem 1993;268:27428–39.
[39] Hiesinger M, Wagner C, Schuller HJ. The acetyl-CoA synthetase gene
ACS2 of the yeast Saccharomyces cerevisiae is coregulated with structural genes of fatty acid biosynthesis by the transcriptional activators
Ino2p and Ino4p. FEBS Lett 1997;415:16–20.
[40] Klein RA, Angus JM, Waterhouse AE. Carnitine in Trypanosoma brucei
brucei. Mol Biochem Parasitol 1982;6:93–110.
[41] Cazzulo JJ. Aerobic fermentation of glucose by trypanosomatids. FASEB
J 1992;6:3153–61.
[42] Darling TN, Balber AE, Blum JJ. A comparative study of d-lactate,
l-lactate and glycerol formation by four species of Leishmania and by
F. Bringaud et al. / Molecular & Biochemical Parasitology 149 (2006) 1–9
[43]
[44]
[45]
[46]
[47]
[48]
[49]
Trypanosoma lewisi and Trypanosoma brucei gambiense. Mol Biochem
Parasitol 1988;30:253–7.
Darling TN, Blum JJ. d-Lactate production by Leishmania braziliensis through the glyoxalase pathway. Mol Biochem Parasitol 1988;28:
121–7.
Vickers TJ, Greig N, Fairlamb AH. A trypanothione-dependent glyoxalase I with a prokaryotic ancestry in Leishmania major. Proc Natl Acad
Sci USA 2004;101:13186–91.
Irsch T, Krauth-Siegel RL. Glyoxalase II of African trypanosomes is
trypanothione-dependent. J Biol Chem 2004;279:22209–17.
Coustou V, Besteiro S, Biran M, et al. ATP generation in the Trypanosoma brucei procyclic form: cytosolic substrate level phosphorylation is essential, but not oxidative phosphorylation. J Biol Chem
2003;278:49625–35.
Besteiro S, Barrett MP, Riviere L, Bringaud F. Energy generation in
insect stages of Trypanosoma brucei: metabolism in flux. Trends Parasitol 2005;21:185–91.
Brown GC. Control of respiration and ATP synthesis in mammalian
mitochondria and cells. Biochem J 1992;284:1–13.
Van Hellemond JJ, Tielens AG. Inhibition of the respiratory chain
results in a reversible metabolic arrest in Leishmania promastigotes.
Mol Biochem Parasitol 1997;85:135–8.
9
[50] Stoppani AO, Docampo R, de Boiso JF, Frasch AC. Effect of inhibitors
of electron transport and oxidative phosphorylation on Trypanosoma
cruzi respiration and growth. Mol Biochem Parasitol 1980;2:3–21.
[51] Manning G, Krasnow MA. Development of the Drosophila tracheal
system. In: Martinez-Arias A, Bate M, editors. The development of
Drosophila. Cold Spring Harbor, New York: Cold Spring Harbor Press;
1993. p. 609–85.
[52] El-Sayed NM, Myler PJ, Blandin G, et al. Comparative genomics of
trypanosomatid parasitic protozoa. Science 2005;309:404–9.
[53] Sanchez-Moreno M, Fernandez-Becerra MC, Castilla-Calvente JJ, Osuna
A. Metabolic studies by 1H NMR of different forms of Trypanosoma cruzi as obtained by ’in vitro’ culture. FEMS Microbiol Lett
1995;133:119–25.
[54] Rainey PM, MacKenzie NE. A carbon-13 nuclear magnetic resonance
analysis of the products of glucose metabolism in Leishmania pifanoi
amastigotes and promastigotes. Mol Biochem Parasitol 1991;45:307–15.
[55] Darling TN, Davis DG, London RE, Blum JJ. Products of Leishmania
braziliensis glucose catabolism: release of d-lactate and, under anaerobic
conditions, glycerol. Proc Natl Acad Sci USA 1987;84:7129–33.
[56] Gilroy FV, Edwards MR, Norton RS, O’Sullivan WJ. Metabolic studies
of the protozoan parasite, Crithidia luciliae, using proton nuclear magnetic resonance spectroscopy. Mol Biochem Parasitol 1988;31:107–15.