The revised Trypanosoma cruzi subspecific nomenclature: Rationale,
epidemiological relevance and research applications
Bianca Zingalesa,⇑, Michael A. Milesb, David A. Campbellc, Michel Tibayrencd, Andrea M. Macedoe,
Marta M.G. Teixeiraf, Alejandro G. Schijmang, Martin S. Llewellynb, Eliane Lages-Silvah,
Carlos R. Machadoe, Sonia G. Andradei, Nancy R. Sturmc
aDepartamento de Bioquímica, Instituto de Química, Universidade de São Paulo, Avenida Professor Lineu Prestes 748, 05508-000 São Paulo, SP, Brazil
bThe London School of Hygiene and Tropical Medicine, Keppel Street, London WC1E 7HT, UK
cDepartment of Microbiology, Immunology & Molecular Genetics, David Geffen School of Medicine, University of California at Los Angeles, 10833 Le Conte Ave, Los Angeles,
CA 90095-7065, USA
dMaladies Infectieuses et Vecteurs Ecologie, Génétique, Evolution etContrôle, MIVEGEC/IDVEGEC, UM1-CNRS 5290-IRD 224, IRD Center, BP 64501, 34394 Montpellier Cedex 5, France
eDepartamento de Bioquímica e Imunologia, Instituto de Ciências Biológicas, Universidade Federal de Minas Gerais, CP 486, 30161-970 Belo Horizonte, MG, Brazil
fDepartamento de Parasitologia, Instituto de Biociências, Universidade de São Paulo, Avenida Professor Lineu Prestes 1374, 05508-000 São Paulo, SP, Brazil
gLaboratorio de Biología Molecular de la Enfermedad de Chagas, INGEBI-CONICET, Vuelto Obligado 2490, Buenos Aires 1428, Argentina
hDepartamento de Ciências Biológicas, Universidade Federal do Triângulo Mineiro, Rua Frei Paulino 30, Uberaba, MG, Brazil
iCentro de Pesquisas Gonçalo Moniz, Fundação Oswaldo Cruz, Rua Waldemar Falcão 121, 40295-001 Salvador, Brazil
a r t i c l ei n f o
Received 31 October 2011
Accepted 16 December 2011
Available online 27 December 2011
Trypanosoma cruzi strains
Discrete typing unit
a b s t r a c t
The protozoan Trypanosoma cruzi, its mammalian reservoirs, and vectors have existed in nature for mil-
lions of years. The human infection, named Chagas disease, is a major public health problem for Latin
America. T. cruzi is genetically highly diverse and the understanding of the population structure of this
parasite is critical because of the links to transmission cycles and disease. At present, T. cruzi is parti-
tioned into six discrete typing units (DTUs), TcI–TcVI. Here we focus on the current status of taxon-
omy-related areas such as population structure, phylogeographical and eco-epidemiological features,
and the correlation of DTU with natural and experimental infection. We also summarize methods for
DTU genotyping, available for widespread use in endemic areas. For the immediate future multilocus
sequence typing is likely to be the gold standard for population studies. We conclude that greater
advances in our knowledge on pathogenic and epidemiological features of these parasites are expected
in the coming decade through the comparative analysis of the genomes from isolates of various DTUs.
? 2012 Elsevier B.V. All rights reserved.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241
The concept of discrete typing unit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241
2.1.The clonal model of evolution in T. cruzi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241
2.2.Discrete typing units and clonets. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241
Two major models for the origin of hybrid DTUs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 242
Phylogeography of the DTUs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244
4.1.TcI and its extensive genetic diversity. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244
4.2.TcIII and TcIV. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245
4.3.TcII, TcV and TcVI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245
4.4. An enigmatic T. cruzi genotype from bats (Tcbat). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245
1567-1348/$ - see front matter ? 2012 Elsevier B.V. All rights reserved.
Abbreviations: ITS1 rDNA, internal transcribed spacer 1 of rDNA; MLEE, multilocus enzyme electrophoresis; MLST, multilocus sequence typing; NTS, non-transcribed
spacer; RAPD, randomly amplified polymorphic DNA; RFLP, restriction fragment length polymorphism; SNPs, single-nucleotide polymorphisms; SL, spliced leader; SL-IR,
spliced leader intergenic sequence.
⇑Corresponding author. Tel.: +55 11 30912686; fax: +55 11 38155579.
E-mail address: email@example.com (B. Zingales).
Infection, Genetics and Evolution 12 (2012) 240–253
Contents lists available at SciVerse ScienceDirect
Infection, Genetics and Evolution
journal homepage: www.elsevier.com/locate/meegid
Standardizing genotyping for identification of the six T. cruzi DTUs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246
Comparative experimental pathology of the DTUs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246
T. cruzi DTUs and human Chagas disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248
7.1. Clinical presentations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248
7.2. Acute Chagas disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248
7.3. Chronic Chagas disease. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248
7.4.Chagas disease reactivation due to immunosuppression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249
T. cruzi genomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 249
Concluding remarks and perspectives. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 250
Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 250
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 250
Infection with Trypanosoma cruzi is a complex zoonosis, trans-
mitted by many hematophagous triatomine species and sustained
byover 70genera of mammalianreservoirhosts. T. cruzi hasa broad
endemic range that extends from the Southern United States to
Argentinean Patagonia. The human infection, named Chagas dis-
ease in recognition of Carlos Chagas who first discovered American
trypanosomiasis in 1909, is found mostly in South and Central
America, primarily affects poor rural populations, and is considered
to be the most important parasitic infection in Latin America with
serious consequences for public health and national economies.
The spectrum of pathological outcomes associated with acute
and chronic Chagas disease ranges from subclinical infection
through the cardiac and digestive syndromes to death. Specific out-
comes may be determined by a variety of non-exclusive factors
including parasite genetics, host genetics, mixed infections, and
cultural and geographical factors (Macedo et al., 2002, 2004; Bus-
caglia and Di Noia, 2003; Campbell et al., 2004).
The diversity of the T. cruzi genome and multiplicity of its geno-
types and phenotypes is well recognized (Dvorak et al., 1982; Bar-
nabé et al., 2000; Brisse et al., 2000; Devera et al., 2003; Lewis et al.,
2009a). Designation of ecologically and epidemiologically relevant
groups for T. cruzi has oscillated between a few discrete groups
(Miles and Cibulskis, 1986; Souto and Zingales, 1993; Souto
et al., 1996; Zingales et al., 1999) and many (Tibayrenc and Ayala,
1988). Currently, six discrete typing units (DTUs) are assigned
(Brisse et al., 2000). In 2009, these DTUs were renamed by consen-
sus as TcI–TcVI (Zingales et al., 2009). Several reviews already de-
scribe how these DTUs correspond with former nomenclatures and
with prospective biological and host associations (Campbell et al.,
2004; Miles et al., 2009; Sturm and Campbell, 2010; Zingales
et al., 2009).
The aim of this review is to explain further the rationale for
naming TcI–TcVI, with reference to their known molecular genet-
ics, eco-epidemiological features and pathogenicity. We also sum-
marize methods for DTU genotyping, and discuss a possible
seventh T. cruzi branch, provisionally named Tcbat. An understand-
ing of the T. cruzi DTUs and their epidemiological implications will
provide new insights to guide research and future interventions
against this devastating infectious disease.
2. The concept of discrete typing unit
Since the late 1970s, T. cruzi has become one of the models for
molecular epidemiologists and population geneticists, and conse-
quently this protozoan parasite is a pathogenic agent for which
evolution and population structure are among the best studied,
although not necessarily the best understood. The emerging pic-
ture is that of a typical pattern of reticulate evolution, similar to
that of many plant species (Avise, 2004).
The concepts of DTUs and clonal evolution have been designed
within the framework of evolutionary research on T. cruzi. Tibay-
renc and co-workers devised descriptive concepts and terminology
to make such research and its implications accessible to non-spe-
cialists, including medical professionals and epidemiologists, and
to bypass certain demands of classical evolutionary biology defini-
tions. Classical cladistic and population genetics approaches
imperfectly depict the biological realities of the evolution of path-
2.1. The clonal model of evolution in T. cruzi
In the framework of this model, a ‘‘clonal species’’ refers to all
cases where descendant multilocus genotypes are virtually identi-
cal to the founding genotype. The main parameter focused on in
this scenario is the inhibition of genetic recombination. The term
‘‘clone’’ in this context refers to the population structure of the spe-
cies under study, not to its precise mating system. Different meth-
ods of propagation can generate genetic clones, including classical
cell division, several cases of parthenogenesis and gynogenesis.
Following this definition (Tibayrenc et al., 1990), selfing and ex-
treme inbreeding are not alternative hypotheses to clonality (Rou-
geron et al., 2009), but rather a particular case of it. Selfing refers to
mating between identical genotypes, which can be issued from the
same clone (Tibayrenc et al., 2010). Extreme inbreeding refers to
mating between extremely similar genotypes. The result is a lack
or extreme limitation of genetic recombination, hence genetic
Stating that T. cruzi is a basically clonal species means neither
that recombination is totally absent in the parasite’s natural pop-
ulations, nor that it does not have an impact on the evolutionary
scale, but rather that it is too rare to break the prevalent pattern
of clonality. The potential for genetic exchange is still present
(Gaunt et al., 2003). Moreover, some localized transmission cycles
suggest that genetic recombination does occur within DTUs of
T. cruzi (Carranza et al., 2009; Ocaña-Mayorga et al., 2010). The
possibility of limited genetic exchange between DTUs is also under
debate (Lewis et al., 2011). However, the species T. cruzi consid-
ered as a whole shows all the signs for a typical clonal population
structure: departures from panmictic expectations, strong linkage
disequilibrium (non-random association of genotypes at different
loci) within and especially between DTUs, and division into
discrete genetic clusters (see DTUs, below).
2.2. Discrete typing units and clonets
Often the genetic subdivisions identified by evolutionary stud-
ies in pathogen species do not fulfill the criteria demanded by rig-
orous cladistic analysis. The main reason is that even in
predominantly clonal species such as T. cruzi there is a certain
amount of genetic recombination that clouds the distinction of
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
phylogenetic subdivisions. By definition, a clade represents an evo-
lutionary unit that is strictly isolated from other evolutionary
units. Moreover, some pathogen lineages may have a hybrid origin:
in other words, two ancestors. This is the case of several T. cruzi ge-
netic subdivisions (Sturm and Campbell, 2010). Thus the clade con-
cept is not applicable in the case of T. cruzi.
However, even when some genetic exchange occurs, discrete
and stable subdivisions can be identified reliably in many in-
stances. Thus the term ‘‘discrete typing unit’’ (Tibayrenc, 1998)
was proposed to describe sets of stocks that are genetically more
similar to each other than to any other stock, and are identifiable
by common molecular markers sometimes referred to as ‘‘tags’’.
The DTUs constitute reliable units for analysis for molecular epide-
miology and experimental studies of evolution. Genetic clusters
within T. cruzi perfectly fit this definition, and thus the DTUs TcI–
TcVI have been assigned (Brisse et al., 2000; Zingales et al., 2009).
Within these DTUs T. cruzi stocks that share profiles for a given
panel of molecular markers are not necessarily genetically identi-
cal, and can often be distinguished with additional markers. Thus
strains within DTUs should be considered as families of closely re-
lated clones, not as a single clone. Tibayrenc and Ayala (1991)
coined the term ‘clonet’ to refer to sets of stocks that appear to
be indistinguishable with a given set of genetic markers in a basi-
cally clonal species, such as T. cruzi. The clonets are relevant units
of analysis for molecular epidemiology, for example, with multilo-
cus sequence types (MLSTs) or karoytypes derived from pulsed-
field gel electrophoresis. However, it is crucial to keep in mind that
the most recent common ancestor of a given clonet can be either a
few weeks or hundreds of years old, depending on the marker’s
power of resolution and rate of evolutionary change (molecular
clock). The latter parameter cannot be known a priori and may
have considerable epidemiological relevance.
3. Two major models for the origin of hybrid DTUs
T. cruzi is predominantly diploid (El-Sayed et al., 2005) and the
known cell replication method is binary fission, i.e. it is an asexual
process. Under the clonal model (above) new DTUs evolve with the
accumulation of discrete mutations, unaffected by rare events of
genetic exchange. However, consistent with the caveat that some
genetic exchange events may occur, evidence for T. cruzi heterozy-
gosity in nature emerged through the study of individual genes
(Chapman et al., 1984; Bogliolo et al., 1996; Carrasco et al., 1996;
Souto et al., 1996; Brisse et al., 1998). The pronounced heterozy-
gosity observed in natural isolates of TcV and TcVI suggested that
these DTUs are hybrids and derived from TcII and TcIII (Sturm
et al., 2003; Sturm and Campbell, 2010). The remaining DTUs,
TcI, TcII, TcIII, TcIV (and Tcbat, see below) show substantial allelic
homozygosity. While the tenet of the clonal theory may still ex-
plain the common mode of T. cruzi population expansion, newer
models incorporate hybridization events to explain the extant pop-
ulation structure that includes hybrid DTUs (Westenberger et al.,
2005; Freitas et al., 2006).
The ‘Two-Hybridization’ model (Westenberger et al., 2005) and
the ‘Three Ancestor’ model (Freitas et al., 2006) both incorporate
two hybridization events (Fig. 1). In the Three Ancestor model
the two recent genetic exchange events between TcII and TcIII
yield TcV and TcVI. The Two-Hybridization model invokes one an-
cient genetic exchange event between TcI and TcII, with loss of het-
erozygosity among progeny to produce TcIII and TcIV, followed by
a second more recent hybridization event between TcII and TcIII to
yield both TcV and TcVI.
Analysis of single-nucleotide polymorphisms (SNPs) among the
six DTUs by multilocus sequence typing (MLST) revealed four
rather than six distinct DNA sequence classes, termed haplogroups
(Machado and Ayala, 2001; Sturm et al., 2003; Broutin et al., 2006).
Two of the four haplogroups were always present in TcV and TcVI,
confirming the predominantly heterozygous nature of their alleles.
Despite being similar by most standards TcV and TcVI are distin-
guishable by isoenzyme electrophoresis (Chapman et al., 1984;
Barnabé et al., 2000), ribosomal RNA markers (Souto et al., 1996),
restriction fragment length polymorphism (RFLP) assays (Rozas
et al., 2008), some MLST markers (Yeo et al., 2011), and microsat-
ellite analysis (Lewis et al., 2011). The nucleotide patterns in each
TcV/TcVI haplogroup closely resemble TcII and TcIII alleles (Mach-
ado and Ayala, 2001; Brisse et al., 2003; Westenberger et al., 2005;
Freitas et al., 2006; Yeo et al., 2011; Lewis et al., 2011), confirming
that TcII and TcIII as the most likely parental types of TcV and TcVI.
However, different MLST data indicated that the TcIII parental cells
included characters derived from TcI and TcII (Sturm et al., 2003;
Elias et al., 2005; Tomazi et al., 2009). Thus, TcIII, as well as TcV
and TcVI, could be the product of a hybridization event (Westen-
berger et al., 2005; Ienne et al., 2010).
The major difference between the Two Hybridization and the
Three Ancestor models is therefore whether TcV and TcVI are prog-
eny from a single hybridization event incorporating TcI alleles ac-
quired via TcIII (Westenberger et al., 2005) or progeny of two
Fig. 1. Comparison of (A) the Two-Hybridization and (B) the Three Ancestor models for the roles of genetic exchange during the clonal evolution of T. cruzi. Rectangles
indicate the distinct DTUs. Fusion of two cells and genetic exchange is indicated by the ovals, with parental contribution indicated by the red arrows. The mitochondrial clades
are shown by fill colors: Blue = clade A; Green = clade B; orange = clade C.
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
hybridization events excluding TcI (Freitas et al., 2006) (Fig. 1). The
participation of TcI in the generation of the extant heterozygous
lines is supported by maxicircle sequences (Westenberger et al.,
2006a; Ruvalcaba-Trejo and Sturm, 2011), MLST data (Tomazi
et al., 2009), RFLP data (Rozas et al., 2008), and 195-bp satellite
DNA sequences and distribution (Elias et al., 2005; Ienne et al.,
2010). Conversely, maxicircle-encoded genes cytochrome oxidase
subunit II and NADH dehydrogenase subunit 1 (Machado and Aya-
la, 2001; Freitas et al., 2006), microsatellite analyses (Freitas et al.,
2006) and some other nuclear markers (Machado and Ayala, 2002;
Rozas et al., 2007) did not detect participation of TcI. The compar-
ison of the new genome sequence of the Sylvio X10/1 strain (Fran-
zen et al., 2011), representative of TcI, with CL Brener, a TcVI hybrid
that encompasses both TcII and TcIII genomes (El-Sayed et al.,
2005), has confirmed that TcIII has an average higher genetic sim-
ilarity at the genome sequence level with TcI than TcII. This obser-
vation is not inconsistent with the previous conclusion that TcIII
may be the product of an ancient hybridization between TcI and
TcII, as depicted in the Two Hybridization model of Fig. 1 (Elias
et al., 2005; Westenberger et al., 2005). However, the possibility
Fig. 2. Approximate geographical distribution of T. cruzi DTUs in domestic and silvatic transmission cycles.
Summary of ecotope, host, vector and disease associations of T. cruzi DTUs.a
Genotype Ecotope/nicheSilvatic hostsSilvatic
palms (e.g Attalea),
rocky; terrestrial in
Primary: arboreal, semi-arboreal;
especially Didelphis, other didelphids,
arboreal rodents, primates, Tamandua
Secondary: terrestrial rodents
South, Central and
North of the Amazon, sporadic in
known; rare in
Atlantic forest primates, didephids,
Armadillos, especially Dasypus,
Primates, Nasua nasua,
sporadic further North
Atlantic and Central Brazil.
Rare in humans (also domestic dogs).
Acute cases in Amazonian Brazil. Clinical
presentation poorly known
Secondary cause of Chagas disease in
Venezuela, sporadic elsewhere in South
TcIV Arboreal, and some
terrestrial hosts in
Rare in silvatic cycles
North and South
TcV Incompletely known: Dasypus, Euphractus,
Southern Cone, greater
Gran Chaco, extreme
South of Brazil
Southern Cone, greater
TcVI Rare in silvatic cyclesIncompletely knownIncompletely
aEcotope host and vector associations are not exclusive.
bSee map in Fig. 2.
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
that characteristics shared by TcI and TcIII are ancestral states that
have diverged in TcII may need further consideration.
4. Phylogeography of the DTUs
Setting aside the theoretical origins of the DTUs, divergent geo-
graphical and biological characteristics are apparent, as is their rel-
evance to understanding of the eco-epidemiology of Chagas
disease (Fig. 2).
It is not surprising that, given the current level of sampling, the
ecological history for all T. cruzi DTUs cannot yet be fully discerned.
Relationships have been obscured by massive changes, from mam-
mal migrations between the Americas, to climate induced retrac-
tion and expansion of habitats, and dramatic recent habitat
destruction and urbanization by humans. A summary of ecotope,
host, and vector associations of T. cruzi DTUs is given in Table 1,
as has been extensively reviewed elsewhere (Miles et al., 2009).
Here we will provide primarily more information on the phyloge-
ography and the extensive genetic diversity of TcI, for which there
has been recent rapid knowledge progress. In terms of propensity
to cause severe Chagas disease, all six DTUs are known to be infec-
tive to humans, and clinical aspects are described in more detail in
4.1. TcI and its extensive genetic diversity
TcI is the most abundant and widely dispersed of all the T. cruzi
DTUs in the Americas. It is found throughout the range of triato-
mine vector distribution, and can be associated with silvatic and
domestic cycles. Human infection with TcI is concentrated in the
north of South and Central America, and is associated with chaga-
sic cardiomyopathy. There are only disparate reports of infection
and disease south of the Amazon basin. Wild TcI isolates exist from
Alabama in the United States (Roellig et al., 2008) at 32? North, to
Limari, Chile (Apt et al., 1987) at 30? South. Review of the literature
reveals 52 mammalian genera naturally infected with this DTU,
with representatives from Marsupialia, Rodentia, Primata, Chirop-
tera, Xenartha, Carnivora, and Artiodactyla in order of abundance,
as well as all major genera of triatomine bugs (Llewellyn, unpub-
lished records, updated 05/01/2010). The divergence date between
TcI and TcII is ill defined, estimated between 88 and 37 million
years ago, based on small subunit rDNA (Briones et al., 1999;
Kawashita et al., 2001) and between 16 and 3 million years ago,
based on dihydrofolate reductase-thymidylate synthase and trypa-
nothione reductase genes (Machado and Ayala, 2001). Unsurpris-
ingly for a parasite so ancient and dispersed, significant genetic
diversity has accumulated within TcI. The earliest recognition of
TcI heterogeneity is manifest in the isoenzyme clonet typing
scheme proposed by Tibayrenc and co-workers whereby 25 geno-
types are assigned to TcI, with a much lower number assigned to
any other of the other five DTUs (Tibayrenc et al., 1986). Sample
size may have represented an early confounder, and we now know
that substantial diversity has also accumulated within other
lineages (Machado and Ayala, 2001; Westenberger et al., 2006b;
Llewellyn et al., 2009a; Marcili et al., 2009a). Access to new,
high-resolution genotyping techniques has seen a resurgence of
interest in the delineation of TcI intra-DTU diversity.
Saravia et al. (1987) examined genetic diversity among 54
Colombian T. cruzi isolates collected from silvatic and domestic
localities at several foci in Meta, Casanare and Cundinamarca prov-
inces using 13 isoenzyme markers. Of those isolates examined,
most were TcI (‘Z1-like’). Furthermore, among TcI isolates, marked
genetic subdivision was observed between strains from domestic
and silvatic transmission cycles, largely independent of geographic
origin. This biological observation is now supported by spliced
leader (SL, also known as mini-exon) intergenic region (SL-IR) se-
quence data from western Colombia (Herrera et al., 2007, 2009;
Falla et al., 2009). Several other molecular methods have been ap-
plied to study TcI heterogeneity in Colombia, including molecular
karyotypic analysis (Triana et al., 2006), SL probe hybridization
(Triana et al., 2006) and minicircle random RFLP (Jaramillo et al.,
1999). More recently, low-stringency single primer PCR (Rodriguez
et al., 2009) provided important insight into the dynamics of TcI
transmission among several communities in the Sierra Nevada de
Santa Marta. Outside Colombia, multilocus enzyme electrophoresis
(MLEE) and randomly amplified polymorphic DNA (RAPD) were
used to reveal putative hybrid and parental strains at a focus of
silvatic TcI transmission in Carajás, Pará State, Brazil (Carrasco
et al., 1996). Using the same parental strains, an extant capacity
for genetic exchange was demonstrated in vitro (Gaunt et al.,
2003). Polymorphic microsatellites can resolve T. cruzi inter-spe-
cific variability (Oliveira et al., 1998), and Llewellyn et al. (2009a)
demonstrated their efficiency in revealing TcI intra-DTU diversity
at a continental scale. Crucially, the use of a multilocus typing sys-
tem permits inference of linkage disequilibrium between markers
within parasite populations and the extent of clonal vs. sexual
reproduction. Thus, when the same markers were employed to
analyze the molecular epidemiology of TcI transmission at re-
stricted geographic foci in Ecuador (Ocaña-Mayorga et al., 2010),
the first population genetic evidence for genetic exchange in TcI
was uncovered among isolates not subdivided in space or time.
The central observation of almost all analyses of TcI diversity in
northern South America is the apparent subdivision between
domestic and silvatic cycles of transmission (Saravia et al., 1987;
Herrera et al., 2007; Falla et al., 2009; Llewellyn et al., 2009a,b;
Ocaña-Mayorga et al., 2010). However, compiling data from these
studies is frustrated by constraints of the different genotyping
techniques employed. Certain types of population genetic data,
especially those derived from microsatellites and isoenzyme loci,
are difficult to standardize between studies. Sequence data, on
the other hand, are a more versatile population genetic currency.
Cura et al. (2010) demonstrated this in an ambitious multi-centric
study using the SL-IR, and corrected the erroneous use of the term
‘haplotype’ in the context of these data (Cura et al., 2010). On the
basis of this dataset, the authors delineated several discrete TcI
groups, some widely dispersed, as well as instances of mixed infec-
tions of genotypes in humans and vectors.
Whilst the SL-IR represents an accessible marker as it is diverse,
easy to amplify directly from biological samples, and straightfor-
ward to sequence with no internal primers required, there are sev-
eral limitations associated with its use. First, it is a multicopy gene.
Non-identical copies are tandemly repeated hundreds of times
throughout the T. cruzi genome, and orthology between samples
is impossible to ascertain. Second, polymorphic microsatellites lo-
cated at the 50end introduce numerous ambiguous alignments,
with an adverse effect on phylogenetic stability (Tomasini et al.,
2011). Third, significant insertions and/or deletions (indels) in this
region with respect to other T. cruzi DTUs (Souto et al., 1996) pro-
hibit the identification of a suitable outgroup. Perhaps the most
important criticism, however, is not intrinsic to the SL-IR per se.
Gene trees are not genome trees. The use of a single genetic locus
to describe genetic diversity in an organism limits conclusions that
can be drawn, especially in the context of genetic recombination,
which may occur not only in TcI (Ocaña-Mayorga et al., 2010)
but also other DTUs.
Typing strategies must be improved and standardized if further
progress is to be made. Sequence data are the ideal genotypic for-
mat for swift comparison. However, new, low copy number, highly
discriminatory markers must be identified, aided by the publication
of the Sylvio X10/1 genome (Franzen et al., 2011). Furthermore, the
T. cruzi mitochondrial genome, used for bar-coding so many other
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
species, should not be ignored (Machado and Ayala, 2001; Freitas
et al., 2006; Spotorno et al., 2008; Carranza et al., 2009), although
some incongruence between nuclear and mitochondrial markers
is likely due to introgression events. Accordingly, and if intra-DTU
genetic recombination is common, the delineation of fixed genetic
groups within TcI represents at best a distraction. Studies targeted
at TcI diversity must be designed with a specific biological or epide-
miologicalhypothesisin mind, not undertakenmerely to categorize
diversity for its own sake. Irrespective of the genotyping system in-
volved, studies like those of Saravia et al. (1987), Falla et al. (2009),
Rodriguez et al. (2009) and Ocaña-Mayorga et al. (2010) all provide
significant insight into the epidemiology of local transmission. Thus
their conclusions inform future interventions to the benefit of the
4.2. TcIII and TcIV
TcIII is mostly associated with the silvatic cycle in Brazil and
adjacent countries, and documented human infections are rare.
Silvatic TcIII is associated with the terrestrial niche and with Dasy-
pus novemcinctus, over a vast range from western Venezuela to the
Argentine Chaco (Llewellyn et al., 2009a; Marcili et al., 2009a). TcIII
is also isolated occasionally from domestic dogs (Chapman et al.,
1984; Cardinal et al., 2008).
TcIV, shows a similar pattern of distribution in South America to
TcIII, with the exception of the Chaco, where it appears to be ab-
sent. Unlike TcIII, TcIV occurs fairly frequently in humans and is
a secondary cause of Chagas disease in Venezuela (Miles et al.,
1981). Five new isolates of TcIV from primates and eight from
Rhodnius brethesi in the Amazon basin were recovered (Marcili et
al., 2009b), confirming earlier indications (Yeo et al., 2005) that
TcIV can have an arboreal ecotope. Evidence is accumulating that
TcIV is split into distinct South and North American lineages (Lewis
et al., 2009b; Marcili et al., 2009b). Further research is required to
understand the history of TcIV and these complex ecological
4.3. TcII, TcV and TcVI
TcII is found predominantly in the southern and central regions
of South America, but its true extent is not yet clear. Within its
main geographic distribution TcII is associated with cardiac mani-
festations, and concomitant megaesophagus and megacolon may
be present. It has been isolated mostly from domestic transmission
cycles. The natural hosts and vectors of TcII have proven elusive
and most of the reported isolations have been made in remaining
fragments of the Atlantic forest of Brazil, from primates and spo-
radically from other mammal species (Fernandes et al., 1999; Zin-
gales et al., 1999; Lisboa et al., 2007).
TcV and TcVI are two similar hybrid DTUs associated with Cha-
gas disease in southern and central South America. Even more so
than TcII, TcV and VI are virtually unknown as silvatic isolates.
Comparative molecular genetics have proven that TcV and TcVI
are hybrids of TcII and TcIII (see above). Until recently genetic
markers have not been of sufficient resolution to determine firstly,
whether TcV and TcVI are the products of independent hybridiza-
tion events (Freitas et al., 2006) or a single hybridization event fol-
lowed byclonal divergence,
hybridization(s) were evolutionarily ancient (Tibayrenc and Ayala,
2002; Brisse et al., 2003) or recent events (Machado and Ayala,
2001; Westenberger et al., 2005). Two recent studies have at-
tempted to address these issues: Flores-López and Machado
(2011) analyzed the evolution of 31 nuclear genes to show the
hybridization events occurred less than 1 million years ago, con-
cluding a single event prior to arrival of humans in the Americas.
Lewiset al. (2011) analyzed
higher resolution maxicircle
sequences and multiple microsatellite loci and found evidence
for two independent events dated to within the last 100,000 years,
concluding that hybridization may conceivably have occurred as a
result of human activities. TcII and TcIII may have met and hybrid-
ized as co-infections emerged in humans, peridomestic mammals,
or domestic Triatoma infestans.
Nevertheless, understanding of the ecology of TcII, V and VI is as
yet vulnerable to the limited sampling of silvatic hosts and vectors.
The paradigms may change. It has been suggested that TcII, V and
VI are more widespread geographically than currently understood
and might be found much further North (Zafra et al., 2008). If the
known TcV and TcVI hybrids are also found in Central and North
America it will most likely imply recent migration with humans
or other carriers; if genetically distinct TcII/TcIII hybrids are ob-
served, hybridization may be an ongoing phenomenon where such
mixed infections occur. Indeed in some endemic areas, notably
parts of Bolivia, mixed DTU infections are common. Microsatellite
analysis reveals that even within a single mammal, there may be
a remarkable range of mixed genotypes (Llewellyn et al., 2011).
4.4. An enigmatic T. cruzi genotype from bats (Tcbat)
Silvatic cycles of T. cruzi transmission are numerous and com-
plex. DTUs circulate in relatively independent cycles with particu-
lar ecological niches and preferentially or opportunistically
determined mammals and vectors. However, members of the same
DTU can infect mammals of distinct species and orders, indicating
that host-switching may be common among sympatric hosts
(Gaunt and Miles, 2000; Yeo et al., 2005; Marcili et al., 2009a,b;
Miles et al., 2009).
Several species of the genus Trypanosoma occur in species of
Chiroptera throughout the world, with more than 30 trypanosome
species recorded from more than 100 species of bats. Insectivorous
bat species are infected more frequently and can harbor stercorar-
ian (subgenera Herpetosoma, Schizotrypanum and Megatrypanum)
and salivarian (Trypanosoma evansi of the subgenus Trypanozoon)
trypanosomes. An extensive summary of the prevalence of bat try-
panosomes worldwide is available (Cavazzana et al., 2010). The
strong association between bats and all Schizotrypanum spp. except
T. cruzi suggests a long shared evolutionary history. However, the
evolutionary processes that have led to the current phylogenetic
structure of Schizotrypanum trypanosomes are understood poorly.
Phylogenetic studies of chiropteran stercorarian species can en-
hance understanding of host-parasite interactions and reconstruc-
tion of T. cruzi evolutionary history (Stevens et al., 2001; Barnabé
et al., 2003; Cavazzana et al., 2010).
Brazilian bats infected with T. cruzi are reported from the Ama-
zonian rainforest to urban areas of Central, Northeast and South-
east Brazil (references cited in Marcili et al., 2009c). To date,
most in vitro adapted isolates from bats belong to the subgenus
Schizotrypanum. Identification of T. cruzi from bats requires careful
analysis; Schizotrypanum species are morphologically indistin-
guishable and generically named as T. cruzi-like. However, T. cruzi
can be confirmed by the ability to infect mice. Since T. cruzi isolates
from wild mammals may induce very low parasitemias in mice, as
is the case for bat isolates, infections must be evaluated using
immunocompromized mice and sensitive parasitological methods
such as PCR and multiple haemocultures.
Analysis of SSU rDNA, gGAPDH and cytochrome b sequences al-
lows separation of T. cruzi from other trypanosomes infecting Bra-
zilian bats, including Trypanosoma cruzi-marinkellei, Trypanosoma
dionisii-like and Trypanosoma rangeli (Maia da Silva et al., 2009;
Marcili et al., 2009c; Cavazzana et al., 2010). Traditional genotyp-
ing methods based on the SL (Fernandes et al., 2001) and LSU rDNA
(Souto et al., 1996) markers placed four bat isolates from Amazonia
within TcI. However, 11 bat isolates from other Brazilian regions
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
yielded a new combination of genotypes, with a TcII-SL pattern and
a novel LSU rDNA product (Marcili et al., 2009c). This group of iso-
lates earned the provisional title of ‘Tcbat’ and awaits further char-
acterization for definitive DTU assignment (Marcili et al., 2009c),
potentially as a seventh DTU: TcVII.
Tcbat is distinguished from the six DTUs by PCR-RFLP analysis
of the internal transcribed spacer 1 of rDNA (ITS1 rDNA) (Marcili
et al., 2009c). The method of fluorescent fragment length barcod-
ing, developed to identify species of trypanosomes on the basis
of polymorphisms of regions of the rDNA locus, when applied to
T. cruzi DTUs showed a unique barcoding pattern for Tcbat (Ham-
ilton et al., 2011). Karyotype (Marcili et al., 2009c) and sequence
analyses of SL gene repeats (D.A.C. and N.R.S., unpublished results)
corroborated that Tcbat diverges from the known DTUs. All phylo-
genetic analyses using sets of molecular markers point to the
placement of Tcbat in a distinct cluster, closer to TcI, but clearly
separated from clusters comprising all the other DTUs (Marcili
et al., 2009c; Cavazzana et al., 2010). MLST analysis corroborates
some affinity of Tcbat to TcI (Teixeira and Yeo, unpublished
Tcbat develops within mammalian cells in vitro, similar to other
T. cruzi DTUs (Marcili et al., 2009c). Unlike isolates of the six DTUs,
Tcbat does not develop in the commonly available triatomine spe-
cies reared in laboratory colonies: T. infestans, Rhodnius prolixus
and Panstrongylus megistus. The Tcbat insect vector is unknown.
Possible vectors are triatomine species encountered in bat refuges,
or cimicids, vectors of T. dionisii in Europe, or bat ectoparasites
(Cavazzana et al., 2010).
Some other T. cruzi isolates also display unusually complex
combinations of molecular markers (Lewis et al., 2009b; Marcili
et al., 2009a,b,c). Thus, Tcbat is but one indicator that the complex-
ity of T. cruzi is higher than currently defined, and will require revi-
sion of DTU relationships as more silvatic isolates are genotyped.
Ideally, criteria for establishing new DTUs could be considered,
for example using MLST data and a specified degree of divergence
from existing DTUs.
5. Standardizing genotyping for identification of the six T. cruzi
The standardized nomenclature for the six T. cruzi DTUs will im-
prove scientific communication and guide future research on com-
parative epidemiology and pathology. To achieve this aim a
straightforward and reproducible genotyping strategy is required
for DTU identification, manageable in any laboratory and adopted
by the T. cruzi research community.
Over the years, numerous approaches have been used to charac-
terize the biochemical and genetic diversity of T. cruzi isolates. No
single genetic target allows complete DTU resolution, and reliance
on a single target is also inadvisable because of the potential influ-
ence of genetic exchange.
A PCR assay system based on the amplification of particular re-
gions of the SL gene and 24Sa rDNA (Souto and Zingales, 1993;
Souto et al., 1996) and 18S rDNA (Clark and Pung, 1994) was pro-
posed (Brisse et al., 2001) in which the size polymorphisms of the
amplification products were suitable for T. cruzi assignment into
each of the six DTUs (Table 2). However, assignments based on
the absence rather than the presence of PCR products are problem-
atic, thus an alternative set of criteria is preferable for a gold stan-
dard typing method.
A multilocus PCR-RFLP analysis of genetic polymorphism of 12
loci was proposed for DTU genotyping (Rozas et al., 2007), several
of which demonstrated inter-DTU differences, and a combination
of one, two or three of these assays allowed identification of the
complete DTU set. The major limitation of this strategy is the com-
plexity of the analysis.
A three-marker sequential typing strategy (Fig. 3A) was pro-
posed (Lewis et al., 2009b) consisting of PCR amplification of the
24Sa rDNA (Souto and Zingales, 1993; Souto et al., 1996) and
PCR-RFLP of the heat shock protein 60 (HSP60) and glucose-6-
phosphate isomerase (GPI) loci (Westenberger et al., 2005). The
combined application of the three PCR-RFLP markers was sufficient
to discriminate the six DTUs in 45 out of 48 analyzed strains (Lewis
et al., 2009b).
Another three-step assay (D’Avila et al., 2009) is outlined in
Fig. 3B. PCR-RFLP analysis of the COII gene (Freitas et al., 2006) al-
lows discrimination of TcI and TcII from the other DTUs; amplifica-
tion of the non-transcribed spacer (NTS) of SL genes (Burgos et al.,
2007) of the unclassified strains defines two distinct clusters, one
formed by TcIII and TcIV and another by TcV and TcVI; amplifica-
tion of 24Sa rDNA (Souto et al., 1996) then resolves the four DTUs.
A third scheme using nested-hot-start PCR assays is technically
more demanding but has the potential advantage of allowing di-
rect DTU typing in biological (Cardinal et al., 2008; Marcet et al.,
2006) and clinical (Burgos et al., 2007, 2010) samples (Fig. 3C),
and has been improved by the use of four sequential multiplex Real
Time PCR assays using TaqMan probes (Duffy et al., 2010). The first
NTS-SL based PCR employs primers recognizing regions flanking a
50-bp insertion characteristic of TcIII and TcIV intergenic regions.
The second NTS-SL based PCR is a hemi-nested reaction using
TCC-TC1 and TCC-TC2 primers (Souto et al., 1996). The A10 PCR
uses primers (Burgos et al., 2007) that recognize a dimorphic
region within the A10 nuclear fragment (Brisse et al., 2000). At
present the revised assay is being tested on biological samples.
An approach based on fluorescent-labeled fragment barcoding
that detects PCR products from four rDNA domains has also been
devised (Hamilton et al., 2011). This technique was able to differ-
entiate many trypanosome species from South American mam-
mals. Some T. cruzi DTUs (including Tcbat) could be clearly
identified, however, TcV and TcVI could not be distinguished from
TcIII and TcII, respectively.
6. Comparative experimental pathology of the DTUs
Since the discovery of Chagas disease in 1909, heterogeneity of
parasite strains has been considered one factor implicated in dif-
ferent clinical presentations of the disease. Andrade (1974) at-
tempted to discriminate several distinct T. cruzi morphobiological
and behavioral phenotypes in murine models using the criteria of
virulence (capacity of multiplication in the host) and pathogenicity
(ability to produce tissue lesions and immunological responses).
These studies defined three main strain phenotypes (Andrade,
1974; Andrade et al., 1983), subsequently designated as ‘biodemes’
I–III (Andrade and Magalhães, 1997), as follows:
Biodeme type I: strains with rapid multiplication rates, maxi-
mum parasitemia, and mortality from 7 to 11 days after infection;
predominance of slender forms, and macrophagotropism during
the early phase of infection. Neuronal alterations are more
Size of PCR product (in bp) of T. cruzi DTUs.a
DTU24Sa rDNA SL 18S rDNA
aBrisse et al. (2001).
bTcIII and TcIV DTUs can be detected by multiplex PCR of the SL gene (Fernandes
et al., 1998; 2001; Burgos et al., 2007).
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
frequent and intense in biodeme type I infections. Zymodeme
patterns of this biodeme correspond to zymodeme Z2b strains, a
variant of Z2.
Biodeme type II: strains with slow multiplication rates and
irregular parasitemia peaks 12–20 days after infection, when mor-
tality rates reach a maximum; predominance of broad forms, myot-
ropism, with predominant myocardial involvement. Zymodeme
patterns of this biodeme type correspond to zymodeme Z2 strains.
According to the 1999 revised nomenclature, this biodeme should
be classified into the major group T. cruzi II (Anonymous, 1999).
Biodeme type III: slow multiplication strains with late and high
parasitemia peaks 20–30 days following infection and late mortal-
ity, usually from day 30 after infection; predominance of broad
forms and myotropism, with myocardial and skeletal muscle
involvement. Biodeme type III corresponds to zymodeme Z1
strains and to the major group T. cruzi I (Anonymous, 1999).
Due to the intense chronic myocarditis, strains of biodeme type
III are considered the most pathogenic in mice (Andrade, 1974).
Nevertheless, the degree of strain virulence may vary within the
same biodeme and between clones of the same strain (Andrade,
1974; Postan et al., 1987). Further studies in murine experimental
models support and expand these observations (Andrade et al.,
1985) and clearly indicate that both the parasite and host geno-
types are important in determining the tissue distribution, physio-
pathology and eventual outcome of T. cruzi infection. For example,
simultaneous infection of four mouse lineages with the Colombi-
ana Col1.7G2 clone (TcI) and the JG clone (TcII) showed identical
tissue distributions in chronic phase infections of BALB/c and
DBA-2 mice, but very different distributions in C57BL/6 (H-2b)
and outbred Swiss mice (Andrade et al., 1999, 2002). Inoculation
of Sylvio X10/4 clone (TcI) into the C3H/HePAS mouse strain
caused intense cardiac inflammatory lesions, whereas in A/J mice
chronic inflammatory lesions were found in the liver and skeletal
muscle without detectable cardiac pathology (Marinho et al.,
2004). Huge differences in virulence in experimental infections
were reported for TcI isolates from Chile (Andersson et al., 2003),
Fig. 3. Typing approaches for DTU assignment. Panel A: triple-assay proposed by Lewis et al. (2009b). Exceptions pointed by the authors for two T. cruzi IV strains from North
America:⁄characteristic 130 bp 24Sa rDNA PCR product;⁄⁄two bands instead of 3 for GPI-HhaI PCR-RFLP. Panel B: triple-assay proposed by D’Avila et al. (2009). Panel C:
Heminested-PCR assay proposed by Marcet et al. (2006), Burgos et al. (2007, 2010). See details in the text.
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
Mexico (Espinoza et al., 2010) and the United States (Roellig and
Tibayrenc and co-workers undertook long-term comparative
studies on the association between T. cruzi subspecific genetic
diversity and the parasite’s biological properties, including behav-
ior in axenic and mammalian cell culture, drug sensitivity in vitro,
transmissibility through the insect vector and pathogenicity in
mice (for example, see Laurent et al., 1997; de Lana et al., 1998;
Revollo et al., 1998). The general pattern is that different DTUs ex-
hibit statistically different biological properties, but with some
overlap between different DTUs. Interestingly, in several cases
mixtures of two clones behaved differently from a simple summa-
tion of their behavior, suggesting interaction between the geno-
types (Pinto et al., 1998).
As one approach to the underlying molecular basis of biological
differences between DTUs, Telleria et al. (2010) performed phylo-
genetic character mapping of gene expression (proteomic diver-
sity) between the six T. cruzi DTUs. The authors found a
correlation between genetic distances measured by various mark-
ers (MLEE, RAPDs, MLST) and proteomic differences between DTUs,
showing a tight correlation between genetic evolution and proteic
divergence, and they identified several proteins with DTU-specific
Overall these studies indicate extensive intra-DTU phenotypic
diversity, complicating the identification of genetic determinants
of pathogenesis and virulence and requiring higher resolution in-
tra-DTU genetic markers or methods to cross experimentally
strains with different virulence and to analyze the genotypes and
phenotypes of resultant progeny.
7. T. cruzi DTUs and human Chagas disease
7.1. Clinical presentations
T. cruzi is transmitted to humans mainly by triatomine insect
vectors, blood transfusion, infected mothers during pregnancy,
and oral infection by consumption of food contaminated with tri-
atomines or their feces. Following infection, a short acute phase
is recognized only in 1–2% of the infected individuals, character-
ized by an abundant parasitemia and mild symptoms that sponta-
neously decline after 4–8 weeks. The disease proceeds to a chronic
phase with scarce parasitemia and an unpredictable clinical course.
Most of the chronic individuals are asymptomatic and show no
electrocardiographic or radiologic alterations in the heart, esopha-
gus or colon. The individuals present positive serological tests for T.
cruzi infection and in many the xenodiagnosis and PCR results may
be repeatedly positive for many years. These persons with the
‘‘indeterminate’’ form will remain asymptomatic for decades, if
not the rest of their lives. Each year approximately 3% will develop
lesions in the heart or gastrointestinal tract (Dias, 2006). Chronic
cardiomyopathy, or chronic Chagas heart disease is the most com-
mon and severe manifestation in humans, affecting approximately
30% of the patients. In endemic areas, it represents the main cause
of disability and mortality. The basic lesions of chronic Chagas
heart disease are focal or extensive myocardial fibrosis, which re-
sult from myocardial cell destruction due to direct parasite action,
inflammatory response, and neuronal involvement. The gastroin-
testinal manifestations consist of progressive enlargement of the
esophagus or colon caused by chronic inflammation and destruc-
tion of parasympathetic neurons. Great regional diversity of Cha-
gas disease severity and the nature of the chronic infection has
been reported, attributed to a set of complex interactions among
the genetic make-up of the parasite, the host immunogenetic back-
ground, and environmental factors (reviewed by Campbell et al.,
2004; Macedo et al., 2004). A goal of T. cruzi taxonomic studies is
to identify links between the infecting DTUs and the clinical pre-
sentation of disease. No proven associations are evident at present.
The search for these associations will drive the ultimate criterion
for defining the clinically meaningful number of biological subdivi-
sions within this species.
7.2. Acute Chagas disease
The control of vector and blood transfusion transmission in sev-
eral Latin American countries has promoted the steady reduction
of acute infections. In recent years, most of these cases are linked
to oral and congenital transmission, blood transfusion or labora-
tory accidents (Coura, 2006, 2007). Several acute cases were docu-
mented in the Amazon region, most caused by TcI and, less
frequently, by TcIII and TcIV (Coura, 2007). In northern countries
of South America TcI is also the major cause of human acute cases
(Miles et al., 1981; Añez et al., 2004; Llewellyn et al., 2009b and ci-
Acute cases resulting from oral contamination have been docu-
mented for outbreaks in different localities, most frequently in the
Amazon region. Most of these cases were due to TcI, with rare cases
due to TcIII and TcIV (Coura, 2007; Marcili et al., 2009a,b; Valente
et al., 2009), with TcI in Venezuela and French Guiana (Alarcón de
Noya et al., 2010; Cura et al., 2010) and TcII in southern Brazil (Ste-
indel et al., 2008). The morbidity and mortality may vary depend-
ing on the parasite burden and parasite genotype ingested.
The incidence of congenital transmission is estimated at more
than 15,000 cases annually in the Americas and is one of the main
modes of transmission in non-endemic countries. The risk factors
determining transmission of the parasite to the fetus are largely
unknown. Cases of congenital infection with all DTUs except TcIV
were reported in Argentina, Bolivia, Chile, Colombia, and Paraguay
(for examples, see García et al., 2001; Svoboda et al., 2005; Virreira
et al., 2006; Burgos et al., 2007; Corrales et al., 2009; Del Puerto
et al., 2010). The prevalence of specific DTUs among congenital
cases appears to be in accordance with their presence in the in-
fected population. However, there seems to be a disparate preva-
lence of congenital cases in endemic regions, with few cases
reported, for example from Venezuela and Brazil, with the excep-
tion of southern Brazil (Carlier and Truyens, 2010).
7.3. Chronic Chagas disease
TcI is implicated with human disease in Amazonia, the Andean
region, Central America, and Mexico (Bosseno et al., 2002; Montilla
et al., 2002; Añez et al., 2004; Higo et al., 2004; Sánchez-Guillén
et al., 2006). Clinical presentations of TcI include chagasic cardio-
myopathy and in immunocopromized hosts severe cases of
In the Southern Cone region, where T. infestans is the main vec-
tor, TcII, TcV and TcVI are the main causes of Chagas disease. TcII
predominates in eastern and central Brazil, TcV in Argentina, Boli-
via, and Paraguay, and TcVI in the Gran Chaco (Chapman et al.,
1984; Zingales et al., 1999; Brenière et al., 2002; Diosque et al.,
2003; Higo et al., 2004; Coronado et al., 2006; Burgos et al.,
2007; Cardinal et al., 2008; Carranza et al., 2009; Del Puerto
et al., 2010). Throughout the Southern Cone region chagasic cardio-
myopathy can be severe, and a proportion of cases may develop
megaesophagus and megacolon (Luquetti et al., 1986; Freitas
et al., 2005; Lages-Silva et al., 2006). The disparate geographical
distribution of the megasyndromes may reflect the divergent phy-
logeographies of T. cruzi DTUs (Miles et al., 1981), a hypothesis
supported by circumstantial evidence. Chagasic megaesophagus
and megacolon are considered rare in northern South America
and Central America (Miles et al., 2009).
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
As mentioned above, TcIII is virtually absent in chronic infec-
tions, although it is found occasionally in domestic dogs in Para-
guay and Brazil and in peridomestic Triatoma rubrofasciata in Rio
Grande do Sul, Brazil (Yeo et al., 2005; Marcili et al., 2009a; Miles
et al., 2009; Câmara et al., 2010). Consequently, DTU TcIII may yet
become another source of human Chagas disease. TcIV is the sec-
ondary cause of Chagas disease in Venezuela (Miles et al., 1981),
and has been identified in oral transmission outbreaks (Ramirez
et al., 2010). Comparisons of the clinical histories for TcI and TcIV
infections are required in Venezuela, where both DTUs are endemic
and sympatric. A summary of the geographic distribution of DTUs
associated with human Chagas disease and the characteristics of
the prevalent clinical manifestations is included in Table 1.
7.4. Chagas disease reactivation due to immunosuppression
Co-infection with HIV/AIDS and immunosuppressant therapies
can bring about acute and unusual clinical manifestations of Cha-
gas disease, such as cutaneous lesions, involvement of central ner-
vous system, and/or serious cardiac lesions. Genotyping of
parasites recovered from the blood of Chagas disease patients with
HIV and in both blood and tissue lesions from patients presenting
clinical reactivation due to AIDS revealed differential tissue tro-
pism of the infecting DTUs (Burgos et al., 2005, 2008; Bisio et al.,
2009). In 18 Argentinean patients, TcV was found in almost all
blood samples, in agreement with previous findings in this region.
In two cases, mixed infections by TcV and TcI were observed and in
one of these patients the cerebrospinal fluid sample amplified only
TcI (Burgos et al., 2008).
A more complex scenario was seen in late-stage Chagas heart
disease patients undergoing heart transplants, manifesting TcI,
TcV, or TcVI in the bloodstream, in endomyocardial biopsies of
the implanted heart and in skin tissues, provoking myocarditis
and skin reactivation after immunosuppressive post-transplanta-
tion treatment, respectively (Burgos et al., 2010).
Conclusions and comparisons of clinical manifestations and
parasite genotype are complicated for Chagas disease for several
reasons. Isolates from blood do not necessarily reveal the full com-
plement of infecting parasite lineages in individual patients, as one
or several distinct T. cruzi strains may be sequestered in the tissues
(Vago et al., 2000; D’Avila et al., 2009; Burgos et al., 2010; Câmara
et al., 2010). Asymptomatic patients may have sub-clinical cardiac
or digestive alterations detectable only by imaging studies. Addi-
tionally, parasite selection may occur during isolation procedures
for genetic analysis due to the preferential proliferation of certain
clones. A theoretical solution to these problems is to design pep-
tides or recombinant proteins for DTU-specific serology that could
be used to provide a current and historical profile of all the T. cruzi
DTUs infecting an individual patient. DTU-specific serology would
greatly facilitate comparisons of virulence and pathogenesis (Bhat-
tacharyya et al., 2010; Risso et al., 2011).
8. T. cruzi genomics
When the TriTryp genome projects published their initial find-
ings in 2005, researchers were just coming to terms with the latest
hitch in T. cruzi genetics. Contrary to prevailing expectations, T. cru-
zi DTUs TcV and TcVI are both largely heterozygous in their nuclear
content (Westenberger et al., 2005). The strain CL Brener chosen to
represent T. cruzi (Zingales et al., 1997; El-Sayed et al., 2005) is a
member of DTU TcVI, derived from T. infestans. This complication
resulted in several consequences: (1) the final coverage level for
CL Brener was 7-fold in depth, versus the expected target 15-fold;
(2) the heterozygous genome could not be accurately assembled,
and thus was presented in its fragmented state; (3) the Esmeraldo
strain from DTU TcII was sequenced at 2.5? coverage to aid in the
CL Brener assembly as a representative of a ‘parental’ contributor
to TcVI heterogeneity.
The sequencing effort revealed that CL Brener genome contains
?22,000 protein-encoding genes, and that over 50% is represented
by repetitive sequences, consisting mostly of large gene families of
surface proteins, retrotransposons, subtelomeric repeats (El-Sayed
et al., 2005) and the T. cruzi-specific 195-bp satellite DNA (Martins
et al., 2008). Putative function could be assigned to approximately
half of the predicted protein-coding genes on the basis of signifi-
cant similarity to previously characterized proteins or known func-
tional domains. Thus around 6,000 proteins hold promise for new
areas of investigation.
The decision to use a shotgun sequencing approach for the
gathering of genomic data in T. cruzi led to the accumulation of
linked 400–900 bp sequences. In the assembly process, elements
repeated over a specific threshold were excluded, leading to the
absence of the maxicircle genome in the initial report. Both the
CL Brener and Esmeraldo maxicircles were reconstructed indepen-
dently in their entirety from the primary sequence reads, each
with a coverage of approximately 50? (Westenberger et al.,
2006a). The size of the sequence reads and the depth of coverage
allowed the assembly of the non-coding variable region of the
mitochondrial genomes that are comprised of highly repetitive se-
quence motifs. A relatively low number of minicircle fragments
was also found among the primary sequence reads (Thomas
et al., 2007). To fulfill the complement of mitochondrial genomes,
the Sylvio X10/1 strain maxicircle DNA sequence was assembled
through an ordered amplification strategy (Ruvalcaba-Trejo and
The many multicopy nuclear genes common in T. cruzi were rel-
egated to the same fate as the maxicircle, resulting either in their
exclusion or compression within the assembly, and skewing their
initial representation. Analysis of the repeated nuclear protein-
coding genes nearly doubles the total number of genes emerging
from the T. cruzi genomic analysis (Arner et al., 2007), highlighting
another level of genetic complexity in this ancient pathogen. A few
multicopy RNA gene families have been studied individually, as
represented by the SL RNA genes (Thomas et al., 2005) and 5S
rRNA genes (Westenberger et al., 2006b). The accurate assembly
of any large tandem array is problematic, even among those of
smaller periodicities such as the SL RNA gene array in Leishmania
Currently an assembled version of CL Brener is available to the
community through TriTrypDB (Weatherly et al., 2009). A steady-
state transcriptome analysis has been performed (Minning et al.,
2009). At the protein level, multiple studies are emerging to com-
plement the initial proteome study (Atwood et al., 2005) that ap-
peared alongsidethe TriTryp
organelles (Ferella et al., 2008) and ribosomes (Ayub et al., 2009).
Most of the available genome, transcriptome and proteomic
data have been obtained for the CL Brener strain. As discussed pre-
viously, several experimental lines indicate that T. cruzi DTUs dis-
play differential virulenceand
however no genetic markers are linked with the severity of the
infection. TcI, TcII, TcV and TcVI are the main agents of human Cha-
gas disease in the Americas, and all are capable of causing cardio-
myopathies, however, only DTUs TcII, TcV and TcVI have been so
far associated to chronic digestive syndromes. The comparative
analysis of the genomes from isolates of various DTUs may shed
light on pathogenic and epidemiological features of these parasites,
and promote the development of new DTU-specific diagnostic
tests. The sequence of the TcI reference strain Sylvio X10/1 has re-
cently been published (Franzen et al., 2011), and a second TcI se-
quence (JR cl4) and TcII (Esmeraldo cl3) sequence have also
entered the public domain, via the TriTryp database.
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
9. Concluding remarks and perspectives
The revised subspecific nomenclature for T. cruzi (Zingales et al.,
2009) recognized that T. cruzi strains should be assigned to one of
six DTUs. The important change in the new nomenclature was that
TcII was no longer divided into five subgroups (TcIIa-e) (Brisse
et al., 2000) but each of those subgroups became independent
DTUs (TcII–VI). The rationale for this change provides the underly-
ing theme for the above review and is abundantly clear from sev-
The apparent affinities between TcI and TcIII and TcIV were
noted when they and the other three T. cruzi subspecific groups
were first described decades ago on the basis of MLEE: in particular
overlapping isoenzyme profiles between what are now designated
as TcI and TcIII (Miles et al., 2009). These affinities have been con-
firmed repeatedly by other molecular markers and phylogenetic
analysis, perhaps most notably by MLST studies and now by com-
parisons of the genome sequences of the hybrid TcVI (CL Brener
strain) and TcI (Sylvio X10/1 strain), as cited above. Furthermore,
the known phylogeographical and eco-epidemiological associa-
tions of TcIII and TcIV and perceptions of their evolutionary origins,
also described above, do not sit comfortably with them being sub-
divisions of TcII. The revised nomenclature therefore provides a
more suitable and valuable framework for future research.
The decade between the two meetings on the nomenclature of
T. cruzi has seen major advances in the understanding of this
important pathogen, at many levels. Consistent with the theme
of this review, here we have focused on population structure, geno-
typing, emergent comparative genomics, and the association of
DTU with features of natural and experimental populations. Mech-
anisms of parasite-host interactions and immune responses to
infection have not been addressed here. It is now proven experi-
mentally that T. cruzi has an extant capacity for genetic exchange.
The extent and mechanisms of genetic exchange in natural popula-
tions are not understood. TcV and TcVI are of special interest, be-
cause they are recent, rapidly spreading and epidemiologically
important inter-DTU hybrids of TcII and TcIII (Lewis et al., 2011).
There are indications, from inter-DTU mitochondrial introgression
and apparent panmixia within localized populations of TcI, that ge-
netic exchange is more widespread than hitherto appreciated.
However, the broad integrity of the DTUs and their validity for
population and eco-epidemiological studies is not disrupted. Nev-
ertheless, the DTU nomenclature is inevitably a dynamic structure
as research progresses and more discoveries are made. More in-
tense and widespread sampling of T. cruzi isolates is required from
silvatic populations, and repeated isolation from individual hosts
to resolve multiclonality.
As described above, straightforward genotyping methods to
identify the DTUs are now available for widespread use in endemic
areas, and research is in progress to optimize sensitivity and sim-
plify techniques so that they may more easily be applied directly
to clinical and biological samples. For the immediate future MLST,
which provides reproducible and transferable data, is likely to be
the gold standard for population studies. Microsatellites provide
valuable high-resolution markers to produce less robust datasets
that have specialized population genetic objectives. Identification
of the genetic determinants of pathogenesis is extraordinarily chal-
lenging and they remain elusive. However, current rate of progress
with new generation low cost sequencing technologies is astonish-
ing and relevant. Although problems remain with assembly of
highly repetitive regions, which are so abundant in T. cruzi, it is
now realistic to obtain genome sequence from many T. cruzi iso-
lates simultaneously, which will certainly illuminate the biology
of this important pathogen. Fundamental to such rapid research
progress is the ability of the scientific community to collaborate
effectively, particularly between molecular biology laboratories
and field research in endemic regions. An integrated but focused
approach, not neglecting drug discovery, is important for improve-
ment of control strategies that may reduce the public health bur-
den of Chagas disease.
BZ thanks the Fundação de Amparo à Pesquisa do Estado de São
Paulo (FAPESP) and Ministério de Ciência e Tecnologia/Conselho
Nacional de Desenvolvimento Científico e Tecnológico/Ministério
da Saúde (MCT/CNPq/MS-SCTIE-DECIT-Edital de Doenças Negli-
genciadas) for financial support. MAM and MSL thank the Well-
come Trust (UK) and European Union Seventh Program Grant
223034 (ChagasEpiNet) for financial support. DAC and NRS are
supported by NIH award AI056034. We thank Michael Lewis and
Matthew Yeo for comments on the manuscript.
Alarcón de Noya, B., Díaz-Bello, Z., Colmenares, C., Ruiz-Guevara, R., Mauriello, L.,
Zavala-Jaspe, R., et al., 2010. Large urban outbreak of orally acquired acute
Chagas disease at a school in Caracas. Venezuela. J. Infect. Dis. 201, 1308–1315.
Andersson, J., Orn, A., Sunnemark, D., 2003. Chronic murine Chagas’ disease: the
impact of host and parasite genotypes. Immunol. Lett. 86, 207–212.
Andrade, L.O., Machado, C.R.S., Chiari, E., Pena, S.D.J., Macedo, A., 1999. Differential
tissue distribution of diverse clones of Trypanosoma cruzi in infected mice. Mol.
Biochem. Parasitol. 100, 163–172.
Andrade, L.O., Machado, C.R.S., Chiari, E., Pena, S.D.J., Macedo, A., 2002. Trypanosoma
cruzi: a role of the host genetic background in the differential tissue distribution
of parasite clonal populations. Exp. Parasitol. 100, 269–275.
Andrade, S.G., 1974. Caracterização de cepas de Trypanosoma cruzi isoladas do
Recôncavo Baiano. Rev. Patol. Trop. 3, 65–121.
Andrade, S.G., Magalhães, J.B., 1997. Biodemes and zymodemes of Trypanosoma
cruzi. Rev. Soc. Bras. Med. Trop. 30, 27–35.
Andrade, V., Barral-Netto, M., Andrade, S.G., 1985. Patterns of resistance of inbred
mice to Trypanosoma cruzi are determined by parasite strain. Br. J. Med. Biol.
Res. 18, 499–506.
Andrade, V., Brodskyn, C., Andrade, S.G., 1983. Correlation between isoenzyme
patterns and biological behaviour of different strains of Trypanosoma cruzi.
Trans. R. Soc. Trop. Med. Hyg. 76, 796–799.
Añez, N., Crisante, G., da Silva, F.M., Rojas, A., Carrasco, H., Umezawa, E.S., et al.,
2004. Predominance of lineage I among Trypanosoma cruzi isolates from
Venezuelan patients with different clinical profiles of acute Chagas’ disease.
Trop. Med. Int. Health 9, 1319–1326.
Anonymous, 1999. Recommendations from a Satellite Meeting. Mem. Inst. Oswaldo
Cruz 94(Suppl. II), 429–432.
Apt, W., Aguilera, X., Arribada, A., Gomez, L., Miles, M.A., Widmer, G., 1987.
Epidemiology of Chagas disease in Northern Chile – isozyme profiles of
Trypanosoma cruzi from domestic and sylvatic transmission cycles and their
association with cardiopathy. Am. J. Trop. Med. Hyg. 37, 302–307.
Arner, E., Kindlund, E., Nilsson, D., Farzana, F., Ferella, M., Tammi, M.T., Andersson,
B., 2007. Database of Trypanosoma cruzi repeated genes: 20, 000 additional
gene variants. BMC Genomics 8, 391.
Atwood 3rd, J.A., Weatherly, D.B., Minning, T.A., Bundy, B., Cavola, C., Opperdoes,
F.R., Orlando, R., Tarleton, R.L., 2005. The Trypanosoma cruzi proteome. Science
Avise, J.C., 2004. Molecular markers, Natural History and Evolution second ed.
Chapman & Hall, New York, London.
Ayub, M.J., Atwood, J., Nuccio, A., Tarleton, R., Levin, M.J., 2009. Proteomic analysis of
the Trypanosoma cruzi ribosomal proteins. Biochem. Biophys. Res. Commun.
Barnabé, C., Brisse, S., Tibayrenc, M., 2000. Population structure and genetic typing
of Trypanosoma cruzi, the agent of Chagas disease: a multilocus enzyme
electrophoresis approach. Parasitology 120, 513–526.
Barnabé, C., Brisse, S., Tibayrenc, M., 2003. Phylogenetic diversity of bat
trypanosomes of subgenus Schizotrypanum based on multilocus enzyme
electrophoresis, random amplified polymorphic DNA, and cytochrome b
nucleotide sequence analyses. Infect. Genet. Evol. 2, 201–208.
Bhattacharyya, T., Brooks, J., Yeo, M., Carrasco, H.J., Lewis, M.D., Llewellyn, M.S.,
Miles, M.A., 2010. Analysis of molecular diversity of the Trypanosoma cruzi
trypomastigote small surface antigen reveals novel epitopes, evidence of
positive selection and potential implications for lineage-specific serology. Int.
J. Parasitol. 40, 921–928.
Bisio, M.M.C., Cura, C.I., Duffy, T., Altcheh, J., Giganti, S.O., Bergher, S., et al., 2009.
Trypanosoma cruzi discrete typing units in Chagas disease patients with HIV co-
infection. Rev. Biom. 20, 166–178.
Bogliolo, A.R., Lauria-Pires, L., Gibson, W.C., 1996. Polymorphisms in Trypanosoma
cruzi: evidence of genetic recombination. Acta Trop. 61, 31–40.
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
Bosseno, M.F., Barnabé, C., Magallon, G.E., Lozano, K.F., Ramsey, J., Espinoza, B.,
Brenière, S.F., 2002. Predominance of Trypanosoma cruzi lineage I in Mexico. J.
Clin. Microbiol. 40, 627–632.
Brenière, S.F., Bosseno, M.F., Noireau, F., Yacsik, N., Liegeard, P., Aznar, C.,
Hontebeyrie, M., 2002. Integrate study of a Bolivian population infected by
Trypanosoma cruzi, the agent of Chagas disease. Mem. Inst. Oswaldo Cruz 97,
Briones, M.R.S., Souto, R.P., Stolf, B.S., Zingales, B., 1999. The evolution of two
Trypanosoma cruzi subgroups inferred from rRNA genes can be correlated with
interchange of American mammalian faunas in the Cenozoic and has
implication to pathogenicity and host specificity. Mol. Biochem. Parasitol.
Brisse, S., Barnabé, C., Bañuls, A.L., Sidibe, I., Noel, S., Tibayrenc, M., 1998. A
phylogenetic analysis of the Trypanosoma cruzi genome project CL Brener
reference strain by multilocus enzyme electrophoresis and multiprimer random
amplified polymorphic DNA fingerprinting. Mol. Biochem. Parasitol. 92, 253–
Brisse, S., Dujardin, J.C., Tibayrenc, M., 2000. Identification of six Trypanosoma cruzi
phylogenetic lineages by sequence-characterised amplified region markers.
Mol. Biochem. Parasitol. 111, 95–105.
Brisse, S., Henriksson, J., Barnabé, C., Douzery, E.J., Berkvens, D., Serrano, M., et al.,
2003. Evidence for genetic exchange and hybridization in Trypanosoma cruzi
based on nucleotide sequences and molecular karyotype. Infect. Genet. Evol. 2,
Brisse, S., Verhoef, J., Tibayrenc, M., 2001. Characterisation of large and small
subunit rRNA and mini-exon genes further supports the distinction of six
Trypanosoma cruzi lineages. Int. J. Parasitol. 31, 1218–1226.
Broutin, H., Tarrieu, F., Tibayrenc, M., Oury, B., Barnabé, C., 2006. Phylogenetic
analysis of the glucose-6-phosphate isomerase gene in Trypanosoma cruzi. Exp.
Parasitol. 113, 1–7.
Burgos, J.M., Altcheh, J., Bisio, M., Duffy, T., Valadares, H.M., Seidenstein, M.E., et al.,
2007. Direct molecular profiling of minicircle signatures and lineages of
Trypanosomacruzi bloodstream populations
disease. Int. J. Parasitol. 37, 1319–1327.
Burgos, J.M., Begher, S., Silva, H.M., Bisio, M., Duffy, T., Levin, M.J., Macedo, A.M.,
et al., 2008. Molecular identification of Trypanosoma cruzi I tropism for central
nervous system in Chagas reactivation due to AIDS. Am. J. Trop. Med. Hyg. 78,
Burgos, J.M., Begher, S.B., Freitas, J.M., Bisio, M., Duffy, T., Altcheh, J., et al., 2005.
Molecular diagnosis and typing of Trypanosoma cruzi populations and lineages
in cerebral Chagas disease in a patient with AIDS. Am. J. Trop. Med. Hyg. 73,
Burgos, J.M., Diez, M., Vigliano, C., Bisio, M., Risso, M., Duffy, T., et al., 2010.
Molecular identification of Trypanosoma cruzi discrete typing units in end-stage
chronic Chagas heart disease and reactivation after heart transplantation. Clin.
Infect. Dis. 51, 485–495.
Buscaglia, C.A., Di Noia, J.M., 2003. Trypanosoma cruzi clonal diversity and the
epidemiology of Chagas’ disease. Microbes Infect. 5, 419–427.
Câmara, A.C.J., Varela-Freire, A.A., Valadares, H.M.S., Macedo, A.M., D’Avila, D.A.,
Machado, C.R., et al., 2010. Genetic analyses of Trypanosoma cruzi isolates from
naturally infected triatomines and humans in northeastern Brazil. Acta Trop.
Campbell, D.A., Westenberger, S.J., Sturm, N.R., 2004. The determinants of Chagas
disease: connecting parasite and host genetics. Curr. Mol. Med. 4, 549–562.
Cardinal, M.V., Lauricella, M.A., Ceballos, L.A., Lanati, L., Marcet, P.L., Levin, M.J.,
et al., 2008. Molecular epidemiology of domestic and sylvatic Trypanosoma cruzi
infection in rural northwestern Argentina. Int. J. Parasitol. 38, 1533–1543.
Carlier, Y., Truyens, C., 2010. Maternal-fetal transmission of Trypanosoma cruzi. In:
Telleria, J., Tibayrenc, M. (Eds.), American Trypanosomiasis Chagas disease one
hundred years of research. Elsevier Inc.
Carranza, J.C., Valadares, H.M.S., D’Avila, D.A., Baptista, R.P., Moreno, M., Galvão,
L.M.C., et al., 2009. Trypanosoma cruzi maxicircle heterogeneity in Chagas
disease patients from Brazil. Int. J. Parasitol. 39, 963–973.
Carrasco, H.J., Frame, I.A., Valente, S.A., Miles, M.A., 1996. Genetic exchange as a
possible source of genomic diversity in sylvatic populations of Trypanosoma
cruzi. Am. J. Trop. Med. Hyg. 54, 418–424.
Cavazzana, M., Marcili, A., Lima, L., da Silva, F.M., Junqueira, A.C.V., Veludo, H.H.,
et al., 2010. Phylogeographical, ecological and biological patterns shown by
nuclear (ssrRNAand gGAPDH) and
trypanosomes of the subgenus Schizotrypanum parasitic in Brazilian bats. Int.
J. Parasitol. 40, 345–355.
Chapman, M.D., Baggaley, R.C., Godfreyfausset, P.F., Malpas, T.J., White, G., Canese, J.,
Miles, M.A., 1984. Trypanosoma cruzi from the Paraguayan Chaco – isoenzyme
profiles of strains isolated at Makthlawaiya. J. Protozool. 31, 482–486.
Clark, C.G., Pung, O.J., 1994. Host specificity of ribosomal DNA variation in sylvatic
Trypanosoma cruzi from North America. Mol. Biochem. Parasitol. 66, 175–179.
Coronado, X., Zulantay, I., Albrecht, H., Rozas, M., Apt, W., Ortiz, S., et al., 2006.
Variation in Trypanosoma cruzi clonal composition detected in blood patients
and xenodiagnosis triatomines: Implications in the molecular epidemiology of
Chile. Am. J. Trop. Med. Hyg. 74, 1008–1012.
Corrales, R.M., Mora, M.C., Negrette, O.S., Diosque, P., Lacunza, D., Virreira, M., et al.,
2009. Congenital Chagas disease involves Trypanosoma cruzi sub-lineage IId in
the northwestern province of Salta. Argentina Inf. Genet. Evol. 9, 278–282.
Coura, J.R., 2007. Chagas disease: what is known and what is needed – a background
article. Mem. Inst. Oswaldo Cruz 102, 113–122.
Coura, J.R., 2006. Transmission of chagasic infection by oral route in the natural
history of Chagas disease. Rev. Soc. Bras. Med. Trop. 39, 113–117.
Cura, C.I., Mejia-Jaramillo, A.M., Duffy, T., Burgos, J.M., Rodriguero, M., Cardinal,
M.V., et al., 2010. Trypanosoma cruzi I genotypes in different geographical
regions and transmission cycles based on a microsatellite motif of the
intergenic spacer of spliced-leader genes. Int. J. Parasitol. 40, 1599–1607.
D’Avila, D.A., Macedo, A.M., Valadares, H.M., Gontijo, E.D., de Castro, A.M., Machado,
C.R., Chiari, E., Galvão, L.M., 2009. Probing population dynamics of Trypanosoma
cruzi during progression of the chronic phase in chagasic patients. J. Clin.
Microbiol. 47, 1718–1725.
de Lana, M., da Silveira, P.A., Barnabé, C., Quesney, V., Noel, S., Tibayrenc, M., 1998.
Trypanosoma cruzi: compared vectorial transmissibility of three major clonal
genotypes by Triatoma infestans. Exp. Parasitol. 90, 20–25.
Del Puerto, F., Sanchez, Z., Nara, E., Meza, G., Paredes, B., Ferreira, E., Russomando, G.,
2010. Trypanosoma cruzi lineages detected in congenitally infected infants and
Triatoma infestans from the same disease-endemic region under entomologic
surveillance in Paraguay. Am. J. Trop. Med. Hyg. 82, 386–390.
Devera, R., Fernandes, O., Coura, J.R., 2003. Should Trypanosoma cruzi be called
‘‘cruzi’’ complex? A review of the parasite diversity and the potential of
selecting population after in vitro culturing and mice infection. Mem. Inst.
Oswaldo Cruz 98, 1–12.
Dias, J.C.P., 2006.The treatmentof
trypanosomiasis). Ann. Int. Med. 144, 772–774.
Diosque, P., Barnabé, C., Padilla, A.M., Marco, J.D., Cardozo, R.M., Cimino, R.O., et al.,
2003. Multilocus enzyme electrophoresis analysis of Trypanosoma cruzi isolates
from a geographically restricted endemic area for Chagas’ disease in Argentina.
Int. J. Parasitol. 33, 997–1003.
Duffy, T., Bisio, M., Altcheh, J., Burgos, J.M., Diez, M., Levin, M.J., et al., 2010. Accurate
real-time PCR strategy for monitoring bloodstream parasitic loads in Chagas
disease patients. PLoS Negl. Trop. Dis. 3, e419.
Dvorak, J.A., Hall, T.E., Crane, M.S., Engel, J.C., Mcdaniel, J.P., Uriegas, R., 1982.
Trypanosoma cruzi – flow Cytometric Analysis.1. Analysis of total DNA organism
by means of mithramycin-induced fluorescence. J. Protozool. 29, 430–437.
Elias, M.C.Q.B., Vargas, N., Tomazi, L., Pedroso, A., Zingales, B., Schenkman, S.,
Briones, M.R.S., 2005. Comparative analysis of genomic sequences suggests that
Trypanosoma cruzi CL Brener contains two sets of non-intercalated repeats of
satellite DNA that correspond to T. Cruzi I and T. cruzi II types. Mol. Biochem.
Parasitol. 140, 221–227.
El-Sayed, N.M., Myler, P.J., Bartholomeu, D.C., Nilsson, D., Aggarwal, G., Tran, A.N.,
et al., 2005. The genome sequence of Trypanosoma cruzi, etiologic agent of
Chagas disease. Science 309, 409–415.
Espinoza, B., Rico, T., Sosa, S., Oaxaca, E., Vizcaino-Castillo, A., Caballero, M.L.,
Martínez, I., 2010. Mexican Trypanosoma cruzi T. cruzi I strains with different
degrees of virulence induce diverse humoral and cellular immune responses in
a murine experimental infection model. J. Biomed. Biotechnol. 2010, 890672.
Falla, A., Herrera, C., Fajardo, A., Montilla, M., Vallejo, G.A., Guhl, F., 2009. Haplotype
identification within Trypanosoma cruzi I in Colombian isolates from several
reservoirs, vectors and humans. Acta Trop. 110, 15–21.
Ferella, M., Nilsson, D., Darban, H., Rodrigues, C., Bontempi, E.J., Docampo, R.,
Andersson, B., 2008. Proteomics in Trypanosoma cruzi – localization of novel
proteins to various organelles. Proteomics 8, 2735–2749.
Fernandes, O., Sturm, N.R., Derré, R., Campbell, D.A., 1998. The mini-exon gene: a
genetic marker for zymodeme III of Trypanosoma cruzi. Mol. Biochem. Parasitol.
Fernandes, O., Mangia, R.H., Lisboa, C.V., Pinho, A.P., Morel, C.M., Zingales, B.,
Campbell, D.A., Jansen, A.M., 1999. The complexity of the sylvatic cycle of
Trypanosoma cruzi in Rio de Janeiro state (Brazil) revealed by the non-
transcribed spacer of the mini-exon gene. Parasitology 118, 161–166.
Fernandes, O., Santos, S.S., Cupolillo, E., Mendonça, B., Derre, R., Junqueira, A.C.,
et al., 2001. A mini-exon multiplex polymerase chain reaction to distinguish the
major groups of Trypanosoma cruzi and T. rangeli in the Brazilian Amazon. Trans.
R. Soc. Trop. Med. Hyg. 95, 97–99.
Flores-López, C.A., Machado, C.A., 2011. Analyses of 32 loci clarify phylogenetic
relationships among Trypanosoma cruzi lineages and support a single
hybridization prior to human contact. PLoS Negl. Trop. Dis. 5, e1272.
Franzen, O., Ochaya, S., Sherwood, E., Lewis, M.D., Llewellyn, M.S., Miles, M.A.,
Andersson, B., 2011. Shotgun sequencing analysis of Trypanosoma cruzi I Sylvio
X10/1 and comparison with T. cruzi VI CL Brener. PLoS Negl.Trop. Dis. 5, e984.
Freitas, J.M., Augusto-Pinto, L., Pimenta, J.R., Bastos-Rodrigues, L., Gonçalves, V.F.,
Teixeira, S.M.R., et al., 2006. Ancestral genomes, sex, and the population
structure of Trypanosoma cruzi. PLoS Pathogens 2, 226–235.
Freitas, J.M., Lages-Silva, E., Crema, E., Pena, S.D.J., Macedo, A.M., 2005. Real time PCR
strategy for the identification of major lineages of Trypanosoma cruzi directly in
chronically infected human tissues. Int. J. Parasitol. 35, 411–417.
García, A., Bahamonde, M., Verdugo, S., Correa, J., Pastene, C., Solari, A., et al., 2001.
Trypanosoma cruzi transplacental infection: situation in Chile. Rev. Med. Chil.
Gaunt, M., Miles, M., 2000. The ecotopes and evolution of triatomine bugs
(triatominae) and their associated trypanosomes. Mem. Inst. Oswaldo Cruz
Gaunt, M.W., Yeo, M., Frame, I.A., Stothard, J.R., Carrasco, H.J., Taylor, M.C., et al.,
2003. Mechanism of genetic exchange in American trypanosomes. Nature 421,
Hamilton, P.B., Lewis, M.D., Cruickshank, C., Gaunt, M.W., Yeo, M., Llewellyn, M.S.,
et al., 2011. Identification and lineage genotyping of South American
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
trypanosomes using fluorescent fragment length barcoding. Infect. Genet. Evol.
Herrera, C., Bargues, M.D., Fajardo, A., Montilla, M., Triana, O., Vallejo, G.A., Guhl, F.,
2007. Identifying four Trypanosoma cruzi I isolate haplotypes from different
geographic regions in Colombia. Inf. Gen. Evol. 7, 535–539.
Herrera, C., Guhl, F., Falla, A., Fajardo, A., Montilla, M., Adolfo, V.G., Bargues, M.D.,
2009. Genetic variability and phylogenetic relationships within Trypanosoma
cruzi I isolated in Colombia based on miniexon gene sequences. J. Parasitol. Res.
2009, article ID 897364.
Higo, H., Miura, S., Horio, M., Mimori, T., Hamano, S., Agatsuma, T., Yanagi, T., et al.,
2004. Genotypic variation among lineages of Trypanosoma cruzi and its
geographic aspects. Parasitol. Int. 53, 337–344.
Ienne, S., Pedroso, A., Ferreira, R.C.E., Briones, M.R.S., Zingales, B., 2010. Network
genealogy of 195-bp satellite DNA supports the superimposed hybridization
hypothesis of Trypanosoma cruzi evolutionary pattern. Infect. Genet. Evol. 10,
Jaramillo, N., Moreno, J., Triana, O., Arcos-Burgos, M., Munoz, S., Solari, A., 1999.
Genetic structure and phylogenetic relationships of Colombian Trypanosoma
cruzi populations as determined by schizodeme and isoenzyme markers. Am. J.
Trop. Med. Hyg. 61, 986–993.
Kawashita, S.Y., Sanson, G.F.O., Fernandes, O., Zingales, B., Briones, M.R.S., 2001.
Maximum-likelihood divergence date estimates based on rRNA gene sequences
suggest two scenarios of Trypanosoma cruzi intraspecific evolution. Mol. Biol.
Evol. 18, 2250–2259.
Lages-Silva, E., Ramirez, L.E., Pedrosa, A.L., Crema, E., Cunha Galvão, L.M., Pena, S.D.J.,
et al., 2006. Variability of kinetoplast DNA gene signatures of Trypanosoma cruzi
II strains from patients with different clinical forms of Chagas’ disease in Brazil.
J. Clin. Microbiol. 44, 2167–2171.
Laurent, J.P., Barnabé, C., Quesney, V., Noel, S., Tibayrenc, M., 1997. Impact of clonal
evolution on the biological diversity of Trypanosoma cruzi. Parasitology 114,
Lewis, M.D., Llewellyn, M.S., Gaunt, M.W., Yeo, M., Carrasco, H.J., Miles, M.A., 2009a.
Flow cytometric analysis and microsatellite genotyping reveal extensive DNA
content variation in Trypanosoma cruzi populations and expose contrasts
between natural and experimental hybrids. Int. J. Parasitol. 39, 1305–1317.
Lewis, M.D., Llewellyn, M.S., Yeo, M., Acosta, N., Gaunt, M.W., Miles, M.A., 2011.
Recent, independent and anthropogenic origins of Trypanosoma cruzi hybrids.
PLoS Negl. Trop. Dis. 5, e1363.
Lewis, M.D., Ma, J., Yeo, M., Carrasco, H.J., Llewellyn, M.S., Miles, M.A., 2009.
Genotyping of Trypanosoma cruzi: systematic selection of assays allowing rapid
and accurate discrimination of all known lineages. Am. J. Trop. Med. Hyg. 81,
Lisboa, C.V., Pinho, A.P., Monteiro, R.V., Jansen, A.M., 2007. Trypanosoma cruzi
(kinetoplastida Trypanosomatidae): biological heterogeneity in the isolates
derived from wild hosts. Exp. Parasitol. 116, 150–155.
Llewellyn, M.S., Lewis, M.D., Acosta, N., Yeo, M., Carrasco, H.J., Segovia, M., Vargas, J.,
Torrico, F., Miles, M.A., Gaunt, M.W., 2009a. Trypanosoma cruzi IIc: phylogenetic
and phylogeographic insights from sequence and microsatellite analysis and
potential impact on emergent Chagas disease. PLoS Negl. Trop. Dis. 3, e510.
Llewellyn, M.S., Miles, M.A., Carrasco, H.J., Lewis, M.D., Yeo, M., Vargas, J., Torrico, F.,
Diosque, P., Valente, V., Valente, S.A., Gaunt, M.W., 2009b. Genome-scale
multilocus microsatellite typing of Trypanosoma cruzi Discrete Typing Unit I
reveals phylogeographic structure and specific genotypes linked to human
infection. PLoS Pathogens 5, e1000410.
Llewellyn, M.S., Rivett-Carnac, J.B., Fitzpatrick, S., Lewis, M.D., Yeo, M., Gaunt, M.W.,
Miles, M.A., 2011. Extraordinary Trypanosoma cruzi diversity within single
mammalian reservoir hosts implies a mechanism of diversifying selection. Int. J.
Parasitol. 41, 609–614.
Luquetti, A.O., Miles, M.A., Rassi, A., de Rezende, J.M., de Souza, A.A., Povoa, M.M.,
Rodrigues, I., 1986. Trypanosoma cruzi: zymodemes associated with acute and
chronic Chagas’ disease in central Brazil. Trans. R. Soc.Trop. Med. Hyg. 80, 462–
Macedo, A.M., Oliveira, R.P., Pena, S.D., 2002. Chagas disease: role of parasite genetic
variation in pathogenesis. Expert. Rev. Mol. Med. 4, 1–16.
Macedo, A.M., Machado, C.R., Oliveira, R.P., Pena, S.D.J., 2004. Trypanosoma cruzi:
genetic structure of populations and relevance of genetic variability to the
pathogenesis of Chagas disease. Mem. Inst. Oswaldo Cruz 99, 1–12.
Machado, C.A., Ayala, F.J., 2002. Sequence variation in the dihydrofolate reductase-
thymidylate synthase (DHFR-TS) and trypanothione reductase (TR) genes of
Trypanosoma cruzi. Mol. Biochem. Parasitol. 121, 33–47.
Machado, C.A., Ayala, F.J., 2001. Nucleotide sequences provide evidence of genetic
exchange among distantly related lineages of Trypanosoma cruzi. Proc. Natl.
Acad. Sci. USA 98, 7396–7401.
Maia da Silva, F., Marcili, A., Lima, L., Cavazzana, M.Jr., Ortiz, P.A., Campaner, M.,
et al., 2009. Trypanosoma rangeli isolates of bats from Central Brazil: genotyping
and phylogenetic analysis enable description of a new lineage using spliced-
leader gene sequences. Acta Trop. 109, 199–207.
Marcet, P.L., Duffy, T., Cardinal, M.V., Burgos, J.M., Lauricella, M.A., Levin, M.J., et al.,
2006. PCR-based screening and lineage identification of Trypanosoma cruzi
directly from faecal samples of triatomine bugs from northwestern Argentina.
Parasitology 132, 57–65.
Marcili, A., Lima, L., Cavazzana, M.J., Junqueira, A.C.V., Veludo, H.H., da Silva, F.M.,
et al., 2009a. A new genotype of Trypanosoma cruzi associated with bats
evidenced by phylogenetic analyses using SSU rDNA, cytochrome b and Histone
H2B genes and genotyping based on ITS1 rDNA. Parasitology 136, 641–655.
Marcili, A., Lima, L., Valente, V.C., Valente, S.A., Batista, J.S., Junqueira, A.C., et al.,
2009b. Comparative phylogeography of Trypanosoma cruzi TCIIc: new hosts,
association with terrestrial ecotopes, and spatial clustering. Infect. Genet. Evol.
Marcili, A., Valente, V.C., Valente, S.A., Junqueira, A.C., da Silva, F.M., Pinto, A.Y., et al.,
2009c. Trypanosoma cruzi in Brazilian Amazonia: Lineages TCI and TCIIa in wild
primates, Rhodnius spp. and in humans with Chagas disease associated with oral
transmission. Int. J. Parasitol. 39, 615–623.
Marinho, C.R.F., Bucci, D.Z., Dagli, M.L.Z., Bastos, K.R.B., Grisotto, M.G., Sardinha, L.R.,
et al., 2004. Pathology affects different organs in two mouse strains chronically
infected by a Trypanosoma cruzi clone: a model for genetic a studies of Chagas’
disease. Inf. Immun. 72, 2350–2357.
Martins, C., Baptista, C.S., Ienne, S., Cerqueira, G.C., Bartholomeu, D.C., Zingales, B.,
2008. Genomic organization and transcription analysis of the 195-bp satellite
DNA in Trypanosoma cruzi. Mol. Biochem. Parasitol. 160, 60–64.
Miles, M.A., Cedillos, R.A., Povoa, M.M., de Souza, A.A., Prata, A., Macedo, V., 1981. Do
radically dissimilar Trypanosoma cruzi strains (zymodemes) cause Venezuelan
and Brazilian forms of Chagas’ disease? Lancet 1, 1338–1340.
Miles, M.A., Cibulskis, R.E., 1986. Zymodeme characterization of Trypanosoma cruzi.
Parasitol. Today 2, 94–97.
Miles, M.A., Llewellyn, M.S., Lewis, M.D., Yeo, M., Baleela, R., Fitzpatrick, S., et al.,
2009. The molecular epidemiology and phylogeography of Trypanosoma cruzi
and parallel research on Leishmania: looking back and to the future.
Parasitology 136, 1509–1528.
Minning, T.A., Weatherly, D.B., Atwood, J., Orlando, R., Tarleton, R.L., 2009. The
steady-state transcriptome of the four major life-cycle stages of Trypanosoma
cruzi. BMC Genomics 10, 370.
Montilla, M., Guhl, F., Jaramillo, C., Nicholls, S., Barnabé, C., Bosseno, M.F., Brenière,
S.F., 2002. Isoenzyme clustering of Trypanosomatidae Colombian populations.
Am. J. Trop. Med. Hyg. 66, 394–400.
Ocaña-Mayorga, S., Llewellyn, M.S., Costales, J.A., Miles, M.A., Grijalva, M.J., 2010.
Sex, subdivision, and domestic dispersal of Trypanosoma cruzi lineage I in
Southern Ecuador. PLoS Negl. Trop. Dis. 4, e915.
Oliveira, R.P., Broude, N.E., Macedo, A.M., Cantor, C.R., Smith, C.L., Pena, S.D., 1998.
polymorphic microsatellites. Proc. Natl. Acad. Sci. USA 95, 3776–3780.
Pinto, A.S., de Lana, M., Bastrenta, B., Barnabé, C., Quesney, V., Noel, S., Tibayrenc, M.,
1998. Compared vectorialtransmissibility
genotypes of Trypanosoma cruzi in Triatoma infestans. Parasitol. Res. 84, 348–
Postan, M., McDaniel, J.P., Dvorak, J.A., 1987. Comparative studies of the infection of
Lewis rats with four Trypanosoma cruzi clones. Trans. R. Soc. Trop. Med. Hyg. 81,
Ramirez, J.D., Guhl, F., Rendon, L.M., Rosas, F., Marin-Neto, J.A., Morillo, C.A., 2010.
Chagas cardiomyopathy manifestations and Trypanosoma cruzi genotypes
circulating in chronic Chagasic patients. PLoS Negl. Trop. Dis. 4, e899.
Revollo, S., Oury, B., Laurent, J.P., Barnabé, C., Quesney, V., Carriere, V., Noel, S.,
Tibayrenc, M., 1998. Trypanosoma cruzi: impact of clonal evolution of the
parasite on its biological and medical properties. Exp. Parasitol. 89, 30–39.
Risso, M.G., Sartor, P.A., Burgos, J.M., Briceno, L., Rodriguez, E.M., Guhl, F., et al.,
2011. Immunological identification of Trypanosoma cruzi lineages in human
infection along the endemic area. Am. J. Trop. Med. Hyg. 84, 78–84.
Rodriguez, I.B., Botero, A., Mejia-Jaramillo, A.M., Marquez, E.J., Ortiz, S., Solari, A.,
Triana-Chavez, O., 2009. Transmission
determined by low-stringency single primer polymerase chain reaction and
Southern blot analyses in four indigenous communities of the Sierra Nevada de
Santa Marta. Colombia. Am. J. Trop. Med. Hyg. 81, 396–403.
Roellig, D.M., Brown, E.L., Barnabé, C., Tibayrenc, M., Steurer, F.J., Yabsley, M.J., 2008.
Molecular typing of Trypanosoma cruzi isolates, United States. Emerg. Infect. Dis.
Roellig, D.M., Yabsley, M.J., 2010. Short report: infectivity, pathogenicity, and
virulence of Trypanosoma cruzi Isolates from sylvatic animals and vectors, and
domestic dogs from the United States in ICR strain mice and SD strain rats. Am.
J. Trop. Med. Hyg. 83, 519–522.
Rougeron, V., De Meeus, T., Hide, M., Waleckx, E., Bermudez, H., Arevalo, J.L., et al.,
2009. Extreme inbreeding in Leishmania braziliensis. Proc. Natl. Acad. Sci. USA
Rozas, M., De Doncker, S., Adaui, V., Coronado, X., Barnabé, C., Tibyarenc, M., et al.,
2007. Multilocus polymerase chain reaction restriction fragment-length
polymorphism genotyping of Trypanosoma cruzi (Chagas disease): taxonomic
and clinical applications. J. Infect. Dis. 195, 1381–1388.
Rozas, N., De Doncker, S., Coronado, X., Barnabé, C., Tibyarenc, M., Solari, A.,
Dujardin, J.C., 2008. Evolutionary history of Trypanosoma cruzi according to
antigen genes. Parasitology 135, 1157–1164.
Ruvalcaba-Trejo, L.I., Sturm, N.R., 2011. The Trypanosoma cruzi Sylvio X10 strain
maxicircle sequence: the third musketeer. BMC Genomics 12, 58.
Sánchez-Guillén, M.C., Barnabé, C., Tibayrenc, M., Zavala-Castro, J., Totolhua, J.L.,
Méndez-López, J., et al., 2006. Trypanosoma cruzi strains isolated from human,
vector, and animal reservoir in the same endemic region in Mexico and typed as
T. cruzi I, discrete typing unit 1 exhibit considerable biological diversity. Mem.
Inst. Oswaldo Cruz 101, 585–590.
Saravia, N.G., Holguin, A.F., Cibulskis, R.E., D’Alessandro, A., 1987. Divergent
isoenzyme profiles of sylvatic and domiciliary Trypanosoma cruzi in the
eastern plains, piedmont, and highlands of Colombia. Am. J. Trop. Med. Hyg.
of pureand mixed clonal
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253
Souto, R.P., Fernandes, O., Macedo, A.M., Campbell, D.A., Zingales, B., 1996. DNA Download full-text
markers define two major phylogenetic lineages of Trypanosoma cruzi. Mol.
Biochem. Parasitol. 83, 141–152.
Souto, R.P., Zingales, B., 1993. Sensitive detection and strain classification of
Trypanosoma cruzi by amplification of a ribosomal RNA sequence. Mol. Biochem.
Parasitol. 62, 45–52.
Spotorno, O., Cordova, L., Solari, I., 2008. Differentiation of Trypanosoma cruzi I
subgroups through characterization of cytochrome b gene sequences. Inf.
Genet. Evol. 8, 898–900.
Steindel, M., Pacheco, L.K., Scholl, D., Soares, M., de Moraes, M.H., Eger, I., et al., 2008.
Characterization of Trypanosoma cruzi isolated from humans, vectors, and
animal reservoirs following an outbreak of acute human Chagas disease in
Santa Catarina State. Brazil. Diagn. Microbiol. Infect. Dis. 60, 25–32.
Stevens, J.R., Noyes, H.A., Schofield, C.J., Gibson, W., 2001. The molecular evolution
of trypanosomatidae. Adv. Parasitol. 48, 1–56.
Sturm, N.R., Campbell, D.A., 2010. Alternative lifestyles: the population structure of
Trypanosoma cruzi. Acta Trop. 115, 35–43.
Sturm, N.R., Vargas, N.S., Westenberger, S.J., Zingales, B., Campbell, D.A., 2003.
Evidence for multiple hybrid groups in Trypanosoma cruzi. Int. J. Parasitol. 33,
Svoboda, M., Virreira, M., Torrico, F., Truyens, C., Alonso-Vega, C., Solano, M., Carlier,
Y., 2005. Detection of the molecular heterogeneity of Trypanosoma cruzi. Rev.
Soc. Bras. Med. Trop. 38, 77–83.
Telleria, J., Biron, D.G., Brizard, J.P., Demettre, E., Seveno, M., Barnabé, C., Ayala, F.J.,
Tibayrenc, M., 2010. Phylogenetic character mapping of proteomic diversity
shows high correlation with subspecific phylogenetic diversity in Trypanosoma
cruzi. Proc. Natl. Acad. Sci. USA 107, 20411–20416.
Thomas, S., Martinez, L.L., Westenberger, S.J., Sturm, N.R., 2007. A population study
of the minicircles in Trypanosoma cruzi: predicting guide RNAs in the absence of
empirical RNA editing. BMC Genomics 8, 133.
Thomas, S., Westenberger, S.J., Campbell, D.A., Sturm, N.R., 2005. Intragenomic
spliced leader RNA array analysis of kinetoplastids reveals unexpected
transcribed region diversity in Trypanosoma cruzi. Gene 352, 100–108.
Tibayrenc, M., 1998. Genetic epidemiology of parasitic protozoa and other
infectious agents: the need for an integrated approach. Int. J. Parasitol. 28,
Tibayrenc, M., Ayala, F.J., 1988. Isozyme variability in Trypanosoma cruzi, the agent
of Chagas-disease – genetic, taxonomical, and epidemiological significance.
Evolution 42, 277–292.
Tibayrenc, M., Ayala, F.J., 1991. Towards a population genetics of microorganisms:
the clonal theory of parasitic protozoa. Parasitol. Today 7, 228–232.
Tibayrenc, M., Ayala, F.J., 2002. The clonal theory of parasitic protozoa: 12 years on.
Trends Parasitol. 18, 405–410.
Tibayrenc, M., Barnabé, C., Telleria, J., 2010. Reticulate evolution in Trypanosoma
cruzi: Medical and epidemiological implications. In: Telleria, J., Tibayrenc, M.
(Eds.), American Trypanosomiasis Chagas Disease One Hundred Years of
Research. Elsevier Inc.
Tibayrenc, M., Kjellberg, F., Ayala, F.J., 1990. A clonal theory of parasitic protozoa -
the population structures of Entamoeba, Giardia, Leishmania,
Plasmodium, Trichomonas, and
taxonomical consequences. Proc. Natl. Acad. Sci. USA 87, 2414–2418.
Tibayrenc, M., Ward, P., Moya, A., Ayala, F.J., 1986. Natural populations of
Trypanosoma cruzi, the agent of Chagas disease, have a complex multiclonal
structure. Proc. Natl. Acad. Sci. USA 83, 115–119.
medicaland their and
Tomasini, N., Lauthier, J.J., Rumi, M.M., Ragone, P.G., D’Amato, A.A., Brandan, C.P.,
et al., 2011. Interest and limitations of spliced leader intergenic region
sequences for analyzing Trypanosoma cruzi I phylogenetic diversity in the
Argentinean Chaco. Infect. Genet. Evol. 11, 300–307.
Tomazi, L., Kawashita, S.Y., Pereira, P.M., Zingales, B., Briones, M.R., 2009. Haplotype
distribution of five nuclear genes based on network genealogies and Bayesian
inference indicates that Trypanosoma cruzi hybrid strains are polyphyletic.
Genet. Mol. Res. 8, 458–476.
Triana, O., Ortiz, S., Dujardin, J.C., Solari, A., 2006. Trypanosoma cruzi: variability of
stocks from Colombia determined by molecular karyotype and minicircle
Southern blot analysis. Exp. Parasitol. 113, 62–66.
Vago, A.R., Andrade, L.O., Leite, A.A., Reis, A.D., Macedo, A.M., Adad, S.J., et al., 2000.
Genetic characterization of Trypanosoma cruzi directly from tissues of patients
with chronic Chagas disease. Am. J. Pathol. 156, 1805–1809.
Valente, S.A., Valente, V.C., Neves Pinto, A.Y., Barbosa César, M.J., Santos, M.P.,
Miranda, C.O., et al., 2009. Analysis of an acute Chagas disease outbreak in the
Brazilian Amazon: human cases, triatomines, reservoir mammals and parasites.
Trans. R. Soc.Trop. Med. Hyg. 103, 291–297.
Virreira, M., Alonso-Vega, C., Solano, M., Jijena, J., Brutus, L., Bustamante, Z., et al.,
DNA polymorphism of Trypanosoma cruzi. Am. J. Trop. Med. Hyg. 75,
Weatherly, D.B., Boehlke, C., Tarleton, R.L., 2009. Chromosome level assembly of the
hybrid Trypanosoma cruzi genome. BMC Genomics 10, 255.
Westenberger, S.J., Barnabé, C., Campbell, D.A., Sturm, N.R., 2005. Two hybridization
events define the population structure of Trypanosoma cruzi. Genetics 171, 527–
Westenberger, S.J., Cerqueira, G.C., El-Sayed, N.M., Zingales, B., Campbell, D.A.,
Sturm, N.R., 2006a. Trypanosoma cruzi mitochondrial maxicircles display
species- and strain-specific variation and a conserved element in the non-
coding region. BMC Genomics 7, 60.
Westenberger, S.J., Sturm, N.R., Campbell, D.A., 2006b. Trypanosoma cruzi 5S rRNA
arrays define five groups and indicate the geographic origins of an ancestor of
the heterozygous hybrids. Int. J. Parasitol. 36, 337–346.
Yeo, M., Acosta, N., Llewellyn, M., Sanchez, H., Adamson, S., Miles, G.A.J., et al., 2005.
Origins of Chagas disease: didelphis species are natural hosts of Trypanosoma
cruzi I and armadillos hosts of Trypanosoma cruzi II, including hybrids. Int. J.
Parasitol. 35, 225–233.
Yeo, M., Mauricio, I.L., Messenger, L.A., Lewis, M.D., Llewellyn, M.S., Acosta, N., et al.,
2011. Multilocus sequence typing for lineage assignment and high resolution
diversity studies in Trypanosoma cruzi. PLoS Negl. Trop. Dis. 5, e1049.
Zafra, G., Mantilla, J.C., Valadares, H.M., Macedo, A.M., Gonzalez, C.I., 2008. Evidence
of Trypanosoma cruzi II infection in Colombian chagasic patients. Parasitol. Res.
Zingales, B., Andrade, S.G., Briones, M.R.S., Campbell, D.A., Chiari, E., Fernandes, O.,
et al., 2009. A new consensus for Trypanosoma cruzi intraspecific nomenclature:
second revision meeting recommends TcI to TcVI. Mem. Inst. Oswaldo Cruz 104,
Zingales, B., Pereira, E.S., Oliveira, R.P., Almeida, K.A., Umezawa, E.S., Souto, R.P.,
et al., 1997. Trypanosoma cruzi genome project: biological characteristics and
molecular typing of Clone CL Brener. Acta Trop. 68, 159–173.
Zingales, B., Stolf, B.S., Souto, R.P., Fernandes, O., Briones, M.R., 1999. Epidemiology,
biochemistry and evolution of Trypanosoma cruzi lineages based on ribosomal
RNA sequences. Mem. Inst. Oswaldo Cruz 94, 159–164.
inBolivia is notassociated with
B. Zingales et al./Infection, Genetics and Evolution 12 (2012) 240–253