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Zoonotic helminths affecting the human eye


Nowaday, zoonoses are an important cause of human parasitic diseases worldwide and a major threat to the socio-economic development, mainly in developing countries. Importantly, zoonotic helminths that affect human eyes (HIE) may cause blindness with severe socio-economic consequences to human communities. These infections include nematodes, cestodes and trematodes, which may be transmitted by vectors (dirofilariasis, onchocerciasis, thelaziasis), food consumption (sparganosis, trichinellosis) and those acquired indirectly from the environment (ascariasis, echinococcosis, fascioliasis). Adult and/or larval stages of HIE may localize into human ocular tissues externally (i.e., lachrymal glands, eyelids, conjunctival sacs) or into the ocular globe (i.e., intravitreous retina, anterior and or posterior chamber) causing symptoms due to the parasitic localization in the eyes or to the immune reaction they elicit in the host. Unfortunately, data on HIE are scant and mostly limited to case reports from different countries. The biology and epidemiology of the most frequently reported HIE are discussed as well as clinical description of the diseases, diagnostic considerations and video clips on their presentation and surgical treatment.

Homines amplius oculis, quam auribus credunt

Seneca Ep 6,5

Men believe their eyes more than their ears


Blindness and ocular diseases represent one of the most traumatic events for human patients as they have the potential to severely impair both their quality of life and their psychological equilibrium. Although it is highly unusual, blindness has always been of great interest in human medicine. For example, the evaluation of the emotional and quality of life impacts in patients with some diseases causing blindness (e.g., macular degeneration) gave results similar to those found in diseases such as AIDS, chronic obstructive pulmonary disease, cardiac disorders and leukemia [1]. In addition, blindness has profound human and socio-economic consequences with high costs for the individual, and society, linked to lost productivity and rehabilitation estimated at $42 USD billion per year in 2000, and predicted to reach as high as $110 USD billion per year in 2020 [2].

There are many causes of blindness and those induced by parasitic agents (i.e., Protozoa, Helminths and Diptera) are of major public health concern in developed and developing countries. For example, eye disease caused by river blindness (Onchocerca volvulus), affects more than 17.7 million people inducing visual impairment and blindness elicited by microfilariae that migrate to the eyes after being released by female adult worms in the subcutaneous tissues [3]. Several parasites localize in human eyes as an effect of a specific neurotropism (e.g., Toxoplasma gondii in the foetuses), larval migration (e.g., ascarids, Dirofilaria spp., Trichinella spp.) and, in a few cases, as a primary localization being released directly into the eyes (e.g., Thelazia callipaeda eyeworm and some oestrid fly larvae causing myiasis) [4].

The present article focuses on those zoonotic helminths transmitted from animals to humans that affect the human eye. Undoubtedly, the parasitic zoonotic diseases and their epidemiology have been changing as a result of complex factors including abiotic (e.g., increasing temperatures) and biotic (e.g., demographical changes, political upheaval and land- use practices) that render this topic of great interest for the scientific community [5]. In addition, the impact of zoonotic diseases may vary in relationship to the socio-economic context and to the public health systems in different geographical areas [6], and for some infections, a greater threat exists for populations in developing countries [7]. Zoonotic helminths infecting eyes (HIE) include those transmitted by vectors (i.e., vector borne zoonosis, VbZ), by food consumption (i.e., food-borne zoonosis, FbZ) and those at direct transmission from the environment (i.e., water, soil, etc.) also known as environmentally-borne zoonosis (EbZ). A list of those helminths along with their route of transmission, geographical distribution, localization in the eye and definitive host species is provided in Table 1. Unfortunately, data on HIE are scant and mostly limited to case reports from different countries. Therefore a broad view of these infections on public health is lacking and ophthalmologists have difficulties managing HIE caused diseases and providing a clear diagnosis and therapeutic option for them. The present article focuses on those zoonotic helminths naturally infecting animals but which, occasionally, are transmitted to humans and affect the eye. Recent advances in the diagnosis and control of these parasitic infestations are also discussed on the basis of their distribution in different geographical areas and of interest in travel medicine.

Table 1 Classification (Order, Family and Species) of zoonotic helminths causing human blindness divided according their route of transmission (Vector borne zoonosis, VbZ, food consumption, FbZ, and those at direct transmission from the environment, EbZ), geographical distribution, localization in the eyes and zoonotic relevance.

Biology and pathogenic effects

Helminths at the adult and/or larval stages may infect human ocular tissues externally (i.e., eyelids, conjunctiva sacs, subconjunctiva, and lachrymal glands) or the ocular globe (i.e., optical nerve, intravitreous retina, anterior and posterior chamber). Several parasitic helminths adapted a tropism for animal eyes and related tissues when migrating throughout the host body mainly during their immature stages. This is the case of ascarids and strongylids, causing ocular larva migrans, filarioid species, and larvae of Trichinella, as well of trematode and cestode parasites. Nonetheless, human ocular infestations by zoonotic helminths may also be caused by the parasitic adult stages as in the case of thelaziids (eye worm infestation) and filarioid species including those belonging to the genera Dirofilaria and Onchocerca[810]. Thus, ocular localization of helminths is mainly caused by aberrant migration in host tissues and, only in one case (i.e., T. callipaeda), by direct inoculation into the eyes. What is equally unclear is the route that most follow to gain entry into the eye. It is supposed that some migrate along and follow the optic nerve but others may enter the bloodstream and be carried to the eye in that manner; however, it is not known if these are the preferred or aberrant routes, or even which is the most common route that helminths follow to reach the eye. Once in the eye, larvae likely find it to be a more protected site from host immune responses, but it is not clear that a directed migration into the eye had occurred.

Consequently, ocular alterations caused by zoonotic helminths vary considerably causing mild to severe clinical signs, including lacrimation, epiphora, conjunctivitis, keratitis, corneal ulcers, or retinal lesions, resulting in vision loss (Table 2). For example, in ascarid infections, visual impairment or blindness results from larval migration, with destruction of the visual cortex. In addition, larvae might develop inside the patient's eye (e.g., in baylisascariasis) progressively impairing the vision. However, blindness might be also an effect of the immune reaction the parasites elicit in the host body, or of a combined effect of both presence of the parasite and antibody-mediated reaction. This is the case of ascarids in which ocular signs are related to inflammation because of the presence of larvae and a local immune reaction to them in the retina [11, 12].

Table 2 Ocular tissue affected and symptoms caused by zoonotic helminths (Genus and/or Species) at different stage [41, 63].

The most commonly reported HIE


There are many nematode parasites that can be found in the orbit or within the eye proper (Table 1). Although most nematode infections of the eye are rare, some are more frequently reported than others. In this section, we will discuss those zoonotic nematodes that are most likely to be encountered and reported, by examining their aetiology, case reports and epidemiology.


Trichinellosis (Trichuroidea, Trichinellidae) has a cosmopolitan distribution, but is generally less important as an infection of humans in the tropics than in more temperate regions of the world. Once thought to be a single species (Trichinella spiralis), there are now at least eight distinct species recognized. Each of these species has a slightly different geographical distribution and host range, and only Trichinella zimbabwensis of crocodiles in Tanzania, has not been reported from humans to date. The most striking feature of this group of parasites is their obligatory transmission by ingestion of infected meat containing larvae, either in typical cysts or unencapsulated in the case of several species [13]. The clinical course is characterized by two phases, the enteric/enteral phase, when adult worms are present in the intestinal mucosa, and the parenteral phase, when the released larvae invade the host muscles [14]. During the parenteral phase, which follows the enteral, a typical syndrome of fever, myalgia, periorbital edema and eosinophilia occur. In addition to periorbital and facial edema, conjunctivitis is also frequent. The cause of the orbital and facial edema is not well known. but probably includes some component of an allergic response. Periorbital edema often appears early in the parenteral phase, and typically begins to wane after several weeks. Larvae also affect the macula and retina, causing hemorrhage and other damage as they migrate through and into these ocular tissues [15]. The diagnosis can be suggested from clinical history of ingesting raw or inadequately cooked meat, and the ophthalmologist is often the first contact because of the swollen eyelids and conjunctivitis. Demonstration of larvae in muscle biopsies of patients or in frozen sample of the ingested meat, if available, is still standard procedure to confirm infection. Good serological tests exist and are also very useful in confirming infection, especially in cases with low-level infection where symptoms may be minimal and the number of larvae in muscle may be low. Some areas of the world, such as the United States and Europe, have effectively controlled the infection in humans by removing the parasite from the domestic pig cycle through heightened food safety regulations regarding inspection and feeding practices. In other areas of the world, the domestic pig cycle continues to be responsible for human infection, and in all areas, human infection continues to occur when infected wild game meat is ingested without proper cooking. Because of the wide range of animals that can harbour infection with Trichinella larvae, proper handling and cooking of all meats is recommended.


Ascarids (Ascaridida, Ascaridiidae) occur worldwide infecting various mammals, including humans [16, 17]. Many of these nematodes are causative agents of zoonoses transmitted to humans via contaminated soil. Within the ascarids, Toxocara canis, Toxocara cati and Baylisascaris procyonis are zoonotic parasites of dogs, cats and raccoons, respectively, and they are among the most widespread causes of neural and ocular larva migrans. Indeed, larvae of T. canis are probably the most common nematode infection of the human eye, also known as ocular larva migrans (OLM), and infection in humans occurs worldwide [11]. Infection occurs through the ingestion of infective eggs, most often from soil or other environmental surfaces that have been contaminated with faeces from infected animals. Examination of soil or sand from parks and playgrounds often demonstrates infective Toxocara eggs, which might remain infectious for long periods of time (even years) in the environment [18]. When ingested, the eggs hatch and larvae migrate in the tissues, most often to the liver, but on occasion to other sites such as the eye and central nervous system (CNS). The wandering larvae cause a syndrome, called visceral larva migrans (VLM), of marked eosinophilia, hepatomegaly, fever, cough, and pulmonary infiltrates. The severity of symptoms is often related to the number of larvae acquired, and can range from asymptomatic to acute, with a fatal outcome. The ability of Toxocara larvae to cause OLM was recognized about 60 years ago [19, 20]. OLM occurs most typically in older children (mean 8 yr versus 2 yr for VLM), generally have no other evidence of organ involvement, and hypereosinophilia, hepatomegaly, and pulmonary symptoms are absent, there is no history of pica, and evidence suggests that OLM is caused by a single larva entering the eye. Antibodies to Toxocara tend to be lower in cases of OLM, possibly as a result of fewer infective larvae, and there is experimental evidence that somewhat different immune responses occur between OLM and VLM [21]. Hundreds of cases have been reported and described and untold thousands of cases have probably occurred, even in developed countries, as evidenced by seropositivity in population-based surveys [22, 23]. Worldwide, cases continue to be reported in the literature, including descriptions of lesions, effective treatments, and new/modified methods to observe the infection in the eye [2437]. Visual observation of motile larvae in the eye is possible, although accurate diagnosis is difficult; serodiagnosis continues to be very useful in detecting and confirming cases [38, 39]. Toxocara larvae are approximately 400 by 20 μm and a larva of this size in the eye is highly suggestive. In this presentation, destruction of the larva by photocoagulation is recommended, and prognosis is favourable when recognized early and prompt treatment is provided [40]. After an undefined period of wandering in the tissues, but probably for several weeks or longer, larvae become encapsulated, including those in the eye. These cases, typically present with unilateral visual deficits, with or without ocular pain, and a raised white retinal mass that presents difficulty in distinguishing from retinoblastoma. Unfortunately, in these situations, loss of visual acuity, blindness, and even enucleation of the eye may result. Toxocara larva seen in biopsy specimens or surgically resected tissues are rather easily identified based on size and morphological features. Generally the larva will be enclosed in a granuloma, coiled, and one or more sections of the larva evident. In tissue sections, larvae measure 15-21 μm in diameter and are characterized by a single prominent lateral ala, non- patent gut, and large excretory columns [41]. The prevention of toxocariasis, including OLM, is based on good personal hygiene, including washing hands, and the proper disposal of pet waste, including and specifically not letting pets and stray animals defecate in public places where children and others play and could come in contact with infective eggs. Rubinsky-Elefant and colleagues [11] recently reviewed the subject.

Ocular disease in baylisascariasis occurs in association with severe neural and VLM and, only rarely, alone. Among the ascarids, Baylisascaris species are most often implicated in serious cases of neural and OLM [4245]. Baylisascaris procyonis, a common ascarid of raccoons in many parts of the United States, Europe, and Japan, has demonstrated potential for extensive larval migration in rodents and birds or other accidental hosts where it can produce a fatal eosinophilic meningoencephalitis. This nematode, different from other causes of larva migrans, has an aggressive somatic migration with larval invasion of the central nervous system and capability for continued larval growth within intermediate hosts [46]. Recent serological studies have indicated that significantly more exposure to this parasite is occurring than previously thought, and consequently, low level infections may also be more common than believed [46]. As we recognize increased exposure, the potential exists for more cases of serious visceral and CNS disease, including migration into the eye (Figure 1). Baylisascariasis is acquired in the same manner as other ascarids, through the ingestion of infective eggs from the soil or other environmental sources that have been contaminated by raccoon faeces. Similar to other visceral or OLM, cases of baylisascariasis undoubtedly span the spectra of asymptomatic to serious, often fatal infections. Baylisascaris spp. differ from Toxocara spp. in that the larvae continue to grow, often reaching 1-2 mm in length (by 50 - 60 μm in diameter) in the tissues, including the eye. The migration of such relatively large larvae can result in significantly more pathology than that of Toxocara. The main symptoms are represented by chorioretinitis, optic neuritis, or optic nerve atrophy (Table 2) and examination occasionally may reveal motile larvae migrating within the retina [11, 47] and vitreous humour [48]. Ocular signs are related to the inflammation and local immune reaction of the retina, retinal vasculature, and optic nerve to the larvae. Prevention of baylisascariasis is directed at avoiding ingestion of infective eggs. This is best accomplished by reducing the environmental contamination with raccoon faeces through not feeding wild animals and not encouraging them to live in close proximity to humans, prompt and safe clean up of raccoon faeces/latrines, and not letting children play in areas that have been contaminated with raccoon faeces [46].

Figure 1
figure 1

Nematode larva in the retinal fundus. Larva, presumably of Baylisascaris procyonis, in the retinal fundus of a patient. The infection was presumed to have been acquired in Connecticut, USA. Larva measures approximately 1.4 mm and was treated with retinal lasar. (Original; courtesy of Drs. Caplivski, Bhatnagar, and Goldberg, Mount Sinai School of Medicine).


Gnathostoma spp.

Another form of larva migrans is caused by spirurid worms in the genus Gnathostoma (Spirurida, Gnathostomatidae) [49]. The definitive hosts include dogs, cats, wild carnivores, raccoons and otters. Due to the large range of intermediate (e.g., fish, frogs, chickens and other birds, snakes, pigs, lizards, some crabs, monkeys, hamsters, rats, mice, squirrels, and guinea pigs) and paratenic (birds, snakes, and frogs) hosts, the control of this parasitic infection is particularly difficult. The zoonotic infections occur much more commonly as cutaneous larva migrans (CLM) or VLM, but on rare occasions can invade the eye [5059]. Ocular gnathostomiasis can involve both invasion of surrounding tissues or the eye itself by a wandering larva. In the former, edema and hemorrhage of the eyelid or inflammation of the orbit may occur. When a larva enters the tissue of the eye, corneal ulceration, iris perforation or retinal artery occlusion may occur, with pronounced uveitis, vitritis, vitreal hemorrhage, or secondary glaucoma. Infections are acquired through accidental ingestion of infected copepods that harbour 2nd stage larvae, but more often through the ingestion of poorly-cooked infected fish or other paratenic hosts such as frogs, snakes, birds or even other mammals. Migration of larvae directly from infected animal tissues used as poultices may also occur and place the larvae in the immediate vicinity of the eye. Four species of Gnathostoma have been reported from humans, including Gnathostoma spinigerum, Gnathostoma hispidum, Gnathostoma doloresi, and Gnathostoma nipponicum. However, many other species occur in nature and may pose a threat of zoonotic infection to man. These are relatively stout larvae, and the 3rd stage larva in human tissue can measure between 1 and 5 mm or more in length by 200-600 μm in diameter. The larval stages have many morphological features in common with adult worms, including the prominent head bulb and cuticular spines. In sectioned tissue, the size of the larva is again distinguishing, and occasionally both the head bulb and cuticular spines may be seen, but are absent in many sections. The cuticle can vary between thin and thick, and the muscle cells are numerous and well defined; the lateral chords tend to be large and very prominent, and the gut is distinctive in nature, being composed of many cuboidal cells, each with multiple nuclei, and a luminal brush border microvilli [41]. The intestine and its lumen can be round or variously shaped but is generally a prominent structure noted inside the larva. Serology has proven useful in diagnosing cases [49, 60, 61]. Because of the size of the larva, and their ability to migrate readily in the tissue, its location within the eye is generally of concern, although there are reports of successful recovery of larva from the eye with restoration of visual acuity (Figure 2) [50]. Prevention is primarily through the avoidance of eating poorly-cooked foods or use of raw flesh as poultice.

Figure 2
figure 2

Gnathostoma spinigerum larva in the anterior chamber of the eye. Gnathostoma spinigerum larva in the anterior chamber of the eye of patient in Thailand. (From Teekhasaenee C, Richt R, Kanchanaranya. Ocular parasitic infection in Thailand. Rev Inf Dis. 1986, 8:350-356).


There are a number of different filarioids that have been reported infecting the eye or the conjunctiva, and those reports date back several hundred years, making them one of the oldest groups of parasites known to occur in or on the eye. Indeed, besides the well-known (but not zoonotic) Wuchereria bancrofti, Brugia malayi and Loa loa, some filarioids from domestic and wild mammals (e.g., Dirofilaria spp., Onchocerca spp., Acanthocheilonema (Dipetalonema) spp., Brugia spp., and Loaina spp.) have a zoonotic origin and may infect human eyes [8, 62, 63]. In addition, a number of yet incompletely identified filarioids have been described in human eyes in the Amazon forest regions [64, 65]. For many of them, the life cycle and animal reservoir hosts are poorly known. Animal filarioids occur globally, in many different forms, and all filarioid infections are transmitted by various bloodsucking arthropods; the majority of them, including zoonotic infections, by mosquitoes, although blackflies, culicoids, and others may be involved. Most persons worldwide are at some risk, and those who are more likely to be exposed to the vectors may be at increased risk, but given the worldwide occurrence of animal filaria, there are probably other undefined risk factors.

The largest number of filarioid eye infections is caused by species of Dirofilaria and these result in a worm that migrates across the conjunctiva or is encapsulated in a nodule on the conjunctiva or eyelid. These cases were referred to as Filaria conjunctivae[66], for decades until they were properly aligned with known animal species [67]. Hundreds of these cases have been reported and new cases continue to be reported from wide geographic areas, including areas that have not previously reported such cases [6876]. Undoubtedly, many more cases occur and are either not recognized or reported. In the United States, these are most often caused by Dirofilaria tenuis, a common parasite of raccoons, and in Europe and other parts of the world, by D. repens, a common parasite of dogs and other canids (See additional file 1: Movie1 Surgical removal of Dirofilaria repens from patient's conjunctiva) [77]. Other Dirofilaria species, such as Dirofilaria ursi of bears, Dirofilaria subdermata of porcupines, Dirofilaria striata of wild cats, and others have been isolated from humans on occasion [8]. These species have not been reported to involve the eye, but they certainly could in the future. In worms removed intact or broken from the conjunctiva or seen in histological sections of nodules, the morphologic features of most Dirofilaria make them relatively easy to identify to genus level [8]. They tend to be large, robust worms, and they have distinctive longitudinal and circular cuticular ridging that gives the external cuticle a beaded or corn-row appearance. This can be seen easily in gross specimens that have been removed intact, and is one of the more prominent features noted in histological section as well. Additionally, in sections, the worms have numerous strong muscle cells (polymyarian and coelomyarian) giving a strong body wall. Determining the species is more difficult, especially if a male worm is not present, and final diagnosis is often based on the presumed location of acquisition (i.e., D. tenuis if in the United States, D. repens elsewhere). Once removed, clinical signs quickly resolve and there are no residual sequelae.

Additional File 1: Surgical removal of Dirofilaria repens from patient's conjunctiva. This video shows the surgical removal of Dirofilaria repens from the patient's conjunctiva, after topical anaesthesia. The palpebral fissure was maintained open by using the blefarostat, the nematode was extracted after incision of the conjunctiva membranes and it was collected. A shortened version of a video in Otranto D, Brianti E, Gaglio G, Dantas-Torres F, Azzaro S, Giannetto S. Human ocular infestation by Dirofilaria repens (Ralliet and Henry, 1911) in a canine dirofilariosis-endemic area. Am Jour Trop Med Hyg 2011 (in press). (MP4 3 MB)

Worms identified as Dirofilaria have also been reported from within the eye, either the anterior chamber or vitreous. Some of these cases have been attributed to D. immitis, the dog heartworm, D. repens, or Dirofilaria roemeri of kangaroos [8]. A small number of them were successfully removed and identified based on morphology. A case of intravitreal dirofilariasis was recently reported from Turkey [78] and a case of human intraocular dirofilariasis has been reported from northern Brazil (See additional file 2: Movie2 Surgical removal of Dirofilaria immitis like nematode) [65]. The nematode from Brazil was morphologically and phylogenetically close to D. immitis but genetically distinct from reference sequences, including those of D. immitis collected from infected dogs in the same area. The possible existence of a closely related zoonotic Dirofilaria species in Brazil and its implications have been discussed and serve to highlight the high number of yet unknown species infecting wild mammals that have potential to cause zoonotic infections. Different from the above reported case, worms were generally of modest size, making removal rather than photocoagulation the preferred method of treatment. Dirofilaria spp. worms are often motile, and noticed by the patient because of interference with vision. Removal is curative and full visual acuity is generally restored with no long term sequelae.

Additional File 2: Surgical removal of a Dirofilaria immitis -like nematode. This video shows the surgical removal of a Dirofilaria immitis -like female nematode from the anterior eye chamber of a patient from Parà, Brasil. Eye was clipped and the cornea incised with a crescent Beaver corneal knife. The nematode was extracted alive with forceps and Fukasacu hook. The patient recovered without complications after the surgery. A shortened version of a video in Otranto D, Diniz DG, Dantas-Torres F, Casiraghi M, de Almeida INF, de Almeida LNF, Nascimento dos Santos J, Penha Furtado A, de Almeida Sobrinho AF, Bain O Human intraocular filariasis caused by Dirofilaria sp., Brazil. Emerg Infect Dis 2011 (in press). (MP4 4 MB)

In addition to Dirofilaria, several cases of small Acanthocheilonema -like worms within the eye have also been reported (Figure 3) [63]. Although removed and examined, it was not possible to positively identify these worms to species. In a single case report, Acanthocheilonema (Dipetalonema) reconditum, a subcutaneous filarial infection in dogs worldwide, was removed from the subconjunctiva in a patient from Australia [79]. Two other unusual infections caused by Macacanema, a parasite of monkeys, have been reported where worms were removed from the conjunctiva of humans [80, 81].

Figure 3
figure 3

Acanthocheilonema - Mansonella -like worm. Anterior end of an Acanthocheilonema - Mansonella -like worm removed from the posterior chamber of the vitreous of a patient in Kansas, USA. Scale bar = 500 μm. Original; courtesy of DPDx, CDC.

The zoonotic role of filarial infection in humans in some regions is far from understood. Recently, a mature male filaria extracted from the iris fibers of a man from the Amazon region of Brazil was identified as belonging to the genus Loaina or Pelecitus (See additional file 3: Movie 3 Surgical removal of Pelecitus sp. from the iris fibers of a patient) [64]. This human case and a previous one from Colombia [62] were of unknown origin and both occurred in the tropical Amazon region but little is known about the source of the infection. Vectors of species of Loaina and Pelecitus are mosquitoes, mallophagans or tabanids, as shown with the three cycles elucidated [8284]. Unfortunately, despite some studies on these genera [85, 86], information on these taxa is scant. In the same manner, worms removed from the eye and identified as Brugia are often hard to identify as B. malayi of human origin or Brugia spp. of animal origin [87].

Additional File 3: Surgical removal of Pelecitus sp. from the iris fibers of a patient. This video shows the surgical removal of a Pelecitus sp. male nematode (approximately 4 mm in length) from the iris fibers of a patient from the Amazon region, Brasil. After peribulbar anesthesia and corneal incision of 2 mm. The nematode was extracted by aspiration and the surgery had no complication. A shortened version of a video in Bain O, Otranto D, Diniz DG, Nascimento dos Santos J, Pinto de Oliveira N, Negrão Frota de Almeida I, Negrão Frota de Almeida R, Negrão Frota de Almeida L, Dantas-Torres F, Frota de Almeida, Sobrinho E: Human intraocular filariasis caused by Pelecitus sp., Brazil. Emerg Infect Dis 2011 (in press). (MP4 3 MB)


Taenia crassiceps

Human ocular infection by Taenia crassiceps occurs when individuals accidentally ingest eggs in contaminated food or water. Indeed, T. crassiceps is a tapeworm closely related to Taenia saginata and Taenia solium and adult stages live in the intestine of carnivores and pass eggs with the faeces. In the intermediate hosts, primarily rodents, the immature cestodes (i.e., cysticerci) develop in the peritoneal cavity [88]. In humans, larvae invade the bloodstream and reach subcutaneous and muscular tissues of immune- compromised individuals [88, 89]. Interestingly, larval T. crassiceps in the anterior eye chamber or subretinally (Figure 4) have been reported in immunocompetent humans, in the United States [90, 91] and Europe [92]. The infection by cysticerci of T. crassiceps may be asymptomatic or cause iridocyclitis and/or retinitis [88, 92]. Surgical intervention on the anterior chamber or the subretinal space is successful in curing the infection [88, 92]. Although not a zoonosis, cysticercosis caused by Taenia solium, has been reported from the human eye [93].

Figure 4
figure 4

Cysticercus cyst of Taenia crassiceps in situ. Retinal photograph showing budding cysticercus of Taenia crassiceps in situ. (From the collection of Herman Zaiman, "A Presentation of Pictorial Parasites").

Coenurus cerebralis

Coenurus cerebralis is the larval stage of the tapeworm T. multiceps (syn. Multiceps multiceps), which develops in the small intestine of dogs, foxes, and other canids (definitive hosts). After ingestion by the intermediate hosts, the oncospheres penetrate the intestinal mucosa, enter the bloodstream, and reach the brain where they develop into the infective cystic coenuri [94, 95]. Coenurosis rarely occurs in humans through accidental ingestion of eggs, causing mainly cerebral lesions but also localizing in the eyes [92]. C. cerebralis ocular lesions cause severe anterior uveitis, retrolental or orbital cystic tumor-like masses, and subretinal lesions (Figure 5). Subconjunctival localization may also occur after accidental direct inoculation with infective eggs. The onset of inflammatory responses result in a red and painful eye, followed by development of glaucoma, retinal fibrosis, and ultimately blindness as the final result of the infection [96, 97]. Surgical removal of accessible cysts (Figure 6). is the only choice to cure the infection [96, 98].

Figure 5
figure 5

Coenurus cyst behind displaced retina. Sagittal section of eye from Ghanaian showing coenurus cyst with multiple protoscoleces lying behind displaced retina. (From Parasites In Human Tissues, Orihel and Ash, ASCP Press, 1995).

Figure 6
figure 6

Coenurus cyst after surgical removing from the eye. Intact coenurus cyst removed from subconjunctival tissue an Ugandan child showing multiple protoscoleces. (Original by Paul Beaver).

Spirometra spp.

Analogously, adult Spirometra cestodes live in the small intestine of carnivores where they release eggs which reach the environment with the host faeces. Larvae of Spirometra spp. tapeworms infect domestic animals and humans. Humans are dead-end hosts given that they become infected mostly by drinking polluted water (via ingesting the immature procercoid), or eating inected intermediate hosts (i.e., frogs, birds, snakes, that, along with rats and mice are infected with the larval stages) and assuming the plerocercoid larvae. Once ingested, the larvae ("spargana") may invade muscles, subcutaneous tissue, urogenital and abdominal viscera, and, sometimes, the central nervous system and the eyes [99]. Human ocular sparganosis has been reported from South America [100], Central Europe [101] and Asia [102105]. Spargana usually infect subconjunctival and conjunctival tissues causing symptoms varying from simple itching due to local granulomata to more serious signs represented by local pain, epiphora, chemosis, and ptosis [105, 106]. Conjunctival infection may also be characterized by irritation, continued foreign body sensation, redness [104] and mimic signs and symptoms of orbital cellulitis, with exophthalmia and corneal ulcers. When the immature cestode invades the orbit it may cause acute anterior uveitis and iridocyclitis [101] and severe inflammation with blindness [107]. Unfortunately, surgery is the only effective treatment [104, 105].

Echinococcus granulosus, Echinococcus multilocularis and Echinococcus oligarthrus

Echinococcus granulosus, Echinococcus multilocularis and Echinococcus oligarthrus are tapeworms that occur worldwide. Adult stages of E. granulosus and E. multilocularis infect mainly dogs or wild canids (e.g., wolves, jackals, coyotes and foxes) while E. oligarthrus adults infect wild felines [108111]. Along with several other animal species, human may act as (accidental) intermediate hosts of these cestodes by ingesting food contaminated by their eggs [110, 111]. When a human being inadvertently ingests eggs, the larvae hatch and disseminate via the bloodstream into different organs and viscera (mostly liver or lungs but also heart), where they produce a typical hydatid cyst (E. granulosus, E. oligarthrus) or many alveolar small cysts (E. multilocularis) causing a major zoonotic disease [108111].

Although not very common, ocular infection by larval Echinococcus spp. may thus occur as a consequence of bloodborne dissemination of the oncospheres. Ocular localization by the larval form of E. granulosus accounts for 1 to 2% of all reports. Intra-orbital hydatid cysts by E. granulosus may cause severe exophthalmia [112] pain and blindness as the hydatids have the ability to fill the vitreous cavity [113] or severe inflammation of orbital structures and acute eyesight loss due to the rupture of intraorbital hydatids [114]. Ocular alveolar hydatidosis caused by E. multilocularis may occur after spreading of the larval cestodes to other sites. For instance a choroidal eye mass has been reported in a patient with history of visceral alveolar hydatid disease with cerebral metastasis [115]. Human infection by E. oligarthrus is very rare with only a few cases published in the international literature, two of which involve the eye [116]. Nonetheless, the ocular localization of E. oligarthrus has a relevant clinical impact since it causes the presence of a single orbital, retro-ocular cyst in the orbit [117] or the occurrence of a retroocular cystic tumor-like mass inducing exophthalmia, chemosis, palpebral ptosis, and blindness [108].


Fascioliasis, also known as liver fluke, is caused by Fasciola hepatica and Fasciola gigantica, trematodes which localize in the biliary ducts of the definitive hosts, grass-grazing domestic and wild ruminants (i.e., cattle, sheep, goats, buffaloes) and also horses and rabbits. This parasite develops through various larval stages in water snails of the genus Limnaea which release cercariae that encyst as metacercariae on aquatic vegetation. Infection occurs when animals ingest freshwater plants or water containing encysted metacercariae [118]. Within the last decade, reports of human cases of fascioliasis have increased [119]. Although migrating immature F. hepatica flukes in humans have been mainly reported in blood vessels, lung, subcutaneous tissue, and ventricles of the brain [119], they have also been recovered from the anterior chamber of a patient in Iran [120].

Alaria americana (syn. canis) is a three-host trematode that lives as adults in the intestine of the dog definitive host. Eggs are passed in faeces and hatch in water, releasing miracidia which penetrate the helisomid snails (first intermediate host) and develop through the sporocyst stage into cercariae [121]. Cercariae released from snails actively penetrate the second intermediate host (tadpoles) becoming infective mesocercariae in about two weeks. In the tadpole or in the frogs (following the metamorphosis), mesocercariae accumulate and may be ingested by a number of paratenic hosts (e.g., other frogs, snakes) or directly by the definitive host. Cases of human intraocular infection with mesocercariae of A. americana and other Alaria mesocercariae have been recorded in patients who had ingested undercooked contaminated frogs legs [122]. Both patients presented with pigmentary tracks in the retina, areas of active or healed retinitis and signs of diffuse unilateral subacute neuroretinitis (Figure 7).

Figure 7
figure 7

Alaria sp. in the eye. Freely moving Alaria mesocercaria on the retina of the eye. The anterior sucker is evident on the left side of the organism. (From the collection of Herman Zaiman, "A Presentation of Pictorial Parasites").

The trematode Philophthalmus lacrimosus (Philophthalmidae), as adults, parasitize the eyes of birds (definitive host). Eggs containing miracidia hatch in the water, miracidia penetrate snails (intermediate hosts) and develop into redia and cercariae. When the metacercariae encyst on surfaces of food for birds the infection of a new definitive host can take place by entering the eye or by oral intake [123]. Human cases of philophthalmosis have been reported in Europe (Yugoslavia), Israel, Asia (Thailand, Sri Lanka, Japan) and America (i.e., Mexico, and the United States) [124].

New or reemerging zoonotic helminths infecting human eyes

Over the last decade, parasitological knowledge has been considerably refined and enhanced by the use of sophisticated technologies and molecular tools, and by the interdisciplinary approach in many fields of the human and veterinary medical sciences. Increasing awareness of physicians on previously poorly known diseases likely is an important part of this process. Reports of several of these infections of the eye have been increasing, but whether this is due to a higher awareness or increasing rates of infection is unclear. Possible reasons for the highest number of reports may include changing epidemiological patterns in the natural definitive hosts, leading to increased exposure of humans, and new geographic range because of spreading into new areas. Three key examples, namely infections by Angiostrongylus cantonensis, Thelazia callipaeda and Onchocerca spp., are discussed below.


Metastrongylids encompass a large group of nematodes (Strongylida, Metastrongyloidae) infecting organs and tissues of different vertebrates [125]. A. cantonensis, also known as rat lungworm, is a well recognized zoonotic infection and, as such, is the primary cause of eosinophilic meningitis in Southeast Asia. The infection has spread widely to many other areas of the world, including the Caribbean and Americas [126130]. The parasite also enters the eye with some frequency. In a review of 484 cases of eosinophilic meningitis, Punyagupta and colleagues [131] noted that 47 (16%) of the cases had reported ocular involvement, and in 7 cases an actively motile worm (most probably A. cantonensis) was visualized and removed from the anterior chamber of vitreous of the eye [132, 133]. Human ocular infections by larval rat lungworm have been reported in several countries in Southeast Asia [134143] and, they likely will continue to be reported wherever the parasite occurs, including in new geographical areas such as the Caribbean [144]. Ocular lesions by A. cantonensis may either occur alone or may accompany other symptoms such as meningitis [142, 145]. Often these long and slender worms reach considerable size in the eye, and are up to a centimeter or more in length [141]. The female worm has a distinctive helical pattern of dark intestine intertwined with light coloured reproductive tubes; male worms have a copulatory bursa and very long (> 1 mm) spicules. These features make it fairly simple to recognize and identify a large worm removed from the eye as A. cantonensis.

Infections are acquired through the ingestion of infected intermediate snail or slug hosts, or a variety of paratenic hosts such as amphibians, reptiles, and some crustaceans that have become infected by ingestion of snails and/or slugs. Migrating larvae in the human host make their way to the CNS, and occasionally into the eye, possibly along the optic nerve. Caution in handling and not eating raw or poorly cooked intermediate or paratenic hosts should prevent most human infections. There is some evidence that people can also be infected by ingestion of produce that has been contaminated with slime trails of snails or slugs into which infective larvae have been shed. Washing produce may help reduce the risk of infection but probably does not remove all risk in endemic areas with high level of transmission. Although in a number of cases, worms are successfully removed, ocular disease caused by larval A. cantonensis may vary from blurred vision due to intraretinal haemorrhage [144] to severe optic neuritis due to the presence of the nematode in the vitreous cavity [143]. Other signs include visual disturbances and impairments, extraocular muscular paralysis and a wide range of ocular inflammatory conditions [141, 146, 147]. Diagnosis is based on the identification of a (usually single) living worm, in any eye localization (e.g., anterior chamber, vitreous cavity, and subretinal space) (Figure 8). The treatment regimen relies on the surgical removal or laser therapy, accompanied by oral benzimidazole (e.g., mebendazole) and corticosteroids in the case of inflammatory manifestatios such as retinitis or optic neuritis [147]. Ocular damage caused by A. cantonensis can be severe and may be permanent, thus in some patients the outcome is poor and depends on the initial visual acuity [141, 147].

Figure 8
figure 8

Angiostrongylus cantonensis in the anterior chamber of the eye. Angiostrongylus cantonensis in the anterior chamber. (Original by John Cross, courtesy of Lawrence Ash).


Thelazia callipaeda (Spirurida, Thelaziidae) represents a good example of HIE that is both spreading and new to the scientific community in western countries. Along with Thelazia californiensis that has been reported to infect humans occasionally in the United States [148], T. callipaeda is the only helminth transmitted by secretophagous flies directly into the orbit of humans [149]. This nematode primarily affects the eyes of domestic dogs and cats and wild carnivores (e.g., foxes, wolves, beech martens and wild cats) (See additional file 4: Thelazia callipaeda infecting the eye of a dog) [150]. Since its first description at the beginning of the previous century, this nematode has been known as the "oriental eye-worm" for its distribution in the former Soviet Union [151] and the Asian continent, including China [10], Korea [152], Japan [153], Indonesia [154], Thailand [155], Taiwan [156] and India [157]. Human thelaziasis may cause mild to severe clinical signs (including lachrymation, epiphora, conjunctivitis, keratitis and/or even corneal ulcers) [125]. The worm is transmitted by various secretophagous flies which feed on lachrymal secretions of infected animals and/or humans, thus ingesting Thelazia 1st stage larvae and, after obligate development in the fly, depositing 3rd stage larvae directly back into the orbit. The competence of drosophilid flies of the genus Phortica (Diptera, Drosophilidae) as vectors of T. callipaeda has recently been elucidated under both laboratory and natural conditions [158, 159]. We now recognize that T. callipaeda infection is widespread throughout Italy with infection prevalence as high as 60% in dogs from some municipalities (Figure 9) [160] and also in southwestern France (Dordogne area) [161, 162] and Switzerland [163]. In addition, four cases of human thelaziasis have been diagnosed in patients coming from an area of north-western Italy and south-eastern France [164]. Infected patients present with exudative conjunctivitis, follicular hypertrophy of the conjunctiva, foreign body sensation, excessive lachrymation, itchiness, congestion, hypersensitivity to light and keratitis, depending on the number of nematodes present in the eye [10]. Children and the elderly seem to be at higher risk. Thelazia worms are generally removed intact from the eye, and there are several morphological features that assist in identifying them from other worms that might occur in the orbit, including filaria such as Loa or Dirofilaria. The morphological identification of T. callipaeda has been reviewed [165]. The adult worms measure from 5 to 20 mm in length by 250 - 800 μm in diameter (males are smaller than females). They have a distinct buccal capsule and the cuticle has typical, regularly spaced distinct transverse striations giving the cuticle a ridged appearance (Figure 10). In addition, adult females of T. callipaeda are characterized by the position of the vulva located anterior to the oesophagus-intestinal junction, and the males possess five pairs of postcloacal papillae. Poor living conditions and low socio-economic standards seem to be risk factors for acquiring infection, and better hygiene would probably contribute to prevention. The adults and larvae of T. callipaeda can be removed mechanically by rinsing the conjunctival sac with sterile physiological saline whereas adults can also be isolated with forceps or cotton swabs [10].

Figure 9
figure 9

Thelazia calllipaeda in a heavily infected dog. Heavy infection by Thelazia callipaeda nematodes in the conjunctiva of a dog from Italy.

Figure 10
figure 10

Thelazia californiensis from a human patient. Posterior end of a female Thelazia californiensis from the conjunctiva of a human patient in New Hampshire, USA showing cuticle serration. Scale bar = 50 μm. Original; courtesy of DPDx, CDC.

Additional File 4: Thelazia callipaeda infecting the eye of a dog in Basilicata region (southern Italy). This video shows Thelazia callipaeda nematodes floating in the eye of an infected dog in an endemic area of Italy. Conjunctivitis and lacrymation were the main symptoms observed. In the second part, numerous T. callipaeda specimens have been collected by an ocular swab. (MP4 1 MB)


As previously noted, the vast majority of filarioid infections of the eye occur on the conjunctiva, and are caused by species of Dirofilaria. However, there is an increasing number of reports of zoonotic Onchocerca infections, and several of these have been either within the eye or associated with the conjunctiva or connective tissue of the orbit (See additional file 5: Onchocerca sp. infecting the anterior chamber of a human patient). Of the 15 clinical cases reported to date [166, 167], five have been associated with the eye; 3 involved the conjunctiva and 2 involved the cornea. These have been reported from Crimea, the United States, Albania, Hungary, and Turkey [168172]. The species causing infections of the eye have tentatively been attributed to Onchocerca gutturosa or Onchocerca cervicalis[169, 173], Onchocerca reticulata[170], Onchocerca spp. [171], and, Onchocerca lupi[172]. This last species, O. lupi, is of particular interest because it affects dogs and it induces acute or chronic ocular disease characterized by conjunctivitis, photophobia, lacrimation, ocular discharge and exophthalmia [166]. In most cases, zoonotic Onchocerca spp. are encased in a nodular granuloma, are resected and sectioned, and the worms identified morphologically (Figure 11a). In section of Onchocerca the distinctive muscle anatomy, composed of few, low, poorly developed cells, and the characteristic structures of the cuticle are often apparent, including the circular ridges and inner cuticular striae, making the identification straightforward. The distances between the prominent, undulated annular ridges and the number of transverse striae in the internal layer represent the morphological characters for differentiating filarioids belonging to the Onchocerca genus (Figure 11a). The apparent increase in number and range of zoonotic Onchocerca infections including those affecting the eye, is noteworthy but difficult to fully explain. Recent cases in both the US and Europe highlight this trend. Case reports of canine ocular onchocerciasis by O. lupi[166] have also increased in Europe, including in Greece, Portugal, Germany, Hungary, and Switzerland [174177]. The number of cases of canine ocular onchocerciasis have also increased in the United States but the species of parasite in the United States has not been established [178180]. The role played by dogs as reservoir of this zoonotic agent deserves to be investigated further to establish both the primary definitive hosts as well as the vectors that serve to transmit the infection naturally and to humans.

Figure 11
figure 11

Zoonotic Onchocerca from human ocular connective tissue. Zoonotic Onchocerca sp. from a nodular granuloma of the eye in a patient from Ohio, USA. a) Transverse section of female worm shown in Fig. 5a encased in a nodular granuloma. Low cuticular ridges and inner striae, 2 per ridge, are evident Hematoxylin and eosin stain. Scale bar = 50 μm. (Original; courtesy of Drs. Yassin and Hariri, University of Pittsburg Medical Center). b) Short piece of female Onchocerca sp. removed from granuloma tissue before fixation, showing characteristic, diagnostic structures of the cuticle with circular ridges and inner cuticular striae. Scale bar = 150 μm. (Original; courtesy of Drs. Yassin and Hariri, University of Pittsburg Medical Center).

Additional File 5: Onchocerca sp. infecting the anterior eye chamber of a human patient. This video shows the occurrence of Onchocerca sp. in the anterior chamber of a patient from Colorado, United States. The nematode was surgically removed, extracted alive and identified as Onchocerca. The patient recovered without complications after the surgery. A video from the case presented in Burr WE, Brown MF, Eberhard ML: Zoonotic Onchocerca (Nematoda: Filarioidea) in the cornea of a Colorado resident. Ophthalmology 105:1494-1497, 1998. Video courtesy of Dr. W.E. Burr. (MP4 483 KB)

Diagnosis and cure

The diagnosis of the causative agent is usually only possible after surgery and extraction of the worm or tissue containing the worm, and often requires the assistance of a specialist with an appreciation of the microscopic features of helminths. Generally, in those cases where the parasite is amenable to obliteration with photocoagulation or laser surgery, only a tentative diagnosis is possible. However, surgery remains the only option available for treating a number of the HIE. Although invasive, and often requiring sophisticated devices and advanced medical expertise (not always available in developing countries), ocular surgery is often curative and also allows the recovery of helminths to identify them (Table 3). At the time of ophthalmological examination, observation of motile larvae in the eye is occasionally possible, although a diagnosis at species level can be difficult. Indeed, in some cases it is possible to tentatively identify helminths by measuring larvae in the retina. This is the case with B. procyonis larvae which are larger (1 to 2 mm by 50 to 60 μm) than those of Toxocara spp. (< 400 by 15 to 20 μm) [47]. This larger size often allows a quick differentiation between Toxocara and Baylisascaris larvae when seen either in tissue sections or within the eye. Although Baylisascaris larvae share several morphologic features in common with Toxocara, including single lateral cuticular alae and paired excretory columns, they differ markedly not only in size but in the fact that the gut is patent (has a distinct lumen) in Baylisascaris spp. Other types of nematode larvae, such as those of Gnathostoma spp., are relatively easy to identify because of their stout, robust size and distinctive head bulb whether they are visualized in the eye or removed. Similarly, A. cantonensis worms often reach considerable size in the eye, but are long and slender worms, up to a centimetre or more in length, and are not easily confused with ascarid or Gnathostoma larvae, being more likely mistaken for a filarioid. Microscopic examination, however, would quickly allow separation of Angiostrongylus from a filaria. Other HIE, such as Alaria mesocercaria could be distinguished on the basis of its shape, size (500 × 150 μm), and movement [122]. Their localization on the conjunctiva surface, size, and morphologic features would allow a relatively easy diagnosis of T. callipaeda.

Table 3 Informative morphological characters and measurements of different stage of helminths infecting human eyes [41, 63].

On the other hand, when HIE are enclosed in a granuloma or within the subconjunctiva, the identification is more difficult. Furthermore, in certain situations, removal may increase the risk to the patient, as in the case of the cystic forms of Echinococcus spp. which could induce anaphylactic immunoreactions when disturbed. Indeed, although rare, the localization of Echinococcus spp. cysts in the eye are always cause of severe disease, thus the careful surgical removal of the cysts is the only option.

Where appropriate tests exist, serological diagnosis can often contribute to a definitive diagnosis of infection, such as in the case of some ascarids for which serological testing using a sensitive, specific enzyme immunoassay (EIA or ELISA) is available. Serological testing is available for baylisascariasis and can be very helpful in identifying and confirming infection, and, like for toxocariasis, in conducting serosurveys to document the degree of exposure in different populations. Unfortunately, individuals may not mount a measurable immune response during the early phases of acute infection and serologic testing will not provide conclusive evidence to help guide treatment, hence the need for aggressive presumptive treatment in cases with solid exposure history [46]. Serologic assays can be very helpful to confirm infections caused by Gnathostoma, especially in cases where no larva or tissue is available to examine. Conversely, seropositivity to spargana in IFAT or ELISA tests always needs to be confirmed by histological examination [104, 105].

Unfortunately, surgery is often the only effective treatment for many HIE (e.g., ocular sparganosis, A. cantonensis) and this is one of the reasons why these infections represent a traumatic event for the patients and treatment is not a particularly cost-effective manner in which to manage the infection. In some cases, such as the typical zoonotic filarial infection, only a single worm is present and the surgical removal is both therapeutic and curative. In other instances, most notably OLM or larval tapeworms, there is some likelihood that additional larval stages may exist and chemotherapy may be indicated with corticosteroids in the case of inflammatory conditions such as retinitis or optic neuritis [147].

It should be noted that the use of photocoaggulation and laser ablation continue to prove useful in a number of cases infection with HIE, e.g., ascarid larvae, Alaria mesocercaria, and often result in improved visual outcome while at the same time destroying the invading helminth in situ[122].

Concerns, ignorance and new avenues linked to HIE

Ophthalmologists and physicians often lack an in- depth knowledge of parasites, rendering it difficult for them to correctly address the etiological identification, treatments and control strategies for many HIE. In addition, the scientific information on HIE available in the international literature is scarce and limited to single case reports in which a clear comparative differentiation among helminth infections is not considered. The main limitation for correctly identifying the etiological agent is that often helminths are not removed, or they are seriously damaged during the surgical procedures thus rendering an accurate morphological identification difficult, if not impossible. The microscopic identification of helminths at the species level often relies on the examination of key morphological characters, not all of which are present on any given specimen or not recognized by the person making the examination, sometimes resulting in an incorrect diagnosis. Accurate identification is crucial to understanding both the source of infection and environmental risks, as well as prescribing correct treatment options.

There are several cases in the literature in which helminths were erroneously identified; for example, cases of Trichinella sp. in the vitreous of a woman and Toxocara sp. in the retina of a man both from Germany, and a case Angiostrongylus sp. recovered from the anterior chamber of a man from Sri Lanka were all incorrectly identified as filaria (reviewed in [63]). Twenty-eight cases of human dirofilariasis from the Old World were erroneously attributed to D. immitis, subsequently reviewed and correctly attributed to D. repens[181]. Recently, the helminth causing a case of human intraocular infestation in Japan was erroneously identified as T. callipaeda although the picture published in the article portrayed a filarioid [182]. In the same article, the authors stated that the life cycle of T. callipaeda remains unclear and discussed the possibility of human infection through the skin or by drinking untreated water. This somewhat implausible hypothesis was already dispelled in the late 1990s [183]. The scant attention of medical researchers towards human thelaziasis may also be attributable to the difficulties in its clinical diagnosis and differentiation from allergic conjunctivitis, particularly when small numbers of adult or larval stages are present in affected patients. More recently, the advent of molecular biological techniques has largely supplemented and enhanced knowledge of parasitologists in areas such as systematics (taxonomy and phylogeny), population genetics and molecular identification, diagnosis and control of some HIE [184]. Indeed, the advent of PCR made it possible to study damaged and incomplete specimens, or fragments of specimens encysted in tissues which otherwise would not be morphologically identifiable [185]. The importance of molecular identification and barcoding approach (by the specific PCR-amplification of the cox 1 and 12S genes) for the rapid identification of specimens has been emphasized, including for either recognized or yet unknown species. Recently, an integrated DNA barcoding of cox 1 and 12S markers and morphology approaches was shown to be a powerful tool for the taxonomical identification of many filarioid species even if small nematode fragments were available [184]. In addition, the delineation of Molecular Operational Taxonomical Units (MOUTS) was useful to infer potential new species [184].

Basic parasitological research in this field is often fragmentary due to the fact that experimental human infections are rarely done, and the retrieval of helminths from the patients' eyes may be an infrequent occurrence during the ophthalmologic examination. For a number of these helminths, poor experimental models exist, or, if good models exist, the infections generally do not affect the eye in the same way that occurs in aberrant human infections. Thus, scientific knowledge in this field, as well the information on helminth migration patterns is limited, and often has been gained from studies of the same parasites in other animal models. All the above concerns need to be addressed through basic and applied research. For example, many nematode species have not yet been described and even those that are known are often poorly studied such that there is a lack of basic information on the helminth fauna of wild animals (e.g., O. lupi). This is particularly true, but not restricted to, regions of the world, such as the Brazilian Amazon forest, where there is wide biodiversity and a large amount of animal and plant species yet to be described [186]. Consequently, species identification of some groups of HIE, such as filarial nematodes, can be difficult if not impossible. Another example of insufficient information is represented by the unknown risk of zoonotic infection, such as other species of Baylisascaris (in addition to B. procyonis), that may be considered as potential zoonotic agents [46]. For example Baylisascaris transfuga, infecting bears worldwide [45, 187] has been reported to produce visceral, neural, or OLM syndromes in mice [188190], gerbils [45, 191], and guinea pigs [192]. In addition, cases of fatal neurological diseases have been reported in a colony of Japanese macaques (Macaca fuscata fuscata) housed with American black bears in a safari-zoo in Japan [193]. However, the zoonotic role of this parasite for humans has never been demonstrated. Since bears are frequently kept in zoos and game parks and often have high prevalence of the infection in the population (up to 50-100% of bears harbour this parasite) studies on the zoonotic capacity of this parasitic species would be pivotal for a better understanding of the public health risk [194]. Overall, a better understanding of the biology of a number of HIE is crucial for addressing their prevention.

Better awareness among physicians (including ophthalmologists) in the field of parasitology and more active collaboration with parasitologists would be very helpful in proper diagnosis, control and prevention of HIE. This would also allow a better knowledge of the potential risks for being infected by an HIE agent in a given area as well as exposure when travelling in endemic areas. Physicians and ophthalmologists need increased awareness about the existence of a range of zoonotic helminths other than those natural parasites of humans that might be expected to be found in patients' eyes.

Unfortunately, there is a lack of knowledge about many parasites in the local fauna and limited basic research studies are carried out. Monitoring and periodic surveillance for the infections of both domestic and wild animals is important to provide a better understanding of what potential pathogens exist locally, and to prevent the HIE. This is the case with B. procyonis which is an emerging infection in raccoons in the southeastern United States, an area traditionally considered to be at low risk [195, 196]. An increasing appreciation of onchocerciasis in domestic and wild animals in Europe and the United States is needed to accurately understand what species exist, what the natural definitive host is and, ultimately, what the risks for human infection are. Veterinarians, physicians, and public health officials all share the need to be alert to the possibility of zoonotic infections inside and outside of traditional high-risk areas. Lastly, we need a better understanding of why some parasites migrate to and occasionally enter the eye, especially given that none of these helminths typically resides in or around the eye.


Despite scientific advances and new methods for treating helminth infections in the human eye, therapies available to patients are somewhat limited and can only be applied in specific cases. This will lead to improvements in the clinical outcome in some cases, but for the foreseeable future, a number of these HIE have complex clinical presentations that still hold potential for serious outcome, including blindness or death, such as in the case of B. procyonis infections, where, despite treatment, neurological outcome is dismal in the overwhelming majority of documented cases [46]. However, many cases of these zoonotic helminth infections are preventable by relatively simple measures of improved health and sanitation conditions and awareness on the part of both public and health care providers. Risks for toxocariasis and baylisascariasis could be significantly reduced through better hygiene and reduction of the amount of animal waste in areas where people, especially children, might come in contact with it. For the foodborne zoonoses, such as angiostrongyliasis, gnathostomiasis and others, proper handling and preparation of foods would minimize the risk of infection. For the vector-borne zoonotic infections, control and prevention is likely going to be much harder, as it involves not only the control of the infection in the definitive animal host, but a concerted control of vectors, which is often outside the control of any individual but almost always done at the community or regional level.


  1. 1.

    Williams RA, Brody BL, Thomas RG, Kaplan RM, Brown SI: The psychosocial impact of macular degeneration. Arch Ophthalmol. 1998, 116: 514-520.

    CAS  PubMed  Article  Google Scholar 

  2. 2.

    Blindness and poverty. []

  3. 3.

    World Health Organization 2003. Onchocerciasis. []

  4. 4.

    Ryan E, Durrand M: Ocular Disease. Tropical Infectious Diseases Principals, Pathogens, and Practices. Edited by: Guerrant RL, WalkerDH, Weller PF. 2005, Philadelphia: Churchill Livingstone, 1554-1600. 2

    Google Scholar 

  5. 5.

    Cutler SJ, Fooks AR, van der Poel WH: Public health threat of new, reemerging, and neglected zoonoses in the industrialized world. Emerg Infect Dis. 2010, 16 (1): 1-7. 10.3201/eid1601.081467.

    PubMed Central  PubMed  Article  Google Scholar 

  6. 6.

    Jones KE, Patel NG, Levy MA, Storeygard A, Balk D, Gittleman JL, Daszak P: Global trends in emerging infections diseases. Nature. 2008, 451: 990-994. 10.1038/nature06536.

    CAS  PubMed  Article  Google Scholar 

  7. 7.

    Irwin PJ, Jefferies R: Arthropod-transmitted diseases of companion animals in Southeast Asia. Trends Parasitol. 2004, 1: 27-34. 10.1016/

    Article  Google Scholar 

  8. 8.

    Orihel TC, Eberhard ML: Zoonotic filariasis. Clin Microbiol Rev. 1998, 11: 366-381.

    PubMed Central  CAS  PubMed  Google Scholar 

  9. 9.

    Pisella PJ, Assaraf E, Rossaza C, Limon S, Baudouin C, Richard-Lenoble D: Conjunctivitis and ocular parasitic diseases. J Fr Ophtalmol. 1999, 22: 585-588.

    CAS  PubMed  Google Scholar 

  10. 10.

    Shen J, Gasser RB, Chu D, Wang Z, Yuan X, Cantacessi C, Otranto D: Human thelaziosis a neglected parasitic disease of the eye. J Parasitol. 2006, 92: 872-875. 10.1645/GE-823R.1.

    PubMed  Article  Google Scholar 

  11. 11.

    Rubinsky-Elefant G, Hirata CE, Yamamoto JH, Ferreira MU: Human toxocariasis: diagnosis, worldwide seroprevalences and clinical expression of the systemic and ocular forms. Ann Trop Med Parasitol. 2010, 104: 3-23. 10.1179/136485910X12607012373957.

    CAS  PubMed  Article  Google Scholar 

  12. 12.

    Cortez RT, Ramirez G, Collet L, Giuliari GP: Ocular parasitic diseases: a review on toxocariasis and diffuse unilateral subacute neuroretinitis. J Pediatr Ophthalmol Strabismus. 2010, 28: 1-9.

    Google Scholar 

  13. 13.

    Gottstein B, Pozio E, Nöckler K: Epidemiology, diagnosis, treatment, and control of trichinellosis. Clin Microbiol Rev. 2009, 22 (1): 127-145. 10.1128/CMR.00026-08.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  14. 14.

    Bruschi F, Murrell KD: Trichinellosis. Tropical Infectious Diseases Principals, Pathogens, and Practices. Edited by: Guerrant RL, WalkerDH, Weller PF. 2005, Philadelphia: Churchill Livingstone, 1225-1230. 2

    Google Scholar 

  15. 15.

    Kociecki J, Kociecka W: Visual system involvement in selected zoonotic diseases. II Trichinellosis. Klin Oczna. 2004, 106 (3): 371-375.

    PubMed  Google Scholar 

  16. 16.

    World Health Organization: Report of the WHO Informal Consultation on the use of chemotherapy for the control of morbidity due to soil-transmitted nematodes in humans Geneva. 1996

    Google Scholar 

  17. 17.

    O'Lorcain P, Holland CV: The public health importance of Ascaris lumbricoides. Parasitology. 2000, 121: S51-S71.

    PubMed  Article  Google Scholar 

  18. 18.

    Carden SM, Meusemann R, Walker J, Stawell RJ, MacKinnon JR, Smith D, Stawell AM, Hall AJ: Toxocara canis: egg presence in Melbourne parks and disease incidence in Victoria. Clin Experiment Ophthalmol. 2003, 31 (2): 143-146. 10.1046/j.1442-9071.2003.00622.x.

    PubMed  Article  Google Scholar 

  19. 19.

    Wilder HC: Nematode endophthalmitis. Trans Am Acad Ophthalmol and Otolaryngology. 1950, 55: 99-109.

    CAS  Google Scholar 

  20. 20.

    Brown DH: Ocular Toxocara canis. II. Clinical review. Journal of Pediatric Ophthalmology. 1970, 7: 182-191.

    Google Scholar 

  21. 21.

    Schantz P: Toxocara larva migrans now. Am J Trop Med Hyg. 1989, 41: 21-34.

    CAS  PubMed  Google Scholar 

  22. 22.

    Logar J, Soba B, Kraut A, Stirn-Kranjc B: Seroprevalence of Toxocara antibodies among patients suspected of ocular toxocariasis in Slovenia. Korean J Parasitol. 2004, 42 (3): 137-140. 10.3347/kjp.2004.42.3.137.

    PubMed Central  PubMed  Article  Google Scholar 

  23. 23.

    Won Y, Kruszon-Moran D, Schantz PM, Jones JL: National seroprevalence and risk factors for zoonotic Toxocara spp. infections. Am J Trop Med Hyg. 2008, 79 (4): 552-557.

    PubMed  Google Scholar 

  24. 24.

    Sabrosa NA, de Souza EC: Nematode infections of the eye: toxocariasis and diffuse unilateral subacute neuroretinitis. Curr Opin Ophthalmol. 2001, 12 (6): 450-454. 10.1097/00055735-200112000-00010.

    CAS  PubMed  Article  Google Scholar 

  25. 25.

    Moraes LR, Cialdini AP, Avila MP, Elsner AE: Identifying live nematodes in diffuse unilateral subacute neuroretinitis by using the scanning laser ophthalmoscope. Arch Ophthalmol. 2002, 120 (2): 135-138.

    PubMed  Article  Google Scholar 

  26. 26.

    Higashide T, Akao N, Shirao E, Shirao Y: Optical coherence tomographic and angiographic findings of a case with subretinal Toxocara granuloma. Am J Ophthalmol. 2003, 136 (1): 188-190. 10.1016/S0002-9394(03)00079-5.

    PubMed  Article  Google Scholar 

  27. 27.

    Altcheh J, Nallar M, Conca M, Biancardi M, Freilij H: Toxocariasis: clinical and laboratory features in 54 patients. An Pediatr (Barc). 2003, 58 (5): 425-431. 10.1157/13046522.

    CAS  Article  Google Scholar 

  28. 28.

    Yokoi K, Goto H, Sakai J, Usui M: Clinical features of ocular toxocariasis in Japan. Ocul Immunol Inflamm. 2003, 11 (4): 269-275. 10.1076/ocii.

    PubMed  Article  Google Scholar 

  29. 29.

    Garcia CA, Gomes AH, Garcia Filho CA, Vianna RN: Early-stage diffuse unilateral subacute neuroretinitis: improvement of vision after photocoagulation of the worm. Eye (Lond). 2004, 18 (6): 624-627. 10.1038/sj.eye.6700742.

    Article  Google Scholar 

  30. 30.

    Suzuki T, Joko T, Akao N, Ohashi Y: Following the migration of a Toxocara larva in the retina by optical coherence tomography and fluorescein angiography. Jpn J Ophthalmol. 2005, 49 (2): 159-161. 10.1007/s10384-004-0157-9.

    PubMed  Article  Google Scholar 

  31. 31.

    Stewart JM, Cubillan LD, Cunningham ET: Prevalence, clinical features, and causes of vision loss among patients with ocular toxocariasis. Retina. 2005, 25 (8): 1005-1013. 10.1097/00006982-200512000-00009.

    PubMed  Article  Google Scholar 

  32. 32.

    Rasquin F, Waterschoot MP, Termote H, Carlier Y: Diffuse unilateral subacute neuroretinitis in Africa. Ocul Immunol Inflamm. 2006, 14 (1): 59-62. 10.1080/09273940500224629.

    CAS  PubMed  Article  Google Scholar 

  33. 33.

    Bertelmann E, Velhagen KH, Pleyer U, Hartmann C: Ocular toxocariasis. Diagnostic and therapeutic options. Ophthalmologe. 2003, 100 (11): 950-954. 10.1007/s00347-003-0815-1.

    CAS  PubMed  Article  Google Scholar 

  34. 34.

    Mori K, Ohta K, Murata T: Vasoproliferative tumors of the retina secondary to ocular toxocariasis. Can J Ophthalmol. 2007, 42 (5): 758-759. 10.3129/i07-137.

    PubMed  Article  Google Scholar 

  35. 35.

    Acar N, Kapran Z, Utine CA, Büyükbabani N: Pars plana vitrectomy revealed Toxocara canis organism. Int Ophthalmol. 2007, 27 (4): 277-280. 10.1007/s10792-007-9078-1.

    PubMed  Article  Google Scholar 

  36. 36.

    Urban B, Bakunowicz-Łazarczyk A, Michał S: Clinical features, the effectiveness of treatment and function of vision organ in children and adolescents with ocular toxocariasis. Klin Oczna. 2008, 110 (10-12): 364-366.

    PubMed  Google Scholar 

  37. 37.

    Sivaratnam D, Subrayan V, Ali NA: Transvitreal migration of a Toxocara larva resulting in a second chorioretinal granuloma. Jpn J Ophthalmol. 2008, 52 (5): 416-417. 10.1007/s10384-008-0569-z.

    PubMed  Article  Google Scholar 

  38. 38.

    Bertelmann E, Velhagen KH, Pleyer U: Ocular toxocariasis. From biology to therapy. Ophthalmologie. 2007, 104: 35-39. 10.1007/s00347-006-1465-x.

    CAS  Article  Google Scholar 

  39. 39.

    de Visser L, Rothova A, de Boer JH, van Loon AM, Kerkhoff FT, Canninga-van Dijk MR, Weersink AY, de Groot-Mijnes JD: Diagnosis of ocular toxocariasis by establishing intraocular antibody production. Am J Ophthalmol. 2008, 145 (2): 369-374. 10.1016/j.ajo.2007.09.020.

    PubMed  Article  Google Scholar 

  40. 40.

    De A Garcia CA, Gomes AHB, de A Garcia Filho CA, Vianna RNG: Early-stage diffuse unilateral subacute neuroretinitis: imporvmenet of vision after photocoagulation of the worm. Eye. 2004, 18: 624-627. 10.1038/sj.eye.6700742.

    Article  Google Scholar 

  41. 41.

    Ash LR, Orihel TC, (Eds): Ash & Orihel's Atlas of Human Parasitology. 2007, American Society for Clinical Pathology, 5

  42. 42.

    Moertel CL, Kazacos KR, Butterfield JH, Kita H, Watterson J, Gleich GJ: Eosinophil-associated inflammation and elaboration of eosinophil-derived proteins in 2 children with raccoon roundworm (Baylisascaris procyonis) encephalitis. Pediatrics. 2001, 108: 93-10.1542/peds.108.5.e93.

    Article  Google Scholar 

  43. 43.

    Sorvillo F, Ash LR, Berlin OG, Morse SA: Baylisascaris procyonis: an emerging helminthic zoonosis. Emerg Infect Dis. 2002, 8 (4): 355-359. 10.3201/eid0804.010273.

    PubMed Central  PubMed  Article  Google Scholar 

  44. 44.

    Mets MB, Noble AG, Basti S, Gavin P, Davis AT, Shulman ST, Kazacos KR: Eye findings of diffuse unilateral subacute neuroretinitis and multiple choroidal infiltrates associated with neural larva migrans due to Baylisascaris procyonis. Am J Ophthalmol. 2003, 135 (6): 888-890. 10.1016/S0002-9394(02)01539-8.

    PubMed  Article  Google Scholar 

  45. 45.

    Sato H, Matsuo K, Osanai A, Kamiya H, Akao N, Owaki S, Furuoka H: Larva migrans by Baylisascaris transfuga: fatal neurological diseases in Mongolian jirds, but not in mice. J Parasitol. 2004, 90: 774-781. 10.1645/GE-3330.

    PubMed  Article  Google Scholar 

  46. 46.

    Gavin PJ, Kazacos KR, Shulman ST: Baylisascariasis. J Clin Microbiol. 2005, 18: 703-718. 10.1128/CMR.18.4.703-718.2005.

    Article  Google Scholar 

  47. 47.

    Goldberg MA, Kazacos KR, Boyce WM, Ai E, Katz B: Diffuse unilateral subacute neuritis. Morphometric, serologic and epidemiologic support for Baylisascaris as a causative agent. Ophthalmology. 1993, 100: 1695-1701.

    CAS  PubMed  Article  Google Scholar 

  48. 48.

    Brasil OF, Lewis H, Lowder CY: Migration of Baylisascaris procyonis into the vitreous. Br J Ophthalmol. 2006, 90: 1203-1204. 10.1136/bjo.2006.095323.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  49. 49.

    Herman JS, Chiodini PL: Gnathostomiasis, another emerging imported disease. Clin Microbiol Rev. 2009, 22 (3): 484-492. 10.1128/CMR.00003-09.

    PubMed Central  PubMed  Article  Google Scholar 

  50. 50.

    Biswas J, Gopal L, Sharma T, Badrinath SS: Intraocular Gnathostoma spinigerum. Clinicopathologic study of two cases with review of literature. Retina. 1994, 14 (5): 438-444. 10.1097/00006982-199414050-00009.

    CAS  PubMed  Article  Google Scholar 

  51. 51.

    Kannan KA, Vasantha K, Venugopal M: Intraocular gnathostomiasis. Indian J Ophthalmol. 1999, 47 (4): 252-253.

    CAS  PubMed  Google Scholar 

  52. 52.

    Qahtani F, Deschênes J, Ali-Khan Z, Maclean JD, Codère F, Mansour M, Burnier M: Intraocular gnathostomiasis: a rare Canadian case. Can J Ophthalmol. 2000, 35 (1): 35-39.

    CAS  PubMed  Article  Google Scholar 

  53. 53.

    Xuan le T, Rojekittikhun W, Punpoowong B, Trang le N, Hien TV: Case report: intraocular gnathostomiasis in Vietnam. Southeast Asian J Trop Med Public Health. 2002, 33 (3): 485-489.

    PubMed  Google Scholar 

  54. 54.

    Baquera-Heredia J, Cruz-Reyes A, Flores-Gaxiola A, López-Pulido G, Díaz-Simental E, Valderrama-Valenzuela L: Case report: Ocular gnathostomiasis in northwestern Mexico. Am J Trop Med Hyg. 2002, 66 (5): 572-574.

    PubMed  Google Scholar 

  55. 55.

    Basak SK, Sinha TK, Bhattacharya D, Hazra TK, Parikh S: Intravitreal live Gnathostoma spinigerum. Indian J Ophthalmol. 2004, 52 (1): 57-58.

    PubMed  Google Scholar 

  56. 56.

    Bhende M, Biswas J, Gopal L: Ultrasound biomicroscopy in the diagnosis and management of intraocular gnathostomiasis. Am J Ophthalmol. 2005, 140 (1): 140-142. 10.1016/j.ajo.2004.12.031.

    PubMed  Article  Google Scholar 

  57. 57.

    Barua P, Hazarika NK, Barua N, Barua CK, Choudhury B: Gnathostomiasis of the anterior chamber. Indian J Med Microbiol. 2007, 25 (3): 276-278. 10.4103/0255-0857.34775.

    CAS  PubMed  Article  Google Scholar 

  58. 58.

    Bhattacharjee H, Das D, Medhi J: Intravitreal gnathostomiasis and review of literature. Retina. 2007, 27 (1): 67-73. 10.1097/01.iae.0000224943.98423.e3.

    PubMed  Article  Google Scholar 

  59. 59.

    Chuenkongkaew W, Chirapapaisan N, Hongyok T, Yoolek A: Isolated optic neuritis from an identified Gnathostoma spinigerum. Eur J Ophthalmol. 2007, 17 (1): 130-132.

    CAS  PubMed  Google Scholar 

  60. 60.

    Anantaphruti MT, Nuamtanong S, Dekumyoy P: Diagnostic values of IgG4 in human gnathostomiasis. Trop Med Int Health. 2005, 10 (10): 1013-10.1111/j.1365-3156.2005.01478.x.

    CAS  PubMed  Article  Google Scholar 

  61. 61.

    Sawanyawisuth K, Chlebicki MP, Pratt E, Kanpittaya J, Intapan PM: Sequential imaging studies of cerebral gnathostomiasis with subdural hemorrhage as its complication. Trans R Soc Trop Med Hyg. 2009, 103 (1): 102-104. 10.1016/j.trstmh.2008.09.011.

    PubMed  Article  Google Scholar 

  62. 62.

    Botero D, Aguledo LM, Uribe FU, Esslinger JH, Beaver PC: Intracoular filarial, a Loaina species, from man in Colombia. Am J Trop Med Hyg. 1984, 33: 578-582.

    CAS  PubMed  Google Scholar 

  63. 63.

    Beaver PC: Intraocular filariasis: a brief review. Am J Trop Med Hyg. 1989, 40: 40-45.

    CAS  PubMed  Google Scholar 

  64. 64.

    Bain O, Otranto D, Diniz DG, Nascimento dos Santos J, Pinto de Oliveira N, Negrão Frota de Almeida I, Negrão Frota de Almeida R, Negrão Frota de Almeida L, Dantas-Torres F, Frota de Almeida , Sobrinho E: Human intraocular filariasis caused by Pelecitus sp., Brazil. Emerg Infect Dis. 2011,

    Google Scholar 

  65. 65.

    Otranto D, Diniz DG, Dantas-Torres F, Casiraghi M, de Almeida INF, de Almeida LNF, Nascimento dos Santos J, Penha Furtado A, de Almeida Sobrinho AF, Bain O: Human intraocular filariasis caused by Dirofilaria sp., Brazil. Emerg Infect Dis. 2011,

    Google Scholar 

  66. 66.

    Addario C: Su di un nematode dell'occhio umano. Ann Oftalmol. 1885, 14: 135-147.

    Google Scholar 

  67. 67.

    Orihel TC, Beaver PC: Morphology and relationship of Dirofilaria tenuis and Dirofilaria conjunctivae. Am J Trop Med Hyg. 1965, 14: 1030-1043.

    CAS  PubMed  Google Scholar 

  68. 68.

    Mittal M, Sathish KR, Bhatia G, Chidamber MS: Ocular dirofilariasis in Dubai, UAE. Indian J Ophthalmol. 2008, 56 (4): 325-326. 10.4103/0301-4738.41415.

    PubMed Central  PubMed  Article  Google Scholar 

  69. 69.

    Jamshidi A, Jamshidi M, Mobedi I, Khosroara M: Periocular Dirofilariasis in a Young Woman: A Case Report. Korean J Parasitol. 2008, 46 (4): 265-267. 10.3347/kjp.2008.46.4.265.

    PubMed Central  PubMed  Article  Google Scholar 

  70. 70.

    Eccher A, Dalfior D, Gobbo S, Martignoni G, Brunelli M, Decaminada W, Bonetti F, Rivasi F, Barbareschi M, Menestrina F: Periorbital subcutaneous tumor-like lesion due to Dirofilaria repens. Int J Surg Pathol. 2008, 16 (1): 101-103. 10.1177/1066896907307040.

    PubMed  Article  Google Scholar 

  71. 71.

    Fodor E, Fok E, Maka E, Lukáts O, Tóth J: Recently recognized cases of ophthalmofilariasis in Hungary. Eur J Ophthalmol. 2009, 19 (4): 675-678.

    PubMed  Google Scholar 

  72. 72.

    Khechine-Martinez R, Doucet C, Blanchard S, Rouland JF, Labalette P: Subconjunctival dirofilariasis: a case report. J Fr Ophtalmol. 2009, 32 (5): 350-10.1016/j.jfo.2009.04.003.

    PubMed  Article  Google Scholar 

  73. 73.

    Rohela M, Jamaiah I, Hui TT, Mak JW, Ithoi I, Amirah A: Dirofilaria causing eye infection in a patient from Malaysia. South Asian J Trop Med Public Health. 2009, 40 (5): 914-918.

    CAS  Google Scholar 

  74. 74.

    Wesolowska M, Kisza K, Szalinski M, Zielinski M, Okulewicz A, Misiuk-Hojlo M, Szostakowska B: First case of heterochthonous subconjunctival dirofilariasis described in Poland. Am J Trop Med Hyg. 2010, 83 (2): 210-10.4269/ajtmh.2010.10-0084.

    PubMed Central  PubMed  Article  Google Scholar 

  75. 75.

    Hasler S, Grimm F, Thiel MA, Müller NJ, Eberhard R, Bosch MM: Swiss patient with a subconjunctival Dirofilaria repens. Klin Monbl Augenheilkd. 2010, 227 (4): 332-333. 10.1055/s-0029-1245245.

    CAS  PubMed  Article  Google Scholar 

  76. 76.

    Avellis FO, Kramer LH, Mora P, Bartolino A, Benedetti P, Rivasi F: A Case of Human conjunctival dirofilariosis by Dirofilaria immitis in Italy. Vector Borne Zoonotic Dis. 2010,

  77. 77.

    Otranto D, Brianti E, Gaglio G, Dantas-Torres F, Azzaro S, Giannetto S: Human ocular infestation by Dirofilaria repens (Ralliet and Henry, 1911) in a canine dirofilariosis-endemic area. Am Jour Trop Med Hyg. 2011,

    Google Scholar 

  78. 78.

    Gungel H, Kara N, Pinarci EY, Albayrak S, Baylancicek DO, Uysal HK: An uncommon case with intravitreal worm. Intravitreal Dirofilaria infection. Br J Ophthalmol. 2009, 93 (5): 573-574. 10.1136/bjo.2008.138842.

    CAS  PubMed  Article  Google Scholar 

  79. 79.

    Huynh T, Thean J, Maini R: Dipetalonema reconditum in the human eye. Br J Ophthalmol. 2001, 85: 1391-10.1136/bjo.85.11.1384i.

    CAS  PubMed  Google Scholar 

  80. 80.

    Lau LI, Lee FL, Hsu WM, Pampiglione S, Fioravanti ML, Orihel TC: Human subconjunctival infection of Macacanema formosana: the first case of human infection reported worldwide. Arch Ophthalmol. 2002, 120 (5): 643-647.

    PubMed  Google Scholar 

  81. 81.

    Natarajan R: Another case of human subconjunctival infection by Macacanema formosana. Arch Ophthalmol. 2003, 121 (4): 584-585. 10.1001/archopht.121.4.584-b.

    PubMed  Article  Google Scholar 

  82. 82.

    Spratt DM: Natural occurrence, histopathology and developmental stages of Dirofilaria roemeri in the intermediate host. Int J Parasitol. 1972, 2: 202-208.

    Google Scholar 

  83. 83.

    Bartlett CM: Development of Dirofilaria scapiceps (Leidy, 1886) (Nematoda: Filarioidea) in Aedes spp. and Mansonia perturbans (Walker) and responses of mosquitoes to infection. Can J Zool. 1984, 62: 112-129. 10.1139/z84-019.

    Article  Google Scholar 

  84. 84.

    Bartlett CM, Anderson RC: Pelecitus fulicaeatrae (Nematoda: Filarioidea) of coots (Gruiformes) and grebes (Podicipediformes): skin-inhabiting microfilariae and development in Mallophaga. Can J Zool. 1987, 65: 2803-2812. 10.1139/z87-423.

    Article  Google Scholar 

  85. 85.

    Pinto RM, Vicente JJ, Noronha D: Nematode parasites of Brazilian psittacids birds, with emphasis on the genus Pelecitus Railiet & Henry, 1910. Mem Inst Oswaldo Cruz. 1993, 88: 279-284. 10.1590/S0074-02761993000200016.

    Article  Google Scholar 

  86. 86.

    Vicente JJ, Rodrigues HOR, Gomes DC, Pinto RM: Nematóides do Brasil. Parte V: nematóides de mamíferos. Rev Bras Zool. 1997, 14: 1-452. 10.1590/S0101-81751997000500001.

    Article  Google Scholar 

  87. 87.

    Rao NG, Mahapatra SK, Pattnayak S, Pattnaik K: Intravitreal live adult Brugian filariasis. Indian J Ophthalmol. 2008, 56 (1): 76-78. 10.4103/0301-4738.37610.

    PubMed Central  PubMed  Article  Google Scholar 

  88. 88.

    Heldwein K, Biedermann HG, Hamperl WD, Bretzel G, Löscher T, Laregina D, Frosch M, Büttner DW, Tappe D: Subcutaneous Taenia crassiceps infection in a patient with non-Hodgkin's lymphoma. Am J Trop Med Hyg. 2006, 75 (1): 108-111.

    PubMed  Google Scholar 

  89. 89.

    Maillard H, Marionneau J, Prophette B, Boyer E, Celerier P: Taenia crassiceps cysticercosis and AIDS. AIDS. 1998, 20: 1551-1552. 10.1097/00002030-199812000-00019.

    Article  Google Scholar 

  90. 90.

    Shea M, Maberley AL, Walters J, Freeman RS, Fallis AM: Intraocular Taenia crassiceps (Cestoda). Trans Am Acad Ophthalmol Otolaryngol. 1973, 77: OP778-OP783.

    CAS  PubMed  Google Scholar 

  91. 91.

    Chuck RS, Olk RJ, Weil GJ, Akduman L, Benenson IL, Smith ME, Kaplan HJ: Surgical removal of a subretinal proliferating cysticercus of Taeniaeformis crassiceps. Arch Ophthalmol. 1997, 115: 562-563.

    CAS  PubMed  Article  Google Scholar 

  92. 92.

    Arocker-Mettinger E, Huber-Spitzy V, Auer H, Grabner G, Stur M: Taenia crassiceps in the anterior chamber of the human eye. A case report. Klin Monbl Augenheilkd. 1992, 201: 34-37. 10.1055/s-2008-1045865.

    CAS  PubMed  Article  Google Scholar 

  93. 93.

    Sinha S, Sharma BS: Neurocysticercosis: a review of current status and management. J Clin Neurosci. 2009, 16 (7): 867-876. 10.1016/j.jocn.2008.10.030.

    PubMed  Article  Google Scholar 

  94. 94.

    Gauci C, Vural G, Oncel T, Varcasia A, Damian V, Kyngdon CT, Craig PS, Anderson GA, Lightowlers MW: Vaccination with recombinant oncosphere antigens reduces the susceptibility of sheep to infection with Taenia multiceps. Int J Parasitol. 2008, 38: 1041-1050. 10.1016/j.ijpara.2007.11.006.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  95. 95.

    Sabattani S, Marliani A, Roncaroli F, Zuccheli M, Zini A, Calbucci F, Chiodo F: Cerebral coenurosis. J Neurosurg. 2004, 100: 964-10.3171/jns.2004.100.5.0964.

    PubMed  Article  Google Scholar 

  96. 96.

    Williams PH, Templeton AC: Infection of the eye by tapeworm Coenurus. Brit J Ophthal. 1971, 55: 766-769. 10.1136/bjo.55.11.766.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  97. 97.

    Manschot WA: Coenurus infestation of eye and orbit. Arch Ophthalmol. 1976, 94: 961-964.

    CAS  PubMed  Article  Google Scholar 

  98. 98.

    Melhorn H: Encyclopedia of Parasitology. 2008, Heidelberg: Springer-Verlag

    Book  Google Scholar 

  99. 99.

    Ou Q, Li SJ, Cheng XJ: Cerebral sparganosis: A case report. Biosci Trends. 2010, 4 (3): 145-147.

    CAS  PubMed  Google Scholar 

  100. 100.

    Leon LA, Almeida R, Mueller JF: A case of ocular sparganosis in Ecuador. J Parasitol. 1972, 58 (1): 184-185. 10.2307/3278274.

    CAS  PubMed  Article  Google Scholar 

  101. 101.

    Rehák M, Kolárová L, Kohnová I, Rehák J, Mohlerová S, Fric E, Chrapek O: Ocular sparganosis in the Czech Republic-a case report. Klin Mikrobiol Infekc Lek. 2006, 12: 161-165.

    PubMed  Google Scholar 

  102. 102.

    Zhong HL, Shao L, Lian DR, Deng ZF, Zhao SX, Gao PZ, He LY, Yun CF, Pan JY: Ocular sparganosis caused blindness. Chin Med J. 1983, 96 (1): 73-75.

    CAS  PubMed  Google Scholar 

  103. 103.

    Wiwanitkit VA: Review of human sparganosis in Thailand. Int J Infect Dis. 2005, 9 (6): 312-316. 10.1016/j.ijid.2004.08.003.

    PubMed  Article  Google Scholar 

  104. 104.

    Subudhi BN, Dash S, Chakrabarty D, Mishra DP, Senapati U: Ocular sparganosis. J Indian Med Assoc. 2006, 104: 529-530.

    CAS  PubMed  Google Scholar 

  105. 105.

    Yang JW, Lee JH, Kang MS: A case of ocular sparganosis in Korea. Korean J Ophthalmol. 2007, 21 (1): 48-50. 10.3341/kjo.2007.21.1.48.

    PubMed Central  PubMed  Article  Google Scholar 

  106. 106.

    Kittiponghansa S, Tesana S, Ritch R: Ocular sparganosis: a cause of subconjunctival tumor and deafness. Trop Med Parasitol. 1988, 39 (3): 247-248.

    CAS  PubMed  Google Scholar 

  107. 107.

    Sen DK, Muller R, Gupta VP, Chilana JS: Cestode Larva (Sparganum) in the anterior chamber of the eye. Trop Geogr Med. 1989, 41: 270-273.

    CAS  PubMed  Google Scholar 

  108. 108.

    Basset D, Girou C, Nozais IP, D'Hermies F, Hoang C, Gordon R, D'Alessandro A: Neotropical echinococcosis in Suriname: Echinococcus oligarthrus in the orbit and Echinococcus vogeli in the abdomen. Am J Trop Med Hyg. 1998, 59: 787-790.

    CAS  PubMed  Google Scholar 

  109. 109.

    Murthy R, Honavar SG, Vemuganti GK, Naik M, Burman S: Polycystic echinococcosis of the orbit. Am J Ophthalmol. 2005, 140: 561-563. 10.1016/j.ajo.2005.03.048.

    PubMed  Article  Google Scholar 

  110. 110.

    Lightowlers MW: Cestode vaccines: origins, current status and future prospects. Parasitology. 2006, 133: S27-S42. 10.1017/S003118200600179X.

    PubMed  Article  Google Scholar 

  111. 111.

    Kern P: Clinical features and treatment of alveolar echinococcosis. Curr Opin Infect Dis. 2010, 23: 505-412. 10.1097/QCO.0b013e32833d7516.

    PubMed  Article  Google Scholar 

  112. 112.

    Chaabouni M, Ben Zina Z, Ben Ayez H, Tounsi R, Trigui A, Ben Mansour H: Hydatid orbital cyst: a unique intra-orbital locality. A case report. J Fr Ophtalmol. 1999, 22: 329-334.

    CAS  PubMed  Google Scholar 

  113. 113.

    Sinav S, Demirci A, Sinav B, Oge F, Sullu Y, Kandemir B: A primary intraocular hydatid cyst. Acta Ophthalmologica. 1991, 69: 802-804. 10.1111/j.1755-3768.1991.tb02065.x.

    CAS  PubMed  Article  Google Scholar 

  114. 114.

    Ozek MM, Pamir MN, Sav A: Spontaneous rupture of an intraorbital hydatid cyst. A rare cause of acute visual loss. J Clin Neuroophthalmol. 1993, 13: 135-137.

    CAS  PubMed  Article  Google Scholar 

  115. 115.

    Williams DF, Williams GA, Caya JG, Werner RP, Harrison TJ: Intraocular Echinococcus multilocularis. Arch Ophthalmol. 1987, 105: 1106-1109.

    CAS  PubMed  Article  Google Scholar 

  116. 116.

    D'Alessandro A, Rausch RL: New aspects of neotropical polycystic (Echinococcus vogeli) and unicystic (Echinococcus oligarthrus) echinococcosis. Clin Microbiol Rev. 2008, 21: 380-401.

    PubMed Central  PubMed  Article  Google Scholar 

  117. 117.

    Lopera RD, Melendez RD, Fernandez I, Sirit J, Perera MP: Orbital hydatid cyst of Echinococcus oligarthrus in a human in Venezuela. J Parasitol. 1989, 75: 467-470. 10.2307/3282609.

    CAS  PubMed  Article  Google Scholar 

  118. 118.

    Möhl K, Grosse K, Hamedy A, Wüste T, Kabelitz P, Lücker E: Biology of Alaria spp. and human exposition risk to Alaria mesocercariae-a review. Parasitol Res. 2009, 105 (1): 1-15.

    PubMed  Article  Google Scholar 

  119. 119.

    Mas-Coma S, Valero MA, Bargues MD: Fasciola, lymnaeids and human fascioliasis, with a global overview on disease transmission, epidemiology, evolutionary genetics, molecular epidemiology and control. Adv Parasitol. 2009, 69: 41-146. full_text.

    PubMed  Article  Google Scholar 

  120. 120.

    Dalimi A, Jabarvand M: Fasciola hepatica in the human eye. Trans R Soc Trop Med Hyg. 2005, 99: 798-800. 10.1016/j.trstmh.2005.05.009.

    PubMed  Article  Google Scholar 

  121. 121.

    Nithiuthai S, Anantaphruti MT, Waikagul J, Gajadhar A: Waterborne zoonotic helminthiases. Vet Parasitol. 2004, 126 (1-2): 167-193. 10.1016/j.vetpar.2004.09.018.

    PubMed  Article  Google Scholar 

  122. 122.

    McDonald HR, Kazacos KR, Schatz H, Johnson RN: Two cases of intraocular infection with Alaria mesocercaria (Trematoda). Am J Ophthalmol. 1994, 117: 447-455.

    CAS  PubMed  Article  Google Scholar 

  123. 123.

    Diaz MT, Hernandez LF, Bashirullah AK: Experimental life cycle of Philophthalmus gralli (Trematoda: Philophthalmidae) in Venezuela. Rev Biol Trop. 2002, 50: 629-641.

    PubMed  Google Scholar 

  124. 124.

    Waikagul J, Dekumyoy P, Yoonuan T, Praevanit R: Conjunctiva philophthalmosis: a case report in Thailand. Am J Trop Med Hyg. 2006, 74 (5): 848-849.

    PubMed  Google Scholar 

  125. 125.

    Anderson RC: Nematode Parasites of Vertebrates. Their development and transmission. 2000, Guilford: CABI, 2

    Book  Google Scholar 

  126. 126.

    Lindo JF, Waugh C, Hall J, Cunningham-Myrie C, Ashley D, Eberhard ML, Sullivan JJ, Bishop HS, Robinson DG, Holtz T, Robinson RD: Enzootic Angiostrongylus cantonensis in rats and snails after an outbreak of human eosinophilic meningitis, Jamaica. Emerg Infect Dis. 2002, 8 (3): 324-326. 10.3201/eid0803.010316.

    PubMed Central  PubMed  Article  Google Scholar 

  127. 127.

    Caldeira RL, Mendonça CL, Goveia CO, Lenzi HL, Graeff-Teixeira C, Lima WS, Mota EM, Pecora IL, Medeiros AM, Carvalho Odos S: First record of molluscs naturally infected with Angiostrongylus cantonensis (Chen, 1935) (Nematoda: Metastrongylidae) in Brazil. Mem Inst Oswaldo Cruz. 2007, 102 (7): 887-889. 10.1590/S0074-02762007000700018.

    CAS  PubMed  Article  Google Scholar 

  128. 128.

    Lima AR, Mesquita SD, Santos SS, Aquino ER, Rosa Lda R, Duarte FS, Teixeira AO, Costa ZR, Ferreira ML: Alicata disease: neuroinfestation by Angiostrongylus cantonensis in Recife, Pernambuco, Brazil. Arq Neuropsiquiatr. 2009, 67 (4): 1093-1096.

    PubMed  Article  Google Scholar 

  129. 129.

    Dorta-Contreras AJ, Magraner-Tarrau ME, Sánchez-Zulueta E: Angiostrongyliasis in the Americas. Emerg Infect Dis. 2009, 15 (6): 991-10.3201/eid1506.071708.

    PubMed Central  PubMed  Article  Google Scholar 

  130. 130.

    Chikweto A, Bhaiyat MI, Macpherson CN, Deallie C, Pinckney RD, Richards C, Sharma RN: Existence of Angiostrongylus cantonensis in rats (Rattus norvegicus) in Grenada, West Indies. Vet Parasitol. 2009, 162 (1-2): 160-162. 10.1016/j.vetpar.2009.02.020.

    CAS  PubMed  Article  Google Scholar 

  131. 131.

    Punyagupta S, Juttijudata P, Bunnag T: Eosinophilic meningitis in Thailand. Clinical studies of 484 typical cases probably caused by Angiostrongylus cantonensis. Am J Trop Med Hyg. 1975, 24: 921-31.

    CAS  PubMed  Google Scholar 

  132. 132.

    Prommindaroj K, Leelawongs N, Pradatsundarasar A: Human angiostrongyliasis of the eye in Bangkok. Am J Trop Med Hyg. 1962, 11 (6): 759-761.

    Google Scholar 

  133. 133.

    Kumar V, Kyprianou I, Keenan JM: Ocular angiostrongyliasis: removal of a live nematode from the anterior chamber. Eye. 2005, 19: 229-230. 10.1038/sj.eye.6701442.

    CAS  PubMed  Article  Google Scholar 

  134. 134.

    Jindrak K: Angiostrongyliasis cantonensis (eosinophilic meningitis, Alicata's disease). Contemp Neurol Ser. 1975, 12: 133-166.

    CAS  PubMed  Google Scholar 

  135. 135.

    Dissanaike AS, Ihalamulla RL, Naotunne TS, Senarathna T, Withana DS: Third report of ocular parastrongyliasis (angiostrongyliasis) from Sri Lanka. Parassitologia. 2001, 43 (3): 95-97.

    CAS  PubMed  Google Scholar 

  136. 136.

    Thu TP, Nguyen NX, Lan le T, Küchle M: Ocular Angiostrongylus cantonensis in a female Vietnamese patient: case report. Klin Monbl Augenheilkd. 2002, 219 (12): 892-895. 10.1055/s-2002-36945.

    PubMed  Article  Google Scholar 

  137. 137.

    Patikulsila D, Ittipunkul N, Theerakittikul B: Intravitreal angiostrongyliasis: report of 2 cases. J Med Assoc Thai. 2003, 86 (10): 981-985.

    PubMed  Google Scholar 

  138. 138.

    Dissanaike AS, Cross JH: Ocular parastrongyliasis (= angiostrongyliasis): probable first report of human infection from a patient in Ceylon (Sri Lanka). Parassitologia. 2004, 46 (3): 315-316.

    CAS  PubMed  Google Scholar 

  139. 139.

    Malhotra S, Mehta DK, Arora R, Chauhan D, Ray S, Jain M: Ocular angiostrongyliasis in a child-first case report from India. J Trop Pediatr. 2006, 52 (3): 223-225. 10.1093/tropej/fmi092.

    CAS  PubMed  Article  Google Scholar 

  140. 140.

    Liu IH, Chung YM, Chen SJ, Cho WL: Necrotizing retinitis induced by Angiostrongylus cantonensis. Am J Ophthalmol. 2006, 141 (3): 577-579. 10.1016/j.ajo.2005.09.033.

    PubMed  Article  Google Scholar 

  141. 141.

    Sawanyawisuth K, Kitthaweesin K, Limpawattana P, Intapan PM, Tiamkao S, Jitpimolmard S, Chotmongkol V: Intraocular angiostrongyliasis: clinical findings, treatments and outcomes. Trans R Soc Trop Med Hyg. 2007, 101: 497-501. 10.1016/j.trstmh.2006.07.010.

    CAS  PubMed  Article  Google Scholar 

  142. 142.

    Baheti NN, Sreedharan M, Krishnamoorthy T, Nair MD, Radhakrishnan K: Eosinophilic meningitis and an ocular worm in a patient from Kerala, south India. J Neurol Neurosurg Psychiatry. 2008, 79: 271-10.1136/jnnp.2007.122093.

    CAS  PubMed  Article  Google Scholar 

  143. 143.

    Sawanyawisuth K, Kitthaweesin K: Optic neuritis caused by intraocular angiostrongyliasis. Southeast Asian J Trop Med Public Health. 2008, 39: 1005-1007.

    PubMed  Google Scholar 

  144. 144.

    Mattis A, Mowatt L, Lue A, Lindo J, Vaughan H: Ocular angiostrongyliasis-first case report from Jamaica. West Indian Med J. 2009, 58: 383-385.

    CAS  PubMed  Google Scholar 

  145. 145.

    Ramirez-Avila L: Eosinophilic Meningitis due to Angiostrongylus and Gnathostoma Species. Emerging Infections. 2009, 48: 322-327.

    Google Scholar 

  146. 146.

    Koo J, Pien F, Kliks MM: Angiostrongylus (Parastrongylus) eosinophilic meningitis. Rev Infect Dis. 1988, 10: 1155-1162. 10.1093/clinids/10.6.1155.

    CAS  PubMed  Article  Google Scholar 

  147. 147.

    Sawanyawisuth K, Sawanyawisuth K: Treatment of angiostrongyliasis. Trans R Soc Trop Med Hyg. 2008, 102: 990-996. 10.1016/j.trstmh.2008.04.021.

    PubMed  Article  Google Scholar 

  148. 148.

    Doezie AM, Lucius RW, Aldeen W, Hale DV, Smith DR, Mamalis N: Thelazia californiensis conjunctival infestation. Ophthalmic Surgery and Lasers. 1996, 27: 716-771.

    CAS  PubMed  Google Scholar 

  149. 149.

    Otranto D, Traversa D: Thelazia eyeworm: an original endo- and ecto-parasitic nematode. Trends Parasitol. 2005, 21: 1-4. 10.1016/

    PubMed  Article  Google Scholar 

  150. 150.

    Otranto D, Dantas-Torres F, Mallia E, DiGeronimo PM, Brianti E, Testini G, Traversa D, Lia P: Thelazia callipaeda (Spirurida, Thelaziidae) in wild animals: report of new host species and ecological implications. Vet Parasitol. 2009, 166: 262-267. 10.1016/j.vetpar.2009.08.027.

    PubMed  Article  Google Scholar 

  151. 151.

    Miroshnichenko VA, Desiaterik MP, Novik AP, Gorbach AP, Papernova N: A case of ocular thelaziasis in a 3-year-old child in Russian. Vestn Oftalmol. 1988, 104-64.

    Google Scholar 

  152. 152.

    Min S, Jae RY, Hyun YP: Enzooticity of the dogs, the reservoir host of Thelazia callipaeda in Korea. Kor J Parasitol. 2002, 40: 101-103. 10.3347/kjp.2002.40.2.101.

    Article  Google Scholar 

  153. 153.

    Koyama Y, Ohira A, Kono T, Yoneyama T, Shiwaku K: Five cases of thelaziasis. Br J Ophthalmol. 2000, 84 (4): 441-10.1136/bjo.84.4.439c.

    CAS  PubMed  Google Scholar 

  154. 154.

    Kosin E, Kosman ML, Depary AA: First case of human Thelaziasis in Indonesia. Southeast Asian. J Trop Med Public Health. 1989, 20: 233-236.

    CAS  Google Scholar 

  155. 155.

    Yospaiboon Y, Sithithavorn P, Maleewong V, Ukosanakarn U, Bhaibulaya M: Ocular thelaziasis in Thailand: a case report. J Med Assoc Thai. 1989, 72: 469-473.

    CAS  PubMed  Google Scholar 

  156. 156.

    Cheung WK, Lu HL, Liang CH, Peng ML, Lee HH: Conjunctivitis caused by Thelazia callipaeda infestation in a woman. J Formos Med Assoc. 1998, 97: 425-427.

    CAS  PubMed  Google Scholar 

  157. 157.

    Singh TS, Singh KN: Thelaziasis: report of two cases. Br J Ophthalmol. 1993, 77: 528-10.1136/bjo.77.8.528.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  158. 158.

    Otranto D, Lia RP, Cantacessi C, Testini G, Troccoli A, Shen JL, Wang ZX: Nematode biology and larval development of Thelazia callipaeda (Spirurida, Thelaziidae) in the drosophilid intermediate host in Europe and China. Parasitology. 2005, 131: 847-855. 10.1017/S0031182005008395.

    CAS  PubMed  Article  Google Scholar 

  159. 159.

    Otranto D, Cantacessi C, Testini G, Lia RP: Phortica variegata is an intermediate host of Thelazia callipaeda under natural conditions: evidence for pathogen transmission by a male arthropod vector. Int J Parasitol. 2006, 36 (10-11): 1167-1173. 10.1016/j.ijpara.2006.06.006.

    CAS  PubMed  Article  Google Scholar 

  160. 160.

    Otranto D, Ferroglio E, Lia RP, Traversa D, Rossi L: Current status and epidemiological observation of Thelazia callipaeda (Spirurida, Thelaziidae) in dogs, cats and foxes in Italy: a "coincidence" or a parasitic disease of the Old Continent?. Vet Parasitol. 2003, 116: 315-325. 10.1016/j.vetpar.2003.07.022.

    PubMed  Article  Google Scholar 

  161. 161.

    Dorchies P, Chaudieu G, Siméon LA, Cazalot G, Cantacessi C, Otranto D: First reports of autochthonous eyeworm infection by Thelazia callipaeda (Spirurida, Thelaziidae) in dogs and cat from France. Vet Parasitol. 2007, 149: 294-297. 10.1016/j.vetpar.2007.08.005.

    PubMed  Article  Google Scholar 

  162. 162.

    Ruytoor P, Déan E, Pennant O, Dorchies P, Chermette R, Otranto D, Guillot J: Ocular thelaziosis in dogs. Emerg Infect Dis. 2010, 16: 1943-1945.

    PubMed Central  PubMed  Article  Google Scholar 

  163. 163.

    Malacrida F, Hegglin D, Bacciarini L, Otranto D, Nägeli F, Nägeli C, Bernasconi C, Scheu U, Balli A, Marengo M, Togni L, Deplazes P, Schnyder M: Emergence of canine ocular thelaziosis caused by Thelazia callipaeda in southern Switzerland. Vet Parasitol. 2008, 157: 321-327. 10.1016/j.vetpar.2008.07.029.

    CAS  PubMed  Article  Google Scholar 

  164. 164.

    Otranto D, Dutto M: Human thelaziasis, Europe. Emerg Infect Dis. 2008, 4: 647-649. 10.3201/eid1404.071205.

    Article  Google Scholar 

  165. 165.

    Otranto D, Lia RP, Traversa D, Giannetto S: Thelazia callipaeda (Spirurida, Thelaziidae) of carnivores and humans: morphological study by light and scanning electron microscopy. Parassitologia. 2003, 45: 125-133.

    CAS  PubMed  Google Scholar 

  166. 166.

    Sréter T, Széll Z: Onchocercosis: a newly recognized disease in dogs. Vet Parasitol. 2008, 151: 1-13.

    PubMed  Article  Google Scholar 

  167. 167.

    Uni S, Boda T, Daisaku K, Ikura Y, Maruyama H, Hasegawa H, Fukuda M, Takaoka H, Bain O: Zoonotic filariasis caused by Onchocerca dewittei japonica in a resident of Hiroshima Prefecture, Honshu, Japan. Parasitol Int. 2010, 59: 477-80. 10.1016/j.parint.2010.05.006.

    PubMed  Article  Google Scholar 

  168. 168.

    Azarova NS, Miretski OI, Sonin MD: 1st case of human infection by the nematode Onchocerca Diesing, 1841 in the USSR. Med Parazitol. 1965, 34 (2): 156-158.

    CAS  Google Scholar 

  169. 169.

    Burr WE, Brown MF, Eberhard ML: Zoonotic Onchocerca (Nematoda:Filarioidea) in the cornea of a Colorado resident. Ophthalmology. 1998, 105: 1494-1497. 10.1016/S0161-6420(98)98035-6.

    PubMed  Article  Google Scholar 

  170. 170.

    Pampiglione S, Vakalis N, Lyssimachou A, Kouppari G, Orihel TC: Subconjunctival zoonotic Onchocerca in an Albanian man. Ann Trop Med Parasitol. 2001, 95: 827-832. 10.1080/00034980120111163.

    CAS  PubMed  Article  Google Scholar 

  171. 171.

    Sallo F, Eberhard ML, Fok E, Baska F, Hatvani I: Zoonotic intravitreal Onchocerca in Hungary. Ophthalmology. 2005, 112 (3): 502-504. 10.1016/j.ophtha.2004.10.036.

    PubMed  Article  Google Scholar 

  172. 172.

    Otranto D, Sakru N, Testini G, Gürlü VP, Yakar K, Lia RP, Dantas-Torres F, Bain O: First Evidence of Human Zoonotic Infestation by Onchocerca lupi (Spirurida, Onchocercidae). Am J Trop Med Hyg. 2011, 84 (1): 55-8. 10.4269/ajtmh.2011.10-0465.

    PubMed Central  PubMed  Article  Google Scholar 

  173. 173.

    Beaver PC, Horner GS, Bilos JZ: Zoonotic onchocercosis in a resident of Illinois and observations on the identification of Onchocerca species. Am J Trop Med Hyg. 1974, 23 (4): 595-607.

    CAS  PubMed  Google Scholar 

  174. 174.

    Széll Z, Erdélyi I, Sréter T, Albert M, Varga I: Canine ocular onchocercosis in Hungary. Vet Parasitol. 2001, 97: 245-251.

    Article  Google Scholar 

  175. 175.

    Komnenou A, Eberhard ML, Kaldrymidou E, Tsalie E, Dessiris A: Subconjunctival filariasis due to Onchocerca sp. in dogs: report of 23 cases in Greece. Vet Ophthalmol. 2002, 5: 119-126. 10.1046/j.1463-5224.2002.00235.x.

    PubMed  Article  Google Scholar 

  176. 176.

    Hermosilla A, Hetzel U, Bausch M, Grübl J, Bauer C: First autochthonous case of canine ocular onchocercosis in Germany. Vet Rec. 2005, 154: 450-452.

    Article  Google Scholar 

  177. 177.

    Sréter-Lancz Z, Széll Z, Sréter T: Molecular genetic comparison of Onchocerca sp. infecting dogs in Europe with other spirurid nematodes including Onchocerca lienalis. Vet Parasitol. 2007, 148: 365-370.

    PubMed  Article  CAS  Google Scholar 

  178. 178.

    Orihel TC, Ash LR, Holshuh HJ, Santenelli S: Onchocerciasis in a California dog. Am J Trop Med Hyg. 1991, 44: 513-517.

    CAS  PubMed  Google Scholar 

  179. 179.

    Eberhard ML, Ortega Y, Dial S, Schiller CA, Sears W, Greiner E: Ocular Onchocerca infections in western United States. Vet Parasitol. 2000, 90: 333-338. 10.1016/S0304-4017(00)00252-1.

    CAS  PubMed  Article  Google Scholar 

  180. 180.

    Zarfoss MK, Dubielzig RR, Eberhard ML, Schmidt KS: Canine ocular onchocerciasis in the United States: two new cases and a review of the literature. Vet Ophthalmol. 2005, 8: 51-57. 10.1111/j.1463-5224.2005.00348.x.

    PubMed  Article  Google Scholar 

  181. 181.

    Pampiglione S, Rivasi F, Gustinelli A: Dirofilarial human cases in the Old World, attributed to Dirofilaria immitis: a critical analysis. Histopathology. 2009, 54 (2): 192-204. 10.1111/j.1365-2559.2008.03197_a.x.

    PubMed  Article  Google Scholar 

  182. 182.

    Kim HW, Kim JL, Kho WG, Hwang SY, Yun IH: Intraocular infestation with Thelazia callipaeda. Jpn J Ophthalmol. 2010, 54 (4): 370-372. 10.1007/s10384-010-0822-0.

    CAS  PubMed  Article  Google Scholar 

  183. 183.

    Zakir R, Zhong-Xia Z, Chiodini P, Canning CR: Intraocular infestation with the worm, Thelazia callipaeda. Br J Ophthalmol. 1999, 83: 1194-1195. 10.1136/bjo.83.10.1194a.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  184. 184.

    Ferri E, Barbuto M, Bain O, Galimberti A, Uni S, Guerrero R, Ferté H, Bandi C, Martin C, Casiraghi M: Integrated taxonomy: traditional approach and DNA barcoding for the identification of filarioid worms and related parasites (Nematoda). Front Zool. 2009, 6: 1-10.1186/1742-9994-6-1.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  185. 185.

    Bimi L, Freeman AR, Eberhard ML, Ruiz-Tiben E, Pieniazek NJ: Differentiating Dracunculus medinensis from D. insignis, by the sequence analysis of the 18S rRNA gene. Ann Trop Med Parasitol. 2005, 99 (5): 511-517. 10.1179/136485905X51355.

    CAS  PubMed  Article  Google Scholar 

  186. 186.

    Barlow J, Gardner TA, Araujo IS, Avila-Pires TC, Bonaldo AB, Costa JE, Esposito MC, Ferreira LV, Hawes J, Hernandez MI, Hoogmoed MS, Leite RN, Lo-Man-Hung NF, Malcolm JR, Martins MB, Mestre LA, Miranda-Santos R, Nunes-Gutjahr AL, Overal WL, Parry L, Peters SL, Ribeiro-Junior MA, da Silva MN, da Silva Motta C, Peres CA: Quantifying the biodiversity value of tropical primary, secondary, and plantation forests. Proc Natl Acad Sci USA. 2007, 104: 18555-18560. 10.1073/pnas.0703333104.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

  187. 187.

    Foster GW, Cunningham MW, Kinsella JM, Forrester DJ: Parasitic helminths of black bear cubs (Ursus americanus) from Florida. J Parasitol. 2004, 90 (1): 173-175. 10.1645/GE-127R.

    PubMed  Article  Google Scholar 

  188. 188.

    Papini R, Casarosa L: Observations on the infectivity of Baylisascaris transfuga eggs for mice. Vet Parasitol. 1994, 51 (3-4): 283-288. 10.1016/0304-4017(94)90166-X.

    CAS  PubMed  Article  Google Scholar 

  189. 189.

    Papini R, Renzoni G, Malloggi M, Casarosa L: Visceral larva migrans in mice experimentally infected with Baylisascaris transfuga (Ascarididae: Nematoda). Parassitologia. 1994, 36 (3): 321-329.

    CAS  PubMed  Google Scholar 

  190. 190.

    Papini R, Renzoni G, Lo Piccolo S, Casarosa L: Ocular larva migrans and histopathological lesions in mice experimentally infected with Baylisascaris transfuga embryonated eggs. Vet Parasitol. 1996, 61 (3-4): 315-320. 10.1016/0304-4017(95)00825-X.

    CAS  PubMed  Article  Google Scholar 

  191. 191.

    Cho S, Egami M, Ohnuki H, Saito Y, Chinone S, Shichinohe K, Suganuma M, Akao N: Migration behaviour and pathogenesis of five ascarid nematode species in the Mongolian gerbil Meriones unguiculatus. J Helminthol. 2007, 81 (1): 43-47. 10.1017/S0022149X07212118.

    CAS  PubMed  Article  Google Scholar 

  192. 192.

    Matoff K, Komandarev S: Comparative studies on the migration of the larvae of Toxascaris leonina and Toxascaris transfuga. Z Parasitenkd. 1965, 25: 538-555. 10.1007/BF00259600.

    Article  Google Scholar 

  193. 193.

    Sato H, Une Y, Kawakami S, Saito E, Kamiya H, Akao N, Furuoka H: Fatal Baylisascaris larva migrans in a colony of Japanese macaques kept by a safari-style zoo in Japan. J Parasitol. 2005, 91 (3): 716-719. 10.1645/GE-3374RN.

    CAS  PubMed  Article  Google Scholar 

  194. 194.

    Sprent JF: Notes on Ascaris and Toxascaris, with a definition of Baylisascaris gen. nov. Parasitology. 1968, 58 (1): 185-198. 10.1017/S0031182000073534.

    CAS  PubMed  Article  Google Scholar 

  195. 195.

    Eberhard ML, Nace EK, Won KY, Punkosdy GA, Bishop HS, Johnston SP: Baylisascaris procyonis in the metropolitan Atlanta area. Emerg Infect Dis. 2003, 9 (12): 1636-1637.

    PubMed Central  PubMed  Article  Google Scholar 

  196. 196.

    Blizzard EL, Yabasley MJ, Beck MF, Harsch S: Geographic expansion of Baylisascaris procyonis roundworms, Florida, USA. Emerg Inf Dis. 2010, 16 (11): 1803-1804.

    Article  Google Scholar 

  197. 197.

    Pampiglione S, Canestri Trotti G, Rivasi F: Human dirofilariasis due to Dirofilaria (Nochtiella) repens in Italy: a review of word literature. Parassitologia. 1995, 37: 149-193.

    CAS  PubMed  Google Scholar 

  198. 198.

    Beaver PC, Meyer EA, Jarroll EL, Rosenquist RC: Dipetalonema from the eye of a man in Oregon, U.S.A. A case report. Am J Trop Med Hyg. 1980, 29: 369-372.

    CAS  PubMed  Google Scholar 

  199. 199.

    Koehsler M, Soleiman A, Aspöck H, Auer H, Walochnik J: Onchocerca jakutensis filariasis in humans. Emerg Infect Dis. 2007, 13: 1749-1752.

    PubMed Central  CAS  PubMed  Article  Google Scholar 

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The authors are grateful to Alessio Giannelli (University of Bari, Italy) and Donato Traversa (University of Teramo, Italy), for their support during the preparation of the manuscript. We are grateful to Emanuele Brianti (University of Messina, Italy) and Riccardo Paolo Lia (University of Bari, Italy) for their assistance with the video editing and figure preparation. The authors thank Susan Montgomery, CDC, and of two anonymous reviewers for their helpful suggestions. DO and MLE would like to thank their wives Irene Canfora and Sandra Eberhard for their continuous support and assistance during the preparation of this article.

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DO and MLE equally contributed in writing the article.

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Otranto, D., Eberhard, M.L. Zoonotic helminths affecting the human eye. Parasites Vectors 4, 41 (2011).

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  • Optic Neuritis
  • Definitive Host
  • Echinococcus
  • Zoonotic Infection
  • Fascioliasis