Larval development of Toxocara canis in dogs

Veterinary Parasitology 175 (2011) 193–206

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Veterinary Parasitology journal homepage: www.elsevier.com/locate/vetpar

Review

Larval development of Toxocara canis in dogs Thomas Schnieder ∗ , Eva-Maria Laabs, Claudia Welz Institute for Parasitology, University of Veterinary Medicine, Buenteweg 17, D-30559 Hannover, Germany

a r t i c l e

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Article history: Received 25 June 2010 Received in revised form 7 October 2010 Accepted 12 October 2010 Keywords: Toxocara canis Toxocarosis Infection routes Larval development Prenatal infection Zoonosis

a b s t r a c t The parasitic roundworm Toxocara canis is present in dog populations all over the world. Due to its zoonotic potential, this roundworm is of special interest not only for veterinarians, but also for medical practitioners. In the present review, current knowledge of infection routes and the subsequent development of larvae within the canine host is summarised. Furthermore, information about the clinical, pathological, enzymatic, haematological and histopathological changes was collected, giving a broad overview of current knowledge of the infection. Although the data collected over the years give an idea of what happens during the larval development of T. canis, many questions remain open. Nevertheless, it is important that we continue our efforts to further understand the biology of this versatile and compelling parasite and try to improve and optimise strategies to prevent the infection in dogs and thereby to protect humans from this infection. © 2010 Elsevier B.V. All rights reserved.

Contents 1. 2.

3.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Larval development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Oral infection with T. canis eggs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.1. The infective stage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2. The first part of the migration route . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.3. The impact of age and acquired immunity on larval migration route . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.4. The impact of gender on prevalence of patent infections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.5. The influence of the infection dose on the course of infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.6. Development into adults . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.7. Somatic migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Oral infection through paratenic hosts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Prenatal infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.1. Reactivation of larvae during pregnancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.2. Migration and development of larvae in the puppies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Lactogenic infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4.1. Distribution of larvae in the mammary glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. Oral infection with juvenile intestinal larvae and periparturient immunosuppression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5.1. The putative impact of hormonal changes in bitches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pathological changes in the host during larval migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Clinic and gross changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

∗ Corresponding author. Tel.: +49 511 953 8711; fax: +49 511 953 8870. E-mail address: [email protected] (T. Schnieder). 0304-4017/$ – see front matter © 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2010.10.027

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3.2.

4.

5.

Haematological and enzymatic changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1. Anaemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2. Eosinophilia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.3. Increased liver enzyme levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Histopathological changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Evasion of the immune system . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1. The antigenic coat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2. The interaction of TES molecules with the immune system . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Toxocara canis (Werner, 1782) is one of the most important gastrointestinal helminths in dogs, with infections reported from all parts of the world. In Argentina, 16.35% of 1944 fresh dog faecal samples collected from both urban and rural areas were positive for T. canis (Soriano et al., 2010). A study in Nigeria in 2007 reported 33.8% of 269 household dogs to be positive, with a prevalence of 51.4% in puppies up to 6 months (Sowemimo, 2007). Another Nigerian study determined the overall prevalence in 396 dogs to be 41%, with younger dogs being more susceptible to the infection (Ugbomoiko et al., 2008). A similarly high infection rate (45.2%) was reported for 438 slaughtered dogs in abattoirs in a Chinese province, determined by counting the adult worms (Dai et al., 2009). Several studies confirm that T. canis infections are also common in the European canine population. A study performed in Italy diagnosed 33.6% out of 295 dogs going to veterinary clinics for whatever reason as T. canis positive (Habluetzel et al., 2003). In Hannover, Germany, 2.2% of the faecal samples from 1281 dogs examined as part of diagnostic services from 1998 to 2002 were positive for eggs of this parasite (Epe et al., 2004). A similar prevalence (2.1%) was determined for the years 2003–2009 (unpublished data). In Serbia, Nikolic et al. (2008) examined stool samples from 151 household, stray, and military dogs, revealing a prevalence as high as 30.5%, whereas 1159 faecal samples tested in Northern Belgium showed a T. canis prevalence of 4.4% in household dogs (Claerebout et al., 2009). The same study identified a prevalence of 26.3% in breeding kennel dogs. In the UK, Batchelor et al. (2008) included 4526 dogs with clinical signs of gastrointestinal disease in their prevalence study, with 1.4% being positive. In a recent German study, out of 445 dogs taken in by animal shelters 2.5% of adult dogs were positive for T. canis, whereas dogs up to 1 year of age were positive at a rate of 8.8% (Rohen, 2009). In the Netherlands, a coproscopic prevalence of 4.4% of T. canis eggs was determined in 92 clinically healthy dogs (Overgaauw et al., 2009). However, the prevalences determined in the cited studies cannot be compared directly to each other as the examined populations were quite different. Nevertheless, T. canis is undeniably present in the canine population worldwide, especially in young dogs. Working dogs and stray dogs seem to be at a higher risk of getting infected than pet dogs (Nikolic et al., 2008). Nonetheless, not only do excreted eggs in dog faeces and contaminating soil provide a human infection risk, but also do eggs attached to dog hair. Wolfe and Wright (2003)

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examined the hair of 60 dogs in Ireland and the UK, with the result that T. canis eggs were found in 25% of the cases. Of these eggs, 78.9% were viable and 4.2% embryonated, emphasising the potential infection risk from these eggs. These findings were supported by a study which examined the hair of 100 stray dogs in Ireland with the result that 67% of the dogs were found to have T. canis eggs on their hair (Roddie et al., 2008b). These authors examined not only dog hair for T. canis eggs but also fox hair, revealing 28% of 87 foxes to be positive. 61.4% of these eggs were either viable or embryonating (Roddie et al., 2008a). In Turkey, Aydenizöz-Ozkayhan et al. (2008) found T. canis eggs in the hair of 21.56% of 51 dogs. All the eggs were viable, 21% being either embryonating or embryonated. Overgaauw et al. (2009) detected T. canis eggs in 12.2% of 148 fur samples of clinically healthy dogs. In this study, none of the eggs were viable. The actual impact of T. canis eggs within dog fur on human health remains to be determined, but as pets literally share house and bed with their owners, this source of infection might be underestimated. The wide distribution of T. canis and its importance as a zoonotic agent, causing visceral and ocular larva migrans in humans, have led to an increasing interest in the biology and epidemiology of this parasite. For efficient prevention of infections in dogs, understanding its complex life cycle, including infections resulting from ingestion of either embryonated eggs from the environment or larvae from infected paratenic hosts, as well as infections from vertical transmission (intrauterine or lactogenic), is crucial. However, there are still unanswered questions concerning the life cycle of this nematode. In this review, current knowledge regarding larval development in dogs, mechanisms of larval reactivation from the tissue, immune evasion and pathological changes in organs after invasion will be outlined and discussed. As some of the previous studies were published in German only, their inclusion in this review might be of special interest. 2. Larval development 2.1. Oral infection with T. canis eggs 2.1.1. The infective stage Eggs of T. canis are unembryonated when shed with the faeces. Under optimal conditions, i.e. temperatures between 25 and 30 ◦ C and relative humidity of 85–95%, the development of the infective larval stage within the egg requires 9–15 days (Schacher, 1957; Okoshi and Usui, 1968). Nevertheless, depending on soil type and climatic

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conditions, development can vary from 3 to 6 weeks up to several months, and the infective stage in the egg can remain viable for at least 1 year (Overgaauw, 1997). Thereby, the infective larval stage is not, as was believed for many years, the second larval stage but the third one, which emerges after two moults in the egg (Araujo, 1972; Brunaska et al., 1995). 2.1.2. The first part of the migration route After ingestion by the host, eggs hatch in the duodenum within 2–4 h. The released, infective larvae penetrate the mucosa of the intestine (Webster, 1958b). The penetration mechanism is as yet not fully understood. However, an elastase-like protease was detected in larval excretory–secretory (E/S) products (Robertson et al., 1989). This protease might assist larvae in penetrating host tissues, i.e. the intestinal mucosa, liver or kidney parenchyma and the walls of blood vessels. Furthermore, direct mechanical disruption is discussed (Parsons, 1987). After penetrating the intestinal wall, larvae invade lymph vessels and migrate to the mesenteric lymph nodes (Webster, 1958b). Thereafter, they travel in venous capillaries via the portal circulation to the liver (Webster, 1958a). The majority of the larvae reach the liver approximately 24 h post infection (p.i.) (Webster, 1958a). Subsequently, within 12 h most larvae continue to migrate, exiting the liver via Vena cava, passing the heart and reaching the lung via the pulmonary artery. The larvae occur there between 24 and 36 h p.i., and numbers increase up to 96 h (Webster, 1958a). Webster (1958b) demonstrated the passive haematogenous transport by detecting T. canis larvae in the blood of the heart as well as in the pulmonary artery 72 h post infection. Those larvae which are trapped in capillaries and cannot continue the migration remain in the liver. After encapsulation, these larvae cause the mottled appearance and the whitish spots characteristic of T. canis infection (Webster, 1958a). 2.1.3. The impact of age and acquired immunity on larval migration route From the lung, larvae follow two different routes, depending on the age and immune status of the host and the infection dose. Firstly, they can penetrate the alveoli wall and continue their migration via bronchioles and trachea to the pharynx. Here they are swallowed and finally grow into adult worms in the intestine. On the second route, larvae again penetrate the alveoli wall and re-enter the circulatory system, being distributed to the somatic tissue (Webster, 1958b). Greve (1971) compared 3-week-, 3-month- and 1-year-old dogs from an ascarid-naïve colony after experimental subcutaneous infection with T. canis. In 3-week-old puppies almost all larvae had migrated through the “tracheal migration” route. In contrast, in 3month- and 1-year-old dogs most larvae were discovered in granulomes in the tissue. Therefore, it was concluded that the likelihood of somatic migration progressively increases from the age of 3 months onwards. Concurrently, the development of larvae into adult ascarids decreases (Greve, 1971; Oshima, 1976). This phenomenon is referred to as age resistance, which is firstly based on the development of immune competence and secondly on acquired

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immunity (Barriga, 1988). Immunity is assumed to occur on the one hand in the lung as a putative delayed type hypersensitivity response (Dubey, 1978), affecting thirdstage larvae after reinfection (Oshima, 1976). On the other hand, immunity is also located in the gastro-intestinal tract, preventing the infective larval stage from penetrating the intestinal mucosa after reinfection. The restricted penetration might be ascribed to an inflammatory allergic reaction after the release of vasoactive amines by sensibilised mastocytes (Soh and Kim, 1973), this possibly also being the explanation for the catarrhal-haemorrhagic diarrhea described in reinfected dogs (Löwenstein, 1981). This statement is supported by elevated gamma globulin values in the exudate and tissue of the intestine (Soh and Kim, 1973). The degree of inhibition was examined by Löwenstein (1981) who discovered that after reinfection of lactating bitches with embryonated eggs, the distribution of larvae in the somatic tissue and the elimination in the milk were 99% lower than in the group infected only once. Zimmermann (1983) determined that a trickle primary infection during 10 days resulted in a 28% lower distribution of larvae compared to a single infection with the same total number of embryonated eggs, thus demonstrating the rapid development of the immune response. Additionally, studies showed that acquired immunity is stage specific against third stage larvae only. Juvenile intestinal larvae (L4) introduced directly into the intestine of previously immunised dogs were able to grow into adult worms at a rate of 46–60%, whereas infective third stage larvae implanted in a similar manner did not develop into adults. Furthermore, no antibody response was detected after implantation of fourth stage larvae (Fernando et al., 1973). These results were confirmed by a study showing that implantation of juvenile intestinal larval stages in five 1- to 2-year-old male dogs, produced patent infections in all dogs. The same was achieved after a first and a second retransplantation (Brunschön-Hellmich, 1987). Furthermore, the immune system is unable to completely eliminate tissue-arrested parasites. Possible reasons will be discussed later on. 2.1.4. The impact of gender on prevalence of patent infections Some studies suggest that the migration route of the parasite is influenced not only by age and immunity but also by gender. It has been observed that patent T. canis infections occur more frequently in adult male dogs (older than 12 months old) than in females of the same age (Ehrenford, 1957; Turner and Pegg, 1977). This was discussed to be part of a survival strategy in terms of evolution, as females spread the infection via reactivated somatic tissue larvae to their offspring, whereas Toxocara infections in males can only be distributed by patent intestinal infections (Overgaauw, 1997). Webster (1958a) also reported that more larvae migrate in the somatic tissues of female dogs compared to those of male dogs. In a survey with 1324 dogs, Ehrenford (1957) found 32.8% of the males in contrast to only 9.4% of the females to be infected with adult T. canis. Furthermore, it was stated that in male dogs there was no evidence of immunity up to 36 months of age, whereas females showed an

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increasing immunity from 6 to 36 months (Ehrenford, 1957). No protective immunological response was detected in 3 adult male greyhounds, which were repeatedly infected with 100–200 viable eggs and became patent even though serum antibodies to the parasite surface as well as to secreted antigens were detected (Maizels and Meghji, 1984). Maizels and Meghji (1984) discussed as possible reasons firstly a greater susceptibility of this breed to certain parasite infections, secondly the gender and thirdly the infection dose. In contrast, in another, more recent, study, female dogs were reported to have a higher prevalence (Bridger and Whitney, 2009). However, the authors themselves acknowledge that the sample size of 28 dogs with known sex was low. Thus, the results may not be comparable to other studies. However, whether the gender really influences the migration and development of T. canis is doubtful, since most prevalence studies have not seen any difference between male and female dogs (Fontanarrosa et al., 2006; Daryani et al., 2009; Rohen, 2009; Gingrich et al., 2010). 2.1.5. The influence of the infection dose on the course of infection Comparing the course of infection after infection with different numbers of infective eggs, it is obvious that the antigenic stimulus has an impact on the development of protective immunity. Dubey (1978) experimentally infected 45 ascarid-naïve 2-monthold puppies and 6 adult dogs. In 24 out of 25 pups infected with 10–1000 embryonated eggs a patent infection was established. In contrast, no patent infection was detectable in 20 puppies infected with 10,000 eggs. Furthermore, three of the six adult dogs (7, 10 and 52 months old) developed patent infections after being fed with 100 embryonated eggs each (Dubey, 1978). Similarly, a study on 18 beagles confirmed the results of Maizels and Meghji (1984) that low infection doses, i.e. 100 embryonated eggs, cause patent infections also in older dogs (Fahrion et al., 2008). Therefore, adult dogs considered immune against T. canis infections were shown to develop patent infections after reinfection if the number of infective larvae and thus the antigen stimulus is low. Thus, it can be assumed that the number of adult dogs with patent infections and the contamination of the environment with eggs might be higher than so far assumed. 2.1.6. Development into adults As mentioned above, the lung is the crucial organ in which parting of the ways occurs: larvae migrate either towards the intestine to establish a patent infection or towards somatic tissue to become a larva migrans. To complete their way to a patent infection, larvae penetrate the alveoli and reach, after passing the bronchioles, the trachea. After 7–9 days larvae can be detected in the trachea and in the esophagus. The gastrointestinal tract is reached 7–15 days after infection (Sprent, 1958). From early studies it is hard to determine where larval moults actually occur, because these authors considered the infective larvae to be second stage larvae and described a total of three moults. More recent knowledge, however, proves that infective larvae in the egg are already third stage larvae

Fig. 1. T. canis larva leaving a lung vessel (Manhardt, 1980).

and only two moults occur inside the host (Araujo, 1972; Brunaska et al., 1995). According to Webster (1958a) one moult occurs either in the lungs and trachea or in the esophagus, the next in the stomach, and a further one in the duodenum. Schacher (1957) also reported about findings of third and fourth stage larvae in the stomach and about fourth and fifth stage larvae in the duodenum. Although conclusive studies are missing, it may be assumed that the first moult to L4 happens somewhere between entering the bronchioles and passing the stomach. Without any doubt the last moult to the preadult stage (L5) takes place in the small intestine. When preadults have matured, first eggs are shed approximately 4–5 weeks p.i. in experimentally infected puppies (Webster, 1958a), whereas in older hosts prepatency seems to be extended to 40 up to 56 days p.i. (Fahrion et al., 2008). It is believed that each adult ascarid lives for 4 months on average (Parsons, 1987). Experimentally infected dogs are reported to expel 53% of their adult worm burden within a period of 8 days, beginning 1 week after the onset of patency (Burke and Roberson, 1979). 2.1.7. Somatic migration As puppies mature, the disposition for tracheal migration and development of a patent infection falls to a low level. Instead, most larvae re-enter the capillary system to finally perform somatic migration. Many of the earlier studies lack information about whether truly ascarid-naïve animals have been used. To gain a better understanding of the way larvae travel in the lung and thereafter in adult, so far ascarid-naïve dogs, Manhardt (1980) injected 200,000 infective larvae into the Vena cephalica antebrachii of 1- to 3-year-old dogs. After 1 h p.i., 75% of the administered larvae were recovered from the lung, whereas no larvae could be detected in the arterial blood. In histological slides larvae could be detected moving inside the lung vessels as well as leaving them (Fig. 1). Furthermore, they were found migrating freely in the interstitium and finally reaching the lung alveoli (Fig. 2). Somatic migration in the lung leads to many petechial haemorrhages, giving the lung a stippled appearance (Webster, 1958a; Manhardt, 1980). Manhardt’s (1980) findings confirmed Oshima’s (1976) assumption that somatic migration is performed by larvae which are

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Fig. 2. T. canis larva in the lung, interstitial pneumonia (Institute for Parasitology, archive).

trapped in capillaries, causing them to penetrate the walls and migrate through the tissue to re-enter the vascular system. This somatic migration ability leads to the appearance of larvae in organs close to the lung and in the pleural cavity. Furthermore, Manhardt (1980) examined the kidney more closely, an organ commonly highly affected by T. canis larvae, but rarely showing serious organ failure. Larvae leave the blood vessels within the cortex, leaving small haemorrhages under the capsule of the kidney, and start a somatic migration (Fig. 3). Some larvae penetrate the urine canaliculi, being detectable up to the renal medulla and even as far as in the urine itself, whereas most larvae remain after a short somatic migration close under the capsule where they are encapsulated in granulomes (Fig. 4). Shortly after the start of the haematogenous transport, larvae were also discovered between the fibrillae of the heart muscle (Manhardt, 1980). The time period in which most larvae were distributed was determined as 24–72 h (Manhardt, 1980). At the same time, the number of larvae in organs other than the lungs increased correspondingly, as shown in Table 1. A surprising phenomenon was observed for the numbers counted after 48 and 72 h in the front legs, which were higher than in the hind legs at the same time. However, this difference was no longer present at day 28 p.i. Regarding the distribution of T. canis larvae in the body of infected dogs, the vast majority of migrating larvae end

Fig. 3. T. canis larvae close to glomerula in the kidney capsule (Institute for Parasitology, archive).

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Fig. 4. T. canis larva encapsulated under the kidney capsule (Manhardt, 1980).

up in the skeletal muscles and the kidneys, but also the liver and central nervous system are affected (Webster, 1958b; Bosse et al., 1980; Manhardt, 1980). In addition, Manhardt (1980) infected an adult bitch orally with 20,000 embryonated eggs. Based on the number of recovered larvae from tissue samples, she calculated that 78.3% of the larvae were contained within the skeletal muscles and 20.1% within the kidneys. In dogs the brain is in general only affected by low numbers of migrating larvae. After digestion of half of the brain of one 3-month-old dog, Greve (1971) found only 7 larvae, whereas no larvae could be recovered in the 1year-old and the other 3-month-old dogs. This contrasts to the situation in paratenic hosts, where the brain and the eyes, directly connected via the Nervus opticus, are severely affected. 2.2. Oral infection through paratenic hosts Besides direct oral ingestion of eggs, dogs can also be infected after ingestion of paratenic hosts which carry T. canis larvae in their tissue. Paratenic hosts can be, among others, birds (Okoshi and Usui, 1968), rodents, e.g. mice (Sprent, 1953), but also rabbits (Sprent, 1955), pigs (Sasmal et al., 2008), foxes (Saeed and Kapel, 2006), and humans (Ito et al., 1986). Paratenic hosts become infected by ingestion of embryonated eggs. There are reports of larvae staying alive for at least 2 years in the tissue of mice, rats, guinea pigs and rabbits (Beaver, 1956) and surviving several weeks in frozen carcasses of mice (Sprent, 1953). In chickens and pigeons they were found to survive for at least 3 months (Beaver, 1956). A number of studies showed that Toxocara infections clearly induce behavioural changes in mice as a model for paratenic hosts. Infected mice are less active and explorative and less aversive to open areas. As a result they are at a higher risk of being caught by predators than uninfected animals (Meyer zur Heyde, 1984; Cox and Holland, 1998, 2001; Hamilton et al., 2006). After infection of dogs with paratenic hosts, Sprent (1958) observed that in some cases intestinal infections developed directly without tracheal migration. The most likely reason for this was that larvae had already migrated

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Table 1 Number of larvae in organs, body cavities and excretions at different time points after injection of 200,000 T. canis larvae into the V. cephalica antebrachii (Manhardt, 1980). Hours/days p.i. 1h Number of larvae in: Lunga Diaphragma Pleural cavity Arterial blood Skeletal musclesa , b Kidneysa Livera Hearta Braina Peritoneal cavity Tracheal mucus Urine Total number of larvae

12 h

24 h

48 h

72 h

28 days

175,637 0 0 0

135,860 1 0 0

96,531 7 22 4

47,058 353 20 15

6426 86 0 1

790 312 1 0

0 0 0 3 0 0 0 0

0 0 0 0 0 0 1 0

212 422 62 55 0 0 9 0

10,965 10,348 2862 1944 95 3 15 2

16,575 12,100 14,157 1,077 30 0 14 0

45,390 18,136 5943 2196 26 0 0 0

175,640

135,862

97,324

73,680

50,466

72,794

a

The number of larvae was calculated from the artificial digestion of 50 g tissue sample and 200 g skeletal muscle, respectively, in relation to the physiological organ weights of a beagle with a body weight of 15 kg. b The muscle sample (200 g) consisted of 50 g each of the following muscles: M. biceps dexter, M. biceps sinister, M. semitendinosus dexter, M. semitendinosus sinister.

in the tissue of the previous host. Therefore, they had already reached a stage of maturity sufficient for direct development (Overgaauw, 1997). Warren (1969), however, detected larvae also in the liver, lung, trachea, and esophagus, therefore concluding that larvae released from mice tissue also undergo tracheal migration in the dog. Herschel (1981) fed infected mice to beagles and 72 h after ingestion larvae were detected in the liver and lungs, in the arterial blood as well as in other peripheral organs of one dog. After feeding infected mice to dogs, a normal prepatent period of 34–48 days was observed (Herschel, 1981). 2.3. Prenatal infection The most important way of T. canis infection in dogs is prenatal transmission, also known as transplacental or intrauterine transmission. There are several reports stating that puppies post partum are commonly infected with T. canis. This infection can be induced by feeding embryonated eggs (Shillinger and Cram, 1923) or subcutaneously injecting “Belaskaris-larvae” (i.e. T. canis larvae) (Fülleborn, 1921) to pregnant bitches. Nonetheless, parenteral infection of puppies occurs not only after the infection of the pregnant bitch but also through reactivation of somatic tissue larvae from earlier infections (Yutuc, 1949; Webster, 1958b; Koutz et al., 1966). It is not known for sure yet how long larvae can remain and survive in the tissue of dogs. However, in a study on mice, larvae could be recovered from the tissue up to 1 year post infection (Bardón et al., 1994). In two abstract-like publications, Douglas and Baker (1959a,b) refer to a study finding that 241 and 358 days after an infection of the bitch, prenatal infections still occurred. Furthermore, it was observed that a bitch harbouring somatic larvae can infect puppies during three consecutive pregnancies (Soulsby, 1983). Koutz et al. (1966) also observed prenatal infections in foetuses from a bitch which had not been experimentally infected but was supposed to harbour somatic larvae from a previous nat-

ural infection. They further stated that the infection of the bitch has to occur until a short period post conceptionem to lead to prenatal infection of the foetuses. They came to this conclusion as they were not able to detect larvae in the progeny of bitches infected during 11–21 days post conceptionem, but indeed gave no further particulars about these data. In contrast, the group of Bosse et al. (1980) was able to induce prenatal infections in puppies at least until 7 days ante partum. In summary, this transmission mode is of major importance for puppies and may occur after reactivation of somatic larvae from previous infections of the bitch as well as from new infections during pregnancy. After infecting the bitch, larvae can either migrate directly to the offspring or remain in the somatic tissue depending on the time point of infection. Douglas and Baker (1959a,b) stated in two short notes that larvae do not reach the foetuses before the 42nd day of pregnancy. This was confirmed by Scothorn et al. (1965), who could not find larvae in foetuses of a bitch on the 35th day of pregnancy, but in 5 out of 6 foetuses of another bitch on the 56th day of pregnancy. Both bitches were not infected experimentally but assumed to be infected naturally prior to the start of the experiments. In a further study, which aimed to determine the time point of larvae appearing in the foetuses more exactly, the first larva was detected in one of six foetuses of an experimentally infected bitch on the 43rd day of pregnancy (Koutz et al., 1966). From the 47th day onwards, all foetuses of infected bitches were positive. No data are available about the time point when somatic larvae are reactivated and actually start to migrate to the foetuses. Regarding the way of transmission from the bitch to the foetus, Webster (1958a) suggested that larvae reach the placenta via the circulatory system and penetrate the delicate layer of tissue which separates the maternal and foetal blood. In the study by Scothorn et al. (1965), which is at least partially included in the analyses of Koutz et al. (1966), two larvae were found in the umbilical cord

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tissue. Therefore, the umbilical cord is regarded as a central transfer point between bitch and foetus in utero. 2.3.1. Reactivation of larvae during pregnancy The mechanism how larvae are reactivated in the somatic tissue of the bitch is still a matter of debate. However, Webster (1958a) already supposed that this phenomenon is connected to the changing hormone status. In mice, which can also transmit T. canis infections vertically, a clear decrease in tissue-arrested larvae after application of prolactin was demonstrated (Oshima, 1961). These results were confirmed in a recent study by Jin et al. (2008), who observed that migration of larvae to the mammary gland was triggered by prolactin. These findings suggest that a similar path for reactivation might also apply for dogs, although the transmission in mice mainly occurs lactogenically (Baumm, 1980) rather than prenatally. Regarding another also mainly lactogenically transmitted parasite, Ancylostoma caninum, a study showed that after administration of estrogen and progesterone to a lactating bitch more larvae were shed in the milk than beforehand (Stoye, 1973; Krause, 1975). A study on reactivation of tissue-arrested third stage larvae of A. caninum revealed that neither estrogen nor progesterone had a direct effect on feeding/reactivation of these larvae. Prolactin showed a stimulatory effect only at concentrations approximately 1000× higher than physiological levels. However, the two isoforms (1 and 2) of TGF-␤ led to significant stimulatory effects on A. caninum larvae (Arasu, 2001). Therefore, Arasu (2001) concluded that during pregnancy host-derived TGF-␤ could act on a parasite-encoded receptor which then causes the reactivation of somatic tissue larvae. It had previously been shown that there is a strong correlation between estrogen and the expression of TGF-␤2 in rats (Schneider et al., 1996). A recent study confirmed that TGF-␤ also induces the reactivation of dauer larval stages of Caenorhabditis elegans (Gallo and Riddle, 2009). As C. elegans and A. caninum are allocated to the same clade of nematodes, which is determined by the relatedness of their 18S RNA sequences, these results indicate that the reactivation of dauer stages might be preserved at least between nematodes of clade V. Whether the same mechanisms induce reactivation of T. canis, a clade III nematode, remains to be investigated. 2.3.2. Migration and development of larvae in the puppies After reactivation of tissue-arrested larvae, they migrate to the liver of the offspring, where they remain until birth (Scothorn et al., 1965; Koutz et al., 1966; Stoye, 1976). Stoye (1976) additionally found 0.4% of the recovered larvae in organs other than the liver or lungs, namely in muscles, the brain and kidneys. While he explains the presence of larvae in the lung by the already started migration, he interprets the larvae in other organs as somatic larvae after migration or distribution, as described in adult dogs. Post partum migration from the liver continues immediately. Larvae were already detectable in the lungs 30 min post partum (Koutz et al., 1966). As soon as larvae reach the lung they undergo a tracheal migration. Koutz et al. (1966) noticed a variable migra-

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tory behaviour with some larvae reaching the intestine already 2 days post partum and others remaining 2.5 days in the liver before starting migration. Nevertheless, within 3 days the majority of the larvae had reached the lung. In all cases, larvae had reached the intestine by the 7th day post partum. There are different reports concerning the duration of prepatency after prenatal infection: 21–30 days were described by Yutuc (1949) and 25–46 days by Douglas and Baker (1959a,b). Voßmann (1985), however, found a prepatent period of 28 days. Subsequently, more than 70% of the adult worm burden of puppies was expelled spontaneously 9–10 weeks post partum (Voßmann, 1985). 2.4. Lactogenic infection The lactogenic transmission is of rather subordinate importance for the distribution of T. canis. Burke and Roberson (1985a,b) compared the number of prenatally and lactogenically transmitted infections. They experimentally infected bitches 2–4 months before pregnancy. Immediately post partum and before nursing, the puppies were exchanged with newborn puppies of uninfected bitches. After 4 weeks of nursing, the puppies were necropsied and examined for T. canis infections. Therefore, prenatal and lactogenic infections could be differentiated. Only 2.3% of the total infection dose could be recovered, 98.5% of these having been transmitted prenatally and only 1.5% lactogenically (Burke and Roberson, 1985a). After infection at mid-pregnancy, Burke and Roberson (1985b) recovered 29.2% of the infection dose from the pups. 95.5% of these had been transmitted prenatally and only 4.5% were transferred lactogenically. When the bitches were infected 48 h post partum, a time point which excludes the possibility of prenatal infection, 7.9% of the infection dose could be recovered from the puppies. Similar experiments were performed by Stoye (1976), although he determined the number of larvae in the milk instead of examining the puppies. After infection of a bitch on the day of mating, he recovered 8.5% larvae of the initial infection dose in total, 99.2% of which had been transmitted prenatally to the puppies and only 0.8% having been found in the bitch’s milk. Stoye (1976) also infected a bitch on the day of birth, and recovered 9.5% of the infection dose from the milk between the 4th and the 28th day post partum. This was confirmed by Löwenstein (1981), who infected lactating bitches 10 days post partum and recovered 9.5% of larvae up to day 28 after infection. Therefore, the prenatal route appears to be the most important infection route for puppies. However, if a bitch becomes infected post partum, the lactogenic transmission is another possibility for the parasite to infect new hosts. Nevertheless, little is known about the development of larvae in puppies after ingestion of the milk. With a long prepatent period of 27–35 days after ingestion of larvae with the milk (Bosse et al., 1980), a direct intestinal development seems to be unlikely. The course of larval shedding with the milk is widely consistent. First larvae can already be detected within the first days post partum. The number of larvae increases then consistently until a maximum is reached approxi-

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Fig. 5. Larva in mammary gland alveoli (Manhardt, 1980).

mately 7–14 days post partum. During the first 2–3 weeks post partum most infections of puppies occur. However, after high infections shortly before or after birth, larvae appear in the milk approximately 4–7 days after infection and can be found in the milk at least up to day 28 after infection (Stoye, 1976; Manhardt, 1980; Löwenstein, 1981).

2.4.1. Distribution of larvae in the mammary glands To further examine the course of shedding with the milk and the distribution of larvae in the mammary gland, on the day of birth Manhardt (1980) infected bitches orally with embryonated eggs or either intravenously, subcutaneously or intra-peritoneally with larvae. Afterwards, each mammary gland complex was examined at different time points up to 28 days p.i. In all experiments, except after subcutaneous infection, shedding of larvae in the milk started 4 days p.i. However, after subcutaneous infection of larvae into the flank close to the mammary gland, larvae reached the milk and tissue already 1 day p.i., but only in the mammary gland complex nearby. 1, 2, and 3 days later larvae had also reached the neighbouring complexes but never reached all of them. Therefore, Manhardt (1980) calculated the speed of migration of larvae through tissues as 15 cm in 24 h. Hence, she concluded that somatic migration of T. canis larvae directly from the intestine via the peritoneal cavity and abdominal wall might be conceivable to a certain extent. Histological examination of the mammary gland 8 days after oral infection with 100,000 embryonated eggs showed that most larvae (565) were present, more or less curled in the alveoli of the gland (Fig. 5), whereas only few were detected in the mammary ducts (2), in the interstitium (3), and in the peri-glandular conjunctive tissue (4) (Manhardt, 1980). Larvae in the mammary ducts were stretched and in the interstitium either stretched or in a meander shape. Larvae in the peri-glandular conjunctive tissue were already being encapsulated in granulomes (Fig. 6). Larvae were found in all parts of the gland, mostly in the alveoli and no larvae were detected in a vessel. Thus, it was not possible to clarify whether larvae migrated mainly haematogenously or somatically.

Fig. 6. Larva in the mammary gland, encapsulated in periglandular tissue (Manhardt, 1980).

2.5. Oral infection with juvenile intestinal larvae and periparturient immunosuppression In the periparturient period bitches can suffer from patent T. canis infections. This may be due to different reasons. One possibility is that bitches can be infected by juvenile intestinal larvae (L4) shed with the faeces of the infected puppies (Sprent, 1961). While nursing and cleaning their puppies, bitches ingest these larvae, which directly develop into adult worms in the intestine (Lloyd et al., 1983). This is possible because, as mentioned beforehand, immunity in previously infected bitches is stage specific and directed against third stage larvae only. The time needed until eggs appear in the faeces after this kind of infection was shown to be 9–12 days (Brunschön-Hellmich, 1987). Another possibility would be a spurious infection, which is caused by the accidental ingestion of eggs shed in the faeces of the offspring, passing through the intestine of the bitch to reappear in her faeces. Additionally, a phenomenon called periparturient immunosuppression may occur (Lloyd et al., 1983). It is proven that pregnancy and lactation in dogs lead to a reduced immunological responsiveness. This is displayed by a suppressed in vitro transformation of the lymphocytes responding to phytomitogens and also a suppressed T. canis-induced lymphocyte transformation. Furthermore, eosinophilia commonly detected during a T. canis infection is repressed during the periparturient period (Lloyd et al., 1983). The immunosuppression might allow tissue-arrested larvae and larvae of newly acquired infections to perform tracheal migration and eventually intestinal development. The relevance of immunosuppression for the development of patent infections in bitches was confirmed by Lloyd et al. (1983), who were able to induce a patent T. canis infection after administering high doses of corticosteroids to a 6-month-old dog. The hypothesis of immunosuppression as a cause of patent infections was also supported by the finding of egg-producing adult worms in a bitch at a time post partum which was too early to be caused by ingestion of immature stages of the offspring (Sprent, 1961).

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2.5.1. The putative impact of hormonal changes in bitches Evans et al. (1991) observed that bitches in the luteal phase were three times more likely to have patent infections compared to other groups (pregnant, spayed, anoestrus, and previously injected with a progestagen). Nonetheless, this was a field study based on routine visits to veterinarians, with no definite knowledge of the background of the examined dogs. Regarding a possible relation between luteal phase and patent infections, Overgaauw et al. (1998) emphasise that after an infection during the luteal phase, eggs would appear within the faeces 4–5 weeks later due to the prepatency period and therefore would not be detected during the luteal phase. The group (Overgaauw et al., 1998) addressed the impact of prolactin on T. canis infections in a more controlled study. The prolactin levels of cyclic bitches, although increasing during the luteal phase, remained low compared to those of pregnant dogs. A higher risk for patent T. canis infections during the luteal phase was not found. Anyhow, patent infections were also not detected in the two pregnant bitches examined in that study. The authors raise the question whether pseudopregnant bitches, which show a high prolactin level, would also develop patent infections (Overgaauw et al., 1998). 3. Pathological changes in the host during larval migration

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Fig. 7. Adult ascarids penetrating a dog liver (Institute for Parasitology, archive).

some were found in the liver parenchyma and in the bile ducts. The common bile duct was dilated and thickened. A single ascarid was even found penetrating through the pancreatic duct. In the intestine several 100 adults were detected. This phenomenon occurs most often in pups after extensive prenatal infection. Moreover, a massive number of ascarids in the intestine of young dogs after prenatal infection can lead not only to the typical pressure sensitive, drum-shaped abdomen (Fig. 8) but also to developmental

3.1. Clinic and gross changes Larval migration leads to tissue damage in the host (Webster, 1958a). 24–72 h p.i. in some dogs bloodymucous enteritis might occur, caused by the penetration of the intestinal wall by the larvae (Fernando, 1968). The arrival of larvae in the lung can then be accompanied by coughing and dyspnoea (Overgaauw, 1997). The final intestinal colonisation usually does not cause more than a limited to moderate enteritis, but in cases of massive infection it can cause obstructions and ruptures of the intestine (Webster, 1958a). Hayden and Van Kruiningen (1975) compared a control group of dogs naturally infected with T. canis and a group additionally infected three times a week for 1 month with 3500 embryonated eggs each time, totalling 50,000 eggs. The latter group was considered superinfected. Lesions in the intestinal tract of dogs naturally infected with T. canis were minimal, whereas superinfected dogs had several white nodules in the jejunum and petechiae in the pyloric mucosa. A profound colonisation can also cause adult ascarids to invade the bile ducts, perforate liver parenchyma and finally enter the abdominal cavity through the liver capsule (Fig. 7). Naturally, this incident leads to a generalised and incurable peritonitis, which exacerbates if female ascarids in the peritoneal cavity continue to produce and release eggs (Dade and Williams, 1975). In the case report of Dade and Williams (1975) on a 5-week-old puppy with inappetence, depression, diarrhea and weakness, during necropsy 21 adult ascarids were found in the peritoneal cavity, 4 adults were apparently in the process of leaving the liver as they were protruding from the capsule, and

Fig. 8. Typical drum-shaped abdomen of a puppy infected with T. canis (Institute for Parasitology, archive).

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disorders such as cachexia, zero growth and also rachitic symptoms (Bosse et al., 1980; Herschel, 1981; Voßmann, 1985). Voßmann (1985) infected four bitches 10 days ante conceptionem with 20,000 embryonated eggs each. The fifth bitch remained uninfected and therefore its puppies served as a control group. Three of the infected bitches were treated with fenbendazole to reduce the prenatal infection, whereas one infected bitch was not treated and therefore highly infected. All puppies from the untreated bitch showed alternating obstipation and diarrhea, furthermore vomiting, anaemia and rachitis. These puppies subsequently died between the 22nd and the 49th days. Necropsy revealed ascites, cachexia, anaemia and dilatation and partial rupture of the proximal duodenum by a ball of ascarids. The cause of death was most often rupture of the intestinal wall and peritonitis after migration of adult ascarids through the bile duct and parenchyma of the liver. Puppies from treated bitches showed no typical signs of toxocarosis, but with 4 kg, compared to the 7 kg body weight of the uninfected control group at the end of the study, a considerably reduced development of body weight. Necropsy showed slight dilatation of the proximal duodenum, a macroscopically thicker stomach and intestinal wall and an activation of the lymphatic system (Voßmann, 1985). Among macroscopic lesions in other organs, red and white spots in the liver distributed over all lobes occur as well as petechiae and yellowish-white and red lesions in the lungs. Kidneys often show typical white lesions throughout the whole cortex (Hayden and Van Kruiningen, 1975; Manhardt, 1980; Herschel, 1981) (Fig. 9). 3.2. Haematological and enzymatic changes 3.2.1. Anaemia Voßmann (1985) examined haematological changes in puppies after prenatal infection and observed changes in the red blood cell counts. Red blood cell counts are physiologically low in newborn puppies and increase

approximately 6–8 weeks post partum to reach the values in adult dogs. In contrast, puppies heavily infected with T. canis showed further decreasing numbers of erythrocytes, which were mostly caused by severe internal bleeding. Reasons for the occurrence of internal bleeding were preadult larvae migrating through the liver as well as perforation of the intestine, caused by the massive load of adults. Moderately infected puppies showed an increase of erythrocyte counts from the 5th week onwards, but did not reach those values of uninfected animals. In contrast, in adult bitches, no changes in the red blood cell counts were observed after infection with T. canis larvae (Zimmermann, 1983). 3.2.2. Eosinophilia Also characteristic for T. canis infection is the eosinophilia which starts at least on the 7th day post infection, reaching its maximum approximately 14 days post infection (Zimmermann, 1983). A similar time course of eosinophilia is observed in prenatally infected puppies from day 7 post partum on (Voßmann 1985). Voßmann (1985) furthermore reported that the degree of eosinophilia in prenatally infected puppies is almost proportionate to the intensity of the infection. With the commencement of egg shedding in the faeces the number of eosinophils slowly decreased, returning to physiological levels 42 days p.i. These data showed that the course of the eosinophilia in prenatally infected pups is comparable to that in experimentally infected adult dogs (Zimmermann, 1983). 3.2.3. Increased liver enzyme levels During infection with T. canis not only changes in the blood counts are observed but also enzymatic changes. During the liver migration, the liver enzymes glutamate dehydrogenase (GLDH) and alanine transaminase (ALT) increased, reaching a maximum 14 days p.i. (Zimmermann, 1983). Afterwards, ALT remained on a high level for a certain time period, whereas GLDH returned to physiological values after 14 days (Stobernack, 1988). After reinfection, liver enzymes show an increase again even though it is lower than after the primary infection (Zimmermann, 1983). Voßmann (1985) observed these two enzymes in prenatally infected puppies to be elevated already at birth up to 67 U/l for GLDH (physiological value up to 6.0 U/l) and 365 U/l for ALT (physiologically up to 55 U/l) in highly infected puppies. The values returned to normal 1–2 weeks post partum. A second increase caused by adult ascarids migrating through the liver and the peritoneal cavity was detected shortly before the death of highly infected puppies. 3.3. Histopathological changes

Fig. 9. Typical white spots on the kidney (Institute for Parasitology, archive).

Webster (1958a) histologically examined the organs affected by the parasite. Within this study, 72 h p.i. in the liver parenchyma free larvae were detected as well as an infiltration of leucocytes. During the following 24–48 h the number of leucocytes increased continuously. Most leucocytes were identified as eosinophils, monocytes and also polymorphonuclear leucocytes. Webster (1958a) reported a tendency of these cells to gather in nodules,

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the host tissue. Why therefore should the immune system not be able to detect and to successfully eliminate these larvae? Two ways of immune evasion have been suggested. One hypothesis is that larvae reach a state of hypobiosis in the tissue, where expression of the immune system stimulating antigens is widely reduced. The other hypothesis suggests an immunosuppression induced by T. canis larvae which affects the function of T-helper cells. Consequently, the host response to parasite antigens and the production of specific antibodies is reduced (Overgaauw, 1997). 4.1. The antigenic coat

Fig. 10. Granuloma in the skeletal muscle (Institute for Parasitology, archive).

sometimes surrounding a larva in their centre. These structures could be referred to as granulomes. Based on these processes the tissue becomes necrotic, haemorrhages and marked fatty degeneration of liver cells occur. After 10 days first signs of encapsulation marked by a thin fibrous capsule were seen associated with the retreat of some of the leucocytes. Finally, about 3 weeks p.i., Webster (1958a) observed the beginning of a regenerative process with fibrous tissue proliferation. Excessive infiltration of lymphocytes, eosinophils and reticulum cells in the liver was also observed in the study of Hayden and Van Kruiningen (1975). Webster (1958a) further examined the lung and also found an infiltration with leucocytes, mainly eosinophils, forming compact nodules. Infected dogs suffered from lobular pneumonia and vascular congestion. Heavy infections, which are not further specified by Webster (1958a), led to serious pneumonia with exudate mainly comprising mucus, blood cells, epithelial cells as well as larvae in the alveoli and bronchioles. Furthermore, haemorrhages were common and in some cases extensive tissue degeneration was observed. Dense aggregates of eosinophils were also found in the intestinal wall, especially in the duodenum and jejunum but also in the cecum and colon. Even mesenteric lymph nodes had a few capsular and cortical granulomes (Hayden and Van Kruiningen, 1975). Furthermore, Hayden and Van Kruiningen (1975) observed mild to severe pyelitis in the kidney in 3 out of 7 dogs and interstitial infiltrates of lymphocytes, plasma cells, eosinophils and histocytes in the cortex of 2 dogs. Granulomes directly under the capsule of the kidney were also described by Herschel (1981). Therefore, it can be said that eosinophil infiltration and granulomes were most pronounced in the liver, kidney and lung (Hayden and Van Kruiningen, 1975). It is known that larvae in skeletal muscles of paratenic hosts are enclosed in granulomes (Parsons, 1987). Manhardt (1980) also made this observation in the definitive host (Fig. 10), the central nervous system being the only exception. 4. Evasion of the immune system As outlined above, larvae migrate for several weeks, and arrested larvae remain alive for at least several months in

Many in vitro studies have been conducted to further elucidate the mechanisms this parasite uses to evade the host’s immune system. As hatched larvae can be maintained in vitro and excretory–secretory (E/S) antigens can be collected, this stage is regarded as a model for studies on host–parasite-interactions (Maizels et al., 1987). Smith et al. (1981) found that larvae are not recognised by antibodies against E/S antigens at 37 ◦ C but are so when kept at 2 ◦ C or at 37 ◦ C after preincubation with certain antimetabolites. Both, low temperatures and antimetabolites, led to a metabolic inhibition, which was displayed by the ability of the antibodies to bind to the surface of the larvae. Furthermore, when the temperature was increased to 37 ◦ C or the antimetabolites were removed, binding of fluorescence-labelled antibodies was again not detected. Due to the fact that E/S antigens could be detected all over the surface of metabolically inhibited larvae but not on the surface of larvae kept at 37 ◦ C, it was concluded that the whole outer larval surface was involved in the release of E/S products (Smith et al., 1981). Smith et al. (1981) then hypothesised that the ability to repeatedly shed E/S products on the entire surface could enable the larvae to continuously remove antibodies attached to the surface. Therefore, antibody-dependent cell adhesion reactions of the host would effectively be prevented. This hypothesis was confirmed by several authors. Badley et al. (1987) found out that absorption of immune sera with E/S antigens removed surface IgG but also C3, which implied that E/S antigens have the ability to remove antibodies and serum components necessary for direct complement fixation. Maizels and Page (1990) found larvae to be bound by complements even up to C9. Furthermore, it was observed by using electron microscopy that the single layer of granular epicuticular material of larvae incubated with immune serum for at least 30 min, thickened and condensed and that the denser portion of the epicuticle frequently detached from the surface. Together with the sheath-like layer also eosinophils, containing vacuoles opening to the larval surface, were detached. Therefore, no gross damage to larvae could be seen after 30 min or after 12 h in culture with cells and serum (Badley et al., 1987). The ability to repeatedly shed attached antibodies and cells is maintained by high production and expression of antigens, which contribute to the surface layer, i.e. E/S antigens. Badley et al. (1987) reported that the E/S antigen production totals quantities of up to 8 ng per larvae and day. Maizels et al. (1984) reported that within 1 h 25% of the surface antigens were released, indicating the rapidity of this process.

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Maizels et al. (1984) further characterised the E/S components from T. canis (TES). Five antigenic components were identified and named after their molecular weights in kDa: TES-32, TES-55, TES-70, TES-120 and TES-400. All TES-molecules were found to contain more than 40% carbohydrates, the majority of which were Nacetylgalactosamine and galactose (Meghji and Maizels, 1986). All TES molecules were first secreted and then formed the larval surface coat except for TES-400. Page et al. (1992) examined the organs which might be connected to the secretion of the major glycoproteins of T. canis, using immunogold electron microscopy. It was found that they are derived from two specialised organs within the nematode organism. The two primary locations were found to be the esophageal glands, which secrete their products via the esophagus in the oral orifice, as well as a large branched secretory cell, which has its opening to the cuticle at an anterior secretory pore. Additionally, it was found that TES-32 is synthesised separately in the cuticle (Page et al., 1992). 4.2. The interaction of TES molecules with the immune system Loukas et al. (1999) found that TES-32 is a C-type lectin having greater similarity to mammalian C-type lectins than to the respective orthologues in C. elegans. As the mammalian C-type lectins are involved in the immune response, the authors supposed an immune evasive strategy of the parasite based on the similarity to host immune cell receptors. Subsequently, Loukas et al. (2000) showed that binding of TES-70 and other TES to host proteins from a canine cell line occurs in a calcium-dependent manner. The interaction of T. canis E/S antigens with the immune system was further documented in a recent study by Giacomin et al. (2008a). Injected E/S antigens enhanced the migration of Nippostrongylus cantonensis in IL-5 transgenic mice, which have an innate resistance to N. cantonensis. The authors confirmed the activation and consumption of the complement factor C3 reported by Badley et al. (1987). Additionally, they surmise a complement-independent mechanism modulating the function of eosinophils, as the resistance of the mice is not based on the complement system (Giacomin et al., 2008b). However, the exact mechanism of the interactions of TES and eosinophils and other immune cells remains unclear. 5. Conclusions Holland and Smith (2006) referred to Toxocara as “The Enigmatic Parasite”. There is no better way to precisely characterise this fascinating nematode, and the present state of knowledge even after decades of intense research, still leaves many questions unanswered. Our present, more stringent animal welfare considerations certainly exclude any new attempts to solve the remaining mysteries of larval development inside the host and help us to accept a certain level of incomplete knowledge. More important however is that despite a large number of highly efficient anthelmintics, this zoonotic parasite still represents the most prevalent nematode in dogs and that we understand

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