FEMS Microbiology Reviews 29 (2005) 987–1020
www.fems-microbiology.org
Purine and pyrimidine transport in pathogenic protozoa:
From biology to therapy
Harry P. de Koning *, Daniel J. Bridges, Richard J.S. Burchmore
Institute of Biomedical and Life Sciences, Division of Infection and Immunity, Joseph Black Building, University of Glasgow, Glasgow G12 8QQ, UK
Received 21 October 2004; received in revised form 22 March 2005; accepted 24 March 2005
First published online 1 July 2005
Abstract
Purine salvage is an essential function for all obligate parasitic protozoa studied to date and most are also capable of efficient
uptake of preformed pyrimidines. Much progress has been made in the identification and characterisation of protozoan purine
and pyrimidine transporters. While the genes encoding protozoan or metazoan pyrimidine transporters have yet to be identified,
numerous purine transporters have now been cloned. All protozoan purine transporter-encoding genes characterised to date have
been of the Equilibrative Nucleoside Transporter family conserved in a great variety of eukaryote organisms. However, these protozoan transporters have been shown to be sufficiently different from mammalian transporters to mediate selective uptake of therapeutic agents. Recent studies are increasingly addressing the structure and substrate recognition mechanisms of these vital
transport proteins.
Ó 2005 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
Keywords: Nucleoside transporter; Nucleobase transporter; Chemotherapy; Pathogenic protozoa; Transporter structure; Purine salvage
Contents
1.
2.
3.
*
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Purine transport activities in various protozoan species. . . . . . . . . . . . . . . . . . . . . . .
2.1. Trypanosoma spp. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.2. Leishmania spp. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.3. Crithidia spp. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.4. Plasmodium spp. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.5. Toxoplasma gondii. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.6. Other protozoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Molecular biology of protozoan nucleoside and nucleobase transporters . . . . . . . . . .
3.1. Trypanosoma brucei. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.2. Leishmania spp. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.3. Plasmodium falciparum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.4. Toxoplasma gondii. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.5. Crithidia fasciculata. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.6. Subcellular transporters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
3.7. General comments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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Corresponding author. Tel./fax: +44 141 330 3753.
E-mail address: [email protected] (H.P. de Koning).
0168-6445/$22.00 Ó 2005 Federation of European Microbiological Societies. Published by Elsevier B.V. All rights reserved.
doi:10.1016/j.femsre.2005.03.004
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988
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4.
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6.
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
Protozoan nucleoside and nucleobase transporters in chemotherapy. . . . . . . . . . . . . . . . . . . . .
4.1. Transport or diffusion? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.2. The TbAT1/P2 transporter of T. brucei. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.3. Nucleobase transporters of Leishmania and Trypanosoma spp. . . . . . . . . . . . . . . . . . . . .
4.4. Nucleoside antimetabolites and transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.5. Use of protective nucleoside transporter inhibitors. . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.6. Chiral nucleosides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
4.7. Possible resistance mechanisms. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Structure–activity relationships of purine transporters. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.1. Topology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.2. Domains involved in substrate recognition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.3. A preliminary model for helix packing of TbAT1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
5.4. Models for transporter–permeant interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Concluding remarks. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Appendix A. Supplementary data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction
Protozoan purine transporters have attracted much
interest in recent years. As a result, purine transporters
have been identified in many of the most important
pathogenic protozoa. Until very recently, all this work
was performed on intact parasites, by measuring transport of radiolabelled purines and pyrimidines. Possibly
the first such study was by Tracy and Sherman, who reported the uptake of radiolabelled purines by the avian
malaria parasite Plasmodium lophurae in 1972 [1],
though other researchers had already started to study
protozoan purine and pyrimidine metabolism (e.g. [2])
The studies on purine metabolism revealed that, unlike
most mammalian cells, protozoan parasites are unable
to synthesise the purine ring de novo and rely solely
on salvage mechanisms for these essential nutrients
[3,4]. This dependence potentially makes protozoa vulnerable to inhibitors of the purine salvage pathways
[5,6]. In contrast, parasitic protozoa are fully capable
of synthesising the pyrimidine ring de novo, with the
exception of Giardia lamblia, Tritrichomonas foetus
and Trichomonas vaginalis [4,7], yet are also capable of
salvaging pyrimidines such as thymidine or uracil (e.g.
[8–11]).
Using biochemical techniques and live parasites,
numerous purine and pyrimidine transport activities
have been identified in a variety of protozoan species
[12,13]. One important early observation was that
many protozoan species, including Giardia intestinalis
[14,15], Leishmania donovani [9,16], Crithidia luciliae
[17], and Trypanosoma brucei [18] possess at least
two nucleoside transport activities in some of their life
cycle stages, identified by their kinetic parameters and
selectivity profile. In addition, they may express one or
more nucleobase transporters [8,19–24]. The overlap in
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1004
1004
1005
1006
1008
1008
1009
1009
1010
1010
1010
1012
1012
1013
1013
1013
1014
specificity between various purine and/or pyrimidine
transporters often makes it hard to study them
in situ, as specific inhibitors or permeants are often
lacking.
However, starting in 1998, the cloning and heterologous expression of a number of protozoan nucleoside
transporters from L. donovani [25,26], T. brucei [27,28],
Toxoplasma gondii [29] and Plasmodium falciparum
[30,31] were reported in quick succession. These breakthroughs allowed the characterisation of protozoan
nucleoside transporters in isolation for the first time,
and generally confirmed the earlier experiments with intact parasites. Heterologous expression in a specially selected and well-characterised system also greatly
facilitates the study of transporters that are otherwise
not easily available for biochemical analysis, due to,
for example, expression in life cycle forms that cannot
be obtained in sufficient numbers or expression at an
intracellular location. Yet, it needs to be stressed that
in situ and heterologous approaches are complementary,
as the observation of nucleoside transport in, for instance, oocytes of Xenopus laevis or a yeast cell, does
not necessarily yield the same kinetic parameters as in
the original cell, nor does it reveal much in itself about
the physiological role of the transporter in the parasiteÕs
biochemistry and physiology. Nonetheless, several specific nucleoside transporter genes have now been unambiguously coupled to a particular transport activity in
the parasite [25–29]. The genes encoding nucleobase
transporters, on the other hand, have proved more elusive, with the first cloning of genuine nucleobase transporter genes reported only in 2003 [32,33]. These first
nucleobase carriers were cloned from T. brucei brucei
and turned out to be of the Equilibrative Nucleoside
Transporter family (ENT), as are all the protozoan
nucleoside transporters cloned to date. It can be ex-
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
pected that, with the completion of more protozoan genomes, the current momentum of discovery will further
increase, and the cloning of a first nucleobase transporter from Leishmania was reported in early 2004
[21]. Even so, it is not yet clear whether all protozoan
nucleobase and nucleoside transporters are encoded by
ENT family genes.
Beyond the undoubted importance of protozoan
nucleoside/nucleobase transporters for the salvage of
essential nutrients, these proteins can also mediate the
uptake of chemotherapeutic agents by the parasite. This
can add a layer of selectivity over and above any selectivity on the level of the drug target. Indeed, in some
cases selective uptake may be the only or main level of
selective action, as in the case of melaminophenyl arsenicals that are efficiently accumulated through the T. b.
brucei TbAT1/P2 transporter [18,34]. In this case, the
melamine ring provides the haptophore, which can be
coupled to various toxophores to selectively introduce
cytotoxic agents into the trypanosome [35–38]. More
generally, purine transporters can be used to selectively
mediate the uptake of purine or pyrimidine antimetabolites [11,39,40]. Such antimetabolites are used widely as
antiviral and anticancer drugs, but limited effort has
been made to develop similar strategies for antiprotozoal agents. Even though various nucleoside and nucleobase analogues display good activity in vitro and/or
in vivo (e.g. [41–45]) only the pyrimidine antifolate pyrimethamine is widely used clinically, particularly in combination with sulphadoxine against malaria [46], and
also against toxoplasmosis [47,48]. In addition the purine nucleobase allopurinol is used in combinations
against leishmaniasis [49,50].
As both the efficacy and the selectivity of nucleoside
antimetabolites may be dependent on the mechanisms
of uptake by the parasite, a rational approach to drug
design must address the structural determinants that allow efficient transport. Ideally, antimetabolites would
be taken up by more than one transporter, as drug
resistance in protozoa is often associated with the loss
of a single transport activity [51,52]. An intimate
knowledge of the numbers of transporter (sub)-types,
expression levels in the relevant life cycle stages and
substrate specificity should therefore be considered as
part of drug development strategies. Techniques to
model the interactions between substrate and transporter binding site that determine affinity and selectivity have been developed and shown to have predictive
value [39,53]. They further highlight strong functional
conservation between transporters in different protozoan species, which cannot be readily determined from
primary sequence data [8,22]. The following review will
catalogue the protozoan purine and pyrimidine transport activities identified to date and categorise those
transporter genes that have been functionally confirmed or that are apparent in current genome dat-
989
abases. The uses and limitations of quantitative
structure-activity (QSAR) modelling of the interactions
between these proteins and their substrates, in relation
to protozoan drug uptake and resistance, will be
discussed.
2. Purine transport activities in various protozoan species
2.1. Trypanosoma spp.
Nucleoside transport has been extensively studied in
the trypanosomatidae, particularly in T. b. brucei and
L. donovani, but to a much lesser extent also in Trypanosoma evansi [54–56], Trypanosoma equiperdum [57],
Trypanosoma congolense [58], Trypanosoma vivax [59]
and Trypanosoma cruzi [10].
An early report by Sanchez et al. in 1976 [60] showed
the utilisation of [3H]adenosine nucleotides, particularly
AMP, by T. b. brucei and T. congolense. However, the
incubations over 15 min did not convincingly exclude
the possibility that the nucleotides were being dephosphorylated prior to transport by the trypanosomal 5 0 nucleotidase. In addition, nucleoside transporters are
not commonly believed to transport nucleotides. However, it should be noted that an L. donovani nucleoside
transporter, NT1.1, appears capable of AMP transport,
generating substrate-dependent currents when expressed
in Xenopus oocytes with an apparent Km of 9.1 ± 3.2 lM
[61], though, as before, it is difficult to exclude the possibility of a contribution from a putative endonucleotidase on the surface of the oocyte.
James and Born [58] first demonstrated the uptake of
radiolabelled adenosine, inosine, guanosine, hypoxanthine and adenine in T. b. brucei and T. congolense.
Though they measured uptake over relatively short
times and showed that the rate of uptake of adenosine
was greater than for the other purines, the conclusions
from their inhibition experiments were flawed as they
did not measure initial rates of transport. Their conclusion of a low-affinity adenosine transporter in addition
to a high affinity carrier remains to be confirmed,
though recent unpublished observations do indicate
the presence of such a transporter in T. b. brucei and
T. equiperdum, which is sensitive to adenine and insensitive to hypoxanthine (M.P. Barrett and M.L. Stewart,
personal communication).
2.1.1. Characterisation of the P1/P2 system
Interest in protozoan nucleoside transporters increased dramatically after the seminal observation in
1993 by Carter and Fairlamb [18] that a strain of T. b.
brucei adapted to melaminophenyl arsenicals (one of
the main classes of clinical trypanocides [62]) Ôlacks an
unusual adenosine transporterÕ. They identified two
NEa
NEa
1300
82
NEa
NEb
NEa
>500
NEa
NEb
NEa
60
NEa
NEb
95
0.46
5.0
4.0
194
HA
2.5
7.2
8.8
28.8
28.5
5
1.8
0.36
5.6
12.4
2.6
NEe
3.6
3.2
8.8
8.0
2.6
NEa
NEa
>1000
NEa
NEb
NEa
NEa
NEa
NEc
NEb
NEa
NEa
>500
NEc
NEb
48
>1000
10.9
NEa
>400
4.7
NEa
9.3
0.12
4.7
3.1
0.55
NEa
CYT
NEd [65]
NEb [53]
URA
HPP
>500
260 [53]
>250 [53]
110 [53]
XAN
GUA
NEd [65]
NEc [53]
0.38 [18]
ADE
HYP
NEd [65]
NEd [53]
THD
44 [53]
NEb [53]
CTD
NEc [53]
NEb [53]
GUO
URD
1080 [8]
NEc [53]
0.94 [65]
NEc [53]
0.36 [65]
NEc [18]
2.2
4.3
1.8
>1000
167
NEa
NEb
20
NEa
Values in bold type represent Km values; all others are Ki values. When several values are available from the literature, as a rule those obtained using intact T. brucei rather than heterologous
expression are given, and from the first report, if multiple exist. Numbers in table are references. ADO, adenosine; INO, inosine; GUO, guanosine; URD, uridine; CTD, cytidine; THD, thymidine;
HYP, hypoxanthine; ADE, adenine; GUA, guanine; XAN, xanthine; HPP, allopurinol; URA, uracil; CYT, cytosine; THY, thymine.
NE, no effect on permeant uptake at a concentration of a, 1 mM; b, 400 lM; c, 250 lM; d, 100 lM; e, 25 lM. HA, high affinity.
2.1.3. Hypoxanthine transporters
Very similar approaches established also that T. b.
brucei nucleobase transporters in both the procyclic (insect-borne) and long-slender bloodstream forms are also
proton symporters [19,20]. To date four hypoxanthine
INO
in which R is the gas constant, F is FaradayÕs constant,
T is the absolute temperature and pHo is the extracellular pH. A linear correlation between PMF and the rate
of adenosine uptake was established (r2 = 0.93 over 11
data points) [65]. Furthermore, an adenosine-induced
cytosolic acidification was demonstrated after ÔbaseloadingÕ with NH4Cl and the electrogenic nature of
adenosine and 2-Cl-adenosine transport was demonstrated by measuring their effect on plasma membrane
potential after inhibition of the proton pump [65].
0.15 [18]
0.59 [18]
2
1.4
0.3
348
590
NEa
NEb
860
NEa
ð1Þ
ADO
PMF ¼ V m ð2.3RT =F ÞðpHi pHo Þ;
Table 1
Kinetic parameters of T. b. brucei purine and pyrimidine transporters
2.1.2. Evidence for proton symport
P1 was also the first protozoan nucleoside carrier
shown to be a proton symporter [65]. While this study
was performed before the cloning of the P1 transporter
and was therefore unable to directly demonstrate and
characterise a proton flux by electrophysiology, strong
evidence for proton coupling was presented. The rate
of [3H]adenosine uptake was pH-dependent, and inhibited by ionophores such as CCCP, nigericin and gramicidin as well as inhibitors of the T. b. brucei plasma
membrane P-type H+-ATPase. Dose-dependent effects
of all these conditions were also measured on the plasma
membrane potential (Vm) and the intracellular pH (pHi),
from which the proton-motive force (PMF) was calculated using Eq. (1):
THY
high-affinity adenosine transporters, designated P1 and
P2, which could be specifically inhibited by inosine
and adenine, respectively [18]. Their experiments clearly
implicated P2 in the transport of melaminophenyl arsenicals [18], through interaction with the melamine pharmacophore [53,63], and further demonstrated that this
transporter could also mediate the uptake of pentamidine [64], another widely used trypanocide. The involvement of this transporter in drug uptake will be discussed
in detail in Section 4.2. The P1/P2 system has since been
studied extensively. P1 is a broad specificity purine
nucleoside transporter [18,27,53,65] (see Table 1) with
low affinity for uridine [8]. It has very low affinity for
purine nucleobases such as adenine and hypoxanthine
[27,65]. In contrast, P2 has no measurable affinity for
oxopurines (guanosine, inosine, hypoxanthine, guanine
and allopurinol), but displays submicromolar affinity
for aminopurines, notably adenine and adenosine
(Table 1) [18,53]. The P1/P2 system has since also been
characterised in T. equiperdum [57] and T. evansi [54–
56,66] and displayed very similar characteristics as in
T. b. brucei.
NEb [53]
NEb [53]
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
P1/NT2
P2/AT1
NT5 [67]
NT6 [67]
NT7 [67]
H1 [19]
H2 [20]
H3 [20]
NT8.1 [33]
H4/NBT1 [32]
U1 [11]
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
transporters have been characterised: H1 and H4 in procyclics [19,32] and H2 and H3 in bloodstream forms
[20]. In addition, a high-affinity uracil transporter
[11,40] and a cytosine transporter (De Koning, unpublished observation) have been identified in procyclics.
The hypoxanthine carriers are all broad specificity purine nucleobase transporters with, as a rule, little or no
affinity for pyrimidines or nucleosides (Table 1). Their
Km value for hypoxanthine varies from 123 nM to
9.3 lM and they are clearly distinguishable on this basis
and on their inhibition profile. For instance, H2, but not
H3 or H1, is inhibited by uracil and guanosine [19,20].
H4 is inhibited by both, with similar Ki values as H2
[32], but also displays equally high affinity for guanine,
3-deazaguanine and 7-deazaguanine, whereas H2 displays much lower affinity for these analogues than for
the natural nucleobase [39].
2.1.4. Regulation of purine transport activity
It therefore appears that T. b. brucei expresses at least
three different purine transporters in its procyclic stage
(P1, H1 and H4) and at least four distinct purine transport activities in its long-slender bloodstream stage (P1,
P2, H2, H3), but the situation in other developmental
stages is far less clear. Sanchez et al. [67] identified several additional adenosine transporters in a locus of 6
closely related genes that includes the NT2 gene that encodes P1 [27] (see Section 3.1). mRNA for all six genes
was detected in long-slender bloodstream forms [67].
As trypanosomes can convert any of the natural purine
nucleosides or nucleobases (excluding uric acid) to any
other [7], and thus develop normally on any single purine source, it is not immediately clear why they should
require this many purine transporters.
A partial explanation for this plethora of purine
transport activities may lie in the adaptability of the purine salvage system. First studied in Crithidia spp. (see
Section 2.3), the purine salvage enzymes and transporters of the trypanosomatids can be dramatically up- or
downregulated according to growth stage and availability of a purine source [24,68–74]. In T. b. brucei, adenosine and hypoxanthine transporters in procyclics appear
to be differentially controlled under these conditions:
hypoxanthine transport rates increased to 450% of control within 24 h of purine stress, whereas adenosine
transport capacity was only increased after 48 h [68]. Indeed, of two hypoxanthine transport activities present in
these cells only the higher affinity activity, at the time
presumed to be H2 but now thought to be H4, was
upregulated [68], showing an appropriate physiological
response to low purine levels. Under purine-replete conditions, the high capacity, lower affinity H1 transporter
is dominant [19,68]. A further factor in explaining the
high number of purine transporter activities and genes
may be in differential expression in the various stages
of the Trypanosoma lifecycle. Some differences between
991
procyclic and long-slender bloodstream forms were
mentioned above. In addition, at least one of the purine
transporters is expressed only in short-stumpy bloodstream forms [75].
2.2. Leishmania spp.
Nucleoside transport in Leishmania species, particularly L. donovani, responsible for the most severe form
of human leishmaniasis, has been as thoroughly studied
as the equivalent transporters in T. b. brucei. The L.
donovani nucleoside transporters were also the first protozoan purine or pyrimidine transporters cloned (see
Section 3.2). However, studies on nucleobase transporter activities have hardly been reported until 2003
and almost all studies so far have dealt with the promastigote (insect) stage rather than the far more difficult to
study amastigotes, which reside in phagolysosomes
within mammalian macrophages.
2.2.1. Purine salvage
Like other parasitic protozoa, Leishmania were found
to be auxotrophic for purines, with adenosine and hypoxanthine being preferred for optimal growth. While adenine and inosine were being salvaged as well, they were
first converted to hypoxanthine before incorporation
into the nucleotide pool [76]. In 1982, Hansen et al.
[77] published a study of the transport of various radiolabelled purines in L. braziliensis panamensis and postulated three different Ôtransport lociÕ: for inosine, for
hypoxanthine/adenine and for adenosine. This is still
the current model for purine transport by promastigotes. In addition, promastigotes, like procyclic T. b. brucei [78], are able to transport S-adenosylmethionine
(AdoMet) [79], a metabolite used in polyamine synthesis
and methylation reactions that can also be converted to
adenosine and adenine [79].
2.2.2. Identification of the L. donovani nucleoside
transporters
Ullman and colleagues [80] created cell lines of L.
donovani promastigotes resistant to either of the nucleoside antimetabolites tubercidin and Formycin B (analogues of adenosine and inosine, respectively). The
resistance phenotype was associated with the loss of
either inosine or adenosine and pyrimidine nucleoside
uptake over 60 min. Uptake of the nucleobases hypoxanthine, adenine and uracil was not affected. This
model was still consistent with the three loci model of
Hansen and, using the mutant cell lines both nucleoside
transporters were characterised with respect to kinetic
parameters [16] and eventually cloned using functional
complementation of the deficient phenotypes [25,26].
Two closely related genes were identified encoding adenosine/pyrimidine transporters, LdNT1.1 and LdNT1.2,
with only LdNT1.1 expressed in promastigotes [26].
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
Both transporters displayed submicromolar affinity for
adenosine (Table 2) and somewhat lower affinity for uridine [26]. LdNT1.1 is also believed to mediate thymidine
and cytidine transport [9,16,80,81], whereas NT1.2 is
highly specific for the oxopurine nucleosides, inosine
and guanosine (Table 2), and is present as a single copy
in the genome [25]. The presence of two distinct adenosine transporter genes in the Leishmania genome, however, may explain the results of Ogbunude and
colleagues [81,82] who found significant variations in
adenosine transport activities, but not in inosine transport activity, between various strains of both L. donovani and Leishmania major. It would seem possible
that expression of NT1.2 does occur in some strains or
is induced during certain growth conditions. Under purine stress conditions, for instance, LdNT1-like activity,
but not LdNT2, is upregulated 10-fold [83]. Changes
in apparent adenosine affinity in adenosine-depleted
parasites, and different levels of upregulation for adenosine and uridine, suggest that perhaps a different transporter than LdNT1.1 is being expressed during purine
stress. A similar pattern occurs in T. b. brucei (see
Section 2.1.4).
Nucleoside transport in L. donovani amastigotes, as
in promastigotes, appears to be mediated by at least
two transporters, T1 and T2 [9] (Table 2). T1 was similar
to the LdNT1 activity, in that it is a high-affinity adenosine transporter and sensitive to inhibition by pyrimidine nucleosides but not by inosine or guanosine. The
T2 transport activity, unlike LdNT2, also transports
adenosine in addition to the oxopurine nucleosides [9].
It seems entirely possible that T1 is encoded by LdNT1.1
or LdNT1.2.
Table 2
Overview of Leishmania purine and pyrimidine transporters
2.2.3. Electrophysiology of nucleoside transport
The L. donovani nucleoside transporters expressed in
Xenopus oocytes have also been studied with electrophysiological techniques [61]. It was clearly shown that
transport by all three cloned nucleoside transporters is
electrogenic, in a proton-dependent, sodium-independent manner. However, nucleoside transport maybe
only partially dependent on the proton-motive force as
the study found no significant charge co-transport associated with adenosine uptake by LdNT1.1 and only a
very minor adenosine:proton ratio with LdNT1.2
(0.0035 charges per molecule of adenosine transported).
This ratio was higher when uridine was the substrate
(0.13 and 0.45 charges per molecule transported by
NT1.1 and NT1.2, respectively). This is in contrast to
a 1:1 ratio for the Leishmania proton/myo-inositol symporter [84]. However, Stein et al. [61] argue that the
charge:substrate translocation ratios may be underestimated by the presence of a constitutive proton leak,
blocked more efficiently by adenosine than by uridine.
In any event, it is clear that the L. donovani [61] and
T. b. brucei [19,20,65] nucleoside and nucleobase transporters can exploit the large proton-motive force across
their plasma membranes to transport purines and pyrimidines with high affinity and, if necessary, against a
concentration gradient.
The nucleoside transporters, and probably the nucleobase transporters, are members of the Equilibrative
Nucleoside Transporter family (see Section 3.2), which
facilitates the equilibrative exchange of substrate across
plasma membranes in vertebrates [85]. Concentrative
transport of nucleosides in mammals is dependent on
the sodium rather than the proton gradient and mediated by the CNT family [86], whereas concentrative
nucleobase transport is mediated by sodium-dependent
transporters of an as yet unidentified gene family [87].
Interestingly, the first nucleoside transporter cloned
from a plant, Arabidopsis thaliana, also appears to be
Substrate
Km (lM)
Expression
Refs.
Adenosine
Uridine
Thymidine
Cytidine
Tubercidin
Formycin A
0.17 ± 0.09
5.6 ± 1.8
PM
[26]
[26]
[9,16]
[9,16]
[9,80]
[9,80]
NT1.2
Adenosine
Uridine
0.66 ± 0.15
40 ± 11
PM?
[26]
[26]
NT2
Inosine
Guanosine
Formycin B
0.3 ± 0.1
1.7 ± 0.5
PM
[25]
[25]
[25]
T1
Adenosine
Uridine
Thymidine
Cytidine
Formycin A
1.14 ± 0.05
AM
[9]
[9]
[9]
[9]
[9]
T2
Adenosine
Inosine
Guanosine
Formycin A
Formycin B
2.09 ± 0.13
AM
[9]
[9]
[9]
[9]
[9]
Hypoxanthine
Adenine
Guanine
Xanthine
Allopurinol
0.71 ± 0.07
4.6 ± 0.9
2.8 ± 0.7a
23 ± 8a
54 ± 3
PM
[22]
[22]
[22]
[22]
[22]
NT3
Hypoxanthine
Adenine
Guanine
Xanthine
16.5 ± 1.5
8.5 ± 1.1
8.8 ± 4.0
8.5 ± 0.6
PM
[21]
[21]
[21]
[21]
LmU1
Uracil
5-Fluorouracil
0.32 ± 0.07
0.66 ± 0.14a
PM
[40]
[40]
Species/
Transporter
L. donovani
NT1.1
L. major
NBT1
PM, promastigotes; AM, amastigotes.
a
Ki value (lM).
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
a proton symporter of the ENT family [88] as are CNT
family members from Caenorhabditis elegans [89] and
Candida albicans [90].
2.2.4. Nucleobase transport in Leishmania
The first systematic study of purine nucleobase transport in Leishmania spp. after the initial report by Hansen [77] showed a single high-affinity transporter with
broad specificity for purine nucleobases in L. major
promastigotes, designated LmaNBT1 [22]. Highest affinity was for hypoxanthine, with a Km value of 0.71 lM,
followed by guanine, adenine and xanthine (Table 2).
Interestingly, the antileishmanial hypoxanthine analogue allopurinol was also taken up by LmaNBT1, with
a Km value of 54.3 lM. Strong kinetic evidence, based
on mutual inhibition profiles, was presented that
LmaNBT1 was the only transport activity for [3H]adenine, [3H]hypoxanthine and [3H]allopurinol in these cells
[22]. The LmaNBT1 transport activity was very similar
to the H2 activity in T. b. brucei [22] (see Section 5.4).
The TbU1 [11] and LmU1 uracil transporters [40] were
equally similar in their substrate selectivity profile, with
high affinity for uracil only (Tables 1 and 2).
Recently, the group of Landfear reported the cloning
and heterologous characterisation of an L. major transporter, LmaNT3, which displays high affinity for purine
nucleobases only [21]. Though the Km values obtained in
Xenopus oocytes are rather higher than reported for
LmaNBT1 [22], with a hypoxanthine Km of 16.1 lM
(Table 2), it seems possible that this gene encodes
LmaNBT1 or the equivalent transport activity in
amastigotes. This group also reports the cloning of a
further ENT member from L. major, LmaNT4, which
is currently being characterised [91].
2.3. Crithidia spp.
Crithidia are parasites of insects, and do not require
intermediate (mammalian) hosts as do other kinetoplastids discussed here. Due to the close evolutionary
relationship between these parasites and, in particular,
Trypanosoma and Leishmania spp., and the ease of their
culture in vitro, they have been used quite extensively as
model organisms for the kinetoplastidae.
Nucleoside and nucleobase transport in the two Crithidia species, Crithidia fasciculata and Cr. luciliae, is
very similar (Table 3). The first report of purine transport in this genus was in 1978 by Kidder et al. [69],
reporting both high- and low-affinity uptake of each of
the bases hypoxanthine, adenine and guanine (Table
3). They further claimed that their experiments showed
that each of the bases is transported by a separate carrier, though this seems unlikely as the uptake of each
base was shown to be inhibited by each of the others
[69]. Nucleobase uptake in Cr. luciliae has been studied
993
Table 3
Nucleoside and nucleobase transporters of Crithidia
Transporter
Substrate
Cr. fasciculata
NBT1
Adenine
Hypoxanthine
Guanine
NT1
NT2
Cr. luciliae
NBT1
NT1b
NT2b
Km (lM)
11.8a
3.5
8.0
Adenosine
Uridine
Thymidine
Cytidine
Tubercidin
9.4
Inosine
Guanosine
0.28
Adenine
Hypoxanthine
Guanine
Adenosine
Adenosine
2 0 Deoxyadenosine
Thymidine
Tubercidin
Inosine
Guanosine
2 0 deoxyGuo
Ki (lM)
Ref.
[69]
[68]
18
25
8.1
5.7
[68]
2.3
2
[24]
3.9
3
42
9.3
[17]
18
ND
ND
ND
[17]
10.6
ND
ND, not determined.
a
Only the high affinity values are given.
b
Not inhibited by cytidine and uridine [70].
in much greater detail and was shown to be similar to
purine transporters of the other kinetoplastidae: high
affinity and broad specificity for purine nucleobases
and low if any affinity for the corresponding nucleosides
and for pyrimidine nucleobases (Table 3) [24].
Nucleoside transport in these species is very similar to
the situation in L. donovani: an NT1 transporter with
high affinity for adenosine and pyrimidine nucleosides,
and NT2 with high affinity for inosine and guanosine
(Table 3). The one exception to this consensus seems
to be the reported inability of cytidine and uridine to inhibit either NT1 or NT2 of Cr. luciliae, but it was not
clear at which concentrations [70]. Overall, purine/
pyrimidine transport in Crithidia does appear to be very
similar to that of other kinetoplastids, particularly to
Leishmania promastigotes, which share a similar environment. Very recently, the group of Buddy Ullman reported the cloning (see Section 3.5) and heterologous
expression of two Cr. fasciculata nucleoside transporters, an NT1-type and the probable NT2 transporter [92].
Groundbreaking work has been performed, using Cr.
luciliae and Cr. fasciculata, on the regulation of purine
salvage in response to different environmental conditions, particularly purine availability [24,68,70,71,74].
This situation was later also reported for T. b. brucei
procyclics [68] and Leishmania promastigotes [83]. In
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
Crithidia, it has been shown that purine starvation leads
to a concerted response with upregulation of extracellular 3 0 -nucleotidase [70,72,93], purine nucleoside and
nucleobase transporters [70,71,74], hypoxanthine–guanine phosphoribosyltransferase [71] and adenine deaminase [71] – all proteins involved in the salvage of
purines. This response seems to be specific since transport of amino acids and glucose in Cr. luciliae was unaffected by purine-starvation [24], as was uracil transport
in T. b. brucei, while hypoxanthine and adenosine transport was strongly upregulated [68]. Conversely, depletion of different essential nutrients, arginine and
haemin, did not affect 3 0 -nucleotidase activity in Cr.
luciliae [93]. The upregulation of purine salvage enzymes
appears to require new protein synthesis, as the response
is sensitive to cycloheximide [24,68,70]. It would now appear that this is required for the de novo synthesis of
regulatory proteins rather than the purine salvage enzymes themselves, as Liu et al. [92] did not detect significantly increased levels of CfNT2 mRNA during purine
depletion, though NT2 activity was upregulated >10fold and regulation must therefore be post-transcriptional. The regulation of purine salvage in Crithidia
has been extensively reviewed elsewhere [73,74].
2.4. Plasmodium spp.
Plasmodium parasites, the etiological agents of malaria, are obligate intracellular parasites with a complicated lifecycle. During the course of infection,
sporozoites first infect liver cells, where they multiply
and re-emerge as metacryptozoites that infect erythrocytes. It is the cyclic development in the erythrocyte that
is ultimately responsible for the clinical manifestations
of malaria, and it is on the intraerythrocytic stages that
most research and drug development has focussed.
2.4.1. Purine and pyrimidine salvage by Plasmodium
It has long been known that Plasmodium species,
like other protozoan parasites, are incapable of synthesising purines [94–96], but do synthesise pyrimidines
de novo [95,97]. Unlike most other parasites they cannot rely on the host cell to provide these essential nutrients: erythrocytes equally lack purine and pyrimidine
biosynthetic pathways and, being anucleate, have a limited requirement for nucleotides [94]. Yet, the rapid
development and division of the parasite is accompanied by a dramatic increase in purine and pyrimidine
utilisation and it is has long been known that Plasmodium can utilise preformed purines for nucleic acid
synthesis [2,98]. In contrast, Plasmodium spp. lack the
ability to salvage preformed pyrimidines [98–100]
except orotate [98]. Consequently, inhibitors of pyrimidine biosynthesis such as pyrazofurin [101] and
the hydroxynaphtoquinone BW58C have strong anti-
Plasmodium effects [102]. Pyrazofurin is an inhibitor
of orotate phosphoribosyltransferase and orotidine-5 0 phosphate decarboxylase and BW58C inhibits dihydroorate dehydrogenase.
Whereas the non-infected erythrocyte salvages nucleosides through the equilibrative hENT1 transporter [103]
and purine nucleobases through the facilitative nucleobase transporter hFNT1 [39,104], the plasma membrane
of the infected erythrocyte contains an additional uptake
mechanism for nucleosides and many other nutrients,
which is sensitive to furosemide [105–107]. While the
nature of this unique New Permeation Pathway (NPP)
is still a matter of intense debate, it is clear that its role
is to import the nutrients into the erythrocyte cytosol
rather than into the Plasmodium parasite. The porous
nature of the membrane of the parasitophorous vacuole
(PVM) in which the parasite resides [108,109] makes it
likely that the parasite experiences the same purine
concentrations as in the erythrocyte cytoplasm. An additional mechanism, however, is required for the transport
of the nucleosides and/or nucleobases across the parasite
plasma membrane.
2.4.2. Central role for hypoxanthine uptake
To study purine transport by Plasmodium spp., parasites were released from the infected erythrocytes by
treatment with saponin [1,110,111]. The first such report, using the avian malaria parasite P. lophurae, described rapid and saturable uptake of 14C-labelled
adenosine, inosine, hypoxanthine and guanine [1]. The
apparent mutual inhibition of uptake of each of these
purines led to the assumption of a single purine nucleoside/nucleobase transporter in this species, though it is
noteworthy that purine uptake was not inhibited by adenine [1]. They also did not exclude a separate transporter
for oxopurines, as they reasoned that adenosine might
inhibit an inosine transporter after deamination. Similar
results were obtained by Manandhar and Van Dyke
using Plasmodium berghei, who reported that adenosine
is not only rapidly deaminated to inosine, but subsequently deribosylated to hypoxanthine prior to uptake
by the free parasite [111]. Hansen and colleagues [110]
further confirmed this model by showing inhibition of
[3H]adenosine uptake in saponin-freed P. berghei parasites with 2 0 -deoxycoformycin, a powerful adenosine
deaminase inhibitor.
The consensus of the early studies of purine salvage
in model malaria parasites was thus that purines are
overwhelmingly taken up as hypoxanthine by Plasmodium parasites, as represented in Fig. 1(a). This view
was entirely consistent with the observation that hypoxanthine is by far the preferred substrate for purine biosynthesis in Plasmodium [112]. Indeed, it was shown that
P. falciparum growth was up to 90% inhibited when xanthine oxidase was used to deplete infected erythrocytes
of hypoxanthine [113].
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
a Erythrocyte
cytosol
1
Ado
995
2
Ino
HX
?
Hypoxanthine
transporter
Plasmodium
cytosol
2
Ino
b
3
4
5
I MP
HX
AMP
ATP
AMP
ATP
6
AMP
Erythrocyte
cytosol
I MP
7
7
1
Ado
2
Ino
HX
Hypoxanthine
transporter
Pf(E)NT1
Plasmodium
cytosol
1
Ado
3
2
Ino
HX
4
I MP
5
Fig. 1. Purine salvage by intra-erythrocytic Plasmodium parasites. (a) Early model, showing predominant or sole uptake of hypoxanthine (based in
part on observations from Refs. [110–113]. (b) Current model (using information from Refs. [7,30,31,115], amoung others). HX, hypoxanthine. 1.
adenosine deaminase. 2. inosine phosphorylase. 3. hypoxanthine–guanine–xanthine–phosphoribosyltransferase. 4. adenylosuccinate synthetase. 5.
adenylosuccinate lyase. 6. AMP deaminase. 7. 5 0 nucleotidase.
2.4.3. P. falciparum adenosine transporters
More recently, efforts have begun to clone and study
purine transporters of P. falciparum, the agent responsible for the most serious forms of human malaria, by
expression in oocytes of X. laevis. mRNA isolated from
infected human erythrocytes and injected into oocytes
induced the expression of transport activities for adenosine and hypoxanthine. Mutual inhibition again appeared to indicate a single transporter for both purines
being expressed, but since Km and Ki values were not
determined, this assumption remained unproven [114].
A more thorough characterisation of a P. falciparum
nucleoside transporter became possible with the almost
simultaneous cloning of PfNT1 and PfENT by the
groups of Ullman [30] and Baldwin [31], respectively.
Despite both groups utilising the X. laevis expression
system and the two genes differing by just a single amino
acid (Phe or Leu at position 385), the conclusions of the
two groups differed markedly. For instance, PfENT1
reportedly has a 25-fold lower affinity for adenosine
compared with PfNT1. Furthermore, PfNT1 was reported to transport L-isomers of adenosine and thymidine whereas the P. falciparum adenosine transporter
had previously been reported to be stereoselective
[106,114]. Nor is there agreement about whether nucleobases are [31,114] or are not [30] substrates of the nucleoside transporter, or whether it is sensitive to inhibition
by dipyridamole. Clearly, additional studies with heterologously expressed transporters and with isolated parasites will be required to reconcile these studies. It
does appear certain, however, that Pf(E)NT is expressed
during the intraerythrocytic stages of the lifecycle
[30,31,115] and is present on the parasite plasma membrane rather than on the plasma membrane of the infected erythrocyte [115]. This is consistent with
Pf(E)NTÕs insensitivity to furosemide [30,114], which
blocks nucleoside uptake into infected erythrocytes
[106,107].
The presence of an adenosine transporter on the P.
falciparum plasma membrane changed the model proposed in Fig. 1(a). It is also now clear that adenosine
deaminase and inosine phosphorylase are present both
inside and outside the parasite [7] (Fig. 1(b)). Both studies of the P. falciparum adenosine transporter report
that intraerythrocytic stages of Plasmodium should be
able to take up at least some pyrimidines, notably
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
thymidine. The parasite may not, however, have the
metabolic pathways to utilise them. For instance, a
probable uracil phosphoribosyltransferase can readily
be identified in the T. brucei genome (Tb04.2H8.180;
www.genedb.org) but not in any Plasmodium genome
available to date (http://www.plasmodb.org/). The enzyme activities of the pyrimidine salvage pathway have
not been identified biochemically either [7,112,116].
The contribution of the adenosine transporter to purine salvage in Plasmodium species needs to be reassessed
in the light of the earlier studies indicating a more
important role for hypoxanthine transport and further
studies of purine transport by Plasmodium spp. are urgently required. Purine transport across the normal
and infected erythrocyte plasma membrane is discussed
in detail elsewhere [107] and will not be addressed here.
2.5. Toxoplasma gondii
2.5.1. Source of purines for salvage
As in the related Apicomplexan species, P. falciparum, the questions of purine salvage have largely centred
on what the preferred purine source is and how the
intracellular parasite, within the parasitophorous vacuole, could have access to sufficient levels of it. Although
there are many parallels between Plasmodium and Toxoplasma purine salvage, one major difference is the type
of host cell the parasite inhabits. While Plasmodium infects the metabolically incomplete, anucleate erythrocyte
(no purine biosynthesis), Toxoplasma invades nucleated
cells with the entire metabolic machinery intact. Unlike
the malaria parasite, Toxoplasma is not believed to induce the formation of additional permeation pathways
in the host cell and therefore relies on free purines in
the host cell cytoplasm, which diffuse freely through
pores in the parasitophorous vacuole membrane [117].
The purine transporters need to be able to salvage the
presumably very low concentrations of free nucleosides
and nucleobases, as the parasite is unable to take up
nucleotides directly [118]. Much speculation has centred
on whether efficient purine salvage includes hydrolysis
of host cell ATP to adenosine by a Toxoplasma-encoded
nucleoside triphosphate hydrolase (NTPase) present in
the parasitophorous vacuole [119–121]. The characterisation [122] and cloning [29] of a low affinity, high
capacity adenosine transporter, TgNT1, seemed to fit
well in this model of massive generation of adenosine
from host ATP. However, it was subsequently shown
that less than 5% of NTPase in the vacuole is active –
and that activation leads to rapid depletion of cellular
ATP and exit of the parasite from the host cell [123].
Furthermore, the NTPase hydrolyses ATP only to 5 0 AMP or ADP [119,124] and it was speculated that a
T. gondii ecto-5 0 -nucleotidase would complete the
hydrolysis to adenosine. In a comprehensive study,
Ngô et al. [118] were unable to demonstrate the exis-
tence of such an enzyme activity in the parasitophorous
vacuole. The conclusion of this study, that host cell
nucleotides are not the major source of purines for
T. gondii, and the earlier observation that the parasite
can incorporate hypoxanthine, adenine, guanine, xanthine, guanosine and inosine as well as adenosine
[125,126], triggered a reinvestigation of purine transport
by T. gondii and high affinity transporters, one for hypoxanthine and one with broad specificity for nucleosides,
have now been reported [8].
2.5.2. Characterisation of nucleoside and nucleobase
transporters
All purine and pyrimidine transport studies with Toxoplasma have been performed with isolated tachyzoites,
the replicating stage infective to nucleated mammalian
cells. The first comprehensive investigation of purine uptake in these cells was performed by Schwab et al. [122].
They reported a low affinity, equilibrative adenosine
transporter (apparent Km = 230 lM). These experiments
were performed in an adenosine kinase-deficient strain,
so as eliminate any influence of adenosine metabolism
on the apparent rates of transport. Slightly unusual
for a protozoan nucleoside transporter, adenosine transport was inhibited by the mammalian nucleoside transport inhibitor, dipyridamole, with an IC50 of 0.7 lM.
In contrast, dipyridamole inhibited transport of inosine,
hypoxanthine and adenine only marginally at 10 lM,
suggesting the presence of additional transporters. This
adenosine transporter, TgAT, was subsequently cloned
by insertional mutagenesis and selection on a cytotoxic
adenosine analogue, adenine arabinoside [29]. TgAT
was a member of the ENT family and mediated the uptake of adenosine when expressed in oocytes, with an
apparent Km value of 114 ± 37 lM, and was 60%
inhibited by 50-fold excess inosine and to a lesser extent
by guanosine, hypoxanthine and guanine [29]. While
adenosine transport in the insertional mutant was much
reduced, it retained salvage capabilities for inosine and
purine nucleobases and the authors concluded that
TgAT encodes the sole adenosine transporter of T. gondii. However, on studies with intact tachyzoites we
found that both adenosine and inosine are transported
by an additional high-affinity nucleoside transporter
(Km values of 0.49, 0.12 and 0.77 ± 0.20 lM, respectively) [8]. This transporter, TgAT2, also displayed high
affinity for guanosine, uridine and thymidine. In the
same study, we reported the characterisation of a highaffinity T. gondii hypoxanthine/guanine transporter,
TgNBT1 (Km (hypoxanthine) = 0.91 ± 0.19 lM). Adenine uptake was not saturable up to 1 mM and apparently depended on simple diffusion [8]. The presence of
TgAT2 contradicts the conclusion of Chiang et al. [29]
that TgAT must be the sole adenosine transporter in
T. gondii, though there are several possible explanations
for this apparent discrepancy. These include the possi-
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
bility that both transporters are the product of the same
gene (alternative splicing, post-translational modification, etc.) and the possibility that additional mutations
occurred during selection of the insertional mutant with
adenine arabinoside. Several additional ENT genes can
readily be identified in the T. gondii genomic database
(ToxoDB.org) and have been named TgNT1, TgNT2
and TgNT3 [127].
2.5.3. Localisation of T. gondii adenosine transporters
A recent study by the group of El Kouni [128] reports
that intracellular tachyzoites are able to transport b-Ladenosine, nitrobenzylthioinosine (NBMPR) as well as
other b-L-purine nucleosides. This ability was much reduced in tachyzoites of the DTgAT strain. Surprisingly,
Toxoplasma-infected human fibroblasts similarly acquired the ability to transport these non-natural nucleosides, leading to the conclusion that TgAT must be
present both on the parasite plasma membrane and
the host cell plasma membrane. This would be in contrast to the situation in Plasmodium-infected erythrocytes, where PfNT1 localises exclusively to the parasite
plasma membrane and could not be demonstrated with
immunoelectron microscopy on the host cell membrane
[115].
2.5.4. Pyrimidine salvage in T. gondii
Toxoplasma tachyzoites are capable of de novo
pyrimidine biosynthesis and this seems a vital pathway
for virulence [129]. Disruption of this pathway by deletion of the gene encoding carbamoyl phosphate synthetase II created a uracil auxotrophic cell line, which was
able to invade host cells but needed supplementation of
>20 lM uracil in order to replicate. This study shows
that T. gondii express a uracil transporter, but do not
ordinarily depend on it. In addition, TgAT2 appears
to transport pyrimidine nucleosides, as well purine
nucleosides [8].
997
later paper [130] that the thymidine carrier does not
actually transport thymine, despite the relatively high
affinity, since 50 lM [3H]thymine uptake was inhibited
by only 12 ± 3% by 2 mM thymidine. However, since
the Vmax of thymine on the nucleobase transporter
(ÔType 3Õ) was very much higher than the [3H]thymidine
Vmax on ÔType 1Õ (70 versus 498 pmol min1 (106 cells)1
[15,130]) it seems possible that Type 1 does transport
[3H]thymine but that its contribution is a minor one.
ÔType 2Õ is described as a broad-specificity nucleoside
transporter [14] with moderate affinity for 2 0 -deoxycytidine, adenosine, guanosine, uridine and thymidine (Table 4). Affinity for 2 0 -deoxy and 5 0 -deoxynucleosides
was slightly less than for the ribonucleosides and affinity
for 3 0 -deoxyadenosine was much lower than for adenosine [14], showing the importance of the 3 0 -hydroxyl
group in substrate–transporter interactions and explaining the low affinity for nucleobases of this transporter.
In contrast, ÔType 3Õ appears to be a genuine nucleobase
transporter, of very low affinity but very high capacity,
with Km values of 1.4 ± 0.1 and 1.6 ± 0.4 mM for adenine and thymine, respectively [130]. All these studies
were performed at low temperatures to prevent parasite
attachment to vessel walls and facilitate a rapid sampling technique. A further study at 24 °C broadly supported the above findings, but found that nucleoside
transport was sensitive to the inhibitor dipyridamole
[131], while the earlier study had not [14]. This study
also found a much greater difference in affinity between
adenosine and 2 0 -deoxyadenosine [131]. Both groups
came to the conclusion, through different observations,
that nucleoside and nucleobase transport in Giardia
is non-energy dependent [130,131]. In summary, the
Table 4
Kinetic parameters of nucleoside and nucleobase transporters of
Giardia lamblia
Km or Ki value (lM)
2.6. Other protozoa
A limited number of studies have been performed
characterising purine transporters in other protozoa.
The aerotolerant intestinal parasite G. lamblia (also
known as G. intestinalis) causes a common form of
waterborne diarrhoea. G. lamblia trophozoites (the flagellate forms that infect the duodenum) reportedly express at least three distinct transporters for nucleosides
and/or nucleobases. ÔType 1Õ is described as a thymidine
transporter, with a Km of 50 lM [15]. This transporter
seems to be selective for oxopyrimidines as it was inhibited with similar affinity by uracil, uridine, thymine and
2 0 -deoxyuridine but not by the aminopyrimidines cytosine and cytidine, nor by ribose. The transporter displayed slightly higher affinity for the bases than for the
nucleosides [15]. However, the authors concluded in a
Transporter:
Reference:
Type 1
[15]
Type 2
[14]
Thymidine
Uridine
2 0 -Deoxyuridine
2 0 -Deoxycytidine
Adenosine
2 0 -Deoxyadenosine
Guanosine
2 0 -Deoxyguanosine
Tubercidin
Formycin A
Thymine
Uracil
Hypoxanthine
Adenine
50
64
96
115 ± 5
45 ± 25
93 ± 13
220 ± 116
45 ± 24
89 ± 8
26 ± 5
93 ± 13
Type 2
[131]
Type 3
[130]
54
668
82
260
13
1080
30
45
NS
1610 ± 370
2280
460
1440 ± 80
Numbers in bold type are apparent Km values, in normal type Ki values
or IC50 values, all in lM. NS, not saturable, uptake probably by
passive diffusion.
998
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
transporters of G. lamblia appear to have much lower
substrate affinity, especially for purines, than do the
transporters of the Apicomplexa and Trypanosomatidae, despite Giardia being purine auxotrophs [132]. This
probably reflects the very nutrient-rich environment of
the small intestine.
The purine salvage pathways of T. vaginalis, the etiological agent of sexually transmitted trichomonial vaginitis, are unique, consisting of a purine nucleoside
kinase [133,134] and a bacterial-type purine nucleoside
phosphorylase
[135],
while
lacking
phosphoribosyltransferases [133,134]. This simplified purine salvage system should therefore be eminently susceptible
to a rationally designed chemotherapeutic strategy
[136] and the adenosine analogue Formycin A, which
inhibits both the PNP and the PNK activities, potently
inhibited in vitro growth of T. vaginalis [136]. There
has also been a report of potent in vitro anti-trichomonal action of allopurinol in combination with
dipyridamole [137], despite the reported lack of a
hypoxanthine phosphoribosyltransferase and the inability to incorporate hypoxanthine or inosine into purine
nucleotides [138]. Pyrimidine salvage could be an
equally good target for therapeutic intervention as
T. vaginalis is auxotrophic for both purines [138] and
pyrimidines [139].
Despite the clear potential for purine or pyrimidinebased chemotherapy of this important pathogen, almost
nothing is currently known about nucleoside and nucleobase transporters in this parasite. T. vaginalis can
incorporate adenine and guanine but their transport
has not been studied. Two nucleoside transporters have
been reported, however, with apparent Km values between 2 and 13 lM for adenosine, guanosine and uridine. Thymidine, cytidine and inosine also inhibited
adenosine transport with Ki values < 100 lM, indicating
broad specificity for nucleosides, but no inhibition was
observed with nucleobases [140].
Purine salvage in T. foetus is manifestly different from
that in T. vaginalis, relying almost exclusively on salvage
through a hypoxanthine–guanine–xanthine phosphoribosyltransferase [141]. This makes the HGXPRT an
excellent target for chemotherapy and rationally designed inhibitors have been designed, which inhibit
T. foetus growth in the low micromolar range [6,142].
This reliance on phosphoribosylation is reflected in the
observation that T. foetus expresses two nucleobase
transporters (a hypoxanthine/guanine transporter and
xanthine transporter) but appears to lack nucleoside
transporters and adenine uptake appeared to be by passive diffusion only [23]. Both transporters were low affinity, with Km values of 0.7 ± 0.3 mM (hypoxanthine),
90 ± 20 lM (guanine) and 0.6 ± 0.2 mM (xanthine).
The absence of nucleoside transporters is to our knowledge unique for protozoa. Interestingly, Hedstrom and
Wang [23] also found that a T. foetus strain resistant
to the IMP dehydrogenase inhibitor mycophenolic acid
had lost the hypoxanthine/guanine transporter, apparently relying solely on a low affinity xanthine transporter
and passive adenine uptake for its purine requirements
[23]. The same strain had reduced adenine deaminase,
further contributing to low intracellular hypoxanthine
concentrations and thus aiding the incorporation of
xanthine into the nucleotide pool, bypassing IMP dehydrogenase [143].
Babesia bovis, the parasite that causes babesiosis
in cattle, infects host red blood cells and reportedly
introduces a broad specificity nucleoside/nucleobase
transporter in the erythrocyte membrane [144,145].
However, as the transport experiments were performed
with infected bovine erythrocytes, it is unclear whether
the transporter, which is sensitive to common nucleoside
transport inhibitors such as NBMPR, dilazep and dipyridamole [146], is located on the red cell membrane.
Alternatively, the transporter could be located on the
parasite plasma membrane, with the erythrocyte membrane fully permeabilised to nucleosides by a NPP-like
entity such as Plasmodium induces in human red cells
(see Section 2.4). A number of the adenosine analogues
were shown to be selectively toxic to intraerythrocytic
B. bovis [147], which is clearly dependent on the parasite-induced permeability for nucleosides, as normal bovine erythrocytes do not appreciably take up purine or
pyrimidine nucleosides [144].
Surprisingly little is known about nucleoside or
nucleobase transport in T. cruzi, and no new studies
have come out for over a decade. Finley et al. [10]
described a tubercidin-resistant epimastigote clone deficient in uptake of both [3H]thymidine and [3H]tubercidin, but not [3H]adenosine or [3H]inosine, leading once
more to a two nucleoside transporter model – though
it is certainly surprising that tubercidin (7-deazadenosine) would be taken up by the thymidine transporter
(TcNT1) rather than the proposed transporter(s) for
adenosine and/or inosine (TcNT2). Nonetheless, the
model was underpinned by the observation that [3H]tubercidin uptake was inhibited similarly by thymidine,
cytidine and uridine, but was only partially inhibited
by 500 lM adenosine and not at all by other purine
nucleosides [10]. One possible explanation would be that
the TcNT1, like LdNT1, also recognizes adenosine, but
in this case with low affinity, thus explaining the small
effect of adenosine on [3H]tubercidin transport. In such
a model, TcNT2 must be a genuine purine nucleoside
transporter with high affinity for adenosine, inosine
and guanosine, explaining that adenosine transport
was not significantly reduced in the tubercidin-resistant
clone. The same group later published an additional report on a different tubercidin-resistant epimastigote
clone [148], which displayed reduced transport only of
[3H]uridine and [3H]tubercidin, but not of [3H]thymidine
or [3H]adenosine. Possibly this clone expressed a
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
mutated pyrimidine nucleoside transporter with altered
substrate specificity. Epimastigotes were also able to
transport [14C]allopurinol riboside, [3H]formycin A
and [3H]formycin B, whereas in trypomastigotes lower
adenosine uptake rates correlated with reduced transport rates of formycin A, but not of formycin B or allopurinol riboside, which each inhibited growth rates of
some epimastigote lines [149]. Allopurinol has shown
promising activity against T. cruzi in vitro and in vivo
[150] and even against chronic ChagasÕ disease
[151,152], halving the cardiopathy associated with this
infection [153]. Yet, though the metabolism of allopurinol in T. cruzi has been well studied [154], and the compound must clearly be taken up by the parasite,
nucleobase transport in this parasite has yet to be
investigated.
3. Molecular biology of protozoan nucleoside and
nucleobase transporters
The existence of multiple purine transporters has
been demonstrated for a number of protozoan parasites
by functional assay on whole cells (Section 2), but the
burgeoning genome sequences for protozoan pathogens
reveal that the complexity of available purine transport
systems is, in many cases, still greater than anticipated
from functional assays. Moreover, characterisation of
individual transporter isoforms by heterologous expression has led to some surprises that have relevance to
purine uptake in other eukaryotic systems. Genome sequences are now available for a number of protozoa,
mainly those whose parasitic lifestyle renders them
pathogens of clinical or veterinary importance (http://
www.sanger.ac.uk/Projects/Protozoa/). Many of these
organisms encode multiple purine transporters, and
more candidates are emerging as annotation of newly sequenced genomes proceeds. Given that most protozoan
parasites transit between different hosts and environments, the existence of multiple membrane transporter
isoforms is to be expected, and developmental regulation of transporter expression will permit the parasite
to adapt to changing nutrient availability. Moreover,
the subcellular compartmentalization that is prominent
in many protozoa may require expression of transporters that are adapted to function at intracellular
locations.
All of the protozoan purine transporters that have
been cloned to date are members of the Equilibrative
Nucleoside Transporter (ENT) family, originally identified in mammals but with members throughout the
higher eukaryotes (see [155] for a recent review of the
ENT family). A second distinct group of nucleoside
transporters, the Concentrative Nucleoside Transporter
(CNT) family has members in both prokaryotes and
eukaryotes, but no CNT-like gene has yet been identified
999
from protozoa. The first ENT proteins to be characterised were found to be nucleoside transport facilitators
and were named as such because the group was distinct
from the CNT family. However, some protozoan ENTs
are, in fact, concentrative rather than equilibrative.
Moreover, some ENT family members are also nucleobase transporters [18,156] and recent data shows that
protozoan ENTs include dedicated nucleobase transporters [21,32,33]. This observation explains the
apparent lack of any previously identified nucleobase
transporter families [87] in the complete or emerging
protozoan genome sequences. The broad range of substrate specificities exhibited by ENT members in protozoa may also have relevance in mammals, where no
dedicated nucleobase transporter has yet been cloned.
Our homology searches of current protozoan genome
data reveal that, while ENT homologues are ubiquitous,
there is no evidence for homologues of non-ENT family
nucleoside or base transporters. In the discussion of
nucleoside and nucleobase transporter genes below we
will focus on a relatively small number of protozoan species of which the genome sequence is (almost) complete.
3.1. Trypanosoma brucei
The genome project for this organism is nearing completion but, as of August 2004 (release 3) (http://
www.genedb.org/genedb/tryp/index.jsp), the largest
chromosomes, IX, X and XI are still made up of multiple contigs, with gaps of unknown length between them.
As a consequence, it is not yet clear how many genes of
the ENT nucleoside/nucleobase transporter family are
encoded in the genome, and in some cases temporary
names must still be used, awaiting final assembly of
chromosomes. For example, the chromosomal position
of TbAT1, the gene encoding the P2 transport activity
and the first cloned T. b. brucei purine transporter, is
still unknown. Its closest homologue (Tb03.6N20.700)
is only 66% identical and 80% similar to the published
sequence of TbAT1 (AF152369 [28]).
The TbAT1 gene was first isolated by functional complementation of Saccharomyces cerevisiae naturally deficient in purine nucleoside uptake, using a cDNA library
of T. b. brucei STIB427 [28] and since from numerous
other strains [28] and field isolates [157] as well as from
T. equiperdum (M.L. Stewart and M.P. Barrett, unpublished) and T. evansi [66]. It does not appear in the latest
release of the T. b. brucei genome and it is therefore possible that multiple ENT family sequences are not yet
listed (see also below). The sequences of the T. equiperdum and T. evansi AT1 genes are >99% identical to the
original TbAT1 sequence.
3.1.1. Purine nucleoside transporter genes
In contrast, the position of the NT2 gene, encoding a
P1-type transporter, is very well defined. This gene is
1000
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
part of a cluster of 6 genes on chromosome 2, consisting
of TbNT2, TbNT3, TbNT4, TbNT5, TbNT7 and
TbNT6, in that order, with intergenic regions of 9 kb
between the ORFs [67], each of which contains a copy
of an ORF currently listed as a putative iron/ascorbate
oxidoreductase family protein in GeneDB. NT2 was
originally cloned from T. b. brucei strain EATRO 110
[27], which shares 447/463 (96%) identity at the amino
acid level with the NT2 sequence of the 927 strain used
for the genome project. Conservation within the NT2–
NT7 cluster is also considerable (81–89% of amino acid
sequence, relative to NT2). Despite this, two of the
genes, TbNT3 and TbNT4, appear to encode proteins
that either are non-functional or have a radically different function from the other 4 proteins. Whereas TbNT2,
TbNT5, TbNT6 and TbNT7 appear to be high affinity
P1-type adenosine/inosine transporters, no substrate
was identified for TbNT3 and TbNT4 [67]. It should
be noted that TbNT5, TbNT6 and TbNT7 also displayed a modest ability to mediate hypoxanthine transport when expressed in X. laevis oocytes, with a Km
value of 49 lM for TbNT5 [67] (see Table 5).
Recently, the group of Landfear [75] reported the
cloning of another P1-like transporter, designated
TbNT10, and showed that this was expressed exclusively
in the non-dividing short-stumpy lifecycle form of the
trypanosome. They demonstrated high-affinity uptake
for adenosine, guanosine and inosine, but not for nucleobases, and these results are entirely in agreement with
our own unpublished results with the same sequence,
which we had provisionally named Adenosine Transporter-like B (AT-B). This gene is not part of the NT2
cluster and is located on chromosome 9. Another transporter, AT-D, is also a P1-type (De Koning et al.,
unpublished) and located on chromosome 6. In addition
to mediating the transport of purine nucleosides it is
capable of mediating hypoxanthine transport, but with
very low affinity, as TbAT-D-mediated uptake of
1 lM [3H]hypoxanthine was not completely inhibited
by 2 mM hypoxanthine but was fully inhibited by
2 mM adenosine (Fig. 2). These eight P1-type nucleoside
transporter genes form a subgroup in a phylogenetic tree
of all (known) T. brucei ENT family genes (Group II;
Fig. 3). When ENT genes from additional protozoan
species and the known human ENT genes are included,
group II is shown also to include LdNT2 and CfNT2
(see below), with the other confirmed nucleoside transporters forming a separate group IV (Fig. 3).
3.1.2. Nucleobase transporter genes
A third such group is formed by TbNBT1, the gene
encoding the H4 purine nucleobase transporter [32],
and a number of very closely related copies including
the nucleobase transporter TbNT8.1 [33], which were
Table 5
Overview of ENT family genes in the T. b. brucei genome
Gene descriptor
Systematic name(s)a
Substratesb
Refs.
AT1
NT2
NT2/927
AF152369
AF153409
Tb927.2.6150
XM_340712
Tb927.2.6200
XM_340717
Tb927.2.6220
XM_340719
Tb927.2.6240
XM_340721
Tb927.2.6320
XM_340729
Tb927.2.6280
XM_340725
AF516605
AY204876
Tb06.28F21.780
AC092199
Tb09.160.5480
Ado, Ade, Pent, DA, MelB
Ado, Guo, Ino
Ado, Guo, Ino
[18,28,34,51,64,159]
[27]
[67]
Unknown
[67]
Unknown
[67]
Ado, Guo, Ino, Hyp
[67]
Ado, Guo, Ino (Hyp)
[67]
Ado, Guo, Ino (Hyp)
[67]
Hyp, Xan, Ade, Gua
Hyp, Xan, Ade, Gua, Guo (Ino)
Ado, Guo, Ino, Hype
[33]
[32]
Unpublished, SML, HPdKc,d
Ado, Guo, Ino
[75]; Unpublished, HPdKd
NT3
NT4
NT5
NT6
NT7
NT8.1
NBT1
NT9/AT-D
NT10/AT-B
Additional T. brucei ENT genes, currently of unknown function, include NT11.1/AT-A (Tb09.244.2020) and NT11.2/AT-G (Tb09.218.0180) ([91]
and De Koning, unpublished) as well as AT-E (Tb03.6N20.700) (De Koning, unpublished).
a
Names from GeneDB (www.genedb.org) or EMBL.
b
Ado, adenosine; Ade, adenine; Pent, pentamidine; DA, diminazene aceturate; MelB, melarsoprol; Guo, guanosine; Ino, inosine; Hyp, hypoxanthine; Xan, xanthine; Gua, guanine.
c
Unpublished results from the Landfear group, according to Ref. [91].
d
Unpublished results, De Koning group.
e
See Fig. 2.
Hypoxanthine Uptake
(pmol/107 cells)
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
1001
0.3
I
0.2
0.1
II
0.0
0
1
2
3
Time (min)
4
5
Fig. 2. Uptake of 1 lM [3H]hypoxanthine by S. cerevisiae strain
MG887-1 transformed with vector pDR195/TbAT-D. Yeast expression the AT-D-gene, or transformed with pDR195 without insert (s),
were incubated with [3H]hypoxanthine at a final concentration of 1 lM
for various intervals as indicated, in the presence of 2 mM hypoxanthine (h), 2 mM adenosine (.) or no inhibitor (j). The result shows
that TbAT-D transports hypoxanthine but with low affinity, while
being completely inhibited by adenosine.
cloned independently and concomitantly by our group
and the Landfear group. Both genes, which differ in only
three amino acids (see Table 6), were isolated from the
Ôreference strainÕ TREU 927. Both groups found evidence for multiple copies and Landfear et al. [33] sequenced an additional two copies of this gene, which
were designated NT8.2 and NT8.3. Their Southern blots
suggested the existence of a tandem repeat of related
genes, similar to that described for the TbNT2 cluster.
Considering the very high levels of sequence identify between the isoforms it is likely that each encodes a functional nucleobase transporter and it should be
remembered that T. b. brucei displays at least four different purine nucleobase transporter activities and a least
two distinct pyrimidine nucleobase transport activities
(see Section 2.1). At the moment it is impossible to predict how many of the isoforms listed in Table 6 will be
included in the final genome, and to what extent the
polymorphisms listed have a functional relevance. Even
with minimal differences in amino acid sequence,
TbNBT1 expressed in S. cerevisiae appears to encode
a higher affinity transporter than TbNT8.1 expressed
in the same organism, particularly for inosine and guanosine, for which NT8.1 displayed no affinity [32,33].
Interestingly, the recently cloned nucleobase transporter LmaNT3 and the putative nucleobase transporter
LmaNT4 (see Section 3.2) group together with the
T. brucei nucleobase transporters (Group III, Fig. 3).
3.1.3. TbAT1 and related genes: true nucleoside/
nucleobase transporters?
TbAT1 was cloned by Mäser et al. [28] and was predicted to be a single copy gene on the basis of Southern
blots. The construction of a TbAT1 null mutant by
III
IV
Fig. 3. Phylogenetic tree with protozoan and human ENT family
transporters. The tree was made from an alignment (see on-line
supplementary information) generated by the program DIALIGN
[247] using the blosum62 matrix. The tree itself was made using the
program tree-puzzle [248], which uses the maximum likelihood
method. A maximum parsimony tree using PHYLIP [249] gave the
same result. The bootstrap values are at the nodes of the tree, and are
given as percentages of results from 10000 replicates (puzzling steps).
Branch lengths are representative of phylogenetic distance. Group I,
TbAT1-like genes; Group III, apparent and possible nucleobase
transporter genes; Groups II and IV, nucleoside transporter genes.
Table 6
Polymorphisms of T. b. brucei nucleobase transporter genes
Transporter
Systematic
name
Position
Ref.
3
51
76
90
245
NBT1
NT8.1
NT8.2/AT-F
NT8.3
AT-J
AY204876a
AF516605a
Tb11.02.1105b
–
Tb11.02.1100b
L
L
L
I
L
S
S
A
A
A
T
I
I
I
I
P
S
S
S
S
N
K
K
N
N
[32]
[21]
[21]
[21]
Unpublished
NBT1 and NT8.1 have been cloned and characterised in heterologous
expression systems [21,32]. The NT8.2 and NT8.3 polymorphisms have
been obtained by direct sequencing from T. b. brucei (strain TREU
927) genomic DNA [21].
a
GenBank accession number.
b
www.GeneDB.org accession muber.
1002
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
Table 7
Km and Vmax values for some purines and trypanocides in bloodstream forms of T. b. brucei
Permeant
Transporter
Km (lM)
Vmax (pmol (107 cells)1 s1)
Vmax/Km
Ref.
Hypoxanthine
Hypoxanthine
Adenosine
Adenosine
Adenineb
Pentamidine
Diminazene
H2
H3
P1
P2
P2
P2
P2
0.12 ± 0.02
4.7 ± 0.9
0.38 ± 0.1
0.92 ± 0.06
0.25 ± 0.05
0.26 ± 0.03
0.45 ± 0.11
1.1 ± 0.2
1.1 ± 0.1
2.8 ± 0.4
1.1 ± 0.1
0.47 ± 0.11
0.068 ± 0.007
0.049 ± 0.010
9.2
0.23
7.4
1.2
1.9
0.26
0.11
[20]
[20]
[53]a
[53]a
–
[159]
[51]
a
Very similar values were published first by Carter and Fairlamb [18]. For a consistent comparison of transport rates of adenosine, hypoxanthine
and the diamidines, values were taken from studies by the same group.
b
De Koning et al., unpublished. Average of 3 experiments using 20 nM [3H]adenine in the presence of 100 lM hypoxanthine.
targeted gene deletion recently confirmed this, as the
P2-type adenosine transport activity was not detectable
in the Tbat1/ strain [34].
As mentioned above, TbAT1 cannot as yet be located
in the T. brucei genomic databases, but three related
genes (see legend to Table 5), which we designated
TbAT-A, E and G until a function can be assigned, have
been identified. These genes have 58%, 66% and 58%
identity with TbAT1 at amino acid level, respectively.
With TbAT1, they form a fourth phylogenetic group
of ENT family genes (Group I, Fig. 3). TbAT1 encodes
a highly unusual transporter, P2, which has very high
affinity for both adenine and adenosine, and transports
both with similar efficiency (Table 7), but does not transport oxopurines or pyrimidines [18,53]. It could therefore be described as an aminopurine transporter rather
than a nucleoside or nucleobase transporter. However,
it also efficiently transports a number of non-purine trypanocides of clinical importance, including melaminophenyl arsenicals and diamidines, which share a
structural recognition motif (see Section 4.2). While
the Km value for adenosine and two diamidines (pentamidine and diminazene aceturate) are very similar,
the maximal uptake rate (Vmax) for the trypanocides is
much lower than for aminopurines (Table 7). One question now is the extent to which the related genes TbATA, E and G will prove to be aminopurine transporters or
perhaps diamidine transporters. It has been well documented that trypanosomes express two pentamidine
transport activities in addition to P2: the High Affinity
Pentamidine Transporter (HAPT1) and the low affinity
Pentamidine Transporter (LAPT1) [34,158–160]. Neither is inhibited by high concentrations of purines nor
pyrimidines.
3.2. Leishmania spp.
The first protozoan purine transporter gene to be
characterised, LdNT1, was cloned [26] from a L. donovani cosmid library by functional complementation of
a strain that had been rendered adenosine transportdeficient after selection with the cytotoxic adenosine
analogue tubercidin. Two almost identical genes,
LdNT1.1 and LdNT1.2, were identified by this approach, both clear homologues of the ENT genes that
had recently been identified in mammals. Expression
of LdNT1 in Xenopus oocytes and in adenosine transport-deficient L. donovani facilitated its characterisation
as an adenosine/pyrimidine nucleoside transporter. A
second nucleoside transporter, LdNT2, was cloned
using a similar strategy, complementing an inosine/guanosine transport mutant. LdNT2 encodes a high-affinity
inosine/guanosine transporter [27].
The recent completion of the first Leishmania genome
project has brought to light homologues of the functionally identified transporters LdNT1 and LdNT2. A
recent report [21] describes the functional characterisation of LmaNT3, an LdNT1 homologue that exhibits
high affinity for a range of nucleobases. LmaNT3 has
considerable identity with the trypanosome nucleobase
transporters TbNBT1 [32] and TbNT8.1 [33]. Thus, in
Leishmania as well as trypanosomes, ENT family members are, at least in part, responsible for nucleobase
transport, in addition to nucleoside transport and it
appears that ENTs that transport nucleobases with
high-affinity cluster together in a phylogenetic tree of
protozoan ENTs (Group III, Fig. 3). A further ENT
homologue is apparent in the L. major genome
(LmjF11.0550) and functional characterisation of this
transporter, provisionally named LmaNT4, is reportedly in progress [91]. This gene encodes a 550 amino acid
protein, which appears to have a unique insertion of
180 amino acids. TMPRED, and our multiple alignments, suggest that this insertion is located in an intracellular hydrophilic loop between transmembrane
domains 6 and 7 of an 11 TM topology (see Section
5.1). This uncharacterised transporter also aligns within
the Ônucleobase transporter domainÕ of the phylogenetic
tree (Fig. 3).
3.3. Plasmodium falciparum
The genome sequence of P. falciparum was the first
completed and has thus been the most extensively studied
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
and most comprehensively annotated of the protozoan
genomes. To date, a single P. falciparum ENT homologue
has been identified. The gene was identified concurrently
by two independent groups and named PfNT1 [30] or
PfENT1 [31] (see Section 2.4). Functional characterisation by both these groups revealed a high-affinity nucleoside transporter of broad specificity, with moderate
affinity for some nucleobases. The P. falciparum genome
contains four different ENT genes (S.A. Baldwin, personal communication) and the characterisation of the
additional transporters must be the highest priority for
understanding purine salvage in this parasite.
3.4. Toxoplasma gondii
The first T. gondii purine transporter, TgAT, was
functionally identified by insertional mutagenesis, leading to resistance to a toxic adenosine analogue [29].
Upon heterologous expression in Xenopus oocytes,
TgAT was revealed as an adenosine transporter of moderate affinity. Inhibition studies suggested a broad specificity for nucleosides and nucleobases and the loss of
adenosine transport capacity in the insertional mutant
suggested that TgAT was the sole transporter responsible for adenosine uptake in T. gondii. However, studies
of purine uptake by T. gondii tachyzoites suggested a
multi-component uptake system [8] and several additional ENT genes can be readily identified in the now
substantially complete T. gondii genomic database (ToxoDB.org). These genes have been named TgNT1,
TgNT2 and TgNT3 [127].
3.5. Crithidia fasciculata
Liu et al. [92] very recently cloned two Cr. fasciculata
ENT transporters designated CfNT1 and CfNT2, by
probing a Crithidia library with LdNT1 and LdNT2.
While the two Crithidia genes were only 30% identical,
they were 72% and 73% identical to their L. donovani
counterparts, showing both functional and genetic conservation between the ENT transporters of these species.
Southern blots revealed that, while CfNT2 exists as a
single copy, multiple copies of CfNT1 could be detected.
The observed pattern was not consistent with (only) a
tandem array for CfNT1, and though it is not clear
how many copies exist in the Cr. fasciculata genome,
several more were identified and sequenced using PCR
approaches [92]. While the sequence of the Crithidia
ENT genes may thus be closest to the Leishmania transporters, the organisation is much closer to that of
T. brucei.
3.6. Subcellular transporters
The translocation of purines and pyrimidines across
the intracellular membranes that compartmentalise
1003
eukaryotic cells is poorly understood, and no data exist
for any protozoa. Nevertheless, the high level of subcellular complexity exhibited by protozoa may partially account for the large number of structurally related
transporters identified in the genomes of some parasites
(notably that of T. brucei). Adenosine transport systems
have been characterised in both mammalian lysosomes
[161] and mitochondria [162], though the genes encoding
the proteins that mediate these activities have not yet
been identified. Transporters with discrete subcellular
localisations are likely to be required for organellar purine/pyrimidine metabolism in protozoa as in other
eukaryotes.
Mitochondria, for instance, require purines/pyrimidines for replication of, and transcription from, the
mitochondrial genome. Rat testis mitochondria contain
a high-affinity adenosine transporter distinct from the
ATP/ADP exchanger [162]. In the Apicomplexa, the
membrane-delimited plastid may also require transporters for delivery of the purines and pyrimidines that are
required for replication of the organellar genome [163].
The glycosome, a modified microbody that appears to
be unique to kinetoplastid parasites and which harbours
some of the enzymes of the purine and pyrimidine metabolic pathways [164–168], is likely to require membrane
transporters for the exchange of purines and pyrimidines
from the cytosol.
However, the challenges of obtaining intact organelles in sufficient quantities for the study of transport
processes have so far proved insurmountable. Genome
data have the potential to provide clues to subcellular
localisation, but there is as yet no understanding of
the targeting signals that must act to direct membrane
proteins to discrete intracellular membranes. The use
of fluorescent or epitope tags has the potential to reveal
the subcellular localisation of membrane transporters
and such approaches have been exploited (see below) despite the valid concern that introduction of a tag may
disrupt the normal targeting.
Thus, for several reasons, the mechanisms by which
integral membrane proteins are targeted to discrete subcellular membrane domains are poorly understood. The
limited number of protozoan purine/pyrimidine transporters that have been localised to date are found to
be homogeneously distributed in surface membranes
(LdNT1 isoforms: [169]; LdNT2: [170]; TbNT8.1: [33];
PfNT1: [115]). Indeed the only example of differential
subcellular localisation of membrane transporters
amongst the protozoa is the discrete localisation of hexose transporters to the distinct pellicular and flagellar
surface membranes of kinetoplastid parasites. Despite
extensive studies, the targeting signal is still poorly
understood [171–173], as is the possible function of the
transporter in the flagellar membrane. Nevertheless, it
is possible that some of the many ENT genes in the genomes of some kinetoplastids may encode isoforms with
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
flagellar localisation, where they may contribute to
acquisition of purines and pyrimidines.
Whereas T. brucei encodes a large number of ENT
family transporter genes, some of which may localise to
distinct intracellular locations, other parasites (notably
P. falciparum and L. major) possess very few ENT family
genes, apparently encoding proteins localised to the parasite plasma membrane. Intracellular purine/pyrimidine
transport in these species may thus be mediated by transporters from an as yet unidentified gene family.
3.7. General comments
Characterisation of purine uptake by protozoa has
proceeded over several decades, giving rise to a large
body of data on a diverse range of (mainly pathogenic)
protozoa. Parallel approaches with prokaryotes and
higher eukaryotes have revealed that purine transport
systems are ubiquitous amongst both auxotrophs and
heterotrophs. In the past decade, many genome projects
have been initiated and a number are now complete or
nearing completion. Concurrently, functional cloning
approaches have identified a number of genes that encode purine transporters. These transporters fall into a
variety of families [85–87] and extensive functional characterisation and gene annotation will be required before
the data on purine transport in whole organisms can be
reconciled to the genetic data.
To date only ENT family members have been identified in protozoa: both function- and homology-based
screens have failed to reveal purine or pyrimidine transporters from other families. For those parasites with
complete genome sequences, it therefore appears that
any additional nucleoside or nucleobase transport activity must be encoded by transporters of novel gene families. Yet, no such transporters have been identified to
date. Functional screens for purine transporters have given rise only to ENT-like genes [25–32] and extensive in
silico searches and hybridisation library screens have
failed to identify homologues of non-ENT nucleoside
or nucleobase transporters. At the time of writing, it
seems possible that purine uptake in protozoa may be
entirely mediated by members of the ENT family. It is
not yet clear, however, whether the existing complements of ENT family members in each parasite species
studied are sufficient to account for all the transport
activities identified in the corresponding organism. For
instance, no dedicated pyrimidine transporter genes
have been identified, even though from several protozoa
(nearly) all ENT members have been cloned and characterised. Therefore, some of these transport activities may
be mediated by novel protein families, particularly since
the structure of the binding pocket of the L. major and
T. brucei uracil transporters appears to be remarkably
different from the ENT purine transporters characterised to date [40].
It seems reasonable to postulate a requirement for
multiple purine transporters to sustain a parasite
through its life cycle, and indeed a battery of purine
transport activities has been described in procyclic and
bloodstream form trypanosomes. However, it is at present difficult to reconcile the presence of perhaps as many
as 16 distinct ENT family members in the T. brucei genome while P. falciparum and Toxoplasma gondii, organisms with a no less complex life cycle, can apparently
satisfy their purine requirements with far fewer ENT
genes. However, all protozoan genomes that have been
sufficiently investigated for the presence of nucleoside
transporter genes have now been found to contain several ENT family members.
The recent advances have produced an abundance of
evidence for significant molecular and functional divergence in purine salvage pathways between protozoan
parasites and their mammalian hosts. This raises the
possibility of specific chemotherapeutic intervention in
parasite purine acquisition.
4. Protozoan nucleoside and nucleobase transporters in
chemotherapy
4.1. Transport or diffusion?
In mammalian systems, nucleoside drugs may act on
intracellular or extracellular targets, an example of the
latter being purine receptors. In protozoa, all current
targets for nucleoside or nucleobase antimetabolites
are intracellular, which means that the drug needs to
cross the parasite plasma membrane, and possibly further organellar membranes. In addition, when targeting
an intracellular parasite, the (pro)drug must cross the
host cell plasma membrane and any parasitophorous
vacuole membrane. These requirements are obviously
different for various protozoan species and need to be
taken into consideration when designing any antiprotozoal agents.
The ability of drugs such as nucleosides to diffuse
across biomembranes depends on their lipophilicity,
usually quantified as the octanol–water partition coefficient log P or clog P for calculated log P. Both lipophilic
and hydrophilic drugs have advantages with respect to
targeting parasites. An obvious advantage of very lipophilic drugs is that they will cross any membrane and
reach intracellular parasites and organellar targets perhaps easier than do hydrophilic compounds. Furthermore, they may cross the blood–brain barrier and thus
have activity against parasitic infections of the CNS,
e.g. late-stage African trypanosomiasis or cerebral toxoplasmosis, though lipophilicity is by no means the only
factor for CNS penetration. A potential drawback is
the loss of specificity that results from selective accumulation by a parasite, rather than host cells. Drugs that
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
enter cells by diffusion will also not be accumulated to
free intracellular concentrations higher than the extracellular drug concentration (blood, cerebrospinal fluid,
etc.). While accumulation is still possible when driven
by intracellular modification or high affinity binding to
a target, active transport can accumulate hydrophilic
compounds to free intracellular levels very much higher
than therapeutic plasma concentrations, locally reaching
concentrations not achievable or tolerated in blood.
Furthermore, lipophilic compounds are more likely to
be substrates of ABC-transporters that confer multidrug
resistance [174] and clear Ôforeign substratesÕ from the
CNS [175], though some nucleoside and nucleotide
analogues have been shown to be substrates for the
human MDR4 and MDR5 transporters [174–176].
Most purines and pyrimidines do not diffuse through
biomembranes at appreciable rates, but some nucleoside antimetabolites do, including desciclovir [177],
abacavir [178] and NA-42 (2-cyclopentylamino, N6cyclopentyladenosine), an experimental compound
with submicromolar in vitro activity against African
trypanosomes [179].
4.2. The TbAT1/P2 transporter of T. brucei
The best-researched example of a protozoan transporter involved in chemotherapy is undoubtedly the
P2/TbAT1 aminopurine transporter of African trypanosomes. There is abundant evidence of its ability to transport melaminophenyl arsenicals and diamidines,
essential agents in the treatment of African trypanosomiasis. Though this has been extensively reviewed elsewhere [52,63,180,181], it is appropriate to discuss some
of the key experiments here.
Evidence for P2 involvement in the uptake of melaminophenyl arsenicals followed from the observation by
Carter and Fairlamb [18] that only substrates of this
transporter (adenosine and adenine) were able to abrogate lysis induced by melarsen oxide in vitro. These purines were unable to affect lysis induced by the highly
lipophilic arsenical phenylarsine oxide, showing the effect to be on the level of uptake. Furthermore, a melarsoprol-resistant line, cRU15, was shown to have lost the
P2 transport activity [18]. The expression of TbAT1 in
yeast conferred some sensitivity to melarsen oxide [28]
and P2-mediated adenosine transport is strongly and
competitively inhibited by all melaminophenyl arsenicals [18,28,53]. Perhaps the definitive evidence was presented using a T. b. brucei clone in which both alleles
of TbAT1 had been deleted [34]. In tbat1-null trypanosomes, the rapid in vitro lysis induced by melarsen oxide
or cymelarsan no longer occurred, but a much slower,
delayed lysis was still evident and, in contrast to the rapid phase of lysis, not sensitive to inhibition by adenosine. Lysis with phenylarsine oxide was unchanged in
tbat1-null cells. Consistent with these observations,
1005
resistance to melaminophenyl arsenicals, both in vitro
and in vivo, was minimal [34]. These experiments show
that, even though P2 is the main transporter for melaminophenyl arsenical uptake, a secondary entry system
exists. The additional uptake mechanism was shown to
be highly sensitive to pentamidine and propamidine,
with 50% effective concentrations similar to the pentamidine Km and propamidine Ki values for the HAPT1
pentamidine transporter, respectively [34]. The implication is that loss of P2 function may be a necessary but
not sufficient condition for melaminophenyl arsenical
resistance and that concomitant loss of HAPT1 would
be required for high levels of resistance.
Pentamidine transport by P2 was first demonstrated
by Carter et al. [64], and accumulation of pentamidine
was reduced in cRU15 [64], the melarsoprol-resistant
line without a functional P2 transporter [18]. Yet, this
clone was not resistant to pentamidine [64], a finding
that was later confirmed with the tbat1 null trypanosomes [34]. As with the arsenicals, P2 is the main conduit for pentamidine, but additional transporters are
sufficient for almost normal efficacy of the drug
[34,159,160]. HAPT1 and LAPT1 have not yet been
cloned and their physiological substrate and function
remain unknown. It is certain, however, that these
T. brucei pentamidine transporters are not purine or
pyrimidine transporters [158,159]. Nor are they choline
transporters (De Koning, unpublished) like the pentamidine transporter in P. falciparum [182]. The P2
transporter clearly mediated the high-affinity transport
of [3H]pentamidine in bloodstream trypanosomes
[64,158,159] and tbat1 null trypanosomes lacked adenosine-sensitive pentamidine transport [34]. However,
adenosine transport by TbAT1 expressed in S. cerevisiae
appeared not to be inhibited by pentamidine, leading to
speculation about cofactors or modifications necessary
for diamidine transport [28]. However, a reinvestigation
[51], using the same yeast strain and vector, found that
diamidines, and in particular pentamidine, bind heavily
to the yeast cell surface, possibly with higher affinity
than to TbAT1, leading to very high backgrounds during transport studies and precluding the measurement
of true initial rates of transport. Even so, accumulation
of pentamidine over 1 h is clearly much higher in yeast
expressing TbAT1 than in control cells [51].
Evidence for P2-mediated transport of the related
diamidine diminazene aceturate (DA; Berenil) mostly
parallels that for pentamidine. DA has high affinity for
P2 [51,53,57,64] and a DA-resistant T. equiperdum line
was shown to have lost normal P2 function [57]. The
tbat1 null clone was highly resistant to DA [34] and lost
saturable [3H]DA transport [51]. Furthermore, [3H]DA
was clearly transported by TbAT1 expressed in yeast
[51]. Recently Witola et al. [66] reported that RNAi
silencing of AT1 in T. evansi also confers resistance to
diminazene.
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The Km values for [3H]DA and [3H]pentamidine
transport by P2 are similar to the Km for adenosine (Table 7), though the Vmax values for the diamidines appear
very much lower. The different translocation rates, despite similar affinity, indicate different transport efficiencies for these compounds, expressed as Vmax /Km.
In addition, P2 displays high affinity for the veterinary phenanthridine trypanocide isometamidium (structurally a hybrid of diminazene and homidium
(ethidium), see Fig. 4) [28,52], but at present there is
no evidence that P2 is involved in isometamidium uptake other than a small but significant reduction in
isometamidium sensitivity in T. evansi with induced
RNAi for P2 [66].
It is not currently believed that P2 is the major route
of entry for isometamidium. Yet, the fact that purine
nucleobases, nucleosides, diamidines and phenanthridines could all have very high and almost identical affinity for the same ENT transporter that is otherwise very
selective (no interaction with oxopurines or pyrimidines,
a
H-bonds
NH2
6
N1
2
for example, at 1 mM) is highly unusual. The key to this
lies in the architecture of the P2 binding pocket and
hence in the way it interacts with potential substrates
such as adenosine. The main P2-adenosine interactions
were independently elucidated from the ability of various compounds to antagonise melarsen oxide-induced
lysis of T. b. brucei [63] or through inhibition of P2-mediated [3H]adenosine transport [53]. The presence of an
accessible amidine motif NR1@CR2–NH2, an aromatic
ring, and possibly a N or O residue in the position corresponding to N9 of the purine ring, are essential for
high-affinity binding and all P2 substrates share this motif (Fig. 4). In addition, a similar structural configuration seems to be essential for high affinity binding to P2.
4.3. Nucleobase transporters of Leishmania and
Trypanosoma spp.
The hypoxanthine analogue allopurinol is in clinical
use against various forms of leishmaniasis [49,50,183–186]
b
NH2
possible electrostatic
interactions
7
5
N
4
N δ+
N
N
8
N
H2 N
9
3
N
NH
melarsoprol
O
HO
π-stacking
OH
HO
adenosine
As
S
S
c
d
+
NH2
OH
H2 N
+
N
N
HN
H2N
H2N
isometamidium
O
(CH2)5 O
pentamidine
N
+
H2C
CH3
NH2
NH2
+
NH2
e
H2N
+ NH2
δ+
N N N
H
NH2
+
NH2
diminazene
Fig. 4. Predicted interactions between some actual permeants and a potential permeant (Isometamidium [52] of the T. b. brucei AT1/P2 transporter
and amino acid residues in the P2 translocation pathway, based on information from Refs. [51–53,63,159]). Shaded areas are functional groups
believed to interact with the transporter. Some care was taken to depict the various molecules in likely conformations, accurately reflecting bond
angles, etc., as far as possible. Alternative conformations are possible, due to free rotation around some bonds.
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
and is taken up by hypoxanthine transporters of L. major promastigotes and L. mexicana amastigotes (see Section 2.2). In both cases a single transporter was
responsible for the uptake of the drug, which should
raise some concern about possible drug resistance, as
the hypoxanthine transporters are unlikely to be essential given the presence of two high affinity nucleoside
transporters in amastigotes of L. donovani [9]. It has
been reported that allopurinol resistance can be readily
induced through in vitro exposure of promastigotes
[187]. In contrast, T. b. brucei express multiple nucleobase transporters, both in procyclic and in long-slender
bloodstream forms, each with (relatively) high affinity
for allopurinol [19,20,32]. If, as expected, the speed at
which drug resistance develops in these parasites were
partially dependent on the number of individual transporter activities mediating uptake of the drug, onset of
allopurinol resistance would be much delayed in trypanosomes. This was tested in procyclic T. b. brucei, where
allopurinol transport was clearly mediated by both the
H1 and H4 hypoxanthine transporters. As predicted, attempts to induce transporter-related allopurinol resistance were unsuccessful after >12 months of exposure
1007
to 3 mM allopurinol [188]. Culture conditions, using
inosine as sole purine source, had been chosen to preclude resistance arising through loss of enzymes of the
purine salvage pathways. This experiment showed that
avoidance of at least one major mechanism of resistance
to purine drugs is feasible if the drug is accumulated
through multiple transporters.
Another cytotoxic nucleobase that is actively accumulated by both T. b. brucei and L. major insect forms
is 5-fluorouracil. In both organisms, otherwise very
exclusive uracil transporters (TbU1 and LmU1) displayed high affinity for this drug, with Km values of
3.0 ± 0.8 and 0.66 ± 0.14 lM, respectively [11,40]. Fluorouracil proved to be more effective against L. major
promastigotes in vitro than either allopurinol or aminopurinol, with an ED50 of 5.1 ± 1.6 lM (only 3-fold higher than for pentamidine in the same assay [40]) or
4.5 ± 0.6 lM against L. amazonensis promastigotes
[189]. Crucially, the uracil analogue also displayed
promising activity using L. major-infected macrophages
[40]. Flow cytometric analysis unambiguously shows a
massive reduction in parasite burden after treatment
with just 10 lM 5-fluorouracil (Fig. 5).
Fig. 5. FACS histogram acquired with macrophages infected with L. major V39. The macrophages were infected for 5 h, washed and incubated in
medium without or with 10 lM or 10 nM 5-fluorouracil for a further 72 h before FACS analysis. Figure reproduced from Ref. [40], with permission.
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
A major advantage of the use of purine nucleobase
analogues against trypanosomes or Leishmania is the
fact that substrate recognition by their nucleobase transporters is very different from that of their human counterpart, the facilitative nucleobase transporter (hFNT1)
[39]. This allows efficient uptake of antimetabolites by
the parasite transporters, while excluding them from
normal host cells. While this would seem to be a disadvantage when combating intracellular parasites, purine
uptake systems are generally altered in parasitised host
cells. This usually results in wider access of purine antimetabolites to the intracellular parasite than might be
expected from purine transport by the pre-invasion cell
(see below). Examples of selective recognition of purine
antimetabolites by protozoan nucleobase transporters
are 3-deazaguanine, 6-thioguanine and 6-thiopurine
[39] as well as a range of thieno-separated tricyclic nucleobase analogues [190]. The tricyclic purines retained
very high affinity for the T. b. brucei H2 transporter
(Ki values <2 lM) and the hypoxanthine-like tricyclic,
TRI-B-002 displayed approximately 100-fold higher
affinity for the trypanosome transporter than for
hFNT1 [190]. It needs to be recognised, of course, that
inhibition of a transporter does not necessarily equate
transport of the inhibitor at any significant rate. However, some of these tricyclic nucleobases show appreciable trypanocidal activity in vitro [190], indicating that
they are internalised.
4.4. Nucleoside antimetabolites and transport
A rather large number of nucleoside analogues have
been reported to possess antiprotozoal activity, and it
is certainly not within the scope of this review to give
an exhaustive listing. The major concern with taking
nucleosides into human trials has always been the potential for toxic or teratogenic side-effects (see also Ref.
[191]) and, without doubt, many nucleosides with antiprotozoal activity are also acutely toxic to their hosts.
Selectivity, however, is possible on the grounds of: (1)
selective metabolism by the parasite, (2) selection at
the drug target level and (3) selective accumulation.
Examples of all three mechanisms can be readily
identified.
An example of the first is 9-deazainosine, which is
converted to 9-deazaadenine nucleotides by Leishmania
and trypanosomes but not by mammalian cells
[192,193], due to differences in their adenylosuccinate
synthetase and/or lyase specificities [192,194]. As
pointed out by Bhattacharya et al. [195], the amination
appears to be the key to the selective activity of 9-deazainosine as 9-deazaadenosine is acutely cytotoxic [196].
The second category includes inhibitors of glycolytic
enzymes in T. b. brucei, in particular glyceraldehyde-3phosphate dehydrogenase (GAPDH), as T. brucei spp.
lack a functional citric acid cycle and are totally depen-
dent on glycolysis for their energy metabolism [197].
Large numbers of 2,N6-disubstituted and 2 0 -N6 disubstituted adenosine analogues have been synthesised and
optimised for GAPDH inhibition by structure-based design [43,198], some of which displayed low micromolar
activity against T. b. brucei, while being much less toxic
to mammalian cell lines.
Most protozoa express nucleoside transporters with
much higher apparent substrate affinities than (most)
mammalian nucleoside transporters. One example is
the T. b. brucei P1 transporter, which displays high affinity for a whole range of nucleoside drugs including Formycin A, Formycin B, 2-chloroadenosine, ribavirin
[53]). In contrast, ribavirin, for example, enters human
erythrocytes through the NBMPR-sensitive nucleoside
transporter with a Km of 0.5 mM [199]. Other examples include the uptake of tubercidin and cordycepin
by TbAT1 [53], Formycin A and tubercidin by L. donovani NT1 [26,80] and T1 [9], Formycin B by LdNT2 [25]
and T2 [9] and L-nucleosides by P. falciparum (see
below).
4.5. Use of protective nucleoside transporter inhibitors
Apart from antimetabolite uptake by parasite transporters on the basis of different permeant selectivity, a
further approach to targeted uptake of antimetabolites
is possible by specific inhibition of the host nucleoside
transporters. This approach was extensively reviewed
by El Kouni recently [191], and allows the use of inherently toxic nucleosides, as their uptake into mammalian
cells will be blocked. Again, this approach has been
abundantly validated, but no clinical strategies have resulted. Most protozoan nucleoside transporters have
been shown to be insensitive to the traditional inhibitors
of mammalian nucleoside transport, dipyridamole, dilazep and NBMPR (reviewed in [128]). Co-administration
of, in particular, NBMPR together with nucleoside antimetabolites has been shown to protect the host from
their toxic effects and be efficacious in the treatment of
several parasitic infections including T. b. gambiense
[200], Plasmodium spp. [201,202] and Schistosoma spp.
[203–206]. NBMPR itself displayed no toxic effects in
mice at 25 or even 100 mg/kg [206,207].
It might be anticipated that this approach would not
work with intracellular parasites, as the transport inhibitor would prevent uptake of the nucleoside drug across
the host cell plasma membrane. However, the expression
of the New Permeation Pathway in Plasmodium-infected
erythrocytes (see Section 2.4) and, apparently, of T. gondii-encoded transporters in infected human fibroblasts
[208], ensure NBMPR-insensitive uptake even in parasitised cells. In fact, these infected-cell-specific nucleoside
uptake pathways appear capable of NBMPR uptake,
whereas this drug is not internalised in non-infected
mammalian cells [128]. Together with phosphorylation
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
by the parasite adenosine kinase [191,208], this leads to
selective toxicity of NBMPR against T. gondii [191,208,
209] and P. falciparum [202]. Therefore, NBMPR not
only protects host cells from toxic nucleoside analogues, it also is selectively toxic to the parasite itself.
Similar observations have been made with other mammalian nucleoside transport inhibitors, in particular
dilazep [210], which is already in clinical use as a
vasodilator.
4.6. Chiral nucleosides
The absolute dependence of Plasmodium on an external purine source has led to the identification of purine,
and in particular adenosine, analogues as anti-malarials
[211]. Many of these compounds do not exhibit acceptable parasite selectivity. However, targeting may be
achieved following the observation that non-physiologically relevant L-nucleosides are transported into the parasite but not into mammalian cells [106]. In this study,
both L-and D-adenosine were not only transported into,
but also metabolised in, merozoite and intraerythrocytic
parasites. Intriguingly, L-adenosine uptake was not observed in saponin-freed intraerythrocytic parasites,
although it was in infected erythrocytes treated with sendai-virus [212]. This permeation pathway also remains
enigmatic at the molecular level due to conflicting reports regarding PfENT1/PfNT1 substrate specificity
(see Section 2.4). Nevertheless, the apparent lack of stereospecificity displayed by the parasiteÕs purine salvage
pathways provides a strong rationale for drug design:
only the parasitic enzymes are capable of metabolising
the L-nucleosides [213]. The selective entry and metabolism strongly suggests the chemotherapeutic use of
L-enantiomers of toxic nucleosides, and initial analysis
confirms the strength of this approach [213]. For example, L-isocoformycin (a structural analogue of the potent
adenosine deaminase (ADA) inhibitor 2 0 -deoxy-D-coformycin), has a Ki of 7 pM (90% inhibition) for parasite
ADA, and yet the mammalian homologue neither binds
to the drug in vitro nor is exposed to the drug in vivo
[214]. A second approach is the utilisation of L-nucleosides as ÔcarrierÕ molecules to deliver pro-drugs or established anti-malarial compounds [215], in a piggy-back
approach as has been envisaged for substrates of the
T. b. brucei P2 transporter [35–38].
4.7. Possible resistance mechanisms
The central role of transporters in therapy based on
nucleoside and nucleobase antimetabolites means that
loss of transporter activity may cause resistance to such
drugs [80]. This principle has been utilised to advantage
in the cloning strategies of the Leishmania [25,26] and
Toxoplasma [29] nucleoside transporters. The loss of
1009
the T. brucei TbAT1/P2 transporter has been linked to
resistance to several key trypanocides (see Section 4.2).
Analysis of TbAT1 alleles from isolates of patients that
relapsed following melarsoprol treatment, using RFLP
analysis [216], SSCP (single strand conformation polymorphism) and direct sequencing has revealed a remarkably small number of TbAT1 polymorphisms that differ
in their drug transport abilities, with a common set of
nine mutations found in various geographical locations
and in both human infective subspecies [157]. However,
a number of relapse patients retained the wild-type
TbAT1 gene suggesting that additional factors may be
involved [157]. As discussed in Section 4.3, it is possible
to avoid, or at least delay, transporter-associated resistance if the drug is taken up by several distinct carriers.
Resistance can also be associated with increased
extrusion of the drug from the parasite. Many, if not
all protozoa, are also known to encode ATP-Binding
Cassette proteins (ABC proteins) including P-glycoproteins [217–220] and some of these have been shown to
be involved in (multi-)drug resistance or the efflux of
xenobiotics. Non-exhaustive examples include EhPgp1
and EhPgp5 of Entamoeba histolytica [221], PGPA
of Leishmania spp. [218], verapamil- and cyclosporin
A-sensitive P-glycoproteins in T. gondii [222], T. b. brucei MRPA [223] and P. falciparum mdr1/pgh1 [224,
225]. Such ABC transporters certainly contribute to
resistance to clinically important drugs such as quinine
and mefloquine (malaria) and antimonials (leishmaniasis), but in other cases it is their ability to mediate drug
efflux when experimentally overexpressed that allocates
them a potential role in drug resistance. However, to
our knowledge, little information exists about the ability of protozoan ABC transporters to confer resistance
to therapeutic nucleoside analogues as is the case with
human MRP4 and MRP5 [174]. Expression of L. major
PGPA, for instance, resulted in 10-fold resistance to
arsenite and trivalent antimonials but not to puromycin
[226]. However, Katakura et al. [189] recently reported
that overexpression of LaMDR2 in L. amazonensis
promastigotes leads to 2.5-fold resistance specifically
to 5-fluorouracil, and not to the nucleosides Formycin
B, puromycin and tubercidin, arsenite, cadmium or regular antileishmanials. For recent reviews of protozoan
ABC transporters in drug resistance, see Refs.
[220,227].
A further mechanism of reducing the accumulation of
therapeutic purines was reported in Leishmania spp.
[228–230]. In L. mexicana amazonensis selected for resistance to tubercidin or inosine dialdehyde, uptake of purine nucleosides and nucleobases was reduced [229]. This
was linked to an extrachromosomal DNA of approximately 55 kb in size, which was not present in sensitive
parent strains or revertants [229]. From this DNA a
TOxic nucleoside Resistance (TOR) gene (GenBank
Accession No. AF016581) was identified, which encodes
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
a 478 amino acid protein that contains a possible (but
unconfirmed) trans-membrane domain (www.ch.embnet.org/software/TMPRED_form.html) but displays
similarities to mammalian transcription factors [228],
suggesting a role in the regulation of gene expression.
The TOR locus was independently isolated from an L.
major cosmid library under tubercidin selection and
shown to confer high levels of resistance for allopurinol
and inosine dialdehyde when expressed in L. major or
L. mexicana [230].
Finally, mutations of the target enzyme or the enzymes of purine salvage pathways could render these
compounds ineffective, especially if the purine salvage
pathways have high levels of redundancy and the activation of the prodrug does not. An L. donovani line deficient in adenosine kinase was highly resistant to the
adenosine analogues tubercidin and Formycin A but
not to the inosine analogues Formycin B and allopurinol riboside or to 6-thioguanosine [231]. Similarly, insertional mutagenesis of T. gondii followed by selection on
the toxic nucleoside adenine arabinoside led to the cloning of the Toxoplasma adenosine kinase [232].
5. Structure–activity relationships of purine transporters
5.1. Topology
The identification of transporter genes and gene families is finally opening the way for structural studies of
both mammalian and protozoan ENT-family transporters. Algorithms for the prediction of transmembrane
(TM) helices commonly predict 9–11 such domains.
To our knowledge, the topology of only one ENT transporter has been experimentally verified [233]. By using
native and engineered N-glycosylation sites in combination with immunological approaches, the human ENT1
transporter was shown to contain 11 TM domains, with
an intracellular amino terminus and extracellular carboxyl terminus. Hydrophilic loops between the helices
were short except for a relatively large glycosylated loop
between TM1 and 2, and an even larger intracellular
loop between TM6 and TM7 [233]. Extensive analysis
of the predicted amino acid sequences of many eukaryote ENT family members has shown that this basic
structure is likely to be highly conserved [233,234],
though it should be emphasised that individual models
do need to be experimentally confirmed. The presence
of a large extracellular loop is less conserved than the
central intracellular loop [234]. The structural predictions of protozoan ENT transporters [91] have been entirely consistent with this general model, originally
based on extensive analysis of the human, Drosophila
melanogaster and C. elegans ENT members [233,234].
The topology of one protozoan ENT transporter,
TbAT1, is depicted in Fig. 6(a).
5.2. Domains involved in substrate recognition
The strong topological conservation makes it probable that the basic folding of the ENT transporters is likewise mostly retained and that domains identified in one
transporter to be part of the substrate translocation
channel, or involved in substrate binding, could be relevant to ENT transporters in general. This seems to be
borne out by the identification of four conserved regions
in an alignment of >30 ENT family genes from many
different species [235], consisting of TM1 and adjacent
amino acids (I), TM4/5 (II), TM8 (III) and TM9/10
including the last intracellular loop (IV). Very limited
data exists on the protein folding or the function of each
domain. Experiments using chimeras of rat ENT1 and
ENT2 identified a fragment, stretching from between
TM4 and TM5 to the end of TM6, associated with substrate selectivity [156] and it seems likely that TM5 and/
or TM6 should therefore be part of the substrate translocation channel. This study followed an earlier report
from the same groups, using chimeric constructs of
rENT1 and hENT1, which identified a larger fragment,
incorporating TM3-6, as involved in the recognition of
the competitive nucleoside inhibitors dipyridamole and
dilazep [236]. Furthermore, SenGupta et al. [237] identified two conserved glycine residues in TM5, at positions
179 and 184 of hENT (Gly 161 and 166 of TbAT1
(Fig. 6)), and determined that G184 may have a critical
structural function, since its replacement led to poor targeting to the plasma membrane as well as complete loss
of function. Substitution of G179, on the other hand,
which is completely conserved in an alignment of all
known protozoan and human ENT transporters, lead
to an almost complete loss of transport activity while
retaining proper membrane localization [237]. Yao and
colleagues [238] also identified a cysteine residue in
rENT2 in the outer half of TM4 that was accessible to
the thiol reactive agent PCMBS, yet protected by the
presence of uridine, a permeant for this transporter.
All these studies strongly suggest that the region of
TM4-6 forms part of the substrate translocation
channel.
The importance of this domain has also been evident
from observations with the Leishmania nucleoside transporters and TbAT1, lending further support for substantial structural conservation. Vasudevan et al. [169]
isolated two different dysfunctional LdNT1.1 alleles
from the tubercidin-resistant line TUBA5. One of these
contained a single mutation, C337Y, in TM7 that conferred reduced Vmax [169]. TbAT1 from drug-resistant
field isolates contains a deletion mutation of a highly
conserved phenylalanine in TM7 [157], supporting a role
for TM7 in translocation of substrate. The other allele
displayed a single point mutation to G183, corresponding to G184 of hENT1 and G166 of TbAT1. The mutation, G183D, is located in predicted transmembrane
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
a
I
II
Y KY AQG
Y
E
I F FF P
M
S
F
I
S VT N
V
VS V
MF F G
L T F
V
YV
F I
E
N
A
S
D
F
G
L
M
III
KPDA KP
E
D
WF KP
K
H
M
F
TSY Y MV
I AA
S
L FEV FV
I
VLAL
V
S
M
L
LTL
G
P
I
G
R
R
I SV
G
G
T TE TG
T
A
V
K
I
V
TM
T
V
MI IL
V
A
A
LP IG NA
I
V M
T
L
C
D
A
G
N
A A LI A
V
S MG
A
K
I VI
S
F
S F
VV
G A V
I VW
V
S
Y
T
C
G
S
F
K
T
P
F
P
VI
VII
G GYH N ML
K
I
I
Q
A
I RS
F
Y
Y
F
V
G
A
L
V MF M YP L V
V
F T
Q SI A TF FM
C L L
V F
C F
V
A L
M W
L
P V
L
R
R
W
K H GG F
N P D V
P D D N
Y D T W
A D D V
AYKQ Q K RA M
D G
A N
P V
NGAA KG M T AT V
P
KR A AY R F EA
G
I
D
DK G AD GD EG
NH2
G
V
G
L
R
V
T
IV
VIII
IX
X
A DT
G
D
G
W
Y
P L PY
TL A TG I I LI H V
I A
S V
A G
L AN F RL C L WT FG
L D V P
Y Y
G F
I
Y
L
I
L G G
L
S
M
I
R
R
S
L
A
Q
C
F
I
L
T GIL V Y
Q
W A
FK AL HV SP R P R T
LM Q QS TT IN
E
V
Q
D PD TM K D MD
1011
XI
E GL
N
V AL
A
F
AM
G F
G IL
L
S I
T
P K C OOH
V
G
I
W
N
A A LS
R
E
G
G S L T TA
Cytoplasm
Gly161
Gly166
Gly181
b
XI
VII
F
IX
II
VIII
V
X
III
VI
IV
I
Fig. 6. (a) Predicted topology of TbAT1, partially based on hydropathy plots (TMPRED). The location of some key residues is indicated and the
trans-membrane domains are numbered. (b) Preliminary model of helix packing for TbAT1 (extracellular view). Amphipathic faces are indicated
with blue bars, the substrate translocation pathway with a blue star and the position of a solvent-accessible cysteine with a green diamond. The
extracellular loop between TM-V and TM-VI is indicated with a red line and the intracellular loop between TMs VI and VII with a dotted red line.
region 5 and led to drastically diminished Vmax. Intriguingly, the more conservative substitution of G183A, by
site directed mutagenesis, led to a mutant LdNT1.1 protein that retained the ability to transport adenosine but
no longer accepted uridine, strongly suggesting that this
glycine residue plays a critical role in substrate recognition. This observation stimulated further analysis of the
role of TM5 in LdNT1.1 function [239] by the substituted cysteine accessibility method, a scanning mutagen-
esis approach that permits identification of residues that
are exposed to substrate [240]. This recent study indicated that 3 of the 4 glycines in TM5, including G183,
were required for transport activity. These glycines are
all highly conserved within the ENT family and may
play a role in helix packing [241,242]. Those cysteine
mutants that were accessible to chemical modification
were arranged on the same helical face as those with
hydrophilic side chains and most of these residues were
1012
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
protected from modification by the presence of adenosine. These data indicate that TM5 is not only critical
for transport activity but also contributes to the pathway of substrate translocation.
5.3. A preliminary model for helix packing of TbAT1
To reveal residues and domains that may have a key
function in substrate recognition by the T. brucei transporter TbAT1, we performed helical wheel analyses for
each predicted hydrophobic transmembrane domain.
Polar residues were highlighted, revealing that helices
5, 6 and 8 have substantial amphipathic faces (defined
here as more than two stacked polar residues). The polar
residues aspartic acid, asparagine and arginine are all
stacked on one face of TM8 and are highly conserved
across the ENT family. The hydrophilic face of TM6
is also highly conserved and a body of evidence (see
above) implicates TM5 in substrate recognition. TM10
is one of the less well-conserved transmembrane domains and is generally amphipathic. Considering that
hydrophobic domains are likely to be shielded from
water and that the limited length of several of the predicted hydrophilic loops will impose packing constraints
in the tertiary structure, we generated a model of transmembrane helix packing that defines a putative substrate permeation pathway for TbAT1 (Fig. 6(b)). In
this model, the substrate permeation pathway is defined
by helices 4–8, with the most significant contribution
from the hydrophilic faces of TM5, TM6 and TM8.
The majority of predicted TM helices in TbAT1 have
substantial hydrophobic faces and these are arranged
in the model so that they are oriented toward the lipid
environment. TM10 is amphipathic, is rich in residues
and motifs that are commonly involved in helix packing
(glycine, alanine and threonine) and we have thus buried
this helix in the core region. Relatively short hydrophilic
loops connect TM9/10 and TM2/3. Therefore these helices have been placed adjacent to each other. This
arrangement gives rise to an asymmetric positioning of
the substrate permeation pathway, but makes no prediction regarding the structure or functional role of the larger hydrophilic loops that are likely to be exposed at the
membrane surface.
Comprehensive alignments of ENT proteins reveal
limited but significant sequence identity, and suggest a
conserved topology. While very preliminary, the model
is consistent with the experimental data that has been
obtained with trypanosomal, leishmanial and mammalian ENTs reviewed in the previous section. Polar residues in helices that are predicted to delimit the
substrate permeation pathway are good candidates for
site-directed mutagenesis in a reverse genetics analysis,
and this working model could direct mutagenesis studies
and aid the interpretation of mutations that are revealed
by forward genetic analysis.
5.4. Models for transporter–permeant interactions
An additional approach to studying interactions between the transporter and the permeant is by studying
the binding affinity of various (potential) substrates,
determined as Km, the concentration at which 50% of
transporters is occupied, for a range of structurally
modified analogues. For instance, the P1 transporter
of T. b. brucei bloodstream forms displays a Km of
0.41 ± 0.08 lM (n = 4 [53]) for [3H]adenosine but only
6.5 ± 1.8 lM (n = 3) for [3H]tubercidin (De Koning
et al., unpublished). Since the only difference between
adenosine and tubercidin (7-deazaadenosine) is a @N–
or a @CH– at position 7 of the purine ring, it is reasonable to suppose that a substantial loss of binding energy
would be associated with the concurrent loss of a strong
H-bond acceptor. The Gibbs free energy of binding can
be estimated from a derivation of the Nernst equation,
DG0 = RTln(Km) [53], and the difference in binding energy between adenosine and tubercidin, d(DG0), thus
gives a value in kJ/mol for the H-bond between N7 of
adenosine and one or more amino acid residues in the
substrate translocation pathway of the P1 transporter
(7.2 kJ/mol in the current example). Using this approach
carefully with a sufficient number of well-chosen analogues allows the construction of a model for all the
interactions between substrate and transporter, which
complements the mutagenesis approach in understanding transporter function.
However, the determination of Km values requires
radiolabelled substrates of high specific activity, which
are almost never available for all the various structural
analogues required for such a study. This limitation requires us to use the inhibition constant Ki instead, calculated from the Cheng–Prusoff equation: Ki = IC50/
[1 + (L/Km)], in which IC50 is the inhibitor concentration
causing 50% inhibition of the transport of a permeant,
at permeant concentration L [243], as an approximation.
This equation is only valid for competitive inhibition,
which must therefore be verified, and the Ki value is
not necessarily equal to the Km, particularly if the rates
of translocation of permeant and inhibitor are substantially different [244]. We recently reported one such
example, where the Km of [3H]diminazene for TbAT1
was determined at 0.45 ± 0.11 lM [51], whereas the Ki
had previously been determined at 2.4 ± 0.5 lM in T.
b. brucei [53] and 3.9 lM in T. equiperdum [57] – a discrepancy likely to be the result of the very different
translocation rates of adenosine and diminazene (see
Table 7). This complication could lead to an overestimate of bond energy when using the Ki rather than the
Km value in the Nernst equation given above.
While accepting the above limitations, semi-quantitative models for transporter–permeant interactions can
be constructed using transport kinetics and structural
analogues [8,22,39,40,53,245,246] and we have shown
H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
that such models have predictive value for Ki values of
potential permeants [8,39,40], which is potentially of
great importance for structure-based design of chemotherapeutics. We have already mentioned the structural
motif recognised by TbAT1/P2 (Section 4.2; Fig. 4).
Such a motif, the haptophore, can be coupled to an active substance, the toxophore, to improve translocation
rates and/or specificity [35–38]. Moreover, such motifs
can highlight functional conservation, where an amino
acid sequence would at most show probable conservation of topology and conserved domains. For instance,
we have characterised the main purine nucleobase transporters of T. b. brucei (H2) and L. major (LmaNBT1) in
great detail and found that both interact in a very similar way with natural purine nucleobases [22]. Fig. 7
shows that TbH2 and LmaNBT1 appear to make similar hydrogen bonds with hypoxanthine, through N(1)H,
N3, N7 and N(9)H. However, the two protozoan nucle-
TbH2
12.6
H
6. Concluding remarks
N
H
N
7.7
8.3
O
HO
ee
O
O
H
8.9
H2N
N
H
N
H
12.5
N
rR
HN
e
10.6
7.0
O
NH2
3.1
H2N
5.8
O
H
In the last decade, our understanding of protozoan
nucleoside and nucleobase transporters has increased
tremendously. We have moved from establishing the
presence and number of such transporters in various
organisms through characterisation of their kinetic
properties to characterisation of their genes and finally
towards understanding their structure and mechanism
of action. We thus move that much closer to using this
knowledge to selectively target drugs to these pathogens,
or, in isolated cases, validating the transporters themselves as drug targets. The study of protozoan transporters is thus finally completing its voyage from biology to
therapy. The next decade should see the fulfilment of the
promise for a purine- or pyrimidine-based therapy for at
least some protozoan infections.
O
N
HN
hFNT1
obase transporters form very different interactions with
oxopurines than the human FNT1 nucleobase transporter, except for a similar H-bond at N3 (Fig. 7) [39],
leading to different substrate selectivity such as a 50-fold
higher affinity for 3-deazaguanine at the protozoan
transporters [22,39]. The genes encoding LmaNBT1
and TbH2 have not been identified with certainty, but
it is highly likely that both will be ENT family members
and that identity at amino acid level will be around 50%
(as it is between TbNBT1 and LmNT3, both very similar
hypoxanthine transporters to H2 and LmaNBT1,
respectively [21,32]). This is considered a high level of
conservation, yet a single amino acid change can completely change a transporterÕs function or selectivity
(see previous section) and it must be concluded that at
the current state of knowledge primary sequence data
is a poor guide to transporter function, beyond a rough
classification as Ôprobable nucleoside or nucleobase
transporterÕ. The substrate binding model depicted in
Fig. 7, however, does have predictive qualities and can
be constructed even in the absence of genetic data.
LmaNBT1 and TbH2 were termed Ôfunctional homologuesÕ to distinguish them from genetic homologues
[22].
12.5
N
HN
H2N
LmNBT1
HO
O
O
1013
~10
N
H
N
7.3
H
Fig. 7. Model of interactions between various nucleobase transporters
and guanine [22,39]. Numbers represent estimates of bond strength in
kJ/mol. The shape of the binding sites and any functional groups
thereof are entirely speculative and for illustration purposes only. The
double line around the 2-position amine in TbH2 and LmaNBT1
depicts suspected steric hindrance.
Acknowledgement
The nucleotide transporter phylogeny was supplied
by Janssen Genomics, http://www.janssen-genomics.com.
Appendix A. Supplementary data
Supplementary data associated with this article can
be found, in the online version, at doi:10.1016/
j.femsre.2005.03.004.
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H.P. de Koning et al. / FEMS Microbiology Reviews 29 (2005) 987–1020
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