ﺑﺎﺯﮔﺸﺖ ﺑﻪ ﺻﻔﺤﻪ ﻗﺒﻠﯽ
خرید پکیج
تعداد آیتم قابل مشاهده باقیمانده : 3 مورد
نسخه الکترونیک
medimedia.ir

Miscellaneous nematodes

Miscellaneous nematodes
Literature review current through: Jan 2024.
This topic last updated: Dec 28, 2022.

INTRODUCTION — The nematodes (roundworms) reviewed here include dirofilariasis, capillariasis, trichostrongyliasis, anisakiasis, Angiostrongylus costaricensis, dracunculiasis, and thelaziasis (table 1). Other tissue nematode infections are discussed separately, including gnathostomiasis, baylisascariasis, and Angiostrongylus cantonensis. (See "Eosinophilic meningitis".)

ANGIOSTRONGYLUS COSTARICENSIS — A. costaricensis is a filariform nematode that causes eosinophilic enterocolitis, a granulomatous inflammatory reaction within the intestinal wall. The appendix, distal small bowel, or right colon may be involved. (See "Eosinophilic gastrointestinal diseases".)

The life cycle of A. costaricensis begins with eggs laid by adult worms in the mesenteric arterioles of rats, the definitive hosts (figure 1). The first-stage larvae are passed in the stool and ingested by a snail or slug (intermediate host). The life cycle is completed with mollusk ingestion by the rat, and the third-stage larvae (infective larvae) migrate to the ileocecal area.

Humans can acquire the infection by eating raw or undercooked snails or slugs infected with the parasite; infection may also be transmitted by raw produce contaminated with larva-containing slug secretions. Alternatively, infection can be transmitted by ingestion of infected paratenic animals, such as crab or freshwater shrimp. In humans, worms migrate to the mesenteric arteries and release eggs into the intestinal tissues. Intense endothelial damage with subsequent arteritis and thrombosis can ensue, with necrosis of adjacent enteric tissue. Colonic mass due to suspected intravascular A. costaricensis infection has been described [1]. Humans are dead-end hosts and the parasite usually dies in the gastrointestinal tract; eggs are not shed in the stool.

A. costaricensis has been reported most frequently from Central and South America and the Caribbean, especially from Costa Rica (where it was first discovered) [2]. Cases have also occurred in the United States and elsewhere [3-5]. It most commonly affects school-aged children and young adults [6].

Clinical manifestations — Many infections are asymptomatic. Depending on the severity of the inflammatory reaction within the bowel wall, fever, abdominal pain, anorexia, vomiting, constipation, bowel obstruction, mesenteric ischemia, or perforation can develop [7,8]. Patients often have clinical signs similar to those of acute appendicitis, corresponding to an eosinophilic infiltrate in the ileocecal region. However, the presentation can be indolent, with relapsing episodes of abdominal pain recurring over several months. Presentation with recurrent gastrointestinal bleeding has also been described [9]. A tumor-like mass is frequently palpable on abdominal examination.

Rarely, eggs and larvae can be carried to extraintestinal sites [2,10,11]. When this occurs, mesenteric lymph node enlargement, testicular involvement with pain and erythema, or hepatic lesions causing tender hepatomegaly can result.

The incubation period is uncertain; it is estimated to range from three weeks to several months [6].

Diagnosis — Definitive diagnosis may be established by identifying the organism on histopathologic examination of biopsies or surgical resections (picture 1). Eggs or larvae may be observed within the tissues, or adult worms may be found in the mesenteric arterial lumen or its branches [6]. Characteristic histopathologic findings include eosinophilic infiltration of the intestinal wall, granulomatous reaction, and eosinophilic vasculitis [12].

Adult parasites measure 2 to 3.5 cm. Stool examination is not helpful since neither eggs nor larvae appear in the stool, although stool examination may be useful to exclude other potential parasitic causes, particularly Enterobius vermicularis. Enzyme-linked immunosorbent assays, immunochromatographic testing, and polymerase chain reaction assays have been developed but are not widely available [13-16].

A high-grade peripheral eosinophilia (up to 20 to 50 percent) is often present and may help to differentiate this infection from acute appendicitis.

Other causes of eosinophilic enteritis include anisakiasis, enterobiasis, toxocariasis, and Ancylostoma caninum.

Treatment — Most patients have a self-limited course and usually can be observed with supportive care in the absence of specific medical or surgical therapy [17]. In some cases, surgery is pursued to exclude appendicitis or other causes of pathology.

Antiparasitic agents have not proven efficacious in any clinical trial [2]. It may be best to avoid anthelminthic therapy if the diagnosis is certain, as treatment may lead to erratic migration followed by worsening of the disease [10].

There is no clear evidence of efficacy for anti-inflammatory agents [6].

ANISAKIASIS — Anisakiasis is a zoonotic roundworm infection caused by Anisakis simplex, Anisakis physeteris, Anisakis pegreffii, or Pseudoterranova species. Marine mammals (whales, sea lions, seals, dolphins, porpoises, and walruses) are the natural hosts; humans are incidental hosts [18,19]. The human "equivalent" of anisakiasis is ascariasis.

Anisakiasis has been described in many regions but is most common in Japan, likely because of the frequent ingestion of raw fish. It is also known as "sushi worm," "herring worm" (Anisakis species), or "cod worm" (Pseudoterranova species). The global prevalence of anisakiasis may be underestimated, given diagnostic limitations in diagnostic tools and protean clinical manifestations; reports from the Americas and from Europe are increasing, especially from Italy and Spain [20-22].

The life cycle of anisakiasis begins with passage of unembryonated eggs in the stool of marine mammals (figure 2). In the water, first- and second-stage larvae are formed; subsequently, these are ingested by crustaceans and migrate to muscle tissues. The larvae are transferred to fish and squid via predation, which maintain third-stage larvae that are infective to humans and marine mammals. Upon ingestion by marine mammals, the larvae develop into adult worms, which become embedded in the stomach mucosa and produce eggs shed in the stool.

Humans become infected by eating undercooked, raw, or pickled infected fish. Salmon, herring, cod, mackerel, and squid transmit Anisakis species; halibut, cod, and red snapper transmit Pseudoterranova species. The larvae are grossly visible in the fish; properly trained sushi chefs can detect them. After ingestion, the larva (usually one or two) penetrates the human gastric and intestinal mucosa. Maturation begins, but the parasite dies because it is not in its natural host. The dying organism induces an inflammatory reaction, and a tissue abscess develops with a predominance of eosinophils. In some cases, the larvae perforate the wall of the intestine and form an abscess within the peritoneal cavity.

Thorough cooking to 70°C or adequate freezing to -20°C for a minimum of 72 hours are the best preventive measures.

Clinical manifestations — The major clinical syndromes include gastric, intestinal, extragastrointestinal, and allergic disease [19]. Most symptoms associated with anisakiasis are due to direct tissue damage or an allergic reaction [23].

Immediately after ingestion of infected raw fish, some individuals develop pruritus and tingling of the posterior oropharynx. Severe laryngeal edema has been reported, which can be fatal [24]. Gastric anisakiasis usually develops one to eight hours after ingestion of raw fish and is characterized by acute epigastric pain, nausea, and vomiting [25]. Intestinal anisakiasis usually develops five to seven days following ingestion of raw fish and may be associated with severe abdominal pain, abdominal distension, and a palpable inflammatory mass that causes intestinal obstruction [25,26]. Diarrhea with blood or mucus may also develop [27].

Eosinophilic esophagitis, gastroenteritis, or enterocolitis can also occur. A syndrome mimicking appendicitis may be observed if the ileocecal region is involved [28,29]. Symptoms are vague and the illness can be misdiagnosed as appendicitis, acute abdomen, stomach ulcer, or ileitis. Anisakis larvae occasionally penetrate into the peritoneal cavity or other visceral organs (extraintestinal anisakiasis) and cause eosinophilic granuloma, which may be confused with neoplasm [26].

Allergic reactions ranging from mild urticaria to bronchoconstriction, angioedema, and anaphylactic shock can occur [21,30]. Associated fever and peripheral eosinophilia are common. Symptoms usually arise acutely; a chronic relapsing course can also occur. These symptoms may mimic seafood allergy. Patients with severe allergic reactions are generally warned against further consumption of marine fish or squid given the potential for infection with Anisakis [31]. (See "Seafood allergies: Fish and shellfish".)

The differential diagnosis of small bowel obstruction due to anisakiasis includes tumor, Crohn disease, primary eosinophilic gastroenteritis, other parasitic infections (Strongyloides, Ascaris, Toxocara, Ancylostoma, Gnathostoma), bacterial infections (Yersinia, tuberculosis), intussusception, and ischemia [32].

Diagnosis — The diagnosis of anisakiasis can be made via visualization of the worm recovered from emesis or by endoscopy (approximately 2 to 2.5 cm by 1 to 2 mm) (picture 2) [33,34]. Larvae may be grossly visible to the naked eye of the endoscopist, or endoscopic examination may demonstrate an ulcerated bleeding lesion in the stomach or duodenum with a worm at the center. Barium studies may demonstrate narrowing of the intestinal lumen in areas with mucosal inflammation (image 1). A thread-like filling defect suggesting a worm is visualized in some cases [35]. Computed tomography or ultrasonography may demonstrate edematous wall thickening of the gastric or intestinal mucosa, dilated loops of small bowel, hyperperistalsis, perigastric stranding, and ascites [36-40].

Total and Anisakis-specific immunoglobulin E levels are often elevated, particularly in patients who develop an allergic reaction following infection [41-43]. Enzyme-linked immunosorbent assay (ELISA) tests and immunoblot tests have been developed for diagnosis of anisakiasis but are not widely available [44-46]. An antigen-capture ELISA has been described with reported sensitivity and specificity near 100 percent [47]. Polymerase chain reaction tests have also been developed, but they are not commercially available [48,49].

Treatment — Physical removal of the parasite (by regurgitation, endoscopy, or surgery) is curative. Symptomatic therapy is usually adequate if the worm is in the distal bowel and cannot be retrieved by endoscopy, since Anisakis larvae can only survive for a few days in the human intestinal tract. Surgery may be necessary for worms that have penetrated the intestine, omentum, liver, or pancreas [50].

Successful treatment with albendazole (400 mg orally twice daily for 3 to 21 days) has been described, sometimes with the addition of prednisolone (20 mg/day up to 1 mg/kg/day) [51-53]; data are limited.

CAPILLARIASIS — There are two major clinical syndromes of capillariasis: intestinal disease (caused by Capillaria philippinensis) and hepatic disease (caused by Capillaria hepatica). Intestinal capillariasis occurs most frequently in Thailand and the Philippines, but it has also been observed in other countries [54]. Rare cases of human infections with C. hepatica have been reported worldwide [55].

Intestinal capillariasis — Intestinal capillariasis is caused by C. philippinensis, a parasite of fish-eating birds (which seem to be the natural definitive host) [56]. Adult worms of C. philippinensis reside in the bird and human intestinal tracts (figure 3). Unembryonated eggs are passed in the stool and become embryonated in fresh water, where they are ingested by fish. Subsequently, larvae hatch, penetrate the fish intestine, and migrate to muscle tissues. Ingestion of raw or undercooked fish results in bird and/or human infection.

Unembryonated eggs can also become embryonated in the human intestine; released larvae can cause autoinfection, leading to hyperinfection (a massive number of adult worms) [57]. Consequently, exposure to even a small parasite load can result in massive infection.

Clinical manifestations — Clinical symptoms associated with intestinal capillariasis include chronic watery diarrhea, abdominal pain, weight loss, malabsorption, and wasting. Fever, abdominal pain, and peripheral eosinophilia may also be present. As the parasite burden increases, malabsorption can become very severe, resulting in edema and muscle wasting. Electrolyte abnormalities and protein loss result, leading to marked cachexia and cardiomyopathy. Untreated infection can lead to death within a few months [58].

Diagnosis and treatment — The diagnosis of intestinal capillariasis can be made by detecting characteristic eggs in stool specimens (picture 3). The eggs measure 45 by 20 microns with plugs at each end. Eggs may be shed intermittently, reducing the sensitivity of stool specimens. Larvae may also be found in stool specimens. In some cases, small intestine histopathology is needed to confirm the diagnosis and should be considered in cases of otherwise unexplained severe malabsorption. Enzyme-linked immunosorbent assay (ELISA), western blot, and stool polymerase chain reaction (PCR) tests have been developed but are not widely available [59,60].

Treatment for intestinal capillariasis consists of albendazole (400 mg on empty stomach once or twice daily, often given for 30 days [minimum 10 days]) or mebendazole (200 mg twice daily for 20 to 30 days) [17,61,62]; there are no randomized controlled trials evaluating the approach to therapy or optimal duration. Repeating the stool examination within a day of anthelminthic drug treatment may increase the sensitivity of a stool specimen to confirm the diagnosis [54].

Supportive therapy with fluids and nutritional supplements may be required. Relapse is common, particularly if patients do not complete the full course of therapy; in such cases, retreatment may be necessary [57]. The optimal management of relapse is uncertain; based on case reports, retreatment with albendazole or mebendazole (perhaps for a longer course if not administered for 30 days initially) is reasonable [63-65].

Hepatic capillariasis — Hepatic capillariasis is caused by C. hepatica, a parasite of rodents, dogs, pigs, and other mammals; humans are incidental hosts. Hepatic capillariasis has been described worldwide. Human infection is rare; infection is most common among children <3 years of age and is acquired by ingesting eggs in contaminated food, water, or soil [66,67].

The life cycle begins with ingestion of embryonated eggs by a suitable mammalian host (figure 4). Larvae are released in the intestine and migrate via the portal vein to the liver, where they mature into adults after about four weeks and lay eggs in the parenchyma. The adult worms in the liver are destroyed by host inflammation, but the eggs remain viable in the hepatic parenchyma. Occasionally, larvae migrate to the lungs, kidneys, or other organs.

Eggs are not passed in the stool of the host; they remain in the liver until the animal dies or is eaten by a predator. Eggs ingested by scavengers or predators are passed in the stool of these animals and embryonate in the environment after about 30 days.

Clinical manifestations — The triad of clinical manifestations includes fever, hepatomegaly, and eosinophilia [68]; symptoms also include acute or subacute hepatitis [69]. Large areas of parenchyma may be replaced by mass of eggs, leading to inflammation, granuloma formation, and fibrosis of the liver. The clinical picture may resemble visceral larva migrans [70]. (See "Toxocariasis: Visceral and ocular larva migrans".)

Diagnosis and treatment — The diagnosis of capillariasis in humans is usually made by finding adults and eggs in liver biopsy or autopsy specimens (picture 4). Adult worms are 2.3 to 4.3 mm long with a diameter of ≥50 micrometers [71]. Larva vary in size according to stage. Eggs are 40 to 67 micrometers by 27 to 35 micrometers. Eggs are not passed in the stool of the host; identification of C. hepatica eggs in stool reflects spurious passage of ingested eggs and is not diagnostic of clinical infection.

Imaging with ultrasound, computed tomography, or magnetic resonance imaging examinations show hepatomegaly and nodular hypoechoic or mixed hypo-/hyperechoic liver lesions, often with irregular peripheral enhancement [69]. Serologic tests (indirect fluorescent antibody and ELISA) and PCR have been developed but are not widely available [72].

The optimal treatment is uncertain. Albendazole (400 mg twice daily for at least two weeks) and thiabendazole have been used successfully in some cases [17,73], often in combination with corticosteroids. Albendazole acts on adult worms but is not effective against the eggs, so adding corticosteroids may reduce the associated inflammatory response [68].

DIROFILARIASIS — Dirofilariasis is caused by a zoonotic filarial nematode. The life cycle of dirofilariasis begins when an infected mosquito (Aedes, Culex, Anopheles, Mansonia) takes a blood meal, introducing third-stage (L3) filarial larvae (figure 5). Usually, a domestic dog or coyote is infected, although a wide variety of other animals can be infected including cats, weasels, aquatic mammals, beaver, horses, and humans. The L3 larvae molt into L4 larvae and then adults, which reside in the subcutaneous tissues (Dirofilaria repens) or the heart of the definitive host (Dirofilaria immitis). Humans are incidental hosts; in humans, D. immitis filaria lodge in pulmonary arteries and usually cannot fully mature into gravid worms. D. repens often lodge in subcutaneous or ocular tissue.

In definitive hosts, adult worms can live for 5 to 10 years. The female worms are capable of producing microfilariae that circulate in the peripheral blood and are ingested by a mosquito during a blood meal. In the mosquito's abdomen, the microfilariae develop into L1 and subsequently into L3 larvae, which can infect another host when the mosquito takes a blood meal.

Dirofilariasis is particularly common in the Mediterranean region but has been described in many regions, including the United States, eastern Europe, and central Asia [74]. In the Americas, D. immitis is the main species; in Europe, D. repens is often the causative agent [75-77]. D. immitis has a worldwide distribution; D. repens is found in Europe, Asia, and Africa [78]. Climate change and increases in the movement of reservoirs (mostly infected dogs) have broadened the geographical range of these parasites and the risk for human infection [79].

Clinical manifestations — There are two major clinical syndromes: pulmonary dirofilariasis (caused by D. immitis) and subcutaneous or ocular dirofilariasis (caused by a few different dirofilarial species, particularly D. repens). In addition to these syndromes, there have been reports of human dirofilariasis in other sites, such as the peritoneal cavity, male genital tract, liver, buccal mucosa, or central nervous system [80-84].

A peripheral eosinophilia of approximately 10 percent may be observed [85].

Pulmonary dirofilariasis — Pulmonary dirofilariasis is caused by D. immitis (also known as the dog heartworm since it is a common cause of congestive heart failure in dogs) (figure 5). In definitive hosts, adult worms of D. immitis live in the heart. In humans, larvae lodge in small caliber pulmonary arteries and never mature into fully gravid worms. The organisms can cause pulmonary infarcts or pneumonitis with granuloma formation, which may result in the appearance of nodules or cavities on chest radiography [86,87]. The radiographic appearance is often described as a "coin lesion" that is usually 1 to 3 cm in diameter and can be confused with a lung tumor.

Most human infections are asymptomatic; often, infection is discovered incidentally when chest imaging is performed for some other reason. Some patients develop chest pain, cough, hemoptysis, fever, and malaise [88,89].

Subcutaneous dirofilariasis — Subcutaneous dirofilariasis is caused by a few different dirofilarial species, including D. repens, D. tenuis, and others (figure 6). These species are parasites of dogs and cats (D. repens), raccoons (D. tenuis), or other mammals. Adult worms can develop in humans, but sexual maturity and production of microfilariae do not occur since humans are an incidental host.

Skin lesions consist of a coiled, degenerating worm in subcutaneous tissues, typically around the eye or on the genitalia or limbs (picture 5) [90-92]. Frequently, the worm is encased in dense fibrous tissue. The nodule can be erythematous and tender and may be associated with an abscess. Concomitant allergic symptoms including urticaria and fever may also develop.

D. repens infection has also been reported as a cause of cutaneous larva migrans syndrome, with creeping eruption and elevated sinuous track under the skin [93,94].

Ocular dirofilariasis — Ocular dirofilariasis is usually caused by D. repens [76]. The conjunctiva is the most common site of nodules; involvement of the orbit (palpable mass) and eyelid have also been described [95]. The vitreous tissues may also be affected. Presentation with orbital cellulitis has been described. The vitreous tissues may also be affected causing floaters [96]. Patients often complain of discomfort, ocular pain, grittiness, and redness of the eye.

Concurrent subcutaneous and ocular D. repens infection has been described [97].

Diagnosis — Definitive diagnosis of dirofilariasis requires biopsy of the involved tissue for histopathologic identification. Dirofilaria are often 10 to 30 cm in length and 300 to 400 microns in diameter and characterized by a thick multilayered cuticle [92]. The cuticle of D. repens worms is spiked, whereas that of D immitis worms is smooth [76]. In one series of 60 patients with solitary pulmonary nodules presumed to be caused by D. immitis, 90 percent contained a single worm; occasionally, two or three worms were present in the same nodule [98]. Circulating microfilariae occasionally have been found [77,99,100].  

Serology using either enzyme-linked immunosorbent assay or indirect hemagglutination is not well standardized or widely used [88,101]. Polymerase chain reaction has also been used for diagnosis and species identification but is not widely available.

Subcutaneous lesions may be examined with ultrasonography for visualization of motile worms [102,103].

Treatment — Treatment consists of simple extraction or complete surgical excision of the worm [104]. In general, no specific medical therapy for dirofilariasis is required routinely. However, treatment with ivermectin may be given, particularly in cases where microfilaremia is present. Alternatively, for D. repens infection, doxycycline may be given to target the bacterial endosymbiont Wolbachia [100,105,106]. Often, the lesions calcify without treatment [98].

DRACUNCULIASIS — Dracunculiasis (also known as guinea worm) is caused by Dracunculus medinensis [107]. Infection is mainly transmitted by consumption of unfiltered water containing copepods (small crustaceans) infected with larvae of D. medinensis (figure 7). It may also be transmitted by eating fish or other aquatic animals [108]; these routes may be particularly important in acquisition of infection by dogs. Following ingestion, the copepods die and release larvae that penetrate the host's stomach and intestinal wall; thereafter, they enter the abdominal cavity and retroperitoneal space.

After maturation into adult worms, the males die and the females (70 to 120 cm in length) migrate in the subcutaneous tissues. Approximately one year after infection, the fertilized female worm migrates to the surface of the skin and induces a painful papule (usually on the distal lower extremity but may occur on the genitalia, buttocks, or trunk). One or more worms emerge, and the patient experiences a burning sensation. When the patient soaks the leg in fresh water to relieve the discomfort, the worm releases larvae into the water. The larvae are ingested by a copepod and become infective after two weeks (and two molts). Human ingestion of the copepods completes the cycle.

Clinical manifestations and diagnosis — Just prior to the formation of the skin papule, systemic symptoms can develop including fever, urticaria, pruritus, dizziness, nausea, vomiting, and diarrhea. The papule measures 2 to 7 cm, and pain is severe as the worm emerges (picture 6). This clinical manifestation is the basis for diagnosis. A peripheral eosinophilia may be present.

Rarely, the worm can migrate to ectopic sites, such as the lung, eye, pericardium, or spinal cord and can produce abscesses at these locations. Secondary infections can lead to systemic sepsis. Chronic arthritis and contractures can develop, particularly if the worm migrates through a joint.

In some cases, the worms die before they can emerge through the skin. In such cases, the worms eventually calcify and may be detected incidentally on radiographs or may be palpable beneath the skin [109].

Treatment — Treatment consists of slow extraction of the worm combined with wound care and pain management [17]. The worm should be wound around a stick, extracting a few centimeters each day. It may take many weeks or months for the entire worm to be removed. If the worm is broken or not fully extracted, an intense, painful inflammatory reaction with swelling along the worm tract can develop.

Epidemiology and prevention — Dracunculiasis occurs most commonly among adults in rural settings. In 1986, an estimated 3.5 million cases occurred each year in 20 countries in Africa and Asia. In 2021, 15 cases were reported; in 2022, 13 human cases were reported [110-112]. Infection continues to occur in five countries. Most infections are reported from Chad, but Angola, Ethiopia, Mali, and South Sudan also are reporting cases [108], and Cameroon continues to be affected by imported cases.

Previously, humans were thought to be the only known host of D. medinensis; emergence of infections in animals (predominantly domestic dogs) has complicated eradication efforts. Novel transmission pathways have been postulated; these include consumption of copepods from water by dogs, consumption of infective larvae in the entrails of aquatic animals, or consumption of raw or undercooked fish containing ingested copepods carrying infective larvae [113,114]. The target for eradication of Dracunculiasis has been pushed to 2030 [115].

Case containment to prevent water supply contamination can prevent infection [116]. Community surveillance and education regarding the mode of transmission is important for control. Other strategies include using of nylon filters for drinking water to remove copepods, use of insecticides in drinking water sources to kill copepods, covering papules with occlusive dressings, and covering drinking water sources so that infected individuals do not immerse infectious papules to propagate infection.

However, dogs may act as alternative hosts of the worm and may serve as reservoirs of D. medinensis, which (together with insecurity and civil unrest) may thwart the final steps toward eradication [117,118]. Control efforts are concentrating on containment of infection in dogs, particularly in Chad where most zoonotic infections occur [108].

THELAZIASIS — Thelaziasis, also referred to as the oriental eye worm, is an ocular infection caused by Thelazia species that is transmitted by drosophilid flies that feed on lacrimal secretions. Typically affected animals include dogs, cats, wild carnivores, horses, and cattle.

In humans, thelaziasis is an uncommon zoonotic infection that has been observed in Asia, Europe and North America [119]. Most human infections are due to Thelazia callipaeda. In the western United States, infection due to Thelazia californiensis and Thelazia gulosa has been described [120,121]. Children and older adults in resource-limited settings are most frequently affected [122].

Infection is transmitted by flies that feed on lacrimal secretions of infected animals. Within the flies, larvae develop into infective third-stage larvae that can be introduced from the fly into the conjunctival sac of host. Larvae develop further over one month into adult worms that live in the conjunctival sac or lacrimal apparatus. Adult worms are 1 to 2 cm in length and are cream colored.

Clinical manifestations — As one or more adult worms develop around the eye, infected patients may experience ocular pruritus, lacrimation, exudative conjunctivitis, a foreign body sensation in the eye, hypersensitivity to light, and keratitis [123]. Presentation with periorbital swelling has also been described [124]. In severe cases, corneal ulceration and scarring may lead to blindness [125].

Diagnosis — The diagnosis of thelaziasis is usually made by visualizing one or more worms in the conjunctival sac or lacrimal secretions. Polymerase chain reaction-based diagnosis has also been described [126].

Treatment — Adult worms and larvae can be removed by rinsing the conjunctival sac with sterile saline, and adults can be removed with forceps or cotton swabs. Repeated removal may be needed when several adults reside in the lacrimal secretions [120,125]. The utility and safety of anthelminthic treatments are not documented; physical removal of adult worms is sufficient.

TRICHOSTRONGYLOSIS — Trichostrongylosis is caused by several nematodes of the Trichostrongylus species, which infect sheep, cattle, and other herbivorous mammals worldwide; humans are incidental hosts.

Eggs are passed in the stool of the definitive host (usually a herbivorous mammal), and rhabditiform larvae hatch within several days; they become infective filariform (third-stage) larvae after 5 to 10 days (and two molts) (figure 8). Infection is transmitted to humans by ingestion of these larvae, which mature into adults in the small intestine. Trichostrongylosis is typically transmitted by ingestion of unwashed vegetables fertilized with contaminated manure [127,128].

Occasionally, infection can occur via larval penetration of the skin. Larvae mature to adults in the small intestine, where they embed in the mucosa and cause inflammation.

Clinical manifestations — Most infections with Trichostrongylus spp are asymptomatic. In the setting of heavy infection, abdominal pain, diarrhea, and anemia can develop. Malabsorption and wasting can ensue if mucosal damage is severe. A peripheral eosinophilia may be observed.

Diagnosis — The diagnosis of trichostrongylosis is generally established by identifying characteristic eggs in the stool (picture 7). Stool concentration techniques may be needed, particularly in the setting of light infections. The diagnosis can also by identification of characteristic Trichostrongylus larvae or adult worms on endoscopic evaluation of the duodenum. Polymerase chain reaction assays have been developed but are not widely available [129,130].

Treatment — Treatment of trichostrongylosis consists of mebendazole (100 mg twice daily for three days) or albendazole (400 mg orally once on empty stomach) [127]. Pyrantel pamoate (11 mg/kg orally once; maximum dose of 1 g) can also be used.

Prevention — Fresh vegetables and salads should be washed carefully. Only dried manure should be used as an organic fertilizer [131].

SUMMARY AND RECOMMENDATIONS

Angiostrongylus costaricensis causes eosinophilic enterocolitis. The primary life cycle consists of transmission between rodents and snails or slugs; humans are incidental hosts (figure 1). The diagnosis may be established by identifying the organism on histopathological examination of biopsies or surgical resections (picture 1). Most patients have a self-limited course and usually can be observed with supportive care in the absence of specific medical or surgical therapy. (See 'Angiostrongylus costaricensis' above.)

Anisakiasis causes gastroenteritis or enterocolitis. The primary life cycle consists of transmission between marine mammals, crustaceans, fish, and squid; humans are incidental hosts (figure 2). The diagnosis may be established by visualization of the worm recovered from emesis or by endoscopy (picture 2). Physical removal of the parasite (by regurgitation, endoscopy, or surgery) is curative. (See 'Anisakiasis' above.)

Capillaria philippinensis causes intestinal capillariasis. The primary life cycle consists of transmission between birds and fish; humans are incidental hosts (figure 3). The diagnosis may be established by detecting characteristic eggs in stool specimens (picture 3). We suggest treatment with albendazole or mebendazole (Grade 2C); dosing is outlined above. (See 'Capillariasis' above.)

Dirofilariasis consists of two major clinical syndromes: pulmonary dirofilariasis (caused by Dirofilaria immitis) and subcutaneous/ocular dirofilariasis (caused by a few different species, mainly Dirofilaria repens). The primary life cycle consists of transmission between dogs and mosquitoes; humans are incidental hosts (figure 5 and figure 6). The diagnosis may be established by biopsy of the involved tissue for histopathologic identification. No specific therapy for dirofilariasis is required. (See 'Dirofilariasis' above.)

Dracunculiasis causes guinea worm infection. Humans were thought to be the only host; however, the majority of cases are now reported in dogs. Infection is transmitted by consumption of unfiltered water containing copepods (small crustaceans) infected with larvae of Dracunculus medinensis (figure 7). Transmission may also occur via ingestion of raw fish or other aquatic animals; these routes may be particularly important in dogs. Following ingestion, larvae penetrate the gastrointestinal tract and adult worms migrate to the subcutaneous tissues, where they induce painful papules (picture 6). Treatment consists of slow extraction of the worm combined with wound care and pain management. (See 'Dracunculiasis' above.)

Trichostrongylosis causes asymptomatic infection or gastrointestinal illness. The primary life cycle consists of transmission between herbivorous animals; humans are incidental hosts (figure 8). The diagnosis may be established by identifying characteristic eggs in the stool (picture 7). We suggest treatment with mebendazole or albendazole (Grade 2C); dosing is outlined above. (See 'Trichostrongylosis' above.)

Thelaziasis is an ocular infection caused by Thelazia species that is transmitted by flies that feed on lacrimal secretions. Affected animals include dogs, cats, horses, and cattle. The diagnosis is made by visualizing one or more worms in the conjunctival sac or lacrimal secretions. Treatment consists of physical removal of the adult worms. (See 'Thelaziasis' above.)

  1. Rubin AK, Burk KS, Staller K, et al. Case 30-2018: A 66-Year-Old Woman with Chronic Abdominal Pain. N Engl J Med 2018; 379:1263.
  2. Loría-Cortés R, Lobo-Sanahuja JF. Clinical abdominal angiostrongylosis. A study of 116 children with intestinal eosinophilic granuloma caused by Angiostrongylus costaricensis. Am J Trop Med Hyg 1980; 29:538.
  3. Wu SS, French SW, Turner JA. Eosinophilic ileitis with perforation caused by Angiostrongylus (Parastrongylus) costaricensis. A case study and review. Arch Pathol Lab Med 1997; 121:989.
  4. Kramer MH, Greer GJ, Quiñonez JF, et al. First reported outbreak of abdominal angiostrongyliasis. Clin Infect Dis 1998; 26:365.
  5. Dard C, Nguyen D, Miossec C, et al. Angiostrongylus costaricensis infection in Martinique, Lesser Antilles, from 2000 to 2017. Parasite 2018; 25:22.
  6. Rojas A, Maldonado-Junior A, Mora J, et al. Abdominal angiostrongyliasis in the Americas: fifty years since the discovery of a new metastrongylid species, Angiostrongylus costaricensis. Parasit Vectors 2021; 14:374.
  7. Waisberg J, Corsi CE, Rebelo MV, et al. Jejunal perforation caused by abdominal angiostrongyliasis. Rev Inst Med Trop Sao Paulo 1999; 41:325.
  8. Kröner PT, Argueta V. Abdominal angiostrongyliasis mimicking acute appendicitis. Endoscopy 2015; 47 Suppl 1 UCTN:E179.
  9. Silvera CT, Ghali VS, Roven S, et al. Angiostrongyliasis: a rare cause of gastrointestinal hemorrhage. Am J Gastroenterol 1989; 84:329.
  10. Rodriguez R, Dequi RM, Peruzzo L, et al. Abdominal angiostrongyliasis: report of two cases with different clinical presentations. Rev Inst Med Trop Sao Paulo 2008; 50:339.
  11. Vázquez JJ, Sola JJ, Boils PL. Hepatic lesions induced by Angiostrongylus costaricensis. Histopathology 1994; 25:489.
  12. Quirós JL, Jiménez E, Bonilla R, et al. Abdominal angiostrongyliasis with involvement of liver histopathologically confirmed: a case report. Rev Inst Med Trop Sao Paulo 2011; 53:219.
  13. Geiger SM, Laitano AC, Sievers-Tostes C, et al. Detection of the acute phase of abdominal angiostrongyliasis with a parasite-specific IgG enzyme linked immunosorbent assay. Mem Inst Oswaldo Cruz 2001; 96:515.
  14. Rodriguez R, da Silva AC, Müller CA, et al. PCR for the diagnosis of abdominal angiostrongyliasis in formalin-fixed paraffin-embedded human tissue. PLoS One 2014; 9:e93658.
  15. Ben R, Rodrigues R, Agostini AA, Graeff-Teixeira C. Use of heterologous antigens for the immunodiagnosis of abdominal angiostrongyliasis by an enzyme-linked immunosorbent assay. Mem Inst Oswaldo Cruz 2010; 105:914.
  16. Graeff-Teixeira C, Pascoal VF, Rodriguez R, et al. Abdominal angiostrongyliasis can be diagnosed with a immunochromatographic rapid test with recombinant galactin from Angiostrongylus cantonensis. Mem Inst Oswaldo Cruz 2020; 115:e200201.
  17. Drugs for Parasitic Infections, 3rd ed, The Medical Letter, New Rochelle, NY 2013.
  18. Audicana MT, Kennedy MW. Anisakis simplex: from obscure infectious worm to inducer of immune hypersensitivity. Clin Microbiol Rev 2008; 21:360.
  19. Hochberg NS, Hamer DH. Anisakidosis: Perils of the deep. Clin Infect Dis 2010; 51:806.
  20. Uña-Gorospe M, Herrera-Mozo I, Canals ML, et al. Occupational disease due to Anisakis simplex in fish handlers. Int Marit Health 2018; 69:264.
  21. Cavallero S, Martini A, Migliara G, et al. Anisakiasis in Italy: Analysis of hospital discharge records in the years 2005-2015. PLoS One 2018; 13:e0208772.
  22. Guardone L, Armani A, Nucera D, et al. Human anisakiasis in Italy: a retrospective epidemiological study over two decades. Parasite 2018; 25:41.
  23. Caramello P, Vitali A, Canta F, et al. Intestinal localization of anisakiasis manifested as acute abdomen. Clin Microbiol Infect 2003; 9:734.
  24. Suzuki S, Bandoh N, Goto T, et al. Severe laryngeal edema caused by Pseudoterranova species: A case report. Medicine (Baltimore) 2021; 100:e24456.
  25. Park MS, Kim KW, Ha HK, Lee DH. Intestinal parasitic infection. Abdom Imaging 2008; 33:166.
  26. Nawa Y, Hatz C, Blum J. Sushi delights and parasites: the risk of fishborne and foodborne parasitic zoonoses in Asia. Clin Infect Dis 2005; 41:1297.
  27. Amir A, Ngui R, Ismail WH, et al. Anisakiasis Causing Acute Dysentery in Malaysia. Am J Trop Med Hyg 2016; 95:410.
  28. Gómez B, Tabar AI, Tuñón T, et al. Eosinophilic gastroenteritis and Anisakis. Allergy 1998; 53:1148.
  29. López-Serrano MC, Gomez AA, Daschner A, et al. Gastroallergic anisakiasis: findings in 22 patients. J Gastroenterol Hepatol 2000; 15:503.
  30. Daschner A, Alonso-Gómez A, Caballero T, et al. Gastric anisakiasis: an underestimated cause of acute urticaria and angio-oedema? Br J Dermatol 1998; 139:822.
  31. Adroher-Auroux FJ, Benítez-Rodríguez R. Anisakiasis and Anisakis: An underdiagnosed emerging disease and its main etiological agents. Res Vet Sci 2020; 132:535.
  32. Ramos L, Alonso C, Guilarte M, et al. Anisakis simplex-induced small bowel obstruction after fish ingestion: preliminary evidence for response to parenteral corticosteroids. Clin Gastroenterol Hepatol 2005; 3:667.
  33. Fuchizaki U, Nishikawa M. IMAGES IN CLINICAL MEDICINE. Gastric Anisakiasis. N Engl J Med 2016; 375:e11.
  34. Kondo T. Woe sushi: gastric anisakiasis. Lancet 2018; 392:1340.
  35. Matsui T, Iida M, Murakami M, et al. Intestinal anisakiasis: clinical and radiologic features. Radiology 1985; 157:299.
  36. Takabayashi T, Mochizuki T, Otani N, et al. Anisakiasis presenting to the ED: clinical manifestations, time course, hematologic tests, computed tomographic findings, and treatment. Am J Emerg Med 2014; 32:1485.
  37. Lee JS, Kim BS, Kim SH, et al. Acute invasive small-bowel Anisakiasis: clinical and CT findings in 19 patients. Abdom Imaging 2014; 39:452.
  38. Shibata E, Ueda T, Akaike G, Saida Y. CT findings of gastric and intestinal anisakiasis. Abdom Imaging 2014; 39:257.
  39. Lalchandani UR, Weadock WJ, Brady GF, Wasnik AP. Imaging in gastric anisakiasis. Clin Imaging 2018; 50:286.
  40. Ripollés T, López-Calderón LE, Martínez-Pérez MJ, et al. Usefulness of Ultrasound in the Diagnosis of Intestinal Anisakiasis. J Ultrasound Med 2020; 39:1703.
  41. Moreno-Ancillo A, Caballero MT, Cabañas R, et al. Allergic reactions to anisakis simplex parasitizing seafood. Ann Allergy Asthma Immunol 1997; 79:246.
  42. García M, Moneo I, Audicana MT, et al. The use of IgE immunoblotting as a diagnostic tool in Anisakis simplex allergy. J Allergy Clin Immunol 1997; 99:497.
  43. Desowitz RS, Raybourne RB, Ishikura H, Kliks MM. The radioallergosorbent test (RAST) for the serological diagnosis of human anisakiasis. Trans R Soc Trop Med Hyg 1985; 79:256.
  44. Gutierrez Ramos R, Tsuji M. Detection of antibodies to Anisakis simplex larvae by enzyme-linked immunosorbent assay and immunoelectrophoresis using crude or purified antigens. J Helminthol 1994; 68:305.
  45. Iglesias R, Leiro J, Santamarina MT, et al. Monoclonal antibodies against diagnostic Anisakis simplex antigens. Parasitol Res 1997; 83:755.
  46. Okazaki M, Goto I, Kurokawa I. Studies on the detection of anti-Anisakis larvae antibodies by ELISA kits. Med Pharmacol 1992; 22:971.
  47. Lorenzo S, Iglesias R, Leiro J, et al. Usefulness of currently available methods for the diagnosis of Anisakis simplex allergy. Allergy 2000; 55:627.
  48. Chen Q, Yu HQ, Lun ZR, et al. Specific PCR assays for the identification of common anisakid nematodes with zoonotic potential. Parasitol Res 2008; 104:79.
  49. Mattiucci S, Paoletti M, Colantoni A, et al. Invasive anisakiasis by the parasite Anisakis pegreffii (Nematoda: Anisakidae): diagnosis by real-time PCR hydrolysis probe system and immunoblotting assay. BMC Infect Dis 2017; 17:530.
  50. Sakanari JA, McKerrow JH. Anisakiasis. Clin Microbiol Rev 1989; 2:278.
  51. Pacios E, Arias-Diaz J, Zuloaga J, et al. Albendazole for the treatment of anisakiasis ileus. Clin Infect Dis 2005; 41:1825.
  52. Moore DA, Girdwood RW, Chiodini PL. Treatment of anisakiasis with albendazole. Lancet 2002; 360:54.
  53. Carlin AF, Abeles S, Chin NA, et al. Case Report: A Common Source Outbreak of Anisakidosis in the United States and Postexposure Prophylaxis of Family Collaterals. Am J Trop Med Hyg 2018; 99:1219.
  54. Sadaow L, Sanpool O, Intapan PM, et al. A Hospital-Based Study of Intestinal Capillariasis in Thailand: Clinical Features, Potential Clues for Diagnosis, and Epidemiological Characteristics of 85 Patients. Am J Trop Med Hyg 2018; 98:27.
  55. Fuehrer HP, Igel P, Auer H. Capillaria hepatica in man--an overview of hepatic capillariosis and spurious infections. Parasitol Res 2011; 109:969.
  56. Limsrivilai J, Pongprasobchai S, Apisarnthanarak P, Manatsathit S. Intestinal capillariasis in the 21st century: clinical presentations and role of endoscopy and imaging. BMC Gastroenterol 2014; 14:207.
  57. Grencis RK, Cooper ES. Enterobius, trichuris, capillaria, and hookworm including ancylostoma caninum. Gastroenterol Clin North Am 1996; 25:579.
  58. Cross JH. Intestinal capillariasis. Clin Microbiol Rev 1992; 5:120.
  59. Khalifa MM, Abdel-Rahman SM, Bakir HY, et al. Comparison of the diagnostic performance of microscopic examination, Copro-ELISA, and Copro-PCR in the diagnosis of Capillaria philippinensis infections. PLoS One 2020; 15:e0234746.
  60. Hassan MA, Basyoni MMA, Amer MF, Al-Antably ASA. Antigen Recognition Patterns of Intestinal Capillariasis Using Immunoblot-Based Serodiagnosis. Acta Parasitol 2020; 65:899.
  61. Vasantha PL, Girish N, Leela KS. Human intestinal capillariasis: a rare case report from non-endemic area (Andhra Pradesh, India). Indian J Med Microbiol 2012; 30:236.
  62. Kasher C, Grossman T, Vainer J, et al. First case of imported Capillaria philippinensis in Israel. J Travel Med 2022; 29.
  63. Cross JH, Basaca-Sevilla V. Albendazole in the treatment of intestinal capillariasis. Southeast Asian J Trop Med Public Health 1987; 18:507.
  64. Belizario VY Jr, G Totañes FI, de Leon WU, et al. Intestinal capillariasis, western Mindanao, the Philippines. Emerg Infect Dis 2010; 16:736.
  65. Dronda F, Chaves F, Sanz A, Lopez-Velez R. Human intestinal capillariasis in an area of nonendemicity: case report and review. Clin Infect Dis 1993; 17:909.
  66. Pampiglione S, Gustinelli A. Human hepatic capillariasis: a second case occurred in Korea. J Korean Med Sci 2008; 23:560.
  67. Klenzak J, Mattia A, Valenti A, Goldberg J. Hepatic capillariasis in Maine presenting as a hepatic mass. Am J Trop Med Hyg 2005; 72:651.
  68. Wang ZQ, Cui J, Wang Y. Persistent febrile hepatomegaly with eosinophilia due to hepatic capillariasis in Central China. Ann Trop Med Parasitol 2011; 105:469.
  69. Wang L, Zhang Y, Deng Y, et al. Clinical and laboratory characterizations of hepatic capillariasis. Acta Trop 2019; 193:206.
  70. Kumar V, Brandt J, Mortelmans J. Hepatic capillariasis may simulate the syndrome of visceral larva migrans, an analysis. Ann Soc Belg Med Trop 1985; 65:101.
  71. Manor U, Doviner V, Kolodziejek J, et al. Capillaria hepatica (syn. Calodium hepaticum) as a Cause of Asymptomatic Liver Mass. Am J Trop Med Hyg 2021; 105:204.
  72. Juncker-Voss M, Prosl H, Lussy H, et al. Serological detection of Capillaria hepatica by indirect immunofluorescence assay. J Clin Microbiol 2000; 38:431.
  73. Choe G, Lee HS, Seo JK, et al. Hepatic capillariasis: first case report in the Republic of Korea. Am J Trop Med Hyg 1993; 48:610.
  74. Jelinek T, Schulte-Hillen J, Löscher T. Human dirofilariasis. Int J Dermatol 1996; 35:872.
  75. Dantas-Torres F, Otranto D. Dirofilariosis in the Americas: a more virulent Dirofilaria immitis? Parasit Vectors 2013; 6:288.
  76. Simón F, Siles-Lucas M, Morchón R, et al. Human and animal dirofilariasis: the emergence of a zoonotic mosaic. Clin Microbiol Rev 2012; 25:507.
  77. Blaizot R, Receveur MC, Millet P, et al. Systemic Infection With Dirofilaria repens in Southwestern France. Ann Intern Med 2018; 168:228.
  78. Genchi C, Kramer L. Subcutaneous dirofilariosis (Dirofilaria repens): an infection spreading throughout the old world. Parasit Vectors 2017; 10:517.
  79. Capelli G, Genchi C, Baneth G, et al. Recent advances on Dirofilaria repens in dogs and humans in Europe. Parasit Vectors 2018; 11:663.
  80. Fleck R, Kurz W, Quade B, et al. Human dirofilariasis due to Dirofilaria repens mimicking a scrotal tumor. Urology 2009; 73:209.e1.
  81. Tada I, Sakaguchi Y, Eto K. Dirofilaria in the abdominal cavity of a man in Japan. Am J Trop Med Hyg 1979; 28:988.
  82. Kim MK, Kim CH, Yeom BW, et al. The first human case of hepatic dirofilariasis. J Korean Med Sci 2002; 17:686.
  83. Poppert S, Hodapp M, Krueger A, et al. Dirofilaria repens infection and concomitant meningoencephalitis. Emerg Infect Dis 2009; 15:1844.
  84. Momčilović S, Gabrielli S, Golubović M, et al. Human dirofilariosis of buccal mucosa - First molecularly confirmed case and literature review. Parasitol Int 2019; 73:101960.
  85. Flieder DB, Moran CA. Pulmonary dirofilariasis: a clinicopathologic study of 41 lesions in 39 patients. Hum Pathol 1999; 30:251.
  86. Hiroshima K, Iyoda A, Toyozaki T, et al. Human pulmonary dirofilariasis: report of six cases. Tohoku J Exp Med 1999; 189:307.
  87. Saha BK, Bonnier A, Chong WH, et al. Human Pulmonary Dirofilariasis: A Review for the Clinicians. Am J Med Sci 2022; 363:11.
  88. Ro JY, Tsakalakis PJ, White VA, et al. Pulmonary dirofilariasis: the great imitator of primary or metastatic lung tumor. A clinicopathologic analysis of seven cases and a review of the literature. Hum Pathol 1989; 20:69.
  89. Solaini L, Gourgiotis S, Salemis NS, Solaini L. A case of human pulmonary dirofilariasis. Int J Infect Dis 2008; 12:e147.
  90. Khoramnia R, Wegner A. Images in clinical medicine: Subconjunctival Dirofilaria repens. N Engl J Med 2010; 363:e37.
  91. Arvanitis PG, Vakalis NC, Damanakis AG, Theodossiadis GP. Ophthalmic dirofilariasis. Am J Ophthalmol 1997; 123:689.
  92. Tzanetou K, Gasteratos S, Pantazopoulou A, et al. Subcutaneous dirofilariasis caused by Dirofilaria repens in Greece: a case report. J Cutan Pathol 2009; 36:892.
  93. Antolová D, Miterpáková M, Paraličová Z. Case of human Dirofilaria repens infection manifested by cutaneous larva migrans syndrome. Parasitol Res 2015; 114:2969.
  94. Kartashev V, Simon F. Migrating Dirofilaria repens. N Engl J Med 2018; 378:e35.
  95. Kalogeropoulos CD, Stefaniotou MI, Gorgoli KE, et al. Ocular dirofilariasis: a case series of 8 patients. Middle East Afr J Ophthalmol 2014; 21:312.
  96. Rajan RP, Jena S, Ramachandran NO, Kohli P. Rare cause of floaters: A motile live worm in vitreous cavity. Indian J Ophthalmol 2019; 67:1490.
  97. Szostakowska B, Ćwikłowska A, Marek-Józefowicz L, et al. Concurrent subcutaneous and ocular infections with Dirofilaria repens in a Polish patient: a case report in the light of epidemiological data. Parasitol Int 2022; 86:102481.
  98. Ciferri F. Human pulmonary dirofilariasis in the United States: a critical review. Am J Trop Med Hyg 1982; 31:302.
  99. Potters I, Vanfraechem G, Bottieau E. Dirofilaria repens Nematode Infection with Microfilaremia in Traveler Returning to Belgium from Senegal. Emerg Infect Dis 2018; 24:1761.
  100. Huebl L, Tappe D, Giese M, et al. Recurrent Swelling and Microfilaremia Caused by Dirofilaria repens Infection after Travel to India. Emerg Infect Dis 2021; 27:1701.
  101. Glickman LT, Grieve RB, Schantz PM. Serologic diagnosis of zoonotic pulmonary dirofilariasis. Am J Med 1986; 80:161.
  102. Acharya D, Chatra PS, Padmaraj SR, Ahamed A. Subcutaneous dirofilariasis. Singapore Med J 2012; 53:e184.
  103. Alam SI, Nepal P, Lu SC, et al. Imaging Findings of Subcutaneous Human Dirofilariasis. Curr Probl Diagn Radiol 2021; 50:755.
  104. Khurana S, Singh G, Bhatti HS, Malla N. Human subcutaneous dirofilariasis in India: a report of three cases with brief review of literature. Indian J Med Microbiol 2010; 28:394.
  105. Lechner AM, Gastager H, Kern JM, et al. Case Report: Successful Treatment of a Patient with Microfilaremic Dirofilariasis Using Doxycycline. Am J Trop Med Hyg 2020; 102:844.
  106. Frenzen FS, Loewe I, Müller G, et al. Dirofilaria repens infection of the eye with concomitant microfilaremia in a traveller. J Travel Med 2021; 28.
  107. Greenaway C. Dracunculiasis (guinea worm disease). CMAJ 2004; 170:495.
  108. Hopkins DR, Weiss AJ, Roy SL, et al. Progress Toward Global Eradication of Dracunculiasis, January 2020-June 2021. MMWR Morb Mortal Wkly Rep 2021; 70:1527.
  109. Carranza-Rodríguez C, Pérez-Arellano JL. Radiological Detection of Dracunculus Medinensis. Am J Trop Med Hyg 2018; 98:1218.
  110. Burki T. Countries recommit to Guinea worm eradication by 2030. Lancet Infect Dis 2022; 22:597.
  111. Hopkins DR, Weiss AJ, Yerian S, et al. Progress Toward Global Eradication of Dracunculiasis - Worldwide, January 2021-June 2022. MMWR Morb Mortal Wkly Rep 2022; 71:1496.
  112. Hopkins DR, Weiss AJ, Yerian S, et al. Progress Toward Eradication of Dracunculiasis - Worldwide, January 2022-June 2023. MMWR Morb Mortal Wkly Rep 2023; 72:1230.
  113. Thach PN, van Doorn HR, Bishop HS, et al. Human infection with an unknown species of Dracunculus in Vietnam. Int J Infect Dis 2021; 105:739.
  114. Goodwin CED, Léchenne M, Wilson-Aggarwal JK, et al. Seasonal fishery facilitates a novel transmission pathway in an emerging animal reservoir of Guinea worm. Curr Biol 2022; 32:775.
  115. Roberts L. Exclusive: Battle to wipe out debilitating Guinea worm parasite hits 10 year delay. Nature 2019; 574:157.
  116. Hopkins DR. Dracunculiasis: an eradicable scourge. Epidemiol Rev 1983; 5:208.
  117. Galán-Puchades MT. Dogs and Guinea worm eradication. Lancet Infect Dis 2016; 16:770.
  118. Dracunculiasis eradication: global surveillance summary, 2016. Wkly Epidemiol Rec 2017; 92:269.
  119. Otranto D, Eberhard ML. Zoonotic helminths affecting the human eye. Parasit Vectors 2011; 4:41.
  120. Bradbury RS, Breen KV, Bonura EM, et al. Case Report: Conjunctival Infestation with Thelazia gulosa: A Novel Agent of Human Thelaziasis in the United States. Am J Trop Med Hyg 2018; 98:1171.
  121. Bradbury RS, Gustafson DT, Sapp SGH, et al. A Second Case of Human Conjunctival Infestation With Thelazia gulosa and a Review of T. gulosa in North America. Clin Infect Dis 2020; 70:518.
  122. Otranto D, Mendoza-Roldan JA, Dantas-Torres F. Thelazia callipaeda. Trends Parasitol 2021; 37:263.
  123. do Vale B, Lopes AP, da Conceição Fontes M, et al. Thelaziosis due to Thelazia callipaeda in Europe in the 21st century-A review. Vet Parasitol 2019; 275:108957.
  124. Koka K, Tongbram A, Mukherjee B, et al. Periocular thelaziasis presenting as an orbital mass - a case report. Orbit 2019; 38:503.
  125. Sah R, Khadka S, Adhikari M, et al. Human Thelaziasis: Emerging Ocular Pathogen in Nepal. Open Forum Infect Dis 2018; 5:ofy237.
  126. Morgado ACT, do Vale B, Ribeiro P, et al. First report of human Thelazia callipaeda infection in Portugal. Acta Trop 2022; 231:106436.
  127. Boreham RE, McCowan MJ, Ryan AE, et al. Human trichostrongyliasis in Queensland. Pathology 1995; 27:182.
  128. Watthanakulpanich D, Pongvongsa T, Sanguankiat S, et al. Prevalence and clinical aspects of human Trichostrongylus colubriformis infection in Lao PDR. Acta Trop 2013; 126:37.
  129. Mizani A, Gill P, Daryani A, et al. A multiplex restriction enzyme-PCR for unequivocal identification and differentiation of Trichostrongylus species in human samples. Acta Trop 2017; 173:180.
  130. Pandi M, Sharifdini M, Ashrafi K, et al. Comparison of Molecular and Parasitological Methods for Diagnosis of Human Trichostrongylosis. Front Cell Infect Microbiol 2021; 11:759396.
  131. Lattes S, Ferte H, Delaunay P, et al. Trichostrongylus colubriformis Nematode Infections in Humans, France. Emerg Infect Dis 2011; 17:1301.
Topic 5694 Version 37.0

References

آیا می خواهید مدیلیب را به صفحه اصلی خود اضافه کنید؟