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Veterinary Parasitology 175 (2011) 193206

Contents lists available atScienceDirect

Veterinary Parasitology

j o u r n a l h o m e p a g e : w w w . e l s e v i e r . c o m / l o c a t e / v e t p a r

Review

Larval development ofToxocara canisin 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 i n f o

Article history:

Received 25 June 2010Received in revised form 7 October 2010Accepted 12 October 2010

Keywords:

Toxocara canis

ToxocarosisInfection routesLarval developmentPrenatal infectionZoonosis

a b s t r a c t

The parasitic roundworm Toxocaracanis ispresentindogpopulationsallovertheworld.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 infectionroutes and the subsequent development of larvae within the canine host is summarised.Furthermore, information about the clinical, pathological, enzymatic, haematological andhistopathological changes was collected, giving a broad overview of current knowledgeof the infection. Although the data collected over the years give an idea of what happensduring the larval development ofT. canis, many questions remain open. Nevertheless, it isimportant that we continue our efforts to further understand the biology of this versatileand compelling parasite and try to improve and optimise strategiesto prevent the infectionin dogs and thereby to protect humans from this infection.

2010 Elsevier B.V. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1942. Larval development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194

2.1. Oral infection withT. canis eggs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1942.1.1. The infective stage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1942.1.2. The first part of the migration route . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1952.1.3. The impact of age and acquired immunity on larval migration route. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1952.1.4. The impact of gender on prevalence of patent infections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1952.1.5. The influence of the infection dose on the course of infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1962.1.6. Development into adults . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1962.1.7. Somatic migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196

2.2. Oral infection through paratenic hosts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1972.3. Prenatal infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198

2.3.1. Reactivation of larvae during pregnancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1992.3.2. Migration and development of larvae in the puppies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 199

2.4. Lactogenic infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1992.4.1. Distribution of larvae in the mammary glands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 200

2.5. Oral infection with juvenile intestinal larvae and periparturient immunosuppression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2002.5.1. The putative impact of hormonal changes in bitches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201

3. Pathological changes in the host during larval migration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2013.1. Clinic and gross changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201

Correspondingauthor. Tel.: +49511 9538711; fax: +49511 9538870.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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194 T. Schnieder et al. / Veterinary Parasitology 175 (2011) 193206

3.2. Haematological and enzymatic changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2023.2.1. Anaemia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2023.2.2. Eosinophilia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2023.2.3. Increased liver enzyme levels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 202

3.3. Histopathological changes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2024. Evasion of the immune system . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 203

4.1. The antigenic coat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2034.2. The interaction of TES molecules with the immune system . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204

5. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204R e f e r e n c e s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 204

1. Introduction

Toxocara canis (Werner, 1782) is one of the most impor-tant gastrointestinal helminths in dogs, with infectionsreported from all parts of the world. In Argentina, 16.35%of 1944 fresh dog faecal samples collected from both urbanand rural areas were positive for T. canis (Soriano et al.,2010).A study in Nigeria in 2007 reported 33.8% of 269household dogs to be positive, with a prevalence of 51.4%in puppies up to 6 months (Sowemimo, 2007). AnotherNigerian study determined the overall prevalence in 396dogs to be 41%, with younger dogs being more suscepti-ble to the infection (Ugbomoiko et al., 2008).A similarlyhigh infection rate (45.2%) was reported for 438 slaugh-tered dogs in abattoirs in a Chinese province, determinedby counting the adult worms (Dai et al., 2009). Severalstudies confirm that T. canis infections are also commonin the European canine population. A study performed inItaly diagnosed 33.6% out of 295 dogs going to veterinaryclinics for whatever reason as T. canispositive (Habluetzelet al., 2003). In Hannover, Germany, 2.2% of the faecalsamples from 1281 dogs examined as part of diagnosticservices from 1998 to 2002 were positive for eggs of thisparasite (Epe et al., 2004).A similar prevalence (2.1%) wasdetermined for the years 20032009 (unpublished data).In Serbia, Nikolic et al. (2008) examined stool samplesfrom 151 household, stray, and military dogs, revealing aprevalence as high as 30.5%, whereas 1159 faecal samplestested in Northern Belgium showed a T. canisprevalenceof 4.4% in household dogs (Claerebout et al., 2009).Thesame study identified a prevalence of 26.3% in breedingkennel dogs. In the UK, Batchelor et al. (2008) included4526 dogs with clinical signs of gastrointestinal disease intheir prevalence study, with 1.4% being positive. In a recentGerman study, out of 445 dogs taken in by animal shel-ters 2.5% of adult dogs were positive forT.canis,whereasdogs up to 1 year of age were positive at a rate of 8.8%(Rohen, 2009). In the Netherlands, a coproscopic preva-lenceof4.4%ofT. canis eggs was determined in 92 clinicallyhealthydogs(Overgaauw et al., 2009). However, the preva-lences determined in the cited studies cannot be compareddirectly to each other as the examined populations werequite different. Nevertheless, T. canis is undeniably presentin the canine population worldwide, especially in youngdogs. Working dogs and stray dogs seem to be at a higherrisk of getting infected than pet dogs (Nikolic et al., 2008).Nonetheless, not only do excreted eggs in dog faeces andcontaminating soil provide a human infection risk, but alsodo eggs attached to dog hair. Wolfe and Wright (2003)

examined the hair of 60 dogs in Ireland and the UK, withthe result thatT. caniseggs 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 exam-ined the hair of 100 stray dogs in Ireland with the resultthat 67% of the dogs were found to have T. canis eggson their hair (Roddie et al., 2008b).These authors exam-ined not only dog hair for T. canis eggs but also fox hair,revealing 28% of 87 foxes to be positive. 61.4% of theseeggs were either viable or embryonating (Roddie et al.,2008a). In Turkey, Aydenizz-Ozkayhan et al. (2008) foundT. caniseggs in the hair of 21.56% of 51 dogs. All the eggswere viable, 21% being either embryonating or embry-onated.Overgaauw et al. (2009) detected T. canis eggs in12.2% of 148 fur samples of clinically healthy dogs. In thisstudy, none of the eggs were viable. The actual impactofT. canis eggs within dog fur on human health remainsto be determined, but as pets literally share house andbed with their owners, this source of infection might beunderestimated. The wide distribution ofT. canis and itsimportance as a zoonotic agent, causing visceral and ocularlarva migransin humans, have led to an increasing inter-est in the biology and epidemiology of this parasite. Forefficient prevention of infections in dogs, understandingits complex life cycle, including infections resulting fromingestion of either embryonated eggs from the environ-ment or larvae from infected paratenic hosts, as well asinfections from vertical transmission (intrauterine or lac-togenic), is crucial. However, there are still unansweredquestions concerning the life cycle of this nematode. Inthis review, current knowledge regarding larval develop-ment in dogs, mechanisms of larval reactivation from thetissue, immune evasion and pathological changes in organsafter invasion will be outlined and discussed. As some ofthe previous studies were published in German only, theirinclusion 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 withthe faeces. Under optimal conditions, i.e. temperaturesbetween 25 and 30 C and relative humidity of 8595%,the development of the infective larval stage within theegg requires 915 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 upto several months, and the infective stage in the eggcan remain viable for at least 1 year (Overgaauw, 1997).Thereby, the infective larval stage is not, as was believedfor 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 duode-num within 24h. The released, infective larvae penetratethe mucosa of the intestine (Webster, 1958b). The pen-etration mechanism is as yet not fully understood.However, an elastase-like protease was detected in larvalexcretorysecretory (E/S)products (Robertsonet al., 1989).This protease might assist larvae in penetrating host tis-sues, i.e. the intestinal mucosa, liver or kidney parenchymaand the walls of blood vessels. Furthermore, directmechanical disruption is discussed (Parsons, 1987).Afterpenetrating the intestinal wall, larvae invade lymph ves-sels and migrate to the mesenteric lymph nodes (Webster,1958b). Thereafter, they travel in venous capillaries via theportal circulation to the liver (Webster, 1958a). The major-ity of the larvae reach the liver approximately 24 h postinfection (p.i.) (Webster, 1958a). Subsequently, within 12 hmost larvae continue to migrate, exiting the liver via Venacava, passing the heart and reaching the lung via the pul-monary artery. The larvae occur there between24 and 36hp.i., and numbers increase up to 96h (Webster, 1958a).Webster (1958b) demonstrated the passive haematoge-nous transport by detecting T. canis larvae in the bloodof the heart as well as in the pulmonary artery 72 h postinfection. Those larvae which are trapped in capillaries andcannot continue the migration remain in the liver. Afterencapsulation, these larvae cause the mottled appearanceand 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 andthe infection dose. Firstly, they can penetrate the alveoliwall and continue their migration via bronchioles and tra-chea to the pharynx. Here they are swallowed and finallygrow into adult worms in the intestine. On the secondroute, larvae again penetrate the alveoli wall and re-enterthe circulatory system, being distributed to the somatic tis-sue (Webster, 1958b). Greve (1971) compared 3-week-,3-month- and 1-year-old dogs from an ascarid-navecolony after experimental subcutaneous infection with T.canis. In 3-week-old puppies almostall larvaehad migratedthrough the tracheal migration route. In contrast, in 3-month- and 1-year-old dogs most larvae were discoveredin granulomes in the tissue. Therefore, it was concludedthat the likelihood of somatic migration progressivelyincreases from the age of 3 months onwards. Concurrently,the development of larvae into adult ascarids decreases(Greve, 1971; Oshima, 1976). This phenomenon is referredto as age resistance, which is firstly based on the devel-opment of immune competence and secondly on acquired

immunity (Barriga, 1988). Immunity is assumed to occuron the one hand in the lung as a putative delayed typehypersensitivity response (Dubey, 1978),affecting third-stage larvae after reinfection (Oshima, 1976).On the otherhand, immunity is also located in the gastro-intestinaltract, preventing the infective larval stage from penetrat-ing the intestinal mucosa after reinfection. The restrictedpenetration might be ascribed to an inflammatory allergicreaction after the release of vasoactive amines by sensi-bilised mastocytes (Soh and Kim, 1973),this possibly alsobeing the explanation for the catarrhal-haemorrhagic diar-rhea described in reinfected dogs (Lwenstein, 1981).Thisstatement is supported by elevated gamma globulin val-ues in the exudate and tissue of the intestine (Soh andKim, 1973). The degree of inhibition was examined byLwenstein (1981)who discovered that after reinfectionof lactating bitches with embryonated eggs, the distribu-tion of larvae in the somatic tissue and the eliminationin the milk were 99% lower than in the group infectedonly once.Zimmermann (1983)determined that a trickleprimary infection during 10 days resulted in a 28% lowerdistribution of larvae compared to a single infection withthe same total number of embryonated eggs, thus demon-strating the rapid development of the immune response.

Additionally, studies showed that acquired immunityis stage specific against third stage larvae only. Juvenileintestinal larvae (L4) introduced directly into the intestineof previously immunised dogs were able to grow into adultworms at a rate of 4660%, whereas infective third stagelarvae implanted in a similar manner did not develop intoadults. Furthermore, no antibody response was detectedafter implantation of fourth stage larvae (Fernando et al.,1973). These results were confirmed by a study show-ing that implantation of juvenile intestinal larval stagesin five 1- to 2-year-old male dogs, produced patent infec-tions in all dogs. The same was achieved after a first anda second retransplantation (Brunschn-Hellmich, 1987).Furthermore, the immune system is unable to completelyeliminatetissue-arrestedparasites. Possible reasons willbediscussed later on.

2.1.4. The impact of gender on prevalence of patent

infections

Some studies suggest that the migration route of theparasite is influenced not only by age and immunitybut also by gender. It has been observed that patentT. canis infections occur more frequently in adult maledogs (older than 12 months old) than in females of thesame age (Ehrenford, 1957; Turner and Pegg, 1977).Thiswas discussed to be part of a survival strategy in termsof evolution, as females spread the infection via reacti-vated somatic tissue larvae to their offspring, whereasToxocara infections in males can only be distributed bypatent intestinal infections (Overgaauw, 1997). Webster(1958a) also reported that more larvae migrate in thesomatic tissues of female dogs compared to those of maledogs. In a survey with 1324 dogs,Ehrenford (1957)found32.8% of the males in contrast to only 9.4% of the females tobe infected with adultT. canis. Furthermore, it was statedthat in male dogs there was no evidence of immunityup 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 wasdetected in 3 adult male greyhounds, which were repeat-edly infected with 100200 viable eggs and became patenteven though serum antibodies to the parasite surfaceas well as to secreted antigens were detected (Maizelsand Meghji, 1984).Maizels and Meghji (1984) discussedas possible reasons firstly a greater susceptibility of thisbreed to certain parasite infections, secondly the genderand thirdly the infection dose. In contrast, in another,more recent, study, female dogs were reported to have ahigher prevalence (Bridger and Whitney, 2009).However,the authors themselves acknowledge that the sample sizeof 28 dogs with known sex was low. Thus, the results maynot be comparable to other studies. However, whether thegender really influences the migration and developmentofT. canisis doubtful, since most prevalence studies havenot 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 infectionwith different numbers of infective eggs, it is obvi-ous that the antigenic stimulus has an impact on thedevelopment of protective immunity. Dubey (1978)experimentally infected 45 ascarid-nave 2-month-old puppies and 6 adult dogs. In 24 out of 25 pupsinfected with 101000 embryonated eggs a patentinfection was established. In contrast, no patentinfection was detectable in 20 puppies infected with10,000 eggs. Furthermore, three of the six adult dogs (7,10 and 52 months old) developed patent infections afterbeing fed with 100 embryonated eggs each (Dubey, 1978).Similarly, a study on 18 beagles confirmed the results ofMaizels and Meghji (1984) thatlow infection doses, i.e. 100embryonated eggs, cause patent infections also in olderdogs(Fahrionetal.,2008). Therefore, adult dogsconsideredimmune againstT. canisinfections were shown to developpatent infections after reinfection if the number of infec-tive larvae and thus the antigen stimulus is low. Thus, itcan be assumed that the number of adult dogs with patentinfections and the contamination of the environment witheggs might be higher than so far assumed.

2.1.6. Development into adults

As mentioned above, the lung is the crucial organ inwhich parting of the ways occurs: larvae migrate eithertowards the intestine to establish a patent infection ortowards somatic tissue to become a larva migrans. To com-plete their way to a patent infection, larvae penetrate thealveoli and reach, after passing the bronchioles, the tra-chea. After 79 days larvae can be detected in the tracheaand in the esophagus. The gastrointestinal tract is reached715 days after infection (Sprent, 1958).From early stud-ies it is hard to determine where larval moults actuallyoccur, because these authors considered the infective lar-vae to be second stage larvae and described a total ofthree moults. More recent knowledge, however, provesthat infectivelarvae in theegg arealready third stage larvae

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

and only two moults occur inside the host (Araujo, 1972;Brunaska et al., 1995).According toWebster (1958a)onemoultoccurseitherinthelungsandtracheaorintheesoph-agus, the next in the stomach, and a further one in the

duodenum.Schacher (1957)also reported about findingsof third and fourth stage larvae in the stomach and aboutfourth and fifth stage larvae in the duodenum. Althoughconclusive studies are missing, it may be assumed that thefirst moult to L4 happens somewhere between entering thebronchioles and passing the stomach.

Without any doubt the last moult to the preadult stage(L5) takes place in the small intestine. When preadultshave matured, first eggs are shed approximately 45 weeksp.i. in experimentally infected puppies (Webster, 1958a),whereas in older hosts prepatency seems to be extended to40upto56daysp.i.(Fahrion et al., 2008). It is believed thateach adult ascarid lives for 4 months on average (Parsons,

1987).Experimentally infected dogs are reported to expel53% of their adult worm burden within a period of 8 days,beginning 1 week after the onset of patency (Burke andRoberson, 1979).

2.1.7. Somatic migration

As puppies mature, the disposition for tracheal migra-tion and development of a patent infection falls to a lowlevel. Instead, most larvae re-enter the capillary systemto finally perform somatic migration. Many of the earlierstudies lack information about whether trulyascarid-naveanimals have been used. To gain a better understanding ofthe way larvae travel in the lung and thereafter in adult, sofar ascarid-nave dogs,Manhardt (1980)injected 200,000infective larvae into theVena cephalica antebrachiiof 1- to3-year-old dogs. After 1 h p.i., 75% of the administered lar-vae were recovered from the lung, whereas no larvae couldbe detected in the arterial blood. In histological slides lar-vae could be detected moving inside the lung vessels aswell as leaving them (Fig. 1). Furthermore, they were foundmigratingfreelyin the interstitium andfinally reaching thelung alveoli (Fig. 2).

Somatic migration in the lung leads to many petechialhaemorrhages, giving the lung a stippled appearance(Webster, 1958a; Manhardt, 1980). Manhardts (1980)findings confirmed Oshimas (1976) assumption thatsomatic migration is performed by larvae which are

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Fig. 2. T. canislarva in the lung, interstitial pneumonia (Institute for Par-asitology, archive).

trapped in capillaries, causing them to penetrate the wallsandmigrate through the tissueto re-enter thevascularsys-tem. This somatic migration ability leads to the appearanceof larvae in organs close to the lung and in the pleuralcavity. Furthermore,Manhardt (1980)examined the kid-ney more closely, an organ commonly highly affected byT. canis larvae, but rarely showing serious organ failure.Larvae leave the blood vessels within the cortex, leavingsmall haemorrhages under the capsule of the kidney, andstart a somatic migration(Fig.3). Somelarvaepenetratetheurine canaliculi, being detectable up to the renal medullaand even as far as in the urine itself, whereas most larvaeremainafter a short somatic migrationclose under the cap-sule where they are encapsulated in granulomes (Fig. 4).Shortly after the start of the haematogenous transport, lar-vae were also discovered between the fibrillae of the heartmuscle (Manhardt, 1980).

The time period in which most larvae were distributedwas determined as 2472 h (Manhardt, 1980). At the sametime, the number of larvae in organs other than the lungsincreased correspondingly, as shown inTable 1.A surpris-ing phenomenon was observed for the numbers countedafter 48 and 72 h in the front legs, which were higher thanin the hind legs at the same time. However, this differencewas no longer present at day 28 p.i.

Regarding the distribution ofT. canislarvae in the bodyof infected dogs, the vast majority of migrating larvae end

Fig. 3. T. canislarvae close to glomerula in the kidney capsule (Institutefor Parasitology, archive).

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

up in theskeletalmusclesand thekidneys,but also theliverand 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 embry-onated eggs. Based on the number of recovered larvaefromtissue samples, she calculated that 78.3% of the larvae werecontained withinthe skeletal muscles and20.1% withinthekidneys. In dogs the brain is in general only affected bylow numbers of migrating larvae. After digestion of half ofthe brain of one 3-month-old dog, Greve (1971) found only7 larvae, whereas no larvae could be recovered in the 1-year-old and the other 3-month-old dogs. This contraststo the situation in paratenic hosts, where the brain and theeyes, directly connected viathe Nervus opticus, are severelyaffected.

2.2. Oral infection through paratenic hosts

Besides direct oral ingestion of eggs, dogs can also beinfected after ingestion of paratenic hosts which carryT. canis larvae in their tissue. Paratenic hosts can be, amongothers, birds (Okoshi and Usui, 1968),rodents, e.g. mice(Sprent, 1953), but also rabbits (Sprent, 1955), pigs(Sasmaletal.,2008), foxes(SaeedandKapel,2006), andhumans(Itoet al., 1986).

Paratenic hosts become infected by ingestion of embry-onated eggs. There are reports of larvae staying alive for atleast 2 years in the tissue of mice, rats, guinea pigs and rab-bits (Beaver, 1956)and surviving several weeks in frozencarcasses of mice (Sprent, 1953).In chickens and pigeonsthey were found to survive for at least 3 months (Beaver,1956).

A number of studies showed that Toxocara infectionsclearly induce behavioural changes in mice as a model forparatenic hosts. Infected mice are less active and explo-rative and less aversive to open areas. As a result they areat a higher risk of being caught by predators than unin-fected 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 infectionsdeveloped directly without tracheal migration. The mostlikely 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. canislarvae into the V. cephalica antebrachii(Manhardt, 1980).

Hours/days p.i.

1 h 12 h 24 h 48 h 72 h 28 days

Number of larvae in:

Lunga 175,637 135,860 96,531 47,058 6426 790

Diaphragm

a

0 1 7 353 86 312Pleural cavity 0 0 22 20 0 1Arterial blood 0 0 4 15 1 0

Skeletal musclesa,b 0 0 212 10,965 16,575 45,390Kidneysa 0 0 422 10,348 12,100 18,136Livera 0 0 62 2862 14,157 5943Hearta 3 0 55 1944 1,077 2196Braina 0 0 0 95 30 26Peritoneal cavity 0 0 0 3 0 0Tracheal mucus 0 1 9 15 14 0Urine 0 0 0 2 0 0

Total number of larvae 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 thephysiological organ weights of a beagle with a body weight of 15 kg.

b Themusclesample (200 g) consisted of 50g each of thefollowingmuscles: M.biceps dexter, M. biceps sinister, M. semitendinosus dexter, M. semitendinosussinister.

in the tissue of the previous host. Therefore, they hadalready reached a stage of maturity sufficient for directdevelopment(Overgaauw,1997). Warren(1969), however,detected larvae also in the liver, lung, trachea, and esoph-agus, therefore concluding that larvae released from micetissue also undergo tracheal migration in the dog. Herschel(1981) fed infected mice to beagles and 72h after ingestionlarvae were detected in the liver and lungs, in the arterialblood as well as in other peripheral organs of one dog. Afterfeeding infected mice to dogs, a normal prepatent periodof 3448 days was observed (Herschel, 1981).

2.3. Prenatal infection

The most important way ofT. canis infection in dogsis prenatal transmission, also known as transplacental orintrauterine transmission. There are several reports stat-ing that puppies post partum are commonly infected withT. canis. This infection can be induced by feeding embry-onated eggs (Shillinger and Cram, 1923) or subcutaneouslyinjecting Belaskaris-larvae (i.e. T. canis larvae) (Flleborn,1921)to pregnant bitches. Nonetheless, parenteral infec-tion of puppies occurs not only after the infection of thepregnant bitch but also through reactivation of somatic tis-sue larvae from earlier infections (Yutuc, 1949; Webster,1958b; Koutz et al., 1966). It is not known for sure yethow long larvae can remain and survive in the tissue ofdogs. However, in a study on mice, larvae could be recov-ered from the tissue up to 1 year post infection (Bardnet al., 1994). In two abstract-like publications, Douglasand Baker (1959a,b)refer to a study finding that 241 and358 days after an infection of the bitch, prenatal infec-tions still occurred. Furthermore, it was observed that abitch harbouring somatic larvae can infect puppies duringthree consecutive pregnancies (Soulsby, 1983). Koutz et al.(1966) also observed prenatal infections in foetuses from abitch which had not been experimentally infected but wassupposed to harbour somatic larvae from a previous nat-

ural infection. They further stated that the infection of thebitch has to occur until a short period post conceptionemto lead to prenatal infection of the foetuses. They came tothis conclusion as they were not able to detect larvae in theprogeny of bitches infected during1121 days post concep-tionem, but indeed gave no further particulars about thesedata. In contrast,the group ofBosse et al. (1980) wasabletoinduce prenatal infections in puppies at least until 7 daysante partum.

In summary, this transmission mode is of major impor-tance for puppies and may occur after reactivation ofsomatic larvae from previous infections of the bitch as wellas from new infections during pregnancy.

After infecting the bitch, larvae can either migratedirectly to the offspring or remain in the somatic tissuedepending on the time point of infection. Douglas andBaker (1959a,b)stated in two short notes that larvae donot reach the foetuses before the 42nd day of pregnancy.This was confirmed byScothorn et al. (1965),who couldnot find larvae in foetuses of a bitch on the 35th day ofpregnancy, but in 5 out of 6 foetuses of another bitch onthe 56th day of pregnancy. Both bitches were not infectedexperimentally but assumed to be infected naturally priorto the start of the experiments. In a further study, whichaimed to determine the time point of larvae appearing inthe foetuses more exactly, the first larva was detected inone of six foetuses of an experimentally infected bitch onthe 43rd day of pregnancy (Koutz et al., 1966).From the47th day onwards, all foetuses of infected bitches werepositive. No data are available about the time point whensomatic larvae are reactivated and actually start to migrateto thefoetuses. Regardingthe wayof transmissionfrom thebitch to the foetus,Webster (1958a)suggested that larvaereach theplacenta viathe circulatory systemand penetratethe delicate layer of tissue which separates the maternaland foetal blood. In the study by Scothorn et al. (1965),which is at least partially included in the analyses ofKoutzet al. (1966),two larvae were found in the umbilical cord

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tissue. Therefore,the umbilicalcord is regarded as a centraltransfer point between bitch and foetus in utero.

2.3.1. Reactivation of larvae during pregnancy

The mechanism how larvae are reactivated in thesomatic tissue of the bitch is still a matter of debate.However, Webster (1958a) alreadysupposed that this phe-nomenon is connected to the changing hormone status.

In mice, which can also transmit T. canis infectionsvertically, a clear decrease in tissue-arrested larvae afterapplication of prolactin was demonstrated (Oshima, 1961).These results were confirmed in a recent study by Jinet al. (2008),who observed that migration of larvae to themammary gland was triggered by prolactin. These find-ings suggest that a similar path for reactivation mightalso apply for dogs, although the transmission in micemainly occurs lactogenically (Baumm, 1980) rather thanprenatally. Regarding another also mainly lactogenicallytransmittedparasite,Ancylostoma caninum,astudyshowedthat after administration of estrogen and progesterone toa lactating bitch more larvae were shed in the milk thanbeforehand (Stoye, 1973; Krause, 1975).A study on reacti-vation of tissue-arrested third stage larvae ofA. caninumrevealed that neither estrogen nor progesterone had adirect effect on feeding/reactivation of these larvae. Pro-lactin showed a stimulatory effect only at concentrationsapproximately 1000 higher than physiological levels.However, the two isoforms(1 and 2) of TGF- led to signifi-cant stimulatory effects onA. caninum larvae (Arasu, 2001).Therefore,Arasu (2001)concluded that during pregnancyhost-derived TGF- could act on a parasite-encoded recep-tor which then causes the reactivation of somatic tissuelarvae. It had previously been shown that there is a strongcorrelation between estrogen and the expressionof TGF-2in rats (Schneider et al., 1996).A recent study confirmedthat TGF- also induces the reactivation of dauer larvalstages ofCaenorhabditis elegans (Gallo and Riddle, 2009).As C. elegans andA. caninum are allocated to the same cladeof nematodes, which is determined by the relatedness oftheir 18S RNA sequences, these results indicate that thereactivation of dauer stages might be preserved at leastbetween nematodes of clade V. Whether the same mecha-nisms induce reactivation ofT. canis, a clade III nematode,remains to be investigated.

2.3.2. Migration and development of larvae in the

puppies

Afterreactivationof tissue-arrestedlarvae, theymigrateto 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 lar-vae in organs other than the liver or lungs, namely inmuscles, the brain and kidneys. While he explains thepresence of larvae in the lung by the already startedmigration, he interprets the larvae in other organs assomatic larvae after migration or distribution, as describedin adult dogs. Post partum migration from the liver con-tinues immediately. Larvae were already detectable inthe lungs 30min post partum (Koutz et al., 1966). Assoon as larvae reach the lung they undergo a trachealmigration.Koutz et al. (1966) noticed a variable migra-

tory behaviour with some larvae reaching the intestinealready 2 days post partum and others remaining 2.5days in the liver before starting migration. Nevertheless,within 3 days the majority of the larvae had reached thelung. In all cases, larvae had reached the intestine by the7th day post partum. There are different reports concern-ing the duration of prepatency after prenatal infection:2130 days were described by Yutuc (1949) and 2546days byDouglas and Baker (1959a,b). Vomann (1985),however, found a prepatent period of 28 days. Subse-quently, more than 70% of the adult worm burden ofpuppies was expelled spontaneously 910 weeks post par-tum (Vomann, 1985).

2.4. Lactogenic infection

The lactogenic transmission is of rather subordinateimportance for the distribution of T. canis. Burke andRoberson (1985a,b) compared the number of prenatallyand lactogenically transmitted infections. They experi-mentally infected bitches 24 months before pregnancy.Immediately post partum and before nursing, the pup-pies were exchanged with newborn puppies of uninfectedbitches. After 4 weeks of nursing, the puppies werenecropsied 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 andonly 1.5% lactogenically (Burke and Roberson, 1985a). Afterinfection 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 wereinfected 48h post partum, a time point which excludes thepossibility of prenatal infection, 7.9% of the infection dosecould be recovered from the puppies.

Similar experiments were performed byStoye (1976),although he determined the number of larvae in the milkinstead of examining the puppies. After infection of a bitchon the day of mating, he recovered 8.5% larvae of the initialinfectiondoseintotal,99.2%ofwhichhadbeentransmittedprenatally to the puppies and only 0.8% having been foundin the bitchs milk. Stoye (1976) also infected a bitch on theday of birth, and recovered 9.5% of the infection dose fromthe milk between the 4th and the 28th day post partum.This was confirmed byLwenstein (1981),who infectedlactating 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 mostimportant infection route for puppies. However, if a bitchbecomes infected post partum, the lactogenic transmis-sion is another possibility for the parasite to infect newhosts. Nevertheless, little is known about the developmentof larvae in puppies after ingestion of the milk. With a longprepatent period of 2735 days after ingestion of larvaewith the milk (Bosse et al., 1980),a direct intestinal devel-opment seems to be unlikely.

The course of larval shedding with the milk is widelyconsistent. First larvae can already be detected within thefirst days post partum. The number of larvae increasesthen consistently until a maximum is reached approxi-

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

mately 714 days post partum. During the first 23 weekspost partum most infections of puppies occur. However,after high infections shortly before or after birth, larvaeappear in the milk approximately 47 days after infec-tion and can be found in the milk at least up to day 28after infection (Stoye, 1976; Manhardt, 1980; Lwenstein,1981).

2.4.1. Distribution of larvae in the mammary glands

To further examine thecourse of shedding with themilkand the distribution of larvae in the mammary gland, onthe day of birthManhardt (1980)infected bitches orallywith embryonated eggs or either intravenously, subcuta-neously or intra-peritoneally with larvae. Afterwards, eachmammary gland complex was examined at different timepointsupto28daysp.i.Inallexperiments,exceptaftersub-cutaneous infection, shedding of larvae in the milk started4 days p.i. However, after subcutaneous infection of larvaeinto the flank close to the mammary gland, larvae reachedthe milk and tissue already 1 day p.i., but only in the mam-mary gland complex nearby. 1, 2, and 3 days later larvaehad also reached the neighbouring complexes but neverreached all of them. Therefore, Manhardt (1980) calculatedthe speed of migration of larvae through tissues as 15cmin 24 h. Hence, she concluded that somatic migration ofT. canis larvae directly from the intestine via the peritonealcavityand abdominalwallmight be conceivableto a certainextent.

Histological examination of the mammary gland 8days after oral infection with 100,000 embryonated eggsshowed that most larvae (565) were present, more or lesscurled in the alveoli of the gland (Fig. 5), whereas onlyfew were detected in the mammary ducts (2), in the inter-stitium (3), and in the peri-glandular conjunctive tissue(4) (Manhardt, 1980).Larvae in the mammary ducts werestretched and in the interstitium either stretched or ina meander shape. Larvae in the peri-glandular conjunc-tive tissue were already being encapsulated in granulomes(Fig.6). Larvaewerefoundinallpartsofthegland,mostlyinthe alveoli and no larvae were detected in a vessel. Thus, itwas not possible to clarify whether larvae migrated mainlyhaematogenously or somatically.

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

2.5. Oral infection with juvenile intestinal larvae and

periparturient immunosuppression

In the periparturient period bitches can suffer frompatent T. canis infections. This may be due to differentreasons. One possibility is that bitches can be infectedby juvenile intestinal larvae (L4) shed with the faecesof the infected puppies (Sprent, 1961). While nursingand cleaning their puppies, bitches ingest these larvae,which directly develop into adult worms in the intes-tine (Lloyd et al., 1983). This is possible because, asmentioned beforehand, immunity in previously infectedbitches is stage specific and directed against third stagelarvae only. The time needed until eggs appear in the fae-ces after this kind of infection was shown to be 912 days(Brunschn-Hellmich, 1987). Another possibility would bea spurious infection, which is caused by the accidentalingestion of eggs shed in the faeces of the offspring, pass-ing through the intestine of the bitch to reappear in herfaeces.

Additionally, a phenomenon called periparturientimmunosuppression may occur (Lloyd et al., 1983). Itis proven that pregnancy and lactation in dogs lead toa reduced immunological responsiveness. This is dis-played by a suppressed in vitro transformation of thelymphocytes responding to phytomitogens and also asuppressed T. canis-induced lymphocyte transformation.Furthermore, eosinophilia commonly detected during aT. canis infection is repressed during the periparturientperiod (Lloyd et al., 1983).The immunosuppression mightallow tissue-arrested larvae and larvae of newly acquiredinfections to perform tracheal migration and eventuallyintestinal development. The relevance of immunosup-pression for the development of patent infections inbitches was confirmed byLloyd et al. (1983),who wereable to induce a patent T. canis infection after admin-istering high doses of corticosteroids to a 6-month-olddog. The hypothesis of immunosuppression as a causeof patent infections was also supported by the find-ing of egg-producing adult worms in a bitch at a timepost partum which was too early to be caused byingestion 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 lutealphase were three times more likely to have patentinfections compared to other groups (pregnant, spayed,anoestrus, and previously injected with a progestagen).Nonetheless, this was a field study based on routine visitsto veterinarians, with no definite knowledge of the back-ground of the examined dogs. Regardinga possible relationbetween luteal phase and patent infections, Overgaauwet al. (1998)emphasise that after an infection during theluteal phase, eggs would appear within the faeces 45weeks later due to the prepatency period and thereforewould not be detected during the luteal phase. The group(Overgaauw et al., 1998) addressed the impact of prolactinon T. canis infections in a more controlled study. Theprolactin levels of cyclic bitches, although increasingduring the luteal phase, remained low compared to thoseof pregnant dogs. A higher risk for patent T. canis infectionsduring the luteal phase was not found. Anyhow, patentinfections were also not detected in the two pregnantbitches examined in that study. The authors raise thequestion whether pseudopregnant bitches, which show ahigh prolactin level, would also develop patent infections(Overgaauw et al., 1998).

3. Pathological changes in the host during larval

migration

3.1. Clinic and gross changes

Larval migration leads to tissue damage in the host(Webster, 1958a). 2472 h p.i. in some dogs bloody-mucous enteritis might occur, caused by the penetrationof the intestinal wall by the larvae (Fernando, 1968).Thearrival of larvae in the lung can then be accompaniedby coughing and dyspnoea (Overgaauw, 1997).The finalintestinal colonisation usually does not cause more thana limited to moderate enteritis, but in cases of massiveinfection it can cause obstructions and ruptures of theintestine (Webster, 1958a). Hayden and Van Kruiningen(1975) compared a control group of dogs naturally infectedwith T. canis and a group additionally infected threetimes a week for 1 month with 3500 embryonated eggseach time, totalling 50,000 eggs. The latter group wasconsidered superinfected. Lesions in the intestinal tractof dogs naturally infected with T. canis were minimal,whereas superinfected dogs had several white nodules inthe jejunum and petechiae in the pyloric mucosa.

A profound colonisation can also cause adult ascaridsto invade the bile ducts, perforate liver parenchyma andfinallyenter theabdominal cavitythrough theliver capsule(Fig. 7).Naturally, this incident leads to a generalised andincurable peritonitis, which exacerbates if female ascaridsin the peritoneal cavity continue to produce and releaseeggs (Dade and Williams, 1975). In the case report ofDade and Williams (1975) on a 5-week-old puppy withinappetence, depression, diarrhea and weakness, duringnecropsy 21 adult ascarids were found in the peritonealcavity, 4 adults were apparently in the process of leavingthe liver as they were protruding from the capsule, and

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

some were found in the liver parenchyma and in the bileducts. 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 weredetected. This phenomenon occursmost often in pups afterextensive prenatal infection. Moreover, a massive numberof ascarids in the intestine of young dogs after prenatalinfection can lead not only to the typical pressure sensitive,drum-shaped abdomen (Fig. 8)but also to developmental

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 rachiticsymptoms (Bosse et al., 1980; Herschel, 1981; Vomann,1985).Vomann (1985)infected four bitches 10 days anteconceptionem with 20,000 embryonated eggs each. Thefifth bitch remained uninfected and therefore its puppiesserved as a control group. Three of the infected bitcheswere treated with fenbendazole to reduce the prenatalinfection, whereas one infected bitch was not treated andtherefore highly infected. All puppies from the untreatedbitch showed alternating obstipation and diarrhea, fur-thermore vomiting, anaemia and rachitis. These puppiessubsequently died between the 22nd and the 49th days.Necropsy revealed ascites, cachexia, anaemia and dilata-tion and partial rupture of the proximal duodenum by aball of ascarids. The cause of death was most often ruptureof the intestinal wall and peritonitis after migration ofadult ascarids through the bile duct and parenchyma ofthe liver. Puppies from treated bitches showed no typicalsigns of toxocarosis, but with 4 kg, compared to the 7 kgbody weight of the uninfected control group at the endof the study, a considerably reduced development ofbody weight. Necropsy showed slight dilatation of theproximal duodenum, a macroscopically thicker stomachand intestinal wall and an activation of the lymphaticsystem (Vomann, 1985).

Among macroscopic lesions in other organs, red andwhite spots in the liver distributed over all lobes occuras well as petechiae and yellowish-white and red lesionsin the lungs. Kidneys often show typical white lesionsthroughout the whole cortex (Hayden and Van Kruiningen,1975; Manhardt, 1980; Herschel, 1981)(Fig. 9).

3.2. Haematological and enzymatic changes

3.2.1. Anaemia

Vomann (1985)examined haematological changes inpuppies after prenatal infection and observed changesin the red blood cell counts. Red blood cell counts arephysiologically low in newborn puppies and increase

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

approximately 68 weeks post partum to reach the valuesin adult dogs. In contrast, puppies heavily infected with T.canisshowed further decreasing numbers of erythrocytes,whichweremostlycausedbysevereinternalbleeding.Rea-sons for the occurrence of internal bleeding were preadultlarvae migrating through the liver as well as perforation ofthe intestine, caused by the massive load of adults. Moder-ately infected puppies showed an increase of erythrocytecounts from the 5th week onwards, but did not reach thosevalues of uninfected animals. In contrast, in adult bitches,no changes in the red blood cell counts were observed afterinfection withT. canislarvae (Zimmermann, 1983).

3.2.2. Eosinophilia

Also characteristic for T. canis infection is theeosinophilia which starts at least on the 7th day postinfection, reaching its maximum approximately 14 dayspost infection (Zimmermann, 1983). A similar timecourse of eosinophilia is observed in prenatally infectedpuppies from day 7 post partum on (Vomann 1985).Vomann (1985) furthermore reported that the degreeof eosinophilia in prenatally infected puppies is almostproportionate to the intensity of the infection. With thecommencement of egg shedding in the faeces the numberof eosinophils slowly decreased, returning to physiologicallevels 42 days p.i. These data showed that the course of theeosinophilia in prenatally infected pups is comparable tothat in experimentally infected adult dogs (Zimmermann,1983).

3.2.3. Increased liver enzyme levels

During infection with T. canis not only changes in theblood counts are observed but also enzymatic changes.During the liver migration, the liver enzymes glutamatedehydrogenase (GLDH) and alanine transaminase (ALT)increased, reaching a maximum 14 days p.i. (Zimmermann,1983).Afterwards, ALT remained on a high level for a cer-tain time period, whereas GLDH returned to physiologicalvalues after 14 days (Stobernack, 1988).After reinfection,liver enzymes show an increase again even though it islower than after the primary infection (Zimmermann,1983). Vomann (1985) observed these two enzymesin prenatally infected puppies to be elevated already atbirth up to 67U/l for GLDH (physiological value up to6.0 U/l) and 365U/l for ALT (physiologically up to 55 U/l)in highly infected puppies. The values returned to normal12 weeks post partum. A second increase caused by adultascarids migrating through the liver and the peritonealcavity was detected shortly before the death of highlyinfected puppies.

3.3. Histopathological changes

Webster (1958a) histologically examined the organsaffected by the parasite. Within this study, 72 h p.i. in theliver parenchyma free larvae were detected as well as aninfiltration of leucocytes. During the following 2448hthe number of leucocytes increased continuously. Mostleucocytes were identified as eosinophils, monocytes andalso polymorphonuclear leucocytes. Webster (1958a)reported a tendency of these cells to gather in nodules,

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

sometimes surrounding a larva in their centre. These struc-tures could be referred to as granulomes. Based on theseprocesses the tissue becomes necrotic, haemorrhages and

marked fatty degeneration of liver cells occur. After 10days first signs of encapsulation marked by a thin fibrouscapsule were seen associated with the retreat of someof the leucocytes. Finally, about 3 weeks p.i., Webster(1958a)observed the beginning of a regenerative processwith fibrous tissue proliferation. Excessive infiltrationof lymphocytes, eosinophils and reticulum cells in theliver was also observed in the study ofHayden and VanKruiningen (1975).

Webster (1958a)further examined the lung and alsofound an infiltration with leucocytes, mainly eosinophils,forming compact nodules. Infected dogs suffered from lob-ular pneumonia and vascular congestion. Heavy infections,

which are not further specified by Webster (1958a), ledto serious pneumonia with exudate mainly comprisingmucus, blood cells, epithelial cells as well as larvae in thealveoli and bronchioles. Furthermore, haemorrhages werecommon and in some cases extensive tissue degenerationwas observed.

Dense aggregates of eosinophils were also found in theintestinal wall, especially in the duodenum and jejunumbut also in the cecum and colon. Even mesenteric lymphnodes had a few capsular and cortical granulomes (Haydenand Van Kruiningen, 1975).Furthermore,Hayden and VanKruiningen (1975)observed mild to severe pyelitis in thekidney in 3 out of 7 dogs and interstitial infiltrates of lym-

phocytes, plasma cells, eosinophils and histocytes in thecortex of 2 dogs. Granulomes directly under the capsule ofthe kidney were also described byHerschel (1981).

Therefore, it can be said that eosinophil infiltration andgranulomes were most pronounced in the liver, kidney andlung (Hayden and Van Kruiningen, 1975).It is known thatlarvae in skeletal muscles of paratenic hosts are enclosedin granulomes (Parsons, 1987). Manhardt (1980) alsomadethis observation in the definitive host (Fig. 10),the centralnervous system being the only exception.

4. Evasion of the immune system

As outlined above,larvae migrate for several weeks,andarrested larvae remain alive for at least several months in

the host tissue. Why therefore should the immune systemnot be able to detect and to successfully eliminate theselarvae? Twoways of immune evasion have been suggested.One hypothesis is that larvae reach a state of hypobiosis inthe tissue, where expression of the immune system stim-ulating antigens is widely reduced. The other hypothesissuggests an immunosuppression induced by T. canis larvaewhich affects the function of T-helper cells. Consequently,the host response to parasite antigens and the productionof specific antibodies is reduced (Overgaauw, 1997).

4.1. The antigenic coat

Many in vitro studies have been conducted to furtherelucidate the mechanisms this parasite uses to evade thehosts immune system. As hatched larvae can be main-tainedin vitroand excretorysecretory (E/S) antigens canbe collected, this stage is regarded as a model for stud-ies on hostparasite-interactions (Maizels et al., 1987).Smith et al. (1981)found that larvae are not recognised byantibodies against E/S antigens at 37 C but are so whenkept at 2 C or at 37 C after preincubation with certainantimetabolites. Both, low temperatures and antimetabo-lites, led to a metabolic inhibition, which was displayed bythe ability of the antibodies to bind to the surface of thelarvae. Furthermore, when the temperature was increasedto 37 C or the antimetabolites were removed, binding offluorescence-labelled antibodies was again not detected.Due to the fact that E/S antigens could be detected all overthe surface of metabolically inhibited larvae but not on thesurface of larvae kept at 37 C, it was concluded that thewhole outer larval surface was involved in the release ofE/S products (Smith et al., 1981).Smith et al. (1981)thenhypothesised that the ability to repeatedly shed E/S prod-ucts on the entire surface could enable the larvae to contin-uously remove antibodies attached to the surface. There-fore, antibody-dependent cell adhesion reactions of thehost would effectively be prevented. This hypothesis wasconfirmed by several authors. Badleyet al.(1987) foundoutthat absorption of immune sera with E/S antigens removedsurface IgG but also C3, which implied that E/S antigenshave the ability to remove antibodies and serum compo-nents necessary for direct complement fixation. Maizelsand Page (1990) found larvae to be bound by comple-ments even up to C9.Furthermore,it wasobservedby usingelectron microscopy that the single layer of granular epi-cuticular material of larvae incubated with immune serumfor at least 30min, thickened and condensed and thatthe denser portion of the epicuticle frequently detachedfrom the surface. Together with the sheath-like layer alsoeosinophils, containing vacuoles opening to the larval sur-face, were detached. Therefore, no gross damage to larvaecould be seen after 30min or after 12h in culture with cellsand serum (Badley et al., 1987).The ability to repeatedlyshed attached antibodies and cells is maintained by highproduction and expression of antigens, which contributeto the surface layer, i.e. E/S antigens. Badley et al. (1987)reportedthattheE/Santigenproductiontotalsquantitiesofupto 8 ngper larvaeand day. Maizels et al. (1984) reportedthat 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/Scomponents from T. canis (TES). Five antigenic compo-nents were identified and named after their molecularweights in kDa: TES-32, TES-55, TES-70, TES-120 andTES-400. All TES-molecules were found to contain morethan 40% carbohydrates, the majority of which were N-acetylgalactosamine and galactose (Meghji and Maizels,1986). All TES molecules were first secreted and thenformed the larval surface coat except for TES-400.

Page et al. (1992)examined the organs which mightbe connected to the secretion of the major glycoproteinsofT. canis, using immunogold electron microscopy. It wasfound that they are derived from two specialised organswithin the nematode organism. The two primary locationswerefoundtobetheesophagealglands,whichsecretetheirproducts via the esophagus in the oral orifice, as well as alarge branched secretory cell, which has its opening to thecuticle at an anterior secretory pore. Additionally, it wasfound 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 lectinhaving greater similarity to mammalian C-type lectinsthan to the respective orthologues in C. elegans. As themammalian C-type lectins are involved in the immuneresponse, the authors supposed an immune evasive strat-egy of the parasite based on the similarity to host immunecell receptors. Subsequently,Loukas et al. (2000)showedthat binding of TES-70 and other TES to host proteins froma canine cell line occurs in a calcium-dependent manner.

The interaction ofT. canis E/S antigens with the immunesystem was further documented in a recent study byGiacomin et al. (2008a). Injected E/S antigens enhanced themigration ofNippostrongylus cantonensis in IL-5 transgenicmice, which have an innate resistance to N. cantonensis.The authors confirmed the activation and consumption ofthe complement factor C3 reported by Badley et al. (1987).Additionally, they surmise a complement-independentmechanism modulating the function of eosinophils, as theresistance of the mice is not based on the complementsystem (Giacomin et al., 2008b). However, the exact mech-anism of the interactions of TES and eosinophils and otherimmune cells remains unclear.

5. Conclusions

Holland and Smith (2006)referred toToxocaraas TheEnigmatic Parasite. There is no better way to preciselycharacterise this fascinating nematode, and the presentstate of knowledge even after decades of intense research,still leavesmany questionsunanswered. Ourpresent, morestringent animal welfare considerations certainly excludeany new attempts to solve the remaining mysteries of lar-val development inside the host and help us to accept acertain level of incomplete knowledge. More importanthowever is that despite a large number of highly efficientanthelmintics, this zoonotic parasite still represents themost prevalent nematode in dogs and that we understand

that intensive research is still required to improve controlstrategies in dogs and to prevent transmission to humans.

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