Hostname: page-component-7b7567c8c-fvjsw Total loading time: 0 Render date: 2026-11-12T06:55:17.825Z Has data issue: false hasContentIssue false
  • English
  • Français

Larval migration of the ascarid nematode Toxocara canis following infection and re-infection in the gerbil Meriones unguiculatus

Published online by Cambridge University Press:  04 September 2015

M.C. Flecher,
C. Musso,
I.V.F. Martins and
F.E.L. Pereira
M.C. Flecher*
Affiliation:
Veterinary Medicine Course and Medicine Course, Vila Velha University, Vila Velha, ES, Brazil
C. Musso
Affiliation:
Medicine Course, Vila Velha University, Vila Velha, ES, BrazilInfectious Diseases Unit, Federal University of Espirito Santo, Vitoria, ES, Brazil
I.V.F. Martins
Affiliation:
Veterinary Medicine Course, Federal University of Espirito Santo, Alegre, ES, Brazil
F.E.L. Pereira
Affiliation:
Medicine Course, Vila Velha University, Vila Velha, ES, BrazilInfectious Diseases Unit, Federal University of Espirito Santo, Vitoria, ES, Brazil
*
*E-mail: mayraflecher@yahoo.com.br
Rights & Permissions [Opens in a new window]

Abstract

A morphological and immunohistochemical study of larval migration patterns was performed in gerbils that were infected once (primary infected group) or twice (secondary infected group) with 1500 eggs of Toxocara canis. Animals from the primary infected and the re-infected group were killed at different times after infection, and larvae were counted in the intestines, liver, lungs and brain. Fragments of all organs were formalin fixed and paraffin embedded for histology and immunohistochemistry analyses (using polyclonal anti-Toxocara serum raised in rabbits infected with T. canis). In the primary infected group, larvae were more abundant in the intestine at 24 h, in the liver and lungs between 24 and 72 h and in the brain after 96 h; larvae predominated in the brain for up to 60 days after infection. In the re-infected group, an increase in the number of larvae in the liver and a reduction in the number of larvae in the brain was observed up to 60 days after re-infection. Inflammatory reactions were absent or limited. Eosinophils and loose granulomata were observed around the larvae and their antigens in the primary infected group and were more severe. Many eosinophils and typical epithelioid granulomata were observed around larvae in the re-infected group. These results demonstrate that the migration pattern of T. canis larvae in gerbils is similar to that in mice and rats, exhibiting a late neurotropic stage. In the re-infected group, there was histological evidence of an adaptive T-helper 2 (Th-2) response, and larvae were apparently retained within granulomata in the liver, without obvious signs of destruction.

Information

Type
Research Papers
Information
Journal of Helminthology , Volume 90 , Issue 5 , September 2016, pp. 569 - 576
Copyright
Copyright © Cambridge University Press 2015 

Introduction

Toxocara canis is the most common intestinal nematode in dogs. Mammals (including humans) and birds can be infected with eggs of the parasite, which deliver larvae that migrate through tissues without completely maturing. In humans, Toxocara larvae induce systemic manifestations known as larva migrans visceral syndrome. Larvae can be located in the eyes or the central nervous system, producing ocular larva migrans syndrome or eosinophilic meningoencephalitis syndrome, respectively (Kayes, Reference Kayes1997; Pawlowski, Reference Pawlowski2001; Despommier, Reference Despommier2003; Moreira-Silva et al., Reference Moreira-Silva, Rodrigues, Pimenta, Gomes, Freire and Pereira2004).

Studies of the migration of Toxocara canis larvae in paratenic hosts have been performed in mice (Burren, Reference Burren1968; Abo-Shehada & Herbert, Reference Abo-Shehada and Herbert1984; Abo-Shehada et al., Reference Abo-Shehada, Al-Zubaidy and Herbert1984; Parsons & Grieve, Reference Parsons and Grieve1990; Bardón et al., Reference Bardón, Cuéllar and Guillén1994; Epe et al., Reference Epe, Sabel, Schnieder and Stoye1994; Farjat et al., Reference Farjat, Minvielle, Pezzani and Niedfeld1995; Guardis et al., Reference Guardis, Radman, Burgos, Fonrouge and Archelli2002; Pecinali et al., Reference Pecinali, Gomes, Amendoeira, Bastos, Martins, Pegado, Bastos, Bozza and Castro-Faria-Neto2005; Camparoto et al., Reference Camparoto, Fulan, Colli, Paludo, Falavigna-Guilherme and Fernandez2008; Ollero et al., Reference Ollero, Fenoy, Cuéllar, Guillén and Del Aguila2008), rats (Lescano et al., Reference Lescano, Queiroz and Chieffi2004), non-human primates (Aljeboori et al., Reference Aljeboori, Stout and Ivey1970; Glickman & Summers, Reference Glickman and Summers1983), chickens (Galvin, Reference Galvin1964; Maruyama et al., Reference Maruyama, Nino, Yamamoto and Katsube1994; Taira et al., Reference Taira, Permin and Kapel2003a), pigs (Helwigh et al., Reference Helwigh, Lind and Nansen1999; Taira et al., Reference Taira, Saeed, Lind, Murrell and Kapel2003b; Sommerfelt et al., Reference Sommerfelt, Rosa, Duchene, Degregorio, López, Pisanú and De Torres2004) and sheep (Aldawek et al., Reference Aldawek, Levkut, Revajová, Kolodzieyski, Seveiková and Dubinský2002).

In the experimental models cited above, the authors described larvae migrating from the intestine to the liver. When eggs are introduced into the stomach, larval migration occurs mainly via a portal route within 12 h of infection by gavages. Over the next 72 h, larvae are detected in the liver and lungs. One week later, larvae are found in muscles and the central nervous system. Thereafter, larvae accumulate in nervous tissue, and their presence in other organs decreases. For this reason, some authors describe the first week of migration as visceral; thereafter, myotropic–neurotropic migration begins (Burren, Reference Burren1968; Abo-Shehada & Herbert, Reference Abo-Shehada and Herbert1984; Lescano et al., Reference Lescano, Queiroz and Chieffi2004; Cho et al., Reference Cho, Egami, Ohnuki, Saito, Chinone, Shichinohe, Suganuma and Akao2007; Alba-Hurtado et al., Reference Alba-Hurtado, Muñoz-Guzmán, Valdivia-Anda, Tórtora and Ortega-Pierres2009).

Gerbils have recently been used as experimental animals for toxocariasis, especially in studies of ocular lesions and lesions in the central nervous system (Takayanagi et al., Reference Takayanagi, Akao, Suzuki, Tomoda, Tsukidate and Fujita1999; Alba-Hurtado et al., Reference Alba-Hurtado, Tórtora, Tsutsumi and Ortega-Pierres2000; Akao et al., Reference Akao, Tomoda, Hayashi, Suzuki, Shimizu-Suganuma, Shichinohe and Fujita2003; Zanandréa et al., Reference Zanandréa, Oliveira, Abreu and Pereira2008). Little information is available concerning the migration of Toxocara larvae in gerbils (Cho et al., Reference Cho, Egami, Ohnuki, Saito, Chinone, Shichinohe, Suganuma and Akao2007; Alba-Hurtado et al., Reference Alba-Hurtado, Muñoz-Guzmán, Valdivia-Anda, Tórtora and Ortega-Pierres2009), especially during the first hours after infection. There are no reports concerning larval migration after re-infection with the parasite in gerbils. For these reasons, we investigated the migration of Toxocara larvae in gerbils after primary infection and re-infection. We used immunohistochemistry to identify larval antigens in tissue lesions.

Materials and methods

Recovery of Toxocara eggs

Toxocara canis eggs were obtained from adult females eliminated by puppies treated with piperazine (125–180 g/kg body weight). The eggs were maintained in a glass vial containing 0.1 n sulphuric acid. The glass vial was covered with gauze, protected from light and left at room temperature for 6 weeks. After the development of the third larval stage, which usually occurs 6 weeks after collection, the eggs were washed. The concentration of the suspension was adjusted to prepare inocula of 1500 eggs per animal. Inoculations were performed by gavage with appropriate metallic needles.

Experimental infections and larval recovery

Up to 35 gerbils received primary infections and three individuals per group were euthanized with pentobarbital anaesthesia or in a carbon dioxide chamber after 4, 8, 12, 24, 48, 72 and 96 h, and also 7, 15, 30 and 60 days post-infection (primary infected group). Another group of 24 gerbils was re-infected 4 weeks after primary infections and three individuals per group were euthanized 4, 8, 48, 72 h and 7, 15, 30, 60 days after re-infection (re-infected group).

Larval counting in the intestine was performed by taking segments of the organ, which were opened, washed and flattened between two 7.5 ×  5 cm glass slides. The slides were sealed with tape and observed under an optical microscope. The liver was weighed, and fragments from different lobes were weighed and flattened between two 7.5 ×  5 cm glass slides for larval counting (squashed fragments represented 30–40% of the total organ weight). The total number of larvae in the liver was obtained by correcting the number of larvae counted in the fragments according to the total weight of the organ. The remaining fragments were immediately fixed in neutral formalin. The encephalon was removed and separated into two hemispheres. One hemisphere was immediately fixed in buffered formalin. The other hemisphere was flattened between 7.5 ×  5 cm slides for larval counting, which was performed as described for the liver. Because lung fragments were difficult to flatten, the left lung was digested in an acid pepsin solution with a pH of 1.1–1.2 (2.6 g pepsin 1:10,000 with biological activity, 5.0 g NaCl and 7.0 ml HCl in 493 ml of distilled water). After centrifugation (at 1000 rpm for 5 min), the larvae in the sediment were counted. The number of larvae in each organ was expressed as a percentage of the total number of larvae counted in each animal.

Histopathology and immunohistochemistry

Fragments of the liver, intestine, lung, heart, skeletal muscles, encephalon and eyes were fixed in 4% buffered formalin. After eyeball fixation, the lenses were removed. Each globe was sectioned in the frontal plane, and the two halves were examined under a stereoscopic microscope to search for larvae and lesions. All tissues were paraffin embedded. Sections were stained with haematoxylin and eosin and used for immunohistochemistry staining.

Immunohistochemistry for the detection of parasite antigens was performed in sections mounted on silanized slides using a polyclonal anti-Toxocara serum. The latter was obtained from rabbits after four inoculations with 5000 T. canis eggs at 2-week intervals (Smith et al., Reference Smith, Quinn, Bruce and Girdwood1982). One week after the last inoculation, blood was collected by cardiac puncture. The serum immunoglobulin was precipitated with ammonium sulphate. The precipitate was dialysed for 12 h, with three changes of phosphate-buffereds saline (PBS). Immunohistochemistry was performed using avidin–biotin–peroxidase (DAKO, Carpinteria, California, USA) according to the manufacturer's instructions. The primary antibody was used at a dilution of 1:300. No previous adsorption with antigens from other nematodes was performed using the rabbit serum. For this reason, cross-reactivity with other nematode larvae could not be ruled out.

Results

Primary infections

No gross changes were observed in the intestine. In liver and lungs, small, whitish nodules were observed in the organ surface during the second week after infection. Haemorrhagic areas were frequently seen in the lungs up to the seventh day after infection. There were no apparent macroscopic changes in the encephalon.

The number of larvae was greater in the intestine for the first 24 h, then larvae began to be frequent in the liver. After 72 h, larvae were frequently observed in the lungs and began to be found in the encephalon. After 1 week, the number of larvae increased in the encephalon and decreased in liver; however, larvae reappeared in the liver 1 week later. After 30 and 60 days, larvae were more frequently observed in the encephalon. Larvae could also be found in the liver at these time points (fig. 1).

Fig. 1

The proportion (%) of Toxocara canis larvae recovered from primary infections in gerbils up to (A) 96 h and (B) 60 days post-infection; jejunum (closed circle), ileum (open circle), caecum (closed triangle), liver (open triangle), lung (closed diamond) and encephalon (closed square).

In the flattened liver and encephalon fragments, mobile larvae were easily observed under the microscope. Larvae frequently appeared without surrounding inflammatory cells in the intestinal wall, crypts and lymphatic vessels, the liver, skeletal muscles and in the encephalon until the second week after infection (fig. 2a). The focal necrosis of neurons, central chromatolysis and foci of glial proliferation were observed in the cerebellum at 7 days post-infection. Focal eosinophilic infiltrates were progressively more frequent 7 days post-infection in the liver, lungs, kidneys and heart (fig. 2b). Granulomatous reactions appeared after the first week, initially as loose macrophage and eosinophil accumulations correlating with larval antigens or larval fragments (fig. 2c). At approximately 30 days post-infection, the granulomata were more organized; peripheral fibrosis was observed in the liver, lungs, skeletal muscles and kidneys (fig. 2d). Focal areas of haemorrhaging were frequent in the lungs. Immunohistochemistry showed the presence of parasite antigens in areas of inflammatory infiltrates, in the vicinity of larvae and within macrophages making up granulomas in all organs.

Fig. 2

Histopathological changes to the internal tissues of gerbils on day 15 post-infection with Toxocara canis, to show (a) the brain with intact larvae (arrows) without any inflammatory reaction; (b) the kidney with eosinophilic infiltrates between the tubules (*); (c) the liver with granulomata and eosinophils and macrophages adjacent to a larva (arrow) and (d) on day 60 post-infection, lung tissue with granulomata and large numbers of eosinophils.

Between 7 and 30 days post-infection, focal areas of haemorrhaging were observed in the plexiform layer of the retina and vitreous humour of the eyes. Limited eosinophil infiltration at the base of the optic nerve, and focal areas of scleritis and episcleritis with eosinophils, were frequently observed. Focal areas of myositis with eosinophils were observed in eye muscles. These lesions were infrequent at 30 days post-infection.

Secondary infections

Manifestations of injury in motor areas, such as walking in a circle, paresis and paralysis of hind limbs, were commonly observed in re-infected animals at 30 days after re-infection.

Small nodules on the surface of the liver and lungs were more frequent and appeared earlier after re-infection than in primary infected animals. Larval distribution in the re-infected group was similar to that observed in the primary infected group. However, the number of larvae increased in the liver and was reduced in the brain at 15 days after re-infection (fig. 3). Microscopic examination of liver fragments mounted between slides revealed the presence of organized granulomata around larvae. These larvae were mobile and sometimes observed leaving the granulomata.

Fig. 3

The proportion (%) of Toxocara canis larvae recovered in gerbils re-infected 30 days following a primary infection up to (A) 72 h and (B) 60 days post-infection; jejunum (closed circle), ileum (open circle), caecum (closed triangle), liver (open triangle) and encephalon (closed square).

The inflammatory reaction was more severe, with higher number of eosinophils, in all organs compared to those in the primary infected group. Necrosis of Purkinje cells and foci of demyelization with microglia aggregates were observed in the cerebellum. Larvae were observed in the vicinity of the lesions. Focal eosinophilic meningitis was observed in all animals at different times after re-infection. Granulomata were observed in the brain and cerebellum at 15 days after re-infection, some with areas of central necrosis and calcification (fig. 4a). Although loose granulomas were frequently found during the first week after re-infection, all of the granulomata observed after the second week were typical epithelioid granulomas with frequent peripheral fibrosis. Larvae or larval fragments were observed within some granulomata (fig. 4b, c). In the eyes, lesions were similar to those observed in the primary infected group. Immunohistochemistry confirmed the presence of parasite antigens in areas of eosinophilic inflammatory infiltrates, in granulomata and in the vicinity of intact larvae (fig. 4d).

Fig. 4

Histopathological changes to the internal tissues of gerbils re-infected at 30 days following a primary infection with Toxocara canis, to show (a) the encephalon 15 days after re-infection, with microglial granuloma (arrow) and central necrosis; typical epitheliod granuloma with peripheral fibrosis and larva on day 30 after re-infection in (b) liver and (c) kidney; and (d) immunohistochemistry in encephalon at 30 days after re-infection, with larval antigens within microglia.

Discussion

The results in the primary infected group demonstrated that, in gerbils, penetration of the wall of the small intestine by Toxocara larvae was completed in 24 h, confirming reports from other studies (Burren, Reference Burren1968; Abo-Shehada et al., Reference Abo-Shehada, Al-Zubaidy and Herbert1984; Parsons & Grieve, Reference Parsons and Grieve1990; Cho et al., Reference Cho, Egami, Ohnuki, Saito, Chinone, Shichinohe, Suganuma and Akao2007; Alba-Hurtado et al., Reference Alba-Hurtado, Muñoz-Guzmán, Valdivia-Anda, Tórtora and Ortega-Pierres2009).

The presence of larvae within lymphatic vessels in intestinal mucosa at 4 h post-infection is an indirect indication that this route can be used to reach the systemic circulation, as proposed in mice by Abo-Shehada et al. (Reference Abo-Shehada, Al-Zubaidy and Herbert1984). It is interesting to note that there were essentially no inflammatory lesions around larvae in the intestinal walls of infected animals.

Larvae were quickly observed in the liver, where they were numerous, for up to 96 h post-infection. The number of larvae decreased over subsequent days and they were rarely observed in the livers of animals sacrificed at 7 days post-infection. Interestingly, the number of larvae in the liver increased 1 week later and progressively decreased until 60 days post-infection. A reduction in the larval load in the liver starting from the fifth day of infection was previously reported in gerbils (Parsons & Grieve, Reference Parsons and Grieve1990; Cho et al., Reference Cho, Egami, Ohnuki, Saito, Chinone, Shichinohe, Suganuma and Akao2007; Alba-Hurtado et al., Reference Alba-Hurtado, Muñoz-Guzmán, Valdivia-Anda, Tórtora and Ortega-Pierres2009); however, a subsequent increase in the larval load was not mentioned in any of these reports. This fluctuation in larval load has been reported in mice; a reduction was observed after 48 h, and an increase in larval load was observed at 21 days post-infection (Farjat et al., Reference Farjat, Minvielle, Pezzani and Niedfeld1995). This fluctuation in larval load in the liver may be due to a long migration via the lymphatic system, as larvae would have to go through the lymph nodes before reaching the circulation.

The pattern of larval distribution after infection observed in the lungs and brain was similar to that described by other authors in gerbils (Cho et al., Reference Cho, Egami, Ohnuki, Saito, Chinone, Shichinohe, Suganuma and Akao2007; Alba-Hurtado et al., Reference Alba-Hurtado, Muñoz-Guzmán, Valdivia-Anda, Tórtora and Ortega-Pierres2009) and in mice (Burren, Reference Burren1968; Abo-Shehada & Herbert, Reference Abo-Shehada and Herbert1984; Guardis et al., Reference Guardis, Radman, Burgos, Fonrouge and Archelli2002; Pecinali et al., Reference Pecinali, Gomes, Amendoeira, Bastos, Martins, Pegado, Bastos, Bozza and Castro-Faria-Neto2005). The observed pulmonary lesions, especially the haemorrhagic lesions, were similar to those described in mice (Mossalam et al., Reference Mossalam, Atallah and Hosney1972; Zyngier, Reference Zyngier1974). In the encephalon, larvae were found starting at 72 h post-infection, and were more abundant after 1 week, confirming previous reports in gerbils (Cho et al., Reference Cho, Egami, Ohnuki, Saito, Chinone, Shichinohe, Suganuma and Akao2007; Alba-Hurtado et al., Reference Alba-Hurtado, Muñoz-Guzmán, Valdivia-Anda, Tórtora and Ortega-Pierres2009) and mice (Burren, Reference Burren1968; Abo-Shehada & Herbert, Reference Abo-Shehada and Herbert1984; Guardis et al., Reference Guardis, Radman, Burgos, Fonrouge and Archelli2002).

The histological picture observed in the liver was similar to that observed in mice examined at the same time of infection (Zyngier, Reference Zyngier1974). Focal inflammatory lesions and poorly organized granulomata were observed in the liver. These were frequently not associated with larvae, which suggested that the larvae move rapidly and that inflammatory reactions occur in larval tracks. In fact, the granulomata observed in sections of liver mounted between microscope slides showed no larvae or had a larva moving into them. It was sometimes possible to observe larvae leaving the granulomata. Serpiginous granulomata were observed in the liver, suggesting that inflammatory reactions occur in larval tracks in response to antigens being liberated by migrating larva. The presence of loose granulomata is an indirect indication that the cellular immune response to larval antigens is slower and weaker in gerbils. Typical epithelioid granulomata around larvae have been described more frequently in the livers of rats, chickens and guinea pigs at 10 days post-infection (Mossalam et al., Reference Mossalam, Atallah and Hosney1972); these findings likely reflect more effective adaptive immune responses against larval antigens in these species.

Regarding histological changes observed in the brain, inflammatory reactions were small or absent around the larvae during primary infection; these findings support those previously described in mice and gerbils (Mossalam et al., Reference Mossalam, Atallah and Hosney1972; Akao et al., Reference Akao, Tomoda, Hayashi, Suzuki, Shimizu-Suganuma, Shichinohe and Fujita2003). Focal necrosis of neurons in the cerebellum was especially frequent. Cerebellar lesions without inflammatory reactions were also reported in gerbils by Akao et al. (Reference Akao, Tomoda, Hayashi, Suzuki, Shimizu-Suganuma, Shichinohe and Fujita2003). However, such lesions have not been reported in other rodents used as models of T. canis infection.

The lesions observed in kidneys and skeletal muscles were similar to those described in mice (Mossalam et al., Reference Mossalam, Atallah and Hosney1972; Kayes & Oaks, Reference Kayes and Oaks1978).

Contrary to what has been reported previously in gerbils, we did not observe severe lesions in the eyes after primary infection; larvae were rarely observed. We do not have a complete explanation for this observed discrepancy. It is likely due to the age of the gerbils, which were 45 and 60 days old, or the amount of inoculum used; inocula with a low number of larvae would be less effective in inducing immune responses in young animals, favouring the migration of larvae (Glickman & Schantz, Reference Glickman and Schantz1981; Zanandréa et al., Reference Zanandréa, Oliveira, Abreu and Pereira2008).

Immunohistochemistry with polyclonal anti-Toxocara serum demonstrated that the inflammatory infiltrates and granulomata observed in all organs were correlated with larvae or larval antigens. In infected humans, Toxocara granulomata were strongly stained with a polyclonal anti-Toxocara serum; stained Toxocara antigen was observed within macrophages in these granulomas (Musso et al., Reference Musso, Castelo, Tsanaclis and Pereira2007). The inflammatory cells of experimentally infected sheep, especially macrophages in the intestine, liver and lungs, were stained with anti-Toxocara serum raised in mice (Aldawek et al., Reference Aldawek, Levkut, Revajová, Kolodzieyski, Seveiková and Dubinský2002)

In re-infected animals, a dramatic change in larval migration was observed after the first 72 h. This difference was observed in the redistribution of larvae, which remained numerous in the liver 60 days after secondary infection, with a peak in abundance at 48 h. This observation was similar to that reported in re-infected mice (Zyngier, Reference Zyngier1974; Parsons & Grieve, Reference Parsons and Grieve1990; Wnukowska & Dzbenski, Reference Wnukowska and Dzbenski2008). Our findings demonstrate that larvae are retained in the liver for a longer time than in primary infected animals. Histological studies in mice also showed that epithelioid granulomata are larger after re-infection and surrounded by fibrous capsules (Zyngier, Reference Zyngier1974; Wnukowska & Dzbenski, Reference Wnukowska and Dzbenski2008). Upon re-infection, granulomata were present in liver fragments flattened between two slides. Larvae were observed moving inside these granulomata, suggesting that the larvae leave the granulomata and continue their migration. In fact, dead larvae were infrequent in the granulomas. The granulomatous reaction around larvae in the liver would explain their retention, as it would delay larval migration to other sites.

Granulomas with larvae, which were sometimes dead, were found more frequently in the encephalon and cerebellum of the secondary infected group than in those of the primary infected group. This is similar to what was observed in the liver. These granulomata are responsible for the more severe neurological symptoms observed in re-infected animals.

In all tissues, the granulomata around larvae or larval antigens were larger than those observed in the primary infected group. Although an evaluation of the immune response to Toxocara antigens has not been performed, the presence of large epithelioid granulomata in re-infected animals suggests a strong adaptive immune response against Toxocara antigens in this group. In fact, athymic nude mice infected with T. canis develop fewer and looser granulomata than control mice infected with the same larval load. This finding confirms the role of the cell-mediated immune response in the development of granulomas around Toxocara larvae or larval antigens (Sugane & Oshima, Reference Sugane and Oshima1983; Wnukowska & Dzbenski, Reference Wnukowska and Dzbenski2008).

In conclusion, our results demonstrated that larval migration during primary infection in gerbils is similar to that observed in mice. This migration has a visceral initial phase, in which larvae predominate in the liver and lungs. During the later neurotropic phase, larvae concentrate in the central nervous system. After re-infection, the visceral phase is amplified, and a great number of larvae are retained in the liver. Inflammatory lesions, especially granulomata, are more intense after re-infection. This finding suggests that the cellular immune response against larval antigens was greater than that observed during primary infection. Despite this intense granulomatous inflammatory reaction, the destruction of larvae within these granulomata was not frequently observed.

Financial support

This research was supported by intramural funding from the Infectious Diseases Unit at the Health Sciences Center, Federal University of Espirito Santo.

Conflict of interest

None.

Ethical standards

The authors assert that all procedures contributing to this work comply with the ethical standards of the relevant national and institutional guides for the care and use of laboratory animals. Experiments were approved by the Ethics Committee for the Use of Animals at the Federal University of Espirito Santo (n. 007/2008).

References

Abo-Shehada, M.N. & Herbert, I.V. (1984) The migration of larval Toxocara canis in mice. II. Post-intestinal migration in primary infections. Veterinary Parasitology 17, 75–83.CrossRefGoogle ScholarPubMed
Abo-Shehada, M.N., Al-Zubaidy, B.A. & Herbert, I.V. (1984) The migration of larval Toxocara canis in mice. I. Migration through the intestine in primary infections. Veterinary Parasitology 17, 65–73.Google Scholar
Akao, N., Tomoda, M., Hayashi, E., Suzuki, R., Shimizu-Suganuma, M., Shichinohe, K. & Fujita, K. (2003) Cerebellar ataxia due to Toxocara infection in Mongolian gerbils, Meriones unguiculatus . Veterinary Parasitology 113, 229–237.Google Scholar
Alba-Hurtado, F., Tórtora, P.J., Tsutsumi, V. & Ortega-Pierres, M.G. (2000) Histopathological investigation of experimental ocular toxocariasis in gerbils. International Journal for Parasitology 30, 143–147.Google Scholar
Alba-Hurtado, F., Muñoz-Guzmán, M.A., Valdivia-Anda, G., Tórtora, J.L. & Ortega-Pierres, M.G. (2009) Toxocara canis: larval migration dynamics, detection of antibody reactivity to larval excretory–secretory antigens and clinical findings during experimental infection of gerbils (Meriones unguiculatus). Experimental Parasitology 122, 1–5.CrossRefGoogle ScholarPubMed
Aldawek, A.M., Levkut, M., Revajová, V., Kolodzieyski, L., Seveiková, Z. & Dubinský, P. (2002) Larval toxocarosis in sheep: the immunohistochemical characterization of lesions in some affected organs. Veterinary Parasitology 105, 207–214.Google Scholar
Aljeboori, T.I., Stout, C. & Ivey, M.H. (1970) Toxocara canis infections in baboons. II. Distribution of larvae and histopathologic responses. American Journal of Tropical Medicine and Hygiene 19, 815–820.Google Scholar
Bardón, R., Cuéllar, C. & Guillén, J.L. (1994) Larval distribution of Toxocara canis in BALB/c mice at nine weeks and one year post-inoculation. Journal of Helminthology 68, 359–360.Google Scholar
Burren, C.H. (1968) Experimental toxocariasis I. Some observations on the histopathology of the migration of Toxocara canis larvae in the mouse. Zeitschrift für Parasitenkunde 30, 152–161.Google Scholar
Camparoto, M.L., Fulan, B., Colli, C.M., Paludo, M.L., Falavigna-Guilherme, A.L. & Fernandez, M.A. (2008) Initial stage of development and migratory behavior of Toxocara canis larvae in BALB/c mouse experimental model. Genetics and Molecular Research 7, 444–450.Google Scholar
Cho, S., Egami, M., Ohnuki, H., Saito, Y., Chinone, S., Shichinohe, K., Suganuma, M. & Akao, N. (2007) Migration behaviour and pathogenesis of five ascarid nematode species in the Mongolian gerbil Meriones unguiculatus . Journal of Helminthology 81, 43–47.Google Scholar
Despommier, D. (2003) Toxocariasis: clinical aspects, epidemiology, medical ecology, and molecular aspects. Clinical Microbiology Reviews 16, 265–272.Google Scholar
Epe, C., Sabel, T., Schnieder, T. & Stoye, M. (1994) The behavior and pathogenicity of Toxocara canis larvae in mice of different strains. Parasitology Research 80, 691–705.Google Scholar
Farjat, J.A., Minvielle, M.C., Pezzani, B.C. & Niedfeld, G. (1995) Relationship between parasitical inoculum and immunological parameters in experimental toxocariasis. Zentralblatt für Bakteriologie 282, 465–473.Google Scholar
Galvin, T.J. (1964) Experimental Toxocara canis infections in chickens and pigeon. Journal of Helminthology 50, 124–127.Google Scholar
Glickman, L.T. & Schantz, P.M. (1981) Epidemiology and pathogenesis of zoonotic toxocariais. Epidemiologic Reviews 3, 230–250.Google Scholar
Glickman, L.T. & Summers, B.A. (1983) Experimental Toxocara canis infection in cynomolgus macaques (Macaca fascicularis). American Journal of Veterinary Research 44, 2347–2354.Google Scholar
Guardis, M.V., Radman, N., Burgos, L., Fonrouge, R.E. & Archelli, S.M. (2002) Migración larval y eosinofilia en el hospedador paraténico. Parasitologia Latinoamerican 57, 46–49.Google Scholar
Helwigh, A.B., Lind, P. & Nansen, P. (1999) Visceral larva migrans: migratory pattern of Toxocara canis in pigs. International Journal for Parasitology 29, 559–565.CrossRefGoogle ScholarPubMed
Kayes, S.G. (1997) Human toxocariasis and the visceral larva migrans syndrome: correlative immunopathology. Chemical Immunology 66, 99–124.Google Scholar
Kayes, S.G. & Oaks, J.A. (1978) Development of the granulomatous response in murine toxocariasis. Initial events. American Journal of Pathology 93, 277–294.Google Scholar
Lescano, S.Z., Queiroz, M.L. & Chieffi, P.P. (2004) Larval recovery of Toxocara canis in organs and tissues of experimentally infected Rattus norvegicus . Memorias do Instituto Oswaldo Cruz 99, 627–628.Google Scholar
Maruyama, S., Nino, T., Yamamoto, K. & Katsube, Y. (1994) Parasitism of Toxocara canis larvae in chickens inoculated with the ascarid eggs. Journal of Veterinary Medical Science 56, 139–141.Google Scholar
Moreira-Silva, S.F., Rodrigues, M.G., Pimenta, J.L., Gomes, C.P., Freire, L.H. & Pereira, F.E.L. (2004) Toxocariasis of the central nervous system: with report of two cases. Revista da Sociedade Brasileira de Medicina Tropical 37, 169–174.Google Scholar
Mossalam, I., Atallah, O.A. & Hosney, Z. (1972) Experimental studies on visceral larva migrans of Toxocara canis in laboratory animals. Acta Veterinaria Academiae Scientiarum Hungaricae 22, 71–80.Google Scholar
Musso, C., Castelo, J.S., Tsanaclis, A.M.C. & Pereira, F.E.L. (2007) Prevalence of Toxocara-induced liver granulomas, detected by immunohistochemistry, in a series of autopsies at a Children's Reference Hospital in Vitoria, ES, Brazil. Virchows Archive 450, 411–417.Google Scholar
Ollero, M.D., Fenoy, S., Cuéllar, C., Guillén, J.L. & Del Aguila, C. (2008) Experimental toxocariosis in BALB/c mice: effect of the inoculation dose on brain and eye involvement. Acta Tropica 105, 124–130.Google Scholar
Parsons, J.C. & Grieve, R.B. (1990) Kinetics of liver trapping of infective larvae in murine toxocariasis. Journal of Helminthology 76, 529–536.Google Scholar
Pawlowski, Z. (2001) Toxocariasis in humans: clinical expression and treatment dilemma. Journal of Helminthology 75, 299–305.Google Scholar
Pecinali, N.R., Gomes, R.N., Amendoeira, F.C., Bastos, A.C., Martins, M.J., Pegado, C.S., Bastos, O.M., Bozza, P.T. & Castro-Faria-Neto, H.C. (2005) Influence of murine Toxocara canis infection on plasma and bronchoalveolar lavage fluid eosinophil numbers and its correlation with cytokine levels. Veterinary Parasitology 134, 121–130.Google Scholar
Smith, H.V., Quinn, R., Bruce, R.G. & Girdwood, R.W. (1982) Development of the serological response in rabbits infected with Toxocara canis and Toxascaris leonine. . Transactions of the Royal Society of Tropical Medicine and Hygiene 76, 89–94.Google Scholar
Sommerfelt, I.E., Rosa, A., Duchene, A., Degregorio, O., López, C., Pisanú, A. & De Torres, R. (2004) Toxocara canis in experimentally infected pigs: migratory pattern and tissue lesions. Veterinary Parasitology 125, 323–334.Google Scholar
Sugane, K. & Oshima, T. (1983) Trapping of large numbers of larvae in the livers of Toxocara canis-reinfected mice. Journal of Helminthology 57, 91–99.Google Scholar
Taira, K., Permin, A. & Kapel, C.M. (2003a) Establishment and migration pattern of Toxocara canis larvae in chickens. Parasitology Research 90, 521–523.Google Scholar
Taira, K., Saeed, I., Lind, P., Murrell, K.D. & Kapel, C.M. (2003b) Population dynamics of Toxocara canis in pigs receiving a single or multiple infection. Parasitology 127, 593–602.Google Scholar
Takayanagi, T.H., Akao, N., Suzuki, R., Tomoda, M., Tsukidate, S. & Fujita, K. (1999) New animal model for human ocular toxocariasis: ophthalmoscopic observation. British Journal of Ophthalmology 83, 967–972.Google Scholar
Wnukowska, N. & Dzbenski, T.H. (2008) Studies on the development and role of some cell-mediated immune response in experimental toxocariasis in mice. Acta Parasitology 53, 205–210.Google Scholar
Zanandréa, L.I., Oliveira, G.M., Abreu, A.S. & Pereira, F.E.L. (2008) Ocular lesions in gerbils (Meriones unguiculatus) infected with low larval burden of Toxocara canis: observations using indirect binocular ophthalmoscopy. Revista da Sociedade Brasileira de Medicina Tropical 41, 570–574.Google Scholar
Zyngier, F.R. (1974) Histopathology of experimental toxocariasis in mice. Annals of Tropical Medicine and Parasitology 68, 225–228.Google Scholar
Figure 0

Fig. 1The proportion (%) of Toxocara canis larvae recovered from primary infections in gerbils up to (A) 96 h and (B) 60 days post-infection; jejunum (closed circle), ileum (open circle), caecum (closed triangle), liver (open triangle), lung (closed diamond) and encephalon (closed square).

Figure 1

Fig. 2Histopathological changes to the internal tissues of gerbils on day 15 post-infection with Toxocara canis, to show (a) the brain with intact larvae (arrows) without any inflammatory reaction; (b) the kidney with eosinophilic infiltrates between the tubules (*); (c) the liver with granulomata and eosinophils and macrophages adjacent to a larva (arrow) and (d) on day 60 post-infection, lung tissue with granulomata and large numbers of eosinophils.

Figure 2

Fig. 3The proportion (%) of Toxocara canis larvae recovered in gerbils re-infected 30 days following a primary infection up to (A) 72 h and (B) 60 days post-infection; jejunum (closed circle), ileum (open circle), caecum (closed triangle), liver (open triangle) and encephalon (closed square).

Figure 3

Fig. 4Histopathological changes to the internal tissues of gerbils re-infected at 30 days following a primary infection with Toxocara canis, to show (a) the encephalon 15 days after re-infection, with microglial granuloma (arrow) and central necrosis; typical epitheliod granuloma with peripheral fibrosis and larva on day 30 after re-infection in (b) liver and (c) kidney; and (d) immunohistochemistry in encephalon at 30 days after re-infection, with larval antigens within microglia.