enzootiology of trypanosoma evansi in pantanal, brazil
TRANSCRIPT
Enzootiology of Trypanosoma evansi in
Pantanal, Brazil
H.M. Herreraa,*, A.M.R. Davilab, A. Noreka, U.G. Abreuc,S.S. Souzab, P.S. D’Andread, A.M. Jansena
aLaboratorio de Biologia de Tripanosomatıdeos, Departamento de Protozoologia, FIOCRUZ/RJ,
Pavilhao Carlos Chagas 3 Andar, Av Brasil 4365, CEP 21045-900, Rio de Janeiro, BrazilbLaboratorio de Biologia Molecular de Tripanosomatıdeos, Departamento de Bioquımica e
Biologia Molecular, FIOCRUZ/RJ, Pavilhao Carlos Chagas 3 Andar,
Av Brasil 4365, CEP 21045-900, Rio de Janeiro, BrazilcCentro de Pesquisa Agropecuaria do Pantanal, EMBRAPA/Pantanal, Rua 21 de Setembro,
1880, CEP 79320-900, Corumba, MS, BrazildLaboratorio de Doencas Endemicas, Departamento de Medicina Tropical, FIOCRUZ/RJ, Pavilhao
Carlos Chagas 3 Andar, Av Brasil 4365, CEP 21045-900, Rio de Janeiro, Brazil
Received 4 July 2004; accepted 26 July 2004
Abstract
In order to better understand the enzootiology of trypanosomiasis caused by Trypanosoma evansi
in the Brazilian Pantanal we examined domestic and wild mammals by microhematocrit centrifuge
technique (MHCT), immunofluorescence antibody test (IFAT) and polymerase chain reaction (PCR).
T. evansi infection was detected in all species sampled with exception of the sheep and the feral pig.
High parasitemias were observed in capybaras (5/24), coatis (18/115), horses (31/321) and dogs (3/
112). Among these species, only the capybaras did not develop anemia. Low parasitemias, only
detected by PCR, were found in buffaloes (18/43), bovines (29/331), marsupials (1/4), small rodents
(14/67), bats (7/18), and one armadillo (1/8). The highest prevalence of T. evansi infection was
recorded in horses (73%), although no neurological signs in infected horses were observed. Diagnosis
through standard parasitological tests and IFAT should be used with caution since they may overlook
comprovedly infected horses. The relationship between ranch management and T. evansi infection in
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Veterinary Parasitology 125 (2004) 263–275
* Corresponding author. Tel.: +55 21 2598 4324; fax: +55 21 2560 6572.
E-mail address: [email protected] (H.M. Herrera).
0304-4017/$ – see front matter # 2004 Elsevier B.V. All rights reserved.
doi:10.1016/j.vetpar.2004.07.013
horse was investigated. The importance of other transmission mechanisms apart from the tabanids
and reservoir hosts are discussed.
# 2004 Elsevier B.V. All rights reserved.
Keywords: Trypanosoma evansi; Natural infection; Reservoir; Diagnosis; Pantanal; Brazil
1. Introduction
Among the pathogenic trypanosome species, Trypanosoma evansi (Trypanosomatidae,
Kinetoplastida) is known to have a large diversity of mammalian hosts and is the most
important disease producing agent throughout the tropical and subtropical areas of the
world. The main difference with other trypanosomatids is the lack of maxicircles kDNA.
Consequently there is no development of T. evansi in the vector. Indeed, this parasite is
adapted to mechanical transmission and has a broad host range in a wide geographical
distribution (Hoare, 1972; Lun and Desser, 1995).
This flagellate is transmitted by hematophagous flies, mainly Tabanus sp., through oral
infection by ingestion of meat or blood from infected animals and by vampire bats
(Desmodus rotundus) in South America (Hoare, 1965, 1972; Losos, 1986).
In spite of being considered a hemoflagellate, T. evansi (Salivaria, Trypanozoon) is a
protozoan parasite of both intra and extra vascular fluids (Sudarto et al., 1990). One of the
most interesting aspects of T. evansi is its ability to periodically switch its major variant
surface glycoprotein (VSG), producing relapses of parasitemias. Parasitemic waves have
been reported with Brazilian T. evansi isolates in experimental infections (Aquino et al.,
1999; Queiroz et al., 2000; Herrera et al., 2001).
There are considerable differences in the severity of syndromes caused by T. evansi
infections in the different geographical areas of its occurrence, depending on the virulence
of the strain and the susceptibility of the host. Anemia is the main outcome of infection.
Consequently, it has been suggested that the resistance to developing anemia, as well as
control of parasitemia, reflect the degree of tolerance to infection by the hosts (Trail et al.,
1990). Sick animals may display fever, general loss of condition and immunosupression
(Hoare, 1972; Holland et al., 2001).
The diagnosis of T. evansi infection is still difficult because the clinical signs are varied
and non-specific and, in enzootic areas, the natural hosts frequently present mild chronic
forms of the disease (Losos, 1986; Franke et al., 1994a; Arias et al., 1997).
The routine of T. evansi diagnosis is mainly based on finding the flagellates in wet films,
smears or by the michrohematocrit test. These methods are specific but less sensitive,
principally in detecting parasites during low levels of parasitemia (Woo, 1970; Murray et
al., 1981; Monzon et al., 1986).
Serological methods have been used for mass screening, however, antibodies may be
missing due to serological latency (Franke et al., 1994b; Aquino et al., 1999; Herrera et al.,
2001; Wernery et al., 2001). The amplification of repetitive DNA satellite regions of
Trypanozoon have shown to be very specific and sensitive and has been used in large
trypanosomiasis epidemiological approaches in both animals and humans (Moser et al.,
1989; Masiga et al., 1992; Kanmogne et al., 1996; Katakura et al., 1997).
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275264
In the Brazilian Pantanal, T. evansi is enzootic, infecting domestic and wild animals
(Nunes et al., 1993). This parasite causes a severe disease in horses locally called ‘‘Mal de
Cadeiras’’ due to the nervous symptomatology characterized by hind limbs paresis,
resulting in a staggering gait (Silva et al., 1995a). Two forms of the disease are described in
Brazil due to T. evansi infections: acute syndrome that produces early death in horses and
dogs if untreated (Silva et al., 1995c, 1996; Aquino et al., 1999), and chronic, affecting
mainly capybaras (Hydrochaeris hydrochaeris) and coatis (Nasua nasua) (Franke et al.,
1994b; Herrera et al., 2002).
Outbreaks of ‘‘Mal de Cadeiras’’ have occurred periodically among Pantanal livestock
since the beginning of the 19th century (Wilcox, 1992). This disease is a problem of great
economic importance due to high cost of treatment and death of sick animals (Silva et al.,
1995b; Seidl et al., 1998, 2001).
The large seasonal floodplain called Pantanal covers about 140,000 km2. It is located
near the geographical center of South America and its economic activities are mainly based
on cattle ranches. The livestock population is estimated in 4,000,000 cattle, 5000 buffaloes
and 140,000 horses. In spite of not being considered of economic importance, sheep are
also raised in the region (Cadavid Garcia, 1986). Wildlife is abundant in this region
(Lourival and Fonseca, 1997).
The Pantanal wetlands contain 10 sub-regions (Fig. 1) that differ in degree of vegetation,
flooding and physiognomy. During the wet season most of the grasslands are flooded and
during the dry season water remains only in a few pools. The length and severity of flooding
and drought in the Pantanal vary, not only from year to year, but also from sub-region to sub-
region (Adamoli, 1987). Outbreaks of disease caused by T. evansi have been reported in the
last two decades in the study area and are associated mainly with the rainy season, when
tabanids are abundant (Silva et al., 1995b; Davila et al., 1999; Barros and Foil, 1999).
Capybaras and coatis have been considered the main wild reservoirs for ‘‘Mal de Cadeiras’’
(Nunes et al., 1993; Silva et al., 1999). Nevertheless nothing is known about the importance of
other species concerning the T. evansi enzootiology in this area.
In this scenario, many aspects concerning the epizootiology of ‘‘Mal de Cadeiras’’ in the
Pantanal region are still unknown. The aim of this research was to improve the knowledge
of the variables that are involved in the transmission cycle of T. evansi in the southern
Pantanal. The possible role of infected wild and domestic animals as reservoir hosts and the
importance of ranch management in the epidemiology of ‘‘Mal de Cadeiras’’ is discussed.
2. Material and methods
2.1. Study area
The sampled area covered approximately 1000 km2 and is located at the Nhecolandia sub-
region, 100 km west of Corumba city (Fig. 1). In this region, livestock shares the same habitat
with wild animals. During the rainy season a large part of the area is flooded and much of the
non-flooded higher ground (‘‘cordilheira’’) is interspersed throughout the region. The
vegetation includes mixed scrub with grassland (woodland savannah), open grass fields and
‘‘cordilheiras’’, which is are covered by deciduous forests (Adamoli, 1987).
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275 265
2.2. Field procedures
Blood samples were collected during four excursions between February 2000 and
March 2001. Samples from domestic animals (horses, dogs, cows, buffaloes and sheep)
were collected from the jugular vein. Wild mammals were trapped under government
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275266
Fig. 1. Map showing the Pantanal, sub-regions and location of the study area.
authorization of the Brazilian Environment Institute (licenses no. 229/2000 and 228/2000),
immobilized with zolazepan and tiletamina cloridrat (Zoletil1 50), venipunctured
and immediately released after blood sampling and recovery from anesthesia. Blood
was collected into vacutainer tubes or microhematocrit containing ethylenediaminete-
traacetic acid (EDTA) as the anticoagulant, kept on ice and processed on the same day of
collection.
Each animal was sampled on only one occasion. In the field, T. evansi was detected by
microhematocrit centrifuge technique (MHCT) (Woo, 1970). The hematocrit value was
estimated by the packed cell volume (PCV), using the standard microhematocrit method
according to Schalm et al. (1975). Samples for the molecular diagnosis (PCR) were
obtained from 10 ml of whole blood dropped on a filter paper confetti. Samples for
serological test (IFAT) were stored at �4 8C until used.
2.3. Anemia
Packed cell volume (PCV) values were used as an index of anemia. The mean PCV
values of T. evansi for negative animals in the serological, parasitological and molecular
tests were considered as normal values.
2.4. Parasitemia
Animals with positive MHCT tests were considered as having high parasitemias, since
positive MHCT test only detects parasitemias from 104 parasites/ml (Woo, 1970). Animals
of MCHT test negative that displayed positive PCR test were considered as displaying low
parasitemias.
2.5. DNA extraction
DNA extraction was performed with Chelex-1001 1% according to Walsh et al. (1991)
with minor modifications by De Almeida et al. (1997), in brief: 500 ml of Milli-Q water
was used for the initial wash of the confetti during 30 min. After 15 min the tubes were
inverted 2–3 times, then, after 10 min of centrifugation at 12,000 rpm in an eppendorf
centrifuge, supernatant was discarded and 100 ml Chelex-1001 1% added. The tubes were
shaked manually, incubated at 56 8C during 30 min, boiled for 8 min and vortexed for
2 min. After a final 5 min centrifugation at 12,000 rpm, 80 ml of supernatant was placed in
a fresh tube and stored at �20 8C.
2.6. DNA amplification
A polymerase chain reaction (PCR) was performed with primer sets (TBR1 and TBR2)
specific for Trypanozoon satellite DNA regions, according to Masiga et al. (1992).
The PCR assay showed to be sensitive enough to detect 10 fg of T. evansi DNA. The
primer sets TBR1 and TBR2 were tested using T. rangeli, T. cruzi, Leishmania braziliensis,
Crithidia fasciculata and Herpetomonas muscarum as controls. The results showed a
single band expected (164 bp) only for T. evansi (paper in preparation).
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275 267
PCR was performed in Perkin Elmer 96001 PCR machines and for each set of
amplification reactions genomic DNA of T. evansi (10–20 ng) was used as positive control
and double distilled water was used as negative control. Standard PCR amplification was
carried out in 10 ml reaction mixtures, each containing 1 ml of DNA as template, 1.5 mM
MgCl2, 1 mM of each primer, 200 mM of each dNTP, 0.5 unit Taq polymerase, and 5%
DMSO. Amplification conditions were as follows: 95 8C for 5 min (initial denaturation),
then 35 cycles of 95 8C for 1 min (denaturation), followed by 55 8C for 1 min (annealing)
and 72 8C for 1 min (extension), with a final extension step of 10 min at 72 8C. Finally, all
the 10 ml of each amplified sample were analyzed by electrophoresis in 2% agarose gel
containing ethidium bromide 0.5 mg/ml and photographed under ultraviolet light.
2.7. Serological test
Immunofluorescence antibody test (IFAT) for detecting IgG was performed for horses,
dogs and coatis. The IFAT reaction was conducted according to the protocol utilized by
Camargo (1964). Negative control serum samples were obtained from animals located
in areas where T. evansi does not occur and positive serum samples were obtained from
the parasitological positive field animals. Standardization of the protocol was undertaken
in order to establish suitable working dilutions for each specific fluorescein antibody
conjugate. The following conjugates were used: anti-horse IgG (Sigma1) for horses,
anti-dog IgG for dogs and anti-raccoon IgG (Kirkegaard & Perry Laboratories1) for
coatis.
The cut off for each animal species was considered as the lower titer of serum samples in
which parasites could be detected by MHCT or PCR. The reaction was taken as being
positive when the dilution of serum was respectively � 1:10 for horses and � 1:40 for dogs
and coatis. The reaction was performed in twofold dilutions starting from 1:5 to 1: 2560.
2.8. Statistical analysis
To compare between MHCT and PCR positive results for every species sampled the chi-
square (x2) test was employed. The t-test (equivalent variances) was used to evaluate a
possible correlation between high parasitemias, assayed by MHCT, and anemia, evidenced
by the PCV values. The comparison of equine IFAT results with their respective values of
PCV was performed by means of t-test.
3. Results
T. evansi infection was detected by at least one of the three methods employed in all
species sampled, except for feral pigs and sheep. The pattern and prevalence of infection
differed according to the animal species (Table 1).
Infection with low parasitemias as detected by positive PCR test and negative MHCT
test, was noticed in bovines, buffaloes, bats, small mammals and one armadillo. It
is noteworthy that buffaloes displayed high T. evansi prevalence as detected by PCR
(Table 1).
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Parasitemias by positive MHCT test were observed in capybaras, coatis, horses and
dogs (Table 1). Coatis and capybaras showed the higher parasitemias since the number of
animals positive by MHCT and number of animals positive by PCR was not significantly
different (P < 0.05).
Horses presented the highest prevalence of T. evansi infection among all species
sampled. The higher equine seroprevalence levels with parasitemias assayed by MHCT
were observed in farms where no control of equine infectious anemia (EIA) was performed
(Table 2). A strong relationship between horse management and high parasitemias could be
established since animals from ranches where a program of EIA control is conducted, did
not present positive MHCT test and displayed the lower seroprevalence rates (Table 2).
Moreover, means of hematocrit values were significantly different (P < 0.05) in horses
derived from farms that control IEA (33.4%) than horses derived from farms that not
control IEA (30.2%), to anyone of MHCT positive and negative animals.
All infected coatis and capybaras even those with positive with MHCT were in apparent
good health condition. No neurological signs were observed in horses, even in animals that
displayed high parasitemias. Dogs with high parasitemias presented severe clinical
symptoms such as emaciation, conjunctivitis and facial edema.
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275 269
Table 1
Prevalence of T. evansi infection in mammals in the Nhecolandia sub-region, Pantanal
Species Total sampled Number of positives/percentual of positivity
MHCT PCR IFAT
Horse 321 31/9.6 112/34.9 236/73.5
Dog 112 3/2.7 11/9.8 26/23.2
Coati 115 18/15.6 22/19.1 35/30.4
Capybara 24 5/20.8 7/29.2 4/33.3
Cattle 331 0 29/8.8 nd
Buffalo 43 0 18/41.9 nd
Small mammals
Thrichomys sp. 46 0 7/15.2 nd
Clyomys sp. 11 0 2/18.2 nd
Oecomys sp. 7 0 3/42.8 nd
Dasyprocta sp. 3 0 2/66.7 nd
Marsupials
Gracilinanus sp. 2 0 0 nd
Monodelphis sp. 2 0 1/50.0 nd
Bats
Artibeus sp. 6 0 3/50.0 nd
Platyrrhinus sp. 5 0 1/20.0 nd
Carollia sp. 3 0 1/33.3 nd
Myotis sp. 2 0 2/100.0 nd
Noctilio sp. 2 0 0 nd
Armadillos
Euphractus sp. 8 0 1/12.5 nd
Feral pigs
Sus scorfa feral 10 0 0 nd
Sheep 40 0 0 nd
MHCT: microhematocrit centrifuge technique; PCR: polymerase chain reaction; IFAT: immunofluorescence
antibody test; nd: not done.
Anemia, as proved by low PCV values, was observed in all examined dogs independent
of the presence of T. evansi, probably as a consequence of the bad conditions in which they
were maintained. Anemia was not recorded in capybaras since no significative difference
(P < 0.05) was observed between means of hematocrit values from MHCT negatives and
positives animals. A direct correlation between high parasitemias and anemia was observed
in horses and coatis since we found significative difference (P < 0.05) between means of
hematocrit values from MHCT negatives and positives animals. It is important to note that
no correlation between anemia and serum positivity in horses was observed (P < 0.05).
IFAT serological titers ranged from 1:10 to 1:1280 in horses and from 1:40 to 1:640 in
coatis and dogs. Serological negative animals with PCR positive results were observed in
horses (13%), dogs (3%) and coatis (4%) (Fig. 2). Nevertheless, we found serological
positive results with PCR negative in horses (55%), dogs (55%) and coatis (44%) (Fig. 3).
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275270
Table 2
Prevalence of T. evansi infection in horses in the Nhecolandia sub-region, Pantanal
Ranch Total sampled Number of positives/percentual of positivity
MHCT PCR IFAT
R1 48 0 1/2.1 21/43.7
R2 18 0 0 8/44.4
R3 119 26/21.8 68/57.1 85/71.4
R4 47 1/2.1 16/34.0 43/91.5
R5 40 3/7.5 23/57.5 36/90.0
R6 49 0 4/8.2 43/87.7
Total 321 31/9.6 112/34.9 236/73.5
R1 and R2 are ranches that control EIA while R3, R4, R5 and R6 do not control EIA.
Fig. 2. Infections observed by positive PCR tests in serologically negative samples. PCR: polymerase chain
reaction; IFAT: immunofluorescence antibody test.
Fig. 3. Low parasitemias as proved by PCR negatives in serological positive samples. PCR: polymerase chain
reaction; IFAT: immunofluorescence antibody test.
Infection by T. evansi was observed in equines, dogs, coatis, capybaras and small
mammals in both the rainy and dry season.
4. Discussion
Our results showed that in the Pantanal region T. evansi infects a wide range of host
species of domestic and wild mammals and causes various degree of parasitemia and
clinical signs. The distinct habitats and behavior patterns of small rodents, bats and
armadillo suggest that there are still unknown factors underlying the transmission cycles.
Low and cryptic parasitemias observed in animals of all T. evansi infected species
suggest their importance in the maintenance of the parasite in nature. Sub-patent
parasitemias have been widely described in T. evansi enzootic areas and some factors were
associated to this feature: difference in virulence of strains, host susceptibility, chronic
phase or even individual nutritional status (Monzon et al., 1986; Aquino et al., 1999;
Herrera et al., 2001). Considering that stress may result in weakness of the infected animals
and consequently low immunity and exacerbation of parasitemia (Luckins et al., 1979;
Payne et al., 1991), the presented data suggest that all T. evansi infected species may be
involved in the transmission cycle of the parasite.
The higher seroprevalence observed in horses suggests that they are more exposed to T.
evansi infection than the other species sampled. The higher prevalence assessed by IFAT in
comparison with the other two tests employed was expected. Horses that presented IFAT
positive test and negative results by PCR and MHCT probably reflect animals submitted to
chemotherapy. High horse seroprevalence rates have also been reported in other areas of
the Pantanal and South America reinforcing the importance of the enzooty in continent
(Franke et al., 1994a; Monzon et al., 1995; Reyna-Bello et al., 1998).
The high number of infected horses with low parasitemias, only detected by PCR reflect
cryptic infections probably as the result of incorrect treatment or the consequence of a mild
course of infection. These low parasitemias explain why no correlation between low PCV
values (anemia) and serological positive horses was found. In addition, the presented data
showed that treatment of T. evansi infected animals based only on screening of animals
with low hematocrit values will certainly overlook infected animals.
Negative IFAT tests in PCR positive testing animals may reflect serological latency or
false negative results. Indeed, IgG plasmatic concentration reach high titer after 3 weeks
(Reyna-Bello et al., 1998; Aquino et al., 1999; Wernery et al., 2001) and has an important
role in the control of ‘‘Mal de Cadeiras’’ because animals with recent infections are not
detected by serological screening. These findings showed the importance of using a more
sensitive and precocious test such as PCR.
The high prevalence rate of horse infection in all periods of sampling suggest two
situations: these animals may become infected during the winter or they may be developing
a long lasting course of T. evansi infection. These data associated with the absence of
nervous symptomatology of ‘‘Mal de Cadeiras’’ in this study support the importance of
horses themselves as reservoir for T. evansi.
The higher equine prevalences of T. evansi infection in farms without control for EIA,
associated to MHCT positive results uniquely evident in these areas, suggest a synergism
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between these two immunosuppressive diseases. Consequently, it is evident that a good
horse management is fundamental for ‘‘Mal de Cadeiras’’ control.
Although the importance of wild mammals in the maintenance of T. evansi in the natural
environment is recognized, a relationship between equine trypanosomiasis prevalence and
proximity of infected wild mammals was not established. We found low horse
seroprevalences with negative MHCT and PCR in areas where coatis and capybaras
displayed high T. evansi prevalence rates and parasitemias. Similar data were recorded in
the northern Pantanal (Franke et al., 1994a). These data point that overlapping T. evansi
transmission cycles may not occur in the Pantanal.
The rapid death of T. evansi infected dogs may explain the low prevalence rates found
during this study. Indeed, these dogs are stray animals and when sick they are not treated.
Our data concerning coatis indicate that animals with high parasitemias develop
anemia. Anemia, biochemical and pathologic changes have been reported in coatis
naturally and experimentally infected with T. evansi (Silva et al., 1999; Herrera et al., 2001,
2002). Therefore, since high prevalences rates were found in the Pantanal region, this
parasite may be acting as an important agent of selection and density control of the free-
living coatis populations.
The high prevalence of T. evansi infection found in capybaras and coatis during the dry
season suggests that these species indeed develop a long lasting course of infection in
nature. However, transmission due to other mechanisms other than bloodsucking flies
should not be ruled out.
Concerning capybaras, we concluded that this caviomorph rodent plays an important
role in the dispersion and maintenance of T. evansi in the Pantanal environment due to: (a)
high T. evansi prevalence rates; (b) the degree of tolerance to infection due to resistance to
develop anemia associated to high parasitemias; (c) the great population density in the
Pantanal region; (d) long lasting patent parasitemia and (e) the absence of clinical
symptoms. Infected but apparently healthy capybaras were also recorded in the Pantanal
and in Venezuela (Morales et al., 1976; Franke et al., 1994a; Arias et al., 1997).
Furthermore, anemia as evaluated by hematocrit values, was not reported in experimentally
infected capybaras (Franke et al., 1994b).
The oral route may be important in the dispersion of T. evansi infection in dogs, coatis
and capybaras. They may become infected as a consequence of their frequent fights.
Moreover, gregarious species such as coatis and capybaras have an aggressive behavior and
may transmit T. evansi among themselves by the oral route, maintaining the infection in the
social group. Therefore, since dogs remain close to the horseman, during the vector season
they can act as a link between wild and domestic animals playing an important role in the
epizootiology of ‘‘Mal de Cadeiras’’.
The parasitemias detected only by PCR in bovines and buffaloes demonstrate that these
species also play a role in the maintenance of T. evansi in the Pantanal due to their high
population density. These species may serve as source of infection for vampire bats and
therefore should be diagnosed and treated. Low parasitemias in bovines seem to be
common in the Pantanal region since negative MHCT test was also observed in
seropositive bovines in the northern Pantanal (Franke et al., 1994a).
The detection of T. evansi infection in small rodents, marsupials, non-hematophagous
bats and armadillo is puzzling and places new questions concerning the transmission of this
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275272
flagellate in the Pantanal region. Since nocturnal species spend the day in their holes and
Tabanidae has diurnal/crepuscular activity, it is possible that other vectors may be involved
in the transmission of T. evansi.
Considering that natural infection was reported for pigs and sheep in South America and
Asia (Lun et al., 1993; Vokaty et al., 1993), our negative results concerning these species
show that transmission cycle of T. evansi is peculiar for the different enzootic areas.
The present data suggest that for the T. evansi epidemiological survey in the Pantanal
region two or more techniques should be used to diagnose reservoir hosts. Diagnosis
through standard parasitological tests should be used with caution when it is impossible
perform PCR and seronegatives animals shall be re-tested after 30 days due to serology
latency.
In spite of the recent trend in not considering virulence and pathogenicity as a parameter
to evaluate a sign of an ancient host–parasite interaction, the observation of pathogenic
features of T. evansi infection in capybaras and coatis associated with prehistoric
biogeographical events, made us to speculate about the time-scale of their relationships.
African rodent caviomorphs (primitive capybara ancestor), that probably have an
ancient coevolution history with T. evansi, reached the South American continent 37
million years ago (mya). Placental carnivores arrived in South America about 3–3.5 mya
(Flynn and Wyss, 1998). Therefore, the T. evansi association with coatis is probably more
recent than with capybaras and, although these two wildlife species develop chronic
infections, coatis display anemia and pathological changes while infected capybaras do not
have clinical signs and anemia, withstanding high parasitemias (Franke et al., 1994b;
Herrera et al., 2001, 2002). This propose that T. evansi infection in Neotropical mammals
precedes the South American colonization by the Spanish Settlers.
Acknowledgements
To ranches and local people for field assistance. Supported by Conselho Nacional de
Pesquisa e Desenvolvimento; Instituto Oswaldo Cruz/Fiocruz; Programa de Apoio a
Pesquisa Estrategica em Saude/Fiocruz and Conservation Internacional do Brasil.
References
Adamoli, J., 1987. Fisiografia do Pantanal. In: Allem, A.C., Valls, J.F.M. (Eds.), Recursos Forrageiros Nativos do
Pantanal Matograssense. DDT-EMBRAPA, Brasılia, pp. 23–25.
Aquino, L.P., Machado, R.Z., Alessi, A.C., Marques, L.C., de Castro, M.B., Malheiros, E.B., 1999. Clinical,
parasitological and immunological aspects of experimental infection with Trypanosoma evansi in dogs. Mem.
Inst. Oswaldo Cruz. 94, 255–260.
Arias, J.F., Garcia, F., Riveira, M., Lopez, R., 1997. Trypanosoma evansi in capybara from Venezuela. J. Wildl.
Dis. 33, 359–361.
Barros, T., Foil, L., 1999. Seasonal occurrence and relative abundance of Tabanidae (DIPTERA) from the Pantanal
region, Brazil. Mem. Enthomol. Int. 14, 387–396.
Cadavid Garcia, E.A., 1986. Estudo Tecnico Economico da Pecuaria Bovina de Corte do Pantanal Mato-
Grossense. EMBRAPA/CPAP, Docum. 4, 126–127.
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275 273
Camargo, M.E., 1964. Improved technique of indirect immunofluorescence for serological diagnosis for
toxoplasmosis. Rev. Inst. Med. Trop. Sao Paulo 3, 117–118.
Davila, A.M.R., Souza, S.S., Campos, C., Silva, R.A., 1999. The seroprevalence of equine trypanosomosis in the
Pantanal. Mem. Inst. Oswaldo Cruz. 94, 199–202.
De Almeida, P.J., Ndao, M., Van Meirvenne, N., Geerts, S., 1997. Diagnostic evaluation of PCR in goats
experimentally infected with Trypanosoma vivax. Acta Trop. 66, 45–50.
Flynn, J.J., Wyss, A.R., 1998. Recent advances in South American mammalian paleontology. Tree 13, 449–454.
Franke, C.R., Greiner, M., Mehlitz, D., 1994a. Investigations on naturally occurring Trypanosoma evansi
infections in horses, cattle, dogs and capybaras (Hydrochaeris hydrochaeris) in Pantanal de Pocone (Mato
Grosso, Brasil). Acta Trop. 58, 159–169.
Franke, C.R., Greiner, M., Mehlitz, D., 1994b. Monitoring of clinical, parasitological and serological parameters
during an experimental infection of cabybaras (Hydrochaeris hydrochaeris) with Trypanosoma evansi. Acta
Trop. 58, 171–174.
Herrera, H.M., Alessi, A.C., Marques, L.C., Santana, A.E., Aquino, L.P.C.T., Menezes, R.F., Moraes, M.A.V.,
Machado, R.Z., 2002. Experimental Trypanosoma evansi infection in South American coati (Nasua nasua):
hematological, in blood biochemical and histopathological changes. Acta Trop. 81, 203–210.
Herrera, H.M., Aquino, L.P.C.T., Menezes, R.F., Marques, L.C., Moraes, M.A.V., Werther, K., Machado, R.Z.,
2001. Trypanosoma evansi experimental infection in South American coati (Nasua nasua): clinical, para-
sitological and humoral immune response. Vet. Parasitol. 102, 209–216.
Hoare, C.A., 1965. Vampire bats as vectors and hosts of equine and bovine trypanosomes. Acta Trop. 22, 204–216.
Hoare, C.A., 1972. The Trypanosomes of Mammals: A Zoological Monograph, Blackwell Scientific Publications,
Oxford.
Holland, W.G., My, L.N., Dung, T.V., Thanh, N.G., Tam, P.T., Vercruysse, J., Goddeeris, B.M., 2001. The
influence of T. evansi infection on the immuno-responsiveness of experimentally infected water buffaloes. Vet.
Parasitol. 102, 225–234.
Kanmogne, G.D., Asonganyi, T., Gibson, W.C., 1996. Detection of Trypanosoma brucei gambiense, in
serologically positive but aparasitaemic sleeping-sickness suspects in Cameroon, by PCR. An. Trop. Med.
Parasitol. 90, 475–483.
Katakura, K., Lubinga, C., Chitambo, H., 1997. Detection of Trypanosoma congolense and T. brucei subspecies in
cattle in Zambia by polymerase chain reaction from blood collected on a filter paper. Parasitol. Res. 83, 241–
245.
Losos, G.J., 1986. Infectious Tropical Diseases of Domestic Animals, Longman Scientific & Technical,
Essex.
Lourival, R.F.F., Fonseca, G.A.B., 1997. Analise de Sustentebilidade do Modelo de Caca Tradicional, no Pantanal
da Nhecolandia, Corumba, MS. In: Valladares-Padua, C., Bodmer, R.E., Cullen Jr., L. (Eds.), MCT-CNPq,
Manejo e Conservacao de Vida Silvestre no Brasil. Sociedade Civil Mamiraua, pp. 123–172.
Luckins, A.G., Boid, R., Era, P., Mahmoud, M.M., el Malik, K.H., Gray, A.R., 1979. Serodiagnosis of infection
with Trypanosoma evansi, in camels in the Sudan. Trop. Anim. Health Prod. 11, 1–12.
Lun, Z.R., Desser, S.S., 1995. Is the broad range of hosts and geographical distribution of Trypanosoma evansi
attributable to the loss of maxicircle kinetoplast DNA? Parasitol. Today 11, 131–133.
Lun, Z.R., Fang, Y., Wang, C.J., Brun, R., 1993. Trypanosomiasis of domestic animals in China. Parasitol. Today
9, 41–45.
Masiga, D., Smyth, A.J., Hayes, P., Bromidge, T.J., Gibson, W.C., 1992. Sensitive detection of trypanosomes in
tsetse flies by DNA amplification. Int. J. Parasitol. 22, 909–918.
Monzon, C.M., Mancebo, O.A., Brem, J.J., 1986. Evaluacion de tres cepas de Trypanosoma equinum em ratones.
Su aplicacion al diagnostico parasitologico. Vet. Arg. III, 651–658.
Monzon, C.M., Hoyos, C.B., Jara, G.A., 1995. Outbreaks of equine trypanosomiasis caused by Trypanosoma
evansi in Formosa Province, Argentina. Rev. Sci. Tech. Sep. 14, 747–752.
Morales, G.A., Wells, E.A., Angel, D., 1976. The capybara (Hydrochaeris hydrochaeris) as a reservoir for
Trypanosoma evansi. J. Wildl. Dis. 12, 572–574.
Moser, D.R., Cook, G.A., Ochs, D.E., Bailey, C.P., McKane, M.R., Donelson, J.E., 1989. Detection of
Trypanosoma congolense and Trypanosoma brucei subspecies by DNA amplification using the polymerase
chain reaction. Parasitology 99, 57–66.
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275274
Murray, M., Clifford, D.J., Gettinby, G., Snow, W.F., McIntyre, W.I., 1981. Susceptibility to African trypano-
somiasis of N’Dama and Zebu cattle in an area of Glossina morsitans submorsitans challenge. Vet. Rec. 109,
503–510.
Nunes, V.L.B., Oshiro, E.T., Dorval, M.E.C., Garcia, L.A.M., Da Silva, A.A.P., Bogliolo, A.R., 1993. Investigacao
epidemiologica sobre Trypanosoma (trypanozoon) evansi no Pantanal sul-matogrossense. Estudos de reser-
vatorios. Braz. J. Vet. Parasitol. 2, 41–44.
Payne, R.C., Sukanto, I.P., Djauhari, D., Partoutomo, S., Wilson, A.J., Jones, T.W., Boid, R., Luckins, A.G., 1991.
Trypanosoma evansi infection in cattle, buffaloes and horses in Indonesia. Vet. Parasitol. 38, 109–119.
Queiroz, A.O., Cabello, P.H., Jansen, A.M., 2000. Biological and biochemical characterization of isolates of
Trypanosoma evansi from Pantanal of Matogrosso – Brazil. Vet. Parasitol. 92, 107–118.
Reyna-Bello, A., Garcia, F.A., Rivera, M., Sanso, B., Aso, P.M., 1998. Enzyme-linked immunosorbent assay
(ELISA) for detection of anti-Trypanosoma evansi equine antibodies. Vet. Parasitol. 80, 149–157.
Schalm, O.W., Jain, N.C., Carrol, E.J., 1975. Veterinary Hematology, 3rd ed, 807 pp.
Seidl, A., Moraes, A.S., Silva, R.A.M.S., 1998. A financial analysis of treatment strategies for Trypanosoma
evansi in the Brazilian Pantanal. Prev. Vet. Med. 33, 219–234.
Seidl, A.F., Moraes, A.S., Silva, R.A., 2001. Trypanosoma evansi control and horse mortality in the Brazilian
Pantanal. Mem. Inst. Oswaldo Cruz. 96, 599–602.
Silva, R.A.M.S., Arosemena, N.A.E., Herrera, H.M., Sahib, C.A., Ferreira, M.S.J., 1995a. Outbreak of trypa-
nosomosis due to Trypanosoma evansi in horses of Pantanal Mato-grossense, Brazil. Vet. Parasitol. 60, 167–
171.
Silva, R.A.M.S., Herrera, H.M., Barros, A.T.M., 1995b. Trypanosomosis outbreak due to Trypanosoma evansi in
the Pantanal, Brazil. A preliminary approach on risk factors. Revue Elev. Med. Vet. Pays Trop. 48, 315–319.
Silva, R.A.M.S., Herrera, H.M., Domingos, L.B.S., Ximenes, F.A., 1995c. Pathogenesis of Trypanosoma evansi in
dogs and horses: hematological and clinical aspects, Ciencia Rural. Santa Maria 25, 233–238.
Silva, R.A.M.S., Sahib, C.A., de la Rue, M., Herrera, H.M., Ferreira, M.J., Davila, A.M.R., 1996. Coagulopathy
due to Trypanosoma evansi acute infection in a dog. Arq. Bras. Med. Vet. Zoot. 48, 485–489.
Silva, R.A.M.S., Victorio, A.M., Ramirez, L., Davila, A.M.R., Trajano, V., Jansen, A.M., 1999. Hematological and
blood chemistry alterations in coatis (Nasua nasua) naturally infected by Trypanosoma evansi in the Pantanal,
Brazil. Revue Elev. Med. Vet. Pays Trop. 52, 119–122.
Sudarto, M.W., Tabe, I.L.H., Haines, D.M., 1990. Immunohistochemical demonstration of Trypanosoma evansi in
tissues of experimentally infected rats and a naturally infected water buffalo (Bubalus bubalis). J. Parasitol. 76,
162–167.
Trail, J.C., D’Ieteren, G.D., Feron, A., Kakiese, O., Mulungo, M., Pelo, M., 1990. Effect of trypanosome infection,
control of parasitaemia and control of anaemia development on productivity of N’Dama cattle. Acta Trop. 48,
37–45.
Vokaty, S., McPherson, V.O., Camus, E., Applewhaite, L., 1993. Ovine trypanosomosis: a seroepidemiological
survey in coastal Guyana. Ver. Elev. Med. Vet. Pays Trop. 46, 57–59.
Walsh, P.S., Metzger, P.K., Higuchi, R., 1991. Chelex1 100 as a medium for simple extraction of DNA for PCR-
based typing from forensic material. Biotechniques 10, 506–513.
Wernery, U., Zachariah, R., Mumford, J.A., Luckins, T., 2001. Preliminary evaluation of diagnostic tests using
horses experimentally infected with Trypanosoma evansi. Vet. J. 161, 287–300.
Wilcox, R., 1992. Cattle and environment in the Pantanal of Mato Grosso, Brazil, 1870–1970. Agricult. Hist. 66,
232–256.
Woo, P.T.K., 1970. The haematocrit centrifuge technique for the diagnosis of African trypanosomiasis. Acta Trop.
27, 384–386.
H.M. Herrera et al. / Veterinary Parasitology 125 (2004) 263–275 275