Download The broad spectrum of Trichinella hosts: From cold

Veterinary Parasitology 132 (2005) 3–11
www.elsevier.com/locate/vetpar
The broad spectrum of Trichinella hosts:
From cold- to warm-blooded animals
E. Pozio *
Department of Infectious Parasitic and Immunomediated Diseases, Istituto Superiore di Sanità,
viale Regina Elena 299, 00161 Rome, Italy
Abstract
In recent years, studies on Trichinella have shown that the host range is wider than previously believed and new Trichinella
species and genotypes have been described. Three classes of vertebrates are known to act as hosts, mammals, birds and reptiles,
and infected vertebrates have been detected on all continents but Antarctica. Mammals represent the most important hosts and all
Trichinella species are able to develop in this vertebrate class. Natural infections with Trichinella have been described in more
than 150 mammalian species belonging to 12 orders (i.e., Marsupialia, Insectivora, Edentata, Chiroptera, Lagomorpha,
Rodentia, Cetacea, Carnivora, Perissodactyla, Artiodactyla, Tylopoda and Primates). The epidemiology of the infection greatly
varies by species relative to characteristics, such as diet, life span, distribution, behaviour, and relationships with humans. The
non-encapsulated species Trichinella pseudospiralis, detected in both mammals (14 species) and birds (13 species), shows a
cosmopolitan distribution with three distinguishable populations in the Palearctic, Nearctic and Australian regions. Two
additional non-encapsulated species, Trichinella papuae, detected in wild pigs and saltwater crocodiles of Papua New Guinea,
and Trichinella zimbabwensis, detected in farmed Nile crocodiles of Zimbabwe, can complete their life cycle in both mammals
and reptiles. To the best of our knowledge, T. papuae and T. zimbabwensis are the only two parasites known to complete their
entire life cycle independently of whether the host is warm-blooded or cold-blooded. This suggests that these two Trichinella
species are capable of activating different physiological mechanisms, according to the specific vertebrate class hosting them.
# 2005 Elsevier B.V. All rights reserved.
Keywords: Trichinella; Mammals; Birds; Reptiles; Warm-blooded animals; Cold-blooded animals
1. Introduction
Until 1974, nematode worms of the genus
Trichinella were recognised exclusively as parasites
of mammals. At that time, Trichinella larvae had been
described in more than 150 mammalian hosts (both
* Tel.: +39 06 49902304; fax: +39 06 49387065.
E-mail address: [email protected].
natural and experimental infections) belonging to 12
orders (Marsupialia, Insectivora, Edentata, Lagomorpha, Chiroptera, Rodentia, Cetacea, Carnivora, Perissodactyla, Artiodactyla, Tylopoda and Primates)
(Campbell, 1983). Furthermore, although three new
species were described in 1972, Trichinella nativa,
Trichinella nelsoni and Trichinella pseudospiralis
(Britov and Boev, 1972; Garkavi, 1972), only one
species, Trichinella spiralis, was recognised as valid
0304-4017/$ – see front matter # 2005 Elsevier B.V. All rights reserved.
doi:10.1016/j.vetpar.2005.05.024
4
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
(Dick, 1983). At the time, biochemical and molecular
methods capable of distinguishing among sister
species had not yet been developed, and the time
was not yet at hand for the introduction of a
multispecies genus.
Since that time, the use of artificial digestion to
identify animals infected with Trichinella has become
widespread, representing a great improvement over
the use of trichinelloscope, which does not allow nonencapsulated larvae to be detected and has a lower
sensitivity for encapsulated larvae relative to artificial
digestion. Moreover, biochemical and molecular
methods to distinguish among Trichinella isolates
have been developed, and there has been an increasing
interest in zoonotic diseases. All of these factors have
allowed the genus Trichinella and its host spectrum to
be re-evaluated.
2. Taxonomy and distribution of the genus
Trichinella
Parasites of the genus Trichinella can be separated
in two main groups: (1) species with encapsulated
larvae in host muscles: T. spiralis, T. nativa (including
one separate genotype, Trichinella T6), Trichinella
britovi (including two separate genotypes, Trichinella
T8 and T9), Trichinella murrelli and T. nelsoni
(Murrell et al., 2000; La Rosa et al., 2003a); and (2)
species with non-encapsulated larvae in host muscles:
T. pseudospiralis, T. papuae and T. zimbabwensis (La
Rosa et al., 2003a).
The distribution area of T. spiralis has been
strongly influenced by the passive introduction of
this pathogen by domestic pigs and synanthropic rats
on different continents, followed by the transmission
to wildlife. It is now known that this parasite is present
in many countries of North, Central and South
America, Europe and Asia, in Egypt (Africa), and
in Indonesia and New Zealand (Australian region)
(Pozio, 2001).
Freeze-resistant T. nativa larvae are widespread and
are mainly found in carnivores of the Holoarctic
region, and the isotherm 48 in January seems to be
the southern border of distribution. The genotype
Trichinella T6, which is closely related to T. nativa, is
widespread among carnivores along the Rocky
Mountains, from Alaska to Montana and Idaho (La
Rosa et al., 2003b). Trichinella britovi is present in
temperate areas of the Palearctic region, where the
northern border of distribution is the isotherm 68 in
January. The genotype Trichinella T9 is present in
Japan, and the genotype Trichinella T8 is present in
South Africa and Namibia; both of these genotypes are
strictly related to T. britovi (Pozio, 2001); furthermore,
T. britovi has been recently documented in West
Africa (Guinea Conakry). Trichinella murrelli is
widespread among wildlife in the United States
(Pozio, 2001) and T. nelsoni is widespread among
wildlife in Eastern Africa, from Kenya to South Africa
(Pozio, 2001).
Three different populations of the non-encapsulated species T. pseudospiralis have been detected in
the Paleartic, Nearctic and Australian regions (La
Rosa et al., 2001), T. papuae has been detected in
several areas of Papua New Guinea (Pozio et al.,
2004a), and T. zimbabwensis in Zimbabwe (Pozio
et al., 2002) and Mozambique, although the presence
in Mozambique has yet to be confirmed by molecular
identification techniques (see below).
3. Relationship between Trichinella species and
vertebrate classes
The encapsulated species of the genus Trichinella
can only complete their life cycle in mammals in that
they require a host body temperature ranging from 37
to 40 8C (Tables 1 and 2) (Pozio et al., 2004b). The
non-encapsulated species, T. pseudospiralis, shows a
wider host spectrum, which includes mammals and
birds because it can complete its life-cycle at host
body temperatures ranging from 37 to 42 8C (Tables 1
and 2) (Pozio et al., 2004b). The two non-encapsulated
Table 1
Relationship between Trichinella species and vertebrate classes
Trichinella species Mammals Birds Reptiles Amphibians Fishes
T.
T.
T.
T.
T.
T.
T.
T.
spiralis
nativa
britovi
pseudospiralis
murrelli
nelsoni
papuae
zimbabwensis
ND, not done.
Yes
Yes
Yes
Yes
Yes
Yes
Yes
Yes
No
No
No
Yes
No
No
No
No
No
No
No
No
No
No
Yes
Yes
No
ND
ND
ND
ND
ND
ND
ND
No
ND
No
No
ND
ND
No
No
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
5
Table 2
Relationships between vertebrate classes, body temperature and Trichinella species
Birds (40.5–42.5 8C)
Mammals (37.5–40 8C)
Reptiles* (25–32 8C)
*
T. papuae, T. zimbabwensis
T. pseudospiralis
All encapsulated species
Not susceptible
Susceptible
Susceptible
Susceptible
Susceptible
Not susceptible
Not susceptible
Susceptible
Not susceptible
Equatorial reptiles.
species, T. papuae and T. zimbabwensis, can infect
both reptiles (only those living in equatorial regions)
and mammals and require host body temperatures
ranging from 25 to 40 8C (Tables 1–3) (Pozio et al.,
2004b). There is only one report of experimental
infections of amphibians (frogs and axolotls) with T.
spiralis; however, the development of larvae in the
muscles was incomplete (Gaugusch, 1950). All
attempts to infect fish with T. spiralis, T. britovi, T.
pseudospiralis, T. papuae and T. zimbabwensis have
failed (Pozio and La Rosa, 2004c; Guevara Pozo and
Contreras-Pena, 1966; Moretti et al., 1997; Tomasovicova, 1981).
4. Reptiles
In laboratory conditions, some authors were able to
reproduce the cycle of T. spiralis in reptiles
(Phrynosoma cornutum, Agama caucasica, Lacerta
agilis, Vipera ammodytes, Emys orbicularis), yet only
when maintained at temperatures (37–40 8C) higher
than those necessary for breeding these reptiles in
captivity (25–30 8C) (Jordan, 1964; Guevara Pozo and
Contreras-Pena, 1966; Cristeau and Perian, 1999;
Asatrian et al., 2000). Experimental infections of
Caiman sclerops reared at 25–30 8C with all of the
encapsulated species and with the non-encapsulated
species, T. pseudospiralis, failed (Kapel et al., 1998)
(Table 3). In 1995, Trichinella sp. was discovered in
farmed Nile crocodiles in Zimbabwe. In 2002, this
parasite was classified as a new species, Trichinella
zimbabwensis (Pozio et al., 2002). Very recently, T.
papuae was detected in a high number of saltwater
crocodiles (Crocodylus porosus) from Papua New
Guinea (Pozio et al., 2004a).
Experimental infections of reptiles belonging to
three orders (Loricata, Squamata and Chelonide) with
the eight Trichinella species have shown that only T.
papuae and T. zimbabwensis are able to complete their
cycle in equatorial reptiles reared at 25–32 8C (i.e.,
their trophic temperature range). Furthermore, a
Table 3
Experimental and natural infections of reptiles with Trichinella species
Trichinella species
Experimental infections
Natural infections
T. spiralis
Development in Phrynosoma cornutum, Agama caucasica,
Lacerta agilis, Vipera ammodytes, Emys orbicularis only
at 37–40˚ C No development in Caiman sclerops,
Python molurus and Pelomedusa subrufa at 25–32˚ C
No development in C. sclerops, P. molurus and P. subrufa at
No development in C. sclerops, P. molurus and P. subrufa at
No development in C. sclerops, P. molurus and P. subrufa at
No development in C. sclerops, P. molurus and P. subrufa at
No development in C. sclerops, P. molurus and P. subrufa at
High infection in C. sclerops and Varanus exanthematicus at
very low infection in P. molurus and P. subrufa at 28–30˚ C
High infection in C. sclerops and Varanus exanthematicus at
very low infection in P. molurus and P. subrufa at 28–30˚ C
No
T.
T.
T.
T.
T.
T.
nativa
britovi
pseudospiralis
murrelli
nelsoni
papuae
T. zimbabwensis
a
Infections should be confirmed.
25–32˚ C
25–32˚ C
25–32˚ C
25–32˚ C
25–32˚ C
25–32˚ C;
25–32˚ C;
No
No
No
No
No
Crocodylus porosus
of Papua New Guinea
Crocodylus niloticus
of Zimbabwe; C. niloticus
of Mozambiquea;
Varanus niloticus of
Zimbabwe a
6
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
different susceptibility has been observed between
reptile orders and suborders (Table 3). In fact, the
turtle (Pelomedusa subrufa) and the python (Python
molurus) were not found to be suitable hosts where
T. papuae and T. zimbabwensis showed a very low
reproductive capacity in these hosts. In contrast, they
showed a high reproductive capacity in the caiman
(Caiman crocodilus, synonymous of Caiman sclerops)
and in the monitor lizard (Varanus exanthematicus),
which did not exhibit any clinical sign of infection in
spite of a high number of larvae per gram of muscle
tissue (Pozio et al., 2004b). Regarding sylvatic
reptiles, infection with Trichinella sp. has been
reported in monitor lizards (Varanus niloticus) from
several sites in Zimbabwe and in Nile crocodiles
(Crocodylus niloticus) from Lake Cabora Basa in
Mozambique (C. Foggin, personal communication),
although these reports have yet to be genetically
confirmed.
Reptile orders show different levels of susceptibility to Trichinella infection and this is consistent
with the diet and behaviour of the species. Crocodiles
and monitor lizards eat flesh from large animals and
show scavenger behaviour, whereas snakes and turtles
eat only small pray and are not scavengers. The
adaptation of T. papuae and T. zimbabwensis to
crocodiles and monitor lizards is also supported by the
long survival of adult worms (at least 60 days) in the
gut (Pozio et al., 2004b). The longer survival, in
comparison to that in mammalian hosts (of 10–15
days), may be related to the lower metabolism of
reptiles, which require a long period of time to digest
the large amounts of flesh ingested at each feeding.
This results in a concomitantly slow release of muscle
larvae into the gut. Without the capacity to survive for
long periods of time in the gut, these worms would
have few opportunities to mate.
Table 4
Family and species of birds in which Trichinella pseudospiralis
infection has been documented or suspected
Family
Species
Corvidae
Accipitridae
Corvus frugilegus
Aquila rapax
Buteo buteo
Accipiter cooperi
Circus aeruginosus
Accipiter gentiles
Falconidae
Falco peregrinus
Stercorariidae Stercorarius pomarinus
Strigidae
Bubo virginianus
Strix aluco
Athene noctua
Tytanidae
Tyto novaehollandiae
Cathartidae Coragypus atratus
Country
Infection
Kazakhstan
Kazakhstan
Spain, Russia
USA
Tasmania
Kazakhstan
Kazakhstan
USA
USA
Italy
Italy
Tasmania
USA
Documented
Documented
Suspected
Suspected
Documented
Suspected
Suspected
Suspected
Suspected
Documented
Documented
Documented
Documented
documented in 1980 (Shaikenov, 1980). In the last 24
years, nematode larvae similar to that of the genus
Trichinella, have been detected in 13 different bird
species in nature (Pozio et al., 1992; Lindsay et al.,
1995; Pozio et al., 1999; B.L. Garkavi, personal
communication), although T. pseudospiralis has been
identified at the species level in only seven of them
(Table 4). The low genetic variability of T. pseudospiralis isolates present in the Nearctic and Palearctic
regions (La Rosa et al., 2001; Gamble et al., 2005)
could be related to a high gene flow between isolates
favoured by birds of each geographical region. The
low number of reports of T. pseudospiralis in birds
could be due to the low number of examined birds in
comparison to that of mammals and to the use of
trichinelloscope instead of artificial digestion. The
role played by birds in the epidemiology of this
parasite in comparison to that of mammals is still
unknown.
6. Mammals
5. Birds
6.1. Marsupialia
Since 1974, experimental infections of birds (e.g.,
hens, ducks, magpies, pigeons, crows, sparrows,
starlings, partridges, owls, kites and herons) have
shown that these vertebrates are suitable hosts for the
non-encapsulated species, T. pseudospiralis (Bessonov et al., 1976). The first natural infection of birds
(crows, Corvus frugilegus) with T. pseudospiralis was
Encapsulated larvae have been detected in Didelphis marsupialis and Didelphis virginiana from the
United States (Zimmermann and Hubbard, 1969;
Solomon and Warner, 1969; Leiby et al., 1988), where
these animals play the role of reservoir. Three species
of marsupials (Dasyurus viverrinus, Sarcophilus
harrisii and Dasyurus maculatus) were found to be
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
infected with T. pseudospiralis in Tasmania where
they act as a reservoir (Obendorf et al., 1990).
6.2. Insectivora
Nematode larvae with a Trichinella shape have been
detected in more than 10 insectivore species (Rausch,
1970; Merkushev, 1970; Zimmermann, 1971; Kim,
1983); however, no isolates from this order have ever
been identified at the species or genus level. It should be
considered that these animals have a very short life span
(several months), are small in size, and ingest very small
quantities of food at each feeding, so that they may not
have the chance to ingest at least two larvae of both
sexes. Consequently, the role of these mammals in the
epidemiology of Trichinella in still unknown (see also
the section on rodents).
6.3. Edentata
The armadillo (Chaetophractus villosus) has been
found to be infected with T. spiralis in Chile and
Argentina (Neghme and Schenone, 1970; Pozio,
2000). Because of its scavenger behaviour, this animal
could act as a reservoir of T. spiralis.
6.4. Lagomorpha
Nematode larvae with a Trichinella shape have
been detected in two sylvatic species of hare and in
one rabbit in North America and Europe (Beck, 1970;
Rausch, 1970; Zimmermann, 1971). However, no
larva has ever been identified at the species level. On
the basis of the diet of this animal group, it is likely
that infection with Trichinella may only be circumstantial, with no important role in the epidemiology of
these pathogens.
6.5. Rodentia
Dozens of rodent species have been found to be
naturally-infected with nematode larvae considered to
belong to the genus Trichinella (Beck, 1970; Rausch,
1970; Merkushev, 1970; Bessonov, 1981; Kim, 1983;
Holliman and Meade, 1980; Nelson, 1970; Dick and
Pozio, 2001). However, to date, only larvae from three
species of rodents have been identified at the species
level (Shaikenov and Boev, 1983; Dick and Pozio,
7
2001). Several of the above quoted authors argue that
most reports were based upon incorrect identifications. Furthermore, published reports do not always
unequivocally explain whether the examined rodents
were trapped near human settlements or in the wild
(i.e., whether these animals were involved in a
synanthropic-domestic cycle or in a sylvatic cyle);
consequently, the role played by these animals in the
sylvatic cycle is still obscure.
To evaluate the role of rodents as well as that of
other small mammals, such as insectivores, in the
epidemiology of Trichinella, it should be considered
that they have a short life span and ingest a low
quantity of flesh at each feeding. This strongly reduces
their risk of becoming infected. Furthermore, the
length of survival of larvae in decaying muscles of
infected micromammals is lower than that in the
muscles of larger animals due to the small size of the
host where the length of decomposition is more
influenced by environmental factors (temperature and
moisture). It should also be taken into account that
thousands of rodents would need to be collected from
the same locality in order to obtain information on
their role in the sylvatic cycle.
Only larvae from Rattus norvegicus, Rattus rattus
and Bandicota bengalensis have been identified at the
species level. Trichinella spiralis, T. britovi and T.
pseudospiralis have been detected in R. norvegicus
(Dick and Pozio, 2001); T. britovi has been detected in
R. rattus (Dick and Pozio, 2001); and T. pseudospiralis
has been detected in B. bengalensis (Shaikenov and
Boev, 1983). All animals originated from farms or
urbanized areas.
The role of R. norvegicus as a reservoir of T.
spiralis in the domestic habitat has been a topic of
debate (Schad et al., 1987). Recently, a survey carried
out in Croatia suggested that the brown rat acts as
vector or victim rather than as a reservoir (Stojcevic
et al., 2004). In fact, the presence of infected rats has
been strongly influenced by the human practice of
spreading pork scraps in the environment, where
infected rats were detected there were infected pigs
but not visa versa.
6.6. Cetacea
There is only one circumstantial report of nematode
larvae, considered to be Trichinella larvae, in muscles
8
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
of a beluga whale (Delphinapterus leucas) (1/482,
0.2%) (Rausch, 1970). No larva from cetaceans has
ever been identified at the species level. The role of
cetaceans in the epidemiology of Trichinella is still
unknown, but on the basis of the diet of these animals,
we can believe that they do not play any important role
in the epidemiology of these parasites.
four continents, and many isolates have been
identified at the species level (Table 5). To date,
carnivores have been found to host nearly all isolates
(>90%) of T. nativa, T. murrelli and T. nelsoni, 68%
of T. britovi isolates, 30% of T. pseudospiralis
isolates, and 19% of T. spiralis isolates identified at
the International Trichinella Reference Centre
(ITRC) (Table 5).
6.7. Carnivora
6.8. Perissodactyla
This is the order of mammals that plays the most
important role in the epidemiology of Trichinella. All
species of Trichinella are able to complete their life
cycle in these animals (Kapel, 2000; Webster et al.,
2002). Most species, mainly those with scavenger
behaviour, act as a reservoir for encapsulated species.
Their role as a reservoir for non-encapsulated species,
however, is still unknown where the use of the less
sensitive trichinelloscope, instead of artificial digestion to detect these parasites in muscles, may have
prevented their detection in the past. As a general rule,
carnivores are at the top of the food chain and thus
represent the best host for Trichinella. Documented
infections with two or more species of Trichinella
show that these animals can get infected more than one
time during their life span (Pozio, 2000) which is
substantially longer than that of other mammalian
orders (e.g., insectivora, rodents). Carnivores also
ingest a large amount of flesh at each feeding and carry
out their trophic activity on a territory larger than that
of other mammals thereby increasing their risk of
acquiring an infection.
Trichinella larvae have been detected in more
than 80 species of carnivores (Campbell, 1983) from
Table 5
Number of Trichinella isolates from carnivores and swine identified
at the species level on the total of Trichinella isolates identified at the
International Trichinella Reference Center (Rome, Italy)
Trichinella species
No. of isolates from
carnivores/total (%)
No. of isolates from
swine/total (%)
T.
T.
T.
T.
T.
T.
T.
T.
102/521 (19%)
211/215 (98%)
366/538 (68%)
7/23 (30%)
27/28 (96%)
14/15 (93%)
0/5
0/1
380/521 (73%)
3/215 (1.4%)
122/538 (23%)
3/23 (13%)
0/33
1/15 (7%)
3/5
0/1
spiralis
nativa
britovi
pseudospiralis
murrelli
nelsoni
papuae
zimbabwensis
Only domestic horses have been found to be
naturally-infected with T. spiralis (most infections),
T. britovi (three cases) and T. murrelli (one case)
(Pozio, 2001). Experimental and natural infections
suggest that the horse is not a good host for
Trichinella (Soule et al., 1989; Pozio, 2001). In fact,
larvae survive for a short period of time in the
muscles of this animal. All natural infections seem to
be related to improper breeding practices; however,
horse meat is one of the most dangerous sources of
infection for humans (Boireau et al., 2000; Pozio,
2001). This is an example of an animal that does not
act as a reservoir of Trichinella, but can have a
tremendous impact on humans given its role as a
vector.
6.9. Artiodactyla
Of the eight families belonging to this order, only
species of the family Suidae play an important role in
the epidemiology of Trichinella (Kapel, 2000; Pozio,
2000, 2001). Sus scrofa, both the domestic (pig) and
the sylvatic strains (wild boar, wild pigs, etc.), is the
most important host of T. spiralis (73% of identified
isolates), whereas T. britovi, T. pseudospiralis and T.
nativa isolates have only been characterized in 23, 13
and 1.4% of infections, respectively (Table 5). Among
the other families of this order, there are only two
circumstantial reports of Trichinella larvae in one
reindeer (Rangifer tarandus) from Russia (Bessonov,
1981) and in two roe deer (Capreolus capreolus) from
Croatia (Pozio, 2001), although these larvae were not
identified at the species level. A suspected infection in
a cow has been reported in China (Murrell, 1994).
Experimental infections in cows, goats, and sheep
suggest that these animals do not play any role in
Trichinella epidemiology, given the short survival of
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
Trichinella larvae in their muscles (Smith et al., 1990;
Reina et al., 1996; Theodoropoulos et al., 2000). In
China, mutton has been considered as the source of
infection for humans, yet no larva of Trichinella has
ever been isolated from a naturally infected sheep
(Takahashi et al., 2000). A circumstantial infection in
a hippopotamus (Hippopotamus amphibius) from a
zoo has been reported in Africa (Nelson, 1970).
6.10. Tylopoda
There is one circumstantial report of air-dried
camel meat as the putative source of a human outbreak
in Germany; however, neither the origin of the meat
nor the nematode larvae were ever identified (Bommer
et al., 1980).
6.11. Chiroptera
No natural infection has been documented; however, experimental-infections with T. spiralis has
shown that bats (Myotis lucifugus, Myotis keenii,
Pipistrellus subflavus) are susceptible to this parasite
(Chute and Covalt, 1960). The diet of this mammal
group (mainly frugivorous and insectivorous) strongly
suggests that these animals do not play any role in the
epidemiology of Trichinella.
6.12. Primates
Homo sapiens is the only species of Primates that
has been found to be naturally-infected with all
species of Trichinella, except T. zimbabwensis (Pozio,
2001). Fifteen species of experimentally-infected nonhuman primates have been shown to be highly
susceptible to several encapsulated and non-encapsulated species of Trichinella, including T. zimbabwensis
(McCoy, 1932; Welt, 1941; Nelson and Mukundi,
1962; Kociecka et al., 1981; S. Mukaratirwa, personal
communication). Trichinella infection has never been
detected in non-human primates from the wild.
7. Discussion
The number of animals proven to be hosts of
Trichinella is lower than the number of animals from
which nematode larvae with a Trichinella shape have
9
been collected but not yet identified. Thus, caution
should be taken when evaluating the role of specific
animals as Trichinella hosts. Since 1982, when
Trichinella larvae started to be identified unequivocally, first with biochemical analysis (Flockhart
et al., 1982) and then with molecular analysis
(Chambers et al., 1986), the number of confirmed
mammalian hosts has climbed to 1/3 of that
identified only on the basis of recovery of larvae
from muscles.
The specific role played by a given host species in
the epidemiology of Trichinella should also be
determined. To define a species as reservoir of
Trichinella in a given area, at least two criteria must
be met: (1) the area where the host species is
detected should be similar in size or larger than the
area where Trichinella is present; and (2) the host
species should maintain the infection for years,
independently of the presence of Trichinella in other
animals living in the same area. In all other cases, the
host species should be considered as a ‘‘vector’’ of
the parasite to another host (e.g., the horse for
humans, the rat for pigs) or as a victim (e.g., infected
persons, rabbit, hare, whale).
Finally, it must be considered that data in the
literature do not allow the role of each animal species
in the epidemiology of Trichinella to be determined.
There is always a bias due to the species target of an
epidemiological investigation. Most surveys are
carried out on animals hunted for sport, which only
represent a small percentage of the putative hosts for
these parasites. Furthermore, the collection of animals
at the top of the food chain (i.e., mainly carnivores)
strongly facilitates the detection of these parasites in
the investigated area. By contrast, surveys on sylvatic
rodents and insectivores require that thousands of
animals be examined, and these surveys are often
frustrating because they provide negative results.
However, in recent years, many species that can play
an important role in the epidemiology of Trichinella
are protected by national and international legislation
thereby limiting the opportunity to collect samples.
The broad host spectrum of parasites of the genus
Trichinella should be considered when implementing
measures for avoiding the transmission from animals
to humans. Furthermore, before implementing control
measures, the assessment of risk and the role of each
host species should be carefully evaluated.
10
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
Acknowledgement
Work funded by the EU project ‘‘TRICHIPORSE’’
(contract QLK1-CT-2001-01156).
References
Asatrian, A., Movsessian, S., Gevorkian, A., 2000. Experimental
infection of some reptiles with T. spiralis and T. pseudospiralis.
In: Abstract book on the 10th International Conference on
Trichinellosis, 20–24 August, Fontainebleau, France, p. 154.
Beck, J.W., 1970. Trichinosis in domesticated and experimental
animals. In: Gould, S.E. (Ed.), Trichinosis in man and animals.
C.C. Thomas Publisher, Springfield, pp. 61–80.
Bessonov, A.S., 1981. Changes in the epizootic and epidemic
situation of trichinellosis in the USSR. In: Kim, C.W., Ruitenberg, E.J., Teppema, J.S. (Eds.), Trichinellosis. Reedbooks,
Chertsey, England, pp. 365–368.
Bessonov, A.S., Penkova, R.A., Gumenshchikova, V.P., 1976. Trichinella pseudospiralis Garkavi, 1972: Morphological and biological characteristics and host specificity. In: Kim, C.W.,
Pawlowski, Z.S. (Eds.), Trichinellosis. University Press of
New England, pp. 79–93.
Boireau, P., Vallée, I., Roman, T., Perret, C., Mingyuan, L., Gamble,
H.R., Gajadhar, A., 2000. Trichinella in horses: a low frequency
infection with high human risk. Vet. Parasitol. 93, 309–
320.
Bommer, W., Kaiser, H., Mergerian, H., Pottkamper, G., 1980.
Outbreak of trichinellosis in a youth centre in Neidersachsen
by air-dried imported camel meat. In: Proceedings of the 1st
World Congress on Food-borne Infections and Intoxications,
West Berlin, Germany, pp. 441–444.
Britov, V.A., Boev, S.N., 1972. Taxonomic rank of various strains of
Trichinella and their circulation in nature. Vestnik Akademii
Nauk KSSR 28, 27–32. [in Russian].
Campbell, W.C., 1983. Epidemiologi I. modes of transmission. In:
Campbell, W.C. (Ed.), Trichinella and Trichinosis. Plenum
Press, New York and London, pp. 425–444.
Chambers, A.E., Almond, N.M., Knight, M., Simpson, A.J., Parkhouse, R.M., 1986. Repetitive DNA as a tool for the identification and comparison of nematode variants: application to
Trichinella isolates. Mol. Biochem. Parasitol. 21, 113–120.
Chute, R.M., Covalt, D.B., 1960. The effect of body temperature on
development of Trichinella spiralis in bats. J. Parasitol. 46, 855–
858.
Cristeau, G., Perian, A., 1999. Experimental infestation and Trichinella larvae in vipers. Rev. Romana Parasitol. 9, 45.
Dick, T.A., 1983. Species and intraspecific variation. In: Campbell,
W.C. (Ed.), Trichinella and Trichinosis. Plenum Press, New
York, pp. 31–74.
Dick, T.A., Pozio, E., 2001. Trichinella spp. and trichinellosis. In:
Samuel, W.M., Pybus, M.J., Kocan, A.A. (Eds.), Parasitic Diseases of Wild Mammals. second ed. Iowa State University Press,
Ames, pp. 380–396.
Flockhart, H.A., Harrison, S.E., Dobinson, A.R., James, E.R., 1982.
Enzyme polymorphism in Trichinella. Trans. R. Soc. Trop. Med.
Hyg. 76, 541–545.
Gamble, H.R., Pozio, E., Lichtenfels, J.R., Zarlenga, D.S., 2005.
Trichinella pseudospiralis from a wild pig in Texas, USA. Vet.
Parasitol. 132, 147–150.
Garkavi, B.L., 1972. Species of Trichinella isolates from wild
animals. Veterinariya 10, 90–91 [in Russian].
Gaugusch, Z., 1950. Studies on the infection of poikilothermic
animals with T. spiralis. Vet. Med. 5, 95–106.
Guevara Pozo, D., Contreras-Pena, A.J., 1966. Trichinella spiralis
Owen, 1835; Experiencias de infestacion en diversos vertebrados. Revista Iberica de Parasitologia 26, 239–288.
Holliman, R.B., Meade, B.J., 1980. Native trichinosis in wild rodents
in Henrico County Virginia. J. Wildlife Dis. 16, 205–207.
Jordan, C.A., 1964. Experimentally trichinized Texas horned
lizards. J. Parasitol. 50, 24.
Kapel, C.M.O., 2000. Host diversity and biological characteristics of
the Trichinella genotypes and their effect on transmission. Vet.
Parasitol. 93, 263–278.
Kapel, C.M., Webster, P., Bjorn, H., Murrell, K.D., Nansen, P., 1998.
Evaluation of the infectivity of Trichinella spp. for reptiles
(Caiman sclerops) Int. J. Parasitol. 28, 1935–1937.
Kim, C.W., 1983. Epidemiology II. Geographic distribution and
prevalence. In: Campbell, W.C. (Ed.), Trichinella and Trichinosis. Plenum Press, New York and London, pp. 445–500.
Kociecka, W., van Knapen, F., Ruitenberg, E.J., 1981. Trichinella
pseudospiralis and T. spiralis infections in monkeys. In: Kim,
C.W., Ruitenberg, E.J., Teppema, J.S. (Eds.), Parasitological
Aspects Trichinellosis. Reedbooks Ltd., Chertsey, England, pp.
199–203.
La Rosa, G., Marucci, G., Pozio, E., 2003a. Biochemical analysis of
encapsulated and non-encapsulated species of Trichinella
(Nematoda, Trichinellidae) from cold- and warm-blooded animals reveals a high genetic divergence in the genus. Parasitol.
Res. 91, 462–466.
La Rosa, G., Marucci, G., Zarlenga, D.S., Casulli, A., Zarnke, R.L.,
Pozio, E., 2003b. Molecular identification of natural hybrids
between Trichinella nativa and Trichinella T6 provides evidence
of gene flow and ongoing genetic divergence. Int. J. Parasitol. 33,
209–216.
La Rosa, G., Marucci, G., Zarlenga, D.S., Pozio, E., 2001. Trichinella pseudospiralis populations of the Palearctic region and
their relationship with populations of the Nearctic and Australian regions. Int. J. Parasitol. 31, 297–305.
Leiby, D.A., Schad, G.A., Duffy, C.H., Murrell, K.D., 1988. Trichinella spiralis in an agricultural ecosystem. III. Epidemiological investigations of Trichinella spiralis in resident wild and
feral animals. J. Wild. Dis. 24, 606–960.
Lindsay, D.S., Zarlenga, D.S., Gamble, H.R., al-Yaman, F., Smith,
P.C., Blagburn, B.L., 1995. Isolation and characterization of
Trichinella pseudospiralis Garkavi, 1972 from a black vulture
(Coragyps atratus). J. Parasitol. 81, 920–923.
McCoy, O.R., 1932. Experimental trichiniasis infection in monkey.
Proc. Soc. Exp. Biol. Med. 30, 85–86.
Murrell, K.D., 1994. Beef as a source of trichinellosis. Parasitol.
Today 10, 434.
E. Pozio / Veterinary Parasitology 132 (2005) 3–11
Murrell, K.D., Lichtenfels, R.J., Zarlenga, D.S., Pozio, E., 2000. The
systematics of Trichinella with a key to species. Vet. Parasitol.
93, 293–307.
Merkushev, A.V., 1970. Trichinosis in the Union of Soviet Socialist
Republics. In: Gould, S.E. (Ed.), Trichinosis in man and animals.
C.C. Thomas Publisher, Springfield, pp. 449–456.
Moretti, A., Piergili Fioretti, D., Pasquali, P., Mechelli, L., Rossodivita, M.E., Polidori, G.A., 1997. Experimental infection of fish
with Trichinella britovi: biological evaluations. In: OrtegaPierres, G., Gamble, H.R., van Knapen, F., Wakelin, D.
(Eds.), Trichinellosis. Centro de Investigacion y Estudios Avanzados IPN, Mexico City, pp. 135–142.
Neghme, A., Schenone, H., 1970. Trichinosis in Latin America. In:
Gould, S.E. (Ed.), Trichinosis in Man and Animals. C.C. Thomas Publisher, Springfield, pp. 407–422.
Nelson, G.S., 1970. Trichinosis in Africa. In: Gould, S.E. (Ed.),
Trichinosis in Man and Animals. C.C. Thomas Publisher,
Springfield, pp. 473–492.
Nelson, G.S., Mukundi, J., 1962. The distribution of Trichinella
spiralis larvae in the muscles of primates. Wiad. Parazytol. 8,
629–632.
Obendorf, D.L., Handlinger, J.H., Mason, R.W., Clarke, K.P., Forman, A.J., Hooper, P.T., Smith, S.J., Holdsworth, M., 1990.
Trichinella pseudospiralis infection in Tasmanian wildlife. Aust.
Vet. J. 67, 108–110.
Pozio, E., 2000. The domestic, synanthropic and sylvatic cycles of
Trichinella and the flow among them. Vet. Parasitol. 93, 241–
262.
Pozio, E., 2001. Vet. Parasitol. New patterns of Trichinella infections 98, 133–148.
Pozio, E., Foggin, C.M., Marucci, G., La Rosa, G., Sacchi, L.,
Corona, S., Rossi, P., Mukaratirwa, S., 2002. Trichinella zimbabwensis sp. (Nematoda), a new non-encapsulated species
from crocodiles (Crocodylus niloticus) in Zimbabwe also infecting mammals. Int. J. Parasitol. 32, 1787–1799.
Pozio, E., Goffredo, M., Fico, R., La Rosa, G., 1999. Trichinella
pseudospiralis in sedentary night-birds of prey from Central
Italy. J. Parasitol. 85, 759–761.
Pozio, E., La Rosa, G., Evaluation of the infectivity of Trichinella
papuae and Trichinella zimbabwensis for equatorial freshwater
fishes, Vet. Parasitol., this issue.
Pozio, E., Marucci, G., Casulli, A., Sacchi, L., Mukaratirwa, S.,
Foggin, C.M., La Rosa, G., 2004b. Trichinella papuae and Trichinella zimbabwensis induce infection in experimentally infected
varans, caimans, pythons and turtles. Parasitology 128, 333–342.
Pozio, E., Owen, I.L., Marucci, G., La Rosa, G., 2004a. Trichinella
papuae in saltwater crocodiles (Crocodylus porosus) of Papua
New Guinea: A potential source of human infection. Emerg.
Infect. Dis. 10, 1507–1509.
Pozio, E., Shaikenov, B., La Rosa, G., Obendorf, D.L., 1992.
Allozymic and biological characters of Trichinella pseudospir-
11
alis isolates from free-ranging animals. J. Parasitol. 78, 1087–
1090.
Rausch, R.L., 1970. Trichinosis in the Arctic. In: Gould, S.E. (Ed.),
Trichinosis in Man and Animals. C.C. Thomas Publisher,
Springfield, pp. 348–373.
Reina, D., Munoz, O.M., Serrano, F., Molina, J.M., Navarrete, I.,
1996. Experimental trichinellosis in goats. Vet. Parasitol. 62,
125–132.
Schad, G.A., Duffy, C.H., Leiby, D.A., Murrell, K.D., Zirkle, E.W.,
1987. Trichinella spiralis in an agricultural ecosystem: transmission under natural and experimentally modified on-farm
conditions. J. Parasitol. 73, 95–102.
Shaikenov, B., 1980. Spontaneous infection of birds with Trichinella
pseudospiralis Garkavi, 1972. Folia Parasitologica 27, 227–230.
Shaikenov, B., Boev, S.N., 1980. Distribution of Trichinella species
in the old world. Wiad. Parazytol. 24, 595–608.
Smith, H.J., Snowdon, K.E., Finley, G.G., Laflamme, L.F., 1990.
Pathogenesis and serodiagnosis of experimental Trichinella
spiralis spiralis and Trichinella spiralis nativa infections in
cattle. Can. J. Vet. Res. 54, 355–359.
Solomon, G.B., Warner, G.S., 1969. Trichinella spiralis in mammals
at Mountain Lake. Virginia J. Parasitol. 55, 730–732.
Soule, C., Dupouy-Camet, J., Georges, P., Ancelle, T., Gillet, J.P.,
Vaissaire, J., Delvigne, A., Plateau, E., 1989. Experimental
trichinellosis in horses: biological and parasitological evaluation. Vet. Parasitol. 31, 19–36.
Stojcevic, D., Zivicnjak, T., Marinculic, A., Marucci, G., Andelko,
G., Brstilo, M., Pavo, L., Pozio, E., 2004. The epidemiological
investigation of Trichinella infection in brown rats (Rattus
norvegicus) and domestic pigs in Croatia suggests that rats
are not a reservoir at the farm level. J. Parasitol. 90, 666–670.
Takahashi, Y., Mingyuan, L., Waikagul, J., 2000. Epidemiology of
trichinellosis in Asia and the Pacific Rim. Vet. Parasitol. 93,
227–239.
Theodoropoulos, G., Kapel, C.M., Webster, P., Saravanos, L., Zaki,
J., Koutsotolis, K., 2000. Infectivity, predilection sites, and
freeze tolerance of Trichinella spp. in experimentally infected
sheep. Parasitol. Res. 86, 401–405.
Tomasovicova, O., 1981. The role of fresh water fish in transfer and
maintenance of Trichinellae under natural conditions. Biologia
36, 115–125.
Webster, P., Malakauskas, A., Kapel, C.M., 2002. Infectivity of
Trichinella papuae for experimentally infected red foxes (Vulpes
vulpes). Vet. Parasitol. 105, 215–218.
Welt, L.G., 1941. Urinary excretion of trichina antigen in experimental trichinosis. Proc. Soc. Exp. Biol. Med. 48, 587–589.
Zimmermann, W.J., 1971. Trichinosis. In: Davis, J.W., Anderson,
R.C. (Eds.), Parasitic diseases of wild animals. Iowa State
University Press, Ames, pp. 127–139.
Zimmermann, W.J., Hubbard, E.D., 1969. Trichiniasis in wildlife of
Iowa. Am. J. Epidemiol. 90, 84–92.