Chapter 16: Parasites

• Potential Food Safety Hazard
• Nematodes
• Cestodes
• Trematodes
• Control Measures
• FDA Guidelines
• Published Process Studies
• Controlling nematodes
• Controlling trematodes
• Analytical Procedures
• Parasitic animals in foods
• Other analytical procedures
• References

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Potential Food Safety Hazard

Parasites (in the larval stage) consumed in uncooked, or undercooked, unfrozen seafood can present a human health hazard. Among parasites, the nematodes or roundworms (Anisakis simplex, Pseudoterranova decipiens, Eustrongylides spp., and Gnathostoma spinigerum, cestodes or tapeworms (Diphyllobothrium spp.) and trematodes or flukes (Clonorchis sinensis, Opisthorchis spp., Heterophyes spp., Metagonimus spp., Nanophyetes salminicola and Paragonimus spp.) are of most concern in seafood. Some products that have been implicated in human infection are: civiche or cibichi (fish and spices marinated in lime juice); lomi lomi (salmon marinated in lemon juice, onions and tomato); poisson cru (fish marinated in citrus juice, onions, tomatoes and coconut milk); salmon roe; sashimi (chunks of raw fish); sushi (pieces of raw fish with rice and other ingredients); green herring (lightly brined herring); drunken crabs (crabs marinated in wine and peppers); cold-smoked fish; and, undercooked grilled fish (FDA, 1998).

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Nematodes

Anisakiasis is caused by the accidental ingestion of larvae of the nematodes (roundworms) Anisakis simplex and Pseudoterranova decipiens. Adult stages of A. simplex or P. decipiens reside in the stomach of marine mammals, where they are embedded in the mucosa, in clusters. Eggs produced by adult females are passed in the feces, hatch and yield second stage larvae. Upon ingestion by crustaceans, third stage larvae develop that are infective to fish and squid. After ingestion by the fish and squid hosts, the larvae migrate from the intestine to the peritoneal cavity to (upon the host's death) the muscle tissues. Through predation, the larvae are transferred from fish to fish until they are ingested by the marine mammal. In this definitive host, the larvae develop into adults, thus closing the cycle. Humans become infected by eating raw or undercooked marine fish. After ingestion, the anisakid larvae penetrate the gastric and intestinal mucosa, causing the symptoms of anisakiasis.

Within hours after ingestion of infected larvae, violent abdominal pain, nausea, and vomiting may occur. Occasionally the larvae are coughed up. If the larvae pass into the bowel, a severe eosinophilic granulomatous response may also occur, causing symptoms mimicking Crohn's disease 1-2 weeks following infection.

A. simplex and P. decipiens are found worldwide, with higher incidence in areas where raw fish is eaten (e.g., Japan, Pacific coast of South America, the Netherlands). Increasing incidence in the United States due to increased consumption of raw fish (CDC, 1999a).

Anisakiasis is associated with eating raw fish (sushi, sashimi, lomi lomi, ceviche, sunomono, Dutch green herring, marinated fish and cold-smoked fish) or undercooked fish (Ward et al., 1997).

Freezing and cooking may kill A. simplex, but may not protect consumers against allergenic reactions to ingested A. simplex antigens (Audicana et al., 1997).

Eustrongylides spp. larvae are large, bright red nematodes, 25-150 mm long and 2 mm in diameter. They occur in freshwater fish, brackish water fish, and marine fish. The larvae normally mature in wading birds such as herons, egrets, and flamingos. If the larvae are consumed in raw or undercooked fish, they can attach to the wall of the digestive tract and penetrate the gut wall with accompanying severe pain. Infections are extremely rare and have been associated with consumption of live minnows and sashimi in the U.S. (FDA, 1991).

Gnathostoma spinigerum infects vertebrate animals. In the natural definitive host (cats, dogs, wild animals) the adult worms reside in a tumor which they induce in the gastric wall. They deposit eggs that are immature when passed in the feces. After maturation in water, the egg releases a first stage larva (L1). After ingestion by a small crustacean (Cyclops)(first intermediate host), the L1 develops into a L2. Following ingestion of the Cyclops by a fish, frog or snake (second intermediate host), the L2 develops in their flesh into a L3. When the second intermediate host is ingested by a definitive host, the L3 develops into an adult stage parasite in the stomach wall. Alternatively, the second intermediate host may be ingested by another animal (paratenic host) in which the L3 does not develop further, but remains infective to the next predator. Humans become infected by eating undercooked fish or poultry containing L3s, or reportedly by drinking water containing L2-infected Cyclops.

The clinical manifestations in human gnasthostomiasis are caused by migration of the immature worms (L3s). Migration in the subcutaneous tissues causes intermittent, migratory, painful, pruritic swellings (cutaneous larva migrans). Migration in other tissues (visceral larva migrans), can result in cough, hematuria, ocular involvement, with the most serious manifestations being eosinophilic meningitis with myeloencephalitis. High eosinophilia is present. Gnathostoma spinigerum is found in Asia, especially Thailand and Japan (CDC, 1999b).

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Cestodes

Diphyllobothrium latum (the fish or broad tapeworm), is the largest human tapeworm. Several other Diphyllobothrium species have been reported to infect humans, but less frequently; they include D. pacificum, D. cordatum, D. ursi, D. dendriticum, D. lanceolatum, D. dalliae, and D. yonagoensis.

The adult D. latum tapeworm resides in the small intestine where it attaches to the mucosa. It can reach more than 10 m in length, with more than 3,000 proglottids. Immature eggs are discharged from the proglottids (up to 1,000,000 eggs per day per worm) and are passed in the feces. Under appropriate conditions, the egg matures (in 11-15 days), yields an oncosphere which develops into a coracidium. After ingestion by a suitable freshwater crustacean (copepod) (first intermediate host) the coracidium develops into a procercoid larva. Following ingestion of the copepod by a suitable freshwater fish (second intermediate host), the procercoid larva migrates into the fish flesh where it develops into a plerocercoid larva (sparganum). When the smaller infected fish is eaten by a larger one, the sparganum may migrate into the flesh of the larger fish. Humans (the optimal definitive host) acquire the infection by eating raw or undercooked infected fish. Eggs appear in the feces 5-6 weeks after infection. In addition to humans, many other mammals can also be infected.

Diphyllobothriasis can be a long lasting infection (decades). Most infections are asymptomatic. Manifestations may include abdominal discomfort, diarrhea, vomiting, weight loss. Vitamin B12 deficiency with pernicious anemia may occur. Massive infections may result in intestinal obstruction. Migration of proglottids can cause cholecystitis or cholangitis.

Diphyllobothriasis occurs in areas where lakes and rivers coexist with human consumption of raw or undercooked freshwater fish. Such areas are found in the Northern Hemisphere (Europe, ex-USSR, North America, Asia), and in Uganda and Chile (CDC, 1999c).

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Trematodes

Clonorchis sinensis is the Chinese or oriental liver fluke. The adult flukes (10-25 mm by 3-5 mm) reside in small and medium sized biliary ducts. Embryonated eggs are discharged in the biliary ducts and in the stool. After ingestion by the suitable snail intermediate host, the eggs release miracidia which go through several developmental stages (sporocysts, rediae, and cercariae). The cercariae are released from the snail and encyst as metacercariae in the skin and flesh of freshwater fish. Infection of humans occur by ingestion of undercooked, salted, pickled, or smoked freshwater fish. After ingestion, the metacercariae excyst in the duodenum and ascend the biliary tract through the ampulla of Vater. Maturation takes approximately 1 month. Adult flukes can survive 20 to 25 years. In addition to humans, carnivorous animals can serve as reservoir hosts.

Most pathologic manifestations result from inflammation and intermittent obstruction of the biliary ducts. In the acute phase, abdominal pain, nausea, diarrhea, and eosinophilia can occur. In long-standing infections, cholangitis, cholelithiasis, pancreatitis, and cholangiocarcinoma can develop, which may be fatal.

Endemic areas are in Asia including Korea, China, Taiwan, Vietnam. Clonorchiasis has been reported in non endemic areas (including the United States). In such cases, the infection is found in Asian immigrants, or following ingestion of imported, undercooked or pickled freshwater fish containing metacercariae (CDC, 1999d).

Opisthorchiasis is caused by Opisthorchis viverrini (Southeast Asian liver fluke) and O. felineus (cat liver fluke). The adult flukes (O. viverrini: 5 mm –10 mm by 1 mm-2 mm; O. felineus:7 mm – 12 mm by 2 mm – 3 mm) reside in the biliary and pancreatic ducts of the mammalian host, where they attach to the mucosa. They deposit fully developed eggs that are passed in the feces. After ingestion by a suitable snail (first intermediate host), the eggs release miracidia, which undergo in the snail several developmental stages (sporocysts, rediae, cercariae). Cercariae are released from the snail and penetrate freshwater fish (second intermediate host), encysting as metacercariae in the muscles or under the scales. The mammalian definitive host (cats, dogs, and various fish-eating mammals including humans) become infected by ingesting undercooked fish containing metacercariae. After ingestion, the metacercariae excyst in the duodenum and ascend through the ampulla of Vater into the biliary ducts, where they attach and develop into adults, which lay eggs after 3-4 weeks.

Most infections are asymptomatic. In mild cases, manifestations include dyspepsia, abdominal pain, diarrhea or constipation. With infections of longer duration, the symptoms can be more severe, and hepatomegaly and malnutrition may be present. In rare cases, cholangitis, cholecystitis, and chlolangiocarcinoma may develop. In addition, infections due to O. felineus may present an acute phase resembling Katayama fever (schistosomiasis), with fever, facial edema, lymphadenopathy, arthralgias, rash, and eosinophilia. Chronic forms of O. felineus infections present the same manifestations as O. viverrini, with in addition involvement of the pancreatic ducts.

O. viverrini is found mainly in northeast Thailand, Laos and Kampuchea. O. felineus is found mainly in Europe and Asia, including the former Soviet Union (CDC, 1999e).

Heterophyes heterophyes, a minute intestinal fluke causes heterophyiasis. Adult H. heterophyes (1.0 mm - 1.7 mm by 0.3 mm - 0.4 mm) reside in the small intestine, where they are attached to the mucosa. They release fully embryonated eggs that are passed in the feces. After ingestion by a suitable snail (first intermediate host), the eggs hatch and release miracidia which undergo several developmental stages in the snail (sporocysts, rediae, and cercariae). The cercariae are released from the snail and encyst as metacercariae in the tissues of a suitable freshwater fish (second intermediate host). The definitive host becomes infected by ingesting undercooked or salted fish containing metacercariae. After ingestion, the metacercariae excyst, attach to the intestinal mucosa, and mature into adults. In addition to humans, various fish-eating animals can be infected by Heterophyes.

The main symptoms are diarrhea and colicky abdominal pain. Migration of the eggs to the heart, resulting in potentially fatal myocardial and valvular damage, has been reported from the Philippines. Migration to other organs (e.g., brain) has also been reported.

H. heterophyes are found in Egypt, the Middle East and Far East (CDC, 1999f).

Metagonimus yokogawai, a minute intestinal fluke (and the smallest human fluke), causes metagonimiasis. Adult M. yokogawai (1.0 mm - 2.5 mm by 0.4 mm - 0.75 mm) reside in the small intestine, where they are attached to the mucosa. They release fully embryonated eggs that are passed in the feces. After ingestion by a suitable snail (first intermediate host), the eggs hatch and release miracidia which undergo several developmental stages in the snail (sporocysts, rediae, and cercariae). The cercariae are released from the snail and encyst as metacercariae in the tissues of a suitable freshwater fish (second intermediate host). The definitive host becomes infected by ingesting undercooked fish containing metacercariae. After ingestion, the metacercariae excyst, attach to the intestinal mucosa, and mature into adults. In addition to humans, fish-eating mammals and birds can also be infected.

The main symptoms are diarrhea and colicky abdominal pain. Migration of the eggs to extra-intestinal sites (heart, brain) can occur, with resulting symptoms.

M. yokogawai are found mostly in the Far East, as well as Siberia, Manchuria, the Balkan states, Israel and Spain (CDC, 1999g).

Paragonimiasis is an infection in animals and humans caused by more than 30 species of trematodes (flukes) of the genus Paragonimus. Among the more than 10 species reported to infect humans, the most common is P. westermani, the oriental lung fluke. Human infection with P. westermani occurs by eating inadequately cooked or pickled crab or crayfish that harbor metacercariae of the parasite. The metacercariae excyst in the duodenum, penetrate through the intestinal wall into the peritoneal cavity, then through the abdominal wall and diaphragm into the lungs, where they become encapsulated and develop into adults (7.5-12 mm by 4-6 mm). Time from infection to oviposition is 65 to 90 days. The eggs are excreted unembryonated in the sputum, or alternately they are swallowed and passed with the stool. In the external environment, the eggs embryonate, hatch and yield miracidia which enter the first intermediate host, a snail. Cercariae emerge from the snail and invade the second intermediate host, a crustacean (crab of crayfish) where they encyst and become metacercariae. Ingestion of the metacercariae closes the cycle. Infections may persist for 20 years in humans, and occasionally other sites than the lungs are involved. Infection occurs also in many animal species.

The acute phase (invasion and migration) may be marked by diarrhea, abdominal pain, fever, cough, urticaria, hepatosplenomegaly, pulmonary abnormalities, and eosinophilia. During the chronic phase, pulmonary manifestations include cough, expectoration of discolored sputum, hemoptysis, and chest radiographic abnormalities. Extrapulmonary locations of the adult worms result in more severe manifestations, especially when the brain is involved.

While P. westermani occurs in the Far East, other species of Paragonimus are encountered in Asia, the Americas, and Africa (CDC, 1999h).

Nanophyetus salmincola or N. schikhobalowi are the names, respectively, of the North American and Russian troglotrematoid trematodes (or flukes). Nanophyetiasis is the name of the human disease caused by these flukes. At least one newspaper referred to the disease as "fish flu." N. salmincola is responsible for the transmission of Neorickettsia helminthoeca, which causes an illness in dogs that may be serious or even fatal.

Knowledge of nanophyetiasis is limited. The first reported cases are characterized by an increase of bowel movements or diarrhea, usually accompanied by increased numbers of circulating eosinophils, abdominal discomfort and nausea. A few patients reported weight loss and fatigue, and some were asymptomatic. The rickettsia, though fatal to 80% of untreated dogs, is not known to infect humans.

There have been no reported outbreaks of nanophyetiasis in North America; the only scientific reports are of 20 individual cases referred to in one Oregon clinic. A report in the popular press indicates that the frequency is significantly higher. It is significant that two cases occurred in New Orleans well outside the endemic area. In Russia's endemic area the infection rate is reported to be greater than 90% and the size of the endemic area is growing.

Nanophyetiasis is transmitted by the larval stage (metacercaria) of a worm that encysts in the flesh of freshwater fishes. In anadromous fish, the parasite's cysts can survive the period spent at sea. Although the metacercaria encysts in many species of fish, North American cases were all associated with salmonids. Raw, underprocessed, and smoked salmon and steelhead were implicated in the cases to date (FDA, 1999).

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Control Measures

The process of heating raw fish sufficiently to kill bacterial pathogens is also sufficient to kill parasites.

Freezing (-20ºC [-4ºF] or below [internal or external] for 7 d or –35ºC [-31ºF] or below [internal] for 15 h) of fish intended for raw consumption also kills parasites. The Food Code recommends these freezing conditions to retailers who provide fish intended for raw consumption.

Brining and pickling may reduce the parasite hazard in a fish, but they do not eliminate it, nor do they minimize it to an acceptable level. Nematode larvae have been shown to survive 28 d in 80º salinometer brine (21% salt by weight).

Trimming away the belly flaps of fish or candling and physically removing parasites are effective methods for reducing the numbers of parasites. However, they do not completely eliminate the hazard, nor do they minimize it to an acceptable level (FDA, 1998).

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FDA Guidelines

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Published Process Studies

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Controlling nematodes

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Controlling trematodes

Table 16-4. Freezing conditions to kill trematodes (Heterophyes spp.) in frozen mullet (Hamed and Elias, 1970).

Maximum External Temperature		Minimum Time
(ºC)	(ºF)	(h)
-10	14	30
-20	-4	30

Table 16-5. Freezing conditions to kill trematodes (Clonorchis sinensis) in frozen cyprinids (Pseudorasbora parva) (Fan, 1998).

Maximum External Temperature			Infective After	Not Infective After
(ºC)	(ºF)	(d)		(d)
-12	10.4	18		20
-20	-4	7		N/D

N/D = Not determined

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Analytical Procedures

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Parasitic Animals in Foods (Bier et al., 1998)

Humans unknowingly consume microscopic and small macroscopic animals with their food. The intestinal tract is inhospitable to most of these organisms, which are either digested or evacuated in the feces. However, some obligate or facultative parasites may become established in the human body. Although a number of parasites produce no symptoms and are not associated with disease, others may cause mild, moderate, or severely acute illness and even permanent damage. The following methods are used to examine foods and food-contact materials for the presence of parasites. For the most part, these techniques are labor-intensive and tedious; work continues to refine them and to develop additional techniques and rapid methods. Several alternative ways to examine fish and shellfish are presented. However, candling is the only method currently used for regulatory purposes with finfish.

1. Digestion to Select Mammalian Parasites in Edible Flesh

By mimicking the chemical and temperature conditions of the mammalian stomach, this method frees parasites from the surrounding flesh and reduces the background of nonparasitic organisms.

1. Equipment and materials
1. Balance, at least 250 g capacity
2. Stirrer or rotating incubator shaker
3. Water bath, 37 ± 0.5ºC
4. Beakers, 100 and 1500 ml
5. Sedimentation cone and support, 1 L, plastic with removable plug, e.g., Imhoff cone
6. Tubing, amber gum, 2.4 and 9.5 mm diameter
7. Tubing clamp
8. Microscopes, dissecting and inverted
9. Culture dishes, plastic, various sizes
10. Sieve No. 18 (U.S. standard sieve series), 1 mm mesh, 204 mm diameter, 51 mm high; other sizes optional
11. Tray, rectangular, polypropylene, about 325 x 260 x 75 mm
12. Cylinder, graduated, 1 L
13. pH meter
14. Pasteur pipets, or polypropylene eyedroppers
15. Rubber bulb, about 2 ml capacity
16. Spoon
17. Spatula
18. Optional materials: blender, meat grinder, food processor, negative pressure hood, foil, plastic wrap, tweezers, and dissecting needles
2. Reagents
1. Physiological saline (R63)
2. Pepsin, laboratory grade
3. pH reference solutions
4. HCl, concentrated
5. Optional reagents: papain, ethanol, glacial acetic acid, glycerine, lactophenol, phenol, formalin, Lugol's iodine (R40), ether
3. Sampling and sample preparation

From a sample weighing 1 kg, take a subsample (100 g) of beef, pork, or poultry, or 250 g of fish. Subsamples of most mammalian meat, poultry, or fish require no further preparation. They may be torn or separated into 5 or more pieces to increase the surface area. Samples with relatively large amounts of connective tissue are not readily digested; snail meat, for example, is digested very poorly. The following methods improve digestion. A 100 g sample is blended in 750 ml saline. Ten intermittent, instantaneous bursts in a blender will destroy some macroscopic organisms but usually will not affect microscopic organisms. A meat grinder is less destructive, although not suitable for some foods such as snails. Least destructive is initial digestion with papain followed by pepsin digestion.

CAUTION: Pathogens that are easily disseminated may be contained in samples and will be liberated by digestion. Of special concern are macroscopic tapeworm cysts and microscopic cysts of protozoa. When the presence of such pathogens is suspected, carry out the digestion and subsequent sample handling in a negative pressure hood until the suspect digest is placed in a safely closed dish. Handle all utensils as if contaminated, and autoclave or incinerate after use.

4. Digestion, sedimentation, and examination

Adjust incubator-shaker or water bath to 37 ± 0.5ºC. Prepare digestion fluid in 1500 ml beaker by dissolving 15 g pepsin in 750 ml saline, add sample, and adjust to pH 2 with concentrated HCl (about 3 ml). Place in incubator or water bath and stir (about 100 rpm) after equilibration for about 15 min; check and adjust pH again. Cover beaker with aluminum foil (if using stirrer, pierce hole for stirring rod) and continue incubating until digestion is complete. The time required for digestion will vary but should not exceed 24 h.

Carefully pour beaker contents through sieve into tray. Rinse remains with 250 ml saline and add to digest. Examine rinsed contents of sieve and record results. Larger parasites will remain on sieve. Replace plug of sedimentation tube with rubber tubing and clamp folded tubing. Carefully transfer contents of tray to sedimentation cone. Transfer undigested sample or parasites to a petri dish, using spoon, tweezers, or dissecting needle.

After 1 h of sedimentation, remove bottom 50 ml of sediment by releasing clamp and collecting in 100 ml beaker. Transfer fluid to petri dish(es) with eyedropper. (Digests vary in their clarity; if digest is dense, dilute with saline until it is translucent.) Cover dish and examine macroscopically for parasites; then examine with dissecting microscope and finally with inverted microscope (a contrasting background can be helpful). Count, tentatively identify, and record observations. Count total number of organisms and differentiate those that are living (motile) and dead (nonmotile), if possible. Examine complete contents of beaker. Light infections may require repeated sampling to detect parasites.

5. Interpretation and further identification

Further information about recovered organisms is usually required, both to classify them and to decide whether the criterion of movement is valid for determining viability. For example, the eggs of Ascaris must be "embryonated," i.e., allowed to incubate so that moving embryos develop inside. Cysts of some protozoa must be excysted to detect motion; those of Toxoplasma gondii can be judged viable only by the outcome of experimental inoculation into the peritoneal cavity of mice. Brief summaries of fixation and staining methods for frequently recovered parasites are given below and in the following references: invertebrates in general parasitology (Barnes, 1987); animals' parasites (Olsen, 1986); medical aspects of parasitology (Noble and Noble, 1982); food parasitology: methods, references, expert consultants (Fayer et al., 1992; Jackson, 1983); immunology and serology of parasitic diseases (Jackson et al., 1969); protozoa (Kudo, 1977); nematodes (Chitwood and Chitwood, 1974;Yorke and Maplestone, 1969); trematodes (Anderson et al., 1974-83; Schell, 1985); cestodes (Schmidt, 19985); arthropods (FDA, 1981).

6. Fixation and staining

Protozoan cysts and helminth eggs. Fix and stain fresh material with Lugol's iodine solution (R40) or use fluorescent antibody stain (if available) on formalin-fixed material.

Nematodes. Fix in glacial acetic acid overnight and store in 70% ethanol with 10% glycerin. Study nematode morphology in temporary mounts by removing from alcohol and clearing in glycerin, lactophenol, or phenol ethanol. Before returning to storage, rinse away clearing fluid(s) with 70% ethanol. Sectioning and staining may be necessary for detailed identification of nematodes.

Trematodes and cestodes. Before fixation, relax both trematode and cestode flatworms in cold distilled water for 10 min. Fix flukes (trematodes) in hot (60ºC) 10% formalin. Fix tapeworms (cestodes) by adding 10X volume of 70ºC fixative to the relaxing fluid, or dip them in 70ºC water repeatedly; then fix in a mixture of ethanol, glacial acetic acid, and formalin (85:10:5) overnight. Store in 70% ethanol. Flatworms are usually stained and mounted as permanent slides, but some may require sectioning and staining for detailed identification.

Acanthocephala. Place in water to evert the proboscis. (Some proboscises may evert almost immediately; others require several h. Do not extend over 8 h.) Fix in steaming 70% ethanol with a few drops of glacial acetic acid. Store in fixative or 70% ethanol. Acanthocephala may be stained and mounted as permanent slides or, like the nematodes, cleared in phenol or glycerol.

Arthropoda. Fix fleas, lice, mites, copepods, fly larvae, and other parasitic or food-inhabiting arthropods in hot water. Store in 70% ethanol.

Keep parasite specimens in tightly capped vials with identifying data (anatomic location of source in host, geographic origin, date of sample collection, date of parasite collection, collector's name, presumed identification) written in indelible pencil or ink on slip of hardened paper. Place paper in liquid-filled vial with parasite. Assistance in identification may be obtained from J.W. Bier, Office of Seafood, FDA, 1110 Vermont Ave., N.W., Washington, DC 20005; Ann M. Adams, FDA, P.O. Box 3012, Bothell, WA 98041-3012; or Marie Chaput, FDA, 109 Holton St., Winchester, MA 01890. Please send a minimum of three whole parasites of each type found and all head/tail fragments as well as all pertinent information to the nearest of the three individuals named above.

7. Viability determination

The major criterion for the viability of helminths is spontaneous movement. Observe organisms for 10 min to see if spontaneous movement occurs. If autonomous movement is not observed, touch with a dissecting needle and observe to see if movement has been stimulated. Allow specimens from salted products to equilibrate in at least 20X volumes of physiological saline for 3 h before viability determination; osmotic pressure may cause apparent movement. Culturable protozoa should be cultured in vitro to determine viability. If culture is not feasible, dye exclusion is the method of choice for viability determination.

2. Candling to Detect Parasites in Finfish

The following procedures are used to determine parasites in finfish. The candling procedure is applicable to fresh or frozen fish with white flesh processed as fillets, loins, steaks, chunks, or minced fish. The ultraviolet (UV) light procedure is for fish with dark flesh and for breading removed from fish portions.

NOTE: This method is not applicable to dried fish or fish in the round.

8. Equipment and materials

Sharp knife

Candling table. Rigid framework to hold light source below rigid working surface of white, translucent acrylic plastic or other suitable material with 45-60% translucency. Length and width of working surface should be large enough to permit examination of entire fillet, e.g., 30 x 60 cm sheet, 5-7 mm thick.

Light source. "Cool white" with color temperature of 4200 K. At least two 20-watt fluorescent tubes are recommended. Tubes and their electrical connections should be constructed to prevent overheating of light source. Average light intensity above working surface should be 1500-1800 lux, as measured 30 cm above center of acrylic sheet. Distribution of illumination should be in ratio of 3:1:0.1, i.e., brightness directly above light source should be 3 times greater than that of outer field, and brightness of outer limit of visual field should be not more than 0.1 that of inner field. Illumination in examining room should be low enough not to interfere with detection of parasites, but not so dim as to cause excessive eye fatigue.

9. Reagents (for preservation reagents, see section I. B-5, above)
10. Sample preparation

Weigh entire sample and record weight on analytical reporting form.

Fillets. If fillets are large (200 g or larger), use 1 fillet for each of the 15 subsamples. If fillets are small (less than 200 g), randomly select fillets to prepare 15 subsamples of approximately 200 g each. Record actual weight analyzed for each subsample. If fillets are more than 30 mm thick, cut with a sharp knife into 2 pieces of approximately equal thickness (not to exceed 30 mm per fillet). Examine both pieces as described below. If fillets have a thickness of 20 mm or less, examine whole.

Fish blocks. Analyze 15 subsamples randomly selected from 2 thawed and drained blocks. Prepare the subsamples as described for fillets, above. Note separately any parasites observed in minced fish added to block around subsamples.

Steaks, loins, chunks. Prepare as for fillets.

Minced fish. If frozen in blocks, analyze 15 subsamples randomly selected from 2 thawed and drained blocks. Prepare subsamples as described for fillets, above. Select portions from different parts of block. If not in blocks, analyze 15-200 g portions. Do not further shred or chop minced fish.

Breaded fish portions. Thaw frozen products at room temperature in a beaker of appropriate size. After thawing, pour hot (50ºC) solution of 2% sodium lauryl sulfate in water over fish in increments of 100 ml per 300 g of product. Stir with glass rod for 1 min. Let stand for at least 10 min or until breading separates from flesh. Transfer individual portions to No. 10 sieve nested over No 40 sieve. Wash breading through No. 10 sieve with gentle stream of warm tap water. Periodically examine No. 40 sieve containing the breading, using UV light. Parasites will appear fluorescent under this light. Note any parasites detected and record on the analytical reporting form. Discard breading by backflushing the No. 40 sieve with tap water. Examine fish portions by candling, using white light. If the flesh is pigmented, use UV light.

11. Examination

Parasites near the surface will appear red, tan, cream-colored, or chalky white. Parasites deeper in the flesh will appear as shadows. Remove representative types of parasites or other defects found. Record general location, size, identification, and other observations as outlined below. For minced fish, spread portion on light table to depth of 20-30 mm for examination. Select representative parasites for descriptive analysis.

12. Ultraviolet examination of dark-fleshed fish

Visually examine each portion (de-breaded or de-skinned, as necessary) on both sides under a desk lamp or similar light source. A magnifying desk lamp may be used. Report findings as described below. Conduct UV examination in darkened room. Examine each portion on both sides with reflected longwave UV light (366 nm wavelength). Parasites should fluoresce blue or green under light of this wavelength. Fish bones and connective tissues, which also fluoresce blue, may be differentiated by their regular distribution and shape. Bone fragments will be rigid when probed (FDA, 1984).

CAUTION: Never expose unprotected eyes to UV light from any source either direct or reflected. Always wear appropriate eye protection such as goggles with uranium oxide lenses, welder's goggle, etc., when such radiations are present and unshielded. Keep skin exposure to UV radiations to a minimum.

13. Parasite identification. Fix parasites as described in section I. F, above.
3. Compression Candling: Detection of Parasites in Molluscs and other Translucent Foods

Parasites may be detected visually in such translucent foods as white-fleshed fish and shellfish by observing the outline of the organism or its capsule in transmitted light. The method described was developed to examine the viscera and muscle of the surf clam Spisula solidissima for the presence of Sulcascaris sp. nematodes, but is also applicable to other foods and parasites. However, not all parasites are detected (Payne et al., 1980), probably because they are obscured by the shadows produced by connective tissue. The method was compared with two other visual methods for detecting nematodes in the calico scallop, Argopectin gibbus. Compression candling detected more nematodes and yielded fewer false positives than the other two methods.

14. Equipment and materials
0. Hinged Plexiglas plates 305 x 305 mm. To construct plates, attach two 305 mm (about 12 inch [30.5 cm]) squares of 3/8 inch (1 cm) Plexiglas stacked plates to a piano hinge so that they are separated by 3 mm with 6-32 x 5/8 inch (3.5 mm-0.35 mm x 16 mm) machine screws. If proper size piano hinge is not available, a nominal 1 inch (2.5 cm) hinge can be retapped to give proper spacing. Attach a 3 mm spacer to each end of the surface of 1 plate at the edge opposite the piano hinge.
1. Light box
2. Knife
3. Specimen vials or jars
4. Dissecting needles
5. Petri dish
15. Reagents
0. Physiological saline solution, 0.85% (R63)
1. Glacial acetic acid
2. 70% Ethanol
16. Method
0. Distribute portion of sample on inside of plastic plate. Quantity to be examined at 1 time depends on size and thickness of sample. Samples over 100 g cannot be compressed. Cut cylindrical samples (e.g., scallops) in half longitudinally to facilitate compression.
1. Close plate and squeeze outer edges firmly.
2. Examine each side of plate for parasites by placing on light table. Parasites in flesh appear as shadows.
3. Record number of parasites. To confirm that parasites are present, mark plate with wax pencil, open, and check by dissection.
4. Fix representative sample to confirm identity (see section I. F, above).

4. Mechanical Disruption and Sedimentation for Detection of Larval Parasites in Fish Flesh

This method detects larval anisakids in the flesh of fillets. It is not applicable to fish that have been treated with salt without being de-boned, such as pickled herring, and, in general, may not be applicable to species such as herring. Subsamples should not exceed 200 g in the food processor, but may be pooled for final analysis.

17. Materials
0. Food processor; Cuisinart Model DLC 10, Moulinex Model 663, or equivalent.
1. Glass tray, 350 x 25 x 60 mm
2. Beaker, 1000 ml
3. White fluorescent lamp
4. UV lamp, <365 nm, or similar, light box
5. Appropriate eye protection
6. Glass rod
7. Forceps
8. Vials or jars
9. Fixative
18. Method

Fillet and skin fish before weighing; then place in food processor with plastic dough hook in place. Add 35ºC water equal to twice the weight of the fish. Activate food processor intermittently until flesh is dissociated (1-2 min). Pour into beaker and wait 30-60 s before decanting all but 100 ml of the supernatant fluid. Add water and stir; then wait 30-60 s, and decant to 100 ml again (2X).

*Place about 25 ml of sediment in glass tray; dilute until quite translucent or until depth of 10 mm is reached (about 375 ml). Examine, collect, or count parasites. Agitation of sediment with forceps may aid in recovery, and forceps will be useful in collecting parasites. Record parasite movement. Collect and fix a representative portion of the parasites present for identification (see section I. F, above). Examine under high intensity (>500 µW/cm²) shortwave (about 365 µm) light. Parasites will fluoresce blue or yellow-green. Count and record. Repeat from *, above, until sample is complete.

5. Concentration of Helminths and Protozoa from Vegetables

Vegetables may become contaminated with parasitic organisms through contact with animal or human fecal material or through application of sewage-derived fertilizer to croplands (Rude et al., 1984). The method outlined below can be used to examine fresh vegetables for parasites. A similar method recovered Cryptosporidium sp. from 1% of water samples examined. Recovery from vegetables would be estimated at 1% or less. (A sample consists of five 1-kg subsamples.)

19. Equipment and materials
0. Balance
1. Polypropylene beakers, 1 L
2. Sonic bath, about 2 L capacity
3. Centrifuge, large capacity, low speed with swinging bucket
4. Polypropylene centrifuge tubes, 50 ml
5. Eye dropper, polypropylene
6. Culture dish with 2 mm grid
7. Microscope slides
20. Reagents
0. Lugol's iodine (R40)
1. Sheather's fluid (500 g sucrose, 320 ml deionized water, 6.5 g phenol)
2. Detergent solutions Nos. 1, 2, and 3 No. 1—2.5% formaldehyde, 0.1% sodium dodecyl sulfate (SDS), 0.1% Tween 80 No. 2—1% Tween 80, 1% SDS No. 3—1% Tween 80
3. Fluorescent antibody kit
21. Procedure

Store vegetables in refrigerator before analysis. Separate vegetables into units: tight head type (cabbage), remove outer 3 layers of leaves; loose head type (leaf lettuce), separate individual leaves; root type (carrot), no preparation; floret type (cauliflower), separate into florets of about 50 g. *Pour 1-1.5 liters of detergent solution No. 1 into sonic bath and add vegetables loosely to about 250 g; operate bath for 10 min. Remove vegetables individually and drain well. Repeat from *, above, until subsample is completely sonicated.

Transfer detergent to beaker; then dispense all of the material into 50 ml centrifuge tubes. Centrifuge at 1200 x g for 10 min. Remove supernatant to 1.5-2 ml and consolidate sediment into 1 tube with eyedropper or plastic pipet. Rinse each tube twice with 1.5 ml detergent No. 2, and add to consolidation tube. Rinse and centrifuge sediment twice with detergent No. 2. Dilute to 10 ml with detergent No. 3 and sonicate for 10 min. Add 25 ml Sheather's fluid to clean centrifuge tube and layer on detergent suspension from sonic bath. Centrifuge at 1200 x g for 30 min. Remove 7 ml of fluid from interface and transfer to centrifuge tube; fill tube with detergent; then centrifuge at 1200 x g for 10 min. *Remove supernatant and dilute with detergent No. 3. Then centrifuge for 10 min at 1200 x g. Repeat from *, above, 2 times.

For helminth eggs, transfer sediment to gridded petri plate and add 1 ml Lugol's iodine. Dilute and examine entire plate with inverted microscope. For protozoa, dilute sediment sufficiently with detergent No. 3 to make translucent 100 µl thin smears on cleaned polylysine-coated microscope slide cleaned with acid alcohol.

Let slides air-dry. Add positive and negative control samples to separate well or slide and let air-dry. Follow manufacturer's instructions for fluorescent antibody staining. Examine each slide at 200-300X with fluorescent microscope. Record results. If sample is positive, calculate number of cysts present per kg of food specimen by measuring remaining suspension and estimating number. If sample is negative, stain and examine remaining sediment.

Contents

Other analytical procedures

• Parasites in fish muscle: Candling procedure (AOAC, 1995).

Contents

References

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